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MET Module V
Module Summary:
This module gives a solid foundation of theory, principles and clinical practice
to equip the medical equipment technician to maintain laboratory equipment.
The medical laboratory department includes facilities, personnel, and equipment
within the hospital or public/private location. Personal include pathologists,
doctors, clinical specialists, lab technologist and lab technicians. The director of
these facilities is usually a physician.
Why is the laboratory area so important to a hospital or medical facility?
The laboratory plays a key role to aid medical staff in the diagnosis, treatment and
prevention of disease.
Why do medical equipment technicians need a solid background and
understanding of this area?
Medical equipment technicians are the personal who are assisting with the
installation, care and maintenance of laboratory equipment.
Equipment contained in the laboratory includes glassware, centrifuges, suction
devices, and sophisticated instrumentation such as colorimeters,
spectrophotometers, blood cell and gas analyzers, chromatographs, automated
chemistry analyzers, and computer-based record and operation systems.
Please note: students should have completed Module 3 anatomy, physiology and
medical terminology before beginning this course.
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MET Module V
Title page
Intro and Summary of Course
Lab basic theory, blood, fluid, cells and chemistry
Cell Counter
Chemistry Analysis and Instrumentation
Colorimeter, Photometer, Spectrophotometer
Flame Photometer/Fluorometry
Electrochemical analysis
pH and Blood Gas Analyzers
Auto analyzer
Reflectance methodology / DT60/Ortho Clinical
Quality Control
Fibrometer / Coagulation
Hot Air Ovens / Incubators
Reverse Osmosis
Lab Safety
Healthcare Biowaste Management
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1. Know why medical equipment technicians must understand the basics of
laboratory theory.
2. List five working areas of the laboratory.
3. Be able to state the purpose of blood and it’s major purposes
4. Be able to list the components and describe the composition of blood. This
includes red blood cells, white blood cells (five types), platelets, and plasma
5. Understand how blood is separated for testing – know the major difference
between plasma and serum
6. Be able to list and describe blood tests; know the main tests done by cell
counting and performed in hematology; know the difference of tests done in
chemistry. This includes basic chemistry principles, fluid and electrolyte
7. State and understand the basic chemistry measuring principles of:
 Optical – absorption/transmission
Be able to label the wavelength spectrum
 Electrochemical
 Chromatography
 Radioimmunological
8. Be able to give an example of an instrument that use the principles of:
optical measuring – absorption, emission and reflectance
electrochemical measuring
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9. Be able to state and discuss the:
 purpose
 uses
 principles of operation – including major components
 maintenance of instrumentation – including most common
faults found
o example: colorimeter most common fault is a burnt out
light source or bulb
Instrument List: blood cell counter
flame photometer
pH meter/blood gas analyzer (pH, PO2 and PCO2)
fibrometer/coagulation instrument
incubators/hot air oven
refrigeration system
chromatograph instrument
water treatment system/distiller
laboratory support devices such as mixers, water baths, steam
10. Understand the principle and use of pipettes. Be able to list and describe at
least three types.
11. Describe the calibration procedure of pipettes
12. Understand the purpose of Quality Control and why it is run in the laboratory.
13. Define accuracy and precision and be able to identify in examples.
14. Understand the importance of lab safely:
 be able to list and describe personal protective devices,
 understand why hand washing is so critical
 list safety measures when handling gas cylinders, nitrogen and biowaste
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Medical Laboratory Department
The medical laboratory department includes facilities, personnel, and equipment
within the hospital or public/private location. The facilities must include a clean,
safe surrounding with a special area for sterilization of contaminated blood/urine
samples and equipment. Since high-volume blood testing occurs in this
department, sufficient storage and cleaning areas must be designated. In such a
situation, the chance of error (misreading or patient record mix-up) is high.
Medical laboratory personnel includes equipment operators (medical
technologists), supervisors, and physicians. The director of these facilities is
usually a physician.
Equipment contained in the laboratory includes glassware, centrifuges, suction
devices, and sophisticated instrumentation such as colorimeters,
spectrophotometers, blood/cell and gas analyzers, chromatographs, autoanalyzers,
and computer-based record and operation systems.
Record keeping is extremely important. This information is used by physicians as
an aid in diagnosing disease and imbalanced physiological states. Standard cards
with printouts of RBC/WBC count, Ht, MCV, MCHC, and blood chemistry are
presented by most clinical instrumentation.
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A. Self-Evaluation Questions. These questions test your prior knowledge of the
material in this chapter. Look for the answers as you read the text. After you have
finished studying the chapter, try answering these questions and those at the end of
the chapter.
1. Define blood and state its purposes.
2. Name the components of blood (cells and plasma).
3. What is the difference between blood cell tests and blood chemistry analysis?
4. Why is the medical laboratory department of critical importance in diagnosing
and treating the patient?
5. Describe the principle of operation and maintenance of the colorimeter, flame
photometer, spectrophotometer, blood cell analyzer, blood gas analyzer (pH, PO2
and PCO2), chromatograph, and autoanalyzer.
B. Blood (Purpose and Components)
Blood is the fluid that circulates through the heart, arteries, veins and capillaries
carrying nourishment, electrolytes, hormones, vitamins, antibodies, heat, and
oxygen to body tissues and taking away waste matter and carbon dioxide.
Whole blood, as shown in Figure 1, is composed of cells and plasma (fluid
containing dissolved and suspended substances). The blood cell portion consists of
the following elements:
1. Red blood cells (RBC) or erythrocytes – These are concaved discshaped cells (8μ length, 3 μ width) that contain no nucleus and live about
120 days before being replaced by the bone marrow. Their number is 4.5
to 5.5 X 106 cells/mm3. Internally, each RBC contains four iron atoms in
a structure known as the hemoglobin molecule. Oxygen from the lung
alveoli enters the bloodstream and chemically combines with hemoglobin
to form oxyhemoglobin. RBCs transport oxygen to the tissues and pick up
carbon dioxide to form carbaminohemoglobin.
2. White blood cells (WBC) or leukocytes – these are amoeba-like cells (10μin
diameter) that contain a nucleus and live from 13 to 20 days. Their number
is 6 to 10 x 103 cells mm3. They are also present in the lymph fluid and
engulf invading bacteria and foreign substances to destroy the invaders’
effect. For example, bacteria invading the leg are encapsulated by WBCs in
the lymph fluid, transported to the interior vena cava, circulated through the
right atrium/ventricle through the lungs to the left atrium/ventricle, and
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pumped to the kidneys, where they are extracted in the urine. They are then
excreted from the body as harmless cell fragments. Specific antibodies are
also produced to kill the invaders’ antigen (toxin).
3. Platelets – These are cell fragments (3 μ in diameter) that contain no
nucleus. Their number is 200 to 800 x 10 3 cells/mm3. These form a repair
substance that initiates blood coagulation and clotting. A protein, thrombin,
also acts on fibrinogen (soluble protein formed in the liver) to generate
insoluble fibrin. Fibrin deposits as fine threads to form the framework of the
blood clot. Platelets cling to intersections of fibrin threads. As fibrinogen is
used up, serum is secreted. Serum will not clot, as it contains no fibrinogen.
Blood plasma consists of the following elements:
1. Plasma proteins– organic repair substances. These are albumins
(synthesized in the liver) that help regulate plasma/tissue cell osmotic
pressure. Fibrinogen and prothrombin is used in the clotting process.
Globulin substances (alpha, beta, and gamma) are catalysts and aid in the
immunizing (disease protection) process.
2. Plasma nutriments – energy storing substances. These are glucose (blood
sugar), lipids (fats) and amino acids (make up proteins for tissue growth).
3 Regulatory and protective substances – These are enzymes
(catalysts for digestion and cell metabolism), hormones
(stimulatory/inhibitory function to target organs) and antibodies
(providing immunity against infection.
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4. Plasma electrolytes – acid-base and nerve impulse transmission
substances. These are inorganic salts (metal and nonmetal combination) and
pure chemical substances (Na+, K+, Cl+).
5. Metabolic waste substances – These include urea and uric acid waste from
the kidneys and carbon dioxide waste from cellular metabolism.
Figure I: Whole blood composed of cells and plasma. RBC-red blood cells, 4.5 – 5.5 x 106 cells/mm3 , 8 x 3 μ ,
no nucleus, 120 day life. WBC-white blood cells, 6-10 x 103 cells/mm3 10μ diameter, nucleus, 15 day life. Pplatelet cells, 200-800x103 cells/mm3, 3μ diameter, no nucleus. F-fibrinogen protein. Plasma – fluid portion,
inorganic and organic substances dissolved in H2O. Whole blood = RBC + WBC + P + F + plasma. Plasma =
whole blood – (RBC + WBC + P). Serum = plasma – fibrinogen.
Figure 2 shows blood that has been spun in a centrifuge (motor-driven mechanical
device that generates a circular motion). A centrifuge is shown in Figure 3. Since
the RBCs are the heaviest, they sink to the bottom and form 45 percent of the total
by volume. Plasma occupies 55 percent and contains substances as indicate. Blood
plasma does contain some dissolved oxygen, but 97 percent of the transported
oxygen is carried in the RBC hemoglobin molecules. During its passage through
the body, blood hemoglobin still remains 70 percent oxygen saturated. The total
carbon dioxide carried by the blood is 30 percent in the RBCs and 60 percent in the
blood plasma.
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Figure 2: Blood that has been spun in a centrifuge.
The body functions as a biological machine that receives life-giving input
substances of oxygen and food nutriments and gives off waste substances.
All living creatures typically display the characteristics or organization (life
process control), irritability (response to change), contractility (movement),
nutrition (ingestion and digestion of food), metabolism/growth (liberation of stored
chemical energy), respiration (intake of O2 and ventilation of CO2), excretion
(elimination of waste), and reproduction (generation of a new structure).
Metabolism is the sum total of all chemical and biochemical processes in the body.
It involves catabolism (breaking down of complex protein/sugar substances to
simpler ones) and anabolism (building up of complex substances for body use).
Waste products from the digestive process are eliminated in the feces. Toxic
substances that result from metabolic processes are removed from the blood by the
kidneys and excreted in the urine.
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Figure 3: A refrigerated centrifuge
The purpose of medical laboratory instrumentation is to provide a means of
measuring required substances and metabolic waste products in urine and blood.
C. Blood Tests (Cells and Chemistry)
Blood cell tests include the following elements:
1. Red blood cell count (RBC) – accomplished manually (diluting a blood
sample 100 to 2 and counting the cells per mm by use of a microscope) or
automatically (blood cell counting analyzers).
2. White blood cell count (WBC) – accomplished manually (10 to 1
dilution) or automatically (blood cell analyzer).
3. Platelet count – accomplished automatically by a blood cell
4. Hematocrit (Ht) – percentage of total blood volume that is
solid (WBC volume is negligible). This is measured by spinning a blood
sample in a test tube and optically observing the percentage of packed
RBCs (see figure 2). It normally ranges from 45 to 55 percent.
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5. Mean cell (corpuscular) volume (MCV) – average volume of an RBC
measured by a value based on the RBC count (number per mm3). This
volume is measured in femtoliters (10-15 l).
MCV = Ht
6. Mean cell hemoglobin (MCH) – the proportional mass of RBC/100ml the
total number of RBCs is expressed as:
hemoglobiningrams / 100ml
This value is indicated in picograms (10-12 g)
7. Mean cell hemoglobin concentration (MCHC) – hemoglobin color
concentration measured by lysing RBCs (breaking their membranes) to
release hemoglobin. Acid hematin or cyanmethemoglobin can be
generated by hemoglobin chemical reaction. The resultant value is
measured by a colorimeter and normally indicates 32 to 36 percent color
hemoglobiningrams / 100ml
Blood chemical tests check for amounts of acidity – pH normally 7.36 to
7.44 (blood is normally slightly alkaline); glucose – lactic acid, lactose
sugar; non-protein nitrogen substances – amino acids, peptides, urea waste;
and uric acid; lipids – fatty acids of cholesterol and triglycerides; proteins –
Plasma albumin, globulins, and fibrinogen; and enzymes and steroids,
among other elements.
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Blood serological tests involve testing for agglutination (clumping) of cells
due to the addition of antigens (bacterial toxins) to blood serum. This occurs
following the reaction of a specific antibody produced by WBCs in response
to the specific invader.
Blood bacteriological tests include growth of blood bacteria in a petri dish
with appropriate nutriments.
Histological tests are studies of small thin tissue samples under the
microscope. Specimens are obtained by cutting tissues with a precision slicer
known as a microtome.
1. Blood is usually collected into tubes that contain a vacuum (defined as
evacuated tubes) to assist in filling the tube. The vacuum tubes have a synthetic
stopper or cap with an elastic septum that can be punctured by a needle
assembly during blood drawing. Blood may also be collected in other types of
containers, such as for cultures.
2. Blood is usually collected by venipuncture using a needle. On some occasions,
blood may be collected by puncturing the skin with a retractable lancet device.
3. Blood is collected and may then be permitted to clot. When a clot forms, a
liquid, called serum, is produced. This serum is used in many tests. A serum
separator tube (SST) contains a gel or filter that forms a barrier to separate
serum from cells during and after centrifugation.
4. An anticoagulant may be used that prevents clotting. The blood may then be
used in liquid form with the cells suspended in it just as they are in the vein. Or
the cells may be removed by centrifugation and the plasma remaining used for
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Figure 4: Blood collected in test tubes
Collection tubes used for blood samples are “color coded” –the color of the stopper
indicates which type it is. Since color standards may change, it is essential to check
the tube label.
The vacuum in the collection tube allows the correct amount of blood to enter the
tube. Do not underfill or overfill the tubes. Blood collected in anticoagulants must
be promptly and thoroughly mixed with the anticoagulant to prevent clotting. Do
not shake the tube. Mix by inverting the tube 8 to 10 times so that the anticoagulant
is thoroughly mixed with the blood. If mixing is not thorough, a partial clot may
form and seriously interfere with the test results (See Figure 5).
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Figure 5: Mixing blood
E. Anticoagulants
There are many coagulation factors involved
in the clotting of blood.
Anticoagulants are substances that prevent blood
from clotting. There are many different kinds of
anticoagulants, which may be powdered or liquid.
The proper anticoagulant must be used for the
specific procedure to be performed. (See Table I).
Blood collected with one anticoagulant may be
suitable for one test or group of tests but not
for others.
F. Four Frequently used anticoagulants
 EDTA (ethylenediaminetraacetic). This anticoagulant is used
primarily when performing blood cell counts.
 Sodium Citrate. This anticoagulant is used primarily when
performing coagulation studies, e.g. prothrombin time.
 Heparin. This anticoagulant is used for a variety of routine and
specialized tests. Note: Green stoppered tubes containing heparin may
contain either sodium heparin or lithium heparin. In general, lithium
heparin is used for chemistry tests. Sodium heparin is used for
specimens requiring the use of viable lymphocytes, such as HLA
testing. The specific type of heparin will be noted on the tube lab
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 Fluoride-oxalate. This anticoagulant is used primarily for glucose
and alcohol determinations. The enzymes, which may destroy glucose
or alcohol, are inhibited by the contents in the tube.
Table 1: Color codes for blood collection tubes
No anticoagulant
Do not mix.
No anticoagulant/serum
Do not mix.
separator tube (SST)
Mix by inverting
Light blue
Mix by inverting
Royal blue
No anticoagulant, heparin
Do not mix (no anticoagulation)
or EDTA (check label)
Mix by inverting (heparin, EDTA)
Lithium heparin, sodium
Mix by inverting.
heparin, or ammonium
heparin (check label)
Fluoride-oxalate or others
Mix by inverting
(check label)
Sodium polyanethole
Mix by inverting
sulfonate (SPS) (for blood
To understand the significance of the composition of the blood and its pathological
changes, it is important to know the most important functions of the blood:
1. To the cells of the body, the blood transports the products of digestion
from the intestines and oxygen from the lungs.
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2. From the cells of the body, the blood transports the products of
combustion to the excretory organs -- carbon dioxide to the lungs, and a
large number of low molecular weight substances to the kidneys,
intestines and skin.
3. The blood is part of the body’s regulatory system and conveys
information – among other things, through hormones from the glands of
internal secretion – so that the function of the other organs is effectively
coordinated and the internal chemical environment will b optimal for the
cells of the body.
4. The blood is part of the body’s system of defense against infection by
microorganisms and foreign substances.
5. The blood has the ability to coagulate, so as to prevent bleeding to death
and to enable tissue wounds to heal.
H. Blood Plasma and Blood Serum
Blood plasma contains both low and high molecular weight substances. The lowmolecular weight substances consist of organic substances and electrolytes, while
the macromolecular substances consist mainly of proteins.
The plasma is obtained by centrifuging a blood sample that has been prevented
from coagulating by adding an anticoagulant. During centrifugation, the heavy
blood cells become packed at the bottom of the centrifuging tube and the plasma
can be poured off. The plasma is a light yellow, almost clear and somewhat
viscous liquid.
The blood serum is obtained by centrifuging a blood sample that has been allowed
to clot. During coagulation a dissolved macromolecular substance, fibrinogen, is
converted to insoluble fibrin. Fibrin consists of invisible fibers that hold the clot
together. Blood serum is thus blood plasma from which the fibrinogen has been
The most important positive ions in the body are sodium, potassium and calcium.
The most important negative ions are phosphate, chloride, and bicarbonate. The
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correct concentrations of ions are necessary for the normal function of cells and the
The ion concentrations vary over wide limits at different places in the body; for
example, most potassium is located intracellularly and most sodium extracellularly.
I. Clinical Chemistry -- Blood
Sodium is thus the most important extracellular-positive ion. It plays a central role
in the body’s electrolyte and fluid balance. Changes in sodium ion concentration
are usually accompanied by changes in the water balance; this is because the body
tends to keep constant both the fluid volume and the sodium ion concentration in
the extracellular space. The responsible regulatory organs are the adrenal glands,
which control the excretion of sodium and water through the kidneys by means of
a hormone (aldosterone).
An increase of the sodium concentration in the blood serum occurs in dehydration,
in certain types of renal damage that cause edema (accumulation of fluid
interstitially) and in adrenocortical hyperfunction, that is, increased secretion of
hormone by the adrenal glands, as occurs in Cushing’s syndrome.
A reduction in the sodium ion concentration occurs in diarrhea (in which the
organism loses sodium), in types of renal damage that cause increased excretion of
sodium and in adrenocortical hypofunction (inadequate output of hormone) as in
Addison’s disease.
Potassium. The correct potassium ion distribution is of vital importance for cell
function – most potassium is located intracellularly. For example, serum must have
a correct potassium concentration for normal nerve cell conduction and for normal
muscle cell contraction.
Of particular importance is the effect on the heart muscle. If the potassium
concentration in the serum exceeds a certain level, there is risk of cardiac arrest
due to prolonged diastole--asystole.
There is an increased serum potassium concentration, hyperkalemia, in several
diseases, among them adrenocortical hypofunction, shock and acidosis and in
certain types of renal damage.
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A depressed potassium level, hypokalemia, is found in adrenocortical
hyperfunction – thus, this situation is opposite to that for sodium.
Calcium has several functions in the body. Besides being a component of the
skeleton, which includes about 1 kg of calcium, it occurs extracellularly in ionic
form. The correct calcium ion concentration is important for nerve and muscle cell
function – calcium is antagonistic to potassium. If too much calcium ion is given in
an intravenous injection, there is a risk of affecting the heart, with consequent
circulatory arrest – just as in the case of increased potassium ion concentration –
though calcium produces fibrillation whereas potassium produces asystole.
Calcium is also of importance for the permeability of cell membranes and for the
clotting of blood.
Phosphate. Phosphorus metabolism is linked with calcium metabolism. This
would be expected since the substance that endows the skeleton with its hardness is
hydroxyapatite – a phosphate of calcium.
The body tends to keep the product of the calcium and phosphate ions in the
plasma constant. This is accomplished by a hormonal regulatory system involving
parathyroid hormone, a hormone from the parathyroid gland, and by excretion via
the kidneys. In increased excretion of parathyroid hormone, hyperparathyroidism,
the serum calcium level is raised and the serum phosphate level lowered. In
hypoparathyroidism, the opposite situation prevails. Hyperparathyroidism results
in decalcification of the skeleton, and often renal damage through the formation of
kidney stones.
Bicarbonate ions are essential for maintaining the correct hydrogen ion
concentration in the body. Bicarbonate ions are part of a buffer system, which acts
according to the Henderson-Hasselbalch equation. Thus, in acidosis and alkalosis,
it is of great importance to determine the bicarbonate ion concentration. This can
be done by the so-called standard bicarbonate test, which is performed in intensive
Chloride. The determination of the serum chloride concentration is of little clinical
interest. The chloride ion level largely follows changes in sodium and bicarbonate
ion concentrations.
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Table 2: Example of electrolytes in the serum
Normal values Examples of diagnostic use
Elevated values
Lowered values
Renal failure,
Potassium in serum
3.6 – 5.1
Calcium in serum
4.3 – 5.3
Phosphate in serum
1.6 – 2.6
bicarbonate in blood
21 – 25
Renal damage,
shock, acidosis
Metabolic alkalosis
Adrenocortical hyper
Reduced absorption
Metabolic acidosis
*mEq/1 = milliequivalent per liter; equivalent = moleweight/valence
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Figure 6: An automatic blood cell counter
Figure 6: An automatic blood cell counter
Since microscope counting is time consuming, most red blood cell counts are
performed on the automatic blood cell counter (Figure 6). The diluted sample is
drawn through a small orifice (100 μm). The column of saline within the orifice
constitutes the greater part of the resistance between the two electrodes. As a single
red blood cell moves through the orifice, the orifice resistance increases, since the
cell has a high resistance. The resulting impulse is detected by the conductivity
meter and counted on an electronic counter to yield the blood cell count.
Erythrocyte sedimentation rate, ESR. If blood mixed with a substance that prevents
clotting is drawn up in a glass tube and left for an hour or so, the blood cells
gradually settle. Over the cells, plasma collects. The height of the plasma column
constitutes a measure of the rate of sedimentation, which is expressed in
millimeters per hour. The normal range of ESR for men is 1-15 mm/h, and for
women, 1-20 mm/h. The ESR is increased in pregnancy and a number of diseases,
such as infections and often in cancer.
The sedimentation rate is an indirect measure of the tendency of the blood cells to
form so-called rouleaux – clumps of cells like a roll of coins. The more the cells
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are packed together, the greater is the formation of rouleaux and thus the higher the
ESR. The increase in rouleaux formation may be due to a raised concentration of
macromolecular substances in the plasma – for example, fibrinogen, certain
globulins, and plasma substitutes (Dextran).
White blood cells
The number of white blood cells and the relative distribution of the different types
of cells changes in pathological processes. Normally there are 3,500 to 9,000 per
μl; of these; neutrophilic un-segmented rod-shaped granulocytes comprise 2 to 5
%; neutrophilic segmented granulocytes, 45 to 70 %; eosinophilic granulocytes,
1to 4 %; basophilic granulocytes, 0 to 1 %; lymphocytes, 20 to 45%; and
monocytes, 4 to 8 %. The determination of the relative numbers of different white
blood cells is known as the differential count. It is performed by examining a
stained smear of blood on a microscope slide.
In infections, the number of white blood cells is increased; however, the relative
distribution varies depending on whether the infection is acute or chronic.
Neutrophilic leukocytes render certain foreign substances harmless, and their
number is raised in acute infections. Lymphocytes transport and produce certain
antibodies, and they are increased in an increased number of eosinophilic
granulocytes, which renders the antigen-antibody complex harmless.
In leukemia, blood cancer, immature forms of leukocytes appear in the blood. They
are liberated prematurely from the bone marrow. At the same time, the number of
leukocytes is often greatly increased. There are different forms of such
pathological conditions – common to them all is the fact that their cause is largely
unknown. During irradiation with high doses of ionizing radiation, similar
pathological conditions arise that cannot be distinguished from leukemia.
The number of white blood cells may also be reduced, leukopenia. This condition
can occur as a result of toxic effects, for example, in hypersensitivity to drugs. The
patient’s resistance to infection is lowered, as would be expected from the role of
the white cells in the specific infection defense mechanism.
I. Methods of Chemical Analysis
A wide variety of analytical methods are used in clinical chemistry. This is due to
the considerable differences in the substances to be analyzed: inorganic
electrolytes, low molecular weight organic substances and macromolecular
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substances. The substances that are more or less routinely analyzed number more
than one hundred. In addition, the concentration range varies widely, from <0.1
mg/l to > 100 g/l. A particular substance may also display large variations in
properties; for example, an enzyme activity in the blood may rise a thousandfold in
a pathological condition.
The analytical methods used should be specific for the substance analyzed; that is,
there should be no interference from any of the innumerable other substances
present in the sample. In addition, the analytical method should be rapid,
economic, and suitable for automation. The analytical methods in most common
use may be classed as optical, electrochemical, and chromatographic. The
radioimmunological method is also used in chemical chemistry.
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Blood Cell
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Figure 6: An automatic blood cell counter
Figure 6: An automatic blood cell counter
Since microscope counting is time consuming, most red blood cell counts are
performed on the automatic blood cell counter (Figure 6). The diluted sample is
drawn through a small orifice (100 μm). The column of saline within the orifice
constitutes the greater part of the resistance between the two electrodes. As a single
red blood cell moves through the orifice, the orifice resistance increases, since the
cell has a high resistance. The resulting impulse is detected by the conductivity
meter and counted on an electronic counter to yield the blood cell count.
Blood Cell Counter
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Knowledge about the number of particles in blood is important data from which a
physician can diagnose disease. These particles are commonly known as red blood
cells (RBC), white blood cells (WBC), and platelets (PLT). They can be
distinguished from each other by virtue of their size and density, as Table 4 shows.
Because of these distinguishing features, it is possible to use a microscope to count
the particles or to use electronic circuitry to do the task automatically. Notice in the
table that the RBCs are more numerous but smaller than the WBCs. Table 4 shows
that most of the particles in blood are RBCs. To determine relative proportion of
blood volume made up by cell particles, known as the hematocrit (ht), a centrifuge
is used (the centrifuge is sometimes called a hematocrit also). The blood sample is
placed in a test tube, which is spun so that the cells are packed on the bottom under
centrifugal force. The ht value equals the height of the packed cells divided by the
height of the blood in the tube. This is typically 45 %.
Table 1: Blood particles
Density (millions/μ l)
Individual size (μ m)
4.26 to 6.2
6.8 to 7.5
0.15 to 0.40
2 to 4
0.004 to 0.011
6 to 18
The function of the red blood cells is to carry hemoglobin to the cells of the body.
It is important to know the mean volume of each red blood cell (MCV) in liters.
MCV is defined as
(in liters/cell)
Where RBC represents the density of red blood cells in cells per liter. A measure
of hemoglobin is made by destroying the red blood cells with an acid and releasing
the red color hemoglobin into solution 1 in a process called lysing. The
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hemoglobin is then separated and weighed. A measure of the mean cell
hemoglobin (MCH) is then computed by the definition
hemoglobin( g / liter )
RBC (liter  1)
The units of MCH are grams per cell. The concentration of hemoglobin in the
blood is then given by the mean cell hemoglobin concentration (MCHC) in grams
per liter:
hemoglobin( g / liter )
A circuit for electronic measurement of the number of cell particles in the blood is
shown in Figure 36. The transducer consists of an orifice through which the sample
is drawn by a vacuum. Since the blood cells have high-resistance membranes, the
cells in the orifice increase its resistance Rout. That is,
Rout = R + ∆ R
where ∆R is the change in resistance due to a cell in the orifice and the value of the
orifice resistance clear of cells is R. ∆R produced by each white blood cell is larger
than that of a red blood cell or a platelet because its size is
greater. The Wheatstone bridge in Figure 36 produces a voltage Vout due to
changes in Rout as follows:
Vout 
( Rout )
( Rout  R
 VBB  (
( Rout  R) 2
2( Rout  R)
where VBB is the bias voltage on the bridge. Inserting equation 8 into 9 gives
Vout 
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4 R  2R
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where ∆ R « R.
Therefore Vout is proportional to ∆R, which is in turn proportional to the size of the
blood cell in the orifice. In Figure 36, Vout is amplified by a
differential amplifier of gain A. An oscilloscope trace of the output is illustrated in
Figure 37.
Figure 36: A circuit for electronic measurement of blood cell count.
Notice that the highest peaks are fewest in number. These are due to the WBCs,
which are correspondingly largest in size and fewest in number according to Table
4. The RBCs are represented by the peaks between threshold T2 and T1. They are
much greater in number than the WBCs but make less resistance change in the
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orifice. In the operation of the instrument, the threshold is first set to zero and the
counter will read N0, the total number of particles per liter,
Figure 37: A blood cell counter display.
WBC + RBC + PLT = N0
where WBC is the number of white blood cells per liter, RBC the number of red
blood cells per liter, and PLT the number of platelets per liter. The threshold is
then set to T1, and the counter will read those signals that exceed the threshold and
give the number N1:
RBC + WBC = N1
Then the threshold is set to T2 and the counter will read just WBC. The
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RBC is then computed from Equation (11) and the PLT from Equation (10).
Example 3:In Figure 36 the orifice resistance, when there are no blood cells of any
type in it, is 1 kΩ. A single RBC in the orifice increases its resistance to 1.01 kΩ.
What will be the output voltage Vout each time an RBC passes into the orifice? The
bias voltage VBB = 10V.
Solution: The orifice resistance R = 1 kΩ. From Equation (7) we have ∆R = 1.01
kΩ –1 kΩ = 10.0 Ω. Then from Equation (9),
Vout =
= 25 m V
Example 4 A blood cell counter threshold is set to zero, and the output display
reads 5.3100 X 1012 liter –1.
The threshold is then set to T1 in Figure 37, and the
output reading becomes 5.8 X 1012 liter –1. The threshold is then set to T2 and the
output reading becomes 0.18 x 10+12
liter –1. Find RBC, WBC, and PLT in units of cells per liter.
Solution The T2 threshold reading is WBC = 0.18 x 10+12. At threshold T1 yields
RBC + WBC = 5.18 x 1012
Therefore RBC = 5.18 x 1012 – 0.18 x 1012 = 5.0 x 1012 cells per liter.
Equation (10) then is used to find the platelets:
PLT = 5.31 x 1012 – 5.0 x 1012 –0.18 x 1012
= 0.13 x 1012 cells per liter
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The counter must be started and stopped over well-timed periods. A vacuum
fluid arrangement for doing this timing is shown in Figure 38. To operate the
mechanism, valves V and F are both initially opened. This draws a flushing
solution through glass tubing into the waste to get rid of any bubbles. Valves F and
V are then closed. A sample of dilute blood is then placed in the beaker and valve
V is opened. The vacuum thus created will cause the sample to enter the orifice.
Simultaneously the mercury will be drawn up into the bulb so that it is confined
between the levels 1 and 1’. Valve V is then closed, cutting off the vacuum pump
from the sample and mercury. The mercury surface then moves from the levels 1
and 1’ to become confined between the levels marked 2 and 2’ in the figure as it
seeks its own level because of the atmospheric pressure at both ends.
Figure38: Glass tubing for a blood cell counter using a mercury timer.
As the mercury surface travels from position 1-1’ to position 2-2’, it activates first
the start switch to begin the counter. When it arrives at the stop switch, the counter
stops. As the mercury falls it also draws a fixed volume of the sample through the
orifice. The counter therefore reads the numbers of particles per unit volume. This
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mechanism controls the rate of flow through the orifice so the digital display can
be calibrated in units of cells per microliter.
Cell Counters
Cell counters are used for hematological measurement. Semi-automated and
automated cell counting has proved to be much more reliable than microscopic cell
counting, because a far greater number of cells can be counted rapidly in a
specimen by an analyzer system, resulting in greater precision. However, the
improvement in precision does not necessarily imply a simultaneous improvement
in accuracy. For the study of pathological cell types, microscopic examination of a
blood smear by an experienced investigator is still most valuable, and in many
instances the method of choice.
A number of different principles are used in cell analyzers:
--impedance measurement
--light scattering
--centrifugation and quantitative buffy-coat analysis
Using these techniques, all major classes of blood cells (erythrocytes, platelets, and
leukocytes) can be identified and even subclasses (granulocytes and lymphocytes)
can be measured. More sophisticated instruments, which combine measurement of
cell size and cell fluorescence,
or cell size determination and immunofluorescence, allow subclassification.
Impedance measurement is most commonly used for cell counting. This principle
of measurement takes advantage of the fact that blood cells are less conductive
than the diluent electrolyte. The principle of the method is explained below.
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1.An electrical potential is applied to two electrodes dipped into an electrolyte
solution, an electric current can be measured owing to the transport of ions from
one electrode to the other. The magnitude of the current will depend on the
concentration of the ions in solution. It will be constant if the transport of ions is
constant (Figure 39a).
2.If the electrodes are separated by an insulator, the flow of electric current will
drop to zero (Figure 39b)
3.The current will reappear if a small aperture is introduced into the
but the magnitude of the current will be small, because the insulation is still
partially effective (Figure 39 c).
4. A small particle such as a blood cell with a conductivity lower than that of the
electrolyte solution, passing through the aperture from one chamber to another,
will temporarily decrease the current because a smaller volume of the electrolyte
solution is able to pass through the aperture at the same time. The current will
regain its original value when the particle has passed through the aperture (Figure
39 d).
When a cell-containing fluid is sucked through the aperture of an insulator
separating two electrode chambers, each change in current (registered as a pulse)
indicates the passage of a particle through the aperture, thus allowing the cells to
be counted. Furthermore, the magnitude of each pulse is proportional to the size of
the particle. Simple cell counters register only the number of pulses above a certain
threshold. More sophisticated counters also register the magnitude of each pulse
and show the distribution of the pulse magnitudes, thus indicating the distribution
of particle sizes in the population of cells. The concentration of cells in the sample
is measured by counting the number of pulses for a known volume of fluid.
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Figure 39: Measurement of impedance as a method of counting cells.
The principle of measurement implies that each measured pulse is attributed to the
passage of a single particle through the aperture of the separating membrane.
Therefore, a blood specimen having a cell concentration of about 5 million cells
per microliter must be appropriately diluted. (in general, 200- or 250- fold) to
ensure that individual cells are counted. Sometimes errors resulting from the
passage of more than one cell still occur in specimens with high cell
concentrations. These errors can be avoided by:
--Further dilution of the blood specimen and a repeat measurement
--Application of a correction factor to the number of cells counted to allow
for coincidence. This factor varies with the cell concentration and may be taken
from a table provided for the instrument by the manufacturer.
Modern instruments have a built-in calculation program to allow for coincidence.
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Most cell counters cannot be calibrated by the user. Therefore, a standard prepared,
or purchased cell suspension can serve only for control of precision but not for
accuracy. The given cell concentration in commercially available control materials
can be used only for measurement with a specified instrument; different results
will be obtained with instruments with a different threshold limit.
The determination of cell size by impedance measurement is influenced by a
number of technical factors, which may vary from one instrument to another; and
also by the type and shape of he particles being measured. For example, the
magnitude of the signal from a discoid cell, e.g., an erythrocyte, is different from
that from a spherical cell, e.g., a leukocyte, even if the cells have the same volume.
Cell counters, using the impedance principle, count the combined total of red blood
cells, which have a cell volume between 70 and 120 fl (femtoliters) and leukocytes,
which have a cell volume between 100 and 350 fl. The results are given in terms of
red blood cell count, ignoring the negligible proportion of leukocytes (usually only
0.2% ---- 10,000 leukocytes compared with 5 million red cells per ml). However, a
considerable error may occur in specimens with a much higher proportion of
leukocytes, e.g., those from patients with pronounced anemia and high leukocyte
For leukocyte counting, erythrocytes are lysed by an ionic or non-ionic detergent in
solution – a procedure that does not lyse leukocytes within a certain time.
However, the size of the leukocytes, and particularly of the polymorphonuclear
cells, may vary. Erroneously low counts may be obtained at high leukocyte
concentrations, when for example two cells pass through the aperture
simultaneously, while being registered only as a single pulse. Such errors can be
eliminated by greater dilution of the specimen, as mentioned above.
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Measurement of platelet concentration is more prone to error, because of their
small cell size (1-10 fl). It is particularly important that the buffer solution used for
the dilution of the blood specimen is free of dust particles; these solutions must be
filtered through a filter of 1μm pore size. Absence of dust is also important for
counting of red and white blood cells, although the presence of impurities with a
particle size of less than 1 μm is of less importance.
General errors that will affect measurements of cell concentrations are:
Unsatisfactory blood sampling and storage;
Inadequate dilution of the specimen;
Fibrin precipitates or cryoprecipitates in the specimen;
Inadequate lysis of red blood cells when counting white blood cells
Lack of homogeneity in the distribution of blood cells in the dilution.
Technical errors in cell counting may result from:
 Fluctuations in the electric current;
 Incorrect setting of the size threshold of the instrument;
 Dust particles in the diluent;
 Leakage in the suction system of the instrument;
 Partial or total obstruction of the aperture;
 Multiple cell passage at high cell counts;
 Carry-over from one measurement to a subsequent measurement.
To avoid these errors, the following measures must be routinely undertaken:
 The cell counter should be connected to an electrical stabilizer;
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 The aperture between the electrolyte chambers should be checked after each
series of measurements, and cleaned with a small soft brush, if necessary;
 The electrodes must be checked, to ensure that they dip into the electrolyte
solution in both chambers;
 The fittings of removable parts should be coated with silicone grease to avoid
air-leakage in the suction system;
 The bottle of diluent solution should be checked daily, and the waste solution
 The suction system should be cleaned with diluent solution to avoid carry-over
 At the end of each working day, the suction system should be cleaned with
detergent solution and afterwards with the diluent solution.
 The aperture of the glass cuvette must always be kept immersed in diluent
solution to avoid obstruction.
Further maintenance procedures (e.g., cleaning of mercury, etc.), must be carried
out according to the manufacturer’s instructions.
Optical Methods of Cell Counting
The cell counts RBC, WBC, PLT, and MCV may also be determined by measuring
the light scattered from each cell particle as it passes through an aperture, as
illustrated in Figure 40. The blood is heavily diluted to reduce the number of
particles counted to one at a time. A sheath fluid is directed around the blood
stream to confine it to the center of the aperture.
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Figure 40: Optical cell countingThe cell is illuminated by a light source such as a laser. The
angle of scattered light is different for different-size cells. The scattering angles of platelets
and red blood cells are sufficiently separate that these two types of cells can be
distinguished by directing the scattered light to different detectors. To separate white blood
cells from red blood cells, it is necessary to destroy the red blood cells with a lysing agent.
This also frees the hemoglobin so that it can be measured. Instrumentation based on this
principle presents a display of RBC, WBC, PLT, ht, MCV, MCH, and MCHC.
Blood Cell Counter
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Figure 41: Blood cell counter – basic block diagram.
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Each cell that crosses through the aperture produces a pulse. The number of pulses
indicates the concentration of RBCs since a fixed volume is drawn (start-stop
mercury contact/pulse gate control circuit). A gated electronic counter counts
pulses above the threshold amplitude and presents the result to the display panel.
The area under the pulse curve (or peak) indicates the volume of the cell passing
A complete count is made in 20s and is displayed as a digital count and CRT peak
versus time display. This conductivity counting method is known as the Coulter
Counter Method. Two Coulter Counters are shown in Figure 42. These devices
measure the RBC count, the WBC count, the MCV (mean cell volume), and Hgh
(Hemoglobin concentration) and calculate Ht (hematocrit), MCH (mean cell
hemoglobin), and MCHC (mean cell hemoglobin concentration).
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Figure 42: Two blood cell counters
(a) Coulter model F. (b) Coulter model senior.
Results are printed out on a specially prepared report form (multicopy card), as
shown in Figure 43 The printout represents a statistical weighting of counts up to
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Figure 43:Hematology output card.
Maintenance includes frequent calibration checks and replacement of burned-out
lamps. Coulter counters have more mechanical problems than electronic. For
example, the pipette system that draws the sample into the machine must be
frequently cleaned. Tubing also becomes dislodged or obstructed.
4. In Figure 36, the resistance of the orifice is 1 kΩ when there are no blood cells
of any type in it. A single RBC in the orifice increases its resistance to 1.02 kΩ.
What will be the output voltage Vout each time an RBC passes into the orifice? The
bias voltage VBB = 15 V.
5. A blood cell counter threshold is set to zero and the output display reads 6.32 x
1012 per liter. The threshold is then set to T1 in Figure 37 and the output reading
becomes 6.15 x 10 12 per liter. The threshold is then set to T2 and the output reading
becomes 0.015 x 1012 per liter. Find the count of the RBC, WBC, and PLT.
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The physician requires exacting analysis of the body fluids and substances in his
treatment of diseases. These tests and analyses are performed in a laboratory of
some type. It may be a small room attached to his office or in the hospital where a
large, complex laboratory performs many tests and extensive research. Many
actual tests are performed on the fluids, etc., a few of which are sedimentation,
clotting, sugar content, albumin and salinity. The results of these tests enable the
physician to diagnose, treat and follow the development and care of body functions
as well as disorders and diseases.
The analysis of liquids normally requires it to be separated into its components
and/or the materials suspended in it. Blood can be separated into areas of the cells
and plasma. This separation enables studies to be made of each individual
component. One means of causing separation is through the use of gravity. The
substance under study may be permitted to stand and periodically be examined to
determine the rate of settling of the heavier components. This is known as testing
for sedimentation.
When gravity alone is not sufficient, other
methods have been devised to accomplish
the desired tests. The application of suction
to facilitate the movement of liquids through
a filter, leaving the particles or sediment on
the filter has decreased the time required to
accomplish the sedimentation tests. An
example of the equipment required for
this procedure is shown in Figure 56.
Figure 56: Sedimentation test equipment
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When the suspended materials are finely divided, or closely mixed and do not vary
greatly in density, their separation requires greater forces to be exerted. This is
accomplished through the use of centrifugal force. Centrifugal force is defined as
“a force that tends to move substances outwardly from a center of rotation.” The
amount of centrifugal force developed depends upon the speed (revolutions per
minute) at which the object is rotated and the radius of the circle in which it is
rotated. The force required to keep a body moving in a circular path is directly
proportional to the square of the speed (RPM) and the radius of the rotation.
More and more the laboratory technician is becoming familiar with the term
relative centrifugal force. This is a method of expressing the relationship between
the gravitational force on a body and the force required to keep it in circular
rotation. The manufacturers of centrifuges have published a chart on the Relative
Centrifugal Force, Figure 57.
A quick review of these charts and their explanations will permit the laboratory
technician to set up procedures called for in specific tests. Also, these charts enable
the associated laboratories to establish standards for specific analysis, thus they all
perform identical tests.
The centrifuge is an essential unit in any laboratory. A centrifuge is a machine that
employs centrifugal force to separate substances of different density, remove
moisture or simulate gravitational effects.
When a fluid is analyzed it is placed in a container and spun at a specified speed
for a certain length of time. The heavier particles are thrown outward and
congregate at the bottom of the container. Conversely the lighter substances will
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remain at the top of the vessel. Each substance then may be microscopically
There are many types and sizes of centrifuges manufactured. The basis for
selection must be determined by the type of tests and the quantity of them to be
performed. All models, sizes and manufacturers will be found in the medical
facilities today. They range from high speed table models used for microanalysis to
the large refrigerated floor models.
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A centrifuge applies a sustained centrifugal force (i.e., a force due to rotation) to
impel matter outwards from the center of the rotation. This principle is used to
separate out particles in a liquid medium by sedimentation. The physical basis of
the separation is the action of a centrifugal force on the rotating particles, which
increases with the radius of the rotational field and the velocity of the rotation. The
rate of sedimentation is determined by the density of the particles. Dense particles
sediment first, followed by lighter particles. Depending on the conditions, very
light particles may even remain in suspension.
The relative centrifugal force is related to the number of revolutions of the rotor
per minute according to the formula:
RCF = 1.118 X 10 –6 X r X n2
Where RCF = relative centrifugal force (g)
r = radius in millimeters from the centrifuge spindle to point
of tube
n = number of revolutions per minute
The relative centrifugal force can easily be calculated from a nomogram (Figure
58), where the radius is measured from the center of the rotor to the middle of the
tube placed in the radially oriented rotor bucket; e.g., if the radius is 75 mm, the
speed of rotation must be 2500 revolutions per minute to develop a centrifugal
force of 520 g. It is important that the temperature in the centrifuge does not
exceed 37˚C; otherwise degradation of some constituents of the specimen may
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Figure 58: Relative Centrifugal Force
There are two main types of centrifuge: preparative and analytical.
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Preparative centrifuges are used to separate the solids suspended in biological
samples from the supporting fluid. This is the most common type of centrifuge,
and they are fitted with swing-out, or fixed-angle, heads.
Preparative centrifuges vary in their sample capacity and size, from floor standing
to small capacity centrifuges that can be sited on a bench. Some are fitted with
internal windshields to protect the operator from contamination
by any aerosols that may be formed. This is now a mandatory safety requirement in
many countries.
Two types of preparative centrifuges are currently used – mechanical and electrical
– although the majority are electrical centrifuges.
Analytical centrifuges may be used to quantify one or more solid components in a
mixed suspension. The only centrifuge of this type used in medical laboratories is
the microhematocrit centrifuge.
Unpacking, siting, installation, and electrical requirements
Follow the manufacturer’s instructions, if these are available. Remove all packing
and any transit fixings that may have been used.
Check that the equipment voltage is the same as the local supply, and that the fuse
rating is correct. A correct fuse should protect the equipment from serious
electrical damage.
Both bench and floor-standing centrifuges must be sited on a rigid surface, away
from laboratory balances. Bench centrifuges should be at least 20 cm from the
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edge of the bench. If the sample breaks during rotation, considerable “out of
balance” forces are generated, and the centrifuge may move about unpredictably.
Good working practices
 The centrifuge must be positioned exactly horizontally to avoid movement if
the instrument is out of balance during operation.
 It is critically important that the centrifuge load is balanced at all times.
Therefore, tubes should be loaded in matched buckets fitted with rubber
cushions, and should be arranged so that like loads are opposite. A “dummy,”
i.e., a tube containing the appropriate volume of water, must be included when
an odd number of specimens are to be centrifuged. Final balancing should be
carried out by placing paired loads on the two pans of a reasonably sensitive
balance, and balancing by adding water from a bottle or Pasteur pipette; if
possible, add water to the lighter of the two samples, so that it balances the
heavier load. Biological samples should be capped during centrifugation. The
centrifuge should be stopped immediately if it develops an abnormal noise,
indicating that it is not properly balanced.
 After use, the buckets should be inverted to drain dry.
 After any sample spillage, always clean up the buckets and the centrifuge and
disinfect with 70 % (700 ml/l) alcohol immediately.
 Clean and disinfect the centrifuge often because it is one of the most frequently
used instruments.
 Check mountings and replace if necessary.
 Check motor brushes and replace if necessary.
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 Check for corrosion and clean if necessary.
 Never operate a centrifuge with the lid open.
 Do not use the centrifuge at higher speeds than necessary.
Hematocrit centrifuges need not be balanced before use. As the samples are small
capillary tubes, and the forces relatively low, it is only necessary to load the
samples symmetrically. Never run the centrifuge with the lid open. Capillaries
should be plugged at one end with the recommended sealing compound. The
plugged end should always be placed against the sealing gasket. Even with the
above precautions, it is possible that blood may leak from the bottom of the
capillary. After any spillage, the centrifuge chamber must be disinfected and
cleaned immediately with soap solution, and then with 70% (700 ml/l) alcohol.
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Because centrifuges are regularly used to prepare blood and urine samples, it is
recommended that the rotor bowl, centrifuge head, buckets, and trunnion rings be
disinfected before any servicing is carried out.
A general tool kit is satisfactory.
Suppression capacitor (interference filter)
carbon brushes
Rubber feet (bench models)
Most of the centrifuges are adaptable to a large range of procedures. A chart of
recommended centrifugation for different tests is shown in Table 7
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Table 2: Centrifugation for different tests.
Urine, Sediment, Blood, 1500
5 minutes
Casts, etc.
Stool or Feces
2 minutes
30 minutes
5 minutes
10 minutes
VDRL, Flocculation
Blood grouping
Centrifuge to pack cells
Coombs test
2 minutes
Prothrombin time
5 minutes
Clotting time
1000 to 3000
5 minutes
Blood sugar
5 minutes
Blood urea and nitrogen 2000
1 hour
Plasma proteins
5 to 10 minutes
Liver function
Centrifuge 5 minutes
Blood Chlorides
5 minutes
Protein Bound Iodine
10 minutes
Ascorbic Acid in blood
Serum Albumin
5 minutes
From this chart we could readily assume that a centrifuge that rotates between
1000 and 3000 revolutions per minute would suffice for the majority of tests and
analyses. This may be true, but with the advances in technology, greater scientific
accuracy is now being demanded.
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Table Model Centrifuges
We will group our centrifuges into two general categories, i.e., table and floor
models. The large hospitals, clinical and research laboratories and blood banks
have requirements for equipment to run the same test on a large number of
specimens simultaneously. They usually desire the larger floor models in which 64
to 100 specimens may be run. However, the centrifuge used most extensively is the
table model. These small units are capable of processing from 1 to 12 specimens at
a time.
There are several small centrifuges on the market today. They are well balanced
and cushioned to prevent vibration and movement. These units normally rotate
about 1700 to 1800 RPM and can run any of the tests the doctor would desire
within a short period of time. However, the Relative Centrifugal Force is only
about 400.
Examples of the table model centrifuge are illustrated and discussed in the
following figures and diagrams.
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Figure 59: Small laboratory centrifuge, 110 volts, AC-DC
This unit, marketed by several manufacturers, will be found extensively used in the
dispensaries, clinics, wards, and doctors’ offices. It is powered by an international
motor (AC-DC) and has a maximum speed of approximately 4000 RPM. The
electrical circuitry is relatively simple – consisting of the power cord, a
combination on—off switch, speed control rheostat, and the motor. Figure 60 is a
schematic drawing of this electrical circuitry
The principle disadvantage of this unit is the uppermost part, or head, into which
the specimens are placed, is exposed and spins in the open. The laboratory
technician has a tendency to stop the head by placing the palm of the hand against
it and exerting pressure. This may cause the joint between
the motor shaft and the head to flex and become loose or break. It can also bend
the motor shaft.
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Figure 60: Simple electrical circuitry schematic
The centrifuge in Figure 61 has a cast iron body in which the head spins. All
moving parts are enclosed. There is no brake system, and once the specimens have
been spun for the proper time, the head coasts to a stop.
Figure 61: Small Laboratory Centrifuge, 110 volts, 60 Hz, AC
Centrifugal force is generated by a series wired AC motor. The electrical circuitry,
Figure 62, is somewhat different than the preceding unit. It is composed of the
power cord, on—off switch, autotransformer, speed control and motor. The speed
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control is operated in conjunction with the autotransformer, thus applying different
voltages to the motor to obtain a variety of speeds (RPM).
Figure 62: Electrical Circuitry for a series wired AC motor
In serological laboratory procedures, micromethods become very important. Often
the amount of blood available is limited and the micro procedures are mandatory.
These centrifuges are run at much higher speeds (12,500 RPM) with resulting
higher relative centrifugal force. The head for the Micro-Hematocrit Centrifuge,
Figure 63, is a flat disk with grooves cut into it. These grooves hold the capillary
tubes. A cover must be placed over the head to secure the small glass vials.
The electrical circuitry on these units varies with the manufacturer. Some employ
an electrical brake; others do not. The explanation of this circuit, Figure 63, will
include several principles not yet found in the other table model centrifuges.
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Figure 63: A Micro-Hematocrit Centrifuge, 110 volts, 60 Hz, AC
Figure 63: Electrical circuitry with safety switch
First, we have an interlock switch. This switch is activated when the cover is
closed and prohibits operation when the cover is open.
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Secondly, there is an electrical brake. It is used by the technician when he desires
to stop the head quickly after a procedure has been run.
To operate this centrifuge, it is only necessary to close the cover and rotate the
timer knob clockwise. This action closes the points in the timer and applies the 110
volts to the series wired motor. There is no speed control on this centrifuge, and it
will run at maximum speed until the time on the timer has elapsed. During the run
cycle current flow is through the interlock switch, timer points, brake switch, the
armature and the field coils. This causes the head to spin clockwise.
When stopping the head, the technician presses the brake switch. This double-polesingle-throw switch is spring loaded and returns to its normally closed position
when released. When the brake switch is depressed, the points change position.
With this change, the current flow is now through the interlock switch, the two
parallel 75-ohm resistors, the brake switch, the armature, and field coils. A
comparison of this current flow through the armature to the flow in the run cycle
will reveal that it is now in the opposite direction. The head will attempt to turn
counterclockwise, thus giving the braking action. Should the brake switch be held
in the depressed position after the head has stopped it will begin spinning in the
counterclockwise direction.
Although there are many table model centrifuges, the three that have been
discussed are a good representation of this group. With certain exceptions, (i.e.,
bent motor shafts or damaged heads) maintenance consists of cleaning, lubrication,
and changing the motor brushes periodically.
Floor Model Centrifuges
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Floor model centrifuges are used in clinical and research laboratories and blood
banks where large numbers of tests are performed. They are available from several
manufacturers. Consequently, many varieties may be encountered. Some types are
refrigerated, some use ultra-violet and infrared lights to accomplish the procedures
desired. However, the majority of the components, systems, and operations are
very similar and the special systems are only additions to the basic units.
The “Lourdes” Model 10 Clini-fuge, manufactured by Vernitron Medical Products,
Inc. (Figure 64), is available under National Stock Number NSN 6640-00-4129008. It has been chosen as the demonstration model. Other manufacturers’ units
will differ in their electrical circuitry and application; however, if this unit is
understood, little difficulty should be encountered with them. The Model 10 CliniFuge is a medium duty, medium capacity centrifuge designed for a low speed
operation in continuous or intermittent use, Depending on the heads and other
accessories used, maximum allowable speeds are between 1,200 and 5,500 RPM.
Figure 64: The “Lourdes Model IO, Clini-fuge
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The components in the Model 10 Clini-Fuge are basically the same as those in
other units and are used for similar purposes. They are:
A. In the base
The motor is a series wound vertically mounted AC motor of ¾ HP
rating. It is used to spin the head in which the specimens are
mounted. Attached to the bottom end of the armature shaft is a
small generator that produces the electrical signal for the speed
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B. In the tank
The head is the framework upon which the carriers, shield, caps,
etc. are mounted. The specimens are placed in vials in
these accessories. There, horizontal swinging heads are machined
with a 20 degree tapered bore. Each machine is supplied with a ½
inch to 20 degree tapered bore adapter with a nut. Care should be
taken when mounting the head to prohibit thread or taper damage.
C. On the control panel
I. Front view, Figure 65. Proceeding from left to right and in
numerical sequence, the following components and controls
appear and will be discussed:
a. Fuse, 10A, Slo Blo (1). This fuse is in the main power
line and is used to protect the transformer and relay
coil from excessive current.
b. Centrifuge master switch (2). A double-pole, singlethrow switch used in the main power line. It controls
both the “common” and the “hot” lines of the
incoming power.
c. Pilot light (3). This light indicates that power is
available to the relay coil.
d. Brake switch (4). A double-pole, double-throw,
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spring loaded switch used to apply a braking action
which stops the free-running motor. It must be held in
the “on” position as the spring action always returns
it to “off.”
e. Timer indicator (5). This control sets-up the mode in
which the centrifuge is to be run.
1.For an un-timed procedure the pointer will be
placed on “hold.” With all other controls in
their proper settings, the centrifuge will run indefinitely.
2. For a timed procedure, the timer indicator
knob must be rotated at least to the 10 minute
position and then to the length of time desired.
Most timed procedures are usually longer than 10 minutes.
With all other controls in their proper settings, the centrifuge
will spin until the timer runs down to zero. It will then coast to
a stop, or the brake can be applied.
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f. Tachometer (6). The
tachometer is an electrical RPM
meter that indicates the rotation
speed of the centrifuge head. It
has two scales, 0-6,000 and 030,000. As the
centrifuge has a top speed of
5,500 RPM, only the 0-6,000
scale is used. Directly under the
meter is a small hole in the
panel where a screw head can
be seen. This is the calibration
adjustment for the
g. The Speed
control (7). is an autotransformer that determines the
voltage applied to the motor.
arbitrary and do not indicate
voltage, RPM, or anything else.
The operator may determine,
after becoming familiar with
the unit, that a particular setting
Figure 65: Front View of the Front Panel
Model IO, Clini-Fuge
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For example, a setting of 80 may correspond consistently with 2,000 RPM on the
tachometer. Thus, the operator can set the control at 80 and be assured that the
centrifuge will spin at 2,000 RPM.
2 Rear view, Figure 66. Proceeding from left to right and in a numerical sequence,
the following components and controls appear and are discussed.
This view shows the front panel loosened and tilted out in order to see the
Diode (1). This is a rectifier that is used in the
brake circuit.
Relay (2). A double-pole, 120 volt relay which
is energized when the centrifuge is to be run.
Resistor (3). A 5 ohm, 100 watt, wire-wound,
coated resistor used in the brake circuitry.
d. Fuse (4), Centrifuge master switch (5), Pilot
light (6), and Brake switch (7). These
components were discussed in the
explanation of the front view of the panel. However, it should be noted that the
contacts on the two switches are numbered. These numbers are important for the
circuit explanation and for troubleshooting.
e. The timer (8). This timer is driven by an electrical motor which is indicated by
the circle in the lower left of the drawing. Connections “A” and “B” are the leads
which carry the potential to run it. The numbered connections once again
pertain to the tie-points in the electrical circuitry and will be used in
f. The tachometer (9). This component was discussed previously. The numbered
connections are for the electrical signal supplied by the small generator mounted
on the lower end of the motor.
g. Potentiometer (10). The potentiometer is in series electrically with the
tachometer and is used to adjust it to accurately register the speed of the head.
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h. Speed control (11). The speed control was previously discussed. It was referred
to as an autotransformer. This device is also referred to as a “Variac” or a
“Powerstat.” Both are tradenames of manufacturers.
i. Micro switch (12). The micro switch’s normally open points are closed when the
speed control is rotated counterclockwise to “0”. This action energizes the relay
coil and pilot light.
Figure 66: Rear view of the front panel
Model IO, Clini-Fuge
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A. Modes. There are two modes of operation of the centrifuge. These are:
1. Un-timed. When the technician is in the area of the centrifuge and the
procedure does not specify an accurate time period he may use the un-timed
mode. To set the unit up for the procedure, he must first load the head correctly
to include balancing, and closing the cover securely.
He then operates the following controls as indicated:
a. Shift the centrifuge master switch to the “on” position. This energizes the
b. Rotate the timer indicator to the “hold” position.
c. Rotate the speed control counterclockwise to “O.” The relay should
energize and the pilot light should glow.
d. Rotating the speed control clockwise applies voltage, starting at O and
progressing toward 120, to the motor. When approximately 35 volts is
impressed across the motor it will begin to turn (unload head). Continued
rotation of the control applies more voltage, causing higher current flow,
resulting in faster speeds of the head. The technician must determine the
speed and time factors for the procedure. He adjusts the head RPM with
the speed control and stops the centrifuge when the time has elapsed.
e. To stop the centrifuge, the technician may:
1. Switch the centrifuge master switch to the “off” position and let it
coast to a stop.
2. Apply a braking action by pressing the brake switch to the “on”
position and holding it there. This causes the head to stop spinning
in approximately ½ the time it takes to coast to a stop.
2. Timed mode. All actions of the timed mode are identical to the un-timed with
the following exceptions:
a. During the control setup, the time must be set to the time
required for the procedure. This is limited only by the fact
that the timer is capable of a maximum of 120 minutes (2
hours). To set the timer, it must be rotated past the 10minute point in order to activate the timer motor contacts.
Then the time setting desired may be made.
b. When the timer indicator returns to “0,” the electrical circuit
to the relay is opened and the motor is de-energized. The
motor will coast to a stop. Should the technician desire a
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faster stop, he can apply the brake circuit as explained in the
un-timed mode.
Electrical Circuitry
The electrical sequence will be explained in four parts. These will be the four main
circuits of the centrifuge. It must be remembered that these four main circuits also
may be reduced to individual load device circuits.
1. The Primary Circuit:
This major circuit has four individual load devices. These are the
autotransformer, relay, pilot light, and timer motor. The pilot light is
connected directly across the relay, so they will be discussed as one load
a. The autotransformer. The autotransformer is the principle load
device for the primary circuit. It is, however, the most basic of the
circuits in this centrifuge. The incoming line goes to one enters the
unit as a connector shown as J2. The white or “common” lead used
J2-13, then goes to one side of the centrifuge master switch and
then to terminal 5 on the autotransformer. The black or “hot” line
uses J2-14 and continues through F-1, the 10-amp fuse, through the
other half of the centrifuge master switch and to terminal 4 of the
b. The relay/pilot light is composed of several switches and
conductors. To energize these two load devices:
1. Move the centrifuge master switch to the “on”
2. Rotate the timer indicator to the “hold” or the “timed”
position. This closes the points between 1 and 2 in the
3. Rotate the speed control counterclockwise to the “0”
position. This action closes microswitch S3 and
completes the circuit. The current path is from the
“common line” of S1 to autotransformer terminal 5,
then to the timer contacts 1 and 2, and through the
yellow lines to one side of the relay coil and the pilot
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light. The “hot” lines leaves S1 through the black line
to S2, the brake switch. When in the relaxed position,
contacts 2 and 3 are connected. Current now flows
through the white wire to terminal 4 of the relay.
From that point a white wire connects to the “N.O.”
point of the S3. With S3 closed, current flows through
a black wire to the Relay 8 terminal, then through a
black wire to the Relay 6 terminal, which is the other
end of the relay coil. It will energize and shift its
points from the NC to the “NO.” A black wire also
connects the Relay 6 terminal to the pilot light 2
terminal. This completes the pilot light circuit at the
same time as the relay is energized. Switch S3 can
now be opened and the relay and pilot light will
remain energized through the lock-in action of Relay
4 and 8 points.
c. The Timer motor is the last load device in the primary circuit.
When a timed procedure is to be used, rotation of the timer
indicator closes both sets of its contacts. Those between 1 and 2
complete the relay/pilot light circuit; and those between 7 and 8
complete the circuit for the timer motor. As soon as the centrifuge
master switch is closed, the timer motor begins to run. When an
untimed procedure is to be used, the timer indicator is placed on
“hold.” This closes the contacts to the relay and pilot light only.
2. The Secondary Circuit:
This simple series circuit consists of the motor, field windings, relay
contacts, brake switch and associated connectors. Depending upon where
the speed control is set, voltage 0 to 120 volts will be available between
terminal 3 and 4 of the autotransformer. When the relay is energized,
contacts 3 and 7 of the relay are closed. This action provides a path for
current flow through the motor. As the speed control is rotated clockwise,
the brush connected to terminal begins to turn. The path for current flow
is from terminal 3 of the autotransformer to point 3 of the relay, to point
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7 of the relay, through J1-7 to one brush in the motor. Current flow
continues through the armature and out the other brush to J1-8, through
contacts 6 and 5 of S2 (brake switch) through J1-9 to the field windings
and J1-10 to terminal 4 of the autotransformer.
3. The Brake Circuit
a. The brake circuit is composed of two sections – the field winding
circuit and the resistor circuit. To place the brake circuit in operation,
it is necessary to push S2 (brake switch) to the “on” position and hold
it there. This action accomplishes several things:
1. It opens the lock-in circuit and de-energizes the relay
and pilot light.
2. When the relay contacts go to their “NC” position, the
motor circuit is opened and the 5-ohm resistor is
placed parallel to the motor.
3. The section of S2 that is in the secondary circuit
opens the motor circuit in a second place. This switch
completes the field winding circuit of the brake.
b. The Field Winding Circuit. This circuit has as its power source those
windings of the autotransformer between terminals 2 and 4, and a load
device composed of the two field windings. The voltage between
these terminals is 22 volts AC. As current flows through the D-1 (a
rectifier), it is converted to pulsating DC and reduced in magnitude to
approximately 11 volts. All that is left in the circuit now is the field
coils. An 11-volt pulsating DC impressed on these coils creates an
electromagnet. This produces a magnetic field strong enough to cause
the armature to slow and stop.
c. The Resistor Circuit’s power source is the MOTOR ARMATURE. A
quick reminder, to create an EMF, a magnetic field, coils of wire and
relative motion is all that is required. All of these are present when the
brake system is applied. To “load” this generated EMF and aid in the
braking action, a 5-ohm, 100-watt resistor is connected across the
armature when the relay points 1 and 7 close.
d. The Tachometer Circuit. The tachometer is the last electrical circuit to
be discussed. The bottom of the motor armature shaft connects to a
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small generator. As the motor turns, an electrical voltage is generated.
The voltage is linear with the speed of the motor. At maximum speed,
6 volts is measured at the generator. The current from this generator is
the signal that excites the tachometer. Two load devices are series
connected across the generator. A potentiometer is used in calibrating
the tachometer. It is adjusted by turning a small screw located beneath
the tachometer.
B. Tachometer verification
Due to the advances in technology in the last decade, more exacting results for
blood tests, etc., are required. Through the used of the available charts for speeds
and forces, certain requirements are placed on the technicians. If the requirements
for certain tests are followed precisely, accurate results can be supplied.
However, this machine must be able to fulfill its part of the procedures. Should the
timer or the tachometer be in error, the results will not be accurate. Tachometer
verification is a part of all good maintenance programs. The timer’s accuracy may
be checked with any stop watch or a watch with acceptable accuracy.
Recently, the College of America Pathologists, Chicago, Illinois, in their summary
report, Standards for Accreditation of Medical Laboratories, has stipulated that the
revolutions per minute of all centrifuges must be verified. Although this
verification is the responsibility of the laboratory chief, it is anticipated that the
biomedical equipment technician will accomplish it on a scheduled basis.
There are several photo-tachometers available for this use. Even though the
tachometer is checked on units using mechanical tachometers, there is no
adjustment. In those cased, decals noting the date and degree of inaccuracy at
different RPM settings will be attached to each centrifuge. When the unit is
equipped with an adjustable electrical tachometer, it will be calibrated to indicate
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the verified RPM. Decals indicating date, verifier, etc., will be attached to the
1201 and 1205 Yankee Rotors – Single Speed
This machine is for use on 115 volt 60 cycle Alternating Current only.
To Assemble for Use
Attach the top platform to the “H” frame by means of the four screws enclosed in
the envelope tied to the frame. Screw the four rubber feet into the tapped holds
provided at the corners of the bottom of the machine.
To use the automatic timer, first place the switch in the “OFF” position. The timer
is adjustable for periods up to 5 minutes. To set the timer, loosen the thumbscrew
that holds the timer stop plate in position on the front of the rotator. Move the plate
until its upper arrow points at one of the numbers indicating minutes on the bottom
of the name plate. Then tighten screw to lock the plate in position. To operate,
move the timer lever until it hits the plate and then release; the machine will now
run for the desired period of time and then automatically shut off.
For continuous operation, use the “ON-OFF” switch, leaving the timer lever in the
“OFF” position.
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This rotator is factory lubricated and ready for use. If used continuously,
lubrication should be renewed after approximately 30 hours of use or at least once
monthly. For access to lubricating points, the platform must be removed.
The motor oil lines are mounted directly on the motor and the lower one is capped
with a slotted neoprene bulb. To oil the upper bearing, tip the machine onto its left
side, insert the tip of the oilcan into the tube and place about 2 drops of oil in the
tube. For the lower oil line, insert the tip of the oilcan into the slot of the neoprene
bulb and feed the oil directly into the bulb.
Do not over-oil! Too much oil will damage the motor.
The universal joints of the platform legs and the ball bearing near the center of the
flywheel should be re-packed with Vaseline
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Figure 67: Centrifuge parts diagram
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Clinical Instrumentation in the early 1900s was almost nonexistent. Since the
1950s, sophisticated apparatus have been developed to measure blood parameters,
as described earlier. The complex substances appearing in blood serum can be
evaluated for concentration based on chemical color reactions. Blood cells can be
counted by electrical conductivity changes as they pass through a fixed diameter
aperture. The following types of instrumentation are used to analyze blood:
1. Colorimeter or filter photometer is an optical electronic device that
measures the color concentration of a substance in solution (following the
reaction between the original substance and a reagent). The results are
displayed in percent optical color transmittance or absorbance to indicate
hemoglobin concentration, for example. The densitometer is a device
similar to the colorimeter and measures optical transmittance (density) of
particles in fluid suspension.
2. Flame photometer is an optical electronic device that measures the color
intensity of substances (i.e., sodium or potassium) which have been
aspirated into a flame.
3. Spectrophotometer is an optical electronic device that
measures light absorption at various wavelengths for a given liquid
sample. This is a type of sophisticated colorimeter.
4. Blood cell analyzer is an electromechanical device that measures the
number of red and white blood cells per scaled volume. This is
accomplished by noting the changes in electrical conductivity as the cells
pass an aperture of fixed diameter.
5. pH/Blood gas analyzer is an electromechanical device which
measures blood pH (acid-base balance). PO2 (partial pressure of
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blood oxygen, and PCO2 (partial pressure of blood carbon
dioxide. This is accomplished through use of glass electrode transducers.
6. Chromatograph is an electromechanical device used to separate, identify,
and measure the concentration of substances in a liquid medium. Results
are displayed as colored bands in a liquid column or as colored strips on
7. Autoanalyzer is an electromechanical-electronic device that sequentially
measures and displays blood chemistry analysis. This is accomplished by
using mixing tubes and colorimeters arranged in a serial system
Most hospitals have a laboratory area separate from patient areas; it is used solely
for chemical analyses and measurements of body fluids and tissues. Typically,
analysis is done on blood and urine and on body tissue. These measurements are
made to aid physicians in the diagnosis of disease states and to help them monitor
the effects of therapy. For example, when oxygen therapy is given to a patient, it is
necessary to frequently monitor the partial pressure of oxygen and carbon dioxide
in the patient’s blood. Accuracy of measurement is critical in the laboratory. And
because the chemical parameters are sensitive to environmental factors such as
temperature, light, and humidity, equipment calibration must constantly be
monitored. The chemical solutions are often active, and cause aging and wearing
effects that may impair accuracy.
The analytical methods used should be specific for the substance analyzed; that is,
there should be no interference from any of the innumerable other substances
present in the sample. In addition, the analytical method should be rapid,
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economic, and suitable for automation. The analytical methods in most common
use may be classed as optical, electrochemical, and chromatographic. The
radioimmunological method is also used in chemical chemistry.
J. Methods of Optical Analysis
Light can be both absorbed and emitted by materials. This occurs to different
extents depending on the wavelength, and on the chemical and physical properties
of the material. Most of the analytical methods used in clinical chemistry are based
on this fact. Two different methods are described below: absorption photometry
and emission photometry.
1. Absorption photometry
Quantitative absorption photometry is based on the principle that the light
absorption of the test substance bears a known relation to the concentration of the
substance. The sample is usually pre-treated by the addition of reagents. Different
chemical processes are used; for example, the substance may be linked with a
pigment or converted to a suitable colored substance. The color need not be visible
to the naked eye, since absorption measurements can be made using either
ultraviolet or infrared light. Absorption photometry is by far the most common
analytical method in clinical chemistry. For example, it is used for determining the
concentration of practically all low molecular weight organic substances.
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Figure 7: Principles of absorption photometry
Figure 7: Principle of absorption photometry.
A quantitative determination is performed by measuring the light absorption of the
prepared sample (Figure 7). Light from a lamp is made monochromatic by
filtration (or by some other suitable technique) and is passed through a cuvette
(cell) containing the sample solution. By means of a photoelectric circuit, the light
transmission, I1, is measured. The light transmission I0 is also measured with the
cuvette filled with reagent solution containing none of the analyzed substances, a
blank value is thus obtained.
According to Lambert-Beer’s law:
I1= I0e –εxc
Where c is the concentration (g/cm3) x the pathlength of the cuvette (cm) and ε the
extinction coefficient (cm2/g). (We may consider the extinction coefficient to be
the apparent area that one gram of the substance blocks for the light wavelength in
question. The greater this apparent cross-sectional area is for a substance, the
greater is the probability that a photon will be absorbed by a molecule of the
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substance.) The concentration c of the substance is calculated from the absorbance,
also called the extinction.
E = InI0/I1 = ε xc
The choice of wavelength in absorption photometry is critical. Thus, the
wavelength used must be one where the absorption of the analyzed substance is
high relative to that of the reagent solution or other substances present in the
sample. Furthermore, the wavelength should be chosen so that the part of the
absorption curve used is almost horizontal. This avoids errors, due to the fact that
the light is not completely monochromatic. For example, the absorption peak β in
Figure 7 is not used even though it is higher than the peak α. By using the peak α, a
more accurate determination can be made, since it is possible to work mainly at a
plateau of the curve. The relationship between concentration and absorbance is
usually assumed as linear, thereby simplifying calculations.
Non-linear relationships between concentration an absorbance can be due to two
causes: physical and chemical. The physical causes, which are the more common,
are usually due to the light not being sufficiently monochromatic relative to the
shape of the absorption curve; this causes a shift in the peak wavelength, as the
light is selectively filtered by the sample.
The chemically produced non-linear relationship may be due to the fact that the
color reaction is affected by the presence of other substances (salt error, protein
error) or that the color reaction does not behave strictly stoichiometrically, that is,
that side-reactions appear so that the reaction does not follow any simple formula.
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The colorimeter shown in Figure 1 is a filter photometer that measures the color
concentration of a substance in solution. This is accomplished electronically by
detecting the color light intensity passing through a sample containing the reaction
products of the original substance and a reagent. A yellow-colored urine sample,
for example, passes yellow light and absorbs blue and green. For this reason, and
to obtain purity in measurement, optical color filters are used to select a narrow
wavelength spread (bandwidth) of light that shines on the photodetectors.
A colorimeter consists of a light source broken into its spectrum of colors by a
prism or diffraction grating. The individual colors are then passed through the
sample. The amount of each color absorbed is measured to determine the type and
concentration of substances in the sample. Since these measurements are highly
temperature dependent, control of this parameter is necessary.
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Figure 21: Colorimeter -- filter photometer.
Basic colorimeter analysis (Equation 1a) involves the precise measurement of light
intensity. Transmittance is defined as:
T = 11 x 100 percent
I2 = TI1
I2 = T2I0
I0 is initial light intensity
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I1 is first attenuated light intensity
I2 is second attenuated light intensity
T is transmittance in percent
Absorbance (optical density) is defined as:
A = log I1 = log I
A is absorbance
I1 and I2 are as before
If the path length or concentration increases, the transmittance decreases and the
absorbance increases. Essentially, this phenomenon can be expressed by Beers
A = aCL
A is absorbance
L is cuvette path length
C is concentration of absorbing substance
a is absorbtivity related to the nature of the absorbing substance
and optical wavelength (known for a standard solution concentration).
Therefore, the concentration of the unknown solution can be found from the
following relationship:
Cμ = C2 Au
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Cu is unknown concentration
C2 is standard concentration (for calibration)
Au is unknown absorbance
As is standard absorbance
A basic colorimeter schematic is shown in Figure 21b. Observe that light passes
through an optical color filter, is focused by lenses on the reference and sample
cuvettes, and falls on the reference and sample photodetectors.
The difference in voltage between the two detectors is increased by a dc amplifier
and applied to a meter. A calibration procedure is as follows:
1. Ground the amplifier input (V1) and adjust potentiometer (R4) for 0 V + 5
mV at the amplifier output.
2. Remove the ground and place reference concentrations in cuvettes 1 and
2 (empty cuvettes or open spaces may also be used).
3. Adjust potentiometer (R1) for 0 V + 10 mV at the amplifier output.
4. Leave the reference concentration in cuvette 1 and replace cuvette 2 with
a cuvette containing the sample.
5. Read the unbalanced voltage on the meter in percent transmittance or
absorbance units.
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A photo of a colorimeter is shown in Figure 22.
Figure 2: Colorimeter.
EXAMPLE I-1 (Refer to Figure 21b) Given V1 = +mV with reference cuvettes
(step 3 in calibration procedure), V1 = +25 mV with reference and sample cuvettes
(step 5 in calibration procedure). R2 = 2kΩ, R3 = 1k Ω, calculate the voltage read
on the meter display for both conditions of V1.
A1 = 1 + R2
op-amp voltage
Av = 3 = 1 + 2kΩ
Condition 1:
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Av = Vm
Vm = AvV1
= 3 (+1 mV)
Vm = +3 mV
Condition 2:
V1 = + 25 mV
= Av V1
Vm = 3 (+25 mV)
Vm = +75 mV
Note that the sample measurement (balanced) voltage is 25 times larger than the
reference measurement (unbalanced) voltage. This is a desirable low-error
Precipitating reagents are usually mixed with samples to remove substances from
the sample. Table 2 shows common chemical tests and their normal ranges.
Maintenance includes calibration adjustment and replacement of burned-out lamps
and photodetectors. Colorimeters are very reliable and usually do not have frequent
electronic problems
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Table 3: Common Chemical Blood Tests and Normal Ranges.
1. Sodium
2. Potassium
3.5 – 5
95 – 105
4. Total CO2
24 - 32
5. Blood urea nitrogen 8 - 16
6, Glucose
mgN/100 ml
70 - 90
7, Inorganic phosphate 3 – 4.5
8. Calcium
9 – 11.5
9. Creatine
0.6 – 1.1
10. Uric acid
11.Total protein
g/100 ml
12. Albumin
13. Cholesterol
160 – 200 mg/100 ml
14. Bilirubin
0.2 – 1
15. SGOT
20 – 50
1 milli equivalent per liter = 1mEq =
molecular weight
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Photometers and Colorimeters.
A spectrophotometer is an optical electronic device that measures light absorption
at various wavelengths for a given liquid sample. Maintenance is similar to the
colorimeter except for mechanically moving parts (diffraction grating, rotating
The chemical content of biological substances can be determined by measuring
how they either absorb or emit visible light. The colorimeter uses light absorption
to determine blood proteins and iron levels. In order to enhance the color of these
substances in blood serum, it is necessary to mix it with reagents. Measurements of
light emitted by ions, such as sodium or potassium in serum or urine, excited by
heat are made with a flame photometer.
A photometer is an optical instrument that is used to measure absorption of light.
According to the Lambert-Beer Law, the absorption of light by a solution is related
to the concentration of the solute. The photometer is an essential instrument for a
laboratory, and can be used for the determination of a great number of analytes in
body fluids. Photometers differ as regards their light source and the way in which
the monochromatic light is generated. The following types are found:
 Instrument with a light source that emits a spectrum of discrete lines
(cadmium or mercury lamp).
 Instrument with a source that generates a continuous spectrum of light
(white light) (tungsten—halogen lamp), which is split by a prism or a
grating into its different components.
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 Instrument with a source that emits a continuous spectrum of light
(tungsten—halogen lamp) which is filtered to produce light with a
desired wavelength.
 Instrument with a diode lamp that emits monochromatic light.
The instruments may be of the single- or double-beam type. In a single-beam
instrument, light passes from the source through a monochromator or filter and
then via a sample cuvette to the detector. In a double-beam instrument, light passes
through a beam-splitting device and then via separate sample and reference
cuvettes to separate detectors.
The instruments with a tungsten light source and filters are usually referred to as
photometers, or in a simpler version as colorimeters. The more complex
instruments with interference filters, prisms or gratings are referred to as
The essential elements of a photometer or spectrometer are shown in Figure 18.
Figure 3: Photometric assembly.
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The positions of the monochromator and the cuvette are reversed in certain
instruments. All of these elements are prone to defects. The diode-lamp photometer
does not need a filter or prism since the emitted light is already monochromatic.
Light source
The light source in a photometer may generate either light with a continuous
spectrum of wavelengths (tungsten lamp), light with a discrete spectrum of
wavelengths (mercury lamp) or monochromatic light (light-emitting diode). To
cover the entire range of wavelengths, spectrometers generally have two lamps,
one generating light in the ultraviolet range (200 to 400 nm) and the other
generating light in the visible range (400 to 800 nm) (Figure 19).
Lamps are pone to slow but continuous attrition, and need to be checked
periodically. If the lamp is the cause of the instability of the absorption signal, it
should be replaced. After a new lamp has been fitted, the optics of the system
should be realigned as follows, to ensure that the maximum amount of light
reaches the photocell after passing through the cuvette.
1. Place a cuvette filled with distilled water and a filter in position.
2. Set the meter to a mid-scale reading, roughly 0.3 (50 % transmission).
3. Move each optical component, in turn, very slightly, and check whether
the reading is affected.
4. If necessary, adjust the lamp alignment to obtain maximum transmission.
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Figure 4: Spectra of light emitted by spectrometer bulbs.
In some instruments, it is possible to place a white card immediately in front of the
photocell. A clear image of the lamp filament should be seen on the card. If the
image is out of focus, or not vertical, the lamp alignment should be adjusted until
the best image is obtained.
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Monochromators are used to disperse the white light into its different light
components, one of which is selected for photometric measurement. Two types of
monochromator exist: the prism and the grating (Figure 20). The grating is often
referred to as a spectral grating or a defraction grating.
Figure 5: Dispersion of white light by (a) a prism, and (b) a transmission grating.
In spectrometers, the monochromator must be correctly aligned. This can be
checked by observing the absorbance maxima of a known reference solution, or
reference absorbing material. Table 1 shows the specific absorbance
values of a potassium dichromate solution (60 mg in 1 liter of 0.005 mol/liter
aqueous sulfuric acid).
Table 4: Specific absorbance values of a standard potassium dichromate solution.
Wavelength (nm)
Specific absorbance (A11)a
124.5 ( ±1.6)
144.0 ( ±1.6)
48.6 ( ±1.6)
106.6 ( ±1.6)
a. The specific absorbance (A11) is defined as the absorbance of a 1% (10g/liter) solution of
the solute in a cell with 1 cm path-length
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A holmium oxide glass filter can also be used as a reference absorbing material; it
has major peaks at the following wavelengths: 241.5 nm; 279.4 nm; 287.5 nm;
333,7 nm; 360.9 nm 418.4 nm, 453.2 nm, 536.2 nm, and 637.5nm.
Filters absorb light of different wavelengths allowing only light with a narrow
range of wavelengths (band width) to pass through. Filters for a photometer have a
bandwidth of about 2 nm, while filters for a colorimeter
are of lower quality and have a wider bandwidth (20-40 nm). Thus more light will
pass through the filter of a colorimeter and the same sample solution will give a
higher reading than in a photometer. The difference in the quality of the filters
explains why a colorimeter is less sensitive than a photometer, and why the
calibration factor for measurement with a colorimeter is higher than that for a
standardized measurement with a photometer. Calibration factors for a specific
method, using a specific reagent, may be given by the manufacturer or may be
found in the literature. However, calibration factors are usually only valid when the
measurement is made at the specified wavelength in a photometer with narrow
bandwidth. The calibration factor must be determined separately when
measurement is made in a colorimeter, or when measurements are compared with a
calibration curve produced from a different instrument and with a different filter.
Filters may be fixed or removable. If the filters can be removed, they should not be
left in the photometer when it is not in use, but should be stored in a dust-free box
to ensure that they cannot be broken. They should be fitted into position before the
lamp is switched on so that the photocell is not damaged.
Diffraction (Spectral) Gratings
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Colorimeters and spectrophotometers, which are used to measure the light
transmitted and absorbed as it passes through a sample, typically use diffraction
gratings to break the light source into individual colors, as defined by their
frequencies and wavelengths.
A diffraction grating consists of slits, or openings, on an opaque plate, as
illustrated in Figure 23. The slits are spaced D meters apart. For each color there is
an angle (Ø), with respect to the plane of the grating, in the direction of which path
lengths from adjacent slits differ by integer numbers of wavelengths.
Therefore the color experiences constructive interference along that path, which
makes it visible to an observer. In Figure 23, white light containing all colors is
incident on the left side of the plate.
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Figure 6: A diffraction grating showing an incident beam of white light, and a ray of
monochromatic light emerging.
An observer moving around a circle as indicated on the right side of the plate
would see violet, yellow, or red, depending on his or her position. A flame
photometer may be constructed with photodetectors at each of those positions.
They are interconnected to the potassium, sodium, and lithium channels of the
flame photometer.
To find the directional angle Ø, observe that the difference in path length to an
observation point from adjacent slits in Figure 23 may be defined as M, where M is
an integer 1, 2, 3, etc.
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It is left as an exercise (Exercise 7) to show that:
M λ = D cos Ø
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for values of λ for which there is constructive interference at the observation point.
In other words, the angle of Ø at which an observer sees the color having a
wavelength λ may be computed from Equation (6).
In order to cause light of different wavelengths to take a path through the sample
under test to the photodetector, it is possible to rotate the diffraction grating on its
axis, while holding the detector in one position. By measuring the detector output
versus its angle of rotation, it is then possible to record the sample absorption
versus light wavelength, or color. This data then leads to the identity of these
substances and their concentration.
The spectrophotometer, as shown in Figure 29, measures light absorption by a
liquid substance at various wavelengths. From this, the components of an unknown
material can be determined or the concentration of a number of known substances
can be measured.
A monochromator uses diffraction grating or prism to disperse the light from the
lamp (slit S1). The light is broken into its spectral components as it arises from slit
S2 and falls on the sample in the cuvette. Narrower slits give rise to shorter
wavelengths. The angle of the diffraction grating determines light wavelength if all
other parameters are fixed and the mirror reduces equipment size. Light output,
photodetector sensitivity, and sample substance absorption change with
wavelength, and this necessitates zero calibration for each wavelength
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Figure 29: Spectrophotometer – simplified schematic.
The double beam spectrophotometer accomplishes this automatically by beam path
switching (sample to reference) via a mechanical shutter or rotating mirror. The
ratio of path absorbances can then be computed. Figure 30 shows a photograph of a
spectrophotometer with input sample to the left and output graph to the right.
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Maintenance includes calibration adjustment and replacement of light source bulbs
and photodetectors. Also, mechanically rotating assemblies (mirrors, diffraction
grating) will occasionally malfunction. The electronics, however, are very reliable.
Figure 30: Spectrophotometer
A cuvette is the transparent container used to hold the test solution. Cuvettes may
be made out of quartz glass, normal glass, or transparent plastic, such as
polystyrene or polyamide. Most cuvettes are rectangular with an internal pathway
of exactly 1 cm. Occasionally they are cylindrical. Cuvettes made out of quartz
glass may be used with light sources producing visible (wavelength 400 to 800 nm)
or ultraviolet (200 to 400 nm) light. Normal glass and plastic cuvettes may be used
with light of wavelength between 340 nm (preferably 365 nm) and 800 nm, and are
suitable for routine measurements. Plastic cuvettes are usually of poorer quality,
but they are cheaper than glass cuvettes and can be discarded after use. On the
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other hand, glass cuvettes are easier to clean for re-use, and do not deteriorate if
properly handled.
The cuvettes must be scrupulously clean if accurate measurements are to be
obtained. Precipitates on glass cuvettes from protein solutions may be removed by
soaking overnight in a concentrated sulfuric acid/potassium dichromate solution.
Subsequently, they must be thoroughly rinsed with distilled water and left to dry
inverted on a clean piece of absorbent paper. Plastic cuvettes must not be cleaned
in strong acid; use a detergent solution.
These solutions, however, leave a film on the cuvette resulting in an increased
absorbance. Cuvettes should be stored in a dust-free box to prevent scratching.
Remember, the following errors will lead to incorrect measurements:
incorrect size of cuvette,
scratched cuvettes,
damaged cuvettes,
incorrect positioning of the cuvette.
Detector and multiplier
The detector and multiplier are electronic components that may fail as a result of
aging, careless handling of the instrument, or incorrect connection to the electrical
power supply. Faults in the monochromator system or photocell and multiplier
system may be the cause of deviations from linearity in measurements of a dilution
series. Deviations from linearity are also caused by high concentrations of an
absorbent in the solution. Therefore, the correct functioning of a photometer needs
to be checked periodically with special control solutions, such as acid potassium
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The absence of a display response can indicate breakdown of the detector. This can
be confirmed if light can be seen on a white card held in front of the detector. A
new photocell is then required.
Photometric measurement
The relationship between the intensity of light entering and leaving a cuvette filled
with a solution that absorbs light of a given wavelength is described by the
Lambert-Beer law. The fraction of the incident light absorbed is proportional to the
number of solute molecules in the light path, i.e.,
Io = incident light intensity,
I = transmitted light intensity,
c = solute concentration (mol/l),
b = path length (cm),
k = calibration coefficient.
The absorbance is defined as the logarithm of the ratio Io/I/ Theoretically the
absorbance of light may vary from zero (no absorbance) to infinity (complete
absorbance). A photometer is most accurate in the absorbance range between 0 and
1.0, and in good instruments up to 2.0. With a colorimeter, reliable results are
obtained between 0 and 0.7.
The constant k is a fundamental property of the solute, and is dependent on
temperature, wavelength, and solvent.
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All the cuvettes used for measurement and calibration must have the same
absorbance. As a check, all cuvettes to be used for measurement should be filled
with distilled water and placed into the cuvette holder. After zeroing the
instrument, measure the absorbance of each cuvette; the absorbance should not
exceed 0.01. Cuvettes with a higher absorbance must not be used, or must be
matched with other cuvettes having the same absorbance to eliminate “cuvette
The use of a single cuvette for all the measurements on a series of specimens
avoids “cuvette error”. Fill the cuvette with a blank solution appropriate to the type
of test, and set the instrument to zero using this blank. Then discard the blank
solution, turn the cuvette upside down and shake it to remove the last drops of the
blank solution prior to refilling with the first test solution. The level of the solution
in the cuvette must be high enough to ensure that reflection of light from the
surface does not interfere with the measurement of absorbance. Also, any air
bubbles trapped against the walls of the cuvette must be removed by gently tapping
the cuvette with a finger.
A major source of error in photometric measurement is a drift of the zero setting
during determination of a series of specimens. To avoid errors, the zero should be
readjusted after each 5 or 10 measurements, by measuring the light absorption by a
cuvette filled with distilled water or reagent solution. Never check the zero with an
empty cuvette.
Light absorption is measured in specimens diluted with buffer and with reagent
solution for reaction. All solutions, i.e., the specimen, the buffer, the reagent
solution, and the final mixture after incubation, should be clear. If the final solution
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is turbid, the transparency of the cuvette must be checked and the investigation
The presence of stray light can be determined by measuring the light absorbance
by a substance that has infinite absorbance at that wavelength. The following
solutions/liquids have infinite absorbance over the stated wavelength ranges, and
are suitable for measuring stray light:
Aqueous potassium chloride (12g/l)
175 – 200 nm
Aqueous sodium bromide (10 g/l)
195 – 223 nm
Aqueous sodium iodide (10 g/l)
210 – 259 nm
250 – 320 nm
Aqueous sodium nitrite (50 g/l)
300 – 385 nm
Set the spectrometer at the wavelength to be checked for stray light. Select an
appropriate solution or liquid. Adjust the instrument to infinite absorbance. Change
the wavelength setting to a longer wavelength and set to zero absorbance. Return
to the original wavelength setting and measure the absorbance. Any reading less
than infinity is due to stray light.
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 Never use a cuvette without having run a blank determination. It is possible that
at high concentrations of the analyte the limits for linear measurement are
 When the reaction time in a cuvette is prolonged, the cuvette should be sealed
to avoid evaporation of the solvent; otherwise, the concentration of the analyte
will increase and result in higher light absorption.
 Cuvettes have two optical walls, where the light beam passes through and two
non-optical walls. They should be held only by the non-optical walls.
 The surface of the solution to be measured in the cuvette must be above the
height of the light beam passing through the cuvette; otherwise, light scattering
at the surface of the liquid may change the magnitude of the absorption signal.
 When using semi-micro and micro cuvettes, they must be correctly positioned
in the light path, otherwise light will be partially reflected and a false reading
will be obtained.
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These are portable devices in which the intensity of color developed in a solution
of unknown concentration is visually compared with that of a known
concentration. A good source of natural light is required in order to carry out the
color matching.
When not in use, store the comparator and the standard filter discs in a dark, dustfree container.
Unpacking, siting, installation
Unpack the instrument carefully, and assemble according to the instruction manual
supplied. Keep specialized packaging for possible future use.
Set the instrument on a firm and level bench that is free from vibration and away
from any strong lighting (especially sunlight). The environment should be free of
dust, fumes, and smoke. Tobacco smoke can be a serious cause of deteriorating
optical performance.
Check that the voltage and fuse rating are in accordance with the manufacturer’s
Good practice
 Handle optical components only by the sides, to avoid contamination.
 Use of clean and matched cuvettes is vital if the full performance of
photometers and comparators is to be realized.
 Occasionally soak the cuvettes in mild detergent for a few hours. Rinse
thoroughly with distilled water and invert to dry.
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 Store cuvettes in a dust-free box, and ensure that they cannot scratch each other
by contact.
 For filter photometers and comparators, keep spare filters in a dust-free box,
and ensure that they cannot be broken or scratched.
 Any spillage on or around the instrument should be cleaned up immediately.
 Turn off the lamp after use, to ensure maximum life.
 Do not leave cuvettes in the instrument.
 Ensure that a filter is in position (in filter photometers) when the lamp is turned
on to avoid damaging the photocell.
If good working practices are observed, there are no special precautions.
Special tools/requirements/spares
A general tool kit and lens tissues are required.
source lamps
lens cleaning kit for cleaning optical components
A calibrating filter is required to check the wavelength accuracy of spectrometers.
When the instrument is cool, and with the electricity turned off:
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1. Clean the filters and optical windows with lens tissue.
2. Keep the cuvettes clean.
The window and/or front surface of the photodetector should be inspected
periodically, and cleaned with lens tissues.
Check lamp alignment.
Wavelength calibration (spectrometers only)
By inserting a calibrating filter in the cuvette compartment in place of a normal
cuvette, the wavelength calibration may be checked as follows:
1. Turn the wavelength control slowly and identify the peaks described in
the data sheet accompanying the filter
2. If the instrument is more than 5 nm off calibration, apply the
manufacturer’s instructions for recalibration.
1. If there is no display response, but light is passing through the system,
then change the photocell.
2. If there is no light passing through the system change the lamp. This may
also be necessary if the light signal does not remain constant during
measurement (as indicated by un-reproducible results of extinction
obtained from repeated measurements using the same cuvette.
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2. Emission photometry
Light can be emitted in different ways. Two analytical procedures used in clinical
chemistry are based on light emission. In a method known as flame photometry, the
sample is heated, and in fluorometry, the sample is irradiated with ultraviolet light.
In flame photometry, the sample is heated in an open flame, which results in the
emission of certain narrow-band wavelengths from the excited ions (Figure 8). The
sample is divided into fine droplets by passing oxygen past the opening of a
capillary tube containing the sample – as occurs in an atomizer spray nozzle; a
combustible gas, such as acetylene, is then added. The mixture is burned and the
light emitted is filtered so that only a limited wavelength range, corresponding to
the emission line of the analyzed substance, is incident on a photocell. The current
output of the cell is proportional to the concentration of the substance in the
Flame photometry is used mainly for analyzing sodium, potassium, calcium, and
lithium. Sometimes lithium is used as the calibration substance in the analysis of
the other three substances; in this case, a known amount of lithium is added to the
sample and the light intensity of the substance under analysis is measured relative
to that of the lithium. In this way, errors due to, for example, varying flame
temperature are eliminated.
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Figure 8: In flame photometry, the sample is heated in an open flame and the intensity of the emitted light is
determined in a narrow wavelength range.
Flame Photometers
A flame photometer is used to analyze urine or blood in order to determine the
concentration of potassium (K), sodium (Na), and lithium (Li). Sodium and
potassium are present in normal urine. Lithium is used as a calibrating substance,
unless it appears in the serum or urine because of medication.
The first use of flame as a spectroscopic source is attributed to Bunsen and
Kirchhoff in 1860. Automated flame photometers came into use after 1945.
A flame photometer operates on the same principle as that used for chemical
analysis with a Bunsen burner. A liquid sample, as shown in Figure 24, is aspirated
into the flame by the gas and air mixture.
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Depending upon the chemical content of the sample, the flame has different colors
resulting from high-temperature thermal collisions that force atoms into excited
The actual intensity of the color is quite variable because of changes in the flow
rate of the gas that draws the sample into the flame. Electronic circuitry is used to
compensate for the variations.
The light from the flame in the flame photometer is filtered, and then directed
along individual channels for the three major substances, Li, Na, and K. The
simplest filter consists of colored glass.
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Figure 7: A flame photometer functional diagram.
For medical analysis, the commonly analyzed ions, and the respective colors they
make in the flame as the excited atoms return to the ground state, are as follows:
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Table 5: Commonly analyzed ions.
Wavelength (Å)
Potassium, K
Sodium, Na
Lithium, Li
Violet glass, for example, passes violet color in the flame but rejects the other
colors. It may be considered an optical filter. Glass filters generally cause
relatively high attenuation of the light and tend to get hot. Flame photometers
typically use diffraction gratings to filter the light, as described earlier.
The flame photometer illustrated in Figure 24 has a separate photodetector for each
channel. The photodetector is a reverse-biased diode for which current increases as
intensity of light incident upon it increases. Calibration potentiometers in each
channel are used to calibrate the instrument with known, standard solutions.
Flame photometers are used routinely for the measurement of lithium (Li), sodium
(Na) and potassium (K) in body fluids. More sophisticated instruments can also
measure calcium (Ca).
The output of the Na and K channel is compared with that of the Li channel. In
normal use, the sample to be tested in this instrument should not contain an
unknown amount of Li. Rather, a known, standard amount of Li is added to the
sample. The output Na and K concentrations are calibrated in terms of
Differences with the known Li, called a stock standard. This procedure
compensates for variations in the flame that are common to all channels.
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figure 25: A simplified flame photometer calibration circuit.
EXAMPLE 2: In a flame photometer, the flame and aspirator have to be
calibrated with lithium. The example calibration circuit in Figure 25 is adjusted
such that the voltage Vout is zero when a known concentration of lithium is
aspirated into the flame. In this case, the lithium causes a photodetector output of
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+2V. Calculate the fraction needed to make Vout =0, where the resistance of the
potentiometer from the wiper arm down is α Rout.
Solution: Since Vout must be zero, we can calculate I = 2/50 A = 0.04 A. because Vout = 0, the
current through the 10-kΩ resistor is zero, so I1 = I = 0.04 A. The voltage V1 = –– I (100) = –
4V. Then Kirchhoff’s current law (KCL) at node 2 gives
V2 – (– 4) + V2 – (– 10) +
α (100)
V2 –10
(1– α) (100) = 0
and KCL at node 1 gives
– 4 – V2 + (– 0.04) + – 4 = 0
Solving this equation for V2 gives V2 = – 4.6 V. This value in Equation 7 gives
– 4.6 + 4 + – 4.6 + 10 + – 4.6 – 10 = 0
100 α
100 (1– α)
Solving this for α we have
0.06α2 – 0.260 α + 0.054 = 0
+0.260 + √0.2602 ─ 4 (0.06) (0.054)
2 (0.06)
= 0.2183 or 4.1
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Since the solution 4.1 is greater than 1, it cannot be the correct value, so α =
0.2183. If you repeat this exercise for the case that the resistor between the pot and
V1 is increased to 100 Ω, you will find that no value less than 1 exists. This means
the circuit cannot be balanced. A faulty resistor that changed its value to 100 Ω
could cause such a trouble.
The flame photometer shown in Figure 26 measures the color intensity of a flame
that is supported by oxygen and a specific substance. The basic schematic shows
that a reference gas containing a lithium salt causes a red color to shine on the
reference photodetector through the reference optical filter. A yellow or violet light
from sample sodium or potassium falls on the sample photodetector. Basically, the
flame photometer is calibrated in a manner similar to the colorimeter. However,
continuous calibration can be accomplished by inspiration of air (oxygen to
support combustion) and lithiums. The output is read in units of sodium or
potassium concentration. A photo of a flame photometer is shown in Figure 27.
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Figure 26: Flame photometer -- simplified schematic.
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Maintenance includes calibration adjustment
and replacement of bulbs and
photodetectors. Aspiration devices and
flame chambers occasionally require
cleaning. Electronic failures are usually
Figure 8: Flame photometer.
In flame photometry, an aqueous salt solution is dispersed in air. The salt in the
dispersed droplets is transferred into a gaseous state by heating with a flame, and
then quickly disintegrates into gaseous atoms. Above a critical temperature the
atoms absorb energy which excites the electrons into higher energy states. When
the excited electrons return to their original state, they emit the absorbed energy as
light. The wavelength of the light emitted by each metal is characteristic for that
element. The intensity of the light emitted at the given wavelength is proportional
to the number of excited metal atoms and can be measured with a suitable optical
filter and photodetector.
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This principle – also called flame emission photometry – can be used to measure
more than 50 elements. However, it is mainly used for determination of the alkali
metals, the excitation of which requires only a low energy input from a lowtemperature flame (propane/air or butane/air).
Under ideal circumstances, there is a linear correlation between the concentration
of the element in dispersion and the light intensity at a specific wavelength.
However, in practice, there will be some degree of ionization, depending on the
concentration of the element. Additionally, the presence of other elements may
suppress ionization. Both atoms and ions of an element in the gaseous state can be
excited, but the emission spectra from atoms and ions are different. Therefore, it is
necessary to choose conditions of measurement in which only atom emission
spectra are obtained. The optimal conditions must be defined for each element.
When measuring potassium in a specimen, for example, the addition of another
element, such as lithium, to the solution not only suppresses potassium ionization,
but can at the same time provide internal calibration, if the lithium concentration is
measured with a reference detector. A flame photometer with a reference detector
for lithium therefore compensates for fluctuations in the energy input from the
A flame photometer consists of the following essential parts (Figure 28):
filter and optics
detector and multiplier
air compressor
gas supply.
The specimen solution is mixed with a diluent, which may contain a lithium salt at
a defined concentration. Many flame photometers have a dilutor as an integral part
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of the instrument. This diluted solution is dispersed by compressed air in the
nebulizer chamber, and heated in a gas flame. The emitted light passes through the
optical system to the detector.
The filter wavelengths for measurement are:
Li – 671 nm
Na – 589 nm
K – 768 nm
With a well-maintained flame photometer, the coefficient of variation in
measurement can be as low as 1.5% for Li and 0.5 % for Na and K.
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Figure 28: Flame photometer assembly
In addition to a reliable electricity supply, flame photometers require:
 compressed air at a pressure of about 100 kPa, from either an electrically
powered air compressor or a gas cylinder (but this is expensive);
 a gas cylinder connected to the flame photometer through an air filter, or
a gas supply (city gas, butane, or propane), as specified by the
 distilled water;
 drainage point.
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Unpacking, siting, installation
Read the manufacturer’s instructions carefully.
Remove the instrument from its packing and wipe the external surfaces with
cloth. Keep the packing for possible future use.
Metal gas lines should not be used for acetylene, except at the expressed
recommendation of the gas supplier. Stainless steel is acceptable, whereas copper
pipes and soldered joints are not. Narrow-bore lines should be used, as acetylene is
less stable in wide-bore pipes.
Operating gas pressure should not exceed 100 kPa. Only reducing valves that are
specifically approved for acetylene should be used.
Special care must be taken with the storage and transportation of cylinders.
Acetylene is so dangerous that it should not be used in the hospital environment
unless there is no alternative available.
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Fluorometry is performed by illuminating the sample at a wavelength that is
absorbed by it (absorption peak) and measuring the excited intensity at a different
wavelength (fluorescence peak). At low concentrations, the intensity of the emitted
light is proportional to that of the absorbed light; that is, the ratio between
fluorescing and absorbed light intensity is independent of concentration; at higher
concentrations, however the relationship is more complex. The calculation of the
concentration of the substance under analysis is therefore more complicated than in
flame photometry. Usually, determinations are made after calibration with standard
solutions of known concentrations.
Fluorometry is used for analyzing substances present only in low concentrations,
for example, certain hormones and vitamins.
Figure 9: Principle of fluorometry. The filter F1 transmits only the wavelength band at which the sample has
an absorption peak. The filter F2 transmits only the wavelength band corresponding to the fluorescence peak.
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3. Review Questions:
1. Two methods of optical analysis often used in clinical chemistry are
____________________________ and _____________________.
2. In absorption photometry, the substance under analysis takes part in
a chemical reaction, in which a compound is formed that has a high
____________________ within a certain wavelength range.
3. The wavelengths usually lie within the _________________range,
but they can also lie within the ______________ or ____________
wavelength range.
4. Two types of emission photometry are used in clinical chemistry,
___________________ and ________________.
5. In flame photometry, the sample is sprayed into a flame and the
light emission is determined within a narrow ________________.
6. The light intensity in flame photometry bears the following
mathematical relation to the concentration of the substance:
7. In fluorometry, a determination is made of the light intensity
emitted by the sample, when illuminated with light of a different
8. Fluorometry is used mostly for analyzing fluorescing substances
present in low concentrations, for example, _________________
and __________________________.
9. Flame photometry is used for analysis of ______________,
_________________, _______________ and _________________.
10. The analytical method in most common use in clinical chemistry
is __________________________.
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K. Methods of Electrochemical Analysis
Among the methods of electrochemical analysis are potentiometry,
polarography, and electrophoresis.
Potentiometry. One form of potentiometry is the earlier described method
for pH measurement in which the potential is determined across a glass
electrode with a selective sensitivity for hydrogen ions.
In the same way, several other ions, such as calcium and potassium, can be
analyzed with ion specific membranes. The measuring circuit employed is similar
to that shown in Figure 7. For each decade change in ion activity, a potential
change of about 60 mV is obtained for monovalent ions, and about 30 mV for
divalent ions.
An advantage of the method is that of the total concentration only the
physiologically active part is measured. A disadvantage is that the specificity of the
membrane is not ideal, that is, other ions than that measured add to the potential,
resulting in a certain error.
Some organic compounds can also be analyzed by such a potentiometric method
by detecting an ionic product of a specific enzyme reaction. For example, urea can
be determined by measuring with an ammonium electrode the ammonium ion
released during breakdown by the enzyme urease.
Polarography. A polarographic method for determining oxygen tension was
mentioned earlier. It is based on measurement of the current when a certain
potential is applied to a platinum electrode in contact with the sample.
A similar method can be utilized for analyzing a number of organic compounds by
means of specific enzyme reactions. A coenzyme or a reaction product is
determined polarographically on a noble metal electrode, and they are an indirect
measure of the substrate itself. Thus, the redox reaction accompanying the
enzymatic breakdown of the substrate causes a change in the current at the
electrode. The change in current is a linear function of the concentration of the
analyzed compound if the enzymatic reaction is made to proceed in a first order
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For example, glucose can be analyzed by means of glucose oxidase and lactate by
cytochrome b2 as enzymes. The method has the advantage of requiring little
preparation of the blood sample and this results in a short analysis time; this is of
importance in certain clinical conditions involving critically ill patients.
is a method for separating and analyzing macromolecular substances such as
plasma proteins. The method is based on the fact that the molecules carry electric
charges and therefore migrate in an electric field. One form of the procedure is
zone electrophoresis, which is illustrated in Figure 10. The charges are due to
basic and acidic groups on the surface of the protein molecules which, in a given
environment, give the molecules a certain net charge. The magnitude and sign of
the net charge determines to some extent the migration velocity in an applied
electric field. The migration velocity is also dependent on the viscosity of the
solvent. The pH of the solution has an indirect influence, since it determines the
degree of dissociation of the charged groups and thus also the net charge on the
protein molecules.
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Figure 10: Principles of zone electrophoresis.
A substance 1 migrates in time t through a distance 1 in a field of field strength E:
1 = u iE • t
where ui is a constant, characteristic of each substance, the electrophoretic
mobility. Experimental measurement of this quantity can be used for classifying,
for example, plasma proteins.
Electrophoresis can be performed in different ways. In free electrophoresis only
buffer solution is used as a medium for the electric field. One disadvantage is that
the separation is countered by convection movements in the liquid, caused by local
density gradients. The convection can be reduced by having the electrophoresis
occur in thin layers – for example, in a filter paper saturated with buffer solution,
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so-called paper electrophoresis, or in a gel, gel electrophoresis. These procedures
have the advantage that when separation is complete, the substances are easily
drawn off from the paper strips or collected fractions of the gel.
Another variant of the procedure is immunoelectrophoresis, in which the separated
macromolecular substances are identified by antigen-antibody reactions.
Review Questions:
1. Three methods of electrochemical analysis in clinical chemistry are
_________________, ______________, and _________________
2. Calcium ion activity determination with an ion specific membrane
is an example of ______________________.
3. In polarography, a quantitative measurement is performed by
measuring the _________________ (current passing through,
potential across, resistance of) an electrode immersed in the test
4. In electrophoresis, macromolecular substances are analyzed by
determining their _____________________ in an electric field.
5. The macromolecular molecules migrate through the electric field
because they have a net charge due to the presence of __________
and ______________ groups.
6. The net charge is dependent on the _____________ of the solution.
7. In electrophoresis, the distance that a substance migrates is proportional
to the ______________, _________________ and
8. Free electrophoresis has the disadvantage that the separation process can
be disturbed by ____________________.
9. To reduce convection, two other forms of electrophoresis can be used,
namely _______________ and _______________.
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L. Chromatography
Chromatography is a method for separating closely related chemical
substances. It is based on differences in the migration velocity of the
substances between a stationary phase and a mobile phase. The difference
in migration velocity is due to a difference in solubility in the two phases.
The stationary phase consists of a solid substance or a liquid, which is rendered
immobile by means of a porous solid medium. The mobile phase is either a liquid,
liquid chromatography, or a gas, gas chromatography. The two methods are used
for separating substances that can be brought into solution or into gaseous form.
The sample is added to the mobile phase; the various components will travel faster
in the mobile phase if their solubility is lower in the stationary phase. The different
components can be characterized by the rate of flow:
Rf =
where vi is the velocity for the component I, and vs the velocity for the mobile
The principle of chromatography is shown in Figure 11. The column may consist
of a glass tube packed with a substance capable of taking up a large amount of the
stationary phase. In liquid chromatography, water is generally used as the
stationary phase; this can be supported by absorption in a suitable medium – for
example, cellulose.
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Figure 11: Principle of chromatography. The components in the sample separate as a result of their different
migration velocities when the mobile phase passes over the stationary phase.
In gas chromatography, the stationary phase is a liquid with a high vaporization
temperature in an inactive porous medium. In liquid chromatography, the mobile
phase is an organic solvent, and in gas chromatography it is usually helium. After
the sample is introduced and passes through the system, the components are
separated according to their relative solubility in the mobile and solid phases. The
process can be followed by means of a detector placed in the outflow of the
column. There are various detection techniques – for example, those based on
measurement of thermal conductivity or density, or on photometric principles.
With liquid chromatography, many substances can be analyzed, for example,
amino acids and sleeping pills (intoxication). Gas chromatography is used for
analyzing substances that can be vaporized, for example, steroids and aromatic
acids in the urine, which appear in many diseases (for example, phenylketonuria, a
congenital disorder that must be diagnosed at birth so that treatment can be
introduced before brain damage results).
Review Questions
1. In chromatography, methods of separation and analysis are based
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on the difference in ________________ between a mobile and a
stationary phase.
2. Differences in migration velocity are due to the difference in
__________________ of substances under analysis in the stationary and
mobile phases.
3. A quantitative measure of migration velocity is the quantity Rf
defined as the ratio between the velocities for the __________
and the _________________.
4. Based on the form in which the sample travels through the column,
two forms of chromatography are distinguished: _________ and
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Most hospitals have a laboratory area separate from patient areas that is used
solely for chemical analyses and measurements of body fluids and tissues.
Typically, analysis is done on blood and urine and on body tissue. These
measurements are made to aid physicians in the diagnosis of disease states
and to help them monitor the effects of therapy. For example, when oxygen
therapy is given to a patient, it is necessary to frequently monitor the partial
pressure of oxygen and carbon dioxide in the patient’s blood. Accuracy of
measurement is critical in the laboratory. And because the chemical
parameters are sensitive to environmental factors such as temperature, light,
and humidity, equipment calibration must constantly be monitored. The
chemical solutions are often active, and cause aging and wearing effects that
may impair accuracy. These effects are most clearly evident in the chemical
Chemical Electrodes
Chemical electrodes produce a potential that depends on the ionic concentration of
fluids under test. These potentials are determined by the Goldman equation
(Equation 1) or the Nernst equation. These equations show that these potentials are
directly proportional to temperature. Ions that are of particular interest in body
fluid analysis include calcium, potassium, sodium, lithium, and the chlorides.
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Chemical electrodes range in complexity from a simple solid in contact with the
solution under test to multiple-element structures. A chloride membrane electrode,
for example, consists of a silver chloride membrane on the tip of a
glass tube. A wire from one side of the membrane connects to a high-impedance
voltmeter. The membrane potential is a function of the chloride content of
perspiration on the skin in contact with the electrode.
The Goldman equation, Equation 1, shows that electrolyte membrane potentials are
proportional to the logarithm of the ion concentration. In a solution containing the
hydrogen ion, a membrane separating two solutions has a potential proportional to
the hydrogen [H+] ion concentration. For example, at 25 degrees C,
Vm = -60 log [H+] + C
(in mV)
where C is a constant. Usually, pH meters are calibrated so that the effect of this
constant is cancelled. The pH is a measure of the hydrogen ion concentration and is
defined as
pH = -log [H+]
Vm = 60 pH + C
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In a pH meter, illustrated in Figure 1, the constant C is compensated for by
calibration so that the meter scale is proportional to pH. Vm is also proportional to
the absolute temperature, T, and ranges from zero to several tenths of a volt. The
pH electrode consists of a reference terminal and an active terminal. The reference
terminal uses a metal, in this case silver-silver chloride (Ag-AgCl), in a potassium
chloride electrode. A salt bridge consisting of a fiber wick saturated with KCl is
inert to the solution under test. However, it maintains the KCl at the potential of
the solution and keeps the reference terminal potentials essentially the same
regardless of the solution under test. The active terminal is sealed with common
glass except for a tip made of pH-sensitive glass. The pH-sensitive glass consists
of a hydrated gelatinous glass layer. Its membrane potential is proportional to the
log [H+] and therefore is proportional to the pH of the solution under test. The pHsensitive glass very slowly dissolves in solution, taking as long as several years to
become ineffective. In general, all boundary potentials in the electrode except
those across the pH sensitive glass are independent of the solution, so long as the
temperature remains constant. The temperature is held constant in a pH meter by
thermal compensation circuits that control the temperature of the sample chamber,
as shown in Figure 1.
The care of pH electrodes includes cleaning to avoid contamination by the sample
solution. They must be replaced periodically because they are a wearing element.
To compensate for unknown variables, the meter is calibrated with a solution of
known pH when used for tests in the clinical setting.
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Figure 9: A pH meter
pH electrodes are wearing elements that may fail because of aging
The pH electrode is also a building block of other chemical electrodes. In
particular, it is a component of blood gas analyzer electrodes.
The concentration of hydrogen ions in a solution, which is conveniently expressed
in terms of its negative decadic logarithm, pH, is often measured during the
preparation of reagent solutions or buffer systems, and also in clinical blood gas
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For the measurement of hydrogen ion concentration, a glass electrode is used in
most pH meters. Glass electrodes are made from special types of glass that allow
hydrogen ions to be absorbed and to penetrate into deeper layers of the glass. Other
small ions such as lithium and sodium ions may also be absorbed to some extent
and change the properties of the electrode, causing the so-called “salt error.” Glass
electrodes are suitable for measurement in the range of pH 0-11. In solutions with a
pH above 11, the salt error becomes important and results in readings of pH that
are too low. In an ideal electrode, the potential difference is 59.1 mV/pH unit at
25˚C. This value is used to calibrate a pH meter in terms of pH units. Since
potential difference depends on temperature, pH measurements should be made at
the calibration temperature; otherwise, a correction factor must be applied.
Calomel (dimercury chloride) electrodes are used as reference electrodes. They
establish a constant potential in an aqueous solution, which is independent of the
pH in the solution. Their potential alters only at very low pH (pH<1), and then only
For the calibration of a pH meter, special buffer solutions must be used; the pH of
the buffer should be near the pH of the solution to be measured. Phosphate buffers
and acetate buffers are preferable. Problems may occur with alkaline buffers, since
their pH may decrease with the absorption of CO2 from the air. This is why all
calibration buffers must be sealed during storage.
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Blood gas analyzers and pH meters should be located in a clean environment, away
from any area where dangerous or corrosive chemicals are stored. For the
calibration of blood gas analyzers, supplies of oxygen and carbon dioxide are
required, and they should therefore be sited appropriately.
Electrodes should be used as recommended for the specified purpose, and the
recommended procedure for equilibrations should be followed when new
electrodes are installed. New glass electrodes must be soaked in a buffer solution
(pH 4-8) for at least 24 hours before use to obtain a stable potential. They should
be calibrated at two values, using the manufacturer’s calibration materials or
solutions prepared from pH buffer tablets. If these are not readily available, buffer
solutions should be prepared for the purpose. However, these solutions should be
checked from time to time against the manufacturer’s calibrants or solutions
prepared from pH buffer tablets.
Manufacturer’s calibration solutions should be used to calibrate blood-gas
analyzers and the usual clinical/chemical quality control procedures should be used
to monitor day-to-day performance.
Keep instrument clean.
Cover after use.
Rinse electrode after use. For short-term storage, it may be kept in a plastic
beaker filled with distilled water to prevent damage.
Check contact between electrode, plug, and instrument.
Avoid contact between electrode and glass beaker.
If applicable, remove the rubber stopper during measurement and refit to
electrode after use.
Make sure that the electrode is always filled with electrolyte according to the
manufacturer’s instructions.
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 Do not touch the electrode membrane, since it can be easily damaged.
 Glass electrodes should be kept immersed in a standard salt solution during
storage for short intervals, and should be stored dry, with a protective cap, for
long-term storage. Glass electrodes that have been stored dry must be soaked in
0.1 mol/liter HCL for at least 4 hours. Thereafter, they must be carefully
washed with distilled water.
 Calomel electrodes must be kept in a potassium chloride (KCL) buffer solution
after use. They must always contain some KCL crystals.
 Protein precipitates on the electrode must be carefully removed by digestion
with pepsin solutions at pH 2 for a few hours. Thereafter, the electrode must be
rinsed thoroughly with distilled water.
Glass electrodes will usually maintain their properties for many years if used
appropriately and stored correctly. Aging of an electrode is indicated when a
constant potential does not develop within a few seconds after insertion into an
ionic solution.
A calomel electrode should be considered unsatisfactory if two calomel electrodes,
when filled with the same KCL solution, differ by more than 5 mV when measured
in the same pH meter.
In modern blood-gas analyzers, in which the electrodes are an integral part of the
instrument, some of these precautions will not be applicable.
If electrodes are stored incorrectly the membrane may dry out. It is sometimes
possible to restore performance by removing the outer layer of the membrane with
a smooth flat file.
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If electrodes are used for measuring biological fluids, they must be cleaned and
disinfected according to the manufacture’s recommendations.
Electrodes are very fragile and must therefore be packed correctly, following the
manufacturer’s instructions. Care is needed in removing the packaging.
pH/Blood Gas Analyzers
Acid-base balance of the blood is generated by body electrolytes and measured by
a pH meter. The respiratory system provides an immediate buffer to sudden blood
pH changes and the renal system provides a slower, more long-range balance
adjustment (protection).
An early pH meter is shown below in Figure 10. An electrode is dipped into the
solution under test (bottom) and read on a meter nulled by a precision dial. The pH
reading is taken from the dial.
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Figure 10: An early pH meter
The glass pH electrode is the heart of the pH meter. Acidity or alkalinity is
indicated by the concentration of hydronium ions (H3O+) in solution. This
gives rise to hydrogen ions (H+). pH is a measure of this ion concentration and is
defined as:
pH = log10 1 = log10 (H+)
pH is acidity
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(H+) is hydrogen ion concentration
Pure water has a pH of 7 (neutral, [H+] = 10 –7). Stomach acid has a pH of 1.5
(greatly acid, [H+] = 10 –1.5). Blood has a pH of 7.36 to 7.44 (slightly alkaline. [H+]
= 10 –7.4 average).
The glass electrode shown in Figure 3, consists of a platinum wire immersed into a
highly acidic buffer solution contained within a thin glass bulb. This bulb wall is
0.1 mm thick and has an electrical resistance of 1000 M Ω. It passes hydrogen ions
only and thus acts as a membrane for separating out these ions. Also immersed into
the test solution is a calomel reference cell (mercurous chloride). The platinum
wire electrode generates a half-cell electrical potential that acts in combination
with the stable reference calomel half-cell. The test solution is in common with
both half-cells. The resultant voltage is amplified by a high input impedance
amplifier such as a MOSFET input op-amp (10 12 Ω) differential input impedance).
Op-amp baseline drift (dc voltage stability) is important since the glass electrode
signal is low-level dc (50 mV). These amplifiers are often chopper stabilized,
which returns the amplifier to ground potential at a relatively high rate (1 KHz).
Blood gas analyzers measure the partial pressure of oxygen and carbon dioxide
(PO2 and PCO2) in solution. This gives an indication of respiratory function. The
carbon dioxide electrode is known as the Severinghaus electrode. It has a thin
Teflon CO2 permeable membrane over the glass bulb of the pH electrode. Final
readings are reached quickly. pH values are measured and compared to pH values
of standard calibration solutions with partial pressures of 60 and 30 Torr (mm Hg).
This is done on a nomograph and is called the Astrup method. Essentially, partial
pressures correspond to specific pH values and, thus, PCO2 electrodes are actually
modified pH electrodes.
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The oxygen electrode is composed of a thin platinum wire and a reference silversilver chloride (Ag-Ag CL) electrode. The magnitude of a small current generated
by a battery connected across the electrodes is proportional to the oxygen
concentration of the solution. Modern measurement techniques use the Clark PO2
electrode. This consists of a platinum wire and Ag-Ag CL electrode mounted
inside a glass housing containing a saturated potassium chloride (K-CL) solution.
A polythene
membrane, semi-permeable to oxygen molecules, covers an opening at the bottom.
A current driven by a battery indicates PO2.
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Figure 11: Basic pH glass electrode.
A combination electrode can also be designed which measures blood pH and gases.
This is known as a Clark-Severinghaus electrode assembly.
Modern blood gas analyzers are precision devices that typically measure blood pH,
PCO2 and PO2. One is shown in Figure 4. The blood micro system is a thermostated
unit incorporating a micro pH electrode (40 µl sample), a microtonometer for four
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disposable equilibrium tubes, and a high-value suction pump. The glass electrode
is seen at the middle and the suction nozzle at the upper left.
Figure 12: Blood gases/blood analysis system.
A Blood Gas Analyzer
A major function of blood is to carry oxygen to the cells and carbon dioxide to the
lungs for expiration. These gases mix with the blood to form a partial pressure in
the blood.
A blood gas analyzer is used to measure the partial pressure of oxygen PO2, the
partial pressure of carbon dioxide, PCO2, and the blood pH. The fundamental
measurement from which the others are derived is the pH measurement. In normal
blood this must be maintained between 7.34 and 7.44, slightly basic. pH readings
from 0 to 7 are acidic, and readings from 7 to 14 are basic. Since pH measurements
are very dependent on temperature, that parameter is carefully regulated to normal
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body temperature, 37º C. A pH electrode, as illustrated in Figure 3, may be used in
a blood gas analyzer. Again, because errors in measurement can cause a dangerous
misdiagnosis of disease, careful calibration procedures are followed.
The CO2 Electrode
The pH electrode is used as a component of a PCO2 electrode to measure the
partial pressure of CO2 by the arrangement shown in Figure 5. Blood or
another fluid to be measured enters a sample chamber and comes in contact
with a Teflon or silicon rubber membrane. This membrane separates the fluid
from a sodium solution but is permeable to the CO2 in the solution. The CO2
combines with water so as to produce free hydrogen ions in the sodium
solution. This changes the solution pH in proportion to the partial pressure of
CO2 in the blood.
figure 13: A cell for measuring PCO2.
A chemical reaction in the electrode is
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H2O + CO2 = H2CO3 → H+ + HCO3
Notice here that the CO2 is proportional to the hydrogen ion, H+. The pH meter
than measures pH-sensitive membrane voltage, Vm, which in turn is proportional to
the pH. The voltmeter is calibrated in units of CO2. The electrode is maintained at
body temperature with temperature-regulating circuits. The input impedance of the
electrode ranges from 50 to 1000 MΩ; therefore a high-input-impedance voltmeter
is required. One form of the CO2 electrode is called a Severinghaus electrode,
named for its developer.
The O2 Electrode
The O2 electrode, known as a Clark electrode, in honor of its inventor, is an
oxygen sensor for blood. It consists of two chambers separated by a
polypropylene membrane that is permeable to oxygen. The blood sample is
injected into the lower sample chamber, as illustrated in Figure 6. The upper
chamber contains the electrode. The O2 in the blood permeates the polypropylene
membrane and reacts chemically with a phosphate buffer contained in the upper
chamber. The buffer maintains the solution pH at a constant level. The O2
combines with water in the buffer, producing electrons in proportion to the number
of oxygen molecules according to the formula
O2 + 2H2O + 4e ¯ → 4OH¯
The electron current, I, is measured by the ammeter A in Figure 6.The electron
current is proportional to the PO2. The electrons on the left side of the equation that
drive the reaction are provided by a source voltage, Vs, that polarizes the electrode
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and has a value between 0.4 and 0.8 V. The need for this polarizing voltage gives
rise to the name polarographic electrode for the Clark electrode. The meter scale is
then calibrated in units of partial pressure of oxygen (PO2) in the blood.
This electrode depends on current flow rather than membrane potential, as was the
case with the pH-electrode-based devices. As with the other electrodes, procedures
for calibration for environmental effects, electrode aging, corrosion, and
contamination must be developed and followed.
A blood gas analyzer that measures blood serum pH, PCO2 and PO2 is illustrated in
Figure 7.
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Figure 3: The Clark electrode for O2
Figure 14: The Clark Electrode for PO2
Figure 15: A blood gas analyzer with an electronic display and hard-copy printout.
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The sample is drawn into the instrument through a capillary. The measured
quantities appear on a front panel display and are printed on paper to provide hard
copy. In order to calibrate the instrument, two known solutions of pH 7.384 and
6.840 are used. After each sample, a flushing solution cleans the tubing and sample
chambers and ejects the waste into the container shown. The laboratory technician
in Figure 8 is injecting samples for automatic chemical analysis.
Non-invasive Blood Gas Monitoring
The blood gas analyzer described in the previous section makes measurements on
blood samples drawn from a patient and carried to the instrumentation. The process
has the disadvantages of discomfort to the patient in drawing the blood and a time
delay in obtaining the data. Real-time, immediately available measurement of PO2
and PCO2, as well as oxyhemoglobin saturation SaO2, can be achieved by use of
non-invasive transducers applied directly to the surface of the skin. Such
monitoring instrumentation is especially valuable in the OR, where immediate
knowledge of the patient’s oxygen and carbon dioxide is of critical importance.
Figure 16: A laboratory technician
analyzing samples.
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It has long been known that both O2 and CO2 diffuse through the skin, as well as
through the alveoli of the lung. Although the diffusion is minimal, it can be
increased significantly by heating the skin. Therefore, to measure the PO2, the
Clark electrode in Figure 6 can be modified by placing it in a heating coil that
heats the skin. The polypropylene membrane is placed against the skin, and O2
passes through it into the electrode, where it can be measured as described earlier.
In a similar manner, the PCO2 electrode may also be modified. Its silicon rubber
membrane is then placed against the patient’s skin so that real-time PCO2
measurement can be made.
The PO2 can also be measured optically by using the fact that oxygenated blood
tends to be red, and low-oxygen blood tends to be blue. About 98 % of oxygen in
the blood combines with hemoglobin (Hb) to form oxyhemoglobin (HbO2). The
ratio of HbO2 to Hb in the blood is called the percentage of oxyhemoglobin
saturation (SaO2). SaO2 also is related in a known manner to PO2 for given values
of blood pH and temperature. To measure SaO2 optically, two light-emitting diodes
are used side by side to illuminate the tissue, such as a fingertip. One light is red
and the other is near infrared. The absorption of the red light is very dependent on
the SaO2, and the infrared light is independent of this value. Therefore the ratio of
the light intensity as detected on corresponding photodetector diodes can be used
to drive an output display calibrated to give the SaO2 value. A finger transducer for
monitoring blood gas is shown in Figure 9.
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Figure 17: The saturated blood oxygen (SaO2) and pulse rate are measured noninvasively
through a finger.
In addition to measuring the SaO2, the instrument shown in Figure 10 measures the
end tidal CO2 (ETCO2). This instrument is called a pulse oximeter, end tidal
Figure 18: A device for monitoring O2 and CO2
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The pH blood gas monitor gives accurate pH, PCO2 and PO2 determinations on the
blood sample or other body fluids. It is designed such that less frequently used
controls are covered by a hinged front panel. Essentially, the blood sample can be
measured for a preset (delay function) time following the push of the measure
button. The unit will measure pH (15 s typically), PCO2 (30 s typically), and PO2 (50
s typically) and then display the values. The values shown in the photograph are
within normal ranges. Calibration can be accomplished easily. This unit contains a
combination of stable analog amplifiers and many digital control and storage
This unit measures hemoglobin concentration and oxygen saturation. Whole blood
or erythrocyte concentrate is aspirated into the system, where hemolysis (breaking
open of RBCs by 40 kHz ultrasonic energy), measurement, calculation, and rinsing
are performed automatically. A precision colorimeter is used to measure
hemoglobin concentration.
Neonatal oxygen monitors are commonly found in nurseries to continuously
monitor environmental oxygen concentration inside the incubator. The one shown
in Figure 12 monitors infant ECG, heart rate, and incubator PO2.
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A fully automated photometric analyzer is shown in Figure 11.
Figure 19: Hemoximeter photometric analyzer.
A transducer is placed next to the child and a manually selected alarm setting
permits warning of low or high PO2.
Figure 20: Gaseous oxygen analyzer.
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Maintenance of pH meters and blood gas analyzers includes frequent calibration
adjustment and replacement of glass electrodes. These electrodes age and require
increasing times to produce accurate readings. For continuous PO2 monitors,
electrodes must be periodically cleaned and occasionally replaced. Electronic
failures are relatively
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The autoanalyzer sequentially measures blood chemistry and displays this on a
graphical readout. As shown in Figure 14, this is accomplished by mixing, reagent
reaction, and colorimetric measurement in a continuous stream.
The system includes the following elements:
1.Sampler – aspirates samples, standards, and wash solutions to the autoanalyzer
2.Proportioning pump and manifold – introduces (mixes) samples with reagents to
effect the proper chemical color reaction to be read by the colorimeter. It also
pumps fluids at precise flow rates to other modules, as proper color development
depends on reaction time and temperature.
3.Dialyzer – separates interfacing substances from the sample material by
permitting selective passage of sample components through a semi-permeable
4.Heating bath – heats fluids, continuously to exact temperature (typically 37
degrees C incubation equivalent to body temperature). Temperature is critical to
color development.
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Figure 21: Autoanalyzer system.
5.Colorimeter – monitors the changes in optical density of the fluid stream flowing
through a tubular flow cell. Color intensities (optical densities) proportional to
substance concentrations, are converted to equivalent electrical voltages.
6.Recorder – converts optical density electrical signal from the colorimeter into a
graphic display on a moving chart.
The heart of the analyzer system is the proportioning pump. This consists of a
peristaltic (occluding or roller) pump. Air segmentation in the mixing tube
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separates the sample/reagent mixture from the cleaning fluid and other samples
(Figure 14). As these air-separated fluids traverse the coil of the mixing tube,
effective mixing action is achieved.
The Technicon SMA 12/60, shown in Figure 15, is a sequential multiple analyzer
that performs 12 different tests on 60 samples per hour. It is a continuous flow
process that produces a chemical profile read on a graphic chart.
Figure 22: Technicon SMA 12/60.
Figure 15: Technicon SMA 12/60.
A later computerized version is shown in Figure 16. This is the Technicon SMAC.
Up to 40 different tests can be performed on an individual serum sample.
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Figure16: Technicon SMAC.
One problem with automatic analyzers is certain identification of samples. Patient
data can be intermixed with other patients if care is not taken.
Maintenance on autoanalyzers includes frequent calibration adjustment. Most
problems are mechanical (tubes, moving pump pats) and electrical (switches,
motors). Electronic failures are few. Sophisticated autoanalyzer system
maintenance and repair requires that the BMET has gone through manufacturer’s
schools. Operation and service manuals must always be consulted. A patient’s life
may hinge on accurate measurement results obtained by clinical instrumentation.
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Pipettes, autopipettes, and dispensers
Pipettes are instruments that are used for transferring a predetermined volume of
liquid from one vessel to another. They are not connected to a reservoir. There are
so many types of pipette that is difficult to discuss the subject systematically. It
should be noted that the replacement of broken conventional calibrated pipettes is
often very costly, and that it may be cheaper in the long run to use mechanical
Mechanical pipettes
Mechanical micropipettes (Figure 31) can only be recommended where a reliable
supply of new disposable tips is readily available. They are used for the delivery
and/or dilution of biological samples in the volume range 5 – 1000 µl. They are
usually of air displacement (indirect) or direct displacement design. To avoid
contamination between consecutive samples, most pipettes have a disposable tip
that is discarded after each delivery. This greatly increases the cost per test. The
practice of washing and reusing disposable tips is not recommended, as any
cleaning procedures will change the “wettability” of the plastic. In addition, drying
at only slightly elevated temperatures may distort the tip, and prevent a good
pneumatic seal with the pipette body.
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Figure 31: A mechanical micropipette
Alternative sample pipettes
Any system that requires mouth pipetting of biological samples is unacceptable
because of the high risk of infection from accidental aspiration of contaminated
material. Thus, the traditional shell-backed pipettes used with a hemocytometertype tube and mouthpiece should never be used.
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Sanz pipettes
A pipette that meets the requirements of safe handling and precision is the Sanz
pipette. It is available in two forms; one is for the accurate measurement of
samples (Figure 32a), the other is for the repeated delivery of reagents in the range
5 – 100 µl (Figure 32b).
Figure 32: Sanz pipettes
Sanz pipettes have a high precision (coefficient of variation 0.5% in the range 5—
100 µl), are very robust, and can be made locally.
Dispensers are instruments for delivering predetermined volumes of liquid from a
reservoir. The reservoir may be an integral part of the instrument, or connected
externally. Dilutors are instruments for taking up different liquids (e.g., sample and
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diluent) and delivering them together in a predetermined ratio and/or
predetermined volume. The reservoirs or the diluent may be an integral part of the
instrument, or connected externally. (Figure 33)
Maintenance and repair
It is virtually impossible to give helpful general advice on the maintenance and
repair of dispensers and autopipettes because there are so many different types.
The manufacturer’s instructions and recommendations should be followed.
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When the specimen is being mixed with a reagent and buffer solution, the
appropriate pipette (or pipette tip) must be used for each individual step of the
procedure. Pipetting by mouth should be forbidden because of the biological and
chemical hazards. A small rubber bulb (Peleus ball) with two valves (Figure 34)
should be fixed to the top of the pipette. The pipette is held vertically while being
filled by suction. The position of the bottom of the meniscus on the pipette scale
indicates the exact volume (Figure 35). When the solution is expelled, the pipette
must also be held vertically. It should be kept in this position for 5 seconds after
the outflow of the last drop. After use, semi-automated pipettes must be kept in an
upright position and thoroughly cleaned periodically.
Figure 34: Peleus ball
Figure 35: Use of a glass pipette
Semi-automated pipettes should be calibrated every 3 months, using appropriate
dye solutions.
Plastic pipette tips must be tested for air-tightness. If there is not a good seal,
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the solution will leak out. Cheap pipette tips are often of poor quality, and
therefore useless. A single pipette tip can be used for serial pipetting of a
solution. Although the same tip may be used repeatedly for the same
solution, it must be replaced before different solutions or samples are
Testing and calibration
Distilled water
Analytical balance
Thermometer (readable to 0.1ºC)
Barometer ( ± 25 mbar)
Weighing vessel (10 – 50 times the test volume, with cover or cap).
When verifying the performance of an instrument, pipetting must be
repeated at least 10 times to estimate accuracy and at least 30 times to
estimate within-run precision. For subsequent control evaluations, the
estimate for within-run precision should be made after pipetting at least 10
times, and the estimate of accuracy after pipetting at least 4 times. The
general procedure is based on gravimetric analysis of water samples
delivered by the instrument. The values are corrected for evaporation. True
mass and volume are then calculated simultaneously, based on the density of
water at specific temperatures, and corrections for air buoyancy.
Note: For safety reasons the use of mercury for gravimetric calibration
should be discouraged.
1. Deliver a total of n samples into a covered weighing vessel and weigh
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each sample after delivery. Replicate as precisely as possible all motions
time intervals in each sampling cycle. Use a randomly selected pipette tip either
only once for each sample weighing or repeatedly for the n weighings.
2. Measure a control blank by dispensing a pre-tared volume of water into
the covered weighing vessel, to estimate the degree of evaporation under the
experimental conditions. Then duplicate all motions and time
intervals as in normal pipetting, with the exception that no more liquid is added
to the weighing vessel. Use the resultant mean loss of weight as the correction
value for evaporation.
3. Measure and record the temperature of the water to 0.1°C before and after the
weighing procedure. The temperature (t) is the average of the
two measurements of water temperature, rounded to the nearest 0.5 C.
Calculate the mean volume (V) delivered at the test temperature (t) from the
mean weighing result (w) by adding the mean evaporation (e) and correcting
the sum by an appropriate factor that allows for density and buoyancy
corrections when water is weighed in air, at the test temperature and
pressure, and standard humidity.
1. Calculate the individual weighing results (wi) by subtracting the tare reading
from the sample reading for each sample.
2. Calculate the mean weight (  ) from the individual weighings (wi)
 wi
where n = number of samples.
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Quality Control
What is QC?
Process of error detection; monitor allowable amount of error; using material with known assayed values
or unknown unassayed values ie assayed-the commercial chemistry control with a pkg insert of known
values for your equipment listed; unassayed-my cbc for the daily control today-must match within %-10%
the values obtained the first time it was run today- may run every four hours or with every 20 patients.
criteria set by lab
Why do we do QC?
To know if results we are releasing are valid; is our equipment working in the expected way
Usually QC is performed daily for chem/hematology; determined by number of samples run; number of
operators; changes in the instrument
Always after calibration and maintenance and if equipment is relocated
correct number with correct rules; most labs use multi-rule Westgard system ie 2 2s, 1 3s, R 4s, most
commonrules used with two levels (normal and abnormal) of control
Determine the expected distribution- normally run 1-2X daily to determine where they should fall;
referred to as normal distritbution
calc mean and SD from data
mean= average
SD= standard deviation from the average
-95% within 2SD 99.7 within 3SD
-the graphing is done on Levy-Jennings chart
-continue to plot values over time to monitor unexpected values
-ideal up to 20 days to establish mean and 2SD limits
-use Westgard rules to monitor performance and accept or reject a set of results.
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- don't make controls values unrealistic; remember what will be medically relevant; same with the rules
- NB keep records
- best if can also participate in external programs
example of out of control chart
What causes errors?
Systematic or random
Systematic error will be detected as changes that occur gradually over time. Random error, on the other
hand, will show an increased scatter about the established mean. The points will be bouncing around on
both sides of the mean outside 3s control limits or 13s control rule)
Accuracy-how close are you to the true value
Precision- how often can you repeat the value
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precise and accurate
Systematic errors may be caused by factors such as a change in reagent lot, change in calibrator lot,
wrong calibrator values, improperly prepared reagents, deterioration of reagents, deterioration of
calibrator, inadequate storage of reagents or calibrators, change in sample or reagent volumes due to
pipettor misadjustments or misalignment, change in temperature of incubators and reaction blocks,
deterioration of a photometric light source, and change in procedure from one operator to another.
Random errors may be caused by factors such as bubbles in reagents and reagent lines, inadequately
mixed reagents, unstable temperature and incubation, unstable electrical supply, and individual operator
variation in pipetting, timing, use of controls, etc.
Systematic errors are most often related to reagent or calibration problems. A sudden shift is usually due
to a recent event such as replacement of reagent, introduction of a new reagent lot number, a recent
calibration, or change in calibrator lot number. When a shift is identified, the operator should inspect the
reagent, calibration, and maintenance records for clues to resolving the problem. For example, if the shift
occurred immediately following a reagent replacement, verify that the lot number is correct and has been
checked out or calibrated, that it has been prepared properly, that it is indeed the correct reagent.
A trend can be more difficult to resolve than a shift simply because the problem is occurring over a longer
period of time. Trends can be the result of a slowly deteriorating reagent, a calibration shift, a change in
instrument temperature, or a deteriorating filter or lamp. Use a systematic logical troubleshooting
approach in isolating the cause, making only one change at a time and documenting each action taken.
In contrast, problems resulting in increased random error are much more difficult to identify and resolve,
mostly due to the nature of the error, which cannot be predicted or quantified as can systematic error.
Many of the sources of random error can be observed by physical inspection of the analytical method
during operation. Careful inspection of reagents and the sampling/reagent pick-up and dispensing
activities will often identify the cause of the problem. If nothing is observed during the inspection
process, consult troubleshooting guides and manufacturer recommendations. If the run is repeated and the
controls are "in" but you feel that you didn't really do anything to "fix" the problem, you may want to
perform a precision run using a patient sample making 10 back to back determinations. This step may
identify further imprecision problems. Duplicate analysis of patient specimens is also recommended when
monitoring random error problems
After the cause of the problem has been identified, it must be corrected and the solution verified by
retesting all of the controls. Place the controls at the front of a run to assess control status. Once incontrol, patient samples from the out-of-control run should be repeated. The out-of-control event must be
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documented along with the corrective action. Troubleshooting reports should be completed for unusual
problems to aid in future problem-solving.
Certain problems occur more frequently with some systems than others. Basic troubleshooting guides,
based on the system operating characteristics, should be developed for each analytical method/system.
Key operators can often recognize the most common problems with a given system and are more skilled
at problem resolution than the infrequent operator. The knowledge of these key operators should be
tapped to identify logical troubleshooting approaches which can be used by all operators.
Resource: J.O.Westgard
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What is coagulation?
Coagulation is the study of clotting factors in the blood. If people are deficient in any of the
factors or the factors do not respond properly, they may bleed to death if they get cut.
Determines the presence of a fibrin clot in a timed mechanical operation.
Characteristics (Temperature, block, reaction well, reproducibility)
Parts: probe and electrode (elliptical sweep)
EXAMPLE Hook picks up initial fibrin thread. 37.2% ± 5 thermostat light off for proper
temperature, Fibrin strand completes circuit and stops circuit. Make sure the 50 HC unit goes to
GHAWA or Figure 17% difference in reading. Fibroplastin from BBL for calibration. Page 18.
It takes three (3) minutes for the temperature to stabilize.
Autopipette starts timer.
Calibrate Fibrotic control plasma Page 19
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Clicking sound occurs. Stops when fibrin is detected. Control time is 11 to 13 seconds.
Clean: lint free paper to clean electrode and distilled water with 10% phosphoric acid. Keep
electrodes parallel.
Prothrombin true test - PTT
Clean and grease pipetter
Critical levels of sample and cuvettes.
Thermostat monitors temperature
Fall of electrode
Electrode alignment.
Erratic end point can be caused by erratic voltage.
Clean cam if timer does not stop.
Schematic review
Probe example
Pt and PTTs
Coag Analyzer
PT 90/hr Prothrombin
APTT activated partial thromboplastin time
TT Thrombin time
FIB Fibrinogen
Uses optical chopper (look for changes in optical density)
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Microscopes are used in medical laboratories to magnify images of light-transmitting or lightreflecting specimens. Examples would be: to see the macro cellular elements in urine such as
cast; the types of white blood cells in leukemia; the type of bacteria in a wound swab.
There are two main types of microscope: light microscopes and electron microscopes.
Light microscopes use glass optics to achieve magnification, while electron microscopes use
electron beams and cathode ray tubes. For normal laboratory work, light microscopes are
sufficient. Their maximum magnification power is usually 1000 times (microscope
magnification = magnification of objective x magnification of eyepiece). A built-in electrical
illuminator or a mirror to reflect artificial light or sunlight onto or through the specimen is used
as a light source.
The light microscope is one of the most important instruments for laboratories in primary health
care. The microscope needs daily attention to ensure reliable laboratory results.
Microscopes have the following components (Fig 2.18):
Stand: tube, tube support, base (foot), and stage.
Optical system: objectives and eyepiece.
Illumination system: light source (mirror or light bulb), condenser, and iris diaphragm.
Fig. 2.18. Light microscope.
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Microscopes must be installed in a clean environment, away from chemicals. Workplaces
should be well ventilated or permanently air-conditioned (intermittent use of air-conditioners
results in condensation). Humidity and higher temperatures often result in the growth of a
fungus that can corrode optical surfaces. Optical instruments should not be kept for long periods
in closed compartments since these conditions encourage fungal growth.
Cleaning of optics
Optical surfaces (condenser, objectives, eyepieces) must be kept free of dust with a soft camel
hair brush or a blower. If dust is found inside the eyepiece, unscrew the upper lens and clean the
inside with the blower or the soft brush.
Oil residues on the leases should be removed with lens paper or absorbent paper or medical
cotton wool. The optics may be finally cleaned with a special solution, consisting of 40%
petroleum ether, 40 % etanol, and 20% ether.
Ethanol (96%) must not be used for cleaning the lenses, since it dissolved the cement. However,
it can be used for cleaning mirrors.
Cleaning of instrument
Heavy contamination can be removed with mild soap solutions. Grease and oil can be removed
with petroleum ether. The instrument should then be cleaned with a 50/50 mixture of distilled
water and 96% ethanol. This solution is not suitable for cleaning the optics.
The mechanical parts (coarse adjustment, fine adjustment, condenser focusing, and mechanical
stage) should be periodically cleaned and lubricated with a drop of machine oil to make them run
Additional precautions to be taken in hot climates
Dry climates
In hot dry climates the main problem is dust. Fine particles work their way into the threads of
the screws and under the leases. This can be avoided as follows:
Always keep the microscope under an air-tight plastic cover when not in use.
At the end of the day’s work, clean the microscope thoroughly by blowing air over it with
a rubber bulb.
Finish cleaning the lenses with a lens brush or fine paintbrush. If dust particles remain on
the surface of the objective, clean it with lens paper.
If there is a wet season lasting more than a month, take the precautions recommended below for hot humid
Humid climates
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In hot humid climates, fungus may develop on the microscope, particularly on the surface of the
lenses, in the grooves of the screws, and under the paint, and the instrument will soon be useless.
This can be prevented as described below.
Always keep the microscope under an air-tight plastic cover when not in use, with a dish filled
with blue silica to desiccate the air under the cover. (The silica will turn red when it has lost its
capacity to absorb moisture from the air. It can be simply regenerated by heating in a hot air
oven or over a fire.) The microscope must be cleaned daily to get rid of dust.
Use and maintenance
More details and useful advice are given in the document, Function, use and maintenance
of routine microscopes, obtainable from Health Laboratory Technology and Blood
Safety, World Health Organization, 1211 Geneva 27, Switzerland. (This document was
prepared by the Zeiss company, and supplied to WHO for free distribution.)
Never dip the objectives in xylene or ethanol (the lenses would become unstuck).
Never use ordinary paper to clean the lenses.
Never touch the lenses with your fingers.
Never clean the supports or the stage with xylene or acetone.
Never clean the inside lenses of the eyepieces and objectives with cloth or paper (this
might remove the anti-reflective coating); use a fine paintbrush only.
Never leave the microscope without the eyepieces unless the openings are plugged.
Never keep the microscope in a closed wooden box in hot humid countries.
Never press the objective onto the slide, since both slide and objective may break. Take
care when focusing the microscope.
Keep the mechanical stage clean.
Do not dismantle the optical components, as this may cause misalignment. The optics
should be cleaned by using lens-cleaning tissue or soft toilet paper.
Never put the microscope away with immersion oil on the objective. Remove any oil
daily. Mild soap solution is suitable for most cleaning.
Organic solvents should only be used in accordance with the manufacturer’s
When changing a bulb, avoid touching the glass with the fingers, as fingerprints reduce
the intensity of illumination.
The life-span of bulbs is extended considerably by adjusting voltage to give the lowest
required light intensity.
If the main voltage fluctuates excessively, use a voltage stabilizer.
In all repair and maintenance procedures, care must be taken not to confuse the condenser
centring screws with the condenser clamp screws.
Check mechanical stage.
Check focusing mechanism.
Remove fungal growth.
Check diaphragms.
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5. Clean mechanical parts.
6. Lubricate to manufacturer’s specification.
7. Check spring load on specimen clamp. Too high a tension may result in breakage
of slides and damage to clamp.
8. Check optical alignment. Dim appearance of the specimen is often due to
misalignment of the optics rather than to insufficient light.
As microscopes are used to investigate biological tissues and fluids, they must be
decontaminated at regular intervals. The mechanical parts may be decontaminated with 70%
ethanol, and the optical parts should be cleaned according to the manufacturer’s instructions.
These procedures must be carried out regularly, and are essential in conjunction with repair and
maintenance procedures.
Rubber-bulb blower
Paintbrush (fine and soft)
Lens paper or cloth for cleaning lenses and optical surfaces
70% alcohol
Q-tips swabs
Pinions, suitable ball and roller bearings
Eyepieces (must be stored under dry and dust-free conditions)
Objectives (must be stored under dry and dust-free conditions)
Greases, oils (see manufacturer’s recommendations)
Fungal cleaning solution
Illumination mirror
Calibrating the microscope for measurement
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Size is an important criterion for the identification of many parasites, particularly cysts and ova.
Size can be determined using a blood cell counting chamber (Neubauer), or alternatively an
eyepiece micrometer. Using an eyepiece micrometer, the procedure is as follows:
1. The eyepiece is divided into 100 small divisions.
2. The stage micrometer scale consists of 1 mm divided into 0.1 mm divisions and each 0.1
mm is divided in 0.01 mm.
3. Insert the eyepiece scale (a round glass disc) into the eyepiece by removing the
uppermost lens and placing the scale on the field stop.
4. Insert the eyepiece into the microscope.
5. Place the stage micrometer on the microscope stage.
6. Focus the low-power objective on the stage scale.
7. Adjust the stage and eyepiece scales until the eyepiece scale and the stage scale are
8. Note the number of eyepiece divisions and its appropriate stage measurement, e.g., 50
eyepiece divisions = 0.75 mm; 10 eyepiece divisions = 0.15 mm.
9. From this reading, work out the value for one eyepiece division, as follows:
50 eyepiece divisions = 0.75 mm
1 eyepiece division = 0.75/50 = 0.015 mm
10 eyepiece divisions = 0.15 mm
1 eyepiece division = 0.15/10 = 0.015 mm.
10. Change the measurement value from mm to μm), e.g., 0.015 mm = 15μm.
11. Repeat for all objectives and note the reading for each.
12. Calibration need be done only once for each microscope used.
Care of the microscope
1. Do keep the microscope covered with a clean plastic or cloth cover
when it is not in use.
2. Do take special care to protect the microscope from dust in hot dry
3. Do take special care to protect the microscope lenses and prisms from
fungal growth in hot humid periods. This can be done by:
 keeping the microscope in an air-conditioned room,
 storing the microscope in a special dehumidified room – an electric
dehumidifier is about half the price of air-conditioner,
 connecting a number of 15 or 25 watt bulbs inside a cupboard with
tightly fitting doors,
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placing a 15 watt bulb in the individual microscope box which then
acts as a warm cupboard,
in areas without electricity, placing a shelf to hold the microscope
box about 30 cm over the chimney of the gas- or keroseneoperated refrigerator of freezer; and airtight bag and silica gel in its
dry state (as indicated by its blue color) will keep a microscope
sufficiently dry to protect lenses from fungi.
4. Do clean the immersion oil from the immersion objective every day;
use a soft cloth dampened with ethanol/ether (3 ml/7 ml) or
benzine/ethanol/ether (2 ml/2 ml/1 ml) and polish with a clean, lintfree cloth.
5. Do clean the oculars with a soft, lint-free cloth; as an alternative, use
lens tissue or facial tissue, if available.
6. Do use the microscope retaining screw fitted at the base of the
microscope box to prevent damage to the instrument while in transit.
7. Do quote the model number and, if possible, the instrument and part
number when ordering replacement parts.
1. Don’t use the tissue or cloth used for the oil immersion objective to
clean the oculars.
2. Don’t use alcohol to clean painted surfaces of the microscope.
3. Don’t dismantle or try to clean parts of the microscope that are
difficult to reach unless you have been trained to do so.
4. Don’t leave the lens ports empty; use the appropriate cover or some
sticking plaster to cover the empty port.
5. Don’t exchange lenses from microscopes of different manufacture –
even some models by the same manufacturer have different
This manual has been produced in conjunction with a two-part video series to assist laboratory
and biomedical engineering personnel in the routine servicing of standard clinical microscopes.
It is not intended as a reference for major repairs.
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Repairs made to a microscope, by an unauthorized service representative, may invalidate the
manufacturer’s warranty. All microscopes under warranty, must be serviced at a manufacturer’s
approved service center.
This manual contains generalized information and is not meant to provide specific information
relative to any particular make or model of microscope. Any individual utilizing this information
is responsible not to violate any of the manufacturer’s guidelines, recommendations, or
provisions of the manufacturer’s warranty.
Laboratory personnel are responsible to insure that the information in this manual does not
violate the requirements for testing procedures performed in the clinical laboratory.
This manual contains information relating to the use of standard clinical microscopes and is not
intended for use relative to specialized areas of microscopy. The enclosed information does not
take into account the use of any specific microscope accessory or the requirements of specialized
laboratory procedures, nor is it intended as a reference source for models of microscopes
produced after 1996.
Blown Fuse
Most microscopes come equipped with some sort of fuse to protect against a short-circuit within
the microscope. If a fuse blows on a microscope, unplug the microscope immediately and
remove it from service.
The primary concern is that the user would simply replace the fuse without determining the
cause of the blown fuse. Fuses are safety devices designed to prevent damage to the microscope
and prevent injury to the microscope user. DO NOT put a microscope back into service after a
fuse has blown, until the source of the problem has been determined and repaired. Have the
microscope examined by your bio-medical staff or outside service company. Lab personnel
should not attempt to make these types of electrical repairs.
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The most common cause of blown fuses are: lamp-socket short, transformer short, or defective
plug assembly. A trained service person with special equipment will be needed to isolate the
Examine the plug of the microscope and look for soot or evidence of a short within the plug
assembly. Replace the plug if necessary.
Always check to make sure that the correct microscope bulb has been installed in the lampsocket assembly. Very often a bulb, that looks externally identical to the correct type of bulb,
has been installed by mistake. This can easily draw excess current which can damage internal
electronics and/or cause a fuse to blow.
Make sure that the correct fuse has been installed in the microscope. Fuses have very specific
ratings for both voltage and amperes. Carefully check the owners manual to determine the
correct fuse type for the microscope. Should an incorrect fuse be installed, there is a possibility
of serious damage to the microscope and a risk of a fire hazard.
Trained service individuals should first examine the inside of the illuminator housing for
evidence of soot from shorts. This should include the lamp-socket, potentiometer switches,
printed circuit-boards, and all wiring. Conductivity checks should be made through the lampsocket assembly with the correct bulb installed. Resistance checks of any potentiometer /rheostat
may indicate possible shorts. Transformer outputs should be checked. Overheating of the
transformer must also be checked.
At all times, follow the instructions in your owner’s manual. Do not attempt any repair that you
are not qualified to perform. Contact the manufacturer or local service representative to have the
microscope repaired.
Purchasing Bulbs
There are several considerations related to purchasing bulbs;
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Purchasing bulbs through large equipment suppliers is usually not cost effective. Prices are
usually inflated 200% to 600% above the wholesale prices. For example, Microscopy USA
will sell some Bi-Pin Halogaen bulbs for under $3.00 each, while large catalog suppliers will
charge $14.00 for the same bulb.
Do not order bulbs from very small suppliers which may have inventory with older bulbs,
which are more likely to fail.
Maintain a good inventory system that rotates stock and alerts the user before the inventory is
Maintain a bulb usage record for your microscopes. This may help troubleshoot electrical
problems within a microscope, or indicate a bad lot number of bulbs received. This is
especially true for expensive mercury vapor bulbs.
Order large quantities when possible. This will usually allow for quantity discounts. Contact
other departments within your home institution to increase discounts.
There are many larger manufacturers who produce bulbs for all types of microscopes. Do not
be afraid to shop around for the best price. Always follow the manufacturer’s guidelines for
bulb selection and usage.
Cleaning Non-Oil Immersion Objectives
If the tips of dry objectives become smudged with immersion oil, finger prints, dried specimen or
slide mounting media, the image quality will suffer greatly.
Dry objectives must be cleaned properly to prevent etching of the lens surface. Microscopy USA
recommends the following procedure for cleaning dry objectives:
1. DON’T DO IT if the image quality is good. DO NOT make the cleaning of dry objectives
part of any regular daily or weekly maintenance program. This type of constant activity will
etch the objective and create more problems than if you left them alone.
3. If there is a problem with the image quality, proceed to clean the
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4. Carefully take a package of lens paper and toss it into the nearest garbage
can. DO NOT use lens paper. Dust or dirt trapped between the lens paper
and the lens will scratch the lens surface. Lens paper will pick up immersion oil from around the edge of the objective and smear it across optical
5. Remove the objective from the microscope by unscrewing it. It is almost
impossible to adequately clean an objective when it is on the microscope.
Carefully examine the lens located on the tip of the objective with the use
of a magnifying glass. Immersion oil, smudges, dried urine, etc., are easily
seen. (Note: As you remove an objective, one or more very thin round
brass shims may come loose that are used to par-focal some objectives.
Save these shims and re-install them onto the threaded end of the
objective before you return the objective tot he microscope nosepiece
after cleaning.)
Cleaning Non-Oil Immersion Objectives (continued)
6. Set the objective down on a countertop with the tip lens pointing up.
Take three cotton tipped applicators and SLIGHTLY dampen them with
an optical grade lens cleaner (not household lens cleaners). Check with
the manufacturer for the recommended type of lens cleaning solution for
your microscope. Hold one of the dampened applicators on the tip of the
objective and rotate the objective three times under the applicator. Apply
only light pressure to the applicator, do not push hard. Repeat this process
with all three applicators. Take a fourth DRY applicator and rotate the
objective ¼ of a revolution to remove any residual lens cleaning solution.
7. Re-examine the objective tip with the magnifying glass to confirm that the
lens surface is clean. If not, repeat this process a second time.
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8. Slidemounting media may present a particular challenge. Contact the
manufacturer for their recommendations to remove slide media.
Generally, a stronger solvent will be necessary to correct the problem.
9. Remount the objective onto the microscope and firmly seat it into the
nosepiece turret.
10. The image quality should greatly improve. If not, the dry objective may
be damaged internally (usually by accidental use of immersion oil on the
objective). The objective may then require repair or replacement.
Microscopy USA will evaluate any defective objectives at no cost to
determine if repair is possible.
Constant Re-Focusing of Image
Clinical microscopes are carefully aligned at the factory so that all of the optical planes are
completely parallel with each other. The optical image plane of the head must be exactly parallel
with the image plane of the objectives, etc.
Occasionally, the components that hold one of these optical elements may become damaged or
mis-aligned. The result will be constant re-focusing of the image as the specimen is moved
across the stage.
Generally, the user should be able to focus a cell under one of the higher magnification
objectives, and then move the image at least the width of three image fields in any direction
without having to adjust the fine focus mechanism whatsoever. Microscopes perfectly aligned
may allow the user to move the slide equal to the width of ten image fields without re-focusing.
[There may be other possible causes to this problem, rule-out parfocality, parcentration and
mechanical stage problems before continuing.]
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In the absence of a simple solution, your microscope may need to be re-collimated. This is a
procedure that will realign all of the components of your microscope to bring the optical planes
into a parallel configuration and center the optical pathway. Expensive equipment and trained
personnel are needed for such a procedure. The microscope will need to be sent to a qualified
service center.
Always ask for an estimate before authorizing work or issuing a purchase order. Microscopy
USA provides free estimates on all work.
Make sure that the microscope is not under manufacturer’s warranty. If it is, only the
manufacturer may perform major repairs.
Keep Your Microscope Covered
In the rush-rush world of life in the clinical laboratory, the practicability of keeping microscopes
covered during periods of non-use seems highly impractical. The fact remains that laboratories
that struggle to meet this goal have microscopes that last longer and function better.
Immersion oil is a non-drying oil that often will penetrate the bearing tracks of moveable stages.
Immersion oil attracts dust and dirt particles that combine with it to produce “sludge” type of
mixture that causes wear on the rack and pinion gears and bearing tracks of most microscopes.
Over many years, this sludge will wear down these surfaces in an excessive way and cause
chronic problems with the mechanical stages and slideholders associated with movement.
Covers have also prevented microscopes from water and smoke damage during fires or the
tripping on of a sprinkler system.
Covers can carry labels pointing night shift or roaming interns toward using a particular
microscope. This can result in only having one microscope totally out of adjustment on any
given morning.
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Covers should be made out of CLEAR heavy-duty vinyl that will allow the microscope to be
picked up and carried safely when the need arises. Custom covers can be handmade to meet the
shape of any microscope. Microscopy USA will produce custom or standard microscope covers
on request.
There are only a few causes of eyestrain from the use of microscopes. As a start, you must isolate
the source of the problem between the eyepiece occulars and the internal head prism optics.
Take eyepieces from another microscope that is not having problems and put them on the
microscope producing eyestrain. If the eyestrain goes away, the original problem was with the
eyepieces, not the internal optics of the microscope head.
The eyepieces of your microscope have several internal components including: lenses, spacer
rings, and lock rings. There is only one way to position these elements that will produce a
comfortable image. Any other combination will always produce eyestrain. Generally, this
problem occurs when someone has taken an eyepiece apart and reassembled it incorrectly. (It
always seems expedient to blame the “night shift.”)
Occasionally, the threaded parts of an eyepiece may work themselves loose, and the internal
elements will become loose. Gently shake the eyepiece. You should NOT hear any sound or
moving or loose parts inside the eyepiece. If you do, perhaps simply tightening the lock ring or
eyepiece casing will correct the eyestrain problem.
If it becomes necessary to reassemble the internal components of an eyepiece, the best place to
start is to find an identical eyepiece that is not having problems. Carefully disassemble the
“good” eyepiece making careful notations and drawing showing the correct sequence and
position of all the internal parts. Please note that the lenses and spacer rings have an “up” and
“down” facing surface. Use this “good” eyepiece as a model to reconstruct the eyepiece causing
the eyestrain. Some owners manuals will show explosion diagrams of the internal lens elements
which may be useful.
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If any lens element seems jammed or stuck in position, do not force it to move. These lenses will
easily chip or crack, which will require the purcase of a new eyepiece.
As if you don’t have enough problems, the optical surfaces of these lenses must be kept
completely free of dust and fingerprints during this process. You will get the lens elements
assembled correctly, only to find the image field covered with artifacts. Keep some optical grade
lens cleaner and cotton-tipped swabs handy for cleaning the lenses as you go.
If switching the eyepieces does not correct the eyestrain, then the internal head optics of your
microscope may have been knocked out of alignment or damaged. This is almost always caused
as a result of the microscope head being accidentally dropped or suffering a concussion type
blow (i.e., mop handle.)
Realignment of the head optics must NEVER be attempted by laboratory or bio-med personnel.
An attempted repair by an untrained individual will result in additional damage and expense!
The head and eyepieces must be shipped to a qualified service center for realignment which will
cost from $300 to $500, not including parts. DO NOT ATTEMPT TO REPAIR LOOSE OR
If the head of the microscope has been severely damaged, prisms may have broken loose inside
the head. Gently rotate the head and listen for any loose pieces inside the head. These must be
removed prior to shipping to avoid any additional damage to the optical elements. All
microscope heads will generally have a set of 4 to 6 small screws, which when removed, will
give you access to the internal spaces of the head. Open the head and stabilize or remove any
loose prisms. Reassemble the head and carefully package the loose prisms for shipment.
Mis-Aligned Head Optics
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The heads of all microscopes have many internal prisms and/or front surface mirrors. These
elements are carefully aligned at the factory. Misalignment of the head optics is usually caused
by the microscope being dropped.
Before beginning, adjust both eyepieces and/or eyetubes to make sure that they are both in focus
with the image field, also resetting them to the central
matched point of adjustment. This may be marked with a white line circling the eyepiece or
To preliminarily diagnose if your microscope has mis-aligned head optics, you will need an
eyepiece that contains some sort of reticle. This can be a crosshair reticle or miller reticle or
reticle micrometer. View the image field only through the eyepiece that contains the reticle.
Focus a small, easily recognizable artifact underneath the crosshairs of the reticle or a part of the
reticle that has a definite pattern. Without touching any of the microscope controls, carefully
move this eyepiece to the other eyetube without rotating it.
The artifact should fall under the identical reticle position in this second eyetube. We
recommend that you use the 40X objective for this procedure. Repeat the process several times
to confirm that there is a problem. There is very little tolerance for the images not to exactly
match. If they are mis-aligned by as little as 1% of the width of the image field, there may be an
alignment problem. If the mis-alignment is 2% of the width of image field or greater, there will
be minor to significant eyestrain.
Mis-aligned head optics will need to be repaired at a qualified service center. If the microscope is
still under warranty, it must be repaired at an authorized service center, otherwise you can send
the head and eyepieces to Microscopy USA for a free estimate. (Generally, head realignment
should cost between $200 and $400.)
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is still under warranty, it must be repaired at a factory service authorized center.
Urinalysis – Set-up Considerations
Every laboratory department has special considerations related to the proper adjustments needed
for their microscopes. Urinalysis testing ha ssome special considerations. One of the microscopic
elements that needs to be identified is Hyaline Casts. These structures are often missed due to the
improper adjustment of the microscope.
Phase microscopy is the recommended type of procedure to eliminate the possibility of missing
transparent Hyaline Casts. Most laboratories, however, use standard bright-field microscopy for
For bright-field microscopes, it is imperative that the iris-diaphragm located just underneath the
stage be adjusted properly. Of this iris is adjusted so that it is 100 % open, excessive light
reflection and poor depth-of-field will eliminate the possibility of seeing Hyaline casts.
Generally, the sub-stage condenser should be adjusted so that it is at the top of its motion, and
the iris should be adjusted so that it is approximately 30 to 50% open. The light intensity will be
less at this position, therefore it may be necessary to increase the amount of light using the
illuminator control potentiometer knob. This 30 to 50 % open position will produce a greater
depth-of-field and the ability to see hyaline casts. If the iris is closed beyond this range, the entire
image field will become very refractile and difficult to read.
Please check the manufacturer’s manual for information related to the proper settings for your
microscope. Do not violate any of the manufacturer’s instructions.
Immersion Oil
Immersion oil for microscopes is a specially prepared oil that generally can be purchased in two
viscosities, commonly labeled “high” or “low” viscosity, or “A” and “B” viscosity. This oil is
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used to contact the specimen and the oil immersion objective. The oil functions to reduce the
angle of light refraction between the objective and the specimen.
There is no optical difference between the two oils, and they can be mixed in any desired
proportions. Generally manufacturers recommend a 50/50 mixture of high and low viscosity oils.
Follow the manufacturer’s guidelines for the proper viscosity oil for your microscope.
Using 100% high viscosity oil may cause a problem in that the slide may be lifted off the
microscope stage while the focus mechanism of the microscope is being adjusted. This may
result in the sensation that the fine focus mechanism is not responding properly.
The use of 100% low viscosity oil may cause the oil to spread out too quickly on the microscope
slide and therefore lose contact between the objective and the slide. This can result in the loss of
sharp image quality. The addition of more oil will correct the poor resolution.
Immersion oil may become contaminated (usually with yeast) when left exposed for long periods
of time in an open bottle. When this occurs, the oil will become turbid. Discard the oil and
properly clean the container.
Specialized immersion oil for immunofluorescent procedures is sold by many manufacturers.
These oils are designed to produce little background fluorescence and are often filtered to
remove dust particles and artifacts. The advantage of these specialized oils is extremely nominal
for most clinical procedures. Use of a filtered oil for Chlamydia Testing may be advisable due to
the pin-point fluorescence indicating a positive result.
All oils purchased recently are PCB free; however, very old stockpiles of immersion oil may
contain carcinogenic PCBs. These supplies must be discarded in an environmentally safe
Light Intensity Not Constant – Halogen and Tungsten Illuminators
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Before attempting any repair to the illuminator of your microscope, make sure that the
microscope has been unplugged and remains unplugged while repairs are being made.
Step number one is to check to make sure that you have the correct bulb type in the microscope.
The wrong bulb can seriously damage your microscope.
If the light coming from halogen or tungsten illuminator appears to surge up and down in
intensity, you must find and correct the problem before there is permanent damage to the
illuminator electronics and/or the transformer.
If there is any light coming from the illuminator, then the fuse has not blown. If the fuse has
blown, then no light will be produced because there is a short-circuit somewhere within the
microscope that must be corrected before the microscope is put back in service.
The most common cause of surging light intensity is oxidation of the electrical contacts between
the bulb and the socket assembly. With the microscope unplugged, clean these contacts until
shiny metal returns. Make sure that the bulb and lamp socket contacts are tight fitting. Loose
fitting or oxidized contacts can produce arcing which may damage your microscope.
The second most common cause of surging light intensity is a potentiometer switch that has
begun to fail. Corrosion of the metal contacts within the switch can cause dead spots to appear at
particular positions or settings of the switch. This problem may require that the potentiometer
switch be replaced. Do not continue to use a microscope when the lamp intensity does not
remain stable.
The third most common cause of surging lamp intensity is the failure of electrical components
within the microscope that regulate the flow of electricity between the transformer and
potentiometer switch. Replacement of these components will be necessary before the microscope
is put back into service.
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If light is produced from the illuminator, but you cannot adjust the intensity, then there has been
a failure within the potentiometer switch or electrical components within the illuminator.
Replacement of these parts will be necessary.
Attempted repair by non-certified personnel may result in damage to the microscope and the
voiding of the manufacturer’s warranty. Do not attempt any electrical repair unless you are
qualified to do so.
Mercury Bulbs and Installation – Clinical Microscopes
Carefully refer to and follow the manufacturer’s service manual prior to attempting the
replacement of a mercury vapor bulb. Do not violate any of the manufacturer’s instructions.
The use of mercury vapor bulbs requires that you consider several safety issues.
Make sure you are using the correct bulb!
Never try to change a bulb while the transformer is plugged in. Always wait 15 minutes
after unplugging the transformer before changing the bulb (capacitors in the transformer
may still hold a current discharge risk.)
Bulbs and lamp housings are HOT even after they have been turned off. Do NOT attempt
to change a bulb unless the microscope has cooled for 15 minutes from its last use.
Never look directly at an ignited mercury vapor bulb.
Make sure that if you are using an L-1 type bulb, that your transformer is set to the L-1
setting. (L-2 setting for L-2 bulbs).
If you crack the bulb during installation, or hear it “pop” while in use, turn off the
microscope and leave the room immediately. Wait 15 minutes before returning. There is
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mercury vapor in the bulb which can be inhaled. After the wair period, follow the
manufacturer’s instructions for mercury spills.
Never get fingerprints on the glass surface of a new bulb during installation.
Mercury vapor bulbs have a definite TOP and BOTTOM end. Installing the bulb
incorrectly will cause poor ignition. Check your service manual for the proper alignment.
Generally, manufacturers recommend that bulbs be changed after 100 to 150 hours of running
time. Follow your manufacturer’s guidelines for bulb life. Keep an accurate log of cumulative
bulb usage. Labs testing for Chlamydia may have to change bulbs more often because of the pinpoint fluorescence which characterizes positive test results.
During installation, bulbs can be easily cracked or damaged. In general, if you are tightening the
clamping mechanism for the top of the bulb – then hold this top mechanism while tightening.
Likewise, hold the bottom clamping mechanism when tightening the bottom of the bulb. This
will avoid putting a torque pressure on the bulb glass, and reduce the risk of breakage.
Make sure that any vacuum seal mark on the bulb is not obstructing the light pathway. This
especially applies to 50W Mercury bulbs.
Once installed, the illuminator housing will generally have three adjusters, one to move the bulb
east/west; one to move the bulb north/south; and one to position a sliding condensing lens used
to align the image.
Additionally, some microscopes have spring loaded adjusters to position a parabolic mirror
located behind the bulb. This mirror will provide a second reflected image of the arc chamber to
increase the intensity of the UV light.
Every microscope manufacturer has specific procedures for the alignment of the mercury bulb.
Follow these procedures carefully. Proper alignment will produce the maximum available UV
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light for testing. Improperly aligned bulbs may result in false negative test results and weak
positive controls.
Parcentration is the ability to move from one objective to another with a particular specimen
element remaining central to the image field.
A user should be able to place a cell or an artifact in the exact center of the 10X image field and
then move to higher magnifications with the image remaining close to the center of the image
fields. (There may be some slight movement of the cell, however, it should stay close to the
center of each objective image field.)
One of the causes of parcentration problems is the fact that an objective may have become
slightly loose in the microscope nosepiece. This will cause the optical alignment of the objective
to drift. Simply check all of the objectives to make sure that all are firmly seated in the
Another cause of problems may be the excessive use of parfocalling shims which will not allow
an objective to properly seat itself in the nosepiece.
There are other major alignment components, which, if mis-aligned, may cause parenctration
problems. The checking of these components requires an experienced service individual.
Parfocality on a microscope is the ability to move from one objective to another with a very
minimal focus adjustment. Generally, the user should be able to move from the 10X to 40X to
100X objectives, while only adjusting the fine focus know less than ½ of one revolution.
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An important exception to this statement will be the low power scanning objectives, 1X, 2.5X,
and 4X. These scanning objectives have very large ranges of “in focus space,” or depth of field.
This fact means that different users will be able to bring the image into focus within a large range
of focus mechanism adjustment. Because of this fact, manufacturers intentionally design these
scanning objectives to NOT be parfocal with the higher magnification objectives. This is done to
prevent the possibility of damage to these higher magnification objectives should specimens be
brought into focus by under scanning objectives that are too close in parfocality. Higher
magnification objectives will be seriously damaged by striking specimens.
The most common cause of parfocality problems is the moving of objectives from one
microscope to another, especially if the objectives are moved to a different manufacturer’s
microscope or a different model of microscope. Objectives have very specific focal lengths and
are rarely interchangeable with other microscopes.
To make minor adjustments to parfocality, very thin brass shims of different thicknesses can be
added to the threaded end of the objective to alter the distance to the specimen, and therefore
bring the image into parfocality. This should be done by a service professional.
Another possibility of a parfocality problem can be the beginning of internal leakage of
immersion oil into an objective. This oil causes an optical distortion which changes perfocality.
If you suspect that this may be happening, have the objective be evaluated for internal oil
Pointer Bulb Life
Microscopes are sometimes equipped with fiber optic cables that will transfer light directly from
the primary illuminator to the pointer assembly. Many models, however, have separate tungsten
pointer bulbs to illuminate the pointer,
If your microscope has a separate pointer bulb, you may notice that it burns out more frequently
than regular illuminator bulbs.
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There are several possible causes for this:
The pointer bulb is left on when the pointer is not in use.
The pointer may be “on,” but the image may have been moved out of the visual field and
forgotten about.
The pointer bulb intensity always set at 100% intensity. (Bulb life is inversely proportional to
usage intensity.)
Someone may have accidentally gotten fingerprints on the bulb during installation. These
fingerprint oils will cook on the glass and reduce bulb life.
The wrong type of bulb may be in use. Many small bulbs appear identical in shape, but often
have different voltage and wattage ratings.
There may be an electrical short within the socket assembly.
Consult your manufacturer’s instruction manual for proper pointer usage and procedures.
Hazy Image Under Oil Immersion
Reduced resolution or image quality for oil immersion images has several possible causes:
1. There must be adequate amounts of immersion oil contacting the oil
objective with the specimen. Add oil and see if the image quality
dramatically improves.
2. Make sure that you are not using the incorrect viscosity of immersion oil. (Refer to the
section identified as “Immersion Oil” in this manual.)
3. Confirm that only this one objective on the microscope is producing a poor image. If all of
the objectives have poor resolution, the problem may be in the head optics or eyepieces of
the microscope as opposed to having a problem with this one particular objective.
4. Make sure that there is not an iris-diaphragm built into the objective. If the outer casing of
the objective rotates, there may be an iris-diaphragm inside. These are usually found on 50X
Oil Immersion objectives, but may be round on others. As these diaphragms are opened and
closed, they have two effects:
The first is that the amount of light reaching the head optics will be
Changing and will cause the image to brighten as it is opened.
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The second effect is that the depth of field, par-focality and image resolution will greatly
change with different iris-diaphragm settings. Find the setting that provides for the
sharpest possible image and still keeps the microscope reasonably par-focal.
5. Occasionally someone may remove an oil objective from the microscope
to try clean the rear lens elements of the objective. This is a big mistake.
Only professionals should be cleaning such sensitive surfaces. A very common problem is
that someone will accidentally get a fingerprint on the rear lens elements which can
drastically affect the image quality. If there is a very obvious and visible fingerprint or
smudge on the rear lens element, remove the fingerprint with a cotton-tipped applicator that
is VERY SLIGHTLY DAMPENED with an optical lens cleaner (If lens cleaner would run
down into the objective, the objective can be permanently damaged.)
6. The objective may be damaged from immersion oil leaking internally into
the objective. All oil immersion objectives have many internal lens elements that are coated
with fluorite film to reduce reflection and refraction as the image passes through. These lens
elements are carefully mounted in a particular position and the tip of the objective is then
sealed with compound that will block penetration of immersion oil from entering the
objective through capillary action. Oil may begin to leak after many years of usage. This can
be caused by damage to the tip of the objective through repeatedly striking the specimen
slides; by dried-out resins in the sealing compound which cause it to leak; or by manufacturer
error in the production of the objective.
If oil has leaked into an oil immersion objective, the image clarity will be sharply reduced. This
can literally happen overnight as capillary action will allow the oil to cover the internal optics.
Repair of these damaged objectives is difficult because the fluorite coatings have probably been
damaged by immersion oil. Replacement or the objective is usually required.
Mechanical Stage Problems – Diagonal Movement
All clinical microscopes are equipped with some sort of moveable mechanical stage to position
the specimen. The mechanical stage may have over a hundred carefully machined parts and is
very detailed in its design.
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The mechanical stage movement will be controlled by a knob which moves the stage in an eastwest direction, and one that moves it in a north-south direction,
Each of these knobs are connected to a set of rack and pinion gears that allow the stage to move.
Each knob assembly has a way to adjust the overall resistance offered by each knob. Every
microscope manufacturer has a different design as it relates to adjusting this knob resistance.
Check your manufacturer’s owner’s manual for specific adjustment instructions related to your
type of microscope. The knobs may be adjusted to many different levels of resistance; tight,
moderate, mild, etc.
It is rather important to adjust these knobs so that both knobs have approximately the same
amount of resistance. (Ex., both knobs with mild resistance). If one of the knobs is adjusted so
that its resistance is significantly greater than the other knob, then the image will begin to move
diagonally when either knob is used. This makes the microscope difficult to use, and is very
dangerous in applications where the entire slide must be systematically screened to examine all
of the specimen elements (i.e., cytology).
Re-adjust the knob tensions so that they are matched in resistance. Follow your manufacturer’s
owner’s manual for specific instructions to adjust the control knob resistance levels.
Mechanical Stage Problems – Noises/Squeaks
All clinical microscopes are equipped with some sort of moveable mechanical stage to position
the specimen. The mechanical stage may have over a hundred carefully machined parts and is
very detailed in its design.
The mechanical stage movement will be controlled by a knob which moves the stage in an eastwest direction, and one that moves it in a north-south direction.
Each of these knobs is connected to a set of rack and pinion gears that allow the stage to move.
These small gears are carefully aligned to travel across each other with just the correct amount of
spacing between the gear surfaces.
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Additionally, the stage will glide in both directions resting on sets of bearings that allow
movement. There will be four individual bearing racks for each stage assembly.
These bearings will usually ride along sets of metal rails called “gib rails.” The gib rails and
bearings are aligned carefully by the manufacturer to provide for smooth action. Special nondrying grease has been used for lubrication.
Often small slivers or chips of broken glass may break free from the corners of microscope
slides and fall into these bearing tracks. The result will be a grinding or scraping type of noise.
Do not despair, clinical microscopes are designed expecting this problem. Within a day the noise
will disappear as the glass is pulverized by the harder metal bearings or is pushed out of the
bearing track by the bearings.
If the noise does not go away, it may be necessary to have the mechanical stage disassembled,
cleaned and reassembled with new grease. Large glass particles can wear down or damage rack
and pinion gears as well as bearing tracks over a period of time. Do not allow this problem to
continue long term.
DO NOT attempt to rinse out the bearing track with a solvent or light weight oil. This will only
dissolve away any remaining lubricating grease and make the problem worse.
Follow the manufacturer’s owner’s manual guidelines. Do not attempt to disassemble the
mechanical stage unless you are trained to do so. Contact a qualified service individual to make
these repairs.
Mechanical Stage Problems – Rack/Pinion Alignment
All clinical microscopes are equipped with some sort of moveable mechanical stage to position
the specimen. The mechanical stage may hve over a hundred carefully machined parts and is
very detailed in its design.
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The mechanical stage movement will be controlled by a knob which moves the stage in an eastwest direction, and one that moves it in a north-south direction.
Each of these knobs is connected to a set of rack and pinion gears that allow the stage to move.
These small gears are carefully aligned to travel across each other with just the correct amount of
spacing between the gear surfaces.
Periodically, the rack gear may come out of alignment. This can happen from the gear being
bumped, or the set screws that hold the gear in place, may have come loose. The result will be
that the mechanical stage will feel “loose” on one end of its motion track and “bind up” on the
other end of the motion track. To correct this problem, the rack and pinion gears of both knob
assemblies will need to be realigned.
This is not an extremely difficult task, but one that does require a lot of patience and experience.
Do not attempt to make these repairs unless you are experienced and qualified to do so.
Another possible cause of these types of problems, will be damage or wear to the rack and pinion
gear assembly. Many years of heavy use may cause the rack gears of microscopes to wear down,
especially in the middle of the range of motion. The result will be that there will be a “looseness”
in the control knob resistance at that point, and the image field may appear to “jump” or
“bounce” as the specimen is moved.
Follow your manufacturer’s owner’s manual for specific instructions to adjust the control knob
resistance levels.
Transformer Housing or Casing of Microscope Seems Hot
Most bright-field clinical microscopes have a step-down transformer built into the body of the
microscope, or as a separate unit. These transformers will always generate some warmth when
current is flowing through them. If, however, the microscope or transformer housing seems “hot”
to the touch, there may be a problem.
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If a microscope housing feels or smells “hot,” turn off the microscope and unplug
immediately. Inform the laboratory supervisor of the problem. Remove the microscope
from service.
Most commonly, such a problem originates from using an incorrect light bulb in the microscope.
Many bulbs appear identical in design, and may or may not have good markings indicating the
voltage and wattage of the bulb. A bulb rated at a higher wattage than designated for the
microscope, if used, can draw excess current from the internal electronics of the microscope and
overheat the transformer. Of special concern are the 6V-20W and 6V-30W Bi-Pin Halogen
bulbs. These bulbs appear to be identical and are often confused. The 30W bulb if placed in
microscopes that require a 20W bulb, may cause serious damage. Place a sticker on the casing of
your microscopes identifying the proper bulb, and inservice the laboratory personnel on the need
for caution. This is a serious problem that should not be ignored. Serious damage to the
microscope can result. In extreme cases, with older microscopes that lack grounding or internal
fuse protection, there can be a fire danger. Install the correct bulb, and examine again for short
circuits, current leakage, or excess heat. If there are no problems, the microscope may be placed
back into service.
If defects are detected, the microscope must be repaired by qualified service personnel. Problems
may be found within the electronic circuitry, lamp socket housing, wiring, transformer,
potentiometer switch, etc. Most manufacturer warranties will become void if non-certified
personnel work on the instrument. Do not attempt any electrical repair unless you are
qualified to do so.
Preventative Maintenance Service Guidelines for Clinical Microscopes
1 Check the previous service record of the microscope.
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2. Place a slid under the microscope to ascertain the current condition of the scope and identify
any problems. Check resolution, mechanical stage, focus mechanism, illumination, etc.
3. Disassemble the microscope.
4. Clean the microscope externally, starting from the top down. Clean easings of head,
condensers, slide holders, etc.
5. Clean oculars, objectives, condenser, iris diaphragms, and re-install on microscope.
6. Perform the basic alignment procedure.
7. Perform a full function check of the microscope including resolution, mechanical stage and
slide holder, par-centration, par-focality, focus mechanism, and illumination.
8. Return scope to end user – reset necessary settings of the end user.
9. Notify the end user of any and all concerns about the microscope.
10. Complete all service records.
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Microscope Maintenance Log
All non-scheduled repair service and
scheduled preventative maintenance
must be recorded below. Service
personnel are required to initial all work
performed. All service personnel are
required to identify the department or
company that they represent.
All records must be dated.
Preventative maintenance will include:
Serial Number
-Eyepieces: remove, inspect and clean
-Objectives: remove, inspect and clean
-Mechanical stage: movement and lubrication
-Iris/Sub-stage condenser: lubricate, alignment
-Electrical Safety check: plus ambient current
-Focus Controls: check for drift and proper motion
-Phase/Polarizer: Alignment
-Parfocality/Par-centration Check
-Collimation check
-Light source: function
Date of service performed by Type of service Comments _______________________
Date of service performed by Type of service Comments _______________________
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Date of service performed by Type of service Comments _______________________
Manufacturers and Suppliers:
Bristoline Microscopes
Accuscope Inc.
8 Roslyn Drive
Glen Head, NY 11545
Fax 1-516-674-3309
E & L Microscope (Cynmar)
Box 498
131 N. Broad Street
Carlinville, IL 62626
Fax 1-800-754-5154
Eagle Instruments (Chinese)
10 Adrian Court
Burlingame, CA 94010
Fax 1-415-697-9207
Edmund Scientific
Fisher Scientific
Instruments Service 1-800-395-5442
Customer Service 1-800-776-7000
Lamp Technology (Secondary Bulb Supplier)
1645 Sycamore Avenue
Bohemia, NY 11716
Leica, Inc. (Leitz, American Optical, Reichert
111 Deer Lake Road
Deerfield , IL 60015
Fax 1-708-405-0147
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LOMO Instruments (Russian)
GEK Dist.
268 Turkey Ridge Road
Charlottesville, VA 22902
Fax 1-804-293-8867
Meiji Stereoscopes
(Capital Instruments)
Microscopy USA
1708 Sanatoga Road
Pottstown, PA 19464
Fax 1-610-327-1016
Web address:
Nikon, Inc.
Instrument Division
1300 Walt Whitman Road
Melville, NY 11747
Fax 1-516-547-0306
Olympus America, Inc.
Two Corporate Center Drive
Melville, NY 11747
Fax 1-516-844-5112
Park Scientific Instruments
1171 Borregas Avenue
Sunnyvale, CA 94089
6 Denny Road, Suite 109
Wilmington, DE 19809
Fax 1-302-762-2847
Rolyn Optical
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706 Arrowgrand Circle
Covina, CA 91722
Fax 1-818-915-1379
Southern Microscope Instruments
120 Interstate N. Parkway East – Suite #308
Atlanta, GA 30339
Fax 1-404-953-4490
Wolfe/Parco – c/o Carolina Biological
Zeiss Microscope Division
One Zeiss Drive
Thornwood, NY 10594
Fax 1-914-681-7446
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Hot-air ovens
Hot-air ovens are used mainly for drying laboratory equipment and surgical
devices in dry air. There are two types of hot-air ovens, with and without internal
circulation of dry air. Only small hot-air ovens can work without internal air
circulation. Sterilization in dry air is less effective than steam sterilization, despite
the higher temperatures applied (Table 6).
Table 6: Dry air sterilization in a hot-air oven.
Temperature (°C)
Time (minutes)
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Figure 48: Temperature variation during hot air sterilization
Use of hot-air ovens
1. Set the thermostat to the required temperature.
2. If there is a fan, check that it is working.
3. Allow to continue heating for the appropriate time after the temperature reaches
the pre-set value.
4. Switch off the heating when time is up.
5. Wait until the temperature falls to 40°C before opening the door.
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Incubators are used mainly for bacterial culture, but have additional uses within the
laboratory. The incubator must maintain a constant temperature (35 ± 2°C for
bacterial culture). Temperature in incubators should be recorded daily. Like all
laboratory instruments, incubators must be cleaned regularly (at least every 14
days) and immediately after any infective material is spilled. Make sure that the
actual temperature corresponds with the thermostat control when the instrument is
used. In carbon dioxide incubators used for microbial culture, the concentration of
carbon dioxide should be maintained at 5-10% and the humidity at 50 – 100%.
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Blood must be stored only in refrigerators that, by design and capacity, maintain
the required blood storage temperature of 1-6°C throughout their interior space.
They must have a system to monitor temperatures continuously and record them at
least every 4 hours, and an alarm system with an audible signal that activates
before blood reaches unacceptable storage temperatures.
Interiors should be clean, adequately lighted, and well organized. Clearly
designated and segregated areas are needed for: 1) unprocessed blood, 20 labeled
blood suitable for allogeneic transfusion, 3) rejected, outdated, or quarantined
blood, 4) autologous blood, and 5) biohazardous autologous blood. Refrigerators
used for the storage of blood and blood components may also be used for blood
derivatives, tissues, patient and donor specimens, and blood bank reagents.
Refrigerators for blood storage outside the blood bank, as may be found in surgical
suites or emergency rooms, must meet these same standards. Temperature records
are required at all times when blood is present. It is usually most practical to make
blood bank personnel responsible for monitoring these refrigerators.
Blood freezers have the same temperature monitoring and alarm requirements as
blood refrigerators and must also be kept clean and well organized. Freezers
designated for plasma storage must maintain temperatures colder than --18°C
(many function at --30°C or colder); RBC freezers must maintain temperatures
colder than --65° (many maintain temperatures colder than --80°C). Self-defrosting
freezers must maintain acceptable temperatures throughout their defrost cycle.
Freezer alarm sensors should be accessible and located near the door, although
older units may have sensors located between the inner and outer freezer walls
where they are neither apparent nor accessible. In such cases, the location of the
sensor can be obtained from the manufacturer and a permanent mark placed on the
wall at that location. Clinical engineers may be able to relocate the sensor
thermocouple for easier use.
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Liquid nitrogen tanks used for blood storage also have alarm system requirements.
The level of liquid nitrogen should be measured and the sensor placed somewhere
above the minimum height needed.
Components that require 20-24°C temperatures can be stored on a tabletop in any
room with an appropriate ambient temperature, provided the temperature is
recorded every 4 hours during storage. Because room temperatures fluctuate,
“environmental” or “platelet chambers” have been developed to provide
consistent, controlled room temperatures. These chambers are equipped with
circulating fans, temperature recorders, and alarm systems, much like blood
In addition, platelets require gentle continuous agitation during storage to facilitate
gas exchange within the bag and to reduce the formation of aggregates. Elliptical,
circular and flat-bed agitators are available for table top or chamber use. Elliptical
rotators are not recommended for use with storage bags made of polyolefin without
1. Continuous Temperature Monitoring Systems
Most blood refrigerators, freezers, and chambers have built-in temperature
monitoring sensors connected to recording charts and/or digital readout systems for
easy surveillance. Digital recording devices measure the difference in potential
generated by a thermocouple; this difference is then converted to temperature.
Because warm air rises, temperature recording sensors are best placed on a high
shelf. They should be immersed in a volume of liquid no greater than the volume
of the smallest component stored. Either a glass container or a plastic blood bag
may be used. Recording charts and monitoring systems are inspected daily to make
sure they function properly.
When recording charts or tapes are changed, they should be dated inclusively (i.e.,
start and stop dates) and labeled to identify the facility, the specific refrigerator or
freezer, and the person changing the charts. Any departure from normal
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temperature should be explained in writing on the chart beside the tracing or on
another document. A chart with a perfect circle tracing can indicate that the
recorder is not functioning properly or is not sensitive enough to record the
expected variations in temperature that occur in any actively used refrigerator.
Blood banks with many refrigerators and freezers may find it easier to use a central
monitoring-alarm system that monitors all equipment continuously and
simultaneously and prepares a hard-copy tape of temperatures at least once every 4
hours. These systems have an audible alarm that sounds as soon as any connected
equipment reaches its predetermined temperature range and indicates the
equipment in question. Blood storage equipment so monitored does not require a
separate independent recording chart.
2. Thermometers
Visual thermometers in blood storage equipment provide ongoing verification of
temperature accuracy. One should be immersed in the container with the
continuous monitoring sensor. The temperature of the thermometer should be
compared periodically to the temperature on the recording chart. If the two do not
agree within 2°C, both should be checked against a thermometer certified by the
National Institute of Standards and Technology (NIST) and suitable corrective
action should be taken. (A 2°C variation between calibrated thermometers allows
for the variation that may occur between thermometers calibrated against the NIST
Thermometers also help verify that temperature is appropriately maintained
throughout the storage space. Large refrigerators or freezers may require several
thermometers to assess temperature fluctuations. In addition to the one immersed
with the continuous monitoring sensor (usually located on a high shelf), at least
one other in a similar container is placed on the lowest shelf on which blood is
stored. The temperatures in both areas must be within the required range at all
Either liquid-in-glass (or analog) thermometers or electronic and thermocouple (or
digital) devices can be used for assessing storage temperatures, as long as their
accuracy is calibrated against a NIST-certified thermometer or a thermometer with
Updated Dec 2007
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a NIST-traceable calibration certificate. Of equal importance is that they be used as
intended, according to the manufacturer’s recommendations.
3. Alarm Systems
To ensure that alarm signals will activate at a temperature that allows personnel to
take proper action before blood reaches undesirable temperatures, both temperature
of activation and power source are tested periodically. The electrical source for the
alarm system must be separate from that of the refrigerator or freezer; either a
continuously rechargeable battery or an independent electrical circuit served by an
emergency generator is acceptable.
Thermocouple devices that function at freezer temperatures are especially useful
for determining the temperature of activation with accuracy when sensors are
accessible. When they are not, approximate activation temperatures can be
determined by checking a freezer’s thermometer and recording chart when the
alarm sounds after it is shut down for periodic cleaning or maintenance. It can also
be assessed by placing a water bottle filled with cold tap water against the inner
freezer wall where the sensor is located. When the alarm goes off, usually in a
short time, the recording chart can be checked immediately for the temperature of
There must be written instructions for personnel to follow when the alarm sounds.
These instructions should include steps to determine the immediate cause of the
temperature change and ways to handle temporary malfunctions, as well as steps to
take in the event of prolonged failure. It is important to list the names of key
people to be notified and what steps should be taken to ensure that proper storage
temperature is maintained for all blood, components, and reagents.
4. Refrigeration
The laws of partial pressure state that, in a space occupied by a mixture of
gases that do not react chemically together, each gas exerts the pressure that
it would produce if it occupied the space alone, and the total pressure is the
sum of these pressures.
When a mixture of ammonia and water is heated by a flame or an electrical
device in the heating chamber, ammonia and a relatively small amount of
water will evaporate. The ammonia and water vapor enter a percolator where
the water is condensed. This water, containing a low concentration of
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ammonia, passes into the absorption vessel. The gaseous ammonia moves to
the condenser. Air circulating over the fins of the condenser cools the
gaseous ammonia, and it condenses. The liquid ammonia then flows by
gravitational forces into the evaporator where it evaporates under low
pressure at ambient temperature. This process extracts heat from the storage
compartment. The evaporation is accelerated by hydrogen gas passing across
the surface of the ammonia. The ammonia-hydrogen mixture travels to the
absorption chamber, where the ammonia is absorbed by the water. This
process occurs so fast that it keeps the partial pressure of ammonia in the
system low and contributes to accelerating evaporation in the evaporator.
The hydrogen passes through the water without being absorbed and back to
the evaporator.
With the absorption of ammonia the liquid in the absorption chamber
increases in density and flows into the heating chamber, from where the
refrigeration cycle is repeated.
5. Compression
Compression systems are used for cold rooms and for some small
refrigerators and require main electricity. They consist of an evaporator, an
expansion valve or capillary pipe, a condenser, and a compressor
(Figure 13).
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Figure 13: Working principle of a compression refrigerator.
A compressor sucks the coolant liquid from the tubes of the evaporator, which are
located inside the cooling compartment of the refrigerator. The residual coolant
liquid in the evaporator evaporates, and in doing so, take sup latent heat from the
cooling compartment. The vapor is compressed into pipes outside the refrigerator,
where it condenses, liberating heat, which is dissipated to the surrounding air by
the condenser fins. The condensed coolant liquid is forced through the capillary
pipe and expands into the evaporator, from where the refrigeration cycle is
repeated. In some refrigerators, the condensed coolant is circulated back to the
compressor to take up heat from the compressor oil, which again causes
evaporation of the refrigerant. In a second condenser, the coolant is condensed
again prior to passing through the capillary tube for expansion and evaporation,
while the liberated heat of condensation is dissipated to the environment.
6. Installation
Electrical compressor-operated refrigerators and freezers should be used only
where there is a stable and reliable electricity supply. Fluctuations in the voltage,
and frequent power interruptions are likely to result in damage to the compressor.
Absorption refrigerators and freezers are preferred in situations where electricity
supply is unreliable.
Equipment should be installed on a flat, horizontal surface, preferably slightly
elevated (on pallet or feet) to avoid accumulation of water and moisture under the
cabinet. This will prevent the formation of rust and allow easy access for cleaning.
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Good practice
 Keep the surrounding area clean.
 Leave at least 20 cm between the cabinet and the wall and other equipment, and
avoid exposure to heat and sunshine.
 Keep the refrigerator upright and level. If the cabinet needs to be moved, it
should be transported in an upright position.
 Wash the door gasket with soap solution and rub with glycerol, when the
cabinet is defrosted.
 Do not re-open the door immediately after closing.
 Never use sharp instruments to remove ice. Defrosting may be quickened by
placing a container of warm water in the refrigerator or freezer after electrical
 Remove all water from the inside of the refrigerator or freezer after defrosting.
 Do not leave the refrigerator or freezer open unnecessarily.
 Open and close the door gently.
Flammable chemicals must only be stored in cabinets designed for that
purpose. Kerosene-operated refrigerators and freezers should be refilled with
uncontaminated kerosene. The burner, chimney and wick must be cleaned
regularly. The baffle must be inserted into the chimney.
7. Maintenance:
The following general advice may be helpful for maintenance:
 The refrigerator must be placed so that sufficient air can flow past the
condenser (at the back of the refrigerator) for exchange of heat.
 The refrigerator door must seal perfectly to prevent warm outside air from
entering the cool chamber.
 The refrigerator must have good insulating walls.
 For photovoltaic (solar-powered) refrigerators, the collector must be positioned
so as to receive maximum solar radiation; it must be cleaned periodically to
ensure the production of enough electricity.
Daily checks
 Check temperature daily
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 Check the gas bottles or kerosene tank, in the case of gas or kerosene
refrigerators, so that more can be ordered in good time.
Monthly checks
 Clear the cool chamber, and defrost the evaporator once a month
 Swab inside the cabinet with 70% ethanol while it is defrosting.
 Clean the outside of the refrigerator.
 Clean any dust from the condenser.
 Clean the door gasket.
 Clean the burner, and check for gas leakage.
 In photovoltaic refrigerators, check the level of electrolyte solution in the
batteries, and fill up with pure distilled water, if necessary.
Door gaskets
On domestic type refrigerators, the gasket-holding mechanism is the inner shell of
the door This fastens to the outer casing with a ring of screws under the gasket.
When this is disassembled in order to change the gasket, the rigidity of the door
structure is lost. In order to ensure a good seal upon reassembly, the complete door
must first be removed and placed on several boards to keep it as flat as possible.
Then, remove the screws and the old gasket, install the new gasket and replace the
screws before moving the door. Reinstall the door, with the hinge screws snug but
not tight. Shut the door with a piece of paper in the seal, and test for tightness by
pulling on the paper. Do this all around the gasket. The hinges may be adjusted
outwards by closing the door with a folded cloth in the seal or by bumping with a
soft rubber mallet. Adjust until the paper indicates that the door is evenly tight all
around, then tighten the screws in the hinges.
Compressor-type refrigerators and freezers
 Clean the condenser (in the compressor compartment) every 6 months
with a brush or vacuum cleaner.
 Oil the door fittings, locks, and other moving parts.
 Replacement of the compressor, which would require recharging with
refrigerant, should be carried out only by a qualified refrigeration
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Absorption-type refrigerators and freezers
 Check the thermostat.
 Check the heating element.
 If the heating element is working but the refrigerator does not become
cool, remove the burner with the tank, or disconnect the refrigerator from
the main. Place upside down for 12 hours, then upright for another 12
hours and re-start normal operation (if the refrigerator or freezer had been
transported incorrectly or tilted, this action will ensure that the ammonia
Refrigerant flows back into the correct pipes.) If this procedure does not
work, send for a qualified refrigeration engineer.
Changing the heating element
Disconnect the refrigerator from the main.
Remove the heating element from the chimney.
Disconnect from the thermostat.
Connect the new element at the ceramic connector or thermostat, using
the same terminals
 Insert the element into the chimney aperture, making sure it is not placed
beside the aperture.
Note: For security reasons, the refrigerant liquid circuit is sealed by the
refrigerator manufacturer. It should never be opened because of the
hazardous nature of the liquid.
Heating elements
Burner glasses
Vacuum cleaner or brush
8. Tips:
The most important thing you can do to prevent future problems with
refrigerators and freezers is to ensure good air flow over the coils which
radiate heat removed from the interior compartments. If yours has a set of
Updated Dec 2007
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condenser coils on the rear, dusting them off annually should be sufficient,
and even that is not critical. Don’t store paper or plastic bags on top of your
refrigerator which might fall down and block air to these coils, however.
Models of this type do need at least 2-3 inches of uncluttered air space above
Them for proper air flow. Models with coils underneath near the compressor
need to have those coils cleaned at least twice a year.
First, remove the kickplate/grill at the bottom front covering the opening
under the lower door. Clean the kickplate with a garden hose to remove dust
and hair. Next, using either an inexpensive condenser coil brush or vacuum
cleaner with crevice tool on the hose (which doesn’t do as good a job), or an
air compressor (which is VERY messy), clean over, under and through the
coils found usually on the right front half behind the kickplate. Older units
are especially sensitive to air flow blockage. Overheated compressors soon
While under there, on frost-free models, removing and cleaning the plastic
defrost drain pan often found there can prevent nasty odors later. Use warm,
not boiling, water, and/or a garden hose. Be sure when reinstalling the pan not to
jam it against the blades of the condenser cooling fan, on those models
with a fan underneath in the rear (most of those without rear wall mounted
cooling coils.)
Other models may have a kind of “jelly roll” black sheet metal with internal freon
passages in the right rear underneath. The condenser brush is the only
good way to clean these, other than a garden hose or an air compressor used
Be sure, if the refrigerator has a rear cardboard cover, to replace it after
service. If it is damaged, cut a new one from a cardboard box. The open fan
cover grill areas are not critical but the solid ones are vital. These covers
force air to be pulled over the hot condenser, cooling it, rather than being
sucked in from open areas at the rear of the refrigerator. Without this cover it
will overheat and may burn out the compressor.
If you have a non frost-free refrigerator or freezer, defrost the freezing
compartment whenever ¼ to ½ inch of ice has built up. With the greater
efficiency, the compressor will run less and last longer. Never use a sharp or
metal instrument to defrost. Pans of hot water, hair dryers, or just time with
the unit turned off and open are the way to go.
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While the rear condenser fans on some models do have oiling holes on the
motors, it is often a fairly difficult job to remove the fan motor for access.
If accessible, and if it has oil hole(s), use a light machine oil (3 in 1 type) or
A zoom-spout oiler to lubricate this motor. If not, they last a very long time
without oiling.
The only things you can do to prevent problems are to check that the door
gaskets are sealing ALL THE WAY around. Often, if not really ragged,
minor gasket air leaks can be patched with razor cuts and silicone caulk
and/or rolled up pieces of duct tape or newspaper, but new gaskets are also
generally available if yours have serious problems. Major air leaks make
your refrigerator work harder, run longer, use more electricity, and, if not a
frost-free, ice up faster.
Placing a thermometer in each compartment is an excellent idea. Then, if
you notice a significant temperature change in either, you will know that you
need to find out why. Problems caught early are usually cheap to fix; wait a
while and you may have major expense.
Stored blood components are inspected immediately before issue for transfusion or
shipment to other facilities. These inspections must be documented; records should
include the date, donor number, description of any abnormal units, the action
taken, and the identity of personnel involved. Visual inspections cannot always
detect contamination or other deleterious conditions; nonetheless, blood products
that look abnormal must not be shipped or transfused.
Contamination should be suspected if:
Segments appear much lighter in color than that of the bag.
The red cell mass looks purple.
A zone of hemolysis is observed just above the cell mass.
Clots are visible.
The plasma or supernatant fluid is murky, purple, brown, or red. Although a
green hue from light-induced changes in bilirubin pigments need not cause the
unit to be rejected, units with grossly lipemic plasma, identified by its milky
appearance, are usually considered unsuitable for transfusion.
 Blood or plasma is observed in the ports or at sealing sites in the tubing, This
suggests inadequate sealing or closure, and the unit should, at the very least, be
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A red cell unit that is questioned for any reason should be quarantined until a
responsible person decides its disposition. Evaluation might include inverting it
gently a few times to mix the cells with the supernatant fluid because considerable
undetected hemolysis, clots, or other alterations may be present in the undisturbed
red cell mass. If, after resuspension, resettling and careful examination, the blood
no longer appears abnormal, it may be returned to inventory. Appropriate records
should be maintained documenting the actions taken, when, and by whom.
In addition, platelets should be inspected for the presence of excessive aggregates.
Units of FFP and CRYO should be inspected, when removed from frozen storage,
for evidence of thawing and refreezing and for evidence of cracks in the tubing or
plastic bag. Unusual turbidity in thawed components may be cause for discard.
F. Bacteriologic Studies
Bacterial contamination of transfusion components is rare, thanks to the use of
aseptic technique, the availability of closed systems for collection and preparation,
and careful control of storage conditions.
Sterility testing of blood or components plays a role in validating initial production
processes. If a transfusion component has an abnormal appearance, or if an adverse
clinical reaction appears to be related to contaminated donor blood, culturing may
be desirable, and a Gram’s-stained smear of supernatant plasma should be
Your Hospital
Blood Bank
Temperature Control Program
Purpose: The temperature of the items tested in this procedure must be recorded
daily or as otherwise stated. These records must be retained for 5 years.
1. On a daily basis, the temperature of this room, instruments, freezer,
refrigerators, and any other items noted on the log sheets are to be
observed and recorded. The tolerance limits for each of the items is
indicated on the log sheet.
Updated Dec 2007
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2. If the tolerance limits are exceeded, the department supervisor should be
notified. If that supervisor is not available, the laboratory manager or one
of the pathologists should be notified. If necessary, follow the procedure
for evacuation of refrigerator or freezer involved.
3. Correctiv4e actions must be recorded and signed by the supervisor when
temperatures exceed acceptable limits. Please use the forms titled
“Quality Indicator Log “ for this purpose.
4. If blood bank graphs are not working correctly, or blood has to be
evacuated to a refrigerator or freezer that does not have a graph, you must
record the temperature every 4 hours. A chart to record the temperatures
is located in this manual.
5. Once per year, the thermometers will be checked against a standard
thermometer. The procedure is in the Maintenance Manual.
6. New thermometers will be checked against a standardized thermometer
before being put into use.
7. The temperature of blood must be maintained below 10° C during
transport. Therefore, if blood is issued in a cooler to the OR, place a
thermometer bag (stored in blood bank refrigerator B at 1-6°C) with the
units of blood under the ice bag. If blood is returned unused in the cooler,
record the temperature and time returned on the “Surgical Blood Cooler
Temperature” chart. A Safe-T-Vue temperature indicator will be attached
to each unit issued to a cooler. If the indicator has a “white” center, the
blood may be returned to the blood bank for re-issue. See the procedure
for using these indicators in this manual.
8. The blood bank alarm system will be tested weekly. (See procedure.)
20th Standards, AABB, 2000
Signature _____________________________________
Review Date: _________________________________
Form 14: Blood Bank Temperature Control Program
Updated Dec 2007
MET Module V
Your Hospital
Blood Bank Manual
Temperature Recording Charts
Seven-day circular charts are used to record temperatures on the Platelet Incubator, Blood Bank A,
Blood Bank B and the Freezers used for blood storage. A 60-day temperature graph is used to monitor
the blood bank refrigerator.
The circular recording graphs must be changed every Friday. Position the chart to start recording on
the correct date and time. The starting and ending time and date must be noted on the chart. Also write
which refrigerator or freezer the graph is monitoring and technologist initials on the chart.
The blood bank temperature-recording chart will monitor for approximately 60 days. The date is
written on the chart each morning.
Observe all temperature monitor charts each morning to determine the accuracy of date and time.
Notify the supervisor of any unusual occurrences.
All abnormal peaks or trends must be circled on the chart. This must be done as close to the time they
occurred as possible and the internal temperature should be recorded on the chart. An explanation of
the event affecting the temperature should be written on the chart. Notify the supervisor of unusual
Alarm checks and repairs are noted on the charts.
All charts will be reviewed weekly by the blood bank tech and monthly by the laboratory chief.
Any alarm or chart failure may require temperature recording every 4 hours. There are charts to be
used in these circumstances can be found in the temperature control manual.
20th Standards, AABB, 2000
Signature ________________________________
Review Date ______________________________
Review Date ______________________________
Figure 15: Blood Bank Temperature Recording Procedure
Updated Dec 2007
MET Module V
Your Hospital
Blood Bank Temperature Chart
Record temperature of refrigerator or freezer used when blood bank
refrigerator or freezer is not running properly, every 4 hours.
Figure 16: Blood Bank Temperature Chart
Updated Dec 2007
MET Module V
Your Hospital
Blood Bank Temperature QC Manual
Testing Blood Bank Alarms
Purpose: To ensure that the alarm systems work even when power is interrupted. This may also
be a test of the remote alarm system, or the remote alarm may be tested manually.
Equipment: Blood Bank Refrigerators (A, B, C, etc.)
Blood Bank Freezer
Platelet incubator
1. The alarm system will be tested once each week on (day).
2. Unplug each individual appliance separately.
a. Unplug each refrigerator and wait for alarm to sound. Check the remote alarm in
chemistry to make sure it is activated. (Do not push the white alarm button under the
remote alarm.)
b. Unplug the blood bank freezer and push the battery test black button. Make sure
the remote alarm in chemistry is activated.
c. Turn off the platelet incubator and lift lid. Wait for alarm to sound. Turn the
incubator back on.
3. Record on alarm check log in this manual.
Signature ____________________________
Review Date _____________________
Review Date _______________________
Figure 17: Blood Bank Alarm Testing Procedure
Your Hospital
Updated Dec 2007
MET Module V
Refrigerator Alarm Test
Purpose: Sensors on the blood bank refrigerator(s) must be checked quarterly.
1. The day the sensors are to be checked, place 300 ml of water in an appropriate
container in the blood bank refrigerator
2. In the afternoon, remove the sensors from the 10% glycerol and
place in the water along with a blood bank thermometer.
3. Using constant agitation, add 15 ml of 20-degree water to the container which has the
sensors in it. This water should cause a 1-degree rise in temperature.
4. Continue with this procedure until the alarm goes off.
5. Record the temperature on quality control sheet for refrigerator alarm. Also write the
word TEST on the graph to explain the fluctuation in temperature.
6. Add ice chips to the water to cause the temperature to decrease.
7. Continue until the low sensor goes off.
8. Record the temperature on the quality control worksheet and write TEST on the
9. Alarm must sound at no less than 1 degree C and no greater than 6 degrees C to be
Signature ___________________________
Reference: AABB Technical Manual, 11th Edition, 1993
Figure 18: Refrigerator Alarm Test Procedure
Your Hospital
Updated Dec 2007
MET Module V
Refrigerator Alarm Test Log
Refrigerator __________________
Year ________________
Low Alarm
must sound no lower
High Alarm
must sound no higher than 6°
then 1 °
Figure 19: Refrigerator Alarm Test Log
Updated Dec 2007
MET Module V
Your Hospital
Alarm Checks (without power) for Refrigerator
Once a week, the alarm system on the blood bank refrigerators, freezers, and
platelet incubator will be checked to make sure that the alarms function even when
the power is interrupted. Unplug the appropriate refrigerator, freezer, or incubator
and follow procedure. The alarm should ring within approximately one minute.
Year ________________
Blood Bank Refrigerator ______________
Jan Feb
May June
Figure 20: Alarm checks (without power) for Refrigerator
Updated Dec 2007
MET Module V
Your Hospital
Alarm Checks (without power) for Freezer
Once a week, the alarm system on the blood bank refrigerators, freezers, and
platelet incubator will be checked to make sure that the alarms function even when
the power is interrupted. Unplug the appropriate refrigerator, freezer, or incubator
and follow procedure. The alarm should ring within approximately one minute.
Year ________________
Blood Bank Freezer ______________
May June
Figure 21: Alarm checks (without power) for Freezer
Updated Dec 2007
MET Module V
Your Hospital
Alarm checks (without power) for Platelet Incubator
Once a week, the alarm system on the blood bank refrigerators, freezers, and
platelet incubator will be checked to make sure that the alarms function even when
the power is interrupted. Unplug the appropriate refrigerator, freezer, or incubator
and follow procedure. The alarm should ring within approximately one minute.
Year ___________________
Platelet Incubator __________________
May June
Figure 22: Alarm checks (without power) for platelet incubators.
Updated Dec 2007
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Figure 23: Blood Bank Maintenance Chart
Updated Dec 2007
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Figure 24: Instrument Repair Log
Updated Dec 2007
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Updated Dec 2007
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How does reverse osmosis work?
To understand "reverse osmosis," it is probably best to start with normal
osmosis. According to Merriam-Webster's Collegiate Dictionary, osmosis
is the "movement of a solvent through a semipermeable membrane (as of a
living cell) into a solution of higher solute concentration that tends to
equalize the concentrations of solute on the two sides of the membrane."
That's a mouthful. To understand what it means, this picture is helpful:
On the left is a beaker filled with water, and a tube has been halfsubmerged in the water. As you would expect, the water level in the tube is
the same as the water level in the beaker. In the middle figure, the end of
the tube has been sealed with a "semipermeable membrane" and the tube
has been half-filled with a salty solution and submerged. Initially, the level
of the salt solution and the water are equal, but over time, something
unexpected happens -- the water in the tube actually rises. The rise is
attributed to "osmotic pressure."
A semipermeable membrane is a membrane that will pass some atoms or
molecules but not others. Saran wrap is a membrane, but it is impermeable
to almost everything we commonly throw at it. The best common example
of a semipermeable membrane would be the lining of your intestines, or a
Updated Dec 2007
MET Module V
cell wall. Gore-tex is another common semipermeable membrane. Gore-tex
fabric contains an extremely thin plastic film into which billions of small
pores have been cut. The pores are big enough to let water vapor through,
but small enough to prevent liquid water from passing (see this page for
more information on Gore-tex fabric).
In the figure above, the membrane allows passage of water molecules but
not salt molecules. One way to understand osmotic pressure would be to
think of the water molecules on both sides of the membrane. They are in
constant Brownian motion. On the salty side, some of the pores get
plugged with salt atoms, but on the pure-water side that does not happen.
Therefore, more water passes from the pure-water side to the salty side, as
there are more pores on the pure-water side for the water molecules to
pass through. The water on the salty side rises until one of two things
The salt concentration becomes the same on both sides of the
membrane (which isn't going to happen in this case since there is
pure water on one side and salty water on the other).
The water pressure rises as the height of the column of salty water
rises, until it is equal to the osmotic pressure. At that point, osmosis
will stop.
Osmosis, by the way, is why drinking salty water (like ocean water) will kill
you. When you put salty water in your stomach, osmotic pressure begins
drawing water out of your body to try to dilute the salt in your stomach.
Eventually, you dehydrate and die.
In reverse osmosis, the idea is to use the membrane to act like an
extremely fine filter to create drinkable water from salty (or otherwise
contaminated) water. The salty water is put on one side of the membrane
and pressure is applied to stop, and then reverse, the osmotic process. It
generally takes a lot of pressure and is fairly slow, but it works
Membrane Processing
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Membrane processing is a technique that permits concentration and separation without
the use of heat. Particles are separated on the basis of their molecular size and shape
with the use of pressure and specially designed semi-permeable membranes. There are
some fairly new developments in terms of commercial reality and is gaining readily in its
proteins can be separated in whey for the production of whey protein concentrate (WPC)
milk can be concentrated prior to cheesemaking at the farm level
apple juice and wine can be clarified
waste treatment and product recovery is possible in edible oil, fat, potato, and fish
fermentation broths can be clarified and separated
whole egg and egg white ultrafiltration as a preconcentration prior to spray drying
The following topics will be covered in this section:
Principle of Operation
Types of Membrane Processing
o Reverse Osmosis
o Ultrafiltration
o Microfiltration
Hardware Design
Ion Exchange
Principle of Operation
When a solution and water are separated by a semi-permeable membrane, the water
will move into the solution to equilibrate the system. This is known as osmotic pressure
If a mechanical force is applied to exceed the osmotic pressure (up to 700 psi), the
water is forced to move down the concentration gradient i.e. from low to high
concentration. Permeate designates the liquid passing through the membrane, and
retentate (concentrate) designates the fraction not passing through the membrane.
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Membrane Processing
Reverse Osmosis
Reverse osmosis (RO) designates a membrane separation process, driven by a pressure gradient,
in which the membrane separates the solvent (generally water) from other components of a
solution. The membrane configuration is usually cross-flow. With reverse osmosis, the
membrane pore size is very small allowing only small amounts of very low molecular weight
solutes to pass through the membranes. It is a concentration process using a 100 MW cutoff,
700 psig, temperatures less than 40°C with cellulose acetate membranes and 70-80°C with
composite membranes. Please click above link for a schematic diagram of these membrane
Hyperfiltration is the same as RO.
Ultrafiltration (UF) designates a membrane separation process, driven by a pressure gradient, in
which the membrane fractionates components of a liquid as a function of their solvated size and
structure. The membrane configuration is usually cross-flow. In UF, the membrane pore size is
larger allowing some components to pass through the pores with the water. It is a separation/
fractionation process using a 10,000 MW cutoff, 40 psig, and temperatures of 50-60°C with
polysulfone membranes. In UF milk, lactose and minerals pass in a 50% separation ratio; for
example, in the retentate would be 100% of fat, 100% of protein, 50% of lactose, and 50% of
free minerals.
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Diafiltration is a specialized type of ultrafiltration process in which the retentate is diluted
with water and re-ultrafiltered, to reduce the concentration of soluble permeate
components and increase further the concentration of retained components.
Please click above link for a schematic diagram of these membrane processes.
Microfiltration (MF) designates a membrane separation process similar to UF but with even
larger membrane pore size allowing particles in the range of 0.2 to 2 micrometers to pass
through. The pressure used is generally lower than that of UF process. The membrane
configuration is usually cross-flow. MF is used in the dairy industry for making low-heat sterile
milk as proteins may pass through but bacteria do not. Please click above link for a schematic
diagram of these membrane processes.
Hardware Design
Open Tubular:
Tubes of membrane with a diameter of 1/2 to 1 inch and length to 12 ft. are encased in
reinforced fibreglass or enclosed inside a rigid PVC or stainless steel shell. As the feed
solution flows through the membrane core, the permeate passes through the membrane
and is collected in the tubular housing.Imagine 12 ft long straws!
Hollow Fibre:
Similar to open tubular, but the cartridges contain several hundred very small (1 mm
diam) hollow membrane tubes or fibres. As the feed solution flows through the open
cores of the fibres, the permeate is collected in the cartridge area surrounding the fibres.
Plate and Frame:
This system is set up like a plate heat exchanger with the retentate on one side and the
permeate on the other. The permeate is collected through a central collection tube.
Spiral Wound:
This design tries to maximize surface area in a minimum amount of space. It consists of
consecutive layers of large membrane and support material in an envelope type design
rolled up around a perforated steel tube.
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Electrodialysis is used for demineralization of milk products and whey for infant formula and
special dietary products. Also used for desalination of water.
Principles of operation:
Under the influence of an electric field, ions move in an aqueous solution. The ionic mobility is
directly proportioned to specific conductivity and inversely proportioned to number of molecules
in solution. ~3-6 x 102 mm/sec.
Charged ions can be removed from a solution by synthetic polymer membranes
containing ion exchange groups. Anion exchange membranes carry cationic groups
which repel cations and are permeable to anions, and cation exchange membranes
contain anionic groups and are permeable only to cations.
Electrodialysis membranes are comprised of polymer chains - styrene-divinyl benzene
made anionic with quaternary ammonium groups and made cationic with sulphonic
groups. 1-2V is then applied across each pair of membranes.
Electrodialysis process:
Amion and cation exchange membranes are arranged alternately in parallel between an anode
and a cathode (see schematic diagram). The distance between the membranes is 1mm or less. A
plate and frame arrangement similar to a plate heat exchanger or a plate filter is used. The
solution to be demineralized flows through gaps between the two types of membranes. Each type
of membrane is permeable to only one type of ion. Thus, the anions leave the gap in the direction
of the anode and cations leave in the direction of the cathode. Both are then taken up by a
concentrating stream.
Concentration polarization. Deposits on membrane surfaces, e.g. proteins - pH control is
important. Prior concentration of whey, to 20% TS, is necessary before electrodialysis.
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Ion Exchange
Ion exchange is not a membrane process but I have included it here anyway because it is used for
product of protein isolates of higher concentration than obtainable by membrane concentration.
Fractionation may also be accomplished using ion exchange processing. It relies on
inert resins (cellulose or silica based) that can adsorb charged particles at either end of
the pH scale. The design can be a batch type, stirred tank or continuous column. The
column is more suitable for selective fractionation. Whey protein isolate (WPI), with a
95% protein content, can be produced by this method. Following adsorption and
draining of the deproteined whey, the pH or charge properties are altered and proteins
are eluted. Protein is recovered from the dilute stream through UF and drying. Selective
resins may be used for fractionated protein products or enriched in fraction allow
tailoring of ingredients
Anyone who has been through a high school science class will likely be familiar with the
term osmosis. The process was first described by a French Scientist in 1748, who noted that
water spontaneously diffused through a pig bladder membrane into alcohol. Over 200 years
later, a modification of this process known as reverse osmosis allows people throughout the
world to affordably convert undesireable water into water that is virtually free of health or
aesthetic contaminants. Reverse osmosis systems can be found providing treated water
from the kitchen counter in a private residence to installations used in manned spacecraft.
Reverse Osmosis is a technology that is found virutally anywhere pure water is needed;
common uses include:
Drinking Water
Car Wash Water Reclamation
Rinse Waters
Biomedical Applications
Laboratory Applications
Pharmaceutical Production
Kidney Dialysis
Water used in chemcial processes
Animal Feed
Metal Plating Applications
Wastewater Treatment
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Boiler Water
Battery Water
Semiconductor production
How Reverse Osmosis Works
A semipermeable membrane, like the membrane of a cell wall or a bladder, is selective
about what it allows to pass through, and what it prevents from passing. These membranes
in general pass water very easily because of its small molecular size; but also prevent many
other contaminants from passing by trapping them. Water will typically be present on both
sides of the membrane, with each side having a different concentration of dissolved
minerals. Since the water i the less concentrated solution seeks to dilute the more
concentrated solution, water will pass through the membrane from the lower concentration
side to the greater concentration side. Eventually, osmotic pressure (seen in the diagram
below as the pressure created by the difference in water levels) will counter the diffusion
process exactly, and an equilibrium will form.
The process of reverse osmosis forces water with a greater concentration of contaminants
(the source water) into a tank containing water with an extremely low concentration of
contaminants (the processed water). High water pressure on the source side is used to
"reverse" the natural osmotic process, with the semi-permeable membrane still permitting
the passage of water while rejecting most of the other contaminants. The specific process
through which this occurs is called ion exclusion, in which a concentration of ions at the
membrane surface from a barrier that allows other water molecules to pass through while
excluding other supstances.
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Semipermeable membranes have come a long way from the natural pig bladders used in
the earlier osmosis experiments. Before the 1960's, these membranes were too inefficient,
expensive, and unreliable for practical applications outside the laboratory. Modern advances
in synthetic materials have generally solved these problems, allowing membranes to
become highly efficient at rejecting contaminants, and making them tough enough to
withstand the greater pressures necessary for efficient operation.
Even with these advances, the "reject" water on the source side of a Reverse Osmosis (RO)
system must be periodically flushed in order to keep it from becoming so concentrated that
it forms a scale on the membrane itself. RO systems also typically require a carbon prefilter
for the reduction of chlorine, which can damage an RO membrane; and a sediment prefilter
is always required to ensure that fine suspended materials in the source water do not
permanently clog the membrane. Hardness reduction, either through the use of water
softening for residential units or chemical softening for industrial use, may also be desirable
in hard water areas.
Low Pressure (Residential) Systems
Low pressure RO systems generally refer to those systems with a water feed pressure of
less than 100 psig. These are the typical countertop or undersink residential systems that
rely primarily on the natural water pressure to make the reverse osmosis process function;
a typical system is shown schematically below.
Typical Point of Use Reverse Osmosis System
Countertop units typically have an unpressurized storage tank; Undersink units typically
have a pressurized accumulator storage tank where the water pressure tends to increase as
the tank fills. This pressurized system provides sufficient pressure to move the water from
the undersink storage tank to the faucet. Unfortunately, this also creates a back pressure
against the membrane, which can decrease its efficiency. Some units overcome this by
using unpressurized tanks with a pump to get the treated water where it is needed.
Low pressure units typically provide between 2 and 15 gallons per day of water, with an
efficiency of 2-4 gallons of reject water per gallon of treated water. Water purity can be as
high as 95 percent. These systems can be highly affordable, with countertop units starting
at about US $150, and undersink units starting at about US $500. These units produce
water for a cost as low as ten cents per gallon once maintenance and water costs are
factored in. Maintenance usually requires replacing any pre- or postfilters (typically one to
four times per year); and the reverse osmosis cartridge once every two to three years,
depending on usage. Look for the WQA Gold Seal (S-300) to find products that have been
successfully tested to industry performance standards; and to Certified Water Specialists
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(CWS I-VI), Certified Sales Representatives (CSR), and Certified Installers (CI) for advice on
your water needs, and equipment installation.
High Pressure (Commercial/Industrial) Systems
High pressure systems typically operate at pressures between 100 and 1000 psig,
depending on the membranes chosen and the water being treated. These systems are
usually used in industrial or commercial applications where large volumes of treated water
are required at a high level of purity.
Most commercial and industrial systems use multiple membranes arranged in parallel to
provide the required quantity of water. The processed water from the first stage of
treatment can then be passed through additional membrane modules to achieve greater
levels of treatment for the finished water. The reject water can also be directed into
successive membrane modules for greater efficiency (see diagram below), though flushing
will still be required when concentrations reach a level where fouling is likely to occur.
High pressure industrial units typically provide from 10 gallons to thousands of gallons per
day of water with an efficiency of 1-9 gallons of reject water per gallon of treated water.
Water purity can be as high as 95 percent. These systems tend to be larger and more
complicated than low pressure systems, and this is reflected in their costs, which range
from US $1000 through tens of thousands of dollars for a large, multi-module unit capable
of providing desalinated drinking water for a resort facility or water bottling plant.
What Reverse Osmosis Treats
Reverse osmosis can treat for a wide variety of health and aesthetic contaminants.
Effectively designed, RO equipment can treat for a wide variety of aesthetic contaminants
that cause unpleasant taste, color, and odor problems like a salty or soda taste caused by
chlorides or sulfates.
RO can also be effective for treating health contaminants like arsenic, asbestos, atrazine
(herbicides/pesticides). fluoride, lead, mercury, nitrate, and radium. When using
appropriate carbon prefiltering (commonly included with most RO systems), additional
treatment can also be provided for such "volatile" contaminants as benzene,
trichloroethylene, trihalomethanes, and radon. Some RO equipment is also capable of
treating for biological contaminants like Cryptosporidium. The Water Quality Association
(WQA) cautions, however, that while RO membranes typically remove virtually all known
microorganisms and most other health contaminants, design consderations may prevent a
unit from offering foolproof protection when incorporated into a consumer drinking water
When looking for a product to treat for a given health contaminant, care should be used to
find products that have been tested successfully for such purposes at a quality testing
Reverse osmosis is a relatively new, but very effective, application of an established
scientific process. Whether it is used to meet the needs of a typical family of four, or the
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needs of an industrial operation requiring thousands of gallons per day, it can be a cost
effective to provide the required quantity of highly treated water. With continual advances
in system and membrane design that boost efficiency and reliability, RO can be expected to
play a major role in water treatment for years to come.
-------------------------------------------------This article first appeared in the WaterReview Technical Brief, (1995) Volume 10, No. 3;
a publication of the Water Quality Research Council; Copyright 1995 by the WQA. All rights
Updated Dec 2007
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Balances are used to measure the weight or mass of a substance. If a comparison is
made between two objects, one known and one unknown, then the measurement is
mass. If the measurement is made against gravitational pull, then the measurement
is weight.
There are two main categories of balance:
-- Mechanical balances
-- Electromagnetic balances
Balances that are based on other principles of measurement (e.g., piezoelectric
balances, magneto-elastic balances, gyrodynamic balances, string balances) are
less frequently used and will not be discussed here.
A number of factors influence the weighing processes. They become more
important the smaller the mass of the substance to be measured. Therefore, the
weighing of small quantities is more prone to errors than the weighing of large
The following influences can cause errors in measurement:
Moisture (atmospheric humidity)
Electrostatic effects
Gravitational forces
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 Vibration.
Many analytical balances, and particularly those measuring in the microgram
range, are constructed to minimize the effects of as many of these factors as
The collective term “analytical balance” describes a balance suitable for chemical
analysis. The weighing range of certified analytical balances is between 10μg and
50 kg. They may be mechanical or electronic. Optical balances are mechanical
balances equipped with an optical read-out
Mechanical balances can be subdivided into:
Spring balances,
Sliding-weight balances
Parallel-guidance-system balances,
Substitution balances:
--three-knife substitution balances
--two-knife substitution balances
With a spring balance, the force of an object is compared with the known force of a
spring. The calibration of a spring balance depends on the gravitational force on
the object, which varies from one locality to another.
Therefore spring balances must be calibrated at their place of use (Figure 49).
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Figure 49: Spring balance
With a sliding-weight balance, the weight of an unknown object or substance is
determined by a sliding device containing a known weight.
Equilibrium is reached by displacement of the sliding weight along the runner, which is
marked with scale divisions (household balance, Roman beam scale) (Figure 50).
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Figure 50: Sliding-weight balance.
In a parallel-guidance-system balance, the degree of displacement of a beam from
the equilibrium position is taken as a measure for an unknown mass. The parallel
guidance prevents the weighing-pan from tipping over (letter balance) (Figure 51).
Figure 51: Parallel guidance balance.
In substitution balances, the sensitivity remains the same during measurement,
since the beam always carries the same load on both sides, no matter what the load
is. Substitution balances include equal-lever-arm
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balances (three-knife balances), and unequal-lever-arm balances (two-knife
An equal-lever-arm balance has a symmetrical lever and three knife edges – one in
the center and one at each end. The difference between two counter-rotating
torques, generated by a known mass and an unknown mass, keeps the beam in a
deflected position, which is taken as a measure for the unknown mass. (Figure 52).
Figure 52: Equal lever arm balance.
Unequal-lever-arm balances (two-knife balances) have one main knife-edge and a
secondary knife-edge, which support both the load and the mass pieces, while a
fixed counterweight is at the other end of the lever. If an unknown mass is placed
on the pan, the balance beam deflects to that side. An appropriate number of
masses must be removed from the side of the beam with the unknown load, so that
the pointer returns to zero (Figure 53).
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Figure 53: Unequal-lever-arm balance.
Electromagnetic balances
With an electromagnetic balance, the unknown mass is loaded on to a wire that is located
between two poles of a permanent magnet, and to which an electrical circuit is applied
(Figure 54). The wire is mechanically displaced by the load. In order to bring the wire back
to its original position, more current must be applied to the wire. The difference in the
current required to keep the wire in the zero position when it is loaded and unloaded is
proportional to, and taken as a measure of, the mass on the pan (Figure 55).
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Figure 54: Electromagnetic balance at rest.
Figure 55: Electromagnetic
balance under load.
Good working practices – all balances
1. The balance must be zeroed prior to each use.
2. Always weigh chemicals/samples in a container, never directly on the pans.
3. Use the smallest possible container for weighing. Avoid containers made
of plastic, because they can become electrostatically charged. Use glass
vessels or weighing paper if possible. The weighing vessel and the
sample to be weighed should be at the ambient temperature. Never put
your hand into the weighing chamber, as this would cause the chamber to
warm up.
4. Use clean weighing containers. Determine the mass of the weighing
vessel prior to filling.
5. Place the sample in the weighing container, in the middle of the weighing
pan to avoid corner-load error.
6. Keep the working-place, weighing chamber, and weighing-pan clean.
Any spillage must be cleaned up immediately. Otherwise, chemicals may
quickly corrode the balance, and biological materials may be a source of
infection. Balance pans can be disinfected with 70% ethanol (700ml/l).
Read the manufacturer’s instructions carefully:
1. With new or returned equipment, remove the packaging and any transit
fixings that may have been fitted. Keep these safe for future use.
2. All balances should be sited on a solid, vibration-free surface that is free
from draughts, away from sunlight, and at an even temperature. The
atmosphere should be dust-free and chemical-free. Never store reagent
bottles in a room with balances.
3. The instrument must be placed in a precisely horizontal position (level).
This is checked using the spirit level. The air bubble must be in the center
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of the level; if it is not centered, then a correction is made by turning the
levelling-foot of the balance.
If good working practices are observed, there should be no special hazards. When
simple balances have been used for weighing or balancing biological samples, it is
prudent to disinfect the balance pans by swabbing with 70% ethanol (700 ml/l).
Special tools/spares/requirements
Leather cloth
Mini vacuum cleaner
Dust brush
Anti-magnetic tweezers
Watchmaker’s screwdrivers
Dental mirror
Magnifying glass
Set of spanners to the manufacturer’s specifications
Set of calibrating weights
Spare bulbs for optical balances.
Maintenance protocols
For all beam balances:
1. Check balance is leveled.
2. Check that all control knobs are properly fitted.
3. Check zeroing device.
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4. Check positioning of beam.
5. Check with both high and low mass calibrating weights.
For optical balances:
As above for beam balances. Then:
 Make sure all counterbalancing weights are in place.
 Check with an appropriate milligram weight that the optical scale agrees with
counterbalance weight.
For all balances:
1. Tighten nuts and screws where applicable.
2. Check and clean knife-edges.
3. Check light path, clean, align, and focus.
Read the manufacturer’s service manual carefully, if it is available.
With optical balances, remove covers and inspect beam and pivots for freedom of
movement; gentle finger pressure on the pan may reveal a “sticky” linkage or
bumper stop.
Check calibration
After any maintenance, service, repair, or re-siting.
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Water-baths are used for investigations at 25° C, 30° C, 37° C, 42° C, or 56° C. It
is important that the water-bath maintains a constant temperature within a narrow
range (± 0.1°C) during the investigation. Incorrect adjustment of the temperature
and insufficient temperature stability will seriously affect the results of
1. The level of water in the water-bath must be above the level of the solution to
be incubated.
2. Open containers, vials, or tubes must be incubated in a water-bath with the lid
of the water-bath open to avoid contamination and dilution of the incubated
material by condensed water.
3. The water in water-baths must be changed regularly to avoid the growth of
algae and bacteria.
4. Temperature should be checked and recorded daily
5. Maintenance includes cleaning, checking the heating element, and verifying the
temperature is accurate.
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Updated Dec 2007
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Laboratory Safety Manual
Table of Contents:
WHO Material Reference
Student Guide to Infection Prevention and Control
Universal Blood and Bodily Fluid Precautions
Chemical Exposure Introduction
Basic Toxicology
Routes of Entry
Toxic Effects Of Chemical Exposure
Chemical Exposure Determination
Section 6: Controlling Chemical Exposures
6.1 General Principles Personal Protection
Personal Protective Equipment or Devices ( PPEs – PPDs )
Section 7: Chemical Spills
Chemical Spill Procedures
Section 8: Safe Work Practices and Procedures
Flammable Materials
Corrosive Materials
Compressed Gases
Section 9: Safe Work Practices and Procedures
Electrical Safety in the Lab.
Laboratory Equipment Safety
Section 11: Anecdotes
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Biosafety: See WHO Web page for PDF download.
Universal Precautions:
PART I Guidelines
Risk assessment
Basic laboratories – Biosafety Levels 1 and 2
The containment laboratory – Biosafety Level 3
The maximum containment laboratory – Biosafety Level 4
PART II Laboratory equipment
Biological safety cabinets
Equipment-related hazards
Equipment designed to reduce biological hazards
PART III Good microbiological technique
Safe laboratory techniques Biosafety and recombinant DNA technology
Transport of infectious substances
Contingency plans and emergency procedures
Disinfection and sterilization
PART IV Chemical, fire and electrical safety
Hazardous chemicals
Fire in the laboratory
Electrical hazards
PART V Safety organization and training
The biosafety officer and safety committee
Safety rules for support staff
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Student Guide
2002 - 2003
Guide for Infection Prevention and Control
1. Strategies for Infection Control
PUH Medical students are expected to utilize universal precautions in the
clinical setting and for all patients. Plan and use precautions based on the
procedure you will be doing, not the patient’s diagnosis. Study after study
shows that providers cannot identify many of the highest risk patients.
a. Hand washing is performed frequently to protect both patients and
healthcare providers. Hands are washed before touching patients,
performing invasive procedures, and eating and after glove use,
accidental soiling with body substances, and using the bathroom. Skin is
a natural barrier to infectious agents; products that protect and promote
skin integrity should be used.
b. Gloves are worn for anticipated contact with all body substances. Clean
gloves are put on immediately before contact with the patient’s mucous
membranes or open skin. Gloves are changed between patients and
between certain sites of the same patient
(e.g., rectal to oral contact), and contaminated gloves are removed and
discarded into an appropriate waste container.
c. Gowns and/or plastic aprons are used to cover areas of the skin or
clothing that are likely to become soiled with body substances during
patient procedures or care. After use, they should be disposed of into an
appropriate waste container.
d. Facial barriers, including mask, glasses/goggles and face shields are
worn whenever splashing or splatter of body substances into the mouth,
nose, or eyes could occur. Masks are also used for certain airborne
diseases such as meningococcal meningitis, tuberculosis, and pertussis.
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e. Other barriers such as hair covers, shoe covers, and boots may be
used when extensive exposure to body fluids may occur (e.g.,
cystoscopy, vaginal delivery, multiple trauma).
f. Special rooms: For patients with active tuberculosis, a private room
with special ventilation is required. For primary varicella, measles,
mumps, and rubella, a private room with negative air pressure is
g. Sharps Management: Preventing injuries from contaminated sharps
reduces the risk of exposure to blood-borne diseases such as HIV,
Hepatitis B and C. Make every effort to prevent accidental injury to
yourself and co-workers.
Discard all used sharps into rigid impervious containers.
Do not place needles or sharps in wastebaskets or leave them at
patients’ bedsides or on a clinic workspace.
Do not routinely recap contaminated needles. If recapping is
unavoidable, acceptable methods of recapping include the use of
safety devices, which either shield the hand or hold the cap to
facilitate a one-handed technique.
Forceps may be used to carefully remove contaminated needles
or knife blades.
Additional "safe" sharps practices are applicable in the operating
room or during special procedures.
Stopcocks may replace injection ports for intravenous infusion or
Needles should be removed from suture material before tying
Sharp instruments should not be passed from hand to hand.
Place the sharp on an appropriate sterile or clean surface and
announce that it is time for the second person to pick it up.
Whenever possible, inform others as to the location of
contaminated sharps in order to prevent injury to co-workers.
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Universal Blood and Body Fluid Precautions
Because the potential for infectivity of any patient's blood and body fluids cannot be known,
blood and body fluid precautions recommended by the Centers for Disease Control (CDC)
should be adhered to for all specimens submitted to the laboratory and for all patients. These
precautions, called "universal precautions," should be followed regardless of any lack of
evidence of the patient's infection status.
1. Routinely use barrier protection to prevent skin and mucous membrane contamination
with blood or body fluids of all patients and specimens.
2. Wear gloves when engaged in the following:
 Touching blood and body fluids, including during routine laboratory work and
 Touching all laboratory specimens and tissues.
 Touching mucous membranes and nonintact skin of all patients.
 Handling items contaminated with blood or body fluids, including specimen
containers, laboratory instruments, counter tops, etc.
 Performing venipuncture, arterial puncture, skin puncture, and other vascular
access procedures.
Note: All skin defects (cuts, abrasions, ulcers, areas of dermatitis, etc,) should be covered
with an occlusive bandage.
3. Change gloves between each patient.
4. Wear a mask and eye covering, or preferably a face shield, during procedures that are
likely to generate droplets of blood or body fluids to prevent exposure to the mucous
membranes of the mouth, nose, and eyes.
5. Wear a gown, apron, or other covering when there is a potential for splashing or spraying
blood or body fluids.
6. Wash hands or other skin surfaces thoroughly and immediately if contaminated with
blood or body fluids.
7. Wash hands immediately after gloves are removed.
8. Take extraordinary care to avoid accidental injuries caused by needles, scalpel blades,
laboratory instruments, etc, when performing procedures, cleaning instruments, handling
sharp instruments, and disposing of used needles.
9. Place used needles, skin lances, scalpel blades, and other sharp items into a punctureresistant biohazard container for disposal. The container should be located as close as
possible to the work area. Phlebotomists should carry puncture-resistant containers with
10. To prevent needlestick injuries, needles should not be recapped, purposely bent, cut,
broken, removed from disposable syringes, or otherwise manipulated by hand.
11. Place large-bore reusable needles (e.g., bone-marrow needles and biopsy needles) and
other reusable sharps into a puncture-resistant container for transport to the reprocessing
12. Minimize the need for mouth-to-mouth emergency resuscitation procedures. Mouth
pieces, resuscitation bags, or other ventilation devices should be used routinely.
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13. Take care to minimize the formation of droplets, spatters, splashes, and spills of blood or
body fluids.
14. Clean all surfaces exposed to blood and body fluids with a detergent solution followed by
decontamination with an appropriate EPA-approved chemical germicide.
Laboratory workers with exudative lesions or weeping dermatitis should refrain from all patient
contact and from handling patient-care equipment and patient specimens until the condition
NOTE: Alternatively, skin lesions should be covered with an occlusive bandage to prevent
Pregnant women are not known to be at greater risk of contracting blood-born infections than
other laboratory workers. However, if HIV infection develops during pregnancy, the infant is at
risk of infection by perinatal transmission. Therefore, pregnant laboratory workers should be
especially aware of universal precautions.
Special Precautions for Laboratories
1. Laboratory space should be allocated to minimize crowding, which may contribute to
laboratory accidents.
2. Laboratory surfaces, counters, and floors should be made of impervious materials to
facilitate disinfection.
3. Good laboratory practices should be followed, and eating, drinking, and smoking should
not be permitted in the laboratory.
4. Adequate and conveniently located biohazard containers for disposal of contaminated
materials should be provided.
5. Adequate decontaminating containers for reusable supplies should be provided.
6. Written decontamination, disinfection, and sterilization protocols should be developed for
processing reusable supplies, laboratory equipment, laboratory waste, machine efluent,
and environmental surfaces.
7. Facilities for handwashing should be provided in each laboratory area. These should be
separate from those used for washing equipment or for waste disposal.
8. Only authorized personnel should be allowed in the laboratory: casual visitors should not
be admitted. Non-laboratory personnel should be closely supervised and should use
appropriate protective measures to ensure that they do not cause a hazard to themselves
or to the laboratory staff.
9. Monitoring compliance is a major responsibility of the management of the laboratory.
The necessary educational, monitoring, and remedial programs should be defined,
documented in writing, and rigorously enforced. The cooperation of the Institutional
Quality Assurance program should be enlisted.
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Chemical Exposure Introduction
The decisions you make concerning the use of chemicals in the laboratory should be based on an
objective analysis of the hazards, rather than merely the perception of the risks involved. Once
this has been accomplished, a reasonable means of controlling the hazards through experimental
protocol, work practices, ventilation, use of protective clothing, etc., can be determined.
In order to assess the hazards of a particular chemical, both the physical and health hazards of the
chemical must be considered.
Before using any chemical, the material safety data sheet (MSDS) or other appropriate resource
should be reviewed to determine what conditions of use might pose a hazard. Accidents with
hazardous chemicals can happen quickly and may be quite severe. The key to prevention of these
accidents is awareness. Once the hazards are known, the risk of an accident may be reduced
significantly by using safe work practices.
5.1 Basic Toxicology
The health effects of hazardous chemicals are often less clear than the physical hazards. Data on
the health effects of chemical exposure, especially from chronic exposure, are often incomplete.
When discussing the health effects of chemicals, two terms are often used interchangeably toxicity and hazard. However, the actual meanings of these words are quite different. Toxicity is
an inherent property of a material, similar to its physical constants. It is the ability of a chemical
substance to cause an undesirable effect in a biological system. Hazard is the likelihood that a
material will exert its toxic effects under the conditions of use. Thus, with proper handling,
highly toxic chemicals can be used safely. Conversely, less toxic chemicals can be extremely
hazardous if handled improperly.
The actual health risk of a chemical is a function of the toxicity and the actual exposure. No
matter how toxic the material may be, there is little risk involved unless it enters the body. An
assessment of the toxicity of the chemicals and the possible routes of entry will help determine
what protective measures should be taken.
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5.2 Routes of Entry
Skin and Eye Contact
The simplest way for chemicals to enter the body is through direct contact with the skin or eyes.
Skin contact with a chemical may result in a local reaction, such as a burn or rash, or absorption
into the bloodstream. Absorption into the bloodstream may then allow the chemical to cause
toxic effects on other parts of the body. The MSDS usually includes information regarding
whether or not skin absorption is a significant route of exposure.
The absorption of a chemical through intact skin is influenced by the health of the skin and the
properties of the chemical. Skin that is dry or cracked or has lacerations offers less resistance.
Fat-soluble substances, such as many organic solvents, can easily penetrate skin and, in some
instances, can alter the skin’s ability to resist absorption of other substances.
Wear gloves and other protective clothing to minimize skin exposure. See Personal Protective
Equipment for more information. Symptoms of skin exposure include dry, whitened skin,
redness and swelling, rashes or blisters, and itching. In the event of chemical contact on skin,
rinse the affected area with water for at least 15 minutes, removing clothing while rinsing, if
necessary. Seek medical attention if symptoms persist.
Avoid use of solvents for washing skin. They remove the natural protective oils from the skin
and can cause irritation and inflammation. In some cases, washing with a solvent may facilitate
absorption of a toxic chemical.
Chemical contact with eyes can be particularly dangerous, resulting in painful injury or loss of
sight. Wearing safety goggles or a face shield can reduce the risk of eye contact. Eyes that have
been in contact with chemicals should be rinsed immediately with water continuously for at least
15 minutes. Contact lenses should be removed while rinsing—do not delay rinsing to remove the
lenses. Medical attention is necessary if symptoms persist.
The respiratory tract is the most common route of entry for gases, vapors, particles, and aerosols
(smoke, mists and and fumes). These materials may be transported into the lungs and exert
localized effects, or be absorbed into the bloodstream. Factors that influence the absorption of
these materials may include the vapor pressure of the material, solubility, particle size, its
concentration in the inhaled air, and the chemical properties of the material. The vapor pressure
is an indicator of how quickly a substance evaporates into the air and how high the concentration
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in air can become – higher concentrations in air cause greater exposure in the lungs and greater
absorption in the bloodstream.
Most chemicals have an odor that is perceptible at a certain concentration, referred to as the odor
threshold; however, there is no relationship between odor and toxicity. There is considerable
individual variability in the perception of odor. Olfactory fatigue may occur when exposed to
high concentrations or after prolonged exposure to some substances. This may cause the odor to
seem to diminish or disappear, while the danger of overexposure remains.
Symptoms of over-exposure may include headaches, increased mucus production, and eye, nose
and throat irritation. Narcotic effects, including confusion, dizziness, drowsiness, or collapse,
may result from exposure to some substances, particularly many solvents. In the event of
exposure, close containers or otherwise increase ventilation, and move to fresh air. If symptoms
persist, seek medical attention.
Volatile hazardous materials should be used in a well-ventilated area, preferably a fume hood, to
reduce the potential of exposure. Occasionally, ventilation may not be adequate and a fume hood
may not be practical, necessitating the use of a respirator. The Occupational Safety and Health
Administration Respiratory Protection Standard regulates the use of respirators; thus, use of a
respirator is subject to prior review by EHS according to University policy. See Personal
Protective Equipment for more information.
The gastrointestinal tract is another possible route of entry for toxic substances. Although direct
ingestion of a laboratory chemical is unlikely, exposure may occur as a result of ingesting
contaminated food or beverages, touching the mouth with contaminated fingers, or swallowing
inhaled particles which have been cleared from the respiratory system. The possibility of
exposure by this route may be reduced by not eating, drinking, smoking, or storing food in the
laboratory, and by washing hands thoroughly after working with chemicals, even when gloves
were worn.
Direct ingestion may occur as a result of the outdated and dangerous practice of mouth pipetting.
In the event of accidental ingestion, immediately go to McCosh Health Center or contact the
Poison Control Center, at 800-962-1253 for instructions. Do not induce vomiting unless directed
to do so by a health care provider.
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The final possible route of exposure to chemicals is by accidental injection. Injection effectively
bypasses the protection provided by intact skin and provides direct access to the bloodstream,
thus, to internal organ systems. Injection may occur through mishaps with syringe needles, when
handling animals, or through accidents with pipettes, broken glassware or other sharp objects
that have been contaminated with toxic substances.
If accidental injection has occurred, wash the area with soap and water and seek medical
attention, if necessary. Cautious use of any sharp object is always important. Substituting
cannulas for syringes and wearing gloves may also reduce the possibility of injection.
5.3 Toxic Effects Of Chemical Exposure
How a chemical exposure affects a person depends on many factors. The dose is the amount of a
chemical that actually enters the body. The actual dose that a person receives depends on the
concentration of the chemical and the frequency and duration of the exposure. The sum of all
routes of exposure must be considered when determining the dose.
In addition to the dose, the outcome of exposure is determined by (1) the way the chemical enters
the body, (2) the physical properties of the chemical, and (3) the susceptibility of the individual
receiving the dose.
Toxic Effects of Chemicals
The toxic effects of a chemical may be local or systemic. Local injuries involve the area of the
body in contact with the chemical and are typically caused by reactive or corrosive chemicals,
such as strong acids, alkalis or oxidizing agents. Systemic injuries involve tissues or organs
unrelated to or removed from the contact site when toxins have been transported through the
bloodstream. For example, methanol that has been ingested may cause blindness, while a
significant skin exposure to nitrobenzene may effect the central nervous system.
Certain chemicals may affect a target organ. For example, lead primarily affects the central
nervous system, kidney and red blood cells; isocyanates may induce an allergic reaction
(immune system); and chloroform may cause tumors in the liver and kidneys.
It is important to distinguish between acute and chronic exposure and toxicity. Acute toxicity
results from a single, short exposure. Effects usually appear quickly and are often reversible.
Chronic toxicity results from repeated exposure over a long period of time. Effects are usually
delayed and gradual, and may be irreversible. For example, the acute effect of alcohol exposure
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(ingestion) is intoxication, while the chronic effect is cirrhosis of the liver. Acute and chronic
effects are distinguished in the MSDS, usually with more information about acute exposures than
Relatively few chemicals have been evaluated for chronic effects, given the complexity of that
type of study. Chronic exposure may have very different effects than acute exposure. Usually,
studies of chronic exposure evaluate its cancer causing potential or other long-term health
Evaluating Toxicity Data
Most estimates of human toxicity are based on animal studies, which may or may not relate to
human toxicity. In most animal studies, the effect measured is usually death. This measure of
toxicity is often expressed as an LD50 (lethal dose 50) – the dose required to kill 50% of the test
population. The LD50 is usually measured in milligrams of the material per kilogram of body
weight of the test animal. The concentration in air that kills half of the population is the LC50.
To estimate a lethal dose for a human based on animal tests, the LD50 must be multiplied by the
weight of an average person. Using this method, it is evident that just a few drops of a highly
toxic substance, such as dioxin, may be lethal, while much larger quantities of a slightly toxic
substance, such as acetone, would be necessary for the same effect.
Susceptibility of Individuals
Factors that influence the susceptibility of an individual to the effects of toxic substances include
nutritional habits, physical condition, obesity, medical conditions, drinking and smoking, and
pregnancy. Due to individual variation and uncertainties in estimating human health hazards, it is
difficult to determine a dose of a chemical that is totally risk-free.
Regular exposure to some substances can lead to the development of an allergic rash, breathing
difficulty, or other reactions. This phenomenon is referred to as sensitization. Over time, these
effects may occur with exposure to smaller and smaller amounts of the chemical, but will
disappear soon after the exposure stops. For reasons not fully understood, not everyone exposed
to a sensitizer will experience this reaction. Examples of sensitizers include epoxy resins, nickel
salts, isocyanates and formaldehyde.
Particularly Hazardous Substances
The OSHA Laboratory Standard defines a particularly hazardous substance as "select
carcinogens", reproductive toxins, and substances that have a high degree of acute toxicity.
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Further information about working with Particularly Hazardous Substances is outlined in
Particularly Hazardous Substances.
Where To Find Toxicity Information
Toxicity information may be found in Material Safety Data Sheets, under the "Health Hazard
Data" section, on product labels, in the Registry of Toxic Effects of Chemical Substances
(RTECS), or in many other sources listed in the MSDS page.
5.4 Chemical Exposure Determination
OSHA establishes exposure limits for several hundred substances. Laboratory workers must not
be exposed to substances in excess of the permissible exposure limits (PEL) specified in OSHA
Subpart Z, Toxic and Hazardous Substances. PELs refer to airborne concentrations of substances
averaged over an eight-hour day. Some substances also have "action levels" below the PEL
requiring certain actions such as medical surveillance or routine air sampling.
The MSDS for a particular substance indicates whether any of the chemicals are regulated
through OSHA and, if so, the permissible exposure limit(s) for the regulated chemical(s). This
information is also available in the OSHA Table Z list of regulated chemicals.
Exposure Monitoring
Exposure monitoring must be conducted if there is reason to believe that exposure levels for a
particular substance may routinely exceed either the action level or the PEL. EHS and the
principal investigator or supervisor may use professional judgement, based on the information
available about the hazards of the substance and the available control measures, to determine
whether exposure monitoring must be conducted.
When necessary, exposure monitoring is conducted by EHS according to established industrial
hygiene practices. Results of the monitoring are made available to the individual monitored, his
or her supervisor, and the departmental Chemical Hygiene Officer within 15 working days of the
receipt of analytical results.
Section 6: Controlling Chemical Exposures
General Principles
 Engineering Controls
 Work Practice and Administrative Controls
 Protective Equipment
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Fume Hoods and Laboratory Ventilation - separate page
Personal Protective Equipment - separate page
6.1 General Principles
There are three general methods for controlling one's exposure to hazardous substances:
 Engineering Controls
 Work Practices and Administrative Controls
 Personal Protective Equipment
In the laboratory, these methods or a combination of them can be used to keep exposure below
permissible exposure limits.
Engineering Controls
Engineering controls include the following:
 Substitution of a less toxic material
 Change in process to minimize contact with hazardous chemicals
 Isolation or enclosure of a process or operation
 Use of wet methods to reduce generation of dusts or other particulates
 General dilution ventilation
 Local exhaust, including the use of fume hoods
The use of engineering controls is the preferred method for reducing worker exposure to
hazardous chemicals, but with the exception of chemical fume hoods, may not be feasible in the
Work Practice and Administrative Controls
Using good laboratory work practices, such as those outlined in this manual, help to reduce the
risk of exposure to chemicals.
Administrative controls involve rotating job assignments and adjusting work schedules so that
workers are not overexposed to a chemical. Given the nature of work in a research laboratory,
administrative controls are not usually a realistic approach to controlling exposure.
Personal Protective Equipment
When engineering controls are not sufficient to minimize exposure, personal protective
equipment, including gloves, eye protection, respirators and other protective clothing should be
used. See Personal Protective Equipment for more information
Personal protective equipment (PPE) is special gear used to protect the wearer from specific
hazards of a hazardous substance. It is a last resort protection system, to be used when
substitution or engineering controls are not feasible. PPE does not reduce or eliminate the hazard,
protects only the wearer, and does not protect anyone else.
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PPE includes gloves, respiratory protection, eye protection, and protective clothing. The need for
PPE is dependent upon the type of operations and the nature and quantity of the materials in use,
and must be assessed on a case by case basis. Workers who rely on PPE must understand the
functioning, proper use, and limitations of the PPE used.
A. Eye Protection
Safety Glasses
Safety glasses look very much like normal glasses buy have lenses that are impact resistant and
frames that are far stronger than standard streetwear glasses. Safety glasses must have side
shields and should be worn whenever there is the possibility of objects striking the eye, such as
particles, glass, or metal shards. Many potential eye injuries have been avoided by wearing
safety glasses. See Anecdotes for accounts of a few of these incidents.
Standard streetwear eyeglasses fitted with side shields are not sufficient. Workers who are
interested in obtaining prescription safety glasses should contact EHS at 8-5294. Safety glasses
come in a variety of styles to provide the best fit and comfort, including some designed to fit
over prescription glasses.
Safety glasses do not provide adequate protection from significant chemical splashes. They do
not seal to the face, resulting in gaps at the top, bottom and sides, where chemicals may seep
through (see Anecdotes for an actual incident where this occurred). Safety glasses may be
adequate when the potential splash is minimal, such as when opening eppendorf tubes.
Safety glasses are also not appropriate for dusts and powders, which can get by the glasses in
ways similar to those described above. Safety goggles are best used for this type of potential
Chemical Splash Goggles
Chemical Splash Goggles should be worn when there is potential for splash from a hazardous
material. Like safety glasses, goggles are impact resistant. Chemical splash goggles should have
indirect ventilation so hazardous substances cannot drain into the eye area. Some may be worn
over prescription glasses.
Goggles come in a variety of styles for maximum comfort and splash protection. Visorgogs are a
hybrid of a goggle and safety glasses. They offer more splash protection than safety glasses, but
not as much as goggles. They fit close to the face, but do not seal at the bottom as goggles do.
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Face Shields
Face shields are in order when working with large volumes of hazardous materials, either for
protection from splash to the face or flying particles. Face shields must be used in conjunction
with safety glasses or goggles. A few incidents where a face shield would have prevented injury
are described in Anecdotes.
Contact Lenses
Contact lenses may be worn in the laboratory, but do not offer any protection from chemical
contact. If a contact lens becomes contaminated with a hazardous chemical, rinse the eye(s)
using an eyewash and remove the lens immediately. Contact lenses that have been contaminated
with a chemical must be discarded.
This particularly recommendation runs counter to what most of us were taught previously.
However, studies have shown that contact lenses are safe to wear in the laboratory environment.
For more information, see the American Optometric Association guidelines.
B. Protective Clothing & Footwear
Protective Clothing
When the possibility of chemical contamination exists, protective clothing that resists physical
and chemical hazards should be worn over street clothes. Lab coats are appropriate for minor
chemical splashes and spills, while plastic or rubber aprons are best for protection from corrosive
or irritating liquids. Disposable outer garments (i.e., Tyvek suits) may be useful when cleaning
and decontamination of reusable clothing is difficult.
Loose clothing (such as overlarge lab coats or ties), skimpy clothing (such as shorts), torn
clothing and unrestrained hair may pose a hazard in the laboratory.
Shoes should be worn at all times in buildings where chemicals are stored or used. Perforated
shoes, sandals or cloth sneakers should not be worn in laboratories or where mechanical work is
conducted. Such shoes offer no barrier between the laboratory worker and chemicals or broken
Chemical resistant overshoes or boots may be used to avoid possible exposure to corrosive
chemical or large quantities of solvents or water that might penetrate normal footwear (e.g.,
during spill cleanup). Leather shoes tend to absorb chemicals and may have to be discarded if
contaminated with a hazardous material.
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Although generally not required in most laboratories, steel-toed safety shoes may be necessary
when there is a risk of heavy objects falling or rolling onto the feet, such as in bottle-washing
operations or animal care facilities.
C. Gloves
Choosing the appropriate hand protection can be a challenge in a laboratory setting. Considering
the fact that dermatitis or inflammation of the skin accounts for 40-45% of all work-related
diseases, selecting the right glove for the job is important.
Not only can many chemicals cause skin irritation or burns, but also absorption through the skin
can be a significant route of exposure to certain chemicals. Dimethyl sulfoxide (DMSO),
nitrobenzene, and many solvents are examples of chemicals that can be readily absorbed through
the skin into the bloodstream, where the chemical may cause harmful effects.
When Should Gloves Be Worn
Protective gloves should be worn when handling hazardous materials, chemicals of unknown
toxicity, corrosive materials, rough or sharp-edged objects, and very hot or very cold materials.
When handling chemicals in a laboratory, disposable latex, vinyl or nitrile examination gloves
are usually appropriate for most circumstances. These gloves will offer protection from
incidental splashes or contact.
When working with chemicals with high acute toxicity, working with corrosives in high
concentrations, handling chemicals for extended periods of time or immersing all or part of a
hand into a chemical, the appropriate glove material should be selected, based on chemical
Selecting the Appropriate Glove Material
When selecting the appropriate glove, the following characteristics should be considered:
 degradation rating
 breakthrough time
 permeation rate
Degradation is the change in one or more of the physical properties of a glove caused by contact
with a chemical. Degradation typically appears as hardening, stiffening, swelling, shrinking or
cracking of the glove. Degradation ratings indicate how well a glove will hold up when exposed
to a chemical. When looking at a chemical compatibility chart, degradation is usually reported as
E (excellent), G (good), F (fair), P (poor), NR (not recommended) or NT (not tested).
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Breakthrough time is the elapsed time between the initial contact of the test chemical on the
surface of the glove and the analytical detection of the chemical on the inside of the glove.
Permeation rate is the rate at which the test chemical passes through the glove material once
breakthrough has occurred and equilibrium is reached. Permeation involves absorption of the
chemical on the surface of the glove, diffusion through the glove, and desorption of the chemical
on the inside of the glove. Resistance to permeation rate is usually reported as E (excellent), G
(good), F (fair), P (poor) or NR (not recommended). If chemical breakthrough does not occur,
then permeation rate is not measured and is reported ND (none detected).
Manufacturers stress that permeation and degradation tests are done under laboratory test
conditions, which can vary significantly from actual conditions in the work environment. Users
may opt to conduct their own tests, particularly when working with highly toxic materials.
For mixtures, it is recommended that the glove material be selected based on the shortest
breakthrough time.
The following table includes major glove types and their general uses. This list is not exhaustive.
Glove Material
General Uses
Offers the highest resistance to permeation by most gases and
water vapor. Especially suitable for use with esters and
Provides moderate abrasion resistance but good tensile strength
and heat resistance. Compatible with many acids, caustics and
Excellent general duty glove. Provides protection from a wide
variety of solvents, oils, petroleum products and some
corrosives. Excellent resistance to cuts, snags, punctures and
Provides excellent abrasion resistance and protection from
most fats, acids, and petroleum hydrocarbons.
Highly impermeable to gases. Excellent protection from
aromatic and chlorinated solvents. Cannot be used in water or
water-based solutions.
Exceptional resistance to chlorinated and aromatic solvents.
Good resistance to cuts and abrasions.
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Silver Shield
Resists a wide variety of toxic and hazardous chemicals.
Provides the highest level of overall chemical resistance.
Natural rubber
Provides flexibility and resistance to a wide variety of acids,
caustics, salts, detergents and alcohols.
Where to Find Compatibility Information
Most glove manufacturers have chemical compatibility charts available for their gloves. These
charts may be found in laboratory safety supply catalogs such as Fisher Scientific and Lab Safety
Supply. Best Gloves offers copies of their glove compatibility charts upon request. To obtain a
copy, call them directly at 800-241-0323. Best Gloves also has a great deal of information
available on their web site, including a downloadable glove selection program.
Most material safety data sheets (MSDS) recommend the most protective glove material in their
Protective Equipment section. There are MSDSs for many laboratory chemicals available on the
web through the EHS home page.
EHS also has a computer program with glove compatibility information for hundreds of
chemicals. Contact EHS at 258-5294 for more information.
Other Considerations
There are several factors besides glove material to consider when selecting the appropriate glove.
The amount of dexterity needed to perform a particular manipulation must be weighed against
the glove material recommended for maximum chemical resistance. In some cases, particularly
when working with delicate objects where fine dexterity is crucial, a bulky glove may actually be
more of a hazard.
Where fine dexterity is needed, consider double gloving with a less compatible material,
immediately removing and replacing the outer glove if there are any signs of contamination. In
some cases, such as when wearing Silver Shield gloves, it may be possible to wear a tight-fitting
glove over the loose glove to increase dexterity.
Glove thickness, usually measured in mils or gauge, is another consideration. A 10-gauge glove
is equivalent to 10 mils or 0.01 inches. Thinner, lighter gloves offer better touch sensitivity and
flexibility, but may provide shorter breakthrough times. Generally, doubling the thickness of the
glove quadruples the breakthrough time.
Glove length should be chosen based on the depth to which the arm will be immersed or where
chemical splash is likely. Gloves longer than 14 inches provide extra protection against splash or
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Glove size may also be important. One size does not fit all. Gloves which are too tight tend to
cause fatigue, while gloves which are too loose will have loose finger ends which make work
more difficult. The circumference of the hand, measured in inches, is roughly equivalent to the
reported glove size. Glove color, cuff design, and lining should also be considered for some
Glove Inspection, Use and Care
All gloves should be inspected for signs of degradation or puncture before use. Test for pinholes
by blowing or trapping air inside and rolling them out. Do not fill them with water, as this makes
the gloves uncomfortable and may make it more difficult to detect a leak when wearing the
Disposable gloves should be changed when there is any sign of contamination. Reusable gloves
should be washed frequently if used for an extended period of time.
While wearing gloves, be careful not to handle anything but the materials involved in the
procedure. Touching equipment, phones, wastebaskets or other surfaces may cause
contamination. Be aware of touching the face, hair, and clothing as well.
Before removing them, wash the outside of the glove. To avoid accidental skin exposure, remove
the first glove by grasping the cuff and peeling the glove off the hand so that the glove is inside
out. Repeat this process with the second hand, touching the inside of the glove cuff, rather than
the outside. Wash hands immediately with soap and water.
Follow the manufacturer’s instructions for washing and caring for reusable gloves.
Latex Gloves and Related Allergies
Allergic reactions to natural rubber latex have been increasing since 1987, when the Centers for
Disease Control recommended the use of universal precautions to protect against potentially
infectious materials, bloodborne pathogens and HIV. Increased glove demand also resulted in
higher levels of allergens due to changes in the manufacturing process. In addition to skin
contact with the latex allergens, inhalation is another potential route of exposure. Latex proteins
may be released into the air along with the powders used to lubricate the interior of the glove.
In June, 1997, the National Institute of Occupational Safety and Health (NIOSH) issued an alert
Preventing Allergic Reactions to Latex in the Workplace (publication number DHHS (NIOSH)
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Latex exposure symptoms include skin rash and inflammation, respiratory irritation, asthma and
shock. The amount of exposure needed to sensitize an individual to natural rubber latex is not
known, but when exposures are reduced, sensitization decreases.
NIOSH recommends the following actions to reduce exposure to latex:
Whenever possible, substitute another glove material.
If latex gloves must be used, choose reduced-protein, powder-free latex gloves.
Wash hands with mild soap and water after removing latex gloves.
D. Hearing Protection
Most laboratory equipment and operations do not produce noise levels that require the use of
hearing protection, with the exception of some wind tunnels, as described below. Princeton
University has a Hearing Conservation Program in place for individuals who are exposed to
noise levels equal to or exceeding the OSHA action level of 85 decibels (dBA) averaged over
eight hours, per the OSHA Occupational Noise Standard. This program includes workplace
monitoring, personal exposure monitoring, annual audiometric testing, use of hearing protection
and annual training.
Laboratory workers who would like to use hearing protection for noise levels below the action
level may do so without enrollment in the Hearing Conservation Program. Using hearing
protection, such as earplugs, earmuffs or hearing bands, can improve communication or provide
comfort to the worker in a noisy environment.
The most common noisy equipment in the laboratories are ultrasonicators and wind tunnels. EHS
has measured noise levels of several ultrasonicators used in the laboratories and found that noise
levels were well below 85 dBA, averaged over eight hours. Some of the wind tunnels,
particularly the supersonic wind tunnels, are capable of very high noise levels. Users should
check with the principal investigator or EHS to determine whether they need to be enrolled in the
Hearing Conservation Program.
For more information about the Hearing Conservation Program, see Section 5, Noise and
Hearing Conservation, of the Princeton University Health and Safety Guide. Contact EHS at
258-5294 to request noise monitoring.
E. Respiratory Protection
A respirator may only be used when engineering controls, such as general ventilation or a fume
hood, are not feasible or do not reduce the exposure of a chemical to acceptable levels. Since
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the use of a respirator is regulated by the OSHA Respiratory Protection Standard, respirator use
at Princeton is subject to prior review by EHS, according to university policy.
Any worker who believes that respiratory protection is needed must notify EHS for evaluation of
the hazard and enrollment in the Respiratory Protection Program. This program involves
procedures for respirator selection, medical assessment of employee health, employee training,
proper fitting, respirator inspection and maintenance, and recordkeeping.
Self-contained breathing apparatus is available in E-Quad for use by trained individuals for
changing out cylinders of highly toxic gases and cleaning up chemical spills. They are not to be
used for fire safety. Training is offered every April and October in E-Quad. Contact Kelly States
if you are interested and are not on the mailing list for this training.
Section 7: Chemical Spills
Information about cleaning up chemical spills is available in the Emergency Procedures section
of the EHS web page. This section contains information regarding:
 Developing a Spill Response Plan
 Spill Response and Cleanup Procedures
 Recommended Spill Control Material Inventory
Pre-planning is essential. Before working with a chemical, the laboratory worker should know
how to proceed with spill cleanup and should ensure that there are adequate spill control
materials available.
Preventing Spills
Most spills are preventable. The following are some tips that could help to prevent or minimize
the magnitude of a spill:
Place chemical containers in a hood or lab bench in a manner that reduces the possibility
of accidentally knocking over a container.
Plan your movements. Look where you are reaching to ensure you will not cause a spill.
Follow the procedures outlined for transporting chemicals safely.
Place absorbent plastic backed liners on benchtops or in fume hoods where spills can be
anticipated. For volumes of liquid larger than what can be absorbed by liners, use trays.
Followed the guidelines outlined for safe storage of chemicals.
 Spill Response and Clean-up Procedures
Developing a Spill Response Plan
Recommended Spill Control Material Inventory
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Spill Response and Clean-up Procedures
In the event of a chemical spill, the individual(s) who caused the spill is responsible for prompt
and proper clean-up. It is also their responsibility to have spill control and personal protective
equipment appropriate for the chemicals being handled readily available. See Developing a Spill
Response Plan for more information.
The following are general guidelines to be followed for a chemical spill. More detailed
procedures may be available in your Departmental Chemical Hygiene Plan or Spill Response
1. Immediately alert area occupants and supervisor, and evacuate the area, if necessary.
2. If there is a fire or medical attention is needed, contact Public Safety at 911.
3. Attend to any people who may be contaminated. Contaminated clothing must be removed
immediately and the skin flushed with water for no less than fifteen minutes. Clothing
must be laundered before reuse. See First Aid for Chemical Exposures for more
4. If a volatile, flammable material is spilled, immediately warn everyone, control sources
of ignition and ventilate the area.
5. Don personal protective equipment, as appropriate to the hazards. Refer to the Material
Safety Data Sheet or other references for information.
6. Consider the need for respiratory protection. The use of a respirator or self-contained
breathing apparatus requires specialized training and medical surveillance. Never enter a
contaminated atmosphere without protection or use a respirator without training. If
respiratory protection is needed and no trained personnel are available, call EHS at x85294 or Public Safety at 911. If respiratory protection is used, be sure there is another
person outside the spill area in communication, in case of an emergency. If no one is
available, contact Public Safety.
7. Using the chart below, determine the extent and type of spill. If the spill is large, if there
has been a release to the environment or if there is no one knowledgeable about spill
clean-up available, contact EHS at x8-5294 or Public Safety at 911.
Treatment Materials
up to 300 cc
300 cc - 5 liters
chemical treatment or
neutralization or absorption spill kit
absorption spill kit
Large more than 5 liters call Public Safety
outside help
8. Protect floor drains or other means for environmental release. Spill socks and absorbents
may be placed around drains, as needed.
9. Contain and clean-up the spill according to the table above.
Updated Dec 2007
MET Module V
Loose spill control materials should be distributed over the entire spill area,
working from the outside, circling to the inside. This reduces the chance of splash
or spread of the spilled chemical.
 Bulk absorbents and many spill pillows do not work with hydrofluoric acid.
POWERSORB (by 3M) products and their equivalent will handle hydrofluoric
acid. Specialized hydrofluoric acid kits also are available.
 Many neutralizers for acids or bases have a color change indicator to show when
neutralization is complete.
10. When spilled materials have been absorbed, use brush and scoop to place materials in an
appropriate container. Polyethylene bags may be used for small spills. Five gallon pails
or 20 gallon drums with polyethylene liners may be appropriate for larger quantities.
11. Complete a hazardous waste sticker, identifying the material as Spill Debris involving
XYZ Chemical, and affix onto the container. Spill control materials will probably need to
be disposed of as hazardous waste. Contact EHS at 258-5294 for advice on storage and
packaging for disposal.
12. Decontaminate the surface where the spill occurred using a mild detergent and water,
when appropriate.
13. Report all spills to your supervisor or the Principal Investigator.
Back to Top
Developing a Spill Response Plan
An effective spill response procedure should consider all of the items listed below. The
complexity and detail of the plan will, of course depend upon the physical characteristics and
volume of materials being handled, their potential toxicity, and the potential for releases to the
1. Review Material Safety Data Sheets (MSDSs) or other references for recommended spill
cleanup methods and materials, and the need for personal protective equipment (e.g.,
respirator, gloves, protective clothing, etc.)
2. Acquire sufficient quantities and types of appropriate spill control materials to contain
any spills that can be reasonably anticipated. The need for equipment to disperse, collect
and contain spill control materials (e.g., brushes, scoops, sealable containers, etc.) should
also be reviewed. See Recommended Spill Control Materials Inventory for more details.
3. Acquire recommended personal protective equipment and training in its proper use. For
example, if an air purifying respirator or self-contained breathing apparatus are needed,
personnel must be enrolled in the Respiratory Protection Program and attend annual
training and fit-testing.
4. Place spill control materials and protective equipment in a readily accessible location
within or immediately adjacent to the laboratory.
5. Develop a spill response plan that includes:
 Names and telephone numbers of individuals to be contacted in the event of a
 Evacuation plans for the room or building, as appropriate.
 Instructions for containing the spilled material, including potential releases to the
environment (e.g., protect floor drains).
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MET Module V
Inventory of spill control materials and personal protective equipment.
Means for proper disposal of cleanup materials (in most cases, as hazardous
waste) including contaminated tools and clothing.
 Decontamination of the area following the cleanup.
6. Discuss the spill response plans with all employees in the area. EHS offers training for
employees who work directly with chemicals (see Chemical Spills and Waste
Procedures) and who are expected to respond outside their work area to assist with spill
cleanup (see Chemical Emergency Response (HAZWOPER) First Responder Operations Level Training). Contact Robin Izzo at 258-6259 to schedule a session for
your area.
Back to top.
Recommended Spill Control Materials Inventory
Your laboratory or work area should have access to sufficient quantity of absorbents or other
types of materials to control any spill that can be reasonably anticipated. Vermiculite, lined 5gallon pails and limited spill control materials are available at the loading docks of Lewis
Thomas Lab, Frick, and E-Quad. Additional materials may be found in certain laboratories and
the chemical stockrooms.
Personal Protective Equipment
2 pairs chemical splash goggles
2 pairs of gloves (recommend Silver Shield or 4H)
2 pairs of shoe covers
2 plastic or Tyvek aprons and/or Tyvek suits
Absorption Materials
4 3M POWERSORB spill pillows (or equivalent)
1 3M POWERSORB spill sock
2 DOT pails (5 gallon) with polyethylene liners
 1 filled with loose absorbent, such as vermiculite or clay
 1 with minimum amount of loose absorbent in the bottom
Neutralizing Materials
Acid Neutralizer
Caustic Neutralizer
 commercial neutralizers, such as Neutrasorb (for acids) and Neutracit-2 (for
bases) have built in color change to indicate complete neutralization
Solvent Neutralizer
 commercial solvent neutralizers, such as Solusorb, act to reduce vapors and raise
the flashpoint of the mixture
Mercury Spills
Small mercury vacuum to pick up large drops (optional)
Updated Dec 2007
MET Module V
Hg Absorb Sponges - amalgamate mercury residue
Hg Absorb Powder - amalgamates mercury
Hg Vapor Absorbent - reduces concentration of vapor in hard to reach areas
Mercury Indicator - powder identifies presence of mercury
Clean-up Tools
Polypropylene scoop or dust pan
Broom or brush with polypropylene bristles
2 polypropylene bags
sealing tape
pH test papers
waste stickers
floor sign - DANGER Chemical Spill - Keep Away
Section 8: Safe Work Practices and Procedures
8.1 Flammable Materials
Flammable and Combustible Liquids
Storage of Flammable and Combustible Liquids
 Safety Cans and Closed Containers
 Flammable Liquid Storage Cabinets
 Refrigerators
 Storage Considerations
Handling Precautions
Flammable Aerosols
Flammable and Combustible Solids
Catalyst Ignition
A. Properties of Flammable and Combustible Liquids
Flammable and combustible liquids vaporize and form flammable mixtures with air when in
open containers, when leaks occur, or when heated. To control these potential hazards, several
properties of these materials, such as volatility, flashpoint, flammable range and autoignition
temperatures must be understood. An explanation of these terms and other properties of
flammable liquids is available the Laboratory Training Guide. Information on the properties of a
specific liquid can be found in that liquid’s material safety data sheet (MSDS), or other reference
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B. Storage of Flammable and Combustible Liquids
Flammable and combustible liquids should be stored only in approved containers. Approval for
containers is based on specifications developed by organizations such as the US Department of
Updated Dec 2007
MET Module V
Transportation (DOT), OSHA, the National Fire Protection Agency (NFPA) or American
National Standards Institute (ANSI). Containers used by the manufacturers of flammable and
combustible liquids generally meet these specifications.
Safety Cans and Closed Containers
Many types of containers are required depending on the quantities and classes of flammable or
combustible liquids in use. A safety can is an approved container of not more than 5 gallons
capacity that has a spring closing lid and spout cover. Safety cans are designed to safely relieve
internal pressure when exposed to fire conditions. A closed container is one sealed by a lid or
other device so that liquid and vapor cannot escape at ordinary temperatures.
Flammable Liquid Storage Cabinets
A flammable liquid storage cabinet is an approved cabinet that has been designed and
constructed to protect the contents from external fires. Storage cabinets are usually equipped
with vents, which are plugged by the cabinet manufacturer. Since venting is not required by any
code or the by local municipalities and since venting may actually prevent the cabinet from
protecting its contents, vents should remain plugged at all times. Storage cabinets must also be
conspicuously labeled "FLAMMABLE – KEEP FIRE AWAY".
Use only those refrigerators that have been designed and manufactured for flammable liquid
storage. Standard household refrigerators must not be used for flammable storage because
internal parts could spark and ignite. Refrigerators must be prominently labeled as to whether or
not they are suitable for flammable liquid storage.
Storage Considerations:
Quantities should be limited to the amount necessary for the work in progress.
No more than 10 gallons of flammable and combustible liquids, combined, should be
stored outside of a flammable storage cabinet unless safety cans are used. When safety
cans are used, up to 25 gallons may be stored without using a flammable storage cabinet.
Storage of flammable liquids must not obstruct any exit.
Flammable liquids should be stored separately from strong oxidizers, shielded from direct
sunlight, and away from heat sources. See Anecdotes for a description of an incident
involving a flammable material stored near a hot plate.
C. Handling Precautions
The main objective in working safely with flammable liquids is to avoid accumulation of vapors
and to control sources of ignition.
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Besides the more obvious ignition sources, such as open flames from Bunsen burners, matches
and cigarette smoking, less obvious sources, such as electrical equipment, static electricity and
gas-fired heating devices should be considered. Accounts of a few of the fires that have occurred
in our laboratories may be found in Anecdotes.
Some electrical equipment, including switches, stirrers, motors, and relays can produce sparks
that can ignite vapors. Although some newer equipment have spark-free induction motors, the
on-off switches and speed controls may be able to produce a spark when they are adjusted
because they have exposed contacts. One solution is to remove any switches located on the
device and insert a switch on the cord near the plug end.
Pouring flammable liquids can generate static electricity. The development of static electricity is
related to the humidity levels in the area. Cold, dry atmospheres are more likely to facilitate
static electricity. Bonding or using ground straps for metallic or non-metallic containers can
prevent static generation.
Control all ignition sources in areas where flammable liquids are used. Smoking, open
flames and spark producing equipment should not be used.
Whenever possible use plastic or metal containers or safety cans.
When working with open containers, use a laboratory fume hood to control the
accumulation of flammable vapor.
Use bottle carriers for transporting glass containers.
Use equipment with spark-free, intrinsically safe induction motors or air motors to avoid
producing sparks. These motors must meet National Electric Safety Code (US DOC,
1993) Class 1, Division 2, Group C-D explosion resistance specifications. Many stirrers,
Variacs, outlet strips, ovens, heat tape, hot plates and heat guns do not comform to these
code requirements.
Avoid using equipment with series-wound motors, since they are likely to produce
Do not heat flammable liquids with an open flame. Steam baths, salt and sand baths, oil
and wax baths, heating mantles and hot air or nitrogen baths are preferable.
Minimize the production of vapors and the associated risk of ignition by flashback.
Vapors from flammable liquids are denser than air and tend to sink to the floor level
where they can spread over a large area.
Electrically bond metal containers when transferring flammable liquids from one to
another. Bonding can be direct, as a wire attached to both containers, or indirect, as
through a common ground system.
When grounding non-metallic containers, contact must be made directly to the liquid,
rather than to the container.
In the rare circumstance that static cannot be avoided, proceed slowly to give the charge
time to disperse or conduct the procedure in an inert atmosphere.
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D. Flammable Aerosols
Flammable liquids in pressurized containers may rupture and aerosolize when exposed to heat,
creating a highly flammable vapor cloud. As with flammable liquids, these should be stored in a
flammable storage cabinet.
E. Flammable and Combustible Solids
Flammable solids often encountered in the laboratory include alkali metals, magnesium metal,
metallic hydrides, some organometallic compounds, and sulfur. Many flammable solids react
with water and cannot be extinguished with conventional dry chemical or carbon dioxide
extinguishers. See Anecdotes for descriptions of incidents involving such materials.
Ensure Class D extinguishers, e.g., Met-L-X, are available where flammable solids are
used or stored.
Sand can usually be used to smother a fire involving flammable solids. Keep a container
of sand near the work area.
If a flammable, water-reactive solid is spilled onto skin, brush off as much as possible,
then flush with copious amounts of water.
NEVER use a carbon dioxide fire extinguisher for fires involving lithium aluminum
hydride (LAH). LAH reacts explosively with carbon dioxide.
F. Catalyst Ignition
Some hydrogenated catalysts, such as palladium, platinum oxide, and Raney nickel, when
recovered from hydrogenation reactions, may become saturated with hydrogen and present a fire
or explosion hazard.
 Carefully filter the catalyst.
 Do not allow the filter cake to become dry.
 Place the funnel containing moist catalyst into a water bath immediately.
Purge gases, such as nitrogen or argon, may be used so that the catalyst can be filtered and
handled in an inert atmosphere
8.2 Corrosive Materials
 Corrosive Liquids
 Corrosive Gases and Vapors
 Corrosive Solids
Many chemicals commonly used in the laboratory are corrosive or irritating to body tissue. They
present a hazard to the eyes and skin by direct contact, to the respiratory tract by inhalation or to
Updated Dec 2007
MET Module V
the gastrointestinal system by ingestion. Anecdotes offers incidents involving chemical burns
from incorrectly handling corrosives.
A. Corrosive Liquids
Corrosive liquids (e.g. mineral acids, alkali solutions and some oxidizers) represent a very
significant hazard because skin or eye contact can readily occur from splashes and their effect on
human tissue generally takes place very rapidly. Bromine, sodium hydroxide, sulfuric acid and
hydrogen peroxide are examples of highly corrosive liquids. See Chemical Specific Issues for
specific corrosive liquids such as Hydrofluoric Acid and Phenol.
The following should be considered:
1. The eyes are particularly vulnerable. It is therefore essential that approved eye and face
protection be worn in all laboratories where corrosive chemicals are handled.
2. Gloves and other chemically resistant protective clothing should be worn to protect
against skin contact.
3. To avoid a flash steam explosion due to the large amount of heat evolved, always add
acids or bases to water (and not the reverse).
4. Acids and bases should be segregated for storage.
5. Liquid corrosives should be stored below eye level.
6. Adequate quantities of spill control materials should be readily available. Specialized
spill kits for acids and bases are available through most chemical and laboratory safety
supply catalogs.
B. Corrosive Gases and Vapors
Corrosive gases and vapors are hazardous to all parts of the body; certain organs (e.g. the eyes
and the respiratory tract) are particularly sensitive. The magnitude of the effect is related to the
solubility of the material in the body fluids. Highly soluble gases (e.g. ammonia, hydrogen
chloride) cause severe nose and throat irritation, while substances of lower solubility (e.g.
nitrogen dioxide, phosgene, sulfur dioxide) can penetrate deep into the lungs.
1. Warning properties such as odor or eye, nose or respiratory tract irritation may be
inadequate with some substances. Therefore, they should not be relied upon as a warning
of overexposure.
2. Perform manipulations of materials that pose an inhalation hazard in a chemical fume
hood to control exposure or wear appropriate respiratory protection.
3. Protect all exposed skin surfaces from contact with corrosive or irritating gases and
Updated Dec 2007
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4. Regulators and valves should be closed when the cylinder is not in use and flushed with
dry air or nitrogen after use.
5. When corrosive gases are to be discharged into a liquid, a trap, check valve, or vacuum
break device should be employed to prevent dangerous reverse flow.
C. Corrosive Solids
Corrosive solids, such as sodium hydroxide and phenol, can cause burns to the skin and eyes.
Dust from corrosive solids can be inhaled and cause irritation or burns to the respiratory tract.
Many corrosive solids, such as potassium hydroxide and sodium hydroxide, can produce
considerable heat when dissolved in water.
1. Wear gloves and eye protection when handling corrosive solids.
2. When mixing with water, always slowly add the corrosive solid to water, stirring
continuously. Cooling may be necessary.
3. If there is a possibility of generating a significant amount of dust, conduct work in a fume
8.3 Compressed Gases
 Hazards
 Handling Procedures
 Storage of Compressed Gas Cylinders
 Using Compressed Gas Cylinders
 Assembly of Equipment and Piping
 Leaking Cylinders
 Empty Cylinders
 Flammable Gases
 Highly Toxic Gases
 Gases Requiring Special Handling
Compressed gases can be toxic, flammable, oxidizing, corrosive, inert or a combination of
hazards. In addition to the chemical hazards, compressed gases may be under a great deal of
pressure. The amount of energy in a compressed gas cylinder makes it a potential rocket.
Appropriate care in the handling and storage of compressed gas cylinders is essential.
A. Hazards
The following is an overview of the hazards to be avoided when handling and storing
compressed gases:
Asphyxiation: Simple asphyxiation is the primary hazard associated with inert gases.
Because inert gases are colorless and odorless, they can escape into the atmosphere
undetected and quickly reduce the concentration of oxygen below the level necessary to
support life. The use of oxygen monitoring equipment is strongly recommended for
enclosed areas where inert gases are being used.
Updated Dec 2007
MET Module V
Fire and Explosion: Fire and explosion are the primary hazards associated with
flammable gases, oxygen and other oxidizing gases. Flammable gases can be ignited by
static electricity or by a heat source, such as a flame or a hot object. Oxygen and other
oxidizing gases do not burn, but will support combustion of organic materials. Increasing
the concentration of an oxidizer accelerates the rate of combustion. Materials that are
nonflammable under normal conditions may burn in an oxygen-enriched atmosphere.
Chemical Burns: Corrosive gases can chemically attack various materials, including
fire-resistant clothing. Some gases are not corrosive in their pure form, but can become
extremely destructive if a small amount of moisture is added. Corrosive gases can cause
rapid destruction of skin and eye tissue.
Chemical Poisoning: Chemical poisoning is the primary hazard of toxic gases. Even in
very small concentrations, brief exposure to these gases can result in serious poisoning
injuries. Symptoms of exposure may be delayed.
High Pressure: All compressed gases are potentially hazardous because of the high
pressure stored inside the cylinder. A sudden release of pressure can cause injuries by
propelling a cylinder or whipping a line.
Cylinder Weight: A full size cylinder may weigh more than 130 pounds. Moving a
cylinder manually may lead to back or muscle injury. Dropping or dragging a cylinder
could cause serious injury.
B. Handling Precautions
Avoid dropping, dragging or sliding cylinders. Use a suitable hand truck or cart equipped
with a chain or belt for securing the cylinder to the cart, even for short distances.
Do not permit cylinders to strike each other violently. Cylinders should not be used as
rollers for moving material or other equipment.
Cylinder caps should be left on each cylinder until it has been secured against a wall or
bench or placed in a cylinder stand, and is ready for installation of the regulator. Cylinder
caps protect the valve on top of the cylinder from damage if knocked.
Never tamper with pressure relief devices in valves or cylinders.
Use only wrenches or tools provided by the cylinder supplier to remove a cylinder cap or
to open a valve. Never use a screwdriver or pliers.
Keep the cylinder valve closed except when in use.
Position cylinders so that the cylinder valve is accessible at all times.
Use compressed gases only in a well-ventilated area. Toxic, flammable and corrosive
gases should be carefully handled in a hood or gas cabinet. Proper containment systems
should be used and minimum quantities of these products should be kept on-site.
When discharging gas into a liquid, a trap or suitable check valve should be used to
prevent liquid from getting back into the cylinder or regulator.
Where more than one type of gas is in use, label gas lines. This is particularly important
when the gas supply is not in the same room or area as the operation using the gases.
Do not use the cylinder valve itself to control flow by adjusting the pressure.
C. Storage of Compressed Gas Cylinders
All cylinders must be secured to a wall, bench or fixed support using a chain or strap
placed 2/3 of the way up. Cylinder stands are an alternative to straps.
Updated Dec 2007
MET Module V
Cylinders should be strapped individually.
Do not store full and empty cylinders together.
Oxidizers and flammable gases should be stored in areas separated by at least 20 feet or
by a noncombustible wall.
Cylinders should not be stored near radiators or other heat sources. If storage is outdoors,
protect cylinders from weather extremes and damp ground to prevent corrosion.
No part of a cylinder should be subjected to a temperature higher than 125oF. A flame
should never be permitted to come in contact with any part of a compressed gas cylinder.
Do not place cylinders where they may become part of an electric circuit.
Keep the number of cylinders in a laboratory to a minimum to reduce the fire and toxicity
Lecture bottles should always be returned to the distributor or manufacturer promptly
when no longer needed or discarded if at atmospheric pressure.
Ensure that the cylinder is properly and prominently labeled as to its contents.
NEVER place acetylene cylinders on their side.
D. Using Compressed Gas Cylinders
Before using cylinders, read all label information and material safety data sheets (MSDSs)
associated with the gas being used. The cylinder valve outlet connections are designed to prevent
mixing of incompatible gases. The outlet threads vary in diameter; some are internal and some
are external; some are right-handed and some are left-handed. Generally, right-handed threads
are used for fuel gases.
1. To set up and use the cylinder, follow these directions. Do not force the fitting. A poor
fit may indicate that the regulator is not intended for use on the gas chosen.
2. Turn the delivery pressure adjusting screw counter-clockwise until it turns freely. This
prevents unintended gas flow into the regulator.
3. Open the cylinder slowly until the inlet gauge on the regulator registers the cylinder
pressure. If the cylinder pressure reading is lower than expected, the cylinder valve may
be leaking.
4. With the flow control valve at the regulator outlet closed, turn the delivery pressure
adjusting screw clockwise until the required delivery pressure is reached.
5. Check for leaks using Snoop or soap solution. At or below freezing temperatures, use a
glycerin and water solution, such as Snoop, rather than soap. Never use an open flame to
detect leaks.
6. When finished with the gas, close the cylinder valve and release the regulator pressure.
E. Assembly of Equipment and Piping
Do not force threads that do not fit exactly.
Use Teflon tape or thread lubricant for assembly. Teflon tape should only be used for
tapered pipe thread, not straight lines or metal-to-metal contacts.
Avoid sharp bends of copper tubing. Copper tubing hardens and cracks with repeated
Updated Dec 2007
MET Module V
Inspect tubing frequently and replace when necessary.
Tygon and plastic tubing are not appropriate for most pressure work. These materials can
fail under pressure or thermal stress.
Do not mix different brands and types of tube fittings. Construction parts are usually not
Do not use oil or lubricants on equipment used with oxygen.
Do not use copper piping for acetylene.
Do not use cast iron piping for chlorine.
F. Leaking Cylinders
Most leaks occur at the valve in the top of the cylinder and may involve the valve threads valve
stem, valve outlet, or pressure relief devices. Lab personnel should not attempt to repair leaking
Where action can be taken without serious exposure to lab personnel:
1. Move the cylinder to an isolated, well-ventilated area (away from combustible materials
if the cylinder contains a flammable or oxidizing gas).
2. Contact Public Safety at 911.
Whenever a large or uncontrollable leak occurs, evacuate the area and immediately contact
Public Safety at 911.
G. Empty Cylinders
Remove the regulator and replace the cylinder cap.
Mark the cylinder as empty or MT and store in a designated area for return to the
Do not store full and empty cylinders together.
Do not have full and empty cylinders connected to the same manifold. Reverse flow can
occur when an empty cylinder is attached to a pressurized system.
Do not refill empty cylinders. Only the cylinder supplier should refill gases.
Do not empty cylinders to a pressure below 25 psi (172 Kpa). The residual contents may
become contaminated with air.
Lecture bottles should always be returned to the distributor or manufacturer promptly
when no longer needed. Do not purchase lecture bottles that cannot be returned.
H. Flammable Gases
Keep sources of ignition away from the cylinders.
Oxidizers and flammable gases should be stored in areas separated by at least 20 feet or
by a non-combustible wall.
Bond and ground all cylinders, lines and equipment used with flammable compressed
Updated Dec 2007
MET Module V
I. Highly Toxic Gases
Highly toxic gases, such as arsine, diborane, fluorine, hydrogen cyanide, phosgene, and silane,
can pose a significant health risk in the event of a leak. Use of these materials requires written
approval by the Principal Investigator or supervisor, using the Particularly Hazardous
Substances Use Approval form.
The following additional precautions must be taken:
Use and store in a specially ventilated gas cabinet or fume hood.
Use coaxial (double walled) tubing with nitrogen between the walls for feed lines
operating above atmospheric pressure.
Regulators should be equipped with an automatic shut-off to turn off gas supply in the
event of sudden loss of pressure in the supply line.
An alarm system should be installed to check for leaks in routinely used gases with poor
warning properties. The alarm level must be set at or lower than the permissible exposure
limit of the substance.
Self-contained breathing apparatus (SCBA) may be appropriate for changing cylinders of
highly toxic gases. Use of an SCBA requires enrollment in the Respiratory Protection
Program and annual training and fit-testing.
Ensure storage and use areas are posted with Designated Area signage.
J. Gases Requiring Special Handling
The following gases present special hazards either due to their toxicity or physical properties.
Review this information before using these gases.
Hydrogen Cyanide
Section 9: Safe Work Practices and Procedures
Handling Cryogenic Liquids
Protective Clothing
Cooling Baths and Dry Ice
Liquid Nitrogen Cooled Traps
Updated Dec 2007
MET Module V
Cryogenic liquids have boiling points less than -73ºC (-100ºF). Liquid nitrogen, liquid oxygen
and carbon dioxide are the most common cryogenic materials used in the laboratory. Hazards
may include fire, explosion, embrittlement, pressure buildup, frostbite and asphyxiation.
Many of the safety precautions observed for compressed gases also apply to cryogenic liquids.
Two additional hazards are created from the unique properties of cryogenic liquids:
Extremely Low Temperatures –The cold boil-off vapor of cryogenic liquids rapidly
freezes human tissue. Most metals become stronger upon exposure to cold temperatures,
but materials such as carbon steel, plastics and rubber become brittle or even fracture
under stress at these temperatures. Proper material selection is important. Cold burns and
frostbite caused by cryogenic liquids can result in extensive tissue damage.
 Vaporization - All cryogenic liquids produce large volumes of gas when they vaporize.
Liquid nitrogen will expand 696 times as it vaporizes. The expansion ratio of argon is
847:1, hydrogen is 851:1 and oxygen is 862:1. If these liquids vaporize in a sealed
container, they can produce enormous pressures that could rupture the vessel. (See
Anecdotes for an account of such an incident.) For this reason, pressurized cryogenic
containers are usually protected with multiple pressure relief devices.
Vaporization of cryogenic liquids (except oxygen) in an enclosed area can cause
asphyxiation. Vaporization of liquid oxygen can produce an oxygen-rich atmosphere, which
will support and accelerate the combustion of other materials. Vaporization of liquid
hydrogen can form an extremely flammable mixture with air.
A. Handling Cryogenic Liquids
Most cryogenic liquids are odorless, colorless, and tasteless when vaporized. When cryogenic
liquids are exposed to the atmosphere, the cold boil-off gases condense the moisture in the air,
creating a highly visible fog.
Always handle these liquids carefully to avoid skin burns and frostbite. Exposure that
may be too brief to affect the skin of the face or hands may damage delicate tissues, such
as the eyes.
 Boiling and splashing always occur when charging or filling a warm container with
cryogenic liquid or when inserting objects into these liquids. Perform these tasks slowly
to minimize boiling and splashing. Use tongs to withdraw objects immersed in a
cryogenic liquid.
 Never touch uninsulated pipes or vessels containing cryogenic liquids. Flesh will stick to
extremely cold materials. Even nonmetallic materials are dangerous to touch at low
Protective clothing must be worn when handling objects that come into contact with cryogenic
liquids and vapor. Trousers should be worn on the outside of boots or work shoes.
Updated Dec 2007
MET Module V
C. Cooling Baths and Dry Ice
Neither liquid nitrogen nor liquid air should be used to cool a flammable mixture in the
presence of air, because oxygen can condense from the air, leading to an explosion
Wear insulated, dry gloves and a face shield when handling dry ice.
Add dry ice slowly to the liquid portion of the cooling bath to avoid foaming over. Do not
lower your head into a dry ice chest, since suffocation can result from carbon dioxide
D. Liquid Nitrogen Cooled Traps
Traps that open to the atmosphere condense liquid air rapidly. If you close the system, pressure
builds up with enough force to shatter glass equipment. Therefore, only sealed or evacuated
equipment should use liquid nitrogen cooled traps.
Electrical Hazards
 Power Loss
Preventing Electrical Hazards
 Insulation
 Guarding
 Grounding
 Circuit Protection Devices
 Motors
Safe Work Practices
High Voltage or Current
Altering Building Wiring and Utilities
Electrically powered equipment, such as hot plates, stirrers, vacuum pumps, electrophoresis
apparatus, lasers, heating mantles, ultrasonicators, power supplies, and microwave ovens are
essential elements of many laboratories. These devices can pose a significant hazard to
laboratory workers, particularly when mishandled or not maintained. Many laboratory electrical
devices have high voltage or high power requirements, carrying even more risk. Large capacitors
found in many laser flash lamps and other systems are capable of storing lethal amounts of
electrical energy and pose a serious danger even if the power source has been disconnected.
Accounts of incidents on campus that resulted in electrical shock, including a near fatal incident,
are described in Anecdotes.
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A. Electrical Hazards
The major hazards associated with electricity are electrical shock and fire. Electrical shock
occurs when the body becomes part of the electric circuit, either when an individual comes in
contact with both wires of an electrical circuit, one wire of an energized circuit and the ground,
or a metallic part that has become energized by contact with an electrical conductor.
The severity and effects of an electrical shock depend on a number of factors, such as the
pathway through the body, the amount of current, the length of time of the exposure, and
whether the skin is wet or dry. Water is a great conductor of
electricity, allowing current to flow more easily in wet conditions and through wet skin. The
effect of the shock may range from a slight tingle to severe burns to cardiac arrest. The chart
below shows the general relationship between the degree of injury and amount of current for a
60-cycle hand-to-foot path of one second's duration of shock. While reading this chart, keep in
mind that most electrical circuits can provide, under normal conditions, up to 20,000
milliamperes of current flow
1 Milliampere
Perception level
5 Milliamperes
Slight shock felt; not painful but disturbing
6-30 Milliamperes
Painful shock; "let-go" range
50-150 Milliamperes
Extreme pain, respiratory arrest, severe
muscular contraction
1000-4,300 Milliamperes
Ventricular fibrillation
10,000+ Milliamperes
Cardiac arrest, severe burns and probable death
In addition to the electrical shock hazards, sparks from electrical equipment can serve as an
ignition source for flammable or explosive vapors or combustible materials. See Anecdotes.
Power Loss
Loss of electrical power can create hazardous situations. Flammable or toxic vapors may be
released as a chemical warms when a refrigerator or freezer fails. Fume hoods may cease to
operate, allowing vapors to be released into the laboratory. If magnetic or mechanical stirrers fail
to operate, safe mixing of reagents may be compromised.
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B. Preventing Electrical Hazards
There are various ways of protecting people from the hazards caused by electricity, including
insulation, guarding, grounding, and electrical protective devices. Laboratory workers can
significantly reduce electrical hazards by following some basic precautions:
Inspect wiring of equipment before each use. Replace damaged or frayed electrical cords
Use safe work practices every time electrical equipment is used.
Know the location and how to operate shut-off switches and/or circuit breaker panels.
Use these devices to shut off equipment in the event of a fire or electrocution.
Limit the use of extension cords. Use only for temporary operations and then only for
short periods of time. In all other cases, request installation of a new electrical outlet.
Multi-plug adapters must have circuit breakers or fuses.
Place exposed electrical conductors (such as those sometimes used with electrophoresis
devices) behind shields.
Minimize the potential for water or chemical spills on or near electrical equipment.
All electrical cords should have sufficient insulation to prevent direct contact with wires. In a
laboratory, it is particularly important to check all cords before each use, since corrosive
chemicals or solvents may erode the insulation.
Damaged cords should be repaired or taken out of service immediately, especially in wet
environments such as cold rooms and near water baths.
Live parts of electric equipment operating at 50 volts or more (i.e., electrophoresis devices) must
be guarded against accidental contact. Plexiglas shields may be used to protect against exposed
live parts.
Only equipment with three-prong plugs should be used in the laboratory. The third prong
provides a path to ground for internal electrical short circuits, thereby protecting the user from a
potential electrical shock.
Circuit Protection Devices
Circuit protection devices are designed to automatically limit or shut off the flow of electricity
in the event of a ground-fault, overload or short circuit in the wiring system. Ground-fault
circuit interrupters, circuit breakers and fuses are three well-known examples of such devices.
Fuses and circuit breakers prevent over-heating of wires and components that might otherwise
create fire hazards. They disconnect the circuit when it becomes overloaded. This overload
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protection is very useful for equipment that is left on for extended periods of time, such as
stirrers, vacuum pumps, drying ovens, Variacs and other electrical equipment.
The ground-fault circuit interrupter, or GFCI, is designed to shutoff electric power if a ground
fault is detected, protecting the user from a potential electrical shock. The GFCI is particularly
useful near sinks and wet locations. Since GFCIs can cause equipment to shutdown
unexpectedly, they may not be appropriate for certain apparatus. Portable GFCI adapters
(available in most safety supply catalogs) may be used with a non-GFCI outlet.
In laboratories where volatile flammable materials are used, motor-driven electrical equipment
should be equipped with non-sparking induction motors or air motors. These motors must meet
National Electric Safety Code (US DOC, 1993) Class 1, Division 2, Group C-D explosion
resistance specifications. Many stirrers, Variacs, outlet strips, ovens, heat tape, hot plates and
heat guns do not comform to these code requirements.
Avoid series-wound motors, such as those generally found in some vacuum pumps, rotary
evaporators and stirrers. Series-wound motors are also usually found in household appliances
such as blenders, mixers, vacuum cleaners and power drills. These appliances should not be used
unless flammable vapors are adequately controlled.
Although some newer equipment have spark-free induction motors, the on-off switches
and speed controls may be able to produce a spark when they are adjusted because they
have exposed contacts. One solution is to remove any switches located on the devices to
insure it is safe to do so, work with only one hand, keeping the other hand at your side or
in your pocket, away from all conductive material. This precaution reduces the likelihood
of accidents that result in current passing through the chest cavity.
Minimize the use of electrical equipment in cold rooms or other areas where
condensation is likely. If equipment must be used in such areas, mount the equipment on
a wall or vertical panel.
If water or a chemical is spilled onto equipment, shut off power at the main switch or
circuit breaker and unplug the equipment.
If an individual comes in contact with a live electrical conductor, do not touch the
equipment, cord or person. Disconnect the power source from the circuit breaker or pull
out the plug using a leather belt.
D. High Voltage or Current
Repairs of high voltage or high current equipment should be performed only by trained
electricians. Laboratory workers who are experienced in such tasks and would like to perform
such work on their own laboratory equipment must first receive specialized electrical safety
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related work practices training by EHS staff. Contact the University Safety Engineer at 258-5294
for more information.
E. Altering Building Wiring and Utilities
Any modifications to existing electrical service in a laboratory or building must be completed or
approved by either the building facility manager, an engineer from the Facilities department or
the building's Special Facilities staff. All modifications must meet both safety standards and
Facilities Engineering design requirements.
Any unapproved laboratory facilities modifications discovered during laboratory surveys or other
activities are reviewed by EHS and facility staff to determine whether they meet design
11.0 Laboratory Equipment Safety
 Refrigerators and Freezers
 Stirring and Mixing Devices
 Heating Devices
 Ultrasonicators
 Centrifuges
 Rotovaps
 Autoclaves
 Electrophoresis Devices
 Glassware
 Vacuums
A. Refrigerators and Freezers The potential hazards posed by laboratory refrigerators and
freezers involve vapors from the contents, the possible presence of incompatible chemicals and
Only refrigerators and freezers specified for laboratory use should be utilized for the storage of
chemicals. These refrigerators have been constructed with special design factors, such as heavyduty cords and corrosion resistant interiors to help reduce the risk of fire or explosions in the lab.
Standard refrigerators have electrical fans and motors that make them potential ignition sources
for flammable vapors. Do not store flammable liquids in a refrigerator unless it is approved for
such storage. Flammable liquid-approved refrigerators are designed with spark-producing parts
on the outside to avoid accidental ignition. If refrigeration is needed inside a flammable-storage
room, you should use an explosion-proof refrigerator.
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Frost-free refrigerators should also be avoided. Many of them have a drain or tube or hole that
carries water and possibly any spilled materials to an area near the compression, which may
spark. Electric heaters used to defrost the freezing coils can also spark.
Only chemicals should be stored in chemical storage refrigerators; lab refrigerators should not be
used for food storage or preparation. Refrigerators should be labeled for their intended purpose;
labels reading “No Food or Drink to be Stored in this Refrigerator” or “Refrigerator For Food
Only” are available from EHS by calling 8-5294.
All materials in refrigerators or freezers should be labeled with the contents, owner, date of
acquisition or preparation and nature of any potential hazard. Since refrigerators are often used
for storage of large quantities of small vials and test tubes, a reference to a list outside of the
refrigerator could be used. Labels and ink used to identify materials in the refrigerators should be
All containers should be sealed, preferably with a cap. Containers should be placed in secondary
containers, or catch pans should be used.
Loss of electrical power can produce extremely hazardous situations. Flammable or toxic vapors
may be released from refrigerators and freezers as chemicals warm up and/or certain reactive
materials may decompose energetically upon warming.
B. Stirring and Mixing Devices The stirring and mixing devices commonly found in laboratories
include stirring motors, magnetic stirrers, shakers, small pumps for fluids and rotary evaporators
for solvent removal. These devices are typically used in laboratory operations that are performed
in a hood, and it is important that they be operated in a way that precludes the generation of
electrical sparks.
Only spark-free induction motors should be used in power stirring and mixing devices or any
other rotating equipment used for laboratory operations. While the motors in most of the
currently marketed stirring and mixing devices meet this criterion, their on-off switches and
rheostat-type speed controls can produce an electrical spark because they have exposed electrical
conductors. The speed of an induction motor operating under a load should not be controlled by
a variable autotransformer.
Because stirring and mixing devices, especially stirring motors and magnetic stirrers, are often
operated for fairly long periods without constant attention, the consequences of stirrer failure,
electrical overload or blockage of the motion of the stirring impeller should be considered.
C. Heating Devices Most labs use at least one type of heating device, such as ovens, hot plates,
heating mantles and tapes, oil baths, salt baths, sand baths, air baths, hot-tube furnaces, hot-air
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guns and microwave ovens. Steam-heated devices are generally preferred whenever temperatures
of 100o C or less are required because they do not present shock or spark risks and can be left
unattended with assurance that their temperature will never exceed 100o C.
A number of general precautions need to be taken when working with heating devices in the
laboratory. When working with heating devices, consider the following:
· The actual heating element in any laboratory heating device should be enclosed in such a
fashion as to prevent a laboratory worker or any metallic conductor from accidentally touching
the wire carrying the electric current.
· Heating device becomes so worn or damaged that its heating element is exposed, the device
should be either discarded or repaired before it is used again.
· Laboratory heating devices should be used with a variable autotransformer to control the input
voltage by supplying some fraction of the total line voltage, typically 110 V.
· The external cases of all variable autotransformers have perforations for cooling by ventilation
and, therefore, should be located where water and other chemicals cannot be spilled onto them
and where they will not be exposed to flammable liquids or vapors.
Fail-safe devices can prevent fires or explosions that may arise if the temperature of a reaction
increases significantly because of a change in line voltage, the accidental loss of reaction solvent
or loss of cooling. Some devices will turn off the electric power if the temperature of the heating
device exceeds some preset limit or if the flow of cooling water through a condenser is stopped
owing to the loss of water pressure or loosening of the water supply hose to a condenser.
C.1 Ovens
Electrically heated ovens are commonly used in the laboratory to remove water or other solvents
from chemical samples and to dry laboratory glassware. Never use laboratory ovens for human
food preparation.
· Laboratory ovens should be constructed such that their heating elements and their temperature
controls are physically separated from their interior atmospheres.
· Laboratory ovens rarely have a provision for preventing the discharge of the substances
volatilized in them. Connecting the oven vent directly to an exhaust system can reduce the
possibility of substances escaping into the lab or an explosive concentration developing within
the oven.
· Ovens should not be used to dry any chemical sample that might pose a hazard because of acute
or chronic toxicity unless special precautions allow for these procedures and materials are in
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closed environment. If mercury is spilled in the oven, the oven should remain closed and turned
off immediately, and it should remain closed until cool. All mercury should be removed from the
cold oven with the use of appropriate cleaning equipment and procedures in order to avoid
mercury exposure.
C.2 Hot Plates
Laboratory hot plates are normally used for heating solutions to 100o C or above when inherently
safer steam baths cannot be used. Any newly purchased hot plates should be designed in a way
that avoids electrical sparks. However, many older hot plates pose an electrical spark hazard
arising from either the on-off switch located on the hot plate, the bimetallic thermostat used to
regulate the temperature or both. Laboratory workers should be warned of the spark hazard
associated with older hot plates.
In addition to the spark hazard, old and corroded bimetallic thermostats in these devices can
eventually fuse shut and deliver full, continuous current to a hot plate.
· Do not store volatile flammable materials near a hot plate
· Limit use of older hot plates for flammable materials.
· Check for corrosion of thermostats. Corroded bimetallic thermostats can be repaired or
reconfigured to avoid spark hazards. Contact EHS for more info.
C.3 Heating Mantles
Heating mantles are commonly used for heating round-bottomed flasks, reaction kettles and
related reaction vessels. These mantles enclose a heating element in a series of layers of
fiberglass cloth. As long as the fiberglass coating is not worn or broken, and as long as no water
or other chemicals are spilled into the mantle, heating mantles pose no shock hazard.
· Always use a heating mantle with a variable autotransformer to control the input voltage. Never
plug them directly into a 110-V line.
· Be careful not to exceed the input voltage recommended by the mantle manufacturer. Higher
voltages will cause it to overheat, melt the fiberglass insulation and expose the bare heating
· If the heating mantle has an outer metal case that provides physical protection against damage
to the fiberglass, it is good practice to ground the outer metal case to protect against an electric
shock if the heating element inside the mantle shorts against the metal case.
· Some older equipment might have asbestos insulation rather than fiberglass. Contact EHS to
replace the insulation and for proper disposal of the asbestos.
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C.4 Oil, Salt and Sand Baths
Electrically heated oil baths are often used to heat small or irregularly shaped vessels or when a
stable heat source that can be maintained at a constant temperature is desired. Molten salt baths,
like hot oil baths, offer the advantages of good heat transfer, commonly have a higher operating
range (e.g., 200 to 425oC) and may have a high thermal stability (e.g., 540oC).There are several
precautions to take when working with these types of heating devices:
· Take care with hot oil baths not to generate smoke or have the oil burst into flames from
· Always monitor oil baths by using a thermometer or other thermal sensing devices to ensure
that its temperature does not exceed the flash point of the oil being used.
· Fit oil baths left unattended with thermal sensing devices that will turn off the electric power if
the bath overheats.
· Mix oil baths well to ensure that there are no “hot spots” around the elements that take the
surrounding oil to unacceptable temperatures.
· Contain heated oil in a vessel that can withstand an accidental strike by a hard object.
· Mount baths carefully on a stable horizontal support such as a laboratory jack that can be raised
or lowered without danger of the bath tipping over. Iron rings are not acceptable supports for hot
· Clamp equipment high enough above a hot bath that if the reaction begins to overheat, the bath
can be lowered immediately and replaced with a cooling bath without having to readjust the
equipment setup.
· Provide secondary containment in the event of a spill of hot oil.
· Wear heat-resistant gloves when handling a hot bath.
· The reaction container used in a molten salt bath must be able to withstand a very rapid heat-up
to a temperature above the melting point of salt.
· Take care to keep salt baths dry since they are hygroscopic, which can cause hazardous
popping and splattering if the absorbed water vaporizes during heat-up.
C.5 Hot Air Baths and Tube Furnaces
Hot air baths are used in the lab as heating devices. Nitrogen is preferred for reactions involving
flammable materials. Electrically heated air baths are frequently used to heat small or irregularly
shaped vessels. One drawback of the hot air bath is that they have a low heat capacity. As a
result, these baths normally have to be heated to 100oC or more above the target temperature.
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Tube furnaces are often used for high-temperature reactions under pressure. Consider the
following when working with either apparatus:
· Ensure that the heating element is completely enclosed.
· For air baths constructed of glass, wrap the vessel with heat resistant tape to contain the glass if
it should break.
· Sand baths are generally preferable to air baths.
· For tube furnaces, carefully select glassware and metal tubes and joints to ensure they are able
to withstand the pressure.
· Follow safe practices outlined for both electrical safety and pressure and vacuum systems.
C.6 Heat Guns
Laboratory heat guns are constructed with a motor-driven fan that blows air over an electrically
heated filament. They are frequently used to dry glassware or to heat the upper parts of a
distillation apparatus during distillation of high-boiling materials.
The heating element in a heat gun typically becomes red-hot during use and the on-off switches
and fan motors are not usually spark-free. For these reasons, heat guns almost always pose a
serious spark hazard. See Anecdote.
· Household hair dryers may be substituted for laboratory heat guns only if they have a grounded
plug or are double insulated.
· Any hand-held heating device of this type that will be used in a laboratory should have groundfault circuit interrupter (GFCI) protection to ensure against electric shock.
· Never use a heat gun near flammable materials including open containers of flammable liquids,
flammable vapors or hoods used to control flammable vapors.
C.7 Microwave Ovens
Microwave ovens used in the laboratory may pose several different types of
· As with most electrical apparatus, there is the risk of generating sparks that can ignite
flammable vapors. · Metals placed inside the microwave oven may produce an arc that can ignite
flammable materials. · Materials placed inside the oven may overheat and ignite. · Sealed
containers, even if loosely sealed, can build pressure upon expansion during heating, creating a
risk of container rupture.
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To minimize the risk of these hazards,
· Never operate microwave ovens with doors open in order to avoid exposure to microwaves. ·
Do not place wires and other objects between the sealing surface and the door on the oven’s front
face. The sealing surfaces must be kept absolutely clean. · Never use a microwave oven for both
laboratory use and food preparation. · Electrically ground the microwave. If use of an extension
cord is necessary, only a three-wire cord with a rating equal to or greater than that for the oven
should be used. · Do not use metal containers and metal-containing objects (e.g., stir bars) in the
microwave. They can cause arcing. · Do not heat sealed containers in the microwave oven. Even
heating a container with a loosened cap or lid poses a significant risk since microwave ovens can
heat material so quickly that the lid can seat upward against the threads and containers can
explode. · Remove screw caps from containers being microwaved. If the sterility of the contents
must be preserved, use cotton or foam plugs. Otherwise the plug the container with kimwipes to
reduce splash potential. D. Ultrasonicators Human exposure to ultrasound with frequencies
between 16 and 100 kilohertz (kHz) can be divided into three distinct categories: airborne
conduction, direct contact through a liquid coupling medium, and direct contact with a vibrating
Ultrasound through airborne conduction does not appear to pose a significant health hazard to
humans. However, exposure to the associated high volumes of audible sound can produce a
variety of effects, including fatigue, headaches, nausea and tinnitus. When ultrasonic equipment
is operated in the laboratory, the apparatus must be enclosed in a 2-cm thick wooden box or in a
box lined with acoustically absorbing foam or tiles to substantially reduce acoustic emissions
(most of which are inaudible).
Direct contact of the body with liquids or solids subjected to high-intensity ultrasound of the sort
used to promote chemical reactions should be avoided. Under sonochemical conditions,
cavitation is created in liquids, and it can induce high-energy chemistry in liquids and tissues.
Cell death from membrane disruption can occur even at relatively low acoustic intensities.
Exposure to ultrasonically vibrating solids, such as an acoustic horn, can lead to rapid frictional
heating and potentially severe burns.
E. Centrifuges Centrifuges should be properly installed and must be operated only by trained
personnel. It is important that the load is balanced each time the centrifuge is used and that the
lid be closed while the rotor is in motion. The disconnect switch must be working properly to
shut off the equipment when the top is opened, and the manufacturer’s instructions for safe
operating speeds must be followed.
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For flammable and/or hazardous materials, the centrifuge should be under negative pressure to a
suitable exhaust system.
F. Rotary Evaporators Glass components of the rotary evaporator should be made of Pyrex or
similar glass. Glass vessels should be completely enclosed in a shield to guard against flying
glass should the components implode. Increase in rotation speed and application of vacuum to
the flask whose solvent is to be evaporated should be gradual.
G. Autoclaves The use of an autoclave is a very effective way to decontaminate infectious waste.
Autoclaves work by killing microbes with superheated steam. The following are recommended
guidelines when using an autoclave:
· Do not put sharp or pointed contaminated objects into an autoclave bag. Place them in an
appropriate rigid sharps disposal container.
· Use caution when handling an infectious waste autoclave bag, in case sharp objects were
inadvertently placed in the bag. Never lift a bag from the bottom to load it into the chamber.
Handle the bag from the top.
· Do not overfill an autoclave bag. Steam and heat cannot penetrate as easily to the interior of a
densely packed autoclave bag. Frequently the outer contents of the bag will be treated but the
innermost part will be unaffected.
· Do not overload an autoclave. An overpacked autoclave chamber does not allow efficient steam
distribution. Considerably longer sterilization times may be required to achieve decontamination
if an autoclave is tightly packed.
· Conduct autoclave sterility testing on a regular basis using appropriate biological indicators (B.
stearothermophilus spore strips) to monitor efficacy. Use indicator tape with each load to verify
it has been autoclaved.
· Do not mix contaminated and clean items together during the same autoclave cycle. Clean
items generally require shorter decontamination times (15-20 minutes) while a bag of infectious
waste (24" x 36") typically requires 45 minutes to an hour to be effectively decontaminated
· Always wear personal protective equipment, including heat-resistant gloves, safety glasses and
a lab coat when operating an autoclave. Use caution when opening the autoclave door. Allow
superheated steam to exit before attempting to remove autoclave contents.
· Be on the alert when handling pressurized containers. Superheated liquids may spurt from
closed containers. Never seal a liquid container with a cork or stopper. This could cause an
explosion inside the autoclave.
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· Agar plates will melt and the agar will become liquefied when autoclaved. Avoid contact with
molten agar. Use a secondary tray to catch any potential leakage from an autoclave bag rather
than allowing it to leak onto the floor of the autoclave chamber.
· If there is a spill inside the autoclave chamber, allow the unit to cool before attempting to clean
up the spill. If glass breaks in the autoclave, use tongs, forceps or other mechanical means to
recover fragments. Do not use bare or gloved hands to pick up broken glassware.
· Do not to leave an autoclave operating unattended for a long period of time. Always be sure
someone is in the vicinity while an autoclave is cycling in case there is a problem.
· Autoclaves should be placed under preventive maintenance contracts to ensure they are
operating properly.
H. Electrophoresis Devices
Precautions to prevent electric shock must be followed when conducting procedures involving
electrophoresis. Lethal electric shock can result when operating at high voltages such as in DNA
sequencing or low voltages such as in agarose gel electrophoresis (e.g., 100 volts at 25
milliamps).These general guidelines should be followed:
· Turn the power off before connecting the electrical leads
· Connect one lead at a time, using one hand only
· Ensure that hands are dry while connecting leads
· Keep the apparatus away from sinks or other water sources
· Turn off power before opening lid or reaching inside chamber
· Do not override safety devices
· Do not run electrophoresis equipment unattended.
· If using acrylamide, purchase premixed solutions or pre-weighed quantities whenever possible
· If using ethidium bromide, have a hand-held UV light source available in the laboratory. Check
working surfaces after each use.
· Mix all stock solutions in a chemical fume hood.
· Provide spill containment by mixing gels on a plastic tray
· Decontaminate surfaces with ethanol. Dispose of all cleanup materials as hazardous waste.
I. Glassware
Although glass vessels are frequently used in low-vacuum operations, evacuated glass vessels
may collapse violently, either spontaneously from strain or from an accidental blow. Therefore,
pressure and vacuum operations in glass vessels should be conducted behind adequate shielding.
It is advisable to check for flaws such as star cracks, scratches and etching marks each time a
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MET Module V
vacuum apparatus is used. Only round-bottomed or thick-walled (e.g., Pyrex) evacuated reaction
vessels specifically designed for operations at reduced pressure should be used. Repaired
glassware is subject to thermal shock and should be avoided. Thin-walled, Erlenmeyer or roundbottomed flasks larger than 1 L should never be evacuated.
J. Vacuums
Vacuum pumps are used in the lab to remove air and other vapors from a vessel or manifold. The
most common usages are on rotary evaporators, drying manifolds, centrifugal concentrators
(“speedvacs”), acrylamide gel dryers, freeze dryers, vacuum ovens, tissue culture filter flasks and
aspirators, desiccators, filtration apparatus and filter/degassing apparatus.
The critical factors in vacuum pump selection are:
 Application the pump will be used on
 Nature of the sample (air, chemical, moisture)
 Size of the sample(s)
When using a vacuum pump on a rotary evaporator, a dry ice alcohol slurry cold trap or a
refrigerated trap is recommended. A Cold Trap should be used in line with the pump when high
vapor loads from drying samples will occur. Consult manufacturer for specific situations. These
recommendations are based on keeping evaporating flask on rotary evaporator at 400 C.
Operating at a higher temperature allows the Dry Vacuum System to strip boiling point solvents
with acceptable evaporation rates.
Vacuum pumps can pump vapors from air, water to toxic and corrosive materials like TFA and
methylene chloride. Oil seal pumps are susceptible to excessive amounts of gas, corrosive acids
and bases and excessive water vapors. Too much air mass over time s compressed by these
pumps and overheats the oil, cooing it into a viscous sludge that can eventually freeze the
pumping motor. Corrosives can also create sludge by breaking down the oil. Excess water thins
the lubricating properties of the oil, which then overheats. Proper trapping (cold trap, acid trap)
and routine oil changes greatly extend the life of an oil seal vacuum.
Diaphragm pumps are virtually impervious to attack from laboratory chemical vapors. They are
susceptible to physical wearing of the membrane if excessive chemical vapors are allowed to
condense and crystallize in the pumping chambers. A five minute air purge either as part of the
procedure or at day’s end will drive off condensed water vapors and further prolong pump life.
Hazardous chemicals can escape from the vacuum pump and pump should be place in the hood.
Cold traps and acid traps can be helpful, but if allowed to thaw or saturate, they can lose their
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Section 11: Anecdotes
Accidents do happen in Princeton University laboratories. The following are accounts of a
few incidents that help to illustrate the need for the safety precautions outlined in this
Improper Shelving
Chemical Burns
Glass Vessel Ruptures
Incidents Involving Reactive Materials
Electrical Shock
Improper Shelving
Wall-Mounted Shelves Collapse
There have been several incidents where wall-mounted shelves detached and fell onto
desks and other work surfaces, dumping the shelving and books all over the work area. In
one case, a person working nearby was injured as a result. In each instance, the shelves
were heavily loaded and either exceeded the load capacity of the shelving or was
incorrectly installed..
Wall-mounted shelving should have heavy-duty brackets and standards and
should be attached to studs or solid blocking.
For books and periodicals, bookcases are preferable to wall-mounted shelving.
See Safe Work Practices - General for more information.
Shelf of Chemical Storage Cabinet Collapses
The bottom shelf of an organic chemical storage cabinet spontaneously collapsed. This
shelf was not a moveable shelf, but a bottom panel contributing to the structural integrity
of the cabinet. Fortunately, the drop was only a few inches and none of the bottles of
chemicals were broken.
The cabinet was constructed of thin plywood with particleboard shelves attached to a
pressed paperboard backing. This type of cabinet is not appropriate for chemical storage.
Only sturdy wood or metal cabinets should be used for chemical storage.
Be sure to check the shelf load capacity before using any storage cabinets or
shelving units.
See Safe Work Practices - General for more information.
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Fire/Burn from Heating Flammable Solvent with Heat Gun
A laboratory worker was using a heat gun to heat approximately 0.5 liters of heptane in a
Pyrex beaker by hand over an open bench. A splash of heptane came in contact with the
elements of the heat gun, igniting the heptane and causing him to toss the beaker away
from him. The sleeve of the worker's shirt caught fire. The flaming beaker landed on
another work surface, spreading the fire to his computer. The worker immediately used a
safety shower to put out the fire on his clothing, then used a dry chemical fire
extinguisher to put out the other fire.
The worker received burns to his hand. The computer containing his thesis was destroyed
by the powder from the extinguisher.
Flammable liquids should be handled in a fume hood to prevent accumulation of
Heat guns and other equipment capable of igniting flammable vapors should not
be used to heat flammable vapors.
Heating operations should not be carried out by hand. Instead, a lab stand and
clamps should be used for this type of work.
Carbon dioxide extinguishers should be used around sensitive equipment. Dry
powder extinguishers can damage such equipment.
If clothing is on fire, smother the flame by rolling on the ground or use a safety
shower to extinguish the fire, as was done in this incident.
See Safe Work Practices - Flammable Materials and Electrical Safety for more
Hood Fire Involving Unattended Operation with Hexane Near Hot Plate
A fire erupted inside a hood containing two reactions running unattended. A laboratory
worker had placed nitrobenzene inside an oil bath atop a hot plate. The hot plate had been
operating for three days, heating the oil bath to 200 C. A plastic squeeze bottle of hexane
was placed next to the hot plate. Eventually, the squeeze bottle warmed enough to
pressurize the container, forcing liquid hexane out of the bottle and onto the hot plate,
where it ignited. Another laboratory noticed the smoke and attempted to put out the fire
using a dry chemical extinguisher. A maintenance worker also noticed the fire and
assisted the laboratory worker. Their attempts were not successful.
The fire department was dispatched. Since the Emergency Information Poster on the door
to the laboratory was inaccurate and there was a significant language barrier between the
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laboratory worker and the fire department personnel, a hazmat response team was
dispatched. Frick, New Frick and Hoyt were evacuated for more than three hours. The
laboratory worker and the maintenance worker were showered and scrubbed by the
hazmat team and their clothing was confiscated (it was later washed and returned to
them). While this was probably an overreaction by the emergency response personnel, it
illustrates the implications of not having an accurate, up-to-date emergency information
Containers of volatile liquids placed near heat sources can become pressurized.
Materials not involved in an experiment should be removed, as possible, to avoid
having them become involved in a fire or other incident.
Keeping the Emergnecy Information Poster up-to-date helps to ensure a
proportionate response by emergency response personnel.
Evaluate the potential problems related to experiments left unattended for days at
a time.
See Safe Work Practices - Flammable Materials and Electrical Safety for more
Chemical Burns
Hydrofluoric Acid Burn from Trifluoracetic Acid
A laboratory worker picked up a container of trifluoroacetic acid with her ungloved hand
to move it. She did not notice that there was a small amount of residue on the glass.
Several hours later, she experienced pain in the palm of her hand and the inside aspect of
her thumb. The result was a serious burn that required skin grafting. She was not aware
that this type of burn could result from handling trifluoracetic acid.
Trifluoracetic acid can form hydrofluoric acid upon contact with moisture. Hydrofluoric
acid can cause deep burns that may not be painful for hours.
Know the hazards of the chemicals involved before handling them.
Always assume containers are contaminated and wear appropriate gloves when
handling chemical containers.
Keep a hydrofluoric acid burn kit in the laboratory when working with
hydrofluoric acid or trifluoracetic acid.
Chemical Splash While Carrying Chemicals Incorrectly
A laboratory worker received burns to the face and chest while carrying chemicals from
one area of the laboratory to another. The worker placed unsealed centrifuge tubes filled
with phenol-chloroform into a Styrofoam centrifuge tube shipping container. The
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Styrofoam broke and the phenol-chloroform splashed onto the worker’s face and dripped
down the chest. The worker immediately flushed the area with a drench hose, but still
suffered from second-degree burns to the face, chest and abdomen. Fortunately, the
worker was wearing chemical splash goggles and did not receive burns to the eyes.
Appropriate eye and face protection helped to minimize the chemical burn.
Wear a closed lab coat when working with hazardous materials.
Use a plastic centrifuge rack instead of a Styrofoam packing container,
particularly when transporting chemicals..
Overpressurization of Gel Column Causes Chemical Splash
A laboratory worker was pouring chloroform though a gel column inside a fume hood.
Due to incorrect equipment configuration, pressure built up in the column and caused the
glassware at the top of the column to break, spraying chloroform out of the hood, onto the
worker’s face, eyes and clothing.
The laboratory worker was wearing safety glasses, rather than chemical splash goggles.
The chloroform seeped through the opening at the top of the glasses and burned both
eyes. The lens of the safety glasses were partially dissolved by the chloroform. The
worker did use a safety shower immediately, but was too embarrassed to remove his
sweater in the presence of other laboratory workers. As a result, he suffered from second
degree burns on both arms where the chloroform soaked through the sweater.
The set-up of the apparatus was changed to allow the hood of the sash to be lowered
when the chloroform is being poured, providing an additional shield between the worker
and the chemical and lowering the potential spray below eye level.
Keep hazardous materials that have the potential for splash below eye level.
Use care when working with pressure or vacuum to avoid pressurizing containers.
Wear a closed lab coat, chemical splash goggles and, if necessary, a face shield
when there is a possibility of a significant chemical splash.
Remove contaminated clothing while rinsing.
Keep the hood sash lowered and/or use shielding when working with pressurized
Failure to Remove Contaminated Clothing Exacerbates Chemical Burns
There have been several incidents, usually involving phenol, where laboratory workers
spilled a chemical on his or her pants. In all cases, the worker bypassed the safety shower
and entered a restroom to remove the pants and rinse the leg. In each case, the worker put
the contaminated pants back on and either went home to rinse further or went to McCosh
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Health Center. All resulted in second degree burns that could have been minimized by
taking off the contaminated clothing and rinsing immediately using a safety shower or
drench hose.
Remove contaminated clothing while rinsing.
Wear appropriate personal protective equipment, including a closed lab coat when
working with hazardous materials.
Do not put contaminated clothing back on.
Wash clothing separately or discard. Many chemicals can permeate leather.
Discard any contaminated leather items..
Mixing Incompatible Wastes
A laboratory worker was cleaning out chemicals from an old refrigerator. Wearing
gloves, chemical splash goggles and a lab coat (over shorts), the worker was segregating
the chemicals into several different waste containers. He found a small bottle of iodine
monochloride, and not knowing the physical properties of the chemical, began to pour it
into a jar with other liquid wastes. The waste container suddenly began fuming
vigorously, startling the worker and causing the worker to drop the bottle of iodine
monochloride. Several drops of the chemical splashed onto the worker's leg, causing a
second degree burn.
The iodine monochloride reacted with a chemical in the waste container. The worker was
fortunate that the reaction did not produce significant amounts of hazardous vapors. Had
the worker been wearing long pants, the burn might have been avoided.
Never mix chemicals unless you are certain of the consequences and are prepared
to control the hazard.
Do not mix incompatible waste chemicals together.
Know the hazards of each chemical before working with it.
Wear pants and a closed lab coat when working with hazardous materials.
See Laboratory Waste Disposal for more information.
Glass Vessel Ruptures
Glass Flask Rupture During Ozonolysis
During an early attempt to scale up a procedure, a laboratory worker introduced ozone
gas into a flask containing a small amount of organic material. The flask was set in a
fume hood in a cooling bath designed to lower the experiment temperature to -85 C, 15
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C below that which is normally used for such experiments. The sash of the fume hood
was completely raised. During the procedure, the worker noticed that a deep blue color
had developed in the flask, an indication that the concentration of ozone was increased.
He attributed it to poor mixing and had started to increase the stir rate when the flask
exploded. Flying glass embedded into his face, neck and safety glasses.
The worker did not experience any injuries to his eyes. Many of the cuts on his face and
neck required stitches. Shards of glass remain in the safety glasses even today.
The sash of the hood might have provided enough of a barrier to avoid injury.
However, most sashes are not designed to protect against explosions.
Shielding should be used around any experiments that might explode.
A face shield would have protected the worker from the cuts on his face and neck.
Carefully evaluate the hazards before proceeding with a scaled-up experiment.
Glass Flask Ruptures, Possibly Overpressurization by Liquid Nitrogen
A 250 ml glass flask became overpressurized and burst, spraying two laboratory workers
with shards of glass.
Approximately 10 grams of styrene and a minute quantity of a drying agent were
immersed in liquid nitrogen to keep the contents frozen. The laboratory worker then
attached the flask to a vacuum pump to evacuate the flask, without success. Thinking the
flask might have developed a crack, the laboratory worker removed the flask from the
vacuum line and was defrosting it under warm water in the sink, holding it and examining
it, when the flask ruptured.
The best guess as to the cause of the rupture is that a small leak, perhaps a pinhole in the
flask, developed while it was being frozen and that some liquid nitrogen entered the flask.
When the flask was warmed, the liquid nitrogen vaporized (expansion ratio 696:1),
overpressurizing the flask and leading to the explosion.
The laboratory worker holding the flask suffered from several lacerations to the face,
hands, chest and abdomen. The other worker, who was standing across the room,
received lacerations to the abdomen. The worker holding the flask noted shards of glass
embedded in his prescription safety glasses.
The procedure was re-written such that under the same conditions, the stopcock will be
unscrewed and the flask set in a catchbucket in the hood to allow the contents to warm up
and vaporize, if volatile.
Appropriate eye protection helped to avoid a potentially serious eye injury.
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Consider shielding operations involving vacuum or pressurization.
Be aware of the potential for pressurization when working with liquid nitrogen.
See Safe Work Practices - Pressure and Vacuum Systems for more information.
Glass Waste Bottle Ruptures, Possible Reaction of Incompatible Chemical
A graduate student sitting at a lab computer was surprised by a chemical waste bottle
which burst and sprayed nitric acid and shards of glass all over the lab.
Approximately 2L of nitric acid waste had been accumulated in a chemical waste bottle
which originally contained methanol. Over the course of 12-16 hours, it is likely that
some residual methanol reacted with the nitric acid waste and created enough carbon
dioxide to overpressurize the container. Two other waste containers in the hood were
severly damaged and several others were cracked or leaking.
Fortunately, the laboratory worker was not injured.
Chemical containers should be triple rinsed and dry before being used for waste
Safety glasses should always be worn while in the laboratory, even while
performing non-laboratory work.
Incidents Involving Reactive Materials
Peroxide Detonation
A laboratory worker attempted to use some anhydrous ethyl ether in a rotary evaporator
extraction. The four-liter container of ether was nearly empty. While pouring the ether
into the apparatus (inside a fume hood), he noticed that the liquid was oily and had a
strange odor, so he decided not to use it. He poured the ether back into the can and went
The next morning, he noticed a white residue inside the rotary evaporator. He used a
metal spatula to scrape the residue from a glass joint, causing a detonation that shattered
the glassware. The flying glass caused severe lacerations to the worker’s hands, face, ear
and scalp. Fortunately, he was wearing safety glasses that protected his eyes from injury.
Shards of glass were embedded in the lenses of the safety glasses. The sash of the hood
was cracked and the light fixture inside the hood shattered.
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The can of ethyl ether was purchased 30 months before the incident and was likely
opened about six months later. The container label clearly warned about the formation of
peroxide in storage, despite the presence of a stabilizer.
Using appropriate personal protective equipment helped to avoid a potentially
serious eye injury.
Discard peroxide forming chemicals six months after opening or one year after
purchase. Unless you plan to use the entire contents within this time period, large
containers such as the one involved in this incident should not be ordered.
Most hood sashes are not explosion-proof. Consider the need for shielding of
reactions that may result in exploding materials.
Do not use metal spatulas with peroxide forming compounds, since contamination
with metals can lead to explosive decomposition. Ceramic, Teflon or wooden
spatulas are recommended.
See Safe Work Practices - Peroxide Forming Compounds and Reactives for more
Lithium Aluminum Hydride Fire
A laboratory worker was attempting to distill tetrahydrofuran (THF) using lithium
aluminum hydride (LAH). THF is a highly flammable liquid that can cause severe eye
irritation and central nervous system depression. LAH is a water-reactive, flammable
The laboratory worker was slowly pouring approximately 1 gram of LAH from a plastic
bag into a flask containing 500 ml of THF inside a fume hood. A small amount of LAH
leaked from a small hole in the bag, onto the surface of the hood and burst into flames,
startling the worker and causing him to drop the remainder of the bag (8-10 grams of
LAH) onto the fire. Concerned about the flask and bottle of THF inside the hood, the
worker immediately removed his lab coat and placed it onto the fire in an attempt to
smother it.
Since the appropriate extinguishing agent was not available, .the worker pulled the
flaming lab coat and LAH out of the hood onto the floor. Once the LAH fire had burned
itself out, the worker used a dry chemical extinguisher to put out the coat fire.
Since the incident, Met-L-X extinguishers were mounted inside the door of the
laboratory. The laboratory worker keeps a supply of sand (in a plastic milk jug with the
top cut off) on the floor at the side of the hood where this work is done.
Know the hazards of the materials, including appropriate extinguishing agents,
before using chemicals.
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Carbon dioxide reacts with LAH explosively; thus, a carbon dioxide extinguisher
could have made the situation worse. A Met-L-X fire extinguisher (for flammable
solids) or dry sand should have been immediately available.
Do not pour solids such as LAH directly from the container into another chemical
or reaction vessel. Measure out what is needed, then pour it.
Potassium Metal Released from Pressurized Container
A laboratory worker received burns to one hand when small pieces of potassium metal
shot out of an alkali jet apparatus when the laboratory worker opened it for cleaning. The
accident occurred because the worker accidentally opened the apparatus while the system
was still under pressure. The burn was exacerbated by the fact that the worker rinsed the
hand with a small amount of mineral oil rather than with copious amounts of water.
To avoid a future occurrence, the worker installed a venting valve with a filter to allow
venting prior to opening the device. In addition, plexiglas shielding is placed around the
apparatus and the workers wear gloves, safety glasses and a face shield when opening the
The proper first aid for skin contact with potassium is to brush away visible
metals and flush with copious amounts of water for at least 15 minutes.
Ensure reaction vessels are at atmospheric pressure before opening them.
Wear gloves, safety glasses and a face shield when working with pressurized
equipment and hazardous materials.
Electrical Shock
Electrical Shock from Laser Power Supply
A laboratory worker noticed condensation on the high voltage power supply for a high
powered laser. With the power still on, he wiped the moisture with a tissue, making
contact with the exposed anode terminal at approximately 17,000 volts DC to ground.
He received a severe electrical shock and second degree burns to his right thumb and
abdomen. Witnesses stated that they heard a loud "snap" and then heard the laboratory
worker scream and stagger out to the hallway. He was immediately met by a secretary,
and told her "I got a shock" as he collapsed into her arms and onto the floor. He had no
pulse and was not breathing. Public Safety officers were nearby and immediately started
CPR. The ambulance crew arrived and was able to restore his heartbeat using a
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Fortunately, the laboratory worker lived to tell his story. He said that he knew that the
power was on but was not aware that contact was possible at the high voltage terminals.
The interlocks had been defeated and guards removed with no alternate guarding or
precautions taken.
Understand the operating characteristics of equipment before use.
Do not defeat machine safety interlocks.
Do not work around energized exposed conductors.
See Safe Work Practices - Electrical Safety for more information.
For information about laser safety, see the Laser Safety Training Guide.
Electrical Shock from Electrophoresis Unit
A laboratory worker received a potentially fatal electrical shock when he accidentally
touched a high voltage electrical connector on an electrophoresis device. The contact
points were on the right elbow and right knee. Had one of the contacts been on the
opposite side of the body, the shock could have been fatal.
The primary cause of this incident was the existence of an exposed high voltage
conductor in the form of a stackable banana plug at the device. When connected to the
male plug on the device, the male connector plug was left exposed with no insulation or
The accident could have been avoided by eliminating all exposed conductors in
connector cords and electrophoresis devices by either
 fitting each electrophoresis with its own set of permanently attached
connector cords to eliminate jacks and plugs entirely at this point; or
 eliminating cords with stackable plugs on both ends by replacing the
stackable plugs on one end with a female only jack (all electrophoresis
devices should be fitted with male only plugs).
For more information, see Electrical Safety and Laboratory Equipment
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This chapter summarizes the waste management practices recommended
in this handbook and selects the options that are especially
suitable for use by establishments that apply minimal programs or in
emergency situations. Typically, these situations include smaller rural
health-care establishments or field hospitals, e.g. in refugee camps. The
selected practices should ensure that health and safety requirements are
met and an acceptable level of hazard protection is achieved. However,
the recommendations should not be viewed as a substitute for the longer term
aim of establishing the more rigorous managerial procedures described
elsewhere in this handbook.
Implementation of the recommendations should be incremental, i.e.
achieved through gradual improvements, but it is of paramount importance
that municipal authorities and managers of health-care establishments
are made fully aware of the need for proper waste management
The first step would be the introduction of waste segregation: too often,
health-care establishments treat hazardous health-care waste in the
same manner as general waste. Separation of sharps may be a good
starting point. Specific methods for the disposal of hazardous health-care
wastes can then be introduced, followed by efforts to encourage waste
minimization and the safe reuse of materials wherever possible.
16.1 Basic principles
The total absence of management measures to prevent exposure to hazardous
health-care waste results in the maximum health risk to the
general public, patients, health-care personnel, and waste workers. It is
therefore emphasized that even very limited waste management measures
can dramatically reduce this risk.
Effective confinement of waste and safe handling measures provide significant
health protection. For example, burning hazardous health-care
waste in open trenches or small furnaces is better than uncontrolled
dumping; reducing the amount of hazardous waste by segregation is
better than accumulating large quantities; good stock management of
chemicals and pharmaceuticals not only reduces waste quantities but
also saves purchase costs; proper identification of waste packages warns
health-care personnel and waste handlers about their contents. All
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these measures to reduce risk are relatively simple and cheap and
should be considered by any health-care establishment. The principle of
“doing something is better than doing nothing” is important and underlies
any effort to initiate a system for the management of health-care
The basic elements of minimal programs of health-care waste management
are represented schematically in Fig. 16.1. At the local level, the
following basic actions should be taken:
• assessment (quantitative and qualitative) of waste production;
• evaluation of local treatment and disposal options;
• segregation of health-care waste from general (or municipal) waste;
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• establishment of internal rules for waste handling (storage, color
coding, collection frequency, etc.);
• assignment of responsibilities within the health-care establishment;
• choice of suitable—or better—treatment and disposal options.
16.2 Health-care waste segregation
16.2.1 The waste categories
Three categories of health-care waste are recognized:
• General (non-risk) waste, including uncontaminated waste similar to
domestic waste; may represent about 80% of the total waste production
from health-care establishments.
• Hazardous health-care waste.
• Highly hazardous health-care waste.
Fig. 16.1 Basic steps in health-care waste management in minimal programs
Minimal programs for health-care waste management
Hazardous health-care waste includes:
• “Usual” infectious waste, excluding sharps but including anatomical
or pathological waste, and waste contaminated with human blood or
other body fluids, excreta, and vomit. This category typically makes up
about 75% of the hazardous health-care waste, or around 15% of the
total waste, produced by health-care establishments.
• Chemical and pharmaceutical residues, e.g. cans, bottles, or boxes
containing such residues, and small quantities of outdated products.
• Non-recyclable and discarded pressurized containers, which are hazardous
only if burned as they may explode. Many undamaged containers
may be refilled.
Highly hazardous health-care waste, which should be given special attention,
• Sharps, especially hypodermic needles.
• Highly infectious non-sharp waste, including microbial cultures, carcasses
of inoculated laboratory animals, highly infectious physiological
fluids, pathological and anatomical waste.
• Stools from cholera patients or body fluids of patients with other
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highly infectious diseases.
• Bulk quantities of outdated hazardous chemicals, such as strong disinfectants,
or significant quantities of waste containing mercury.
• Genotoxic waste, e.g. radioactive or cytotoxic waste, typically used in
cancer chemotherapy but not in district hospitals. If minimal waste
management programs are being applied, genotoxic substances
should not be used in general hospitals, but may be used in the
oncological departments of university hospitals.
16.2.2 Segregation and packaging
Careful segregation and separate collection of hospital waste may be
somewhat onerous for hospital personnel but it is the key to safe, sound
management of health-care waste. Segregation can substantially reduce
the quantity of health-care waste that requires specialized treatment. To
make separate collection possible, hospital personnel at all levels, especially
nurses, support staff, and cleaners, should be trained to sort the
waste they produce.
In any area that produces hazardous waste—hospital wards, treatment
rooms, operating theatres, laboratories, etc.—three bins plus a separate
sharps container will be needed. Recommendations for the segregation of
waste are given in Table 16.1. The following important points should be
• If hazardous and highly hazardous wastes are to be disposed of in the
same way, they should not be collected separately.
• In a health-care establishment using genotoxic products, the safety
procedures applicable to radioactive or genotoxic products should be
• If sharps are to be encapsulated, it is convenient to collect them
directly in the metallic drums or barrels used for encapsulation, which
limits the hazards associated with handling.
• For hazardous waste and highly hazardous waste the use of double
packaging, e.g. a plastic bag inside a holder or container is recommended
for ease of cleaning.
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• Stools of cholera patients should be collected in buckets because of the
need for disinfect ion. Discharge to sewers or to the environment may
contribute to the spread of the disease.
Selection of appropriate packaging is difficult in establishments that
cannot afford disposable plastic bags or containers. In such circumstances,
hazardous waste may also be collected in paper bags, inside a
container that will not be removed. Plastic or metal containers for hazardous
waste should be disinfected, for example with sodium hypochlorite
(bleach), before reuse. The bags should be sealed or containers firmly
closed before they are filled to three-quarters of their capacity. The
equipment should be simple, robust and locally available.
16.2.3 Safe handling and storage
Hospital cleaning personnel should be informed about the potential risks
posed by waste handling. They should be trained in safe handling procedures
and should wear protective aprons and gloves.
The waste should be collected daily. General waste may be stored in
convenient places that facilitate collection by the municipal service, but
hazardous health-care waste should be stored in a closed room. Waste
should not be stored close to patients or where food is prepared. Infectious
waste should be disposed of within the following periods:
temperate climate: maximum 72 hours in winter
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maximum 48 hours in summer
warm climate: maximum 48 hours during the cool season
maximum 24 hours during the hot season
Before containers of hazardous health-care waste are loaded on to a truck
for transport off site, they should be sealed. Waste bags and containers
should also be labeled with the address of the producer and the waste
category. For safety reasons, however, it is strongly recommended that
establishments applying minimal waste management programs in
areas without adequate treatment facilities should dispose of hazardous
health-care waste within their own premises.
16.3 Minimization and safe recycling of health-care waste
16.3.1 Chemicals and pharmaceuticals
Careful and comprehensive management of stores will substantially reduce
the quantities of chemical and/or pharmaceutical waste produced by
health-care establishments. Ideally, the waste in these categories should
be limited to residues of chemical or pharmaceutical products in their
original packaging (bottles, boxes, cans, etc.). Waste minimization will
also give rise to financial savings.
Proper management of chemical or pharmaceutical stores will be supervised
by the Chief Pharmacist of the health-care establishment and
should include the practices listed in Box 16.1.
16.3.2 Pressurized containers
Aerosol cans are not generally recyclable and may be disposed of to
landfills together with general waste. Many undamaged pressurized gas
containers, however, may be easily recycled, and should be returned to
their original supplier for refilling. Pressurized containers must never
be incinerated as they may explode, causing injury to workers and/or
damage to equipment.
16.3.3 Mercury
Metallic mercury is a valuable product. In case of a spill, e.g. from a
broken thermometer, all droplets of mercury should be recovered with a
spoon for later sale or reuse.
16.3.4 Recyclable sharps
Hospitals with very limited resources should use recyclable sharps, such
as glass syringes with needles, and scalpels. Only items that are designed
for reuse, i.e. that withstand the sterilization process, should be recycled
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in this way. Before reuse, scalpels, syringes, needles, and other sharps
must be thoroughly cleaned and sterilized; disinfection alone is inadequate.
Any failure in the sterilization process may result in the transmission
of severe infections. Sterilization may be by chemical means, by
flame exposure, or by autoclaving. Smaller district hospitals that lack
autoclave facilities may consider sending items to the closest general
hospital for sterilization.
16.4 Treatment and disposal of hazardous health-care waste
For health-care establishments with few resources and applying minimal
waste management programs, affordable treatment and disposal
methods for hazardous and highly hazardous waste may be classified into
three categories:
• thermal processes
• chemical processes
• containment processes.
16.4.1 Thermal processes
Static-grate single-chamber incineration
Waste may be burned in a simple furnace, with a static grate and natural
air flow. De-ashing, loading, and unloading operations are carried out
manually. The low heating value of properly segregated health-care
waste is high enough for combustion, but addition of a small quantity of
kerosene may be needed to start the fire and blowing of air may also help
in establishing optimal combustion. The burning efficiency may reach
90–95%, i.e. 5–10% of the material may remain unburnt in the ashes and
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slags. The operating temperature will be around 300C, which will kill
most microorganisms but will be insufficient to destroy thermally resistant
chemicals or pharmaceuticals.
• Good disinfection efficiency.
• Drastic reduction of waste; the weight and volume of residual ashes
and slags are about 20% those of the original waste. The residues may
then be landfilled.
• No requirement for highly qualified operators.
• Relatively low investment and operation costs.
• Generation of significant emissions containing atmospheric pollutants,
including flue gases and fly ash; may produce odours (which can be
limited by not incinerating halogenated plastics).
• Periodic removal of slag and soot necessary.
• Inefficiency in destruction of thermally resistant chemicals and drugs
(e.g. cytotoxics).
Drum or brick incinerators
Where a single-chamber incinerator is not affordable or available, simple
confined burning may be applied. A steel drum or walls of bricks or
concrete can be erected over a screen or fine grate and covered with a
second screen to prevent dispersion of ashes or light material. The waste
is placed inside and burned with the help of manual ventilation and
addition of kerosene if necessary. Constant supervision is essential to
prevent any spread of the fire to the surrounding area. The combustion
efficiency may reach 80–90% and kill 99% of microorganisms. The temperature
of the fire will not exceed 200 C, and this process should be used
only in emergency situations or when other treatment methods cannot be
• Drastic reduction of weight and volume of the waste.
• Very low investment and operating costs.
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• Relatively poor destruction efficiency.
• No destruction of many chemicals and pharmaceuticals.
• Massive emission of black smoke, particulates, and toxic flue
Open-air burning
Open-air burning of infectious waste (excluding pathological waste)
should be carried out only as a last resort, in rural dispensaries, isolated
health posts, or emergency situations. If possible, the burning should
take place in the pit of final disposal (i.e. where the residues will be
buried), and the process should be supervised by the person responsible
for waste management in the health-care facility. It should be performed
downwind of, and as far as possible from, the facility or other communities.
The area within which the burning is carried out should be fenced to
prevent unauthorized persons and animals from entering.
Confined burning, e.g. in a drum incinerator, should always be preferred,
as the risk to personnel of contact with the waste or with partly burned
residues is lower. The advantages and drawbacks of open-air burning are
the same as for drum or brick incinerators, but there is the additional
disadvantage that burning may be incomplete and non-uniform.
16.4.2 Wet thermal disinfection
Wet thermal disinfection is based on exposure of shredded infectious
waste to high-temperature, high-pressure steam. Shredded waste is introduced
into a reacting tank, vacuum conditions are established, and
steam is introduced. Precise operating procedures have to be followed by
qualified technicians for efficient disinfection. Wet thermal disinfection
should be considered only by health-care establishments with sufficient
technical and financial resources and where incineration in single-chamber
or drum/brick incinerators is unacceptable, for example because of air
pollution problems.
• Environmentally sound.
• Reduction in waste volume.
• Relatively low investment and operation costs.
• Shredders subject to breakdown and poor functioning (and are thus
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the weak point of the process).
• Qualified operators essential.
• Inadequate for anatomical, pharmaceutical, and chemical waste, and
waste that is not easily penetrated by steam.
Safe management of wastes from health-care activities
Autoclaving is an efficient wet thermal disinfection process. Typically,
autoclaves are used in hospitals for the sterilization of recyclable items,
and these units allow for the treatment of only limited quantities of
waste. They are therefore generally used only for highly infectious waste,
such as microbial cultures and sharps. Even a general hospital with very
limited resources should be equipped with an autoclave, but a district
hospital may well not have one. The advantages and drawbacks of the
autoclave are similar to those of wet thermal processes.
• Efficient.
• Environmentally sound.
• Relatively low investment and operation costs.
• Qualified operators essential.
• Inadequate for anatomical, pharmaceutical, and chemical waste, and
waste that is not easily penetrated by steam.
• The hospital autoclave used for sterilization has capacity for treatment
of only limited quantity of waste.
16.4.3 Chemical disinfection
Chemical disinfection is an efficient process, but costly if the prices of
disinfectants are high. For safe operation it requires trained technicians
provided with adequate protective equipment and is therefore not recommended
for treating all infectious health-care waste. However, the process
can be useful in specific cases, such as disinfection of recyclable
sharps or disinfection of stools from cholera patients.
Chemical sterilization of recyclable sharps
Chemical sterilization of scalpels, syringes with needles, and other recyclable
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sharps may be considered as an alternative or complementary
method to thermal sterilization. After thorough cleaning and drying, the
sharps are placed in a tank and exposed to a strong disinfecting gas or
liquid, such as ethylene oxide, formaldehyde, or glutaraldehyde.
• Highly efficient (may be more efficient than thermal sterilization).
• Trained operators essential.
• Costly if the chemical disinfectants are expensive.
• Uses hazardous substances that necessitate safety measures.
Chemical disinfection of stools from cholera patients
Vibrio cholerae, the causative agent of cholera, is not very resistant and
its elimination does not require the use of very strong chemical disinfectants.
Buckets containing stools of patients with acute diarrhoea may be
disinfected through addition of chlorine oxide powder or dehydrated lime
oxide (CaO). Other liquid or powder disinfectants may also be used. In
case of a cholera epidemic, hospital sewage must also be treated and
disinfected. Where there is sufficient space, sewage may be treated
through lagooning, followed by effluent disinfection with sodium hypochlorite.
In cholera epidemics in emergency situations these disinfection
Minimal programmes for health-care waste management
measures should also be applied in field hospitals to prevent the spread
of the disease.
• Efficient disinfection.
• No need for highly trained operators.
• Not significant compared with the benefits.
16.4.4 Containment processes
Landfilling in municipal disposal sites
Waste may be landfilled in municipal disposal sites if it cannot be treated
before disposal. However, health-care waste should not be deposited or
scattered on the surface of open dumps. If landfilling is planned, the
following minimal requirements should be met:
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• measures established by a municipal authority for the rational and
organized deposit of municipal wastes that could be used to dispose of
health-care wastes;
• if possible, engineering work instigated by the municipal authority to
prepare the disposal site to retain wastes more effectively;
• rapid burial of the health-care waste, so that human or animal contact
is as limited as possible.
In addition, it is recommended that health-care waste is deposited in one
of the following two ways:
• in a shallow hollow excavated in the mature municipal waste, in the
layer below the base of the working face, where it is immediately
covered by a 2-m layer of fresh municipal waste; scavenging in this
part of the site must be prevented.
• in a deeper pit (1–2 m) excavated in mature municipal waste (at least
3 months since being landfilled) which is then backfilled with the
mature waste that was dug out; again, scavenging in this part of the
site must be prevented.
Alternatively, a specially constructed small burial pit could be prepared
to receive health-care waste only. The pit can be 2m deep and filled to a
depth of 1 m. Each load of waste should be covered with a soil layer 10–
15cm deep. (Lime may be placed over the waste if coverage with soil is
not possible.) In case of a disease outbreak involving especially virulent
pathogens (such as the Ebola virus), both lime and soil cover may be
added. Access to this area should be restricted and closely supervised by
the responsible staff to prevent scavenging. An example of dedicated pit
design is shown in Fig. 8.12 (page 109).
Before health-care wastes are sent for land disposal, it is prudent to
inspect the proposed landfill site to ensure that there is satisfactory
control of waste deposition.
• Low costs.
• Relatively safe if access is restricted and the site is selected according
to the above conditions.
Safe management of wastes from health-care activities
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• Effective biodegradation of the biological components of health-care
waste if landfill operations are properly carried out.
• Access restrictions may not always be guaranteed.
• It may be difficult to assess whether the conditions for safe landfill are
being met.
Safe burying inside premises
In certain health-care establishments in remote locations, temporary
refugee camps, and areas experiencing exceptional hardship, safe burial
of wastes on hospital premises may be the only rational option available
at times. The design and operation of the burial pit is as described above
and illustrated in Fig. 8.12 (page 109). To limit risks to health and of
environmental pollution, some basic rules should be applied:
• Access to the disposal site should be restricted to authorized personnel
• The burial boundary should be lined with a material of low permeability
(e.g. clay), if available.
• Only hazardous health-care waste should be buried.
• Large quantities (over 1 kg) of chemical wastes should not be buried at
the same time; burial should be spread over several days.
• The burial site should be managed in the same way as a landfill, with
each layer of waste being covered with a layer of earth to prevent
development of odors and infestation by rodents and insects.
The safety of waste burial relies critically on operational practices. Safe
on-site burial is practicable for only relatively limited periods of time, e.g.
1–2 years, and for relatively small quantities of waste, say up to 5–10
tons in total. Where these limits are exceeded, a longer-term solution,
involving treatment of the waste or disposal at a municipal solid waste
landfill, will need to be found.
• Less hazardous than letting waste accumulate and remain accessible.
• Low costs.
• Risks of pollution in permeable soils if the waste becomes saturated
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with water.
• It may be difficult to prevent scavenging at all times.
Encapsulation is recommended as the easiest technology for the safe
disposal of sharps. Sharps are collected in puncture-proof and leak-proof
containers, such as high-density polyethylene boxes, metallic drums, or
barrels. When a container is three-quarters full, a material such as
cement mortar, bituminous sand, plastic foam, or clay is poured in until
the container is completely filled. After this material has dried, the
container is sealed and may be landfilled, stored, or buried inside the
hospital premises. It is also possible to encapsulate chemical or pharmaceutical
residues together with sharps.
Minimal programs for health-care waste management
• Simple and safe.
• Low costs.
• Also applicable to chemicals and pharmaceuticals.
• Not recommended for non-sharp infectious waste.
16.5 Management of hazardous health-care waste by
waste categories
16.5.1 Infectious waste and sharps
Most treatment methods outlined in section 16.4 above are suitable for
infectious waste and sharps, except that:
• in the wet thermal process, shredding of sharps is problematic;
• encapsulation is not suitable for infectious waste.
Incineration in single-chamber incinerators should be the method of
choice in establishments that apply minimal waste management
programs. Highly infectious waste, such as cultures and stocks of
infectious agents from laboratory work, should be sterilized by wet thermal
treatment (e.g. autoclaving) at the earliest stage, i.e. inside the
health-care establishment, and soon after production, if possible. For
other infectious health-care waste, disinfection to reduce microbial concentration
is sufficient.
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Sharps should also be incinerated whenever possible and can be incinerated
together with other infectious waste. Encapsulation is also suitable
for disposing of sharps.
Blood should be disinfected before discharge to the sewer (unless there is
an adequate wastewater treatment plant) or may be incinerated.
After incineration or other disinfection process, residues may be
16.5.2 Pharmaceutical waste
Sound management of pharmaceutical products, with a view to waste
minimization (see section 16.2), is of prime importance. Small quantities
of chemical or pharmaceutical waste can be disposed of easily and relatively
cheaply, but large amounts may require special, more costly
treatment, such as high-temperature incineration. Comprehensive management
of pharmaceutical stores should be supervised by the Chief
Pharmacist of the health-care establishment.
Small quantities of pharmaceutical waste are usually collected in yellow
containers together with infectious waste and therefore follow the same
disposal pathway, being either incinerated or safely buried. It should be
noted, however, that temperatures reached in a single-chamber furnace
may be insufficient to disintegrate thermally resistant pharmaceuticals.
Small quantities of pharmaceutical waste, such as outdated drugs (except
cytotoxics and antibiotics), may also be discharged to the sewer but
should not be discharged into natural waters (rivers, lakes, etc.).
Safe management of wastes from health-care activities
Significant quantities of pharmaceutical waste may be disposed of by the
following methods:
• Incineration (if an incinerator able to reach a combustion temperature
of 800 C is available); the incineration residues may be landfilled.
• Discharge to the sewer. Water-soluble, relatively mild pharmaceutical
mixtures, such as vitamin solutions, cough syrups, intravenous solutions,
eye drops, etc., may be diluted with large amounts of water and
then discharged to sewers (where sewerage systems exist). This process
should not be used for antibiotics.
• Encapsulation. When incineration is not feasible and water dispersion
is not recommended, pharmaceutical waste should be encapsulated.
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• Return to the original supplier if possible.
Note: Cytotoxic drug residues and other cytotoxic waste should never be
mixed with other pharmaceutical waste, but should be processed
separately according to the procedure described in this handbook
(section 9.3).
16.5.3 Chemical waste
As for pharmaceutical waste, improved management of chemical waste
starts with waste minimization efforts. The proper management of
chemical stores will be supervised by the Chief Pharmacist of the healthcare
establishment (see section 16.3).
The hospital’s Infection Control Officer, Chief Hygienist, or Chief Pharmacist
should be designated to supervise the use of chemicals throughout
the health-care establishment. The main users of chemical disinfectants,
which are among the most hazardous chemicals used in the establishment,
are likely to be the Infection Control Officer/Chief Hygienist and
his or her staff.
Small quantities of chemical waste will include residues of chemicals in
their packaging, outdated or decomposed chemicals, or chemicals that
are no longer required. These are generally collected in yellow containers,
together with infectious waste, and follow the same disposal pathway
(either incineration or safe burying).
Large quantities of chemical waste should not be collected in yellow
plastic bags or containers. There is no safe and cheap method for their
disposal; the treatment options are the following:
• Incineration under subcontract by a public or private agency equipped
for the safe disposal of hazardous chemical waste. The thermal reactivity
of the waste should be checked; certain solvents will burn and
can therefore be incinerated in simple incineration units, although it
must be remembered that those containing halogens could cause air
• Return to the original supplier (if the supplier has facilities for safe
disposal). In this case, appropriate provisions should be included in the
original purchase contract for chemicals.
• Exportation to a country with the expertise and facilities to dispose
safely of hazardous chemical waste. Shipment of chemical waste
should comply with international agreements, such as the Basel Convention
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and the United Nations Recommendations on the transport of
dangerous goods.
Minimal programs for health-care waste management
All three options are costly and may be impracticable, which makes it
particularly crucial that chemical waste is minimized. The following
recommendations should also be observed:
• Hazardous chemical wastes of different nature should never be mixed.
• Hazardous chemical waste should not be disposed of in sewer systems.
• Large amounts of chemical waste should not be buried as they may
contaminate groundwater.
• Large amounts of chemical disinfectants should not be encapsulated
as they are corrosive and sometimes flammable.
16.5.4 Cytotoxic waste
Cytotoxic drugs are highly hazardous to the health of the individual
and to the environment. Recommendations on cytotoxic safety may be
found in section 12.3. Disposal options, described in section 9.3, are the
• Return to the original supplier.
• Incineration at high temperatures, e.g. in rotary kilns or high performance
double-chamber pyrolytic incinerators (if available).
• Chemical degradation.
The following recommendations should also be observed:
• Residues from cytotoxic drugs or other cytotoxic waste should never be
mixed with other pharmaceutical waste.
• Cytotoxic waste should never be discharged into natural water bodies
or landfilled.
In countries where the above disposal procedures are not feasible, use of
cytotoxic and radioactive products should be restricted to university
research and teaching hospitals.
16.5.5 Radioactive waste
For safety reasons, medical use of radioactive isotopes should be restricted
to university hospitals, and any hospital that uses radioactive
products should appoint a qualified Radiation Officer. The rules for safe
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management of radioactive waste outlined in section 9.7 of this handbook
should be enforced.
16.5.6 Pressurized containers
Undamaged pressurized containers should be returned to the supplier
for refilling, and adequate provision for this should be included in the
original purchase contracts. If return is not possible, containers may be
buried safely. Any residual pressure should be released before disposal.
Aerosol containers cannot usually be refilled and should be buried. Pressurized
containers should never be burned or incinerated because of the
severe risk of explosion.
16.5.7 Used batteries and thermometers
Batteries, thermometers, and various items of measuring equipment
may have a high metal content, including toxic heavy metals such as
mercury or cadmium. Disposal options are as follows:
Safe management of wastes from health-care activities
• Recycling by specialized cottage industries. This is the best disposal
solution when available.
• Exportation to a country with the expertise and facilities to dispose
safely of hazardous chemical waste. Conditions of shipment should
comply with the Basel Convention.
• Encapsulation. If neither of the two options above is feasible, encapsulated
waste may be disposed of in an impermeable landfill (if available)
or other landfill.
This type of waste should not be incinerated because of the toxic metallic
vapors emitted, nor should it be buried without encapsulation as this
may cause pollution of groundwater.
However, if the quantities of wastes with high heavy-metal content are
minimal (similar to the quantities in municipal waste) and there are no
opportunities for reuse of heavy metals within the country, they may also
join the municipal waste stream.
16.6 Workers’ training and safety at work
In health-care establishments and regions that operate minimal management
programs, the health and safety practices described in Chapter
12 and the training outlined in Chapter 15 should be implemented.
This is of particular importance, since minimal programs of waste
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management are likely to result in greater risks of exposure for workers
than the more comprehensive managerial methods described in this
For personnel who handle wastes, including hospital cleaners and technicians,
training in safety measures should cover the following issues:
• packing, handling, and storing of hazardous health-care waste;
• the need to wear protective gloves and aprons when handling waste
• operation of on-site treatment and disposal methods, such as single chamber
furnace operations, encapsulation, and safe burying.
Technicians in charge of chemical disinfection should be trained to implement
appropriate safety precautions and emergency measures and be
informed about chemical hazards. Nurses and cleaning personnel should
be made aware of the occupational risks linked to handling of sharps.
References and suggested further reading
Christen J (1996). Dar es Salaam Urban Health Project. Health care waste
management in district facilities: situational analysis and system development.
St Gallen, Switzerland, Swiss Centre for Development Cooperation in
Technology and Management (SKAT).
WHO. Guidelines for drug disposal after emergencies. Geneva, World Health
Organization (unpublished document, in preparation; will be available from
Department of Essential Drugs and other Medicines, World Health Organization
1211 Geneva 27, Switzerland).
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