Читать книгу Diagnostic Medical Parasitology - Lynne Shore Garcia - Страница 20
ОглавлениеSome of the following information has been adapted from Clinical Microbiology Procedures Handbook, published by the American Society for Microbiology (1). Additional information on the following topics can also be found in that publication. A number of excellent documents are also available for information and guidance (2–4).
Good, clean microscopes and light sources are mandatory for the examination of specimens for parasites (Fig. 11.1). Organism identification depends on morphologic differences, most of which must be seen under stereoscopic microscopes (magnification, ×50) or regular microscopes at low (×100), high dry (×400), and oil immersion (×1,000) magnifications. The use of a 50× or 60× oil immersion objective for scanning can be very helpful, particularly if the 50× oil and 100× oil immersion objectives are placed side by side. This arrangement on the microscope can help avoid accidentally getting oil on the 40× high dry objective. Calibration of the microscope is discussed later in this chapter. Although 5× oculars are acceptable, most laboratories select 10× oculars, preferably with a binocular, tilting head.
Types
Stereoscopic Microscope. A stereoscopic microscope is recommended for larger specimens (arthropods, tapeworm proglottids, and various artifacts). The total magnification usually varies from approximately ×10 to ×45, either with a zoom capacity or with fixed objectives (0.66×, 1.3×, and 3×) that can be used with 5× or 10× oculars. Depending on the density of the specimen or object being examined, you must be able to direct the light source either from under the stage or onto the top of the stage (may require a separate simple laboratory light directed at the top of the stage). Current uses include the examination of larger specimens (arthropods, artifact removed from clinical specimens, whole mounts, etc.).
Figure 11.1 Microscopes routinely used in clinical laboratories. (Top row) Regular compound microscopes. (Second row, left) Microscope with tube for photography; (right) stereomicroscope. (Bottom) Fluorescence microscope. doi:10.1128/9781555819002.ch11.f1
Regular Light Microscope. The light microscope should be equipped with the following:
1. Head. A binocular head is recommended and should be equipped with a diopter adjustment to compensate for focus variation in the eyes. A tilt head is highly recommended to accommodate various users.
2. Oculars. 10× oculars are required; 5× oculars can be helpful but are optional.
3. Objectives. 10× (low power), 40× (high power), 100× (oil immersion). Some laboratories are currently using 50× or 60× oil immersion lenses for screening permanent stained smears. Examination with a combination of the 50× oil or 60× oil and the 100× oil immersion lenses allows screening to proceed more quickly and eliminates the problem of accidentally getting oil on the high dry objective lens when switching back and forth between the 40× (high dry) and 100× (oil immersion) objectives.
4. Stage. A mechanical stage for both vertical and horizontal movement is required. Graduated stages are mandatory for recording the exact location of an organism on a permanent stained smear and recommended for any facility performing diagnostic parasitology procedures. This capability is essential for consultation and teaching responsibilities. However, remember that calibration numbers for the exact location of an organism may not be the same for different microscopes and different stages.
5. Condenser. A bright-field condenser equipped with an iris diaphragm is required; however, an adjustable condenser is not required with the newer microscopes. The condenser numerical aperture should be equal to or greater than the highest objective numerical aperture.
6. Filters. Both clear blue glass and white ground-glass filters are recommended.
7. Light source. The light source, along with an adjustable voltage regulator, is usually contained in the microscope base. This light source should be aligned as specified by the manufacturer. If the light source is external, the microscope must be equipped with an adjustable mirror and an adjustable condenser containing an iris diaphragm. The light source should be a 75- to 100-W bulb.
8. Current Uses. Routine microscopy in the microbiology laboratory, and parasitology in particular, includes the examination of direct wet mounts, concentration sediment and surface flotation layers, stained permanent smears, blood specimens, aspirates and small samples or smears/scrapings from any body site, and cytologic and histologic preparations (stained or unstained).
Fluorescence Microscope. The fluorescence microscope should be equipped with the following:
1. Head. A binocular head is recommended and should be equipped with a diopter adjustment to compensate for focus variation in the eyes. A tilt head is highly recommended to accommodate various users.
2. Oculars. 10× oculars are required; 5× oculars can be helpful but are optional.
3. Objectives. 10× (low power), 40× (high power), 100× (oil immersion).
4. Stage. A mechanical stage for both vertical and horizontal movement is required. Graduated stages are mandatory for recording the exact location of an organism on a permanent stained smear and recommended for any facility performing diagnostic parasitology procedures. This capability is essential for consultation and teaching responsibilities. However, remember that calibration numbers for the exact location of an organism may not be the same for different microscopes and different stages.
5. Light source. Mercury-free microscopy (MFM) provides a new approach that encourages the selection of modern mercury-free light sources to replace more traditional mercury-based arc lamps. Microscope performance is enhanced with new solid-state technologies; they offer a more stable light intensity output and have a more uniform light output across the visible spectrum. Solid-state sources eliminate mercury, eliminate the cost of consumable bulbs (lifetime ~200 hours), use less energy, reduce the instrument downtime when bulbs fail, and reduce the staff time required to replace and align bulbs. With lifetimes of approximately tens of thousands of hours, solid-state replacements can pay for themselves over their lifetime, are sustainable, and comply with institutional and government body mandates to reduce energy consumption, carbon footprints, and hazardous waste. Solid-state light-emitting diode (LED)-based light sources have been used in the laboratory for some time. However, advancements in solid-state "light engine" technology now provide laboratories with mercury alternatives that are straightforward to operate, environmentally friendly, affordable to buy, and better suited for fluorescence microscopy.
6. Fluorescence filter set. Usually housed in cube-shaped optical blocks, fluorescence filter sets include excitation and barrier (emission) filters along with a dichromatic mirror. The filter block is positioned in the vertical illuminator, above the objective, so that incident illumination can be directed through the excitation filter and reflected from the surface of the dichromatic mirror onto the specimen. The fluorescence emitted by the specimen is gathered by the objective and transmitted through the dichromatic mirror to the eyepieces or detector system.
7. Quenching. Quenching refers to any process which decreases the fluorescence intensity of a given substance. Although it is important to store slides in the dark (Giardia/Cryptosporidium FA reagent slides) during incubation and prior to reading, personal use indicates quenching is very minimal; slides may retain fluorescence for several days to weeks.
8. Current uses. There are a number of diagnostic procedures available that rely on fluorescence. Examples include fecal immunoassays for antigen detection (FA) for Cryptosporidium, Giardia, and microsporidia (not FDA approved); demonstration of autofluorescence with Cyclospora; various stains such as calcofluor white, auramine, rhodamine; various serologic methods for antibody detection; and research products for labeling/conjugation/linking various components.
Maintenance (5)
1. Use a camel hair brush to remove dust from all optical surfaces; remove oil and finger marks immediately from the lenses with several thicknesses of lens tissue. Single-thickness lens tissue may permit corrosive acids from the fingers to damage the lens. Do not use any type of tissue other than lens tissue; otherwise you may scratch the lens. Use very little pressure, to prevent removal of the coatings on external surfaces of the lenses.
2. Use water-based cleaning solutions for normal cleaning. If you have to use organic solvents, use them in very small amounts and only if absolutely necessary to remove oil from the lens. Since microscope manufacturers do not agree on solvents to be used, each company’s recommendations should be consulted. One recommended solvent is 1,1,1-trichloroethane; it is good for removing immersion oil and mounting media and does not soften the lens sealers and cements. Xylene, any alcohols, acetone, or any other ketones should never be used as cleaning fluids.
3. After the lamp has been installed into the lamp holder, clean it with lens tissue moistened in 70% isopropyl or ethyl alcohol (to remove oil from fingers). Make sure that the lamp is cool and the switch is in the off position when replacing or removing the lamp.
4. Clean the stage with a small amount of disinfectant (70% isopropyl or ethyl alcohol) when it becomes contaminated.
5. Using petroleum jelly or light grease, clean and lubricate the substage condenser slide as needed.
6. Cover the microscope when not in use. In extremely humid climates (a relative humidity of more than 50%), good ventilation is necessary to prevent fungal growth on the optical elements.
7. At least annually, schedule a complete general cleaning and readjustment to be performed by a factory-trained and authorized individual. If microscopes are in continual use, maintenance should be performed twice a year. Record all preventive maintenance and repair data (date, microscope identification number, names of company and representative, maintenance and/or repairs, part replacement, recommendations for next evaluation, estimated cost if you have such information). This information should be cumulative so that a review for each piece of equipment can be scanned quickly for continuing problems, justification for replacement requests, etc. Depending on the physical site and use of the microscope, laboratories may use different maintenance schedules.
Calibration
The identification of protozoa and other parasites depends on several factors, one of which is size. Any laboratory doing diagnostic work in parasitology should have a calibrated microscope available for precise measurements (6). Measurements are made with a micrometer disk that is placed in the ocular of the microscope; the disk is usually calibrated as a line divided into 50 units (Fig. 11.2). Depending on the objective magnification used, the divisions in the disk represent different measurements. The ocular disk division must be compared with a known calibrated scale, usually a stage micrometer with a scale of 0.1- and 0.01-mm divisions. Although there is not universal agreement, it is probably appropriate to recalibrate the microscope once or more each year. This recommendation should be followed if the microscope has received heavy use or has been bumped or moved multiple times. Often, the measurement of red blood cells (approximately 7.5 µm) is used to check the calibrations of the three magnifications (×100, ×400, and ×1,000). Latex or polystyrene beads of a standardized diameter (Sigma, J. T. Baker, etc.) can be used to check the calculations and measurements. Beads of 10 and 90 µm in diameter are recommended.
Figure 11.2 Ocular micrometer, top scale; stage micrometer, bottom scale. (Illustration by Nobuko Kitamura; modified from references 1, 6, and 55.) doi:10.1128/9781555819002.ch11.f2
Supplies
1. Ocular micrometer disk (line divided into 50 units) (any laboratory supply distributor [Fisher, Baxter, Scientific Products, VWR, etc.])
2. Stage micrometer with a scale of 0.1- and 0.01-mm divisions (Fisher, Baxter, Scientific Products, VWR, etc.)
3. Immersion oil
4. Lens paper (do not use other types of tissues)
5. Binocular microscope with 10×, 40×, and 100× objectives. Other objective magnifications may also be used (50× oil or 60× oil immersion lenses).
6. Oculars of 10×. Some may prefer 5×; however, smaller magnification may make final identifications more difficult.
7. Single 10x ocular to be used to calibrate all laboratory microscopes (to be used when any organism is being measured – must be used on each calibrated microscope (oculars are NOT interchangeable).
Note All measurements should be documented in quality control (QC) records.
Procedure
1. Unscrew the eye lens of a 10× ocular, and place the micrometer disk (engraved side down) within the ocular. Use lens paper (several thicknesses) to handle the disk; keep all surfaces free of dust and lint.
2. Place the calibrated micrometer on the stage, and focus on the scale. You should be able to distinguish the difference between the 0.1- and 0.01-mm divisions. Make sure that you understand the divisions on the scale before proceeding.
3. Adjust the stage micrometer so that the “0” line on the ocular micrometer is exactly lined up on top of the 0 line on the stage micrometer.
4. After these two 0 lines are lined up, do not move the stage micrometer any farther. Look to the right of the 0 lines for another set of lines that is superimposed. The second set of lines should be as far to the right of the 0 lines as possible; however, the distance varies with the objectives being used (Fig. 11.2).
5. Count the number of ocular divisions between the 0 lines and the point where the second set of lines is superimposed. Then, on the stage micrometer, count the number of 0.1-mm divisions between the 0 lines and the second set of superimposed lines.
6. Calculate the portion of a millimeter that is measured by a single small ocular unit.
7. When the high dry and oil immersion objectives are used, the 0 line of the stage micrometer will increase in size whereas the ocular 0 line will remain the same size. The thin ocular 0 line should be lined up in the center or at one edge of the broad stage micrometer 0 line. Thus, when the second set of superimposed lines is found, the thin ocular line should be lined up in the center or at the corresponding edge of the broad stage micrometer line.
Examples:
Example: If a helminth egg measures 15 ocular units by 7 ocular units (high dry objective), using the factor of 2.0 µm for the 40× objective (example C above), the egg measures 30 by 14 µm and is probably Clonorchis sinensis.
Example: If a protozoan cyst measures 23 ocular units (oil immersion objective), using the factor of 0.8 µm for the 100× objective (example D above), the cyst measures 18.4 µm.
Results. For each objective magnification, a factor will be generated (1 ocular unit = certain number of micrometers). If standardized latex or polystyrene beads or a red blood cell is measured with various objectives, the size of the object measured should be the same (or very close), regardless of the objective magnification. The multiplication factor for each objective should be posted (either on the base of the microscope or on a nearby wall or bulletin board) for easy reference. Once the number of ocular lines per width and length of the organism is measured, then, depending on the objective magnification, the factor (1 ocular unit = certain number of micrometers) can be applied to the number of lines to obtain the width and length of the organism. Comparison of these measurements with reference measurements in various books and manuals should confirm the organism identification.
Procedure Notes for Microscope Calibration
1. The final multiplication factors will only be as good as your visual comparison of the ocular 0 and stage micrometer 0 lines.
2. As a rule of thumb, the high dry objective (40×) factor should be approximately 2.5 times more than the factor obtained from the oil immersion objective (100×). The low-power objective (10×) factor should be approximately 10 times that of the oil immersion objective (100×).
Limitations of Microscope Calibration
1. After each objective has been calibrated, the oculars containing the disk and/or the objectives cannot be interchanged with corresponding objectives or oculars on another microscope.
2. Each microscope used to measure organisms must be calibrated as a unit. The original oculars and objectives that were used to calibrate the microscope must also be used when an organism is measured.
3. The objective containing the ocular micrometer can be stored until needed. This single ocular can be inserted when measurements are taken. However, this particular ocular containing the ocular micrometer disk must have also been used as the ocular during microscope calibration.
A table or floor model centrifuge to accommodate 15-ml centrifuge tubes is recommended. It is also helpful to have a centrifuge that can hold 50-ml centrifuge tubes, particularly when some of the commercial concentration systems are used. Regardless of the model, a free-swinging or horizontal head is recommended. With this type of centrifuge, the sediment is deposited evenly on the bottom of the tube and the flat surface of the sediment allows removal of the supernatant fluid from the sediment, particularly when you cannot turn the tube upside down to pour out the supernatant fluid. Most laboratories are using carrier cups that have screw-cap closures; this feature, in addition to capped centrifuge tubes, will minimize any aerosol formation and/or distribution.
Maintenance (5)
1. Before each run, visually check the carrier cups, trunnions, and rotor for corrosion and cracks. If anything is found to be defective, replace it immediately or remove the equipment from service. Check for the presence and insertion of the proper cup cushions before each run.
2. At least quarterly, check the speed at all regularly used speeds with a stroboscopic light to verify the accuracy of a built-in tachometer or speed settings. Remember to record results. Some laboratories perform this function every 6 months or yearly.
3. Following a breakage or spill and at least monthly, disinfect the centrifuge bowl, buckets, trunnions, and rotor with 10% household bleach or phenolic solution. Following disinfection, rinse the parts with warm water and perform a final rinse with distilled water. Thoroughly dry the parts with a clean absorbent towel to prevent corrosion. At least quarterly, brush the inside of the cups with mild warm soapy water and use fine steel wool to remove deposits; the cups should then be rinsed in distilled water and thoroughly dried.
4. Follow manufacturer’s recommendations for preventive maintenance (lubrication).
5. Semiannually, check brushes and replace if worn to 1/4 in. (1 in. = 2.54 cm) of the spring. Also semiannually, check the autotransformer brush and replace if worn to 1/4 in. of the spring.
6. Record all information relating to preventive maintenance and repair (date, centrifuge identification number, names of company and representative, maintenance and/or repairs, part replacement, recommendations for next evaluation, estimated cost if you have such information). This information should be cumulative so that a review for each piece of equipment can be scanned quickly for continuing problems, justification for replacement requests, etc.
Chemical fume hoods should be used when there is risk of exposure to hazardous fumes or splashes while preparing or dispensing chemical solutions. Airflow is generally controlled by a movable sash and should be in the range of 80 to 120 ft/min (1 ft = 30.48 cm). Chemical fume hoods are certified annually. Although a fume hood is not required for diagnostic parasitology work, many facilities keep the staining setup and formalin (see below for a discussion of regulations regarding the use of formaldehyde) in a fume hood. Fume hoods may also be preferred for the elimination of odors. The placement of reagents, supplies, and equipment within the hood should not interfere with the proper airflow.
Maintenance
1. At least yearly, with the sash fully open and the cabinet empty, check the air velocity with a thermoanemometer (minimum acceptable face velocity, 100 ft/min) (7). Also, a smoke containment test should be performed with the cabinet empty to verify proper directional face velocity.
2. Lubricate the sash guides as needed.
The Class II-A1 or II-A2 biological safety cabinet (BSC) is best suited and recommended for the diagnostic laboratory. BSCs operate at a negative air pressure with air passing through a HEPA filter, and the vertical airflow serves as a barrier between the cabinet and the user. Although a BSC is not required for processing routine specimens in a diagnostic parasitology laboratory, some laboratories use class I (open-face) or class II (laminar-flow) BSCs for processing all unpreserved specimens (3). Use of a BSC is recommended if the laboratory is performing cultures for parasite isolation. However, remember that BSCs should not be used as fume hoods. Toxic, radioactive, or flammable vapors or gases are not removed by HEPA filters (8).
Maintenance (5)
1. After each use, disinfect the work area. Since UV radiation has very limited penetrating power, do not depend on UV irradiation to decontaminate the work surface (9). At least weekly, clean UV lamps (in the off position) with 70% isopropyl or ethyl alcohol.
2. At least annually, have class I BSCs certified. They should also be certified after installation but before use and after they have been relocated or moved. Certification should include the following and will be documented by the trained company representative (contracted to handle the BSC inspection).
A. Measurements of the air velocity are taken at the midpoint height approximately 1 in. behind the front opening. Measurements should be made approximately every 6 in.
B. The average face velocity should be at least 75 linear ft/min. A thermoanemometer with a sensitivity of ±2 linear ft/min should be used (10).
C. With the cabinet containing the routine work items, such as a Bunsen burner, test tube rack, bacteriological loop and holder, etc., a smoke containment test should be performed to determine the proper directional velocity.
D. Record the date of recertification, the names of the individual and company recertifying the cabinet, and any recommendations for future service. Any maintenance performed should also be documented in writing.
3. Replace the filters as needed.
4. On installation, have a class II BSC certified to meet Standard 49 of the National Sanitation Foundation, Ann Arbor, MI (10). The cabinet must also be recertified at least annually and/or when it is moved, after filters are replaced, when the exhaust motor is repaired or replaced, and when any gaskets are removed or replaced. Record the date of recertification, the names of the individual and company performing the service, and any recommendations for future service.
Any general-purpose laboratory (non-explosion-proof) or household-type refrigerator-freezer (4 to 6°C) can be used in the parasitology laboratory. Solvents with flash points below refrigeration temperature should not be stored, even in modified (explosion-proof) refrigerators.
Maintenance
1. On a daily basis, monitor and record the temperature of the refrigerator. The thermometer should be placed into a liquid to permit stable temperature recording, or thermocouples may be used.
2. On a daily basis, monitor and record the temperature of the freezer. The thermometer should be placed in antifreeze (any brand with freezing point below that of the freezer, e.g., ethylene-glycol-water solutions, glycerol-water solutions, or Prestone) to permit stable temperature recording. Thermocouples may be used instead.
3. Periodically when the door is opened, check to see if the fan is operational.
4. Monthly, check the door gasket for deterioration, cracks, and proper seal. Seal problems are often seen when ice begins to build up in a freezer or the temperature is not holding. Periodically, petroleum jelly can be rubbed onto the door gasket to lubricate the material and to help maintain flexibility for a tight seal when the door is shut.
5. Semiannually, clean the condenser tubing and air grill with a vacuum cleaner.
6. Semiannually, check to ensure that the drain tubes are kept open.
7. Annually, wash the interior with a warm solution of baking soda and water (approximately 1 tablespoon/qt [ca. 13 to 14 g/0.946 liter]). Rinse with clean water, and dry. Also, wash the door gasket and water collection tray with a mild soap and water. If the gasket accumulates a black mold, scrub with 50% household bleach solution and a small brush. Rinse with clean water, and dry.
Supplies for the diagnostic parasitology laboratory are often identical to those needed for routine work in other areas of microbiology. Although not every size of glassware used is specified, the list below should be helpful for anyone setting up a laboratory for this type of work.
1. Disposable glass or plastic pipettes and bulbs (some sterile for culture work)
2. Pipettes: 1, 5, and 10 ml (some sterile for culture work)
3. Glass slides (1 by 3 in., or larger if preferred). Slides with rounded edges are now available (safety-“sharps”).
4. Coverslips (22 by 22 mm; no. 1 or larger if preferred)
5. Beakers, 250, 500, 1,000, and 2,000 ml
6. Covered Coplin jars or staining dishes (with slide rack)
7. Graduated cylinders, 50, 100, 500, and 1,000 ml
8. Mortar and pestle (range of sizes)
9. Flasks, Erlenmeyer, 500 and 1,000 ml
10. Flasks, volumetric, 500 and 1,000 ml
11. Bottles, brown, 150 to 200 ml
12. Bottles, clear, 100, 500, and 1,000 ml
13. Bottles, airtight, 50 ml
14. Funnel (glass) to hold filter paper
15. Büchner funnel
16. Centrifuge tubes, 15 and 50 ml (some with screw caps)
17. Petri dish, plastic, sterile
18. Tubes, screw-cap, 13 by 100 mm or 16 by 125 mm (some sterile for culture work)
19. Plastic syringe, 15 ml
20. Sterile syringes (glass or plastic), 1, 10, 20, and 50 ml
1. Culture tube racks
2. Gauze (woven or pressed)
3. Applicator sticks (wood)
4. Sterile cryovials or screw-cap vials (to hold 1 ml)
5. Box for vial storage in freezer
6. Filter paper, Whatman no. 1 and Whatman no. 42
7. Sterile filtration system
8. Membrane filters (pore size, 0.22 µm) (to be used with sterile filter)
9. Nuclepore membrane filter, 25-mm, 5-µm, and 3-µm porosity
10. Swinney filter adapter (attaches to syringe, holds filter)
11. Filter paper pad, 25 mm (used to support the membrane filter in the Swinney adapter)
12. Bacteriological loop
13. Sterile syringe needles, 20 and 27 gauge
14. Vaspar
15. Parafilm (American Can Co.) or equivalent
16. Slide boxes for positive-slide storage
17. Forceps and scissors
18. Stage micrometer with scale of 0.1- and 0.01-mm divisions
19. Disk micrometer divided into 50 units
20. Biohazard container with disinfectant for proper disposal of slides, tubes, and pipettes
21. Biohazard container for proper disposal of patient specimens
22. Microscope lens paper
ATCC Quality Control Organisms
1. An 18- to 24-h-old culture of Escherichia coli or Enterobacter aerogenes
2. ATCC 30010 (Acanthamoeba castellanii)
3. ATCC 30133 (Naegleria gruberi)
4. ATCC 30925 (Entamoeba histolytica HU-1:CDC)
5. ATCC 30015 (Entamoeba histolytica HK-9)
6. ATCC 30001 (Trichomonas vaginalis)
7. ATCC 30883 (Leishmania mexicana)
8. ATCC 30160 (Trypanosoma cruzi)
Note None, some, or all of these QC organisms will be required, depending on the culture procedures performed in the laboratory.
Safety: Personnel and Physical Facilities
1. Be careful! All material to be received by or discarded from the laboratory must be considered potentially pathogenic.
2. Smoking, eating, or drinking in the laboratory is not permitted.
3. Do not work with uncovered open cuts or broken skin. Cover them with a Band-Aid, finger cot, or other suitable means, such as rubber or plastic gloves.
4. Mouth pipetting of specimens is not permitted.
5. Do not create aerosols. Remember, infectious diseases, such as infectious hepatitis, may be transmitted by aerosols produced by centrifuges, stirrers, pipettors, etc. Exercise extreme care when using such equipment. Cool inoculating loops or needles before touching colonies on plates or inserting into liquid material.
6. Develop the habit of keeping your hands away from your mouth, nose, and eyes. Wash hands well with soap before leaving the laboratory.
7. Do not lay personal articles, such as eyeglasses, on the bench in your work area.
8. Laboratory coats or gowns are not to be worn outside the laboratory, particularly not to the employee lounge or cafeteria or out of the building.
9. Wipe off benches in your working area with disinfectant before and after each day’s work. Keep your area clean at all times.
10. In case of injury or unusual incident, however slight, the supervisor in charge must be immediately notified, and a report to occupational health facility is required. Also fill out an accident report form. Major accidents must be documented and reported in detail to the supervisor and chief of microbiology. The report should indicate:
A. Cause of the accident
B. Type of contamination or hazard
C. List of personnel possibly exposed and amount of exposure to possible pathogenic material
D. Decontamination procedures taken if pathogenic material was involved
E. Actions taken to prevent recurrence
F. Actions taken to safeguard or monitor employees
11. Infections may be spread by a number of routes in the laboratory. The actual occurrence of an infection depends on the virulence of the infecting agent, susceptibility of the host, route of entry, inoculating dose, etc. (Table 11.1).
A. Airborne. Droplets and aerosols may be formed by simply removing caps or cotton plugs or swabs from tubes. Heating liquids on needles too rapidly may also create an aerosol. Breakages in centrifuges are serious accidents (Table 11.2).
B. Ingestion. Ingestion can occur through mouth pipetting, failure to wash hands after handling specimens or cultures, or smoking, eating, and drinking in the laboratory.
C. Direct inoculation. Scratches, needlesticks, cuts from broken glass, or animal bites may permit direct inoculation.
D. Skin contact. Some very virulent organisms, and others not so virulent, can enter through small cuts or scratches or through the conjunctiva of the eye.
E. Vectors. Flies, mosquitoes, ticks, fleas, and other ectoparasites can be potential sources of infection in the laboratory, especially if animal work is performed.
12. Personnel who display risk-prone behavior or are pregnant, immunocompromised, or immunosuppressed should be restricted from performing work with highly infectious microorganisms and, in some situations, be restricted to a low-risk laboratory (22).
Handwashing is the most important procedure to reduce the duration of exposure to an infectious agent or chemical, to prevent dissemination of an infectious agent, and to reduce overall infection rates in a health care facility. Hand contamination occurs during manipulation of specimens and during contact with work surfaces, telephones, and equipment. Laboratory personnel should wash their hands immediately after removing gloves, after obvious contamination, after completion of work, before leaving the laboratory, and before hand contact with nonintact skin, eyes, or mucous membranes (8).
Handwashing sinks should be located at each entry/exit door; if possible, the faucet should be operated by a knee/foot control. If these controls are not available, the faucet should be turned on and off using a paper towel. Handwashing should be performed using soap or an antiseptic compound, starting at the wrist area and extending down between the fingers and around and under the fingernails, and rinsed from the wrists downward. Recently, the Centers for Disease Control and Prevention recommended that in addition to traditional soap and water handwashing, health care personnel can use alcohol-based gels (8).
Personal Protective Equipment (OSHA 2001 Blood Borne)
Occupational Safety and Health Administration (OSHA) standards for exposure to blood-borne pathogens require the laboratory to provide personal protective equipment (PPE) for its employees. OSHA standards require that PPE be provided, used, and maintained for all hazards found in the workplace, including biological, environmental, and chemical hazards, radioactive compounds, and mechanical irritants capable of causing injury or illness through absorption, inhalation, or physical contact. Employees must be trained in the appropriate use of PPE for a specific task, the limitations of PPE, and procedures for maintaining, storing, and disposing of PPE.
Gloves
Gloves protect the wearer from exposure to potentially infectious material and other hazardous material and are available in material designed for specific tasks. Gloves must be provided by the employer and must be the proper size and appropriate material for the task. Due to latex hypersensitivity in some workers, only powder-free latex gloves should be used or gloves should be manufactured from nitrile, polyethylene, or other material.
Protective Clothing
Laboratory workers should wear long-sleeved coats or gowns that extend below the level of the workbench, and they should be worn fully closed. The material must be fluid resistant if there is any potential for splashing or spraying. Fluid-proof clothing (plastic or plastic lined) must be worn when there is the potential for soaking by infectious material. Laboratory workers should not wear laboratory clothing outside the laboratory. All protective clothing should be changed immediately when contaminated to prevent the potentially infectious material or chemical from contacting the skin. Coats and gowns should not be taken home for cleaning but should be laundered by the institution.
Face and Eye Protection
Face and eye protection should be used when splashes or sprays of infectious material or chemicals may occur. Equipment includes goggles, face shields, and splashguards. Face shields provide the best protection for the entire face and neck, although splashguards provide an alternative method. If only goggles are worn, the user should also wear an appropriate mask to prevent contamination of mucous membranes.
1. Specimens with gross internal contamination must not be accepted. Place the specimen in a plastic bag to protect subsequent handlers. The test request slips should be bagged separately from the specimen (keep paperwork clean and uncontaminated). Wear gloves when handling the specimen, and wash your hands thoroughly when you finish handling the specimen. Notify the supervisor immediately about a contaminated specimen so that further corrective action may be taken.
2. Specimens to be centrifuged must be placed in a sealed container to prevent aerosols.
3. All specimens other than bacterial or fungal blood culture bottles to be set up for isolation of organisms must be processed in a BSC. All “open-system” work is to be done on an absorbent surface (i.e., a towel) in a BSC, using appropriate protective techniques. Towels should be changed daily. We recommend that the immediate working area of the towel be dampened with any recommended disinfectant to handle small spills (Fig. 11.3).
Figure 11.3 Example of a commercially available disinfectant (Solucide) that can be used for countertops, small spills, and any hard surface that requires a disinfectant (www.med-chem.com). doi:10.1128/9781555819002.ch11.f3
1. All specimens are potentially pathogenic—always use careful techniques (Tables 11.3 and 11.4).
2. All discard material used in processing of specimens is considered contaminated.
A. Fluids used in the processing of specimens (buffers, etc.), as well as excess liquid specimens, should be poured into plastic screw-top autoclavable bottles and sterilized prior to disposal. A tongue depressor placed in the bottle can help to prevent splashing if liquid is poured down the slanting stick. These sticks can also be placed in disinfectant before being discarded. Disinfectants and incineration can also be used for decontamination of infectious materials.
B. Reusable items (tissue grinders, bottles, etc.) are placed in an autoclave pan. Once the pan is full, place it (without a cover) in a large brown autoclave bag and staple shut. After being autoclaved, the reusable items can be cleaned.
C. Specimen containers and centrifuge tubes are disposable. These items should be placed in plastic autoclave bags and secured with masking tape for sterilization. Slides can also be placed in containers of liquid disinfectant prior to disposal. Remember, slides should be treated as sharps.
D. Pipettes can be placed in a covered discard pan containing 5% amphyl or placed in autoclavable containers for sterilization before discarding.
1. Disinfectants in use at present
A. Bleach solutions. To be an effective disinfectant, working bleach solutions must contain >0.5% but <2% sodium hypochlorite. Hypochlorite concentration in household bleach varies by manufacturer. Many household bleach solutions contain 5.25% sodium hypochlorite, and a 1:10 dilution (5,000 ppm Cl) will produce a 0.53% hypochlorite solution. Use of bleach solutions with lower hypochlorite concentrations might not provide the proper level of disinfection. Each day, prepare a fresh 1:10 household bleach solution (3, 23).
B. Alcohols. Alcohols generally act by coagulating proteins and as organic solvents. Rapid evaporation may lead to inadequate “killing” time. Although there are some conflicting reports, 70% solutions generally have the best microbicidal activity. Caution is advised because of flammability. DO NOT USE ALCOHOLS OR ALCOHOL-BASED SOLUTIONS ALONE TO DISINFECT SURFACE AREAS; EVAPORATION SUBSTANTIALLY DECREASES EFFICACY.
C. Another option can be seen in Fig. 11.3.
Note Use disinfectants recommended for environmental surfaces, such as Environmental Protection Agency (EPA)-registered disinfectants effective against hepatitis B virus, human immunodeficiency virus (HIV), and other blood-borne pathogens, or use a 1:10 dilution of household bleach. EPA environmental disinfectant product registration information is available at http://www.epa.gov/oppad001/chemregindex.htm (accessed 9/11/2013).
2. Immediate actions
A. Clear the area at once.
B. Notify the supervisor and chief of microbiology.
C. Assess the type of spill and degree of hazard involved.
D. Determine the most effective and least hazardous approach to clean up and decontaminate.
3. “Dry spills” with no significant aerosol formation
A. Flood the area with disinfectant solution.
B. Soak up disinfectant and contaminated material with absorbent material (sand or paper towels), and dispose of it in a plastic biohazard bag or sealed container. Wear gloves for cleanup.
C. If routine cleanup is not possible, the unit may have to be decontaminated with a sterilizing gas such as paraformaldehyde.
D. Thoroughly wash the unit (if possible) after decontamination.
4. Liquid spills on the bench or floor
A. If significant aerosols were formed, the area should be evacuated and not reentered for at least 1 h.
B. Flood the area with disinfectant, and cover the spill with absorbent material (sand or paper towels). Wear gloves during cleanup.
C. Dispose of absorbent and contaminated material in plastic bags or sealed containers, and autoclave.
D. Thoroughly wash the area after cleanup.
5. Centrifuge spills
A. Shut off the instrument, and evacuate the area at once.
B. Do not reenter the area for at least 1 h until aerosols have settled.
C. The person entering the area to clean up should wear protective clothing, gloves, and a mask.
D. If liquids are present, soak up in absorbent material and handle as above. If liquids are not present, clean the instrument and room thoroughly before resuming work.
E. Wipe all surfaces with a recommended disinfectant.
6. Spills in incubators, autoclaves, or other closed areas
A. Flood with disinfectants, and soak up liquids with absorbent material.
B. Dispose of the material as specified above, if possible.
C. If routine cleanup is not possible, the unit may have to be decontaminated with a sterilizing gas such as paraformaldehyde.
D. Thoroughly wash the unit (if possible) after decontamination.
Disposal of Contaminated Materials (3, 24)
1. Autoclave screw-cap tubes before cleaning or discarding.
2. Discard specimens and cultures into containers with double plastic lining. Liners should be changed when the containers are about half full.
3. Place culture plates into “Biohazard” receptacles lined with autoclavable bags.
4. Materials or containers to be reused should be autoclaved before being cleaned. Place them in sealed and clearly labeled containers to minimize hazard to others before sterilization.
5. Any breakage of glass or leakage of contaminated materials should be reported to the supervisor or chief of microbiology at once for instructions on procedures for safe cleanup.
All laboratories should adhere to the concept of standard precautions, which state that all patients and all laboratory specimens are potentially infectious and should be handled accordingly (8, 24–26). This concept arose from the observation that infections are often unrecognized in patients. “Standard precautions” replaces earlier terms such as Blood and Body Fluid Precautions, Universal Precautions, and Body Substance Precautions found in OSHA documents (8, 26). The OSHA documents place the emphasis on blood-borne pathogens such as HIV and hepatitis B and C viruses, whereas the concept of standard precautions recognizes that all infectious agents and all other potentially infectious material, except sweat, pose a risk to the health care worker (8).
OSHA identifies a number of practices that should be implemented to protect the worker from exposure to blood-borne pathogens, including an exposure control and risk assessment plan (7). Methods should be implemented to minimize exposure to infectious agents, to shield the laboratory worker from infectious material through a set of engineering and work practice controls, and to use personal protective equipment. In addition, the OSHA regulations require that employers provide hepatitis B vaccination and postexposure evaluation and follow-up; communicate the hazards to employees; and maintain appropriate records (8). Employees who decline immunization against hepatitis B virus are required to sign a hepatitis B vaccine declination form.
Note Under circumstances in which differentiation between body fluid types is difficult or impossible, all body fluids shall be considered potentially infectious.
1. All job positions with any exposure to blood and body fluids must be listed on an exposure form, which must match up with continuing education training records from the facility.
A. Training must be provided at no cost to the employee and during working hours.
B. Training shall be provided as follows:
a. At the time of initial assignment to job functions in which occupational exposure may occur
b. Within 90 days after the effective date of the standard
c. At least annually thereafter
2. A complete task assessment must be performed for employees who have any exposure; protective measures, including equipment, must be defined.
A. Engineering controls
B. Immunization programs
C. Work practices (hand washing and procedures for handling sharps)
D. Disposal and handling of infectious waste
E. Use of personal protective gloves, gowns, and goggles
F. Use of mouthpieces, resuscitation bags, or other ventilation devices
G. Use of disinfectants
H. Labeling and signs
I. Training and education programs
J. Postexposure follow-up
Some of the specifics are as follows:
A. All procedures shall be performed to minimize splashing, spraying, spattering, and generation of droplets of these substances.
B. When there is occupational exposure, the employer shall provide, at no cost to the employee, PPE such as (but not limited to) gloves (including hypoallergenic gloves, glove liners, powderless gloves, or other alternatives), gowns, laboratory coats, face shields or masks, and eye protection. Other options might include mouthpieces, resuscitation bags, pocket masks, or other ventilation devices.
C. PPE is considered “appropriate” only if it does not permit blood or other potentially infectious materials to pass through or to reach the employee’s work clothes, street clothes, undergarments, skin, eyes, mouth, or other mucous membranes. The type and characteristics of outer garments depend on the tasks and degree of exposure anticipated.
D. All working surfaces shall be cleaned and decontaminated after contact with blood or other potentially infectious materials. All spills shall be immediately contained and cleaned up by appropriate professional staff or others trained and equipped to work with potentially concentrated infectious materials.
E. Specimens of blood or other potentially infectious materials shall be placed in a container which prevents leakage during collection, handling, processing, storage, transport, or shipping (27–29).
F. Mouth pipetting or suctioning of blood or other potentially infectious material is prohibited.
G. Certified BSCs or other combinations of personal protection or physical containment devices shall be used for all activities that may pose a threat of exposure. BSCs shall be certified when installed, whenever they are moved, and at least annually.
3. Key questions that people ask concerning these rules are listed below (30).
A. How does OSHA define “occupational exposure”?
“Reasonably anticipated skin, eye, mucous membrane, or parenteral contact with blood or other potentially infectious materials that may result from the performance of an employee’s duties”
B. What does “reasonably anticipated” mean?
“Actual contact would be expected during an autopsy or surgery. In these cases, blood or other potentially infectious materials come in direct contact with the employee’s gloves or other protective clothing. In other cases, contact may not occur each time the task or procedure is performed, but when blood or other potentially infectious materials are an integral part of the activity, it is reasonable to anticipate that contact may result.”
C. What should the exposure control plan contain?
“The Exposure Control Plan is a key provision because it requires the employer to identify the individuals who will receive the training, protective equipment, vaccination, and other provisions of the standard.”
D. What is considered “appropriate” personal protective equipment?
“Gloves, gowns, lab coats, face shields or masks and eye protection, mouthpieces, resuscitation bags, pocket masks, or other ventilation devices. Such protection will be considered ‘appropriate’ only if it does not permit blood or other potentially infectious materials to pass through to or to reach the employee’s work clothes, street clothes, undergarments, skin, eyes, mouth, or other mucous membranes under normal conditions of use and for the duration of time which the protective equipment will be used. The type and characteristics of outer garments will depend upon the tasks and degree of exposure anticipated.”
E. Does this mean that protective clothing must be “impervious” or “fluid-proof”?
“Selection of personal protective equipment is performance-oriented. Selection of the type and characteristics of necessary personal protective equipment is based upon the exposure anticipated to be associated with the task. In those instances where such equipment was incapable of halting penetration of blood or other potentially infectious materials normally encountered during a procedure, then additional protection such as a plastic apron should be worn or used.”
Note “Fluid-resistant” and “fluid-proof” no longer appear in the final rule.
F. How can the employer ensure that the employee uses appropriate PPE?
“It is not the intent that the employer ‘watch over everyone’s shoulder’ for compliance, but there are reasonable policies and regulations that employees follow.”
G. What does the standard say about accessibility, cleaning, repair, removal, and storage of PPE?
“The employer shall ensure that appropriate personal protective equipment in appropriate sizes is available and is accessible at the worksite; the employer shall clean, launder, and dispose of personal equipment; the employer shall repair or replace when necessary at no cost to the employee. If a garment is penetrated by potentially infectious materials, it shall be removed ASAP and replaced by clean garments. When these garments are removed, they must be placed in a designated area or container for storage, washing, decontamination or disposal.”
H. What is the employer’s responsibility if PPE or the hepatitis B vaccine is refused?
“The employer shall ensure that the employee uses appropriate personal protective equipment unless the employer shows that the employee temporarily and briefly declined to use personal protective equipment when, under rare and extraordinary circumstances, it was the employee’s professional judgment that its use would have prevented the delivery of health care services or would have created an increased hazard to the safety of the worker or co-worker. When an employee makes this decision, the circumstance shall be investigated and documented in order to determine whether changes need to be made to prevent such reoccurrences.”
If the employee declines to accept hepatitis B vaccination, a form indicating that fact must be signed by the employee and held in the file.
I. What training is required for employees?
“Training must be available at the worksite, at no cost to the employee, and during working hours. It shall be provided at the time of initial assignment to tasks where exposure may occur, within 90 days after the effective date of the Standard, and at least annually thereafter.”
J. What type of training is required for personal protective equipment?
“An explanation of the use and limitations of methods that will prevent exposure is required, including engineering controls (biosafety cabinets), work practices, and personal protective equipment. Information on types, proper use, location, removal, handling, decontamination, and disposal of personal protective equipment must be presented, in addition to an explanation of the basis for selection of such controls.”
Hepatitis Exposure Protocol (4)
Laboratory personnel have a high risk of contracting hepatitis for the following reasons:
1. Frequent close personal contact with patients with hepatitis
2. Direct and frequent contact with biological specimens containing the virus
3. Frequent opportunity for accidental puncture wound with contaminated needles and sharp objects
4. Carelessness in handling specimens
5. Inadequate or unsafe disposal of contaminated needles, specimens, or other objects
6. Carelessness in proper and frequent hand washing technique
The prevention of hepatitis is the primary reason for the emphasis on infection control precautionary measures found in any safety manual (e.g., no smoking, eating, drinking, or mouth pipetting, and procedures for safe specimen handling). These same procedures also apply to HIV.
All known hepatitis specimens must be so labeled; however, to minimize exposure to the virus, every specimen should be handled as though it could transmit hepatitis. Studies have shown that hepatitis may be transmitted to laboratory personnel in several ways:
1. Puncture or other wounds (HIV)
2. Abrasions of the skin (HIV)
3. Aerosols (inhaled into the respiratory tract)
4. By mouth (oral route)
5. Direct contact with the patient
6. Splashing material into the eyes
The following protocol should be followed in the event of exposure:
1. Immediately wash the exposed area with soap and water.
2. Report the incident to the supervisor and laboratory personnel office.
3. Consult with a physician in Employee Health Facility concerning possible recommendations (e.g., administration of immune serum globulin).
Dangerous Properties of Industrial Materials
OSHA requires each laboratory to develop a comprehensive, written chemical hygiene plan (CHP). Every hazardous chemical in the laboratory, regardless of the type of risk, volume, or concentration, must be included in the CHP. The plan should include storage requirements, handling procedures, location of OSHA-approved material safety data sheets, and the medical procedures that are to be followed if exposure occurs. The CHP must specify the clinical signs and symptoms of the environmental conditions (such as a spill) that would give the employer reason to believe that exposure had occurred. When such conditions exist, the CHP should indicate the appropriate medical attention required.
Ethyl ether (also known as ether, anesthesia ether, diethyl ether, ethyl oxide, sulfuric ether, and ethoxyethane) is so volatile that dangerous concentrations are readily built up in the laboratory atmosphere. It is highly flammable and has a tendency to form explosive peroxides. Its low flash point, low ignition temperature, wide explosive range, high volatility, and very heavy vapor (which tends to “pocket”) combine to make ethyl ether an extremely serious fire hazard. The spontaneous formation of explosive peroxides presents severe risks to the user who does not apply precautionary practices. Therefore, it is mandatory to follow these safety procedures (31, 32).
1. Date cans of ether immediately upon receipt.
2. Do not attempt to open a can that is 1 year old (after time of receipt); peroxides may form in sealed cans and explode when the cans are opened. Order appropriate quantities to avoid expensive waste.
3. Date the can of ether when opened; do not use after 1 month. Order cans of appropriate size to avoid waste.
4. Store opened (corked) cans of ether close to the floor, in a cool place, on a shelf, or in a cabinet that is not airtight. The heavy vapors can dissipate along the floor, with less opportunity for contact with sources of ignitions such as an open flame and electric sparks.
5. Never store opened cans of ether in a refrigerator, since ether still vaporizes at refrigerator temperatures. If peroxides have formed from ether, merely opening the refrigerator door may trigger an explosion. The heavy vapors that collect in an airtight refrigerator may ignite on contact with the electrical refrigerator motor. This recommendation also applies to “shielded” refrigerators (those in which the electrical systems are protected).
6. Unopened cans may be stored with opened cans (near the floor on an open shelf or nonairtight cabinet), in a storage cabinet for flammable materials, or in a storage room for volatile substances.
7. Outdate and discard cans after storage for 1 month (if opened) or 1 year (if unopened).
8. Flush empty, or almost empty, ether cans with copious amounts of water before discarding. “Empty” cans have been known to explode! Do not flush more than the contents of a 0.25-lb (ca. 0.1-kg) can down the drain; consult the Research and Occupational Safety Office or the local fire department for proper disposal of large quantities. Empty, thoroughly rinsed cans may be discarded in the regular trash.
9. Do not attempt to open any containers of uncertain age or condition or those whose cap or stopper is tightly stuck. Peroxides have been known to form under the threads of caps! These containers should be discarded.
10. Unopened outdated cans must be delivered to Research and Occupational Safety for disposal (each institution may have a comparable office or safety office).
11. Certain chemicals can be replaced by others that are safer (33, 34). Ethyl acetate, with a higher flash point (4.0°C), has replaced ether (flash point of −45°C) in the formalin-ether concentration technique. Hemo-De (Medical Industries, Los Angeles, CA, or Fisher Scientific, Los Angeles, CA) can also replace ethyl acetate in the concentration procedure. Hemo-De has a flash point of 57.8°C and is generally regarded as safe by the U.S. Food and Drug Administration. Xylene, as used in the trichrome or iron hematoxylin staining of polyvinyl alcohol (PVA)-fixed fecal smears, poses potential toxic and fire hazards. Again, Hemo-De has successfully replaced xylene in both the carbol-xylene and xylene steps of the trichrome procedure (33). Other substitutes are available as well (e.g., Hemo-Sol [Fisher Scientific]). Check with your local pathology departments or reagent suppliers for other alternatives.
12. Mercuric chloride, used in PVA and in Schaudinn’s fixative, presents both a toxic hazard and a disposal problem. Copper sulfate has been suggested as a substitute for mercuric chloride (35, 36); however, protozoan morphology is not as clear and precise when this formula is used. The use of zinc rather than copper gives fairly good morphology when the trichrome stain is used (35).
Note Information on other substances is provided in Table 11.5.
Current OSHA Regulations for the Use of Formaldehyde
Formaldehyde has been in use for over a century as a disinfectant and preservative; it is also found in a number of industrial products. There is disagreement about the carcinogenic potential of lower levels of exposure, and epidemiologic studies of the effects of formaldehyde exposure among humans have given inconsistent results. Studies of industry workers with known exposure to formaldehyde report little evidence of increased cancer risk (37). It also appears that persons with asthma respond no differently from healthy individuals following exposure to concentrations of formaldehyde up to 3.0 ppm (38).
OSHA requires all workers to be protected from dangerous levels of vapors and dust. Formaldehyde vapor is the most likely air contaminant to exceed the regulatory threshold in the clinical laboratory (39, 40). Current OSHA regulations require vapor levels not to exceed 0.75 ppm (measured as a time-weighted average [TWA]) and 2.0 ppm (measured as a 15-min short-term exposure). OSHA requires monitoring for formaldehyde vapor wherever formaldehyde is used in the workplace. The laboratory must have evidence at the time of inspection that formaldehyde vapor levels have been measured, and both 8-h and 15-min exposure must have been determined.
If each measurement is below the permissible exposure limit and the 8-h measurement is below 0.5 ppm, no further monitoring is required as long as laboratory procedures remain constant. If the 0.5-ppm 8-h TWA or the 2.0-ppm 15-min level is exceeded, monitoring must be repeated semiannually. If either the 0.75-ppm 8-h TWA or the 2.0-ppm 15-min level is exceeded (unlikely in a clinical laboratory setting), employees must be required to wear respirators. Accidental skin contact with aqueous formaldehyde must be prevented by the use of proper clothing and equipment (gloves and laboratory coats).
The amendments of 1992 add medical removal protection provisions to supplement the existing medical surveillance requirements for employees suffering significant eye, nose, or throat irritation and for those experiencing dermal irritation or sensitization from occupational exposure to formaldehyde. In addition, these amendments establish specific hazard-labeling requirements for all forms of formaldehyde, including mixtures and solutions composed of at least 0.1% formaldehyde in excess of 0.1 ppm. Additional hazard labeling, including a warning label that formaldehyde presents a potential cancer hazard, is required where formaldehyde levels, under reasonably foreseeable conditions of use, may potentially exceed 0.5 ppm. The final amendments also provide for annual training of all employees exposed to formaldehyde at levels of 0.1 ppm or higher (41).
Note The use of monitoring badges may not be a sensitive enough method to correctly measure the 15-min exposure level. Contact the Occupational Health and Safety Office within your institution for monitoring options. Usually, the accepted method involves monitoring airflow in the specific area(s) within the laboratory where formaldehyde vapors are found.
The incidence and evidence of clinical latex sensitivity appear to be increasing since its first description in 1979. Since the introduction of standard precautions, the use of latex gloves by health care workers has increased dramatically (7). Those at risk appear to be health care workers and employees working in latex industries, patients with atopic diathesis, and patients who underwent repeated surgical procedures during childhood (42). The allergic response occurs to one or more natural rubber latex proteins, resulting in contact urticaria, angioedema, allergic rhinitis, asthma, or anaphylaxis. Early recognition of sensitization to latex is crucial to prevent the occurrence of life-threatening reactions in sensitized health care providers or patients (43). Latex allergy is now an important medical, occupational, medicolegal, and financial problem, and it is essential that policies be developed to reduce the problem (44–46). There also seems to be a relationship between allergies to natural latex rubber and allergies to avocados, chestnuts, and bananas. A high incidence of latex sensitivity has also been found in people with spina bifida (47).
The use of cornstarch powder in gloves can sensitize healthy people and exacerbate the symptoms of allergic patients. The powder spreads the latex allergens into the environment (42). Accompanying this increase in latex allergy is evidence of positive circulating specific immunoglobulin E antibodies to latex (48). Unfortunately, the primary treatment for latex allergy is avoidance (49).
The area of quality systems forms an integral part of any clinical laboratory operation. In many cases of clinical laboratory testing, various programs and guidelines for quality assurance procedures (including QC) have been thoroughly outlined. However, specific quality assurance recommendations within the parasitology section of microbiology have not been well defined in the past, and the information in this chapter should help clarify these recommendations. Although the diagnostic procedures in this area seldom yield numerical data, there are still a number of QC measures that can help to ensure accurate results.
The range of services provided will vary from one laboratory to another. To comply with the Health Care Financing Administration (HCFA) guidelines, only two extent classifications (rather than the previous four) are recognized in parasitology by the College of American Pathologists (50).
1. Extent 2. “Limited parasitology performed. The laboratory is able to recognize the presence of parasites, including protozoa in clinical specimens, but may need to refer them for definitive identification. When permanently stained smears for intestinal protozoa are indicated, polyvinyl alcohol fixative (PVA) or other appropriate fixative is available, and such material is placed in a fixative and then referred to a reference laboratory for staining identification.”
2. Extent 3. “Definitive identification of parasites present to the extent required to establish a correct clinical diagnosis and to aid in selection of safe and effective therapy. Laboratories are expected to have a high degree of expertise and should be able to differentiate Plasmodium falciparum from other species of Plasmodium.”
Note There is no longer a College of American Pathologists listing for extent 1 (no parasitologic procedures are performed and all specimens are submitted to a reference laboratory). However, in this situation, the laboratory would not be required to subscribe to outside proficiency testing specimens in this discipline.
Based on the Extent of Services mentioned above and the requirements in the Clinical Laboratory Improvement Amendments of 1988 (CLIA ’88), all (with rare exceptions) clinical laboratories (including physician office laboratories performing other than waived tests) must participate in an approved program of interlaboratory comparison testing that is consistent with the level and complexity of the work performed. The comprehensive microbiology and/or parasitology survey programs are recommended. The state Department of Health must be consulted to confirm which proficiency testing services fulfill the requirements related to a laboratory’s particular accreditation, particularly if the laboratory accepts interstate specimens in parasitology.
There must be evidence of active review of the survey results by the laboratory director or the designated supervisor. There also must be written evidence of evaluation of the results and, if indicated, corrective action when unacceptable results have been reported. Corrective action statements should review the correct answer and specific steps that will be taken to ensure that the same, or similar, error will not occur in the future. On the basis of the regulations within CLIA ’88, HCFA or one of the HCFA-approved agencies (may include some state agencies) will be involved in the process of monitoring laboratory performance and coordinating any possible sanctions that might be involved as a result of unacceptable performance in proficiency testing programs.
Copies of the 28 February 1992 Federal Register containing the final CLIA regulations can be obtained by writing to Government Printing Office, Attn: New Order, P.O. Box 371954, Pittsburgh, PA 15250-7954. Specify the date of the issue requested (28 February 1992) and stock number (069-001-00042-4). A $3.50 check or money order payable to “Superintendent of Documents” or a Visa or MasterCard number and expiration date should be enclosed. Orders may be made by telephone [(202) 783-3238] or fax [(202) 512-2250].
It is recommended that all of the proficiency testing specimens be saved for future review, particularly when students are being trained or technologists, pathologists, and residents are receiving review training. It is just as important to maintain negative specimens as to maintain positive ones, since many errors are made in which artifacts are identified as parasites. Specimens should be periodically checked to make sure that the volume of fixative is sufficient to prevent the specimens from drying out. These specimens should be cataloged for easy reference.
Specific recommendations for the handling and examination of proficiency testing specimens can be found in Table 11.6. These guidelines should assist the participant in the proper approach to these specimens.
CLIA ’88 mandated universal regulation for all clinical laboratory testing sites in the United States, including previously unregulated sites in physician offices. Quality testing was to be achieved through a combination of total quality management and mandated minimum quality practices. Through both internal and external proficiency testing, performance standards were also mandated. After the implementation of CLIA ’88, the percentage of laboratories passing proficiency testing requirements has improved and almost all laboratories are currently using quality practices for a more comprehensive quality system approach (51, 52).
Survey results from the Laboratory Proficiency Testing Program, Toronto, Ontario, Canada, over the course of the program, which began in 1977, indicate a marked improvement in laboratory performance. They send four samples in each of four surveys per year consisting of specimens to be examined for gastrointestinal parasites. Improvement is thought to result from a combination of voluntary withdrawal by laboratories with poor performance and improved performance by the other laboratories. Extensive educational initiatives have also played a significant role in improved performance (53).
Supervision
The QC program should be under surveillance by the chief technologist or section supervisor and should be reviewed at least once each month by the laboratory director, QC supervisor, or other supervisor designate. There should be written evidence of active review of all records (controls for routine procedures, instrument function tests, and equipment checks [temperature, humidity, systems, maintenance]). There should also be written evidence of corrective action when controls do not fall within acceptable limits. A written checking system should be in operation to help detect clerical errors, analytical errors, and/or unusual laboratory results. This system should also provide for error correction within acceptable time frames. With the emphasis on outcome-oriented measures, it is very important to have the capability to track testing through the preanalytical, analytical, and postanalytical phases (specimen integrity, processing, testing, reporting, and consultation), with primary emphasis being placed on accurate, reliable, and timely diagnostic testing for good patient outcome. This approach to laboratory inspections is emphasized by HCFA or other HCFA-designated agencies for outside inspections and should also be emphasized in all in-house quality assurance programs.
Procedure Manuals
Although card files and wall charts are acceptable for quick review, a well-written and complete procedure manual is very important for the routine operation of the laboratory and mandatory for accreditation requirements (Tables 11.7 and 11.8). Instructions contained in manufacturers’ package inserts are helpful but do not fulfill the requirements for a procedure manual. It is highly recommended that the National Committee for Clinical Laboratory Standards (NCCLS; now the Clinical and Laboratory Standards Institute [CLSI]) format be followed for protocol preparation (54). This publication covers the design, preparation, maintenance, and use of technical procedure manuals for the clinical laboratory. The procedure manual should contain for each test: (i) the principle of the test; (ii) acceptable specimens (including rejection criteria) and instructions for proper specimen collection and processing; (iii) preparation of reagents and solutions; (iv) all supplies and equipment necessary to perform the test; (v) QC procedures, results, and interpretation (including corrective action if QC is out of control); (vi) the test method; (vii) possible results; (viii) the correct way to report results; (ix) procedure notes (tips); (x) limitations of the procedure; (xi) additional tables and charts; and (xii) references. Complete parasitology protocols (written according to the NCCLS guidelines) can also be found in the Clinical Microbiology Procedures Handbook (1).
Protocols are divided into three sections: preanalytic, analytic, and postanalytic. The preanalytic phase includes steps taken before the test is actually performed (pre-examination phase). Processes begin with the test request and its format, patient preparation, specimen collection, specimen transportation, specimen receipt in the laboratory, and specimen processing. The analytic phase contains all activities related to the performance of laboratory testing or examination of specimens. The postanalytic phase contains the steps that occur after a laboratory test has been performed. Processes include data review, report formatting, result interpretation and presentation, permission for release, result transmission, and sample storage.
Specimen Handling and Record Storage
All specimens must be recorded in an accession book, log book, worksheet, computer, or other comparable record. If specimens are not examined as fresh material, they should be preserved in appropriate fixatives for later examination. On the basis of current guidelines, it is recommended that specimen and test result records be kept for at least 2 years (longer for pathology records and those from the blood bank). Also, remember that when your laboratory is inspected, you must be able to retrieve those records within no more than a few hours; this is particularly important if the inspector is actually checking this parameter.
Test Request Requisitions
All requisitions should be designed to allow sufficient information for patient and physician identification and any clinically relevant information. Specific items should include (i) patient name, (ii) patient identification number, (iii) name of ordering physician, (iv) source of specimen, (v) specific procedure(s) ordered, (vi) time of specimen collection, and (vii) time of receipt or initial processing by the laboratory. Other helpful information, particularly in the parasitology division, is (i) suspected diagnosis, (ii) travel history, and (iii) recent medication. Since this information is often not on the original requisition, the physician may have to be contacted for clarification.
Procedure QC and Documentation
QC measures for all procedures are included in the individual protocols within this section of the book. Not only must the QC checks be performed as indicated, but also complete documentation should be generated, including specific plans for out-of-control results and problem resolution. Result expectations and normal results or values must be specifically defined and posted on any QC check sheets used in the laboratory. It is not sufficient to merely check a column that QC is within acceptable limits; you must also have on these sheets a definition of what normal or “acceptable limits” means. Dates and space for the initials or name of the person performing the QC checks must also be included. Specific procedure recommendations include the following (1, 6, 55, 56)
1. The examination of formed stools should include a concentration procedure.
2. A permanent stained smear is recommended for every stool specimen submitted for testing (liquid, soft, or formed).
3. A direct wet mount should be performed for fresh stool specimens (liquid or mushy—not semiformed or formed), in addition to the concentration and permanent stained smear. Since the direct wet mount is designed to allow the detection of organism motility, it is not necessary to perform the direct wet mount on preserved specimens or semiformed/formed stools. Consequently, if the stool specimens are received in preservative, proceed directly to the concentration and permanent stained smear.
4. Both thick and thin blood films are prepared for examination for malarial parasites.
5. Stained smears are washed with buffered distilled water (pH 6.8 to 7.2); any of the blood stains can be used (Giemsa, Wright/Giemsa, Field’s, rapid stains).
6. Adequate numbers of oil immersion fields should be examined (at least 300) on both stained blood films and permanent stained fecal smears.
Test Result Reports
With few exceptions, the following methods of reporting are acceptable.
1. All parasites should be reported, regardless of whether they are considered nonpathogenic or pathogenic. The presence of any parasites within the intestine generally confirms that the patient has acquired the organism through fecal-oral contamination.
2. Generally, protozoa and helminths are not quantitated on the laboratory slip. However, the specific stage (i.e., trophozoites, cysts, oocysts, spores, eggs, or larvae) is indicated.
3. Exceptions to quantitation would be Blastocystis spp. (there may be some association between numbers and symptoms) and Trichuris trichiura (light infections may not be treated).
4. The complete genus and species name of the organism identified should be reported.
5. Charcot-Leyden crystals should be reported and quantitated.
6. If the specimen is fresh or freshly preserved, budding yeast cells should be mentioned and quantitated (consultation with the physician recommended).
7. All quantitation should be consistent (Table 11.9).
8. Helminth eggs and larvae should be reported.
9. Special circumstances (additional specimens required to rule out this infection, etc.) require additional comments on the laboratory report.
10. Remember, the name of the laboratory actually performing the procedures must be indicated on the final report or elsewhere.
Reagents
All reagents should be properly labeled with (i) content, (ii) concentration, (iii) date prepared or received, (iv) lot number, (v) date placed in service, and (vi) expiration date. If reagents are prepared in-house, you should also have recorded (i) the name of person preparing the reagent and (ii) the date and results of the QC check.
The specific gravity of certain solutions (e.g., zinc sulfate) should be checked before use (this is particularly important if the solution is used infrequently). The hydrometer should have a scale large enough to differentiate between specific gravities of 1.18 for use with fresh specimens and 1.20 for use with formalinized specimens. This type of solution should be kept in a tightly stoppered bottle.
Stains should be routinely checked with control specimens for correct staining properties before use. Certainly, new lot numbers or new batches of stain should be checked. Also, if reagent use is infrequent, the stains should be checked before use. The same rule generally applies to fixatives.
With these basic recommendations in mind, the following specific QC methods can be used. While not every reagent used in diagnostic parasitology work is reviewed, these methods can be adapted to other reagents.
The examination of fecal material for ova and parasites may reveal, in addition to parasitic organisms, cells of human origin (blood cells). Most of these cells are not usually found in the stool; however, they may be found in the presence of parasites and/or other disease-producing organisms. These cells may occur in the stool without the presence of a specific etiologic agent (allergic conditions, ulcerative colitis, etc.). The buffy coat cells (white blood cells [WBCs]) can also be used as a QC check for various fixatives. The WBCs normally found in the buffy coat portion of whole blood include polymorphonuclear neutrophils, eosinophils, lymphocytes, and monocytes (macrophages). When blood is visible in the stool (indicating the presence of fresh blood), the WBC morphology is very similar to that seen on the stained thin blood film used for a differential. However, if the blood has been in the gastrointestinal tract for some time, the cellular morphology changes slightly.
QC for Schaudinn’s Fixative
1. Collect a fresh tube of blood (lavender top, EDTA anticoagulant), or get a tube from the hematology section (high WBC count, if possible).
2. Centrifuge the blood, and remove the buffy coat (layer containing WBCs).
3. Add the buffy coat to approximately 2 to 4 g of fresh, soft stool, and mix thoroughly but gently.
4. This stool-WBC mixture can be smeared onto several slides and immediately immersed in Schaudinn’s fixative.
5. After staining, if the WBCs appear well fixed and typical morphology is visible, one can assume that any intestinal protozoa placed in the same lot number of Schaudinn’s fixative would also be well fixed, provided that the fecal sample was fresh and fixed within recommended time limits.
QC for Fixative containing PVA
1. Mix approximately 2 g of fresh, soft stool with 10 ml of fixative containing PVA (in solution, ready for use). The ratio of stool to fixative should be approximately 1/3 to 1/5; do NOT add too much stool to the vial.
2. To this fixative-PVA-stool mixture, add several drops of buffy coat cells (concentrated/collected as above) and mix gently.
3. After a 30-min fixation, pour a small amount of the fixative-stool-PVA-cell mixture onto a paper towel to absorb the excess PVA.
4. The material can then be smeared onto several slides, allowed to dry thoroughly (60 min at room temperature or 30 min at 35°C), and stained. After staining, if the WBCs appear well fixed and typical morphology is visible, one can assume that any intestinal protozoa placed in the same lot number of PVA fixative would also be well fixed provided that the fecal sample was fresh and fixed within recommended time limits.
Note Entamoeba coli mature cysts are very difficult to fix properly and may be difficult to identify on the stained smear. For this reason, it is possible to have fixatives that are adequately QC checked but do not always yield good morphology for this particular organism. Use of a longer fixation time (60 min) sometimes produces better morphology after staining.
Note The same approach can be used for other stool preservatives (sodium acetate-acetic acid-formalin [SAF], zinc- and copper-based fixatives, other single-vial collection systems, and the Universal Fixative/TOTAL-FIX).
QC for Stains (Trichrome, Iron Hematoxylin, etc.)
1. Slides may be prepared from a known positive fecal specimen preserved in PVA or SAF. These slides can then be stained and checked for typical organism morphology. Positive PVA-preserved samples can be purchased commercially; many slides can be prepared from one sample. QC slides (already prepared and ready for staining) are also available commercially.
2. Buffy coat cells (after appropriate fixation) may also be used to check the staining properties of each new lot number of stain.
Note Do not prepare too many slides in advance. If dry fecal smears containing PVA are held too long before staining, the organism or buffy coat cell morphology may be poor.
Instruments and Equipment
General requirements include the following: (i) all instruments and equipment should be on a routine preventive maintenance schedule, and records should be maintained in writing; (ii) instrument manuals and maintenance/service records should be available for technical personnel to review; (iii) all thermometers should be checked and calibrated against certified standard thermometers, and the information should be recorded in writing; and (iv) checked and calibrated thermometers should be in every refrigerator, freezer, incubator, water bath, heat block, etc., and temperatures should be recorded on each day of use. As specific requirements, (i) centrifuges should be calibrated for correct speed and the calibration should be recorded on the instrument, and (ii) all microscopes should be calibrated by using a stage and ocular micrometer, the calibrations should be posted on the instrument, and recalibrations should be performed if any ocular or objective is switched on that particular microscope. It is recommended that microscopes be recalibrated once each year, even if the oculars and objectives have not been changed. This is particularly true if the microscopes receive heavy use.
Reference Materials
Reference materials should be available for comparison with unknown organisms, refresher training, and the training of additional personnel. Ideal reference materials include formalin-preserved specimens of helminth eggs, larvae, and protozoan cysts; stained fecal smears of protozoan oocysts, cysts, and trophozoites; and positive blood smears. Color slides and atlases are recommended, although the level of microscopic focus cannot be changed. Reference books and manuals from a number of publishers are available, and selected ones should be part of the parasitology library. It is also recommended that you maintain a list of consultants who can be called in case questions arise.
Patient outcome measures often refer to aspects of quality assurance that are monitored in addition to narrower parameters such as QC. Among the issues that are appropriate to monitor on an ongoing basis are the following:
1. Submission of the appropriate specimen within the correct time frame for collection
2. Condition of the specimen when received by the laboratory (too old, volume not sufficient, contains interfering substances, etc.)
3. Turnaround time for test results
4. Appropriateness of STAT requests
5. Improvements in compliance with established guidelines for the above, particularly after continuing-education efforts
6. Clinical relevance of and documented use of test results
All of these quality assurance parameters can be monitored by using The Joint Commission (formerly JCAHO) 10-step criteria for developing and implementing monitoring and evaluation activities (57; also http://www.jointcommission.org/accreditation/lab_standards_information.aspx [accessed 5/26/2015] and http://www.jointcommission.org/assets/1/18/2012_DSC_Cert_Guide.pdf [accessed 5/26/2015]). These 10 steps are listed below.
1. Assign responsibility. The designated individual identifies and assigns responsibilities to others within the department and ensures that these responsibilities are fulfilled.
2. Delineate scope of care. Delineating the scope of care involves identifying the diagnostic and therapeutic modalities used, times and locations where services are provided, types of personnel providing care, and all clinical services within the laboratory.
3. Identify important aspects of care. To effectively use the institution’s resources in quality assurance, activities selected should be those with the greatest impact on patient care (the function occurs frequently or affects large numbers of patients, serious consequences could occur if services are not provided correctly or within set time frames, and the issue has tended to cause problems for staff and/or patients in the past).
4. Identify indicators. Indicators are well-defined, objective variables that are used to monitor the quality and appropriateness of any selected aspect of patient care. Within the laboratory, indicators could include clinical situations in which it may be inappropriate to use a particular test, situations in which inappropriate tests are often used, correct sequencing of tests, and clinical outcomes that may indicate inappropriate test use for a given situation.
5. Establish thresholds for evaluation. The thresholds for evaluation must relate specifically to the indicator and establish a value below which one does not continue to monitor and above which one continues to monitor and work on improvement and reaching the threshold or moving below.
Note Some monitoring of an indicator usually occurs before a reasonable threshold can be identified. Once the threshold has been determined, monitoring and educational efforts continue, with the goal being to reach or go below the threshold. Once the threshold has been reached, consideration may be given to lowering it even more, thus leading to additional improvements in the indicator being monitored.
6. Collect and organize data. The following must be determined for each indicator: data sources, data collection method(s), sampling system, time frame for data collection, and process for comparing level of performance with set thresholds.
7. Evaluate care. If the data indicate that the acceptable threshold of performance has not been reached, extensive review occurs to identify problems or opportunities for improvement. An analysis of patterns or trends (specific shifts, units, personnel, skills) can help identify specific areas for improvement and changes. If the threshold of acceptance has been reached, it should be reevaluated in terms of keeping it where it is or possibly lowering it further to strive for additional improvements. If the decision is made to leave the threshold where it is, monitoring may be performed on a less frequent basis (every 6 months and then every year), just to verify that the data do not indicate that performance has declined.
8. Take actions to solve identified problems. The laboratory develops a plan of action that specifies who or what is expected to change, what action is appropriate, and when the changes should be complete. Three of the most common causes of problems are insufficient knowledge or poor communication, defects in the process, and poor compliance with process expectations. Frequently, educational presentations by various means (one on one, verbal presentations, written information, newsletters) can be used to help resolve the problems.
9. Assess the actions and document improvement. After sufficient time is allowed for improvements to occur, follow-up assessments are very important in documenting what progress has been made. This process must focus on the problem or opportunity for improvement, not on the action taken. If improvement is found, less frequent monitoring may be necessary; if the problem remains, new action should be taken with subsequent assessment for evidence of improvements.
10. Communicate relevant information to the organization-wide quality assurance program. It is critical that the findings not only be reviewed within the area being examined but also be conveyed and discussed at all levels of management. In addition to these discussions, documentation of such discussions is mandatory; one method often used is meeting minutes.
Continuous quality improvement (CQI) (or total quality management) is a continuous quality improvement process that evaluates processes from a customer satisfaction point of view. The aim is continuous process improvement. This approach is a natural extension and expansion of the quality assurance programs that laboratories have been using for the past few years. CQI uses familiar tools such as check sheets, run charts, and flowcharts in new ways. Issues for review in a quality improvement program include some of the issues involved in a quality assurance program, such as timeliness of response to requests, turnaround time, and effective communication. The problem-solving process in a CQI laboratory is done by broad-based teams with members from all affected groups inside and outside the laboratory. Team members are trained in and use group process methods for identifying problems and generating solutions. Rather than management remaining “top down and autocratic,” it becomes “bottom up and independent.”
CQI laboratories focus on improving customer satisfaction; however, “customers” may be defined differently from the traditional use of the word. A CQI customer can be anyone who uses the products of the production process—a ward clerk responsible for charting results, a patient needing blood drawn for preadmission tests, or a physician waiting for a STAT result. Anyone who is an “end user” of the laboratory process is a customer and should be satisfied with the laboratory’s product.
The Joint Commission uses the term “continuous quality improvement.” The concept of “quality assurance” has been changed to “quality assessment and improvement” and includes the major trends leading to quality improvement. The formal surveillance process is deemphasized in lieu of promoting activities that better reflect the principles of CQI. However, this does not suggest that the Joint Commission will abandon the 10-step process. The use of CQI requires a shift in the organization’s definition of quality from “good enough if it meets standards” to “we must work continuously to improve quality.” This whole process is similar to any recommendation for problem solving; however, there are specific differences to consider.
1. Empowerment of employees to become involved in analyzing, operating, recommending change, and implementing solutions is necessary for CQI to be adopted.
2. CQI focuses on continuous monitoring of processes, not just limited issues.
3. The problems identified almost always cross over multiple departments or units.
4. The key issue always focuses on customer satisfaction and how we define it, measure it, monitor it for success, fix it when things go wrong, etc.
5. Groups who work on these identified problems include members from all areas relevant to the final service product (from initial ordering or product to final delivery).
6. The identification of areas for improvement originates from all levels within the institution.
7. A simplified version of the above is as follows.
A. Identify and choose a process to fix.
B. Analyze all aspects of the issue.
C. Select a possible solution.
D. Correct the process.
E. Monitor the results.
F. If results are satisfactory, solidify the change and move on to other issues.
G. Who does it? A group composed of representatives of all aspects of the process under review.
Another way to look at organizing performance improvement activities can be seen in Table 11.10. This approach merges the 10-step and FOCUS-PDCA approaches; similarities are certainly more numerous than differences.
HCFA began implementing the performance-based survey process in February 1996. This is a self-survey process, and the form used is titled the Alternate Quality Assessment Survey (AQAS). The form is designed to be used in certain laboratories for recertification purposes under the CLIA program in lieu of an on-site survey. Laboratories that were surveyed under the CLIA program via the State Agencies and found to have exceptional performance, i.e., no or few minor deficiencies and satisfactory proficiency testing, are potential candidates for the AQAS. The survey form contains questions that reflect an outcome-oriented, quality improvement type of assessment. Samples of some of the areas included in the questionnaire can be seen in Table 11.11.
Following receipt of the completed form, the State Agency reviews the laboratory’s responses and supplemental documentation. On the basis of this information, the laboratory may receive a telephone call requesting more information or may receive an on-site visit. If no problems are identified, the laboratory receives recertification for a 2-year period, provided that all relevant fees are paid. The certificate is forwarded to the laboratory. No laboratory will go longer than 4 years without an on-site survey. Approximately 5% of the laboratories are visited on-site to verify the effectiveness of this self-survey process.
The performance-based survey is a reward for exceptional performance and an incentive for improvement for those laboratories that have not yet been granted permission to use this self-survey approach. General information regarding the survey process can be obtained by calling Judith Yost (Director, Center for Laboratories, Health Standards and Quality Bureau) at HCFA [(410) 786-3531] (58).
Currently, there are ongoing efforts at both the national and international levels to provide guidelines for clinical laboratories in order to upgrade their QC and quality assurance programs to a broader definition—that of quality systems. Where QC and quality assurance programs monitor performance aspects of very specific operations, a quality system applies universal quality elements throughout the entire operation of the laboratory.
The application of Lean management systems to laboratory medicine is becoming more relevant. Lean is the term used to describe a principle-based CQI management system based on the Toyota production system that has been evolving for the past 70 years. Its origins were heavily influenced by the work of W. Edwards Deming and the scientific method that forms the basis of most quality management systems. Lean has two fundamental components: a systematic approach to process improvement by removing waste in order to maximize value for the end-user of the service and a commitment to respect, challenge, and develop the people who work within the system to create a culture of continuous improvement. Lean principles have been applied to many health care systems throughout the world to improve the quality and cost-effectiveness of services for patients. Also, a number of clinical/pathology laboratories have used Lean to shorten turnaround times, improve quality (reduce errors), and improve productivity. Increasingly, models designed to implement large-scale change in health care systems have evidence-based improvement methodologies (such as Lean CQI) as a core component (59).
ISO (International Organization for Standardization) is the world’s largest developer of standards. Since 2000, the worldwide standard for a quality system in business and industry has been the ISO 9001 guidelines (the fourth edition is published by the ISO). Countries trading products across international boundaries have found the standardized requirements very beneficial in improving quality; this approach has also reduced the time and cost of multiple inspections due to the international nature of their business. In the future, there will be a universal standard for quality management in medical laboratories: ISO 15189 (60, 61). ISO 9001 Quality Management Systems is under review and an updated version is expected by the end of 2015. All ISO standards are reviewed every 5 years to make sure they remain as helpful and relevant (http://www.iso.org, accessed 4/3/2015).
When the large majority of products or services in a particular business or industry sector conform to International Standards, a state of industry-wide standardization can be said to exist. This standardization is achieved through consensus agreements between national delegations representing all the economic stakeholders concerned: suppliers, users, government regulators, and other interest groups, such as consumers. They agree on specifications and criteria to be applied consistently in the classification of materials, in the manufacture and supply of products, in testing and analysis, in terminology, and in the provision of services.
According to their objectives, laboratories will choose a recognition of their quality management system with an ISO 9001 certification or a recognition extended to the technical skills with an ISO 17025 or ISO 15189 accreditation. The contents of these last two documents are very similar, and both integrate requirements of the standard ISO 9001. The standard ISO 17025 is somewhat distant from clinical laboratory needs, requiring many efforts of adaptation, just like the ISO 9001 standard. The standard ISO 15189 seems to be well adapted but more constraining, considering the detailed requirements required. It necessitates a perfect control of the preanalytical phase, which is difficult to acquire in a clinical framework where the clinical specimens are not taken by the laboratory staff.
A number of advantages are associated with the use of ISO. The focus on patients has been reestablished, all processes are identified and subject to continuous improvement, and performance measurements provide an integrated picture of results. Measurements subsequently lead to improvement of quality of care and to quality system improvements. The documentation system can serve the organization’s needs without leading to bureaucracy. Given the need for adequate quality management tools in health care and the need to demonstrate quality, the positive effects suggest that ISO will become more prevalent in health care organizations (62).
ISO provides technical and quality management, which results in benefits observed in daily laboratory practices. Technical requirements can include the addition of formal personnel training plans and detailed records, method development and validation procedures, measurement of method uncertainty, and a defined equipment calibration and maintenance program. Also, an expanded definition of the sample preparation process can be implemented and documented to maintain consistency in sampling. Management quality improvements often emphasize document control to maintain consistent analytical processes, improved monitoring of supplier performance, a periodic contract review process for documenting customer requirements, and a system for handling customer comments and complaints. Continuous improvement is monitored and improved through corrective and preventive action procedures and audits. Quarterly management review of corrective actions, nonconforming testing, and proficiency testing results also identifies more long-term trends. The practical benefits of these technical and management quality improvements can be seen on a daily basis in the laboratory. Faster identification and resolution of issues regarding methods, personnel or equipment, improved customer satisfaction, and overall increased laboratory business are all the result of implementing an effective quality system. Certainly, the ISO system provides one option among various quality improvement approaches (63).
In the 1990s, blood centers and transfusion services within the United States began adopting the concepts of quality systems into their routine operations, primarily based on direction provided by a 1993 FDAQA guideline (64). After a series of revisions, the American Association of Blood Banks (AABB) published its 10 Quality System Essentials (QSEs) (65). These have been incorporated into the most recent edition of the AABB Standards for Blood Banks and Transfusion Services (66).
It became very obvious that the QSEs proposed for the blood bank were applicable to the entire laboratory operation. These essentials also incorporated many of the quality requirements developed by CLIA ’88, The Joint Commission (formerly JCAHO), and the College of American Pathologists (CAP).
Recognizing that standardization would be not only valuable, but also essential for high-quality laboratory operations as a whole, the Clinical and Laboratory Standards Institute (CLSI) (formerly known as the National Committee for Clinical Laboratory Standards [NCCLS]) developed a subcommittee to prepare a guideline encompassing a complete quality system for the clinical laboratory. This document was recently published as QMS01-A4 (GP26-A4), Quality Management System: A Model for Laboratory Services; Approved Guideline—Fourth Edition (67).
Factors Influencing Future Laboratory Practice
The current atmosphere of uncertainty within the health care environment in the United States in the second decade of the 21st century is well recognized. Funding and support for health care are diminishing, and the economic future of health care is hostage to a number of identifiable trends and concerns (Table 11.12).
As consolidation among payers continues through mergers, acquisitions, and other arrangements, providers of care find themselves with fewer buyers of care with whom to negotiate. This requires the development of strategies through which provider leverage can be maintained. Consolidation on the provider side has been one response. Price reductions and exclusive arrangements have been others. Response to these changes also includes affiliation with community providers to increase bargaining strength and aggressive cost reduction to allow more competitive bidding. However, many institutions are now facing the acceptance of marked reductions in reimbursement rates or terminating contracts. As additional changes occur and negotiations continue on these issues, each institution much achieve a balance between the market share realized from each contract, the revenue per service unit, and the total revenue.
Marketplace trends include continued growth in managed care, particularly in the health maintenance organization sector; integration of payers, providers, suppliers, and buyers; mergers and alliances; integration between hospitals and physicians; continued pressure for provider consolidation to address excess bed capacity; continued shift from inpatient to outpatient procedures; and provider alliances based on complementing services (linkage of primary-care facility with the tertiary or quaternary expertise of another institution).
Strategies for survival will include (i) increased ability to capture market share, preferably related to covered lives rather than patient days; (ii) improved information systems to accurately identify costs (fixed, variable by service, procedure) and track the profitability of contracts; and (iii) recognition of the importance of these issues and implementation of proactive measures to structure the organization for survival in the coming years.
With the increase in health care contracts and overall health care reform, a number of potential issues will affect the laboratory of the future. Each institution is actively seeking new markets, with the overall objective being to increase the patient base served by that particular institution. As these contracts increase in number, the reimbursement approach becomes one of capitated payment; the provider takes on the risk previously assumed by the third-party payer. The provider receives a certain amount for each patient per year, the assumption being that the care can be delivered at less than the reimbursement capitated amount; thus, a profit is realized. As these changes occur, there continue to be mergers and the development of large care networks. However, as more large networks are developed, there are fewer contracts on which to bid. Each institution may be forced to consider contracts that pay less than the current costs of service. Obviously, these changes serve as a tremendous incentive to deliver care at continued reduced costs. Once contracts are lost to other bidders, it may be more difficult to recapture that lost market.
Managed care is changing from a cost containment environment to a value purchasing environment. Considerations include careful examination of what is being received for the money spent, increases in requests for outcome data, and case management options. As competitiveness in the managed-care arena increases, long-standing partnerships between institutions and health maintenance organizations will have no inherent merit in guaranteeing continued partnerships. More focused medical management, lower costs, and higher efficiency will determine the continuation or elimination of present agreements. The new environment will demand increased cohesiveness, cooperation, and creativity on the part of all employees within an institution.
An insurance, payment, and delivery system for health care, loosely described as “managed care,” is increasingly viewed as dysfunctional by consumers (i.e., patients), providers, and employers, the last of which constitute the primary source of funding for health insurance for the employed sector of the population. In the current environment of managed-care dominance within the United States, clinical laboratories continue to change their focus of operations and mission in response to the continually changing landscape. Traditional laboratories that are unwilling to change and adapt to this environment will probably not survive. A synopsis of changes since the early 1990s is provided in Table 11.13.
There appears to be a climate of blame in which the various players in the health care arena reproach each other and advance conflicting solutions. Although the “demise” or significant modification of the managed-care system is widely predicted, there is a distinct absence of any consensus about the parameters of politically or economically viable alternatives. Nonetheless, it appears that “something else” will emerge—a return of the “managed-competition” plan proposed in the early years of the Clinton presidency, a national health model resembling the Canadian system, implicit and explicit rationing of resources, or other combinations and approaches. The Affordable Care Act, which was passed during the first term of the Obama administration, contains provisions that radically alter all aspects of health care delivery and financing. However, implementation has only begun, and it is clear, that whatever the outcome of the Affordable Care Act, decreased financial resources available for health care will continue.
Capitated reimbursement has already been mentioned as one of the driving forces behind the necessity to deliver health care at continuing reduced rates. Each year, reimbursement rates decline; currently, leverage lies with the payer, not the provider. Competitive bidding has become an absolute necessity. Also, institutions continue to review their costs in much greater detail than at any time in the past. To survive, each institution must know exact costs so that it can bid appropriately and remain financially viable. Complicating this approach are the various regulatory requirements and the fact that each contract is somewhat different in terms of services and costs. Laboratories are continually reviewing their test menus, methodologies, etc., for the most clinically relevant, cost-effective approach to diagnostic testing.
Current problems related to health care costs include a growing federal deficit and concerns about the future viability of Medicare, the increasing inability of state budgets to fund rising Medicaid expenditures, an aging population that will generate increasing demands on health care resources, the proliferation of costly technological and pharmacological advances in the ability to diagnose and treat disease, and a health care insurance and delivery system unable to provide insurance and predictable access to care for approximately 40 million Americans (3).
Immediately prior to the 21st century, Congress began to focus on legislation and regulation in health care, perhaps the two most striking examples of which are the Clinical Laboratory Improvement Amendments of 1988 (CLIA) and the Health Insurance Portability and Accountability Act of 1996 (HIPAA). The four main regulations that are relevant to pathology and laboratory medicine are CLIA, HIPAA, the Occupational Safety and Health Administration (OSHA) standards for occupational exposure to blood-borne pathogens, and the Stark regulations that prohibit self-referral by health care providers.
CLIA ’88
The Clinical Laboratory Improvement Amendments were enacted in 1988; however, the regulations were not implemented until 1992. Although changes to the regulations continue to be made, the amendments in their current form are well established and are applicable to all testing performed on humans. Specific regulations include personnel standards, patient test management, quality assurance, proficiency testing, provisions for inspection and certification, and both civil and criminal penalties for noncompliance. Inspection and accreditation options are multiple and include government and private-sector organizations such as the College of American Pathologists and the Joint Commission.
HIPAA
The Health Insurance Portability and Accountability Act was passed in 1996. The Privacy Rule standards address the use and disclosure of individuals’ health information (protected health information) by organizations subject to the Privacy Rule (covered entities) as well as standards for individuals’ privacy rights to understand and control how their health information is used. Within the U.S. Department of Health and Human Services, the Office for Civil Rights (OCR) has responsibility for implementing and enforcing the Privacy Rule with respect to voluntary compliance activities and civil money penalties (http://www.hhs.gov/ocr/privacy/hipaa/understanding/summary/index.html; accessed 4/3/2015).
A major goal of the Privacy Rule is to assure that individuals’ health information is properly protected while allowing the flow of health information needed to provide and promote high quality health care and to protect the public’s health. The Rule strikes a balance that permits important uses of information, while protecting the privacy of people who seek care and healing. Given that the health care marketplace is diverse, the Rule is designed to be flexible and comprehensive to cover the variety of uses and disclosures that need to be addressed.
“Individually identifiable health information” is information, including demographic data, that relates to and that identifies the individual or for which there is a reasonable basis to believe it can be used to identify the individual:
• the individual’s past, present, or future physical or mental health or condition,
• the provision of health care to the individual, or
• the past, present, or future payment for the provision of health care to the individual.
OSHA Standards for Occupational Exposure to Blood-Borne Pathogens
The standards for exposure to blood-borne pathogens were developed in 1991 by OSHA of the Department of Labor in response to widespread concerns by unions representing health and hospital workers concerning the inadequacy of protective practices, equipment, and other safeguards for employees with regular and predictable exposure to body fluids and substances with the potential to transmit HIV, hepatitis B virus, and hepatitis C virus. More complete information can be found at http://www.osha.gov/SLTC/bloodbornepathogens/index.html (accessed 4/3/2015).
In order to reduce or eliminate the hazards of occupational exposure to blood-borne pathogens, an employer must implement an exposure control plan for the worksite with details on employee protection measures. The plan must also describe how an employer will use a combination of engineering and work practice controls, ensure the use of personal protective clothing and equipment, and provide training, medical surveillance, hepatitis B vaccinations, and signs and labels, among other provisions. Engineering controls are the primary means of eliminating or minimizing employee exposure and include the use of safer medical devices, such as needleless devices, shielded needle devices, and plastic capillary tubes.
The Stark Laws
The Stark regulations were enacted sequentially over a period of several years and severely limit referral of a wide variety of services (including laboratory testing) to entities in which the referring provider has a financial stake. The regulations are named for Rep. Fortney “Pete” Stark (D.-CA) who was the legislative sponsor of the initial regulations enacted in 1992 (Stark I) that prohibited a physician from making Medicare referrals for clinical laboratory services if the physician or an immediate family member had any financial relationship with the laboratory. In 1993, the Stark II legislation extended the regulations to Medicaid and broadened coverage to 10 additional health services, including, among others, occupational and physical therapy, durable medical equipment, radiology, and radiation therapy. However, the regulations are very complex and difficult to interpret. Enforcement by the government, particularly the Justice Department, is closely linked to other “fraud and abuse” enforcements under a variety of federal anti-kickback and “false claims” statutes (68).
In the last few years, many inpatient procedures have been shifted to the outpatient setting in response to payer per diem cost containment initiatives. Most institutions are actively involved in developing or expanding outreach programs, including laboratory services, teaching, and consultations. Another area that will continue to expand is point-of-care testing (POCT), or testing at the site of the patient (bedside, emergency room, or clinic), or alternative-site testing (home or shopping mall). With this expansion comes new technology that is advertised as “fail-safe and foolproof,” some of which may be true and some of which may be exaggerated. Obviously, this approach is directly linked to patient and overall customer satisfaction related to a short turnaround time for test results. Since more and more patient care is being delivered on an outpatient basis as decentralized care, reductions in length of hospital stay and in cost continue to be issues for review.
In some institutions that service very specific patient groups, POCT may offer distinct advantages; however, in institutions with highly efficient specimen transport systems and rapid response capabilities within the main clinical laboratory, advantages of POCT may be minimal to none. Continued review of regulatory requirements, QC issues, proper and consistent documentation, proficiency testing, performance enhancement, and cost-effectiveness is mandatory for success. As technology moves toward microcomputerization, microchemistry, chip technology, and enhanced test menus, the use of POCT will require ongoing scrutiny (69, 70).
Although the push for decentralized testing may be coming from a variety of sources, it is generally thought that the central laboratory has become more involved in managing the new testing technologies and trying to circumvent anticipated problems with testing quality and patient care.
There are real, continued concerns about test accuracy, precision, training, and QC issues; personnel training and experience; and proficiency testing by those actually performing the testing. To date, there is little scientific evidence for cost reduction and/or reduction in length of stay. There appears to be mounting evidence that in some settings, the cost is higher than that of central laboratory testing.
Large, integrated testing laboratories may become “the laboratory of the future,” particularly considering the issue of economy of scale; i.e., the more we perform large numbers of the same test, theoretically the lower the cost. Although this approach is more applicable to some areas of the laboratory (chemistry), it is being carefully evaluated even in labor-intensive areas with less automation (microbiology). As technology continues to change, the departmental limits within any laboratory (chemistry, hematology, microbiology, or blood bank) continue to become less well defined. Many laboratory operations are now structured around automated and manual methods rather than specific clinical disciplines. This approach will continue and will also affect personnel training and utilization. Review of in-house versus send-out tests will continue, particularly regarding cost containment parameters.
With continued development and implementation of computerized history algorithms, clinical pathways, and case management, more structured support for clinical decision making will also dramatically affect the laboratory. Inappropriate test-ordering patterns and overutilization will become less and less significant. The use of algorithms may also become much more common within the laboratory itself. All of these changes will be developed through the use of multidisciplinary teams, including all members of the health care delivery system (ancillary departments, nursing staff, and physicians).
New technology will continue to be evaluated for a number of parameters, including type of data, technical characteristics, diagnostic characteristics, clinical utility, and cost-benefit analysis. Laboratory design will be another key issue and will be influenced by a number of factors, including consolidation of health care facilities; continued shift to outpatient care and POCT and alternative-site testing; development and use of clinical paths; and development and evaluation of new technologies. Physical plant design should consider a number of factors, such as analysis of institution mission, scope of the project, and specific objectives; written statements of needs; mechanisms for keeping people informed; careful analysis of operational needs (work flow, traffic patterns, etc.); detailed review of current and future space requirements (consolidation of instrumentation, workstations, storage capacity, and equipment location); flexibility of design (modular furniture and utilities); and safety issues.
Currently, hospitals and clinics are not able to meet their expenses without continued decreases in labor costs. During the next few years, the greatest savings in laboratory costs will come from technology that leads to labor reductions. Laboratory automation has the potential to be a highly successful, cost-saving measure and has been implemented in Japan, where many laboratories employ 5 to 10 times fewer full-time equivalents than European and North American laboratories while still achieving similar productivity results. Future expectations include the performance of all but a few esoteric tests by automated systems, the use of mass spectrometry, and performance of most routine testing on a single automated analyzer. It is very likely that specimens will arrive in the laboratory ready for testing, with all processing having been done prior to receipt by the laboratory. The specimen may contain information chips carrying patient medical history and physiologic data at the time of sampling. These chips may also allow wireless encoding and reading of specimen information. Thus, the old specimen-processing area can concentrate on customer service, provision of medical information, and client education.
As an adjunct to the highly automated core laboratory, miniature analyzers and POCT will play a major role in testing. Multianalyte, spectroscopy-based, noninvasive sensors will provide a wide range of tests at the bedside; there is even the possibility of a wireless invasive analyzer that would reside in a device that could be injected or implanted under the skin or elsewhere in the body.
New molecular tools have been developed rapidly, especially with the excitement over the completion of the human genome project. Synthetic antibody-like molecules with high affinity and specificity have been developed from both DNA and RNA. These molecules may replace antibodies in molecular diagnostics, achieving tremendous levels of sensitivity.
Biometrics is the use of sensors to measure natural body features for positive identification. In the future, biometrics will be extremely helpful in confirming that patients and their body fluids or other specimens are correctly identified. A patient’s complete medical history may be available on the Web when the patient simply places a finger into the sensor mechanism.
Nonlaboratory systems, such as robots, will be used much more commonly for delivery functions, saving a significant amount of money as well as improving service. Surgery robots that can perform complex procedures through very small incisions are currently under development, and some have been implemented.
An area of research and laboratory practice that will be critically impacted by complex and interrelated legislative, regulatory, and administrative developments is stem cell research and the application of cloning technology to medical practice in tissue and organ transplantation and reproductive technology. The cloning research issue is closely linked to the highly emotional controversy over abortion and genetic manipulation and has elicited strong and conflicting opinions from various constituencies, including the research and medical community, religious groups, and organizations committed to research and treatment of specific diseases.
A number of approaches are being developed and used to serve as “practice guidelines”—more structured approaches for both information gathering and clinical decision making related to the patient. A clinical pathway is defined as a set of interventions or actions that is used to help a patient move progressively through a clinical experience to an expected, positive outcome. One of the main reasons for developing clinical pathways is to provide a tool to evaluate clinical patient management and outcomes. Once defined sets of expectations are developed, data can be used to monitor variances. The cost of care and utilization of resources can then be monitored more closely.
By using clinical pathways for predictable clinical outcomes (conditions, diseases, and procedures selected are generally very well established and patient outcomes are consistent), many also believe that patient care is more consistent. These clinical pathways can also be used as a teaching tool for all health care personnel. The development of clinical paths must be a multidisciplinary effort, with input from all relevant members of the health care team. Many institutions are also using the process of clinical path development to foster team building and continuous quality improvement initiatives.
Regardless of names or titles, the following are becoming more widely used and accepted. They are listed from the most well-defined and predictable medical situations (clinical pathway) to very complex cases with many underlying factors and complications (case management).
1. Clinical pathways (care maps) (used for conditions, diseases, and procedures for which the patient outcome is very predictable)
2. Clinical guidelines (used for care situations in which the outcome may have a few more variables)
3. Clinical algorithms (used for patient outcome situations in which more options may be required)
4. Case management (generally used for very complex cases, for which it is very difficult to establish a predetermined pathway; the patient may have multiple medical problems and/or complications)
Many personnel issues will continue to be relevant and important. Particular emphasis will probably include (i) the use of unlicensed individuals in relationship to CLIA ’88 and various states developing, implementing, and revising personnel regulations; (ii) cross-training and the use of multiskilled laboratory personnel; (iii) tremendous flexibility in covering fluctuations in workload; (iv) expansion of the use of multidisciplinary teams; (v) continued emphasis on horizontal rather than vertical management structures; and (vi) development and utilization of personnel competency assessment programs (now mandated by regulations).
The aging of the population will have a profound effect on many health care issues. Laboratory testing and monitoring will be no exception. Patient access, alternative-site testing, and clinical decision support options will play a large role in helping manage this expanding population.
Not only do we have to deal with expanding diseases and conditions in the elderly, but also we now are faced with some serious problems related to infectious-disease threats. There is explosive population growth worldwide with expanding poverty and urban migration; international travel is increasing; and technology is rapidly changing. All of these factors affect our risk of exposure to the infectious agents in our environment. In recent years, our antimicrobial drugs have become less effective against many infectious agents, and experts in infectious diseases are concerned about the possibility of a “postantibiotic era.” As a result, our ability to detect, contain, and prevent emerging infectious diseases is in jeopardy.
In the wake of the events of September 11, 2001, credible threats of terrorism that include the potential use of nuclear, chemical, and biological agents have had significant impacts on the funding and costs of providing laboratory services. The identification of clinical cases and environmental contamination caused by deliberate dissemination of Bacillus anthracis through the mail system during the early part of 2002 emphasized this lesson for a number of microbiology laboratories. These events and the fear of additional terroristic use of other infectious agents have not only alarmed the public, the government, and the medical and microbiological community but also focused attention on technical capabilities, biologic safety infrastructure, and staff preparation (71, 72).
Unfunded programs will impact training costs, acquisition and deployment of equipment, facility modification, and other unfunded mandates, such as the CDC-issued rules (August 2002) requiring all laboratories to report to the government whether they are in possession of “select agents” that could be used in a bioterrorist attack. The rule is available at http://www.selectagents.gov/regulations.html (accessed 5/26/2015) (4). However, increased federal concerns about laboratory preparedness have generated some increased funding for specific areas of the laboratory; it is more likely that in the event of actual or suspected biological attacks, the impact on the laboratory will be to significantly increase laboratory services. These increases will focus on unusual/potentially dangerous diagnostic challenges and on the potential massive increase in demand for routine laboratory services required for the care of large numbers of acutely ill patients.
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