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
