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.
