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Sputum, induced sputum and bronchoalveolar lavage (BAL) for Pneumocystis jiroveci
Sputum, induced sputum, and bronchoalveolar lavage (BAL) material are commonly used for diagnosing Pneumocystis jiroveci (previously classified as Pneumocystis carinii) infections.  To examine these specimens for cysts of P. jiroveci, prepare smears of the sediment and examine after staining with the recommended procedure (Giemsa or methenamine silver stain).

Staining Procedures
Stain only one set of smears and leave the duplicates unstained.  The latter will prove useful if a problem occurs during the staining and/or if you wish later to send the smears to a reference laboratory.

Preparation of the specimen
Refer to safety section prior to handling the specimens (link)

1. Treat specimens containing mucus (sputum mainly) with a mucolytic agent (e.g., Sputalysin) by mixing the agent in a 1:1 ratio with the specimen.
2. Centrifuge at 500 × g for 5 minutes.
3. Discard supernatant.
4. Treat sediment containing blood with red cell lytic agent (e.g., saponin*).  If this is the case, centrifuge once more following conditions described above and discard the supernatant.
5. Resuspend the remaining sediment.
6. Place drops of the sediment on the center of a microscopic slide.*
7. Spread the drop on the slide with a pipette to evenly distribute on the slide.
8. Air dry and stain following the required procedure.

*For specimens containing blood, separate an aliquot that has not been subjected to lyses and prepare smears from that as well.

Giemsa Stain

1. Stock 100× Giemsa Buffer	0.67 M
Na2HPO4	59.24 g
NaH2PO4H2O	36.38 g
Deionized water	1000.00 ml
Autoclave or filter-sterilize (0.2 µm pore).  Sterile buffer is stable at room temperature for one year.

2. Working Giemsa Buffer	0.0067 M, pH 7.2
Stock Giemsa Buffer	10.0 ml
Deionized water	990.0 ml
Check pH before use. Should be 7.2.  Stable at room temperature for one month.

3. Triton X-100 5%
Deionized water (warmed to 56°C)	95.0 ml
Triton X-100	5.0 ml
Prewarm the deionized water and slowly add the Triton X-100, swirling to mix.

4. Stock Giemsa stain
Giemsa stain is available commercially, but the following formulation gives more constant results and does not expire

1. Put into a 500 ml brown bottle the glass beads and the other ingredients, in the order listed.  Screw cap tightly.  Use glassware that is clean and dry.
2. Place the bottles at an angle on a shaker; shake moderately for 30 to 60 minutes daily, for at least 14 days.
3. Kept tightly stoppered and free of moisture, stock Giemsa stain is stable at room temperature indefinitely (stock stain improves with age).
4. Just before use, shake the bottle.  Filter a small amount of this stock stain through Whatman #1 filter paper into a test tube.  Pipet from this tube to prepare the working Giemsa stain.

5. Working Giemsa stain (2.5%): make fresh for each batch of smears

Staining

1. Prepare fresh working Giemsa stain in a staining jar, according to the directions above.  (The 40 ml fills adequately a standing Coplin jar; for other size jars, adapt volume but do not change proportions).
2. Pour 40 ml of working Giemsa buffer into a second staining jar.  Add 2 drops of Triton X-100.  Adapt volume to jar size.
3. Place slides into the working Giemsa stain (2.5%) for 45-60 minutes.
4. Remove thin smear slides and rinse by dipping 3-4 times in the Giemsa buffer.  Thick smears should be left in buffer for 5 minutes.
5. Dry the slides upright in a rack.

Note: As alternates to this 45-60 minutes in 2.5% Giemsa stain, the smears could be stained for shorter times in more concentrated stains; one alternate is 10 minutes in 10% Giemsa; the shorter stains yield faster results, but use more stain and might be of less predictable quality.

Staining Procedure: Quality Control
To ensure that proper staining results have been achieved, a positive smear (malaria) should be included with each new batch of working Giemsa stain.

Methenamine silver stain (Gomori’s silver methenamine)
Store reagents at room temperature (25°C) except the silver nitrate, which must be stored at 4°C

1. CrO3 10% solution (stable for 1 year)

PROCEDURE:

1. Add chromic acid 10% solution to cover the smears. Incubate for 10 minutes at room temperature.
2. Prepare working methanine solution by mixing 20 ml of 3% methanine, 1 ml of 5% silver nitrate, 1.5 ml of 5% sodium borate, and 17 ml of distilled deionized H2O.
3. Wash slide with distilled H2O.
4. Cover slides with 1% sodium bisulfate.  Incubate for 1 minute at room temperature.
5. Wash slides with distilled H2O.
6. Place slides in a Coplin jar containing the working methanine solution.  Cover with a cap and place in a microwave oven.
7. Microwave it at 50% power for 35 seconds.  Gently rotate the jar and microwave again for 35 seconds.  Leave the slides in the hot methanine solution for 1-3 minutes.  Microwave on 600 Watts for 35 seconds.  If a microwave oven is not available, use a water bath heated at 80°C.  Prewarm the solution by putting the Coplin jar in the bath in the bath for 6 minutes, prior to adding the slides.  Add the slides and incubate in the water bath for additional 5 minutes.
8. Wash the slides with distilled and then with deinonized H2O.
9. Dip the slides several times in the in the 1% gold chloride solution.
10. Place the slides on a rack, cover with 5% sodium thiosulfate solution and incubate for 1 minute at room temperature.
11. Wash the slides with distilled H2O.
12. Cover the slides with the working light green solution (counterstain) and incubate for 1 minute at room temperature.
13. Wash the slides with distilled H2O. Drain the excess of water and air dry.
14. Examine the slides.

For additional information, call the Division of Parasitic Diseases, at 770-488-4474.