and reports excellent results (27–30). There are several FDA-approved molecular diagnostic tests for this organism. They include the APTIMA Trichomonas vaginalis Assay (GenProbe, San Diego, CA). This assay utilizes the same amplification technology that has been used for years to detect Neisseria gonorrheae and Chlamydia trachomatis. It also uses the same specimen types (i.e., clinician-collected vaginal and endocervical swabs, female urine, and PreservCyt solution). It has reported high sensitivity and specificity, and does not require organism viability or motility. It, as well as other antigen-based assays, represents advances in the detection of this parasite.
The Affrirm VPIII DNA probe technology (Becton, Dickinson and Co., Franklin Lakes, NJ) offers a dependable, rapid means for the early identification of three organisms causing vaginitis: Candida spp., Gardnerella vaginalis, and T. vaginalis. During the test protocol, the assay aligns complementary nucleic acid strands to form specific, double-stranded complexes called hybrids. For each organism, this hybrid is composed of capture and color development single-stranded nucleic acid probes, complementary to the released target nucleic acid analyte. Enzyme conjugate then binds to the captured analyte.
Examination of urinary sediment is indicated in certain filarial infections. The occurrence of microfilariae in urine has been reported with increasing frequency in Onchocerca volvulus infections in Africa. The triple-concentration technique is recommended for the recovery of microfilariae (13) (see chapter 7).
Urine is collected into a bottle, the volume is recorded, and thimerosal (1 ml/100 ml of urine) is added. The specimen is placed in a funnel fitted with tubing and a clamp; this preparation is allowed to settle overnight. The following day, 10 to 20 ml of urine is withdrawn and centrifuged. The supernatant fluid is discarded, and the sediment is resuspended in 0.85% NaCl. This preparation is again centrifuged, and 0.5 to 1.0 ml of the sediment is examined under the microscope for the presence of nonmotile microfilariae. The membrane filtration technique used with blood can also be used with urine for the recovery of microfilariae. Administration of the drug diethylcarbamazine (Hetrazan) has been reported to enhance the recovery of microfilariae from the urine (31).
A membrane filter technique for the recovery of Schistosoma haematobium eggs is also useful (32) (Fig. 5.9 and 5.10).
Figure 5.9 Membrane filtration system. (Upper) Membrane holder, which can be attached to a syringe for filtration of various types of specimens, particularly urine. (Lower) Package of membranes; different sizes with various mesh sizes are available, depending on the clinical specimen and suspected organism size. doi:10.1128/9781555819002.ch5.f9
Figure 5.10 Schistosoma haematobium eggs from a urine filtration. Note the terminal spines on the eggs. Since the specimens were not preserved prior to filtration, determination of the viability of the eggs is possible. The viability information should be conveyed to the physician as a part of the report. doi:10.1128/9781555819002.ch5.f10
Membrane Filter Technique
1. Collect a urine sample in a container (urine should be well mixed before step 2).
2. With a syringe, draw up 1 ml of urine into the syringe barrel.
3. Fill the rest of the barrel volume with air.
4. Attach the filter holder containing an 8-mm-pore-size Nuclepore membrane filter to the syringe.
5. Express the urine through the filter.
6. Remove the filter, and place it on a microscope slide face down.
7. Moisten the filter with 0.85% NaCl.
8. Examine the filter under ×100 power for the presence of eggs.
The efficiency of the polycarbonate membrane filtration technique for detecting S. haematobium eggs in urine is increased by using a pore size of 14 µm and the suction of a water jet pump. Egg concentrations of 1 egg in >1,000 ml of urine can be detected. Viability can be assessed after filtration by staining with trypan blue. This technique is highly recommended in light infections with small numbers of eggs in which previous diagnostic methods have not confirmed the infection (33).
References
1. al-Rufaie HK, Rix GH, Perez Clemente MP, al-Shawaf T. 1998. Pinworms and postmenopausal bleeding. J Clin Pathol 51:401–402. PMID 9708211
2. Arora VK, Singh N, Chaturvedi S, Bhatia A. 1997. Fine needle aspiration diagnosis of a subcutaneous abscess from Enterobius vermicularis infestation. A case report. Acta Cytol 41:1845–1847. PMID 9390155
3. Avolio L, Avoltini V, Ceffa F, Bragheri R. 1998. Perianal granuloma caused by Enterobius vermicularis: report of a new observation and review of the literature. J Pediatr 132:1055–1056. PMID 9627606
4. Maki AC, Combs B, McClure B, Slack P, Matheson P, Wiesenauer C. 2012. Enterobius vermicularis: a cause of acute appendicitis in children. Am Surg 78:E523–524. PMID 23265108
5. Erhan Y, Zekioglu O, Ozdemir N, Sen S. 2000. Unilateral salpingitis due to Enterobius vermicularis. Int J Gynecol Pathol 19:188–189. PMID 10782420
6. Ok YZ, Ertan P, Limoncu E, Ece A, Ozbakkaloglu B. 1999. Relationship between pinworm and urinary tract infections in young girls. APMIS 107:474–476. PMID 10335951
7. Reipen J, Becker C, William M, Hennerlein B, Friedrich M, Salehin D. 2012. Peritoneal enterobiasis causing endometriosis-like symptoms. Clin Exp Obstet Gynecol 39:379–381. PMID 23157050
8. Garcia LS (ed). 2010. Clinical Microbiology Procedures Handbook, 3rd ed. ASM Press, Washington, DC.
9. Liu LX, Chi J, Upton MP, Ash LR. 1995. Eosinophilic colitis associated with larvae of the pinworm Enterobius vermicularis. Lancet 346:410–412. PMID 7623572
10. Brooke MM, Melvin D. 1969. Morphology of Diagnostic Stages of Intestinal Parasites of Man. US Department of Health, Education, and Welfare publication (HSM) 72-8116. US Government Printing Office, Washington, DC.
11.
