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The use of molecular techniques for the diagnosis and epidemiologic study of sexually transmitted infections

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Abstract

Molecular diagnostic tests are more sensitive and, in many cases, more specific than conventional laboratory methods for the detection of sexually transmitted infections. Here, we review recently developed molecular methods for the diagnosis and subtyping of the most common sexually transmitted infections: infections caused by Chlamydia trachomatis, Neisseria gonorrhoeae, human papillomavirus, Trichomonas vaginalis, and the agents of genital ulcer disease (Haemophilus ducreyi, herpes simplex virus, Treponema pallidum, and Calymmatobacterium granulomatis). We also provide an overview of the laboratory diagnostic tests and clinical specimens to use when infection with these agents is suspected.

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References and Recommended Reading

  1. Davis JD, Riley PK, Peters CW, Rand KH: A comparison of ligase chain reaction to polymerase chain reaction in the detection of Chlamydia trachomatis endocervical infections. Infect Dis Obstet Gynecol 1998, 6:57–60.

    Article  CAS  PubMed  Google Scholar 

  2. Pasternack R, Vuorinen P, Pitkajarvi T, et al.: Comparison of manual Amplicor PCR, Cobas Amplicor PCR, and LCx assays for detection of Chlamydia trachomatis infection in women by using urine specimens. J Clin Microbiol 1997, 35:402–405.

    CAS  PubMed  Google Scholar 

  3. Puolakkainen M, Hiltunen-Back E, Reunala T, et al.: Comparison of performances of two commercially available tests, a PCR assay and ligase chain reaction test, in detection of urogenital Chlamydia trachomatis infection. J Clin Microbiol 1998, 36:1489–1493.

    CAS  PubMed  Google Scholar 

  4. Goessens WH, Mouton JW, van der Meijden WI, et al.: Comparison of three commercially available amplification assays, AMP CT, LCx, and COBAS AMPLICOR, for detection of Chlamydia trachomatis in first-void urine. J Clin Microbiol 1997, 35:2628–2633.

    CAS  PubMed  Google Scholar 

  5. Steingrimsson O, Jonsdottir K, Olafsson JH, et al.: Comparison of Roche Cobas Amplicor and Abbott LCx for the rapid detection of Chlamydia trachomatis in specimens from highrisk patients. Sex Transm Dis 1998, 25:44–48.

    Article  CAS  PubMed  Google Scholar 

  6. Chernesky MA, Chong S, Jang D, et al.: Ability of commercial ligase chain reaction and PCR assays to diagnose Chlamydia trachomatis infections in men by testing first-void urine. J Clin Microbiol 1997, 35:982–984.

    CAS  PubMed  Google Scholar 

  7. Stary A, Schuh E, Kerschbaumer M, et al.: Performance of transcription-mediated amplification and ligase chain reaction assays for detection of chlamydial infection in urogenital samples obtained by invasive and noninvasive methods. J Clin Microbiol 1998, 36:2666–2670.

    CAS  PubMed  Google Scholar 

  8. Toye B, Woods W, Bobrowska M, Ramotar K: Inhibition of PCR in genital and urine specimens submitted for Chlamydia trachomatis testing. J Clin Microbiol 1998, 36:2356–2358.

    CAS  PubMed  Google Scholar 

  9. Berg ES, Anestad G, Moi H, et al.: False-negative results of a ligase chain reaction assay to detect Chlamydia trachomatis due to inhibitors in urine. Eur J Clin Microbiol Infect Dis 1997, 16:727–731.

    Article  CAS  PubMed  Google Scholar 

  10. Mahoney J, Chong S, Jang D, et al.: Urine specimens from pregnant and nonpregnant women inhibitory to amplification of Chlamydia trachomatis nucleic acid by PCR, ligase chain reaction, and transcription-mediated amplification: identification of urinary substances associated with inhibition and removal of inhibitory activity. J Clin Microbiol 1998, 36:3122–3126. This was the first study to look comprehensively at inhibitors in urine specimens for all commercially available nucleic acid amplification tests for C. trachomatis.

    Google Scholar 

  11. Hook EW 3d, Smith K, Mullen C, et al.: Diagnosis of genitourinary Chlamydia trachomatis infections by using the ligase chain reaction on patient-obtained vaginal swabs. J Clin Microbiol 1997, 35:2133–2135. This was the first report of the use of vaginal swabs (a less invasively acquired specimen) for detection of C. trachomatis infection. Although their use has not yet cleared by the FDA, vaginal specimens are likely to be well accepted in the field because they seem to be a suitable substitute for endocervical specimens and can be self-collected by women who do not undergo pelvic examinations.

    PubMed  Google Scholar 

  12. Polaneczky M, Quigley C, Pollock L, et al.: Use of self-collected vaginal specimens for detection of Chlamydia trachomatis infection. Obstet Gynecol 1998, 91:375–378.

    Article  CAS  PubMed  Google Scholar 

  13. Thomas BJ, Pierpoint T, Taylor-Robinson D, et al.: Sensitivity of the ligase chain reaction assay for detecting Chlamydia trachomatis in vaginal swabs from women who are infected at other sites. Sex Transm Dis 1998, 74:140–141.

    CAS  Google Scholar 

  14. Ostergaard L, Andersen B, Olesen F, et al.: Efficacy of home sampling for screening of Chlamydia trachomatis: randomised study. BMJ 1998, 317:26–27.

    CAS  PubMed  Google Scholar 

  15. Howell MR, Quinn TC, Brathwaite W, Gaydos CA: Screening women for Chlamydia trachomatis in family planning clinics: the cost-effectiveness of DNA amplification assays. Sex Transm Dis 1998, 25:108–117. This was the first analysis to show that screening for C. trachomatis infection with nucleic acid amplification tests can be cost effective in certain settings.

    Article  CAS  PubMed  Google Scholar 

  16. Kacena KA, Quinn SB, Howell MR, et al.: Pooling urine samples for ligase chain reaction screening for genital Chlamydia trachomatis infection in asymptomatic women. J Clin Microbiol 1998, 36:481–485. This is the first report of the pooling of urine specimens to increase the cost effectiveness of nucleic acid amplification testing in lowprevalence populations.

    CAS  PubMed  Google Scholar 

  17. Peeling RW, Toye B, Jessamine P, Gemmill I: Pooling of urine specimens for PCR testing: a cost saving strategy for Chlamydia trachomatis control programmes. Sex Transm Infect 1998, 74:66–70.

    CAS  PubMed  Google Scholar 

  18. Morre SA, Moes R, Van Valkengoed I, et al.: Genotyping of Chlamydia trachomatis in urine specimens will facilitate large epidemiological studies. J Clin Microbiol 1998, 36:3077.

    CAS  PubMed  Google Scholar 

  19. Dean D, Millman K: Molecular and mutation trends analyses of omp1 alleles for serovar E of Chlamydia trachomatis: implications for the immunopathogenesis of disease. J Clin Invest 1997, 99:475–483. This study took a novel and sophisticated approach to the study of pathogenesis of chlamydial infection through gene sequence analysis of the outer membrane protein gene.

    Article  CAS  PubMed  Google Scholar 

  20. Stothard DR, Boguslawski G, Jones RB: Phylogenetic analysis of the Chlamydia trachomatis major outer membrane protein and examination of potential pathogenic determinants. Infect Immun 1998, 66:3618–3625.

    CAS  PubMed  Google Scholar 

  21. Morre SA, Ossewarde JM, Lan J, et al.: Serotyping and genotyping of genital Chlamydia trachomatis isolates reveal variants of serovars Ba, G, and J as confirmed by omp1 nucleotide sequence analysis. J Clin Microbiol 1998, 36:345–351.

    CAS  PubMed  Google Scholar 

  22. Ciemins EL, Borenstein LA, Dyer IE, et al.: Comparisons of cost and accuracy of DNA probe test and culture for the detection of Neisseria gonorrhoeae in patients attending public sexually transmitted disease clinics in Los Angeles County. Sex Transm Dis 1997, 24:422–428.

    Article  CAS  PubMed  Google Scholar 

  23. Young H, Anderson J, Moyes A, McMillan A: Non-cultural detection of rectal and pharyngeal gonorrhoea by the Gen-Probe PACE 2 assay. Genitourin Med 1997, 73:59–62.

    CAS  PubMed  Google Scholar 

  24. Xu K, Glanton V, Johnson SR, et al.: Diagnosis of Neisseria gonorrhoeae infection by ligase chain reaction testing of urine among adolescent women with and without Chlamydia trachomatis infection. Sex Transm Dis 1998, 25:533–538.

    Article  CAS  PubMed  Google Scholar 

  25. Kehl SC, Georgakas K, Swain GR, et al.: Evaluation of the Abbott LCx assay for detection of Neisseria gonorrhoeae in endocervical swab specimens from females. J Clin Microbiol 1998, 36:3549–3551.

    CAS  PubMed  Google Scholar 

  26. Carroll KC, Aldeen WE, Morrison M, et al.: Evaluation of the Abbott LCx ligase chain reaction assay for detection of Chlamydia trachomatis and Neisseria gonorrhoeae in urine and genital swab specimens from a sexually transmitted disease clinic population. J Clin Microbiol 1998, 36:1630–1633.

    CAS  PubMed  Google Scholar 

  27. Stary A, Ching SF, Teodorowicz L, Lee H: Comparison of ligase chain reaction and culture for detection of Neisseria gonorrhoeae in genital and extragenital specimens. J Clin Microbiol 1997, 35:239–242.

    CAS  PubMed  Google Scholar 

  28. Hook EW 3d, Ching SF, Stephens J, et al.: Diagnosis of Neisseria gonorrhoeae infections in women by using the ligase chain reaction on patient-obtained vaginal swabs. J Clin Microbiol 1997, 35:2129–2132.

    PubMed  Google Scholar 

  29. Farrell DJ: Evaluation of AMPLICOR Neisseria gonorrhoeae PCR using cppB nested PCR and 16S rRNA PCR. J Clin Microbiol 1999, 37:386–390.

    CAS  PubMed  Google Scholar 

  30. Tabrizi SN, Paterson BA, Fairley CK, et al.: Comparison of tampon and urine as self-administered methods of specimen collection in the detection of Chlamydia trachomatis, Neisseria gonorrhoeae and Trichomonas vaginalis in women. Int J STD AIDS 1998, 9:347–349.

    Article  CAS  PubMed  Google Scholar 

  31. Kacena KA, Quinn SB, Hartman SC, et al.: Pooling of urine samples for screening for Neisseria gonorrhoeae by ligase chain reaction: accuracy and application. J Clin Microbiol 1998, 36:3624–3628.

    CAS  PubMed  Google Scholar 

  32. Billings SD, Fuller D, LeMonte AM, et al.: Characterization of DFA-negative, probe-positive Neisseria gonorrhoeae by pulsed field gel electrophoresis. Diagn Microbiol Infect Dis 1997, 29:281–283.

    Article  CAS  PubMed  Google Scholar 

  33. Cooke SJ, de la Paz H, La Poh C, et al.: Variation within serovars of Neisseria gonorrhoeae detected by structural analysis of outer-membrane protein PIB and by pulsed-field gel electrophoresis. Microbiol 1997, 143:1415–1422.

    CAS  Google Scholar 

  34. Harnett N, Brown S, Riley G, et al.: Analysis of Neisseria gonorrhoeae in Ontario, Canada, with decreased susceptibility to quinolones by pulsed-field gel electrophoresis, auxotyping, serotyping and plasmid content. J Med Microbiol 1997, 46:383–390.

    CAS  PubMed  Google Scholar 

  35. Hobbs MM, Alcorn TM, Davis RH, et al.: Molecular typing of Neisseria gonorrhoeae causing repeated infections: evolution of porin during passage within a community. J Infect Dis 1999, 179:371–381.

    Article  CAS  PubMed  Google Scholar 

  36. Poh CL, Ramachandran V, Tapsall JW: Genetic diversity of Neisseria gonorrhoeae IB-2 and IB-6 isolates revealed by whole-cell repetitive element sequence-based PCR. J Clin Microbiol 1996, 34:292–295.

    CAS  PubMed  Google Scholar 

  37. Morse SA: Chancroid and Haemophilus ducreyi. Clin Microbiol Rev 1989, 2:137.

    CAS  PubMed  Google Scholar 

  38. Beck-Sague CM, Cordts JR, Brown K, et al.: Laboratory diagnosis of sexually transmitted diseases in facilities within the United States: results of a national survey. Sex Transm Dis 1996, 23:342–349.

    Article  CAS  PubMed  Google Scholar 

  39. DiCarlo RP, Martin DH: The clinical diagnosis of genital ulcer disease in men. Clin Infect Dis 1997, 25:292.

    CAS  PubMed  Google Scholar 

  40. Htun Y, Morse SA, Dangor Y, et al.: Comparison of clinically directed, disease specific, and syndromic protocols for the management of genital ulcer disease in Lesotho. Sex Transm Infect 1998, 74(suppl 1):S23-S28.

    PubMed  Google Scholar 

  41. Parsons LM, Shayegani M, Waring AL, et al.: DNA probes for the identification of Haemophilus ducreyi. J Clin Microbiol 1989, 27:1441–1445.

    CAS  PubMed  Google Scholar 

  42. Totten PA, Stamm WE: Clear broth and plate media for culture of Haemophilus ducreyi. J Clin Microbiol 1994, 32:2019–2023.

    CAS  PubMed  Google Scholar 

  43. Orle KA, Gates CA, Martin DH, et al.: Simultaneous PCR detection of Haemophilus ducreyi, Treponema pallidum, and herpes simplex viruses types-1 and -2 from genital ulcers. J Clin Microbiol 1996, 34:49–54. This was the first study to describe the use and performance of a multiplex PCR assay that can amplify and detect the major etiologic agents of GUD. The development of this assay was a major advance in our ability to diagnose GUD, and the assay is an important tool for understanding the epidemiology of GUD.

    CAS  PubMed  Google Scholar 

  44. Mertz KJ, Weiss JB, Webb RM, et al.: An investigation of genital ulcers in Jackson, Mississippi, with use of a multiplex polymerase chain reaction assay: high prevalence of chancroid and human immunodeficiency virus infection. J Infect Dis 1998, 178:1060–1066.

    Article  CAS  PubMed  Google Scholar 

  45. Risbud A, Chan-Tack K, Gadkari D, et al.: The etiology of genital ulcer disease by multiplex polymerase chain reaction and relationship to HIV infection among patients attending sexually transmitted clinics in Pune, India. Sex Transm Dis 1999, 26:55–62.

    Article  CAS  PubMed  Google Scholar 

  46. Morse SA, Trees DL, Htun Y, et al.: Comparison of clinical and standard laboratory and molecular methods for the diagnosis of genital ulcer disease in Lesotho: association with human immunodeficiency virus infection. J Infect Dis 1997, 175:583–589.

    CAS  PubMed  Google Scholar 

  47. Mertz KA, Trees D, Levine WC, et al.: Etiology of genital ulcers and prevalence of human immunodeficiency virus coinfection in 10 US cities. J Infect Dis 1998, 178:1795–1798.

    Article  CAS  PubMed  Google Scholar 

  48. Beyrer CK, Jitwatcharanan C, Natpratan R, et al.: Molecular methods for the diagnosis of genital ulcer disease in a sexually transmitted disease clinic population in Northern Thailand: predominance of herpes simplex virus infection. J Infect Dis 1998, 178:243–246.

    CAS  PubMed  Google Scholar 

  49. Chui L, Albritton W, Baster B, et al.: Development of the polymerase chain reaction for the diagnosis of chancroid. J Clin Microbiol 1993, 31:659–664.

    CAS  PubMed  Google Scholar 

  50. West B, Wilson SM, Changalucha S, et al.: Simplified PCR for detection of Haemophilus ducreyi and diagnosis of chancroid. J Clin Microbiol 1995, 33:787–790.

    CAS  PubMed  Google Scholar 

  51. Roesel DJ, Gwanzura L, Mason PR, et al.: Polymerase chain reaction detection of Haemophilus ducreyi DNA. Sex Transm Infect 1998, 74:63–65.

    Article  CAS  PubMed  Google Scholar 

  52. Gu XX, Roussau R, Jannes G, et al.: The rrs(16S)-rrl(23S) ribosomal intergenic spacer region as a target for the detection of Haemophilus ducreyi by a heminested PCR assay. Microbiology 1998, 144:1013–1019.

    CAS  PubMed  Google Scholar 

  53. Parsons LM, Waring AL, Otido J, et al.: Laboratory diagnosis of chancroid using species-specific primers from Haemophilus ducreyi groEL and the polymerase chain reaction. Diagn Microbiol Infect Dis 1995, 23:89–98.

    Article  CAS  PubMed  Google Scholar 

  54. Johnson SR, Martin DH, Cammarata C, et al.: Alterations in sample preparation increase sensitivity of PCR assay for diagnosis of chancroid. J Clin Microbiol 1995, 33:1036–1038.

    CAS  PubMed  Google Scholar 

  55. Larsen SA, Steiner BM, Rudolph AH: Laboratory diagnosis and interpretation of tests for syphilis. Clin Microbiol Rev 1995, 8:1–21.

    CAS  PubMed  Google Scholar 

  56. Kennedy EJ Jr, Creighton ET: Darkfield microscopy for the detection and identification of T. pallidum. In A Manual of Tests for Syphilis, edn 9. Edited by Larsen SA, Pope V,Johnson R, Kennedy EJ Jr. Washington, DC: American Public Health Association; 1998:120–134.

    Google Scholar 

  57. Zoechling N, Schuluepen EM, Soyer HP, et al.: Molecular detection of Treponema pallidum in secondary and tertiary syphilis. Br J Dermatol 1997, 136:683–686.

    Article  CAS  PubMed  Google Scholar 

  58. Horowitz HW, Valsamis MP, Wicher V, et al.: Cerebral syphilitic gumma confirmed by the polymerase chain reaction in man with human immunodeficiency virus infection. N Engl J Med 1994, 331:1488–1491.

    Article  CAS  PubMed  Google Scholar 

  59. Inagaki H, Kawai T, Miyata M, et al.: Gastric syphilis: polymerase chain reaction detection of treponemal DNA in pseudolymphomatous lesions. Hum Pathol 1996, 27:749–750.

    Article  Google Scholar 

  60. Hay PE, Clarke JR, Strungel RA, et al.: Use of the polymerase chain reaction to detect DNA sequences specific to pathogenic treponemes in cerebrospinal fluid. FEMS Microbiol Lett 1990, 68:233–238.

    CAS  Google Scholar 

  61. Sanchez PJ, Wendel GD Jr, Grimprel E, et al.: Evaluation of molecular methodologies and rabbit infectivity testing for the diagnosis of congenital syphilis and neonatal central nervous system invasion by Treponema pallidum. J Infect Dis 1993, 167:148–157.

    CAS  PubMed  Google Scholar 

  62. Centurion-Lara A, Castro C, Shaffer JM, et al.: Detection of Treponema pallidum by a sensitive reverse transcriptase PCR. J Clin Microbiol 1997, 35:1348–1352.

    CAS  PubMed  Google Scholar 

  63. Cone RW, Hobson AC, Palmer J, et al.: Extended duration of herpes simplex virus DNA in genital lesions detected by polymerase chain reaction. J Infect Dis 1991, 164:757–760.

    CAS  PubMed  Google Scholar 

  64. Fodor PA, Levin MJ, Weinberg A, et al.: Atypical herpes simplex virus encephalitis diagnosed by PCR amplification of viral DNA from CSF. Neurology 1998, 51:554–559.

    CAS  PubMed  Google Scholar 

  65. Slomka MJ, Emery L, Munday PE, et al.: A comparison of PCR with virus isolation and direct antigen detection for diagnosis and typing of genital herpes. J Med Virol 1998, 55:177–183.

    Article  CAS  PubMed  Google Scholar 

  66. Cohen BA, Rowley AH, Long CM: Herpes simplex type 2 in a patient with Mollaret’s meningitis: demonstration by polymerase chain reaction. Ann Neurol 1994, 35:112–116.

    Article  CAS  PubMed  Google Scholar 

  67. Hobson A, Wald A, Wright N, et al.: Evaluation of a quantitative competitive PCR assay for measuring herpes simplex virus DNA content in genital tract secretions. J Clin Microbiol 1997, 35:548–552.

    CAS  PubMed  Google Scholar 

  68. Diaz-Mitoma F, Ruben M, Sacks S, et al.: Detection of viral DNA to evaluate outcome of antiviral treatment of patients with recurrent genital herpes. J Clin Microbiol 1996, 34:657–663.

    CAS  PubMed  Google Scholar 

  69. Cullen AP, Long CD, Lorincz AT: Rapid detection and typing of herpes simplex virus DNA in clinical specimens by the Hybrid Capture II signal amplification probe test. J Clin Microbiol 1997, 35:2275–2278.

    CAS  PubMed  Google Scholar 

  70. Goldberg J: Studies on granuloma inguinale. V. Isolation of a bacterium resembling Donovania granulomatis from the feces of a patient with granuloma inguinale. Br J Vener Dis 1962, 38:99.

    CAS  PubMed  Google Scholar 

  71. Packer H, Goldberg J: Studies on the antigenic relationship of D. granulomatis to members of the tribe Eschericheae. Am J Syph Gon Vener Dis 1950, 34:342.

    CAS  Google Scholar 

  72. Bastian I, Bowden FJ: Amplification of Klebsiella-like sequences from biopsy specimens from patients with donovanosis. Clin Infect Dis 1996, 23:1328.

    CAS  PubMed  Google Scholar 

  73. Carter J, Hutton S, Sriprakash KS: Culture of the causative organism of donovanosis (Calymmatobacterium granulomatis) in Hep-2 cells. J Clin Microbiol 1997, 35:2915.

    CAS  PubMed  Google Scholar 

  74. Mittal A, Chandra M: In vitro cultivation of Calymmatobacterium granulomatis in cultured macrophages. Indian J Sex Transm Dis 1992, 13:68.

    Google Scholar 

  75. Kharsany ABM, Hoosen AA, Kiepiela P, et al.: Culture of Calymmatobacterium granulomatis. Clin Infect Dis 1996, 22:391.

    CAS  PubMed  Google Scholar 

  76. Sarafian SK, Woods TC, Knapp JS, et al.: Molecular characterization of Haemophilus ducreyi by ribosomal DNA fingerprinting. J Clin Microbiol 1991, 29:1949–1954. The recently developed ribotyping system for H. ducreyi has become an important tool for investigating the epidemiology of chancroid.

    CAS  PubMed  Google Scholar 

  77. Flood JM, Sarafian SK, Bolan GA, et al.: Multistrain outbreak of chancroid in San Francisco, 1989–1991. J Infect Dis 1993, 167:1106–1111.

    CAS  PubMed  Google Scholar 

  78. Brown TJ, Ison CA: Non-radioactive ribotyping of Haemophilus ducreyi using a digoxigenin labelled cDNA probe. Epidemiol Infect 1993, 110:289–295.

    Article  CAS  PubMed  Google Scholar 

  79. Pillay A, Hoosen AA, Kiepiela P, et al.: Ribosomal DNA typing of Haemophilus ducreyi strains: proposal for a novel typing scheme. J Clin Microbiol 1996, 34:2613–1615.

    CAS  PubMed  Google Scholar 

  80. Miao RM, Fieldsteel H: Genetic relationship between Treponema pallidum and Treponema pertenue, two noncultivable human pathogens. J Bacteriol 1980, 141:427–429.

    CAS  PubMed  Google Scholar 

  81. Noordhoek GT, Hermans PW, Paul AN, et al.: Treponema pallidum subspecies pallidum (Nichols) and Treponema pallidum subspecies pertenue (CDC2575) differ in at least one nucleotide: comparison of two homologous antigens. Microbiol Pathol 1989, 6:29–42.

    Article  CAS  Google Scholar 

  82. Centurion-Lara A, Arroll TA, Castillo R, et al.: Conservation of the 15 kd lipoprotein among Treponema pallidum subspecies, strains, and other pathogenic treponemes: genetic and antigenic analyses. Infect Immun 1996, 65:1440–1444.

    Google Scholar 

  83. Centurion-Lara A, Castro C, Van Voorhis WC, et al.: Two 16S-23S ribosomal DNA intergenic regions in different Treponema pallidum subspecies contain tRNA genes. FEMS Microbiol Lett 1997, 143:235–240.

    Article  Google Scholar 

  84. Pillay A, Liu H, Chen CY, et al.: Molecular subtyping of Treponema pallidum subspecies pallidum. Sex Transm Dis 1998, 25:408–414. This was the first study to conclusively identify genetic polymorphisms in T. pallidum. The importance of this finding with respect to our understanding of the epidemiology of syphilis remains to be determined.

    Article  CAS  PubMed  Google Scholar 

  85. Fraser CM, Norris SJ, Weinstock GM, et al.: Complete genome sequence of Treponema pallidum, the syphilis spirochete. Science 1998, 281:375–388.

    Article  CAS  PubMed  Google Scholar 

  86. Kuzushima K, Kimura H, Kino Y, et al.: Clinical manifestations of primary herpes simplex virus type 1 infection in a closed community. Pediatrics 1991, 87:152–158.

    CAS  PubMed  Google Scholar 

  87. Alam TM, Joncas JH, Ozanne G: DNA polymorphism among isolates from multiple sites of a patient with chronic herpes simplex virus type 1 infection. J Med Virol 1989, 29:186–191.

    Article  CAS  PubMed  Google Scholar 

  88. Sakaoka H, Kurita K, Gouro T, et al.: Analysis of genomic polymorphism among herpes simplex virus type 2 isolates from four areas of Japan and three other countries. J Med Virol 1995, 45:259–272.

    Article  CAS  PubMed  Google Scholar 

  89. Roizman B, Tognon M: Restriction endonuclease patterns of herpes simplex virus DNA: applications to diagnosis and molecular epidemiology. Curr Top Microbiol Immunol 1983, 104:275–286.

    Google Scholar 

  90. Bollen LJ, Tjong A, Hung SP, et al.: Human papillomavirus deoxyribonucleic acid detection in mildly or moderately dysplastic smears: a possible method for selecting patients for colposcopy. Am J Obstet Gynecol 1997, 177:548–553.

    Article  CAS  PubMed  Google Scholar 

  91. Peyton CL, Schiffman M, Lorincz AT, et al.: Comparison of PCR-and hybrid capture-based human papillomavirus detection systems using multiple cervical specimen collection strategies. J Clin Microbiol 1998, 36:3248–3254.

    CAS  PubMed  Google Scholar 

  92. Cope JU, Hildesheim A, Schiffman MH, et al.: Comparison of the hybrid capture tube test and PCR for detection of human papillomavirus DNA in cervical specimens. J Clin Microbiol 1997, 35:2262–2265.

    CAS  PubMed  Google Scholar 

  93. Lie AK, Skjeldestad FE, Hagen B, et al.: Comparison of light microscopy, in situ hybridization and polymerase chain reaction for detection of human papillomavirus in histological tissue of cervical intraepithelial neoplasia. Acta Pathol Microbiol Immunol Scand 1997, 105:115–120.

    CAS  Google Scholar 

  94. Kareen BN, Kalsen F, Holm R, et al.: A novel grid polymerase chain reaction (G-PCR) approach at ultrastructural level to detect target DNA in cell cultures and tissues. J Pathol 1997, 183:486–493.

    Article  Google Scholar 

  95. Clavel C, Rihet S, Masure M, et al.: DNA-EIA to detect high and low risk HPV genotypes in cervical lesions with E6/E7 primer mediated multiplex PCR. J Clin Pathol 1998, 51:38–43.

    CAS  PubMed  Google Scholar 

  96. Gostout BS, Podratz KC, McGovern RM, et al.: Cervical cancer in older women: a molecular analysis of human papillomavirus types, HLA types, and p53 mutations. Am J Obstet Gynecol 1998, 179:56–61.

    Article  CAS  PubMed  Google Scholar 

  97. Gravitt PE, Peyton CL, Apple RJ, et al.: Genotyping of 27 HPV types by using L1 consensus PCR products by a singlehybridization, reverse line blot detection method. J Clin Microbiol 1998, 36:3020–3027.

    CAS  PubMed  Google Scholar 

  98. DeMeo LR, Draper DL, McGregor JA, et al.: Evaluation of a deoxyribonucleic acid probe for the detection of Trichomonas vaginalis in vaginal secretions. Am J Obstet Gynecol 1996, 174:1339–1342. To date, this is the only reported evaluation of the commercial DNA probe for T. vaginalis (Affirm, Becton Dickinson).

    Article  CAS  PubMed  Google Scholar 

  99. Madico G, Quinn TC, Rompalo A, et al.: Diagnosis of Trichomonas vaginalis infection by PCR using vaginal swab samples. J Clin Microbiol 1998, 36:3205–3210.

    CAS  PubMed  Google Scholar 

  100. Witkin SS, Inglis SR, Polaneczky M: Detection of Chlamydia trachomatis and Trichomonas vaginalis by polymerase chain reaction in introital specimens from pregnant women. Am J Obstet Gynecol 1996, 175:165–167.

    Article  CAS  PubMed  Google Scholar 

  101. Heine RP, Wiesenfeld HC, Sweet RL, et al.: Polymerase chain reaction analysis of distal vaginal specimens: a less invasive strategy for detection of Trichomonas vaginalis. Clin Infect Dis 1997, 24:985–987.

    CAS  PubMed  Google Scholar 

  102. Paterson BA, Tabrizi SN, Garland SM, et al.: The tampon test for trichomoniasis: a comparison between conventional methods and a polymerase chain reaction for Trichomonas vaginalis in women. Sex Transm Infect 1998, 74:136–139.

    CAS  PubMed  Google Scholar 

  103. Lin RR, Shaio MF, Liu JY: One-tube, nested-PCR assay for the detection of Trichomonas vaginalis in vaginal discharges. Ann Trop Med Parasitol 1997, 91:61–65.

    CAS  PubMed  Google Scholar 

  104. Vanacova S, Tachezy J, Kulda J, Flegr J: Characterization of trichomonad species and strains by PCR fingerprinting. J Eukaryot Microbiol 1997, 44:545–552.

    Article  CAS  PubMed  Google Scholar 

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  1. Centers for Disease Control and Prevention, Mailstop A12, 1600 Clifton Road NE, 30333, Atlanta, GA, USA

    Carolyn M. Black PhD & Stephen A. Morse MSPH, PhD

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Black, C.M., Morse, S.A. The use of molecular techniques for the diagnosis and epidemiologic study of sexually transmitted infections. Curr Infect Dis Rep 2, 31–43 (2000). https://doi.org/10.1007/s11908-000-0085-x

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