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Br J Ophthalmol. Nov 2006; 90(11): 1425–1429.
Published online Aug 9, 2006. doi:  10.1136/bjo.2006.096420
PMCID: PMC1857507

Detection by broad‐range real‐time PCR assay of Chlamydia species infecting human and animals



Tests available for molecular diagnosis of chlamydial infections detect Chlamydiatrachomatis, but do not find other Chlamydia species associated with genital, ophthalmic, cardiovascular, respiratory or neurological diseases. The routine detection of all Chlamydia species would improve the prognosis of infected people and guide therapeutic choices.


To design and validate a sensitive, specific, reproducible, inexpensive and easy‐to‐perform assay to quantify most Chlamydia species.


Primers and probe were selected using the gene coding for the 16S rRNA. The detection limits were assessed for suspensions of Chlamydia trachomatis, Chlamydia psittaci and Chlamydia pneumoniae. The performance of this test was compared with that of two commercial kits (Amplicor‐Roche and Artus) on 100 samples obtained from children with trachoma.


The detection capacities for Chlamydia trachomatis of the broad‐range real‐time polymerase chain reaction (PCR) were similar or slightly better than those obtained with commercial kits (0.2 copies of DNA/μl). Only the broad‐range PCR identified specimens containing Chlamydia psittaci and Chlamydia pneumoniae. The commercial kits and the broad‐range assay detected Chlamydia species in 5% and in 11%, respectively, of samples from children with trachoma.


This new real‐time PCR offers a sensitive, reproducible assay that produces results in <3 h. With panels of quantified Chlamydia species, this real‐time PCR can be run with all real‐time PCR equipment. Larger trials are needed to confirm the utility of this test in diagnosis and for therapeutic follow‐up.

The Chlamydiaceae (Chlamydia)—two distinct Gram‐negative obligate intracellular bacterial lineages that branch into nine separate clusters—cause cervicitis, urethritis, rectitis, endometritis and salpingitis, inclusion conjunctivitis and cardiorespiratory infections, and are responsible (or are cofactors) for trachoma.1,2,3,4 The Chlamydiaceae, reclassified according to their 16S and 23S ribosomal gene sequences, which were previously known only by the genus Chlamydia, were divided into two genera, Chlamydia and Chlamydophila gen nov. The Chlamydophila gen nov assimilates the current species Chlamydia pecorum,Chlamydia pneumoniae and Chlamydia psittaci, to form Chlamydophila pecorum comb nov, Chlamydophilapneumoniae comb nov and Chlamydophila psittaci comb nov. Three new Chlamydophila species were derived from Chlamydia psittaci: Chlamydophila abortus gen nov, sp nov, Chlamydophila caviae gen nov, sp nov and Chlamydophila felis gen nov, sp nov.5

Chlamydia trachomatis is the most common sexually transmitted agent in non‐gonococcal urethritis and may cause epididymitis.6,7,8,9 Infants may develop eye diseases (inclusion conjunctivitis), pneumonia, and pharyngeal and enteric complications.1,10,11,12Chlamydia associated with poverty triggers trachoma, the prime cause of preventable infectious blindness.13,14,12,15Chlamydia psittaci shed by avian species and by domestic mammals may also cause conjunctivitis and severe pulmonary diseases.16,17,12Chlamydia pneumoniae can induce proliferation of smooth muscle cells and contribute to the aggravation of primary pulmonary symptoms,18,19,4 and is associated with pharyngitis, bronchitis, pneumonia and endothelial disorders.20,21,22

Various molecular approaches have been developed in the nucleic acid amplification tests (NAATs) used for diagnosis of chlamydial infections:

  1. Qualitative PCR (Amplicor Roche‐USA), which amplifies a cryptic plasmid DNA from Chlamydia trachomatis
  2. Quantitative PCR assay (Artus‐Germany), which amplifies a sequence of a major structural protein (ompA)
  3. The AC2 assay (Gen‐Probe‐USA), which targets the Chlamydia trachomatis rRNA and uses a technology based on the transcription‐mediated amplification of isolated target sequences23
  4. The Becton Dickinson ProbeTec (Becton Dickinson Diagnostic Systems‐USA), which targets a sequence within a cryptic plasmid DNA of Chlamydia trachomatis using DNA polymerase and a specific restriction‐enzyme strand‐displacement amplification.24

As the commercial diagnostic tests for these entities are able to detect only Chlamydia trachomatis, we designed a set of primers and a probe that recognises Chlamydiaceae using the real‐time technology. The goal of this work was to develop a sensitive, inexpensive and easy‐to‐perform assay that was able to detect and quantify most of the species of Chlamydia infecting humans and animals. The detection limits for this new assay were assessed with strains of Chlamydia trachomatis, Chlamydiapsittaci and Chlamydiapneumoniae. In addition, the detection capacities of this assay were studied in parallel with two commercial kits (Amplicor and Artus), with samples obtained from children with trachoma.

Material and methods

Human samples

Sampling was carried out after obtaining the informed consent of parents or guardians during epidemiological surveys, and in accordance with the Declaration of Helsinki and the electroencephalogram human experimentation guidelines.

A total of 100 clinical samples obtained from children (boys and girls aged 1–10 years) living in the Kankan area (Haute‐Guinea) who had trachoma diagnosed by experienced ophthalmologists (according to the World Health Organization grading card: [gt-or-equal, slanted]5 follicles in a specific area of the upper eye lid) were obtained by vigorous scraping of the upper conjunctiva with Dacron swabs. To avoid cross contamination, the doctor in charge of sampling was not allowed to handle pens, torches or clinical files, or to register any data. Independent assistants managed the identification and seating of the children and the registration of data. Disposable material was used, and paper covers on the seats were changed after each patient had used them; equipment and surfaces were cleaned with disinfectant and 1 N hydrochloric acid. The operators wore disposable shirts, masks and glasses, and two pairs of gloves, one of which was changed for each new patient. Once clinical diagnosis was confirmed, the upper tarsal conjunctiva was swabbed intensely with Dacron swabs, and the samples were placed in a sterile tube, which was closed tightly. Before introduction of the swabs into the containers, the outside tube surfaces were decontaminated with sterile gauze lightly dampened with 1 N hydrochloric acid. The tubes were kept dry in a container (4–8°C) for <8 h after sampling and then stored at −20°C for DNA extraction. The aliquots were blind tested with the three assays. The Amplicor and Artus PCRs were carried out according to the manufacturers' instructions.

Nucleic acid extraction

Specimens in the Dacron swabs were thawed; 500 μl of phosphate‐buffered saline was added and vortexed intensively. DNA was extracted using the Magnapure (Roche Applied Sciences) from 200 μl of specimens using total nucleic acid isolation. The extracts were recovered in 50 μl of water and analysed by Amplicor and by Artus.

A separate extraction with Magnapure was carried out for the broad‐spectrum Chlamydia PCR. To monitor the extraction and amplification processes, 5 μl of a whole virus preparation of seal herpes virus (a gift from G J van Doornum, Department of Virology Erasmus MC, Rotterdam, The Netherlands) was added to each sample before extraction (final concentration of about 1000 viral particles/ml).

Primers and probe

Alignments of the chlamydial genes coding for the 16S rRNA (rDNA) were carried out to identify the highest conserved regions in Chlamydia pneumoniae, Chlamydia trachomatis and Chlamydia psittaci sequences exhibiting >90% homology.25,26,27,4 The sequences were selected using the primer Express 1.0 software (Applied Biosystems, France). The forward primer (5′TCGAGAATCTTTCGCAATGGAC) and the reverse primer (5′CGCCCTTTACGCCCAATAAA) were BLAST searched using the NCBI Blastn FAQs (ncbi.nlm.nih.gov/Genbank). These primers bracket a highly conserved sequence (multicopy gene) coding for the ribosomal RNA in most sequences of Chlamydia trachomatis, Chlamydia psittaci, Chlamydia pneumoniae, Chlamydia felis, Chlamydia pecorum, Chlamydia caviae, Chlamydia suis and Chlamydia muridarum.28,5,26,27,4 The reporter (5′fluor, 6‐FAM) and the quencher (TAMRA) dyes were attached to the 5′ and 3′ ends of the probe, respectively (FAM‐AAGTCTGACGAAGCGACGCCGC). No cross reactivity with genes of other microorganisms, or with any other mammalian gene was detected for the primers or for the probe.

Real‐time TaqMan broad‐spectrum PCR assay

Different temperatures and annealing or extension times for each step of the reactions were studied in preliminary assays to determine the best experimental conditions in which at least 0.5 copies/μl of Chlamydia trachomatis DNA could be detected. PCR reactions were carried out in a final volume of 50 μl containing 2× TaqMan Universal Mastermix (MNL 430449, Applied Biosystems), forward primer (0.5 μmol/l), reverse primer (0.5 μmol/l), FAM‐TAMRA probe (0.4 μmol/l) and 25 μl of the isolated DNA eluted in distilled water. After incubation for 2 min at 50°C with uracil N‐glycosylase to neutralise potential PCR contaminants from previous reactions, the microtubes were incubated for 10 min at 95°C. The PCR cycling programme consisted of 50 two‐step cycles of 10 s at 95°C, and 65 s at 60°C. The amplification and detection were carried out with the ABI Prism 7000 sequence detector system (Applied Biosystems). Retest and second derivative analysis were carried out with the SmartCyclerII (IL Laboratory, Cepheid‐Sunnyvale, California, USA). The Ct value for each sample was determined according to the fluorescence signal exceeding the background limit of 0.20. Each run contained negative controls with no template.

Calibrated samples containing Chlamydia trachomatis, purchased from the European Union Concerted Action on Quality Control of Nucleic Acid Amplification Program (EQCP, Glasgow, UK), were tested pure and diluted in distilled water before DNA extraction to assess the linearity, sensitivity, reproducibility and detection limits. The introduction into each sample before DNA extraction of a non‐human calibrated virus served to monitor the DNA extraction and to assess PCR inhibition. Dilutions of Chlamydia trachomatis, Chlamydia psittaci and Chlamydia pneumoniae were retested using the SmartCyclerII system, which enabled us to follow the second derivative of the growth curve (rates of change for the curve slopes) in realtime. Here, the highest peak of the second derivative curves representing the point of maximum curvature of the signal curves or the transition from non‐specific signals and background to amplified product fluorescence was used to validate the true relevance of the signals.


Table 11 shows that the broad‐range real‐time PCR assay is able to detect and semiquantify Chlamydia trachomatis. The broad‐range assay detection capacities for Chlamydia trachomatis are similar or slightly better than those obtained with two commercial kits, and as few as 0.2 copies of DNA/μl were detected with this assay in three different experiments. Table 22 shows that only the broad range assay for Chlamydia was able to detect three different species of Chlamydia, whereas the commercial kits did not detect any of the dilutions of either Chlamydia psittaci or Chlamydia pneumoniae. The broad range test was positive for the three samples containing the equivalent of 0.5 CFU/1000 μl or more of Chlamydia psittaci and Chlamydia penumoniae. The linearity and reproducibility of the assay were studied with supernatants of cells infected with Chlamydia psittaci or Chlamydia pneumoniae (commercial quality control panels are not yet available), and the Ct values generating positive signals (confirmed by analysing the second derivative) were similar when tested in different experiments (dispersion always <1.2 Ct).

Table thumbnail
Table 1 Detection capacities of the broad‐range real‐time PCR assay and two commercial kits on Chlamydia trachomatis elementary bodies
Table thumbnail
Table 2 Comparison of the detection capacities of the broad‐range real‐time PCR for Chlamydia with two commercial kits on dilutions of semiquantified culture supernatants of Chlamydia psittaci and Chlamydia pneumoniae

Table 33 shows that when comparing the performances of the Amplicor, the Artus and the broad‐spectrum real‐time PCR assay with samples obtained from children with trachoma, the rates of positivity were similar for Amplicor and Artus, but greater for the broad‐range assay (11 positive v 5 positive for the commercial assays). The six positive samples detected as positive with the broad‐range assay should have been further investigated to determine the sequences of the amplicons, but the specimens were tested in a routine laboratory in which tubes containing amplified products were discarded, suggesting that in future studies additional steps should be carried out to detect the implication of Chlamydia species other than Chlamydia trachomatis in trachoma.

Table thumbnail
Table 3 Comparison of the detection capacities of the broad‐range real‐time PCR for Chlamydia with two commercial kits on 100 clinical samples obtained from children with trachoma


The NAATs are known for being the most sensitive methods for the diagnosis of Chlamydia trachomatis infections and can detect as few as one organism per assay, whereas the limit of detection for the conventional tests is [gt-or-equal, slanted]10 organisms;28,29,30 previous studies with urine samples showed that commercial kits are not significantly different in their ability to detect Chlamydiatrachomatis.23,24

The commercial NAATs have monospecies spectrums and fail to meet the expectations of the clinical microbiology services. It has been reported that these assays may be effective to varying degrees for the detection of Chlamydia trachomatis in samples from patients with conjunctivitis or trachoma, but they may produce negative results for other species or if the plasmid (detected by Amplicor and by the Becton Dickinson Probe Tec test) that is required for chlamydial growth in vitro is altered or is absent.31,32 Hence, the conclusions drawn from amplifying narrow sequences (in the plasmid and only for the species Chlamydia trachomatis) or regions coding for structural proteins that may be altered (Artus and Gen Probe AC2 assays) may underestimate the levels of infection.13,14,33,34,31,32 In the present study, the amplification of the plasmid (Amplicor) and the amplification of the momp‐gene (Artus) showed identical rates of positivity in patients with trachoma, suggesting that the low detection levels may not be due to the loss of the plasmid or due to mutations in the mono‐gene.31,35,27

Previous trials carried out in areas with endemic trachoma showed discrepancies between the clinical diagnosis and PCR (Amplicor), with positivity levels ranging from <10% to 70%. As an example, in high‐prevalence communities with active trachoma in Nepal, 70% of clinically active cases were positive by PCR, whereas only 8% were positive in the low‐prevalence areas. In a hyper endemic region in Africa, PCR was positive in <57%.13,34,35,15,36,37,38 In the present trial, where the prevalence of clinical follicular trachoma was 30%, only 5% were positive using commercial kits, suggesting the need for a comprehensive review of the predictive value of the commercial NAATs.

It was reported that the persistent Chlamydia psittaci infection may have contributed to the development of lymphomas, as was supported by the clinical responses observed with antibiotic treatment39; in patients with ocular adnexal lymphoma (higher prevalence of Chlamydia psittaci infection in both tumour and peripheral blood mononuclear cells), this new broad‐spectrum assay should be a complementary tool for routine diagnosis. In addition, Chlamydia pneumoniae was associated with chronic forms of follicular conjunctivitis, ophthalmia neonatorum and with human choroidal neovascular membranes,19,16,17 suggesting that by inducing chronic inflammation and pro‐angiogenic cytokines it may contribute to the pathogenesis of age‐related macular degeneration20,40 (chlamydial genes coding for cytokines, growth factors and signalling pro‐inflammatory molecules are upregulated as early as 2 h post‐infection).10,41

The load of Chlamydia pneumoniae is also higher in people with aortic stenosis in the calcified and fibrotic regions of the aortic valve,21 and this infection was associated with acute myocardial infarction in young men in the US military. Nevertheless, its direct involvement is under discussion as clinical trials carried out with macrolides do not seem to alter the risk of cardiac events,42 and no benefits for survival in patients with peripheral arterial disease have yet been reported.43,44,45,46

Presently, the NAATs carried out to detect Chlamydia psittaci show poor performance, and the sensitivity of this method could have been enhanced by adding amplification steps (“nested PCR”) or by hybridisation of the amplicons with specific probes, but this technology, useful for research purposes, is impractical in a clinical routine microbiology setting.16,30 Regarding the broad‐range NAATs for Chlamydia species, one PCR assay was reported, but the amplified products were electrophoresed through 2% agarose and the amplicons hybridised (which is reagent‐consuming and time‐consuming) before the results could be assessed as positive or negative by ultraviolet transillumination.25 Because Chlamydiae are not isolated in any of the media used by microbiology laboratories, the NAATs are becoming the reference methods.26,4,47,29 However, we should warn of the risk of misdiagnosis (false negative) when drawing conclusions from results obtained with mono‐specific assays. A broad‐spectrum biological assay should be of great help in understanding the pathophysiology and the association of clinical signs with Chlamydia (trachomatis and others), which may guide the choice towards agents with appropriate activity against intracellular bacteria.

In conclusion, this study shows the results of the first broad‐range real‐time PCR assay, targeting sequences of a conserved region of a bacterial multi copy gene coding for the 16S rRNA shared by most Chlamydia species. This assay is reproducible and results can be obtained within 3 h at considerably lower cost than of those on the market. If quality control panels of quantified Chlamydia are available, the broad‐range real‐time PCR can be easily adapted to different equipments and can be run as a routine test.


CFU - colony forming units

NAAT - nucleic acid amplification test

PCR - polymerase chain reaction


Competing interests: None declared.


1. Barnes R C, Rompalo A M, Stamm M E. Comparison of Chlamydia trachomatis serovars causing rectal and cervical infections. J Infect Dis 1987. 156953–958.958 [PubMed]
2. Boisvert J F, Koutsky L A, Suchland R J. et al Clinical features of Chlamydia trachomatis rectal infection by serovar among homosexually active men. Sex Transm Dis 1999. 26392–398.398 [PubMed]
3. Dean D, Oudens E, Bolan G. et al Major outer membrane protein variants of Chlamydia trachomatis are associated with severe upper genital tract infections and histopathology in San Francisco. J Infect Dis 1995. 721013–1022.1022 [PubMed]
4. Nash P, Krenz M M. In: Balows A, Hausler WJ Jr, Herrmann KL, Isenberg HD, ShadomyHJ, eds. Manual of clinical microbiology. 5th edn. Washington: American Society for Microbiology, 1991
5. Everett K D, Bush R M, Andersen A A. Amended description of the order Chlamydiales, proposal of Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov., each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms. Int J Syst Bacteriol 1999. 2415–440.440 [PubMed]
6. Patton D L, Sweeney Y T, Kuo C C. Demonstration of delayed hypersensitivity in Chlamydia trachomatis salpingitis in monkeys: a pathogenic mechanism of tubal damage. J Infect Dis 1994. 169680–683.683 [PubMed]
7. Stamm W E, Koutsky L A, Benedetti J K. et al Chlamydia trachomatis urethral infections in men. Prevalence, risk factors, and clinical manifestations. Ann Intern Med 1984. 10047–51.51 [PubMed]
8. Wang S P, Grayston J T. Immunologic relationship between genital TRIC, lymphogranuloma venereum, and related organisms in a new microtiter indirect immunofluorescence test. Am J Ophthalmol 1970. 70367–374.374 [PubMed]
9. Workowski K A, Stevens C E, Suchland R J. et al Clinical manifestations of genital infection due to Chlamydia trachomatis in women: differences related to serovar. Clin Infect Dis 1994. 19756–760.760 [PubMed]
10. Belland R J, Scidmore M A, Crane D D. et al Chlamydia trachomatis cytotoxicity associated with complete and partial cytotoxin genes. Proc Natl Acad Sci USA 2001. 9813984–13989.13989 [PMC free article] [PubMed]
11. 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. 99475–483.483 [PMC free article] [PubMed]
12. Kalayoglu M V. Ocular chlamydial infections: pathogenesis and emerging treatment strategies. Curr Drug Targets Infect Disord 2002. 185–91.91 [PubMed]
13. Alexander N D, Solomon A W, Holland M J. et al An index of community ocular Chlamydia trachomatis load for control of trachoma. Trans R Soc Trop Med Hyg 2005. 3175–177.177 [PubMed]
14. Bird M, Dawson C R, Schachter J S. et al Does the diagnosis of trachoma adequately identify ocular chlamydial infection in trachoma‐endemic areas? Infect Dis 2003. 101669–1673.1673 [PubMed]
15. Solomon A W, Peeling R W, Foster A. et al Diagnosis and assessment of trachoma. Clin Microbiol Rev 2004. 4982–1011.1011 [PMC free article] [PubMed]
16. Hartley J C, Stevenson S, Robinson A J. et al Conjunctivitis due to Chlamydophila felis (Chlamydia psittaci feline pneumonitis agent) acquired from a cat: case report with molecular characterization of isolates from the patient and cat. J Infect 2001. 17–11.11 [PubMed]
17. Iwamoto K, Masubuchi K, Nosaka H. et al Isolation of chlamydia psittaci from domestic cats with oculonasal discharge. Jpn J Vet Med Sci 2001. 8937–938.938 [PubMed]
18. Apfalter P, Barousch W, Nehr M. et al Comparison of a new quantitative ompA‐based real‐time TaqMan assay for detection of Chlamydia pneumoniae in respiratory specimens with four conventional PCR assays. J Clin Microbiol 2003. 2592–600.600 [PMC free article] [PubMed]
19. Grayston J T, Kuo C C, Campbell L A. et al Chlamydia pneumoniae strain TWAR. Int J Syst Bacteriol 1989. 3988–90.90
20. Kalayoglu M V, Bula D, Arroyo J. et al Identification of Chlamydia pneumoniae within human choroidal neovascular membranes secondary to age‐related macular degeneration. Graefe's Arch Clin Exp Ophthalmol 2005. 24321 [PubMed]
21. Pierri H, Higuchi‐Dos‐Santos M H, Higuchi M D. et al Density of Chlamydia pneumoniae is increased in fibrotic and calcified areas of degenerative aortic stenosis. Int J Cardiol 2006. 2243–47.47 [PubMed]
22. Wohlschlaeger J, Wimmer M L, Nagler D K. et al Identification of Chlamydia pneumoniae in intracranial and extracranial arteries patients with stroke and in controls: combined immunohistochemical and polymerase chain reaction analyses. Hum Pathol 2005. 4395–402.402 [PubMed]
23. Gaydos C A, Theodore M, Dalesio N. et al Comparison of three nucleic acid amplification tests for detection of Chlamydia trachomatis in urine specimens. JClin Microbiol 2004. 423041–3045.3045 [PMC free article] [PubMed]
24. Koenig M G, Kosha S L, Doty B L. Direct comparison of the BD ProbeTec ET system with in‐house LightCycler PCR assays for detection of Chlamydia trachomatis and Neisseria gonorrhoeae from clinical specimens. Heath DG. J Clin Microbiol 2004. 425751–5756.5756 [PMC free article] [PubMed]
25. Harris K A, Hartley J C. 125 Development of broad‐range 16S rDNA PCR for use in the routine diagnostic clinical microbiology service. J Med Microbiol 2003. 8685–691.691 [PubMed]
26. Lamas C, Eykyn S J. Blood culture negative endocarditis: analysis of 63 cases presenting over 25 years. Heart 2003. 3258–262.262 [PMC free article] [PubMed]
27. Lampe M F, Suchland R J, Stamm W E. Nucleotide sequence of the variable domains within the major outer membrane protein gene from serovariants of Chlamydia trachomatis. Infect Immun 1993. 61213–219.219 [PMC free article] [PubMed]
28. Gerhardt P, Murray R Wood W. et al Methods for general and molecular bacteriology. Manual of methods for general bacteriology. Washington: ASM, 1994
29. Ripa K T, Mardh P A. Cultivation of Chlamydia trachomatis in cycloheximide‐treated McCoy cells. J Clin Microbiol 1977. 6328–331.331 [PMC free article] [PubMed]
30. Trevejo R T, Chomel B B, Kass P H. Evaluation of the polymerase chain reaction in comparison with other methods for the detection of Chlamydia psittaci. J Vet Diagn Invest 1999. 6491–496.496 [PubMed]
31. Farencena A, Comanduci M, Donati M. et al Characterization of a new isolate of Chlamydia trachomatis which lacks the common plasmid and has properties of biovar trachoma. Infect Immun 1997. 72965–2969.2969 [PMC free article] [PubMed]
32. Pickett M A, Everson J S, Pead P J. et al The plasmids of Chlamydia trachomatis and Chlamydophila pneumoniae: determination of copy number and the paradoxical effect of plasmid‐curing. Microbiology 2005. 3893–903.903 [PubMed]
33. Brunham R, Yang C, Maclean I. et al Chlamydia trachomatis from individuals in a sexually transmitted diseases core group exhibit frequent sequence variation in the major outer membrane protein (omp1) gene. J Clin Invest 1994. 94458–463.463 [PMC free article] [PubMed]
34. Dean D. Molecular characterization of new Chlamydia trachomatis serological variants from a trachoma endemic region of Africa. In:Orfila J, Byrne GI, Chernesky MA, Grayston JT, Jones RB, Ridgway GL, Saikku R, Schachter J, Stamm WE, Stephens RS, eds. Chlamydial infections. Bologna, Italy: Societa Editrice Esculapio, 1994. 259–262.262
35. Hayes L J, Bailey R L, Mabey D C. et al Genotyping of Chlamydia trachomatis from a trachoma‐endemic village in the Gambia by a nested polymerase chain reaction: identification of strain variants. J Infect Dis 1992. 51173–1177.1177 [PubMed]
36. Thein J, Zhao P, Liu H. et al Does clinical diagnosis indicate ocular chlamydial infection in areas with a low prevalence of trachoma? Ophthalmic Epidemiol 2002. 4263–269.269 [PubMed]
37. Ward M E. Genotyping of Chlamydia trachomatis from a trachoma‐endemic village in the Gambia by a nested polymerase chain reaction: identification of strain variants. J Infect Dis 1992. 51173–1177.1177 [PubMed]
38. West E S, Munoz B, Mkocha H. et al Mass treatment and the effect on the load of chlamydia trachomatis infection in a trachoma‐hyperendemic community. Invest Ophthalmol Vis Sci 2005. 183–87.87 [PubMed]
39. Ferreri A J, Guidoboni M, Ponzoni M. et al Evidence for an association between Chlamydia psittaci and ocular adnexal lymphomas. J Natl Cancer Inst 2004. 8586–594.594 [PubMed]
40. Taylor H, Guymer R. Exposure to Chlamydia pneumoniae infection and progression of age‐related macular degeneration. Am J Epidemiol 2005. 1611013–1019.1019 [PubMed]
41. Zhong G, Fan P, Ji H. et al Identification of a chlamydial protease‐like activity factor responsible for the degradation of host transcription factors. J Exp Med 2001. 193935–942.942 [PMC free article] [PubMed]
42. Arcari C M, Gaydos C A, Nieto F J. et al Association between Chlamydia pneumoniae and acute myocardial infarction in young men in the United States military: the importance of timing of exposure. Clin Infect Dis 2005. 81123–1130.1130 [PubMed]
43. Andraws R, Berger J S, Brown D L. Effects of antibiotic therapy on outcomes of patients with coronary artery disease: a meta‐analysis of randomised controlled trials. JAMA 2005. 212641–2647.2647 [PubMed]
44. Cannon C P, Braunwald E, McCabe C H. et al Pravastatin or Atorvastatin Evaluation and Infection Therapy‐Thrombolysis in Myocardial Infarction Investigators. Antibiotic treatment of Chlamydia pneumoniae after acute coronary syndrome. N Engl J Med 2005. 161646–1654.1654 [PubMed]
45. Grayston J T, Kronmal R A, Jackson L A. et al ACES Azithromycin for the secondary prevention of coronary. N Engl J Med 2005. 161637–1645.1645 [PubMed]
46. Sodeck G, Domanovits H, Khanakah G. et al The role of Chlamydia pneumoniae in human aortic disease‐a hypothesis revisited. Eur J Vasc Endovasc Surg 2004. 5547–552.552 [PubMed]
47. Centers for Disease Control and Prevention Recommendations for the prevention and management of Chlamydia trachomatis infections. Morb Mortal Wkly Rep 1993. 421–39.39 [PubMed]
48. Zhang J, Lietman T, Olinger L. et al Genetic diversity of Chlamydia trachomatis and the prevalence of trachoma. Pediatr Infect Dis J 2004. 3217–220.220 [PubMed]

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