Development of a New Chlamydiales-Specific Real-Time PCR and Its Application to Respiratory Clinical Samples ^,

Abstract

Originally composed of the single family Chlamydiaceae, the Chlamydiales order has extended considerably over the last several decades. Chlamydia-related bacteria were added and classified into six different families and family-level lineages: the Criblamydiaceae, Parachlamydiaceae, Piscichlamydiaceae, Rhabdochlamydiaceae, Simkaniaceae, and Waddliaceae. While several members of the Chlamydiaceae family are known pathogens, recent studies showed diverse associations of Chlamydia-related bacteria with human and animal infections. Some of these latter bacteria might be of medical importance since, given their ability to replicate in free-living amoebae, they may also replicate efficiently in other phagocytic cells, including cells of the innate immune system. Thus, a new Chlamydiales-specific real-time PCR targeting the conserved 16S rRNA gene was developed. This new molecular tool can detect at least five DNA copies and show very high specificity without cross-amplification from other bacterial clade DNA. The new PCR was validated with 128 clinical samples positive or negative for Chlamydia trachomatis or C. pneumoniae. Of 65 positive samples, 61 (93.8%) were found to be positive with the new PCR. The four discordant samples, retested with the original test, were determined to be negative or below detection limits. Then, the new PCR was applied to 422 nasopharyngeal swabs taken from children with or without pneumonia; a total of 48 (11.4%) samples were determined to be positive, and 45 of these were successfully sequenced. The majority of the sequences corresponded to Chlamydia-related bacteria and especially to members of the Parachlamydiaceae family.

INTRODUCTION

The Chlamydiales order contains obligate intracellular bacteria separated in 7 different families and family-level lineages, the Chlamydiaceae, the Criblamydiaceae, the Parachlamydiaceae, the Piscichlamydiaceae, the Rhabdochlamydiaceae, the Simkaniaceae, and the Waddliaceae (16, 23–25). Some of these bacteria are established pathogens and, for instance, Chlamydia trachomatis, C. psittaci, and C. pneumoniae from the Chlamydiaceae family can cause significant human infections. The other families constitute a group called Chlamydia-related bacteria (also referred to as Chlamydia-like organisms), which has been yet poorly investigated. Like the Chlamydiaceae, these Chlamydia-related bacteria are obligate intracellular bacteria that also exhibit a biphasic developmental cycle. Serological and molecular studies have implicated some species in various human and animal infections. Parachlamydia acanthamoebae is associated with human pneumonia (6, 12, 26, 27) and might cause bovine abortions (5, 38, 39), and Simkania negevensis might be responsible for respiratory infections, especially in children (18, 20, 22, 28, 32, 33, 35), whereas Waddlia chondrophila has been reported to cause abortion in bovines (14, 40) and is strongly suspected to cause miscarriage in humans (3, 4). Some of these newly discovered Chlamydia-related bacteria that resist digestion by several environmental amoebae are also resistant to professional phagocytes of the innate immune system such as macrophages. Considering their potential threat to human health, it is important to be able to detect these obligate intracellular bacteria, since classical culture methods are ineffective. Thus, quantitative real-time PCRs have been developed (6, 21, 26, 31, 42); however, they target specifically only one single species. Moreover, the only “broad-range” quantitative real-time PCR previously developed in the field is a family-specific PCR amplifying DNA from members of the Chlamydiaceae family, which will not allow the detection of Chlamydia-related bacteria (17). Since the biodiversity of Chlamydiales appears to be much larger than previously expected and new chlamydial strains are constantly discovered (7–9, 29, 30, 41), a molecular diagnostic tool able to detect any member of the Chlamydiales order is needed. Such a molecular tool would help in identifying the potential pathogenic role of Chlamydia-related bacteria and in specifying the true diversity of Chlamydiales, which is likely underestimated.

Thus, we developed a Chlamydiales-specific real-time TaqMan PCR (hereafter referred to as pan-Chlamydiales PCR), that we validated using 128 clinical samples available from previous studies. We also applied this new PCR to 422 nasopharyngeal swabs samples taken from children with or without pneumonia to investigate the presence of chlamydial DNA.

MATERIALS AND METHODS

DNA extraction.

Nasopharyngeal swab samples were extracted automatically with the LC automated system (Roche, Rotkreuz, Switzerland) and the MagNA Pure LC DNA isolation kit I (Roche). Extracted DNA was resuspended in 100 μl of the provided elution buffer. One negative extraction control was included for each extraction run (32 wells/extraction run).

Primers and probe.

Based on an alignment of the 16S rRNA sequences available in GenBank database (http://www.ncbi.nlm.nih.gov/GenBank/), specific primers and a probe were designed using the Geneious software 5.0.3 and primer3Plus (37). Locked nucleic acids (underlined in the sequences below) were added to ensure a higher specificity. We chose the primer forward panCh16F2 (5′-CCGCCAACACTGGGACT-3′), the primer reverse panCh16R2 (5′-GGAGTTAGCCGGTGCTTCTTTAC-3′), and the probe panCh16S (5′-FAM [6-carboxyfluorescein]-CTACGGGAGGCTGCAGTCGAGAATC-BHQ1 [Black Hole Quencher 1]-3′), targeting a fragment of about 207 to 215 bp in the 16S rRNA gene (the length was variable according to the species).

Real-time PCR assay.

PCR assays were performed in 20 μl, with iTaq Supermix with ROX (Bio-Rad, Reinach, Switzerland), 0.1 μM concentrations of each primer (Eurogentec, Seraing, Belgium), a 0.1 μM concentration of probe (Eurogentec), molecular-biology-grade water (Sigma-Aldrich, Buchs, Switzerland), and 5 μl of DNA sample. The cycling conditions were 3 min at 95°C, followed by 50 cycles of 15 s at 95°C, 15 s at 67°C, and 15 s at 72°C. The PCR products, tested in duplicate, were detected with a StepOne instrument (Applied Biosystems, Zug, Switzerland) for child nasopharyngeal swabs and with a ABI 7900 (Applied Biosystems) for analytic validation on samples from the retrospective study. Water was used as a negative PCR control.

Quantification and positive recombinant plasmid control.

DNA from Parachlamydia acanthamoebae strain Hall's coccus was isolated from a purified bacterial culture available in our laboratory, using a Wizard Genomic DNA purification kit (Promega, Duebendorf, Switzerland). A PCR was performed using the polymerase AmpliTaq Gold (Applied Biosystems) and the primers Pacstd16SF2 (5′-GCTGACGGCGTGGATGAGGC-3′) and Pacstd16SR2 (5′-CCTACGCGCCCTTTACGCCC-3′). The PCR products were purified with an MSB Spin PCRapace kit (Invitek, Berlin, Germany) and cloned according to the manufacturer's protocol in the pCR2.1-TOPO vector (Invitrogen, Basel, Switzerland) containing ampicillin and tetracycline resistance genes. Isolation of plasmid DNA was performed using a QIAprep spin miniprep kit (Qiagen, Kombrechtikon, Switzerland). The construction was checked by sequencing using primers of the pCR2.1-TOPO vector provided in the kit. Quantification of the recombinant plasmid was done on a Nanodrop ND-1000 (Witech, Littau, Switzerland), and 10-fold dilutions (10⁵ copies to 1 copy/μl) were used as positive controls to establish a standard curve for quantification and to check the reproducibility and efficiency of detection (see below). Negative controls, the standard curve, and samples were all analyzed in duplicate.

Analytical specificity, efficiency, and reproducibility of the PCR.

The specificity of the new quantitative PCR was tested using DNA extracted from different bacteria commonly found in respiratory tract samples (Table 1). DNAs were diluted at 10⁵ copies of the 16S rRNA gene per reaction. Using the positive control plasmid, the analytical sensitivity and the reproducibility of the PCR was assessed on duplicates with 10-fold dilutions (5 × 10⁵ to 5 copies/reaction) in 12 independent runs. The efficiency of detection was performed with the positive control plasmid diluted at 50, 20, 5, 1, and 0.5 DNA copies per reaction; each concentration was tested in 20 replicates. The range of the PCR was also evaluated with chlamydial DNA from 15 different strains (Table 2).

Table 1.

Bacterial and amoebal species used to test the specificity

Speciesa	Source or strain
Bacteria
Bacteroides fragilis*	ATCC25825
Escherichia coli*	ATCC 25922
Haemophilus influenzae*	ATCC 49247
Legionella pneumophila	Clinical specimen
Mycoplasma pneumoniae	Clinical specimen
Pseudomonas aeruginosa*	ATCC 27853
Staphylococcus aureus*	ATCC 25923
Streptococcus mitis	ATCC 6249
Streptococcus pneumoniae*	Clinical specimen
Amoebae
Acanthamoeba castellanii	ATCC 30010
Acanthamoeba comandoni	Strain WBT
Dictyostelium discoideum	DH1-10
Hartmannella vermiformis	ATCC 50237

^a*, Bacterial DNA used in the competition test with P. naegleriophila strain KNic.

Table 2.

Chlamydial DNA used to evaluate the range of the new PCR

Chlamydial species	Source or strain
Chlamydia abortus	Strain S26/3a
Chlamydia pecorum	Strain W73b
Chlamydia pneumoniae	Strain K6c
Chlamydia psittaci	Strain T49/90d
Chlamydia suis	Strain S45/6a
Chlamydia trachomatis	Clinical specimen
Criblamydia sequanensis	Strain CRIB-18
Estrella lausannensis	Strain CRIB-30
Neochlamydia hartmannellae	ATCC 50802
Parachlamydia acanthamoebae	Strain Hall's coccus
Parachlamydia acanthamoebae	ATCC VR-1476 (strain Bn9)
“Candidatus Protochlamydia amoebophila”	ATCC PRA-7 (strain UWE25)
Protochlamydia naegleriophila	Strain KNic
Simkania negevensis	ATCC VR-1471
Waddlia chondrophila	ATCC VR-1470

^aKindly provided by G. E. Jones, Moredun Research Institute, Edinburgh, United Kingdom.

^bKindly provided by J. Storz, Baton Rouge, LA.

^cKindly provided by A. Pospischil, Zürich, Switzerland.

^dKindly provided by R. K. Hoop, Zürich, Switzerland.

Clinical samples.

The new pan-Chlamydiales PCR was validated on 128 clinical samples. Different clinical samples, including urine, cervicovaginal, and anorectal samples and nasopharyngeal swabs, were collected, and DNA was extracted between 2004 and 2010 by the diagnostic laboratory of the Institute of Microbiology, Lausanne, Switzerland (Tables 3 and 4). These samples were originally tested with a real-time PCR specific for Chlamydia trachomatis (113 samples) (2, 13) and with a multiplex real-time PCR (42) detecting specifically Chlamydia pneumoniae but also Mycoplasma pneumoniae and Legionella pneumophila (15 samples). Positive samples for M. pneumoniae (5 samples) or L. pneumophila (3 samples) were included to confirm the high specificity of the new real-time PCR. We then applied our new pan-Chlamydiales PCR to 422 nasopharyngeal swabs prospectively collected between 2008 and 2010 at the University Hospitals of Geneva from children with (n = 265) or without (n = 157) pneumonia. Pneumonia was defined by the presence of at least one of the following symptoms: fever (>38°C), cough, dyspnea, tachypnea, and an infiltrate or a consolidation visible on a lung X-ray image. All samples were systematically tested for the following viruses by PCR: human respiratory syncytial viruses (HRSV) A and B; adenoviruses A, B, C, and E; coronaviruses HKU1, OC43, 229E, and NL63; parainfluenza viruses 1, 2, and 3; human metapneumovirus (HMPV) A and B; enteroviruses A, B, C, and D; rhinoviruses A and B; influenza viruses A and B; and H1N1 virus for some samples during the 2009 epidemics. In addition, the nasopharyngeal samples were tested by PCR for the presence of Streptococcus pneumoniae and by real-time PCR for Mycoplasma pneumoniae and Legionella pneumophila. The children were between 1 and 15 years old; the median ages for the pneumonia and control groups were 4.6 and 6.2 years old, respectively. All DNA samples were tested in duplicate. Positive samples were systematically confirmed in a second run. To test for potential false-negative results due to PCR inhibitors, an inhibition test was systematically performed with 4 μl of clinical DNA samples and 1 μl of the positive control at a concentration of 200 DNA copies/μl. The PCR was considered inhibited when the quantification was less than 50 DNA copies per reaction (4-fold reduction). Moreover, a total of 60 uninoculated swabs (Copan, Brescia, Italy) were used as an additional negative control to confirm that the commercial swabs used in the prospective study were not contaminated with any chlamydial DNA.

Table 3.

Analysis of samples from various origins by pan-Chlamydiales PCR compared to C. pneumoniae PCR

Sample type	No. of samples
	C. pneumoniae (n = 15)		Pan-Chlamydiales (n = 15)
	+	–a	+	–
Nasopharyngeal swab	1	11/0	1	1
Bronchoalveolar lavage	0	62/1	0	6
Bronchial aspirate	0	40/2	0	4
Sputum	1	22/0	1	2
Total	2	13	2	13

^aSuperscript numbers indicate the number of positive Mycoplasma pneumoniae samples/the number of positive Legionella pneumophila samples.

Table 4.

Analysis of samples from various origins by pan-Chlamydiales PCR compared to C. trachomatis PCR

Sample type	No. of samplesa
	C. trachomatis (n = 113)		Pan-Chlamydiales (n = 113)
	+	–	+	–
Vaginal or cervical swab	14	15	14	15
Anorectal swab	14	0	12	2**
Urethral swab	1	0	1	0
Eye swab	1	1	1	1
Urine specimen	32	33	31 (+1)b	32 (+1)*
Ascites liquid	1	1	0	2*
Total	63	50	60	53

^aOne (*) or two (**) sample(s) were positive for C. trachomatis but negative with the pan-Chlamydiales PCR.

^bOne sample was negative for C. trachomatis but positive with the pan-Chlamydiales PCR.

Sequencing of positive samples.

Amplicons of positive samples were purified using an MSB Spin PCRapace kit. A sequencing PCR was performed with the specifically designed inner primers panFseq (5′-CCAACACTGGGACTGAGA-3′) and panRseq (5′-GCCGGTGCTTCTTTAC-3′). The sequencing PCR assay was done using a BigDye Terminator v1.1 cycle sequencing kit (Applied Biosystems). Sequences of positive nasopharyngeal samples taken from children have been deposited on the NCBI website. The accession numbers are HQ721193 to HQ721240.

RESULTS

Sensitivity and specificity of pan-Chlamydiales quantitative PCR.

No cross-reaction was observed with the different bacterial or amoebal strains tested (Table 1). A competition test was also performed with increasing amounts of DNA from Protochlamydia naegleriophila strain KNic (from 0 to 10³ copies of the 16S rRNA gene per reaction) in the presence of an increasing amounts of a mixture of nonchlamydial DNA (Table 1) (from 0 to 10⁶ copies of the 16S rRNA gene per reaction). The amplification of the DNA from P. naegleriophila strain KNic was not affected by competing nonchlamydial DNA at up to 10⁵ copies of the 16S rRNA gene of nontargeted bacteria, demonstrating the high specificity of the PCR. The range of the new PCR was evaluated with 15 DNAs from different chlamydial strains (Table 2). As expected, all of the different members of the Chlamydiales order tested were detected, confirming the large range of the PCR. Despite the presence of one mismatch with the probe in the 16S ribosomal DNA (rDNA) sequence of C. psittaci and C. abortus, both species were successfully amplified. Alignment of all other sequences available from members of the Chlamydiales order demonstrated that one mismatch is also present for C. caviae, C. felis, and “Candidatus Clavochlamydia salmonicola” in the probe or for Rhabdochlamydia porcellionis and R. crassificans in the forward primer. These species are nevertheless likely all amplified with our new pan-Chlamydiales PCR. Indeed, numerous DNAs somehow related to Rhabdochlamydiaceae have been successfully amplified from clinical samples (see below). The only known member of the Chlamydiales order likely not amplified using our pan-Chlamydiales PCR is Piscichlamydia salmonis, since as many as six mismatches are present.

Reproducibility and efficiency of the pan-Chlamydiales real-time PCR.

The inter-run and intra-run reproducibility was assessed, respectively, on 12 independent runs and 72 duplicates (Fig. 1A and B). All duplicates were amplified for 50 or more DNA copies per reaction and 18 replicates of 24 (75%) for 5 DNA copies per reaction. The Bland-Altman graph clearly indicates that differences between duplicates were below one cycle threshold (C_T) for DNA copies above 50 per reaction, demonstrating a high reproducibility. The efficiency of detection was evaluated on 20 replicates for 50, 20, 5, 1, and 0.5 DNA copies per reaction. The PCR showed 100% detection for 50 and 20 DNA copies, and 75, 30, and 5% for 5, 1, and 0.5 DNA copies per reaction, respectively (Fig. 1C).

Fig. 1.

Reproducibility and efficiency of the new real-time PCR. Inter- and intra-run reproducibility was evaluated among 12 different runs representing 72 duplicates of a positive control. (A) Inter-run variability. (B) Bland-Altman graph showing the intra-run variability between duplicates. A bias of 0.36 was calculated, as well as a 95% limit of agreement (indicated by the dashed line). (C) The efficiency was evaluated with 20 replicates of five different plasmid control concentration (50, 20, 5, 1, and 0.5 copies per reaction).

Analytical validation of the new PCR.

Of the 65 samples positive for Chlamydia trachomatis or C. pneumoniae, 61 (93.8%) samples were determined to be positive with the new PCR (Tables 3 and 4). The four discordant samples were originally positive for C. trachomatis, from anorectal swabs (n = 2), a urine specimen (n = 1), and ascites liquid (n = 1). These four samples were tested a second time with the original test (C. trachomatis real-time PCR) and were found to be negative (n = 1) or positive with only 0.2, 6, and 1.2 copies per reaction, which was most certainly below the detection limits of the pan-Chlamydiales PCR. Seven positive samples with the pan-Chlamydiales PCR (C_T = 23.6 to 41.3) were sequenced to confirm the results. All of the sequences obtained showed 100% similarity with the expected species (see Table S1 in the supplemental material), confirming the specificity of the new PCR and the possible identification by sequencing even with later C_T values. Positive samples for Mycoplasma pneumoniae (n = 5) and Legionella pneumophila (n = 3) were all determined to be negative with the new PCR (Table 3). Of the 63 samples found to be negative for C. trachomatis or C. pneumoniae, only 1 was amplified, showing 92% similarity to the closest previously described Protochlamydia naegleriophila strain CRIB 41 (FJ532294.1) (Table 4).

Application of the quantitative PCR.

Application of the new pan-Chlamydiales PCR using 422 nasopharyngeal swabs samples taken from children yielded 48 positive samples: 31 (7.3%) samples with 1/4 positive wells, 6 (1.4%) samples with 2/4 positive wells, 4 (0.9%) samples with 3/4 positive wells, and 7 (1.7%) samples with 4/4 positive wells (see Table S2 in the supplemental material). A correlation between the cycle threshold value (C_T) and the number of positive wells was observed (see Fig. S1 in the supplemental material). Samples with <5 DNA copies per reaction (high C_T values) were amplified in 3/4, 2/4, and 1/4 wells. The 48 positive samples were sequenced, and 48 sequences were obtained from 45 different patients (see Table S2 in the supplemental material). Indeed, the sequencing of three samples failed and for three other samples, two different sequences were obtained (patients GE10169, HE210023, and HE210045, see Table S2 in the supplemental material). Thus, 94% of the positive samples were successfully sequenced. Patients' characteristics and sequencing results for patients with pneumonia are presented in Table 5. Of the 25 patients listed in Table 5, another etiology was identified for only 8.

Table 5.

Sequencing results of nasopharyngeal samples from the pneumonia group positive with the new pan-Chlamydiales PCR

Patient no.	Sexa	Age (yr)	Signs and symptomsb	Other etiology	Underlying condition(s)	% 16S rRNA gene homology with most similar GenBank sequence (corresponding family)c
GE10160	F	3.7	39.0°C, cough, thoracic pain, RDS		coeliakie	100% Chlamydia pneumoniae LPCoLN (Ch)
GE10097	F	2.7	40.6°C, cough, RDS			100% Chlamydia pneumoniae LPCoLN (Ch)
VS30014	M	12.4	38.9°C, cough			100% Chlamydia pneumoniae LPCoLN (Ch)
VS30030	F	12.2	39.5°C, cough			100% Chlamydia trachomatis D-LC (Ch)
GE10098	F	8.0	38.0°C, cough			100% Chlamydia pneumoniae CWL029 (Ch)
GE10014	M	7.6	39.5°C, cough	M. pneumoniae		94% uncultured Neochlamydia sp. strain LTUNC09656 (P)
GE10159	M	3.7	40.0°C, cough			100% Chlamydia pneumoniae LPCoLN (Ch)
GE10169	M	5.6	40.0°C, cough, thoracic pain, tachypnea		Bronchodysplasia, premature birth (28 weeks)	97% “Candidatus Rhabdochlamydia porcellionis” (R); 91% Chlamydiales bacterium cvE38 (S)
GE10179	F	3.6	Dyspnea	S. pneumoniae H1N1 virus		Sequencing failed
HE20032	M	1.5	41.6°C, cough, RDS			94% uncultured “Candidatus Protochlamydia sp.” clone CN823 (P)
VS30003	M	5.8	39.5°C, cough, RDS			95% uncultured Chlamydiales bacterium clone P-5 (P)
GE10027	M	9.8	38.1°C, cough, RDS	M. pneumoniae	asthma	92% uncultured soil bacterium clone 530-2 (Cr)
GE10036	M	1.6	38.0°C, cough, RDS, tachypnea	S. pneumoniae		95% uncultured bacterium clone F5K2Q4C04JDDHX (P)
GE10047	M	2.6	39.5°C, cough, RDS, tachypnea			92% Criblamydia sequanensis (Cr)
GE10072	F	4.8	39.5°C	HMPV A		Sequencing failed
GE10147	M	13.8	38.2°C, cough, RDS	S. pneumoniae		95% uncultured bacterium clone FW1013-189 (P)
GE10193	F	5.1	38.9°C, cough, tachypnea			92% Chlamydiales bacterium cvE38 (S)
HE20008	F	3.3	38.3°C, cough, tachypnea, RDS	HUK1		93% Estrella lausannensis strain CRIB 30 (Cr)
HE20028	F	1.1	39.5°C, cough, RDS, tachypnea	HRSV A		94% uncultured Chlamydiales bacterium clone P-5 (P)
HE20036	M	1.4	40.0°C, RDS			96% Chlamydiales bacterium cvE21 (E6)
HE20074	M	4.5	38.1°C, cough, RDS			94% Criblamydia sequanensis (Cr)
VS30007	M	6.6	38.7°C, cough, RDS			95% “Candidatus Metachlamydia lacustris” strain CHSL (P)
VS30013	M	6.4	38.5°C, cough, tachypnea			94% uncultured Chlamydiales bacterium clone P-9 (P)
VS30044	M	3.8	40.4°C, cough			95% uncultured Chlamydiales bacterium clone P-7 (P)
VS30055	F	4.1	38.5°C, cough, tachypnea			92% “Candidatus Metachlamydia lacustris” strain CHSL (P)

^aF, female; M, male.

^bRDS, respiratory distress syndrome.

^cCh, Chlamydiaceae; P, Parachlamydiaceae; R, Rhabdochlamydiaceae; S, Simkaniaceae; Cr, Criblamydiaceae; E6, novel E6 lineage.

A percentage of similarity for the best BLAST of >90% was observed for all 48 sequences obtained, allowing identification at least at the family level. Of the 48 sequences obtained, 26 belonged to the Parachlamydiaceae family, 7 belonged to the Chlamydiaceae family, 5 belonged to the Simkaniaceae family, 5 belonged to the Criblamydiaceae family, 3 belonged to the Rhabdochlamydiaceae family, 1 seemed to belong to the novel E6 lineage (7, 11), and 1 other sequence corresponded to an unclassified Chlamydiales (see Table S2 in the supplemental material). Of the 7 sequences corresponding to a Chlamydiaceae species, 6 demonstrated 100% similarity with Chlamydia pneumoniae: five samples were taken from children with pneumonia (Table 5), whereas one was taken from an apparently healthy child (see Table S2 in the supplemental material). This latter patient had a history of obstructive bronchitis and chronic otitis media. The remaining Chlamydiaceae sequence showed 100% similarity with C. trachomatis (the sample was taken from a child with pneumonia) (Table 5). Of the 26 sequences corresponding to a member of the Parachlamydiaceae family, 10 (40%) were taken from 10 patients with pneumonia, and 16 (60%) were from 15 patients from the control group (patient HE210023 being positive for two different bacteria). These latter patients were positive in 4/4, 3/4, 2/4, and 1/4 wells, respectively, for 2, 2, 3, and 18 nasopharyngeal swabs (see Table S2 in the supplemental material). Criblamydiaceae species were recovered from four patients with pneumonia (all with 1/4 positive well) and one patient from the control group (4/4 positive wells). Simkaniaceae species were found in five patients (three control patients and two children with pneumonia). Finally, Rhabdochlamydiaceae species were identified in one case of pneumonia and two control subjects. Thus, 17 and 20 children with and without pneumonia, respectively, were positive for a Chlamydia-related bacterium. In addition, a sample taken from a control subject could not be affiliated in any range of family-level lineage (unclassified Chlamydiales).

All 60 uninoculated Copan swabs were determined to be negative with the pan-Chlamydiales PCR, demonstrating that the positive samples were not false positives. Furthermore, no absence of the internal amplification control was observed, excluding false-negative results due to PCR inhibitors.

DISCUSSION

In this study, we developed a new Chlamydiales-specific PCR that was specific to the Chlamydiales order, sensitive for at least 5 DNA copies per reaction of the positive control (with an efficiency of 75%), and highly reproducible. Moreover, its application to clinical samples taken from children with or without pneumonia demonstrated the common exposure of humans to various Chlamydia-related bacteria. This new PCR showed a broad range of targeted species since it detected the 15 different chlamydial strains tested and the DNAs of 36 never-described species-level lineages (<97% similarity of the 16S rDNA sequence) of the Chlamydiales order (>80% similarity of the 16S rDNA sequence) (16, 25) present in nasopharyngeal swabs samples (see Table S2 in the supplemental material). Furthermore, this new PCR could detect chlamydial DNA from samples of various origins (Tables 3 and 4).

Other classical pan-Chlamydiales PCRs have been developed (10, 36, 43), but they detected about 1,000 DNA copies per reaction compared to real-time PCRs that can detect about 200- to 1,000-fold less DNA copies per reaction. This higher sensitivity is likely due to shorter reads (∼200 bp) and readout due to the use of a fluorescent TaqMan probe. Considering this high sensitivity, DNA extraction, real-time PCR, and sequencing reactions were processed in separate rooms to avoid contaminations between samples. In addition, automated DNA extraction at locations outside our research laboratory was preferred. Moreover, since no sequence obtained showed more than 97% similarity to bacteria grown in our laboratory, contamination may not explain the results we obtained. The sequencing of most positive samples was possible, and the results were informative at the family level. A previous study using short sequences of Chlamydiales also successfully identified strains at the family level with sequences of similar lengths (140 to 195 bp) (43). Further identification, at the species level, may be performed using complementary methods (PCR targeting of a more discriminative core gene, such as rpoB or gyrA).

Since previous studies on nasal and/or nasopharyngeal samples have already allowed the recovery of Chlamydia-related bacteria or the amplification of DNA of these obligate intracellular bacteria (1, 12, 15, 36), we chose similar samples for the first application of the new PCR. The sequencing results obtained using these nasopharyngeal swabs confirmed previous studies on the occurrence of Chlamydia-related bacteria in nasal mucosa of healthy individuals (1). They also clearly showed that the biodiversity of Chlamydia-related bacteria is far from established: of the 48 sequences, 36 were from putative new species (using the Everett cutoff of <97% 16S rRNA similarity to define species-level lineages), and all belonged to the Chlamydiales order (>80% similarity of the 16S rDNA sequence) (16, 23, 24). Thus, our work clearly demonstrates the common exposure of children to different Chlamydiales strains, since ca. 11.4% of the patients were found to be positive by the new PCR. When a Chlamydiaceae species was amplified, it was generally from a sample taken from a child with pneumonia (6/7). It is interesting that Criblamydiaceae DNA was also mainly amplified from patients with pneumonia (4/5), whereas other Chlamydia-related bacteria were amplified from nasopharyngeal swabs obtained from children both with and without pneumonia. Thus, although our study demonstrated a common exposure to Parachlamydiaceae (amplified from 5.9% of all samples), this family was not overrepresented in the pneumonia group. Nonetheless, since the sequencing does not allow identification at the species level, a significant correlation with a given species may not be excluded. Similarly, our work did not demonstrate an association of Simkaniaceae with pneumonia in children. This was expected, since initial studies that suggested an association of Simkania negevensis with diverse respiratory infections in children (15, 18–20, 22, 32, 33, 35) were not confirmed in more recent works (34). Further research is now needed to determine the specific pathogenic role of each species in the Chlamydiales order.

In conclusion, we provide here a new diagnostic approach to show the biodiversity and pathogenic role of Chlamydia-related bacteria and highlight the common exposure of children to Parachlamydiaceae.