JOURNAL OF CLINICAL MICROBIOLOGY, Nov. 2003, p. 4915–4923 0095-1137/03/$08.00⫹0 DOI: 10.1128/JCM.41.11.4915–4923.2003 Copyright © 2003, American Society for Microbiology. All Rights Reserved.
Vol. 41, No. 11
MINIREVIEW Molecular Diagnosis of Mycoplasma pneumoniae Respiratory Tract Infections K. Loens,1* D. Ursi,1 H. Goossens,1,2 and M. Ieven1 Medical Microbiology, Universitaire Instelling Antwerpen, B2610 Wilrijk, Belgium,1 and Department of Medical Microbiology, Leiden University Medical Center, Leiden, The Netherlands2 eral NAATs targeting different genes to detect M. pneumoniae have been adequately evaluated. Since NAATs targeting DNA can detect both viable and nonviable organisms, detecting RNA by reverse transcriptase PCR (RT-PCR) or nucleic acid sequence-based amplification (NASBA) may be a useful method to identify productive M. pneumoniae infections. The possible long-term carrier state of M. pneumoniae in the respiratory tract may hinder the evaluation of different diagnostic tests for the diagnosis of acute infections. An overview of the peer-reviewed literature on the use of NAATs to detect M. pneumoniae since 1989 is given. Search combinations were M. pneumoniae and PCR, M. pneumoniae and diagnosis, and M. pneumoniae and amplification. This minireview describes the molecular biology-based amplification methods to detect M. pneumoniae that are currently available. Topics discussed include specimen collection and transport, preparation of nucleic acid from clinical specimens, choice of the target sequence, and detection of the amplicons. Methods to recognize and prevent false-positive and false-negative results, the results of NAATs in comparison with results obtained by conventional diagnostic tests, and clinical applications are also reviewed.
Mycoplasma pneumoniae is responsible for 10 to 20% of the cases of community-acquired pneumonia and has been associated with acute exacerbations of asthma (22). M. pneumoniae is also implicated in mild acute respiratory infections, such as sore throat, pharyngitis, rhinitis, and tracheobronchitis (2). Correct diagnosis of M. pneumoniae infections is important to allow the appropriate antibiotic treatment of patients, since it is impossible to identify a M. pneumoniae infection solely on the basis of clinical signs and symptoms. It should decrease inappropriate use of antibiotics, influence the patient outcome by reduction of morbidity and mortality, and improve our knowledge of the prevalence of the causes of so-called atypical pneumonia. Conventional assays for the detection of M. pneumoniae have their limitations, resulting in the need for more accurate diagnostic methods. Culture is time-consuming and relatively insensitive, because M. pneumoniae grows slowly in vitro, requiring 2 to 5 weeks for colonies to become visible. Serological methods, particularly the complement fixation (CF) test, are most widely used. The sensitivity of these assays depends on whether the first serum sample is collected early or late after the onset of disease and on the availability of paired serum samples collected with an interval of 2 to 3 weeks. Immunoglobulin M (IgM) assays which are more sensitive than the CF test have been developed, but the IgM response may be nonspecific (61) or absent, particularly in adults (70). Hybridization with DNA probes has also been proposed as a rapid and specific procedure to replace culture, but it lacks sensitivity (35). Nucleic acid amplification techniques (NAATs) have the potential to produce rapid, sensitive, and specific results, allowing early appropriate antibiotic therapy. In the absence of a reference method, the so-called “gold standard” for the diagnosis of an M. pneumoniae infection, either an expanded gold standard or the technique of latent class analysis (LCA) should be applied to calculate the sensitivity and specificity of the available diagnostic tests. The technique of LCA can be used if at least three independent techniques can be compared. Thus far, only a few PCR tests and a limited number of studies applying culture, serology, and sev-
TECHNICAL ASPECTS Specimen collection. Specimens suitable for the detection of M. pneumoniae include sputum (74) and bronchoalveolar lavage (BAL) specimens (40, 66), nasopharyngeal and throat swabs (25, 78), nasopharyngeal aspirates (20, 28, 36), tracheal aspirates (1, 26), pleural fluid specimens (52), and transthoracic needle aspirations (17). More unusual specimens, such as nasal polyps (29), open-lung biopsies, and autopsy specimens (65), have also been tested. Gnarpe et al. (25) compared nasopharyngeal and throat swabs for the detection of M. pneumoniae and found throat swabs to be superior to nasopharyngeal swabs. Honda et al. (34) applied capillary PCR to sputum and BAL specimens and throat swabs. Review of the differences in PCR positivity rates as a function of the type of specimens collected showed the highest rate of detection from throat swabs (28.6%). However, there were some problems with proper collection of throat swab specimens due to inadequate scraping of the mucosal surface, resulting in false-negative results due to the collection of an insufficient amount of DNA. The positivity rate was
* Corresponding author. Mailing address: Medical Microbiology, Universitaire Instelling Antwerpen, Universiteitsplein 1 S3, B2610 Wilrijk, Belgium. Phone: 3238202551. Fax: 3238202663. E-mail:
[email protected]. 4915
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21.5% for BAL specimens and 14.2% for sputum specimens. The low positivity rate for sputum specimens was attributed to the absence of sputum in many patients with pneumonia due to M. pneumoniae. Transport of specimens. Specimens should be transported to the laboratory as soon as possible and stored at 4°C or frozen at ⫺70°C. RNA specimens should be processed or frozen at ⫺70°C as soon as possible to prevent RNA degradation (6, 47). Processing of specimens. Different extraction methods with and without sample pretreatment have been described: (i) dilution of the sample with 0.9% sodium chloride, followed by the addition of sodium dodecyl sulfate, extraction with phenolchloroform, and precipitation with ammonium acetate and ethanol (28); (ii) pretreatment with proteinase K (15), followed by phenol-chloroform or phenol-chloroform-isoamyl alcohol extraction and ethanol precipitation (10); (iii) Boom extraction (5) on protease-treated (47) or untreated samples (62); (iv) treatment with Sputazyme (Kobayashi Pharmaceutical Co., Tokyo, Japan), followed by proteinase K digestion (34); (v) incubation with Chelex (Bio-Rad Laboratories, Richmond, Calif.) and sodium azide (40); (vi) treatment by sonication or boiling (7); (vii) phenol-chloroform-isoamyl alcohol extraction, followed by ether extraction (79); and (viii) phenolchloroform extraction and precipitation by sodium acetate or sodium chloride (17). Extraction methods using commercial kits are available: the Boehringer High Pure viral nucleic acid kit (Boehringer, Mannheim, Germany) (28); the QIAamp blood kit (QIAgen, Hilden, Germany) (69); cell lysis with sodium dodecyl sulfate and proteinase K, followed by purification with QIAamp DNA binding columns (QIAgen) (31); and the Amplicor sputum sample preparation kit (Roche Diagnostic Systems Inc., Branchbury, N.J.) (25), which was found to increase the sensitivity of a Chlamydia pneumoniae PCR, due to a more complete lysis of cells in the specimens (24). Fahle and Fischer (16) evaluated six commercially available DNA extraction kits for their ability to recover DNA from various dilutions of cytomegalovirus (CMV) added to BAL, cerebrospinal fluid, plasma, or whole-blood specimens. The kits evaluated included the Puregene DNA isolation kit (Gentra Systems Inc., Minneapolis, Minn.), the Generation capture column kit (Gentra Systems Inc.), the MasterPure DNA purification kit (Epicentre Technologies, Madison, Wis.), the IsoQuick nucleic acid extraction kit (MicroProbe Corp., Bothell, Wash.), the QIAamp blood kit (QIAgen), and the NucliSens extraction kit (Organon Teknika Corp., Durham, N.C.). All six kits evaluated effectively removed PCR inhibitors from each of the four specimen types and produced consistently positive results. However, the NucliSens extraction kit and the Puregene DNA isolation kit had the most consistently positive results at the lowest concentrations of spiked CMV and were the most sensitive methods for extracting CMV DNA from the four different kinds of spiked specimens. In contrast, there are very few studies comparing different methods for M. pneumoniae nucleic acid extraction (1, 36). Abele-Horn et al. (1) compared the efficacy of DNA extraction by cell lysis with proteinase K without further nucleic acid purification and DNA extraction after lysis by phenol-chloroform followed by ethanol precipitation. A 10-fold dilution se-
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ries of M. pneumoniae was used. The researchers reported that the phenol-chloroform DNA extraction technique was timeconsuming and resulted in a 10-fold decrease in sensitivity. In a study by Ieven et al. (36), 371 nasopharyngeal aspirates from children with acute respiratory infections were examined for the presence of M. pneumoniae by culture and several different PCR protocols in two laboratories. Each laboratory applied one sample preparation method: (i) freeze-boiling (method A) or (ii) isothiocyanate treatment, followed by phenol chloroform extraction (method B). Prepared samples were exchanged between the two laboratories. In both laboratories, identical primers were used in a PCR directed at the P1 gene. A specific internal control for P1 amplification was included. After sample preparation method A, laboratory 1 identified 9 positive samples of 13 samples and identified 2 more samples after diluting them 1/10 to eliminate polymerase inhibitors. One additional positive sample was identified after hybridization. Laboratory 2, using the same material, obtained similar results. After sample preparation method B, 12 positive samples were detected with the primers directed at the P1 adhesin gene in laboratory 1, and 13 positive samples were detected in laboratory 2 after hybridization. It was concluded that, provided a specific internal control is used, sample preparation by freeze-boiling could be recommended. Target regions and amplification protocols. Several regions in the M. pneumoniae genome have been used to detect and identify M. pneumoniae by PCR or other amplification techniques (Table 1). Van Kuppeveld et al. targeted a fragment of the 16S rRNA for genus and species identification of M. pneumoniae (73, 74). They compared PCR with RT-PCR, and a 1,000-fold increase in sensitivity was found when rRNA instead of ribosomal DNA (rDNA) was used as a target (73). When the 16S rRNA PCR was compared with culture and serological methods for the diagnosis of M. pneumoniae infections, it was concluded that PCR was the optimal approach (74). Tjhie et al. used the same primers in a comparison of direct PCR and RT-PCR (67), culture, and serology. A positive correlation between the direct PCR and serology, as tested by the microparticle agglutination assay, was found in 88.1% of the cases: PCR and serology gave positive results for 6 of 59 (10.2%) of patients, 3 of 59 (5.1%) patients were positive for M. pneumoniae only by PCR, and 4 of 59 (6.8%) patients were positive for M. pneumoniae only by serology. Kessler et al. designed a PCR targeted at the 16S rDNA; the amplicons were identified by probe hybridization in a nonradioactive microwell plate format (40). When applied to BAL specimens, 12 specimens were found positive by PCR and serology, of which 7 were subsequently confirmed by culture. Bernet et al. selected a specific DNA sequence from a genomic library and chose two oligonucleotides in this sequence to produce an amplicon of 144 bp (3). Analysis of clinical samples showed that PCR was more sensitive than culture for the detection of M. pneumoniae. These results were confirmed by Skakni et al. (62) but were in contrast with those obtained by Falguera et al. (17) who applied the assay to transthoracic needle aspiration specimens. Vekris et al. (75) developed a microtiter hybridization assay to detect ATPase amplicons, generated by the Bernet primers (3), and found it
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TABLE 1. Nucleic acid amplification assays developed in-house for the detection of M. pneumoniae by year of publication Authors
Yr of publication
Reference
Gene target
Product size (bp)
Assay typea
Bernet et al. Jenssen et al. Jenssen et al. Buck et al. Sasaki et al. Ursi et al. Van Kuppeveld et al. Williamson et al. Williamson et al. Cadieux et al. De Barbeyrac et al. Kai et al. Lu ¨neberg et al. Zingangirova et al. Leng et al. Tjhie et al. Fink et al. Ovyn et al. Ramirez et al. Stone et al. Kessler et al. Abele-Horn et al. Narita et al. Corsaro et al. Dorigo-Zetsma et al. Gro ¨ndahl et al. Layani-Milon et al. Tong et al. Hardegger et al. Honda et al. Kong et al. Waring et al. Loens et al. Welti et al.
1989 1989 1989 1992 1992 1992 1992 1992 1992 1993 1993 1993 1993 1993 1994 1994 1995 1996 1996 1996 1997 1998 1998 1999 1999 1999 1999 1999 2000 2000 2000 2001 2002 2003
3 38 38 7 60 72 73 81 81 9 11 39 48 84 46 67 18 55 57 63 40 1 52 10 14 28 45 69 31 34 43 78 47 80
ATPase operon gene P1 gene 16S rRNA gene P1 gene 16S rRNA gene P1 gene 16S rRNA P1 gene 16S rRNA gene P1 gene P1 gene 16S rRNA gene tuf gene P1 gene P1 gene 16S rRNA gene P1 gene 16S rRNA P1 gene 16S rRNA 16S rRNA gene ATPase operon gene ATPase operon gene ATPase operon gene P1 gene 16S rRNA P1 gene P1 gene P1 gene ATPase operon gene P1 gene P1 gene 16S rRNA P1 gene
144 153 584 375 809 209 277 543 290 345 466 88 950 245, 210 631 277 466, 183 190 375 NSb 427 144, 104 144, 108 144 272, 133 277 466 345 76 250 111–154 309–339 190 76
S⫹H S⫹A S⫹A S⫹H S⫹H S⫹A RT ⫹ H S⫹H S⫹H M⫹H S ⫹ H/RE S⫹A S⫹H N⫹A S⫹A S⫹H N⫹A NA ⫹ H S⫹A Q S⫹H N⫹A N⫹A M⫹A N⫹A M⫹H S⫹H M⫹H R C⫹A N⫹A S⫹H NA ⫹ H M⫹R
a Abbreviations: A, Agarose gel electrophoresis; C, capillary PCR; H, hybridization; M, multiplex PCR; N, (semi)nested PCR; NA, NASBA; Q, Q replicase; R, Real-time PCR; RE, restriction enzyme digestion; RT, RT-PCR; S, single-step PCR. b NS, not specified.
to be cheaper, involving fewer steps and allowing easy handling of a large number of specimens. Another frequently used target for M. pneumoniae PCR is the P1 adhesin gene. The target sequence for amplification in the assay designed by Buck et al. is a 375-bp segment of this gene (7). The assay proved to be more sensitive than culture and the Gen-Probe assay. De Barbeyrac et al. generated a 466-bp fragment of the P1 gene with primers MP-P11 and MP-P12. Identification was performed by restriction enzyme digestion (11). Leng et al. (46) designed an assay resulting in a 631-bp amplicon and used the primers described by Bernet et al. (3) for comparison. The assays were applied on throat swabs; the latter protocol was the most sensitive. The PCR protocol described by Ursi et al. (72) was compared with culture and serological tests by Dorigo-Zetsma et al. (15). They concluded that PCR results could be added to the criteria for the diagnosis of M. pneumoniae infections and could even replace culture; PCR and the CF test were found to be complementary. The system developed by Lu ¨neberg et al. (48) is based on the gene encoding the elongation factor Tu (tuf) targeted by primers Mpn 38 and Mpn 39 and detected by probe Mpn 46. Compared with culture and serology, the assay had a sensitivity
of 90 and 83%, respectively. Compared with serology, the specificity was 97%. Studies comparing the performance of PCR methods with different M. pneumoniae target regions and primers are extremely rare. Ieven et al. (36) compared the PCR assay targeted at the P1 adhesin gene described by Ursi et al. (72) with the PCR assay described by Van Kuppeveld et al. (74). To confirm the identity of the amplicons, probe hybridization was performed. The P1 adhesin gene primers were found to be more sensitive than the 16S rRNA primers. This most likely results from the P1 cytadhesin gene being present in multiple copies (64). Three different PCR assays were compared by Abele-Horn et al. (1): (i and ii) the assay originally described by Bernet (3), with and without an additional hybridization step for amplicon detection, and (iii) a newly developed nested PCR format. The PCR was performed directly on the specimens or after overnight incubation in Hayflick broth. All three PCR assays proved to be reliable in detecting M. pneumoniae in respiratory specimens, but the nested format was the most sensitive one. Comparison of sensitivity data from different clinical studies is complicated by differences in sample collection, transporta-
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tion, and extraction procedures, input sample volumes, target genes, primers, cycling parameters, and detection systems. Furthermore, there is great variation in the units applied to measure the detection limits: CFU, color-changing units (CCU) (1 CCU corresponds to 10 to 100 organisms [3]), number of cells, or quantity of DNA. This makes a comparison of the sensitivity of different assays very difficult. For example, Abele-Horn et al. (1) reported the sensitivity of their assay to be 3,000 genome copies, 30 pg of DNA, 19 CFU, or 1.9 ⫻103 organisms. Furthermore, different definitions for 1 CFU are used: for example, in one study, 1 CFU corresponds to 160 organisms (32), whereas in another study, 1 CFU corresponds to 10 to 1,000 organisms (58). More importantly, the lack of a consensus method for appropriate evaluation of the different methods precludes comparisons. In conclusion, standardized highly sensitive and specific NAATs are sorely needed. Multiplex PCR. Multiplex PCRs to detect two or three different respiratory pathogens have been developed by some groups (9, 10, 69). Gro ¨ndahl et al. developed a multiplex RTPCR to detect nine respiratory pathogens in a single tube (28). However, comparisons of monoplex and multiplex PCR assays are rare. Both duplex PCR and the monoplex PCR applied by Corsaro et al. detected 2 CFU of M. pneumoniae in DNA extracts from clinical samples (10). Welti et al. (80) developed a multiplex real-time quantitative PCR assay to detect C. pneumoniae, Legionella pneumophila, and M. pneumoniae in respiratory tract specimens. When dilutions of the three pathogen DNAs cloned in plasmids were tested, no significant differences in the sensitivity of each primer set in both the multiplex and monoplex real-time PCR assays were observed. The comparison of multiplex real-time and conventional PCR assays on 73 respiratory specimens showed an overall agreement of 98.3%, corresponding to 95.8, 100, and 100% agreement for C. pneumoniae, L. pneumophila, and M. pneumoniae, respectively. Tong et al. (69) compared the sensitivity and specificity of three PCRs when applied separately and in a triplex format. Sensitivity decreased by about 1 log unit, when the assays were combined. This result was not unexpected, given the complexity of variables in a multiplex PCR, including different combinations of primer concentrations, magnesium ion concentrations, and annealing temperatures. Surprisingly, the sensitivity of the nine-organism multiplex assay of Gro ¨ndahl et al. is said to detect one target sequence in nucleic acid extracts from a diluted M. pneumoniae stock solution (28). Detection of amplification products. Southern blot hybridization or (semi)nested reamplification are often used to increase the sensitivity and to confirm the specificity of the amplicon. However, such methods are time-consuming and cannot be adapted to process large numbers of specimens. Recently, 5⬘ nuclease assays and real-time PCR formats, allowing automated PCR amplification and detection of different pathogens, have been described (31, 54). The advantages of such a real-time system are higher speed, less handling of PCR products, and decreased risk of false-positive results due to carryover contamination. However, the sensitivity of real-time PCR targeting the P1 gene was slightly lower than that of conventional PCR (31). The reasons for this are not clear but may be related to the different assay formats.
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Alternative amplification techniques. Besides PCR, alternative amplification techniques, such as NASBA (55), transcription-mediated amplification, ligase chain reaction, Q replicase amplification (63), and strand displacement amplification, have been developed. These techniques may be useful alternatives to PCR, but so far, few studies on using these techniques to detect M. pneumoniae in respiratory specimens have been published, except for NASBA. The NASBA assay, targeted at 16S rRNA, followed by an enzyme-linked gel assay (ELGA) was used to type M. pneumoniae (55). Later on, NASBA in combination with ELGA and electrochemiluminescence detection was used to detect M. pneumoniae RNA in nucleic acid extracts from respiratory specimens (47). The Q replicase assay was applied to detect synthetic M. pneumoniae 16S rRNA transcripts and seems to be less sensitive than PCR (63). Quantification of M. pneumoniae nucleic acid. DorigoZetsma et al. (14) investigated the relationship between the M. pneumoniae load and disease severity. The M. pneumoniae load in five throat swabs from M. pneumoniae-positive outpatients and from five M. pneumoniae-positive hospitalized patients was assessed semiquantitatively by performing a nested PCR on a series of dilutions of the nucleic acids extracted from these throat swabs. The calculated load varied from 20 to 3,830 CFU/ml. The mean M. pneumoniae load for samples from hospitalized subjects was significantly higher than the load for samples from nonhospitalized subjects. Quantitative real-time DNA and RNA detection is now also possible by using specialized equipment, such as Perkin-Elmer Taqman (56). PRACTICAL DIFFICULTIES AND IMPORTANCE OF QUALITY CONTROL False-positive results. Interlaboratory comparisons illustrate the need for quality control of NAATs. The objective of the study by Ursi et al. (71) was to evaluate the performance of four different assays in detecting M. pneumoniae by PCR in three laboratories through exchange of DNA extracted from respiratory samples. False-positive PCR results were registered in all three participating laboratories, underscoring the importance of including a sufficient number of negative controls in the amplification runs for the detection of sample-tosample carryover, especially when a high proportion of positive samples is expected. In another study by Ieven et al. (36), contamination of entire amplification runs occurred twice, was visualized after hybridization, and was detected by positive results in the negative-control tubes. These contaminations resulted most probably from strong positive samples present in these runs. False-positive results, undetected by the negative controls, occurred in only 0.2% of the samples and were detected after hybridization. Abele-Horn et al. (1) reported that an increase in sensitivity obtained by a nested PCR format was accompanied by an enhanced risk of contamination resulting in false-positive results. In preliminary assays, contamination, detected by positive results after the second amplification step with randomly incorporated negative samples, and probably caused by carryover, occurred in 10% of the samples. By strictly following the guidelines for PCR procedures, such as performing differ-
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TABLE 2. Application of controls used to monitor inhibition of PCR for M. pneumoniae Reference
Specimen(s)a
No. of specimens
Type of controlb
Inhibitionc
1 4 10 14 26 36 59 62 66 67 68 69 72 78
NA, TA TS BAL, BA, PS TS RPA, TA NA TS, NA NA, BAL BAL TS, Sp, NSw, BAL, NA TS, NSw Sp NA, BA, NSw, BAL, Sp, ETA, TS, PF TS
190 99 163 305 165 371 54, 45 124 103 79 462 279 219 41
-Globin gene -Globin gene IFN-␥ gene EHC G3PDH gene EHC -Globin gene -Globin gene EHC -Globin gene -Globin gene Human mt DNA EHC -Globin gene
10%, 2%d, 0% after dilution 0% 0% 20%, 0% after dilution NS 24.6%, 0% after dilution 0% for TS, 25% for NA 25% 7.8%, 0% after lower sample volume 5.1% 11% without extraction, 1.1% after extraction 3.2%, 0% after dilution 15%, 0% after dilution 0%
a Abbreviations: BA, bronchus aspirate; BAL, bronchoalveolar lavage; ETA, endotracheal aspirate; NA, nasopharyngeal aspirate; NSw, nasopharyngeal swab; PF, pleural fluid; PS, pharyngeal swab; Sp, sputum; TS, throat swab; RPA, rhinopharyngeal aspirate; TA, tracheal aspirate. b Abbrevations: IFN-␥, gamma interferon; EHC, extrinsic homologous control; G3PDH, glyceraldehyde phosphate dehydrogenase; mt, mitochondrial. c Abbreviations: NS, not specified; TS, throat swab; NA, nasopharyngeal aspirate. d When PCR is applied on culture-enhanced Hayflick broth (i.e., specimens were received, immediately inoculated into Hayflick broth, and incubated overnight, after which PCR was applied).
ent steps in separate rooms, contamination was reduced to 0.5% in their study. Confirmation of positive findings by repeating the test, including the extraction procedure on the original specimen, is recommended to detect false-positive results due to sporadic contaminations. False-negative results. Negative or nonreproducible NAAT results can be due to a low target copy number, inhibition of amplification, primer mismatches due to strain variations at the primer recognition site, or technical and methodological errors. Amplification inhibitors occur frequently and may be difficult to eliminate. These inhibitors include heme compounds (33) and polysaccharides in sputum (44), as well as in some reagents (27), and mucolytic agents added to sputum (12). Gibb and Wong found that placing throat swabs in Amies clear medium caused inhibition of PCR, the inhibitory component being agar that is dissolved by DNAzol and subsequently precipitated with the DNA by ethanol (21). Others have noted inhibition of PCR by calcium alginate and aluminum swab shafts (77). Therefore, both sampling devices and transport media should be checked for the presence of inhibitors. Different types of internal controls can be used with NAATs to discriminate between a false-negative reaction and a truly nonreactive sample: a homologous extrinsic control, a heterologous extrinsic control, and a heterologous intrinsic control. The former is a wild-type target-derived control, containing a non-target-derived sequence insert. It is added to each sample prior to nucleic acid extraction and coamplified in a single reaction with the same primers as used for amplification of the target sequence. They can be adapted depending on the detection format chosen. The advantages of the homologous extrinsic control are that sample-specific effects, the primers and probes, and the number of copies added can be monitored. The disadvantages are that it does not identify false-negative results resulting from degradation of nucleic acids in clinical specimens prior to the addition of the internal control and it
does not identify the absence of cellular material in a clinical specimen. A heterologous extrinsic control is a non-target-derived control added to each sample prior to nucleic acid extraction. It requires a duplex amplification of target DNA or RNA and control DNA or RNA in a single reaction. Although this kind of control requires primers and probes different from the target, it will still reveal sample-specific effects. This option requires optimization to prevent inhibition of the target amplification by the control amplification reaction. A heterologous intrinsic control confirms the presence of human nucleic acid and thus cellular material in the sample. An example is the detection of a low-abundance mRNA derived from a human cellular housekeeping gene, encoding the A protein present in the human U1 (U1A) small nuclear ribonucleoprotein (snRNP) particle, as a marker of the overall RNA integrity in clinical specimens when RNA is to be analyzed (53). Other examples are the amplification of the -globin or the gamma interferon gene in PCR assays. The disadvantage of this control is the necessity to perform two separate amplification reactions on the same sample or a duplex amplification, with possible inhibition resulting in the latter (37). This kind of control also allows sample-specific effects to be monitored. It cannot be recommended as a test of inhibition, since the number of copies of the human gene in each sample may be much higher than the number of copies of the M. pneumoniae target gene. Interestingly, the proportion of respiratory specimens revealing inhibition by this type of control is usually much lower than that of the extrinsic homologous control: 0 to 10% versus 15 to 36% (Table 2). Dilution of samples or nucleic acid extracts is a simple method to improve amplification by the reduction of inhibitors; however, the sensitivity may also be reduced as a result of dilution of the target molecules, and consequently, samples with low copy numbers may yield false-negative results. The study of Ursi et al. (71), revealed also that none of the three participating laboratories was free of false-negative re-
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sults: one false-negative result was obtained from 78 samples in two laboratories, and four false-negative results were obtained from 78 samples in the third laboratory. The concordance of the assay results in the three laboratories was 84%. Validation of amplification techniques. In-house NAATs were validated by assessing their analytical specificity, sensitivity, and reproducibility on both simulated and clinical specimens in comparison with the best available conventional methods. To define its analytical specificity, the NAAT developed in-house was applied to cultures of organisms that are taxonomically related to the target organism and to organisms commonly present in the clinical specimens to be examined. The analytical sensitivity of the test is defined as the smallest number of organisms that yield a positive result. In practice, this was determined by testing serial dilutions of the target organism, usually under optimal conditions. By repeatedly applying the NAAT to the same series of specimens, the reproducibility of the test could be elucidated. Finally, the defined NAAT has to be compared with the best traditional diagnostic procedures in the clinical microbiology laboratory (this may actually require a combination of methods, including culture, serology, or other microscopic methods) on a panel of positive and negative specimens. Traditionally, NAATs were validated by comparing the NAAT results on a series of clinical specimens with the results of one or two traditional tests. There are, at present, very few prospective studies comparing the performance of two or more amplification protocols, including different specimen preparation methods, on a large number of unselected specimens. Given the multiple amplification protocols proposed, such studies are clearly needed, but the high cost of multiple amplification protocols limits the number of such studies. Ideally, a newly proposed NAAT should be validated by comparison with a sensitive culture system and at least one validated PCR or another NAAT that targets a different gene or a different sequence of the same gene. A comparison of different methods for the diagnosis of respiratory tract infections by M. pneumoniae is presented in Table 3. Agreement between different methods is frequently low, and PCR findings by one or more alternative methods have not always been confirmed. APPLICATION OF NAAT FOR THE DIAGNOSIS OF M. PNEUMONIAE INFECTIONS IN RESPIRATORY INFECTIONS Waring et al. studied a large outbreak of M. pneumoniae in a closed religious community in New York State (78). Throat swab specimens were collected and processed for culture and PCR. A total of 349 specimens were tested by PCR. Of the 349 specimens, 280 specimens were from the original outbreak and 69 were follow-up specimens. Of the 280 initial specimens, 73 were positive and 207 were negative after PCR and hybridization and 22 were positive by culture. The first specimens were tested by both culture and PCR. Since no PCR-negative specimens were culture positive, the remaining specimens were tested by culture only if they were PCR positive. By this approach, PCR was approximately twice as sensitive as culture. In contrast to these findings, Kai et al. detected M. pneumoniae by
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PCR in only 22 of the 30 throat swabs that were positive by culture (39). In this study, however, culture was considered positive when the medium changed color without contamination of the growth of M. pneumoniae. These researchers themselves stated that their culture method may not have been specific for M. pneumoniae. Ieven et al. (36) detected M. pneumoniae in 3.5% of samples from children with acute respiratory tract infections but detected it significantly more often (6.9%) in samples from children above 2 years of age. M. pneumoniae was the third most common etiologic agent of acute respiratory infections in children, after respiratory syncytial virus and influenza virus. In lower respiratory tract infections, such as bronchopneumonia and pneumonia, M. pneumoniae was found as frequently as respiratory syncytial virus (36). Abele-Horn compared PCR, culture, serology, and the direct antigen test for M. pneumoniae (1) in specimens from patients with acute respiratory complaints. A total of 190 patients were divided into three groups: group I (n ⫽ 90) consisted of immunocompromised patients with respiratory complaints after organ or bone marrow transplantation, group II (n ⫽ 50) were adults with acute respiratory tract disease, and group III (n ⫽ 50) included children with lower respiratory tract infections. Among the 190 patients, 20 (11%) were positive by PCR, 11 (8%) were positive by the direct antigen test, 8 (4%) were culture positive, and 17 (9%) were positive by serology. In group I, there were 6, 1, 0, and 3 positive results, respectively. In group II, there were 6, 5, 4, and 6 positive results, respectively. In group III, there were 8, 5, 4, and 8 positive results, respectively. In this study, the best correlation between culture, serology, and PCR results was observed among patients with current infections of the lower respiratory tract (groups II and III). MacFarlane et al. (49) prospectively studied the incidence, etiology, and outcome of lower respiratory tract infections in adult outpatients by serology, culture, and PCR on throat swabs and sputum specimens collected from 316 patients. Twenty-three patients (7.3%) were found to be M. pneumoniae positive. Surprisingly, all were diagnosed by serology. Kessler et al. prospectively collected BAL specimens for PCR and culture from 116 patients admitted to the hospital with a community-acquired pneumonia (40). Serology for M. pneumoniae was done on both acute- and convalescent-phase sera by the CF test. Twelve samples (10.3%) were PCR positive, and 7 (6.0%) of these were subsequently confirmed by culture. The CF test showed seroconversion for these 12 patients, and the results for all other patients remained negative. In contrast, in a comparison of PCR with serological methods, Menendez et al. (51) found their PCR assay to have a lower sensitivity than serology. Only 3 of 184 community-acquired pneumonia patients were found M. pneumoniae positive (51). M. pneumoniae infections seem to be rare in human immunodeficiency virus (HIV)-infected patients (82). Tarp et al., in a retrospective study, applied PCR to BAL fluids obtained from 103 episodes of pneumonia in 83 patients (66). M. pneumoniae was found in two patients (2%). In both cases, M. pneumoniae was present as a coexisting pathogen. The researchers concluded that M. pneumoniae does not seem to play a major role in lower respiratory tract infections in HIV-infected adults and children.
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TABLE 3. Comparison of methods for the diagnosis of respiratory M. pneumoniae infection Reference
Specimen(s)a
No. of patients
Subject age rangeb
1 4 8 10 11 13 14 15 17 20 25 28 26d 30 31 34 36 39 40 41 45 48 49 50 51 57 62 66 67 68 69 74 76 78 79e 83
NA, TA TS TS BAL, BA, Sp BAL, TS TS, BAL, BA, SP, NSw TS TS TNA NA NSw, TS NA RPA, TA NA NA, TS TS, BAL, Sp NA TS BAL TS NSw TS TS, Sp NSw, TS TS, Sp TS NA, BAL BAL Sp, TS, NSw, BAL, NA TS, NSw Sp Sp TS TS NA TS, NSw
190 99 21 163 75 144 305 92 93 132 66 1,118 165 115 48 197 371 105 116 63 3,897 102 316 557 184 155 100 103 79 462 244 34 473 280 278 168
3–66 ⬍1–81 2–18 1–81 NS 20–93 NS NS NS 3m–14y NS NS 2–15 6m–29m ⬍1–16 NS ⬍1–16 NS 1–80 21–62 NS NS NS 0–94 NS 7–92 ⬍1–16 NS ⬍1–85 0–16 5–95 NS 15–65 NS ⬍1–16.5 ⬍1–16
No. (%) of samples positive by detection methodc: PCR
20 (10.5) 49 (49.5)f 11 (52.4) 13 (8.0) 6 (8.0) 15/18 (10.4) 12 (3.9) 7/9 (7.6) 8 (8.6) 3 (2.3) 7 (10.6) 91 (8.1) 22 (13.3) 30 (26) 31 (64.5) 25/34 (12.7) 13 (3.5) 27/31 (25.7) 12 (10.3) 9 (14.3) 283 (7.3) 35 (34.3) 0 (0) 7 (1.3) 3 (1.6) 8 (5.2) 20 (20) 2 (1.9)f 9/14 (11.4) 11 (2.4) 7 (2.9) 9/10 26.5) 4 (0.8)g 73 (26.1) 10 (3.6) 5 (3.0)
Culture
CF
DAG
Serology
8 (4.2) ND ND 9 (5.5) ND ND ND 6/9 (6.5) ND ND ND ND 13 (7.9) ND ND ND 8 (2.2) 31 (29.5) 7 (6.0) ND ND 21 (20.5) ND ND ND ND 1 (1) ND 2/14 (2.5) ND ND 7/10 (20.1) 2 (0.6)h 22 (7.9)i 8 (2.9) 5 (3.0)
ND ND ND ND ND 10/18 (6.9) ND 7/9 (7.6) ND ND ND ND 5 (3.0) ND ND ND ND ND 12 (10.3) ND ND 40 (39.2) ND ND ND ND 5 (5) ND 9/14 (11.4)j ND ND ND ND ND ND ND
11 (5.8) ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND ND
17 (8.9) 32 (32.3) 13 (61.9) ND ND 4/18 (2.8) ND 7/9 (7.6) 18 (19.3) ND ND ND 12 (7.3) ND 23 (47.9) 31/34 (15.7) ND ND ND ND ND ND 23 (7.3) ND 2 (1.1) 9 (5.8) ND ND 9/14 (11.4) ND ND 9/10 (26.5) 2/4 ND 31 (11.2) 10 (6.0)
a Abbreviations: BAL, bronchoalveolar lavage; NA, nasopharyngeal aspirate; NSw, nasal swab; PS, pharyngeal swab; RPA, rhinopharyngeal aspirate; Sp, sputum; TA, tracheal aspirate; TNA, transthoracic needle aspiration; TS, throat swab. b Age ranges given in years (y) unless specified otherwise. m, months; NS, not specified. c Abbreviations: CF, complement fixation test; DAG, direct antigen test; ND, not done. d Twenty PCR-positive specimens were cultured, but complement fixation and IgM serology were not done for all patients. e PCR and culture were not done for all patients. f Results confirmed by another PCR targeting another region of the P1 gene. g Two of four nasopharyngeal aspirates were PCR positive. h Two of 350 throat swab specimens were culture positive. i Culture was done for a selection of 108 of 280 specimens. j Done for 59 of 79 patients.
PCR also detected M. pneumoniae in specimens from 1 to 3% of healthy subjects (48, 67) or in patients after symptomatic infections and even after antibiotic treatment, raising the possibility of a carrier state or persistence of the organism in the respiratory tract (19, 42). Furthermore, Gnarpe et al. found that in endemic and epidemic situations, 4.6 and 13.6%, respectively, of healthy blood donors had throat swabs positive for M. pneumoniae (23). CONCLUSIONS Numerous in-house PCR assays to detect M. pneumoniae have been developed. Proper validation and standardization are often lacking, and quality control studies have revealed frequent deficiencies resulting in both false-negative and falsepositive results.
Consequently, these tests must be submitted to extensive validation before their introduction in the molecular diagnostic laboratory. Validation must be performed at several levels, including sample preparation, amplification, and detection. Since respiratory samples often contain substances inhibiting amplification, special attention should be paid to the efficiency of the reaction with these samples. Once a test is validated, it should be further evaluated in proficiency testing programs. Whereas quality control is an essential part of quality assurance in molecular diagnostics, proficiency panels for the detection of M. pneumoniae are not readily available yet. They are urgently needed to allow meaningful comparisons between the results obtained in different laboratories. Since it has been reported that application of multiplex NAATs may decrease the sensitivity, these assays have to be
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compared carefully with the corresponding individual assays. In many studies, this comparison is lacking. Given the high sensitivity and specificity of NAATs, NAATs are the preferred diagnostic procedures for the diagnosis of M. pneumoniae infections, provided that the quality of the procedures is controlled. Additional studies on large numbers of patients with respiratory signs and symptoms, including hospitalized and nonhospitalized patients, are necessary to extend our knowledge on the epidemiology of M. pneumoniae. As with most applications of NAATs, various amplification targets and the different methods used to detect the targets must be compared to define the most sensitive and specific tests; these studies remain to be undertaken. ACKNOWLEDGMENT The financial support from the European Commission QLK2-CT2000-00294 is gratefully acknowledged. REFERENCES 1. Abele-Horn, M., U. Busch, H. Nitschko, E. Jacobs, R. Bax, F. Pfaff, B. Schaffer, and J. Heesemann. 1998. Molecular approaches to diagnosis of pulmonary diseases due to Mycoplasma pneumoniae. J. Clin. Microbiol. 36: 548–551. 2. Atmar, R. L., and S. B. Greenberg. 1989. Pneumonia caused by Mycoplasma pneumoniae and the TWAR agent. Semin. Respir. Infect. 4:19–31. 3. Bernet, C., M. Garret, B. de Barbeyrac, C. Be´be´ar, and J. Bonnet. 1989. Detection of Mycoplasma pneumoniae by using the polymerase chain reaction. J. Clin. Microbiol. 27:2492–2496. 4. Blackmore, T. K., M. Reznikow, and D. L. Gordon. 1995. Clinical utility of the polymerase chain reaction to diagnose Mycoplasma pneumoniae infection. Pathology 27:177–181. 5. Boom, R., C. J. A. Sol, M. M. M. Salimans, C. L. Jansen, P. M. E. WerthumVan Dillen, and J. Van der Noordaa. 1990. Rapid and simple method for purification of nucleic acids. J. Clin. Microbiol. 28:495–503. 6. Bruisten, S. M., P. Oudshoorn, P. van Swieten, B. Boeser-Nunnink, P. van Aarle, S. P. Tondreau, and H. T. M. Cuypers. 1997. Stability of HIV-1 RNA in blood during specimen handling and storage prior to amplification by NASBA-QT. J. Virol. Methods 67:199–207. 7. Buck, G. E., L. C. O’Hara, and J. T. Summersgill. 1992. Rapid, sensitive detection of Mycoplasma pneumoniae in simulated clinical specimens by DNA amplification. J. Clin. Microbiol. 30:3280–3283. 8. Buck, G. E., and N. S. Eid. 1995. Diagnosis of Mycoplasma pneumoniae in pediatric patients by polymerase chain reaction (PCR). Pediatr. Pulmonol. 20:297–300. 9. Cadieux, N., P. Lebek, and R. Brousseau. 1993. Use of a triplex polymerase chain reaction for the detection and differentiation of Mycoplasma pneumoniae and Mycoplasma genitalium in the presence of human DNA. J. Gen. Microbiol. 139:2431–2437. 10. Corsaro, D., M. Valassina, D. Venditti, V. Venard, A. Le Faou, and P. E. Valensin. 1999. Multiplex PCR for rapid and differential diagnosis of Mycoplasma pneumoniae and Chlamydia pneumoniae in respiratory infections. Diagn. Microbiol. Infect. Dis. 35:105–108. 11. De Barbeyrac, B., C. Bernet-Poggi, F. Fe´brer, H. Renaudin, M. Dupon, and C. Be´be´ar. 1993. Detection of Mycoplasma pneumoniae and Mycoplasma genitalium in clinical samples by polymerase chain reaction. Clin. Infect. Dis. 17(Suppl. 1):S83–S89. 12. Deneer, H. G., and I. Knight. 1994. Inhibition of the polymerase chain reaction by mucolytic agents. Clin. Chem. 40:171–172. 13. Dorigo-Zetsma, J. W., R. P. Verkooyen, H. P. Van Helden, H. Van der Nat, and J. M. Van den Bosch. 2001. Molecular detection of Mycoplasma pneumoniae in adults with community acquired pneumonia requiring hospitalization. J. Clin. Microbiol. 39:1184–1186. 14. Dorigo-Zetsma, J. W., S. A. J. Zaat, A. J. M. Vriesema, and J. Dankert. 1999. Demonstration by a nested PCR for Mycoplasma pneumoniae that M. pneumoniae load in the throat is higher in patients hospitalized for M. pneumoniae infection than in non-hospitalized subjects. J. Med. Microbiol. 48: 1115–1122. 15. Dorigo-Zetsma, J. W., S. A. J. Zaat, P. M. E. Wertheim-van Dillen, L. Spanjaard, J. Rijntjens, G. van Waveren, J. S. Jensen, A. F. Angulo, and J. Dankert. 1999. Comparison of PCR, culture, and serological tests for diagnosis of Mycoplasma pneumoniae respiratory tract infection in children. J. Clin. Microbiol. 37:14–17. 16. Fahle, G. A., and S. H. Fischer. 2000. Comparison of six commercial DNA extraction kits for recovery of cytomegalovirus DNA from spiked human specimens. J. Clin. Microbiol. 38:3860–3863.
17. Falguera, M., A. Nogues, A. Ruiz-Gonzales, M. Garcia, and T. Puig. 1996. Detection of Mycoplasma pneumoniae by polymerase chain reaction in lung aspirates from patients with community acquired pneumonia. Chest 110: 972–976. 18. Fink, C. G., S. J. Read, and M. Sillis. 1995. Direct sample polymerase chain reaction for the detection of Mycoplasma pneumoniae: a simple system for clinical application. Br. J. Biomed. Sci. 52:9–13. 19. Foy, H. M. 1993. Infections caused by Mycoplasma pneumoniae and possible carrier state in different populations of patients. Clin. Infect. Dis. 17(Suppl. 1):S37–S46. 20. Freymuth, F., A. Vabret, J. Brouard, F. Toutain, R. Verdon, J. Petitjean, S. Gouarain, J.-F. Duhamel, and B. Guillois. 1999. Detection of viral, Chlamydia pneumoniae, and Mycoplasma pneumoniae infections in exacerbations of asthma in children. J. Clin. Virol. 13:131–139. 21. Gibb, A. P., and S. Wong. 1998. Inhibition of PCR by agar from bacteriological transport media. J. Clin. Microbiol. 36:275–276. 22. Gil, J. C., R. L. Cedillo, B. G. Mayagoitia, and M. D. Paz. 1993. Isolation of Mycoplasma pneumoniae from asthmatic patients. Ann. Allerg. 70:23–25. 23. Gnarpe, J., A. Lu ¨ndback, and B. G. Sundelo ¨f. 1993. Prevalence of Mycoplasma pneumoniae in subjectively healthy individuals. Scand. J. Infect. Dis. 24:161–164. 24. Gnarpe, J., and K. Eriksson. 1995. Sample preparation for Chlamydia pneumoniae PCR. APMIS 103:307–308. 25. Gnarpe, J., A. Lunba ¨ck, H. Gnarpe, and B. Sundelo ¨f. 1997. Comparison of nasopharyngeal and throat swabs for the detection of Chlamydia pneumoniae and Mycoplasma pneumoniae by polymerase chain reaction. Scand. J. Infect. Dis. Suppl. 104:11–12. 26. Grattard, F., T. Bourlet, C. Galambrun, C. Berger, J. L. Stephan, B. Lauras, and B. Pozzetto. 1998. Inte´reˆt de l’amplification ge´nique par PCR pour le diagnostic des infections `a Mycoplasma pneumoniae chez l’enfant. Pathol. Biol. 46:464–469. 27. Greenfield, L., and J. T. White. 1993. Sample preparation methods, p. 122– 137. In D. H. Persing, T. F. Smith, F. C. Tenover, and T. J. White (ed.), Diagnostic molecular microbiology. Principles and applications. American Society for Microbiology, Washington, D.C. 28. Gro ¨ndahl, B., W. Puppe, A. Hoppe, I. Ku ¨hne, J. A. I. Weigl, and H.-J. Schmitt. 1999. Rapid identification of nine microorganisms causing acute respiratory tract infections by single tube multiplex reverse transcription PCR: feasibility study. J. Clin. Microbiol. 37:1–7. 29. Gurr, P. A., A. Chakraverty, V. Callanan, and S. J. Gurr. 1996. The detection of Mycoplasma pneumoniae in nasal polyps. Clin. Otolaryngol. 21:269–273. 30. Hallander, H., J. Gnarpe, H. Gnarpe, and P. Olin. 1999. Bordetella pertussis, Bordetella parapertussis, Mycoplasma pneumoniae, Chlamydia pneumoniae and persistent cough in children. Scand. J. Infect. Dis. 31:281–286. 31. Hardegger, D., D. Nadal, W. Bossart, M. Altwegg, and F. Dutly. 2000. Rapid detection of Mycoplasma pneumoniae in clinical samples by real-time PCR. J. Microbiol. Methods 41:45–51. 32. Harris, R., B. P. Marmion, G. Varkanis, T. Kok, B. Lunn, and J. Martin. 1988. Laboratory diagnosis of Mycoplasma pneumoniae infection. 2. Comparison of methods for the direct detection of specific antigen or nucleic acid sequences in respiratory exudates. Epidemiol. Infect. 101:685–694. 33. Higuchi, R. 1989. Simple and rapid preparation of samples for PCR, p. 31–38. In H. A. Erlich (ed.), PCR technology. Principles and applications for DNA amplifications. Stockton Press, New York, N.Y. 34. Honda, J., T. Yano, M. Kusaba, J. Yonemitsu, H. Kitajima, M. Masuoka, K. Hamada, and K. Oizumi. 2000. Clinical use of capillary PCR to diagnose Mycoplasma pneumoniae. J. Clin. Microbiol. 38:1382–1384. 35. Hyman, H. C., D. Yogev, and S. Razin. 1987. DNA probes for detection and identification of Mycoplasma pneumoniae and Mycoplasma genitalium. J. Clin. Microbiol. 25:726–728. 36. Ieven, M., D. Ursi, H. Van Bever, W. Quint, H. G. M. Niesters, and H. Goossens. 1996. Detection of Mycoplasma pneumoniae by two polymerase chain reactions and role of M. pneumoniae in acute respiratory tract infections in pediatric patients. J. Infect. Dis. 173:1445–1452. 37. Ieven, M., and H. Goossens. 1997. Relevance of nucleic acid amplification techniques for diagnosis of respiratory tract infections in the clinical laboratory. Clin. Microbiol. Rev. 10:242–256. 38. Jenssen, J. S., J. Sondergard-Andersen, S. A. Uldum, and K. Lind. 1989. Detection of Mycoplasma pneumoniae in simulated clinical samples by polymerase chain reaction. APMIS 97:1046–1048. 39. Kai, M., S. Kamiya, H. Yabe, I. Takakura, K. Shiozawa, and A. Ozawa. 1993. Rapid detection of Mycoplasma pneumoniae in clinical samples by the polymerase chain reaction. J. Med. Microbiol. 38:166–170. 40. Kessler, H. H., D. E. Dodge, K. Pierer, K. K. Y. Young, Y. Liao, B. I. Santner, E. Eber, M. G. Roeger, D. Stuenzner, B. Sixl-Voigt, and E. Marth. 1997. Rapid detection of Mycoplasma pneumoniae by an assay based on PCR and probe hybridization in a nonradioactive microwell plate format. J. Clin. Microbiol. 35:1592–1594. 41. Khoo, S. H., M. Hajia, C. C. Storey, P. E. Klapper, E. G. L. Wilkins, D. W. Denning, E. M. Dunbar, G. Corbitt, and B. K. Mandal. 1998. Influenza-like episodes in HIV-positive patients: the role of viral and atypical infections. AIDS 12:751–757.
VOL. 41, 2003 42. Kleemola, S. R. M., J. E. Karjalainen, and R. K. H. Ra ¨ty. 1990. Rapid diagnosis of Mycoplasma pneumoniae infection: clinical evaluation of a common probe test. J. Infect. Dis. 162:70–75. 43. Kong, F., S. Gordon, and G. L. Gilbert. 2000. Rapid cycle PCR for detection and typing of Mycoplasma pneumoniae in clinical specimens. J. Clin. Microbiol. 38:4256–4259. 44. Lamblin, G., H. Rahmoune, J. M. Wieruszeski, M. Lhermitte, G. Strecker, and P. Rocussel. 1991. Structure of two sulphated oligosaccharides from respiratory mucins of a patient suffering from cystic fibrosis: a fast-atombombardment m.s. and 1H-n.m.r. spectroscopic study. Biochem. J. 275:199– 206. 45. Layani-Milon, M.-P., I. Gras, M. Valette, J. Luciani, J. Stagnara, M. Aymard, and B. Lina. 1999. Incidence of upper respiratory tract Mycoplasma pneumoniae infections among outpatients in Rho ˆne-Alpes, France, during five successive winter periods. J. Clin. Microbiol. 36:1721–1726. 46. Leng, Z., G. E. Kenny, and M. C. Roberts. 1994. Evaluation of the detection limits of PCR for identification of Mycoplasma pneumoniae in clinical samples. Mol. Cell. Probes 8:125–130. 47. Loens, K., D. Ursi, M. Ieven, P. van Aarle, P. Sillekens, P. Oudshoorn, and H. Goossens. 2002. Detection of Mycoplasma pneumoniae in spiked clinical samples by nucleic acid sequence-based amplification. J. Clin. Microbiol. 40:1339–1345. 48. Lu ¨neberg, E., J. S. Jensen, and M. Frosch. 1993. Detection of Mycoplasma pneumoniae by polymerase chain reaction and nonradioactive hybridization in microtiter plates. J. Clin. Microbiol. 31:1088–1094. 49. MacFarlane, J., W. Holmes, P. Gard, R. MacFarlane, D. Rose, V. Weston, M. Leinonen, P. Saikku, and S. Myint. 2001. Prospective study of the incidence, aetiology and outcome of adult lower respiratory tract illness in the community. Thorax 56:109–114. 50. Meijer, A., C. F. Dagnelie, J. C. De Jong, A. De Vries, T. M. Bestebroer, A. M. Van Loon, A. I. Bartelds, and J. M. Ossewaarde. 2000. Low prevalence of Chlamydia pneumoniae and Mycoplasma pneumoniae among patients with symptoms of respiratory tract infections in Dutch general practices. Eur. J. Epidemiol. 16:1099–1106. 51. Menendez, R., J. Cordoba, P. de la Cuadra, M. J. Cremades, J. L. LopezHontagas, M. Salavert, and M. Gobernado. 1999. Value of the polymerase chain reaction assay in noninvasive respiratory samples for diagnosis of community acquired pneumonia. Am. J. Respir. Crit. Care Med. 159:1868– 1873. 52. Narita, M., Y. Matsuzono, O. Itakura, S. Yamada, and T. Togashi. 1998. Analysis of mycoplasmal pleural effusion by the polymerase chain reaction. Arch. Dis. Child. 78:67–69. 53. Nelissen, R., P. Sillekens, R. Beijer, H. Van Kessel, and W. van Venrooij. 1991. Structure, chromosomal localization and evolutionary conservation of the gene encoding human U1 snRNP-specific A protein. Gene 102:189–196. 54. Oberst, R. D., M. P. Hays, L. K. Bohra, R. K. Phebus, C. T. Yamashiro, C. Paszko-Kolva, J. M. Sargeant, and J. R. Gillespie. 1998. PCR-based DNA amplification and presumptive detection of Escherichia coli O157:H7 with an internal fluorogenic probe and the 5⬘ nuclease (TaqMan) assay. Appl. Environ. Microbiol. 64:3389–3396. 55. Ovyn, C., D. Van Strijp, M. Ieven, D. Ursi, B. Van Gemen, and H. Goossens. 1996. Typing of Mycoplasma pneumoniae by nucleic acid sequence based amplification, NASBA. Mol. Cell. Probes 10:319–324. 56. Pahl, A., U. Kuhlbrandt, K. Brune, M. Rollinghoff, and A. Gessner. 1999. Quantitative detection of Borrelia burgdorferi by real-time PCR. J. Clin. Microbiol. 37:1958–1963. 57. Ramirez, J. A., S. Ahkee, A. Tolentino, R. D. Miller, and J. T. Summersgill. 1996. Diagnosis of Legionella pneumophila, Mycoplasma pneumoniae, or Chlamydia pneumoniae lower respiratory infection using a single throat swab specimen. Diagn. Microbiol. Infect. Dis. 24:7–14. 58. Razin, S. 1994. DNA probes and PCR in diagnosis of Mycoplasma infections. Mol. Cell. Probes 8:497–511. 59. Reznikov, M., T. K. Blackmore, J. J. Finlay-Jones, and D. L. Gordon. 1995. Comparison of nasopharyngeal aspirates and throat swab specimens in a polymerase chain reaction-based test for Mycoplasma pneumoniae. Eur. J. Clin. Microbiol. Infect. Dis. 14:58–61. 60. Sasaki, Y., M. Shintani, T. Shimada, H. Watanabe, and T. Sasaki. 1992. Detection and discrimination of Mycoplasma pneumoniae and Mycoplasma genitalium by the in vitro DNA amplification. Microbiol. Immunol. 36:21–27. 61. Sillis, M. 1993. Modern methods for diagnosis of Mycoplasma pneumoniae pneumonia. Rev. Med. Microbiol. 4:24–31. 62. Skakni, L., A. Sardet, J. Just, J. Landman-Parker, J. Costil, N. Moniot-Ville, F. Bricout, and A. Garberg-Chenon. 1992. Detection of Mycoplasma pneumoniae in clinical samples from pediatric patients by polymerase chain reaction. J. Clin. Microbiol. 30:2638–2643. 63. Stone, B. B., S. P. Cohen, G. L. Breton, R. M. Nietupski, D. A. Pelletier, M. J. Fiandaca, J. G. Moe, J. H. Smith, J. S. Shah, and W. G. Weisburg. 1996. Detection of rRNA from four respiratory pathogens using an automated Q replicase assay. Mol. Cell. Probes 10:359–370. 64. Su, C. J., A. Chavoya, and J. B. Baseman. 1988. Regions of Mycoplasma
MINIREVIEW
65. 66. 67.
68.
69.
70. 71.
72. 73.
74.
75. 76.
77.
78.
79. 80.
81.
82. 83.
84.
4923
pneumoniae cytadhesin P1 structural gene exist as multiple copies. Infect. Immun. 56:3157–3161. Talkington, D. F., W. Z. Thacker, D. W. Keller, and J. S. Jensen. 1998. Diagnosis of Mycoplasma pneumoniae infections in autopsy and open-lung biopsy tissues by nested PCR. J. Clin. Microbiol. 36:1151–1153. Tarp, B., J. S. Jensen, L. Ostergaard, and P. L. Andersen. 1999. Search for agents causing atypical pneumonia in HIV-positive patients by inhibitor controlled PCR assays. Eur. Respir. J. 13:175–179. Tjhie, J. H. T., F. J. M. Van Kuppeveld, R. Roosendaal, W. J. G. Melchers, R. Gordijn, D. M. MacLaren, J. M. M. Walboomers, C. J. L. M. Meijer, and A. J. C. van den Brule. 1994. Direct PCR enables detection of Mycoplasma pneumoniae in patients with respiratory tract infections. J. Clin. Microbiol. 32:11–16. Tjhie, J. H. T., J. W. Dorigo-Zetsma, R. Roosendaal, A. J. C. Van den Brule, T. M. Bestebroer, A. I. M. Bartelds, and C. M. J. E. Vandenbroucke-Grauls. 2000. Chlamydia pneumoniae and Mycoplasma pneumoniae in children with acute respiratory infection in general practice in The Netherlands. Scand. J. Infect. Dis. 32:13–17. Tong, C. Y. W., C. Donnelly, G. Harvey, and M. Sillis. 1999. Multiplex polymerase chain reaction for the simultaneous detection of Mycoplasma pneumoniae, Chlamydia pneumoniae and Chlamydia psittaci in respiratory samples. J. Clin. Pathol. 52:257–263. Uldum, S. A., J. S. Jensen, J. Sondergard-Andersen, and K. Lind. 1992. Enzyme immunoassay for detection of immunoglobulin M (IgM) and IgG antibodies to Mycoplasma pneumoniae. J. Clin. Microbiol. 30:1198–1204. Ursi, D., M. Ieven, G. T. Noordhoek, M. Ritzler, H. Zandleven, and M. Altwegg. 2003. An interlaboratory comparison for the detection of Mycoplasma pneumoniae in respiratory samples by the polymerase chain reaction. J. Microbiol. Methods 53:289–294. Ursi, J. P., D. Ursi, M. Ieven, and S. R. Pattyn. 1992. Utility of an internal control for the polymerase chain reaction. APMIS 100:635–639. Van Kuppeveld, F. J. M., J. T. M. van der Logt, A. F. Angulo, M. J. van Zoest, W. G. V. Quint, H. G. M. Niesters, J. M. D. Galama, and W. J. G. Melchers. 1992. Genus- and species-specific identification of Mycoplasmas by 16S rRNA amplification. Appl. Environ. Microbiol. 58:2606–2615. Van Kuppeveld, F. J. M., K.-E. Johansson, J. M. Galama, J. Kissing, G. Bo ¨lske, E. Hjelm, J. T. van der Logt, and W. J. Melchers. 1994. 16S rRNA based polymerase chain reaction compared with culture and serological methods for diagnosis of Mycoplasma pneumoniae infection. Eur. J. Clin. Microbiol. Infect. Dis. 13:401–405. Vekris, A., F. Bauduer, S. Maillet, C. Be´be´ar, and J. Bonnet. 1995. Improved microplate immunoenzymatic assay of PCR products for rapid detection of Mycoplasma pneumoniae. Mol. Cell. Probes 9:25–32. Wadowsky, R. M., E. A. Castilla, S. Laus, A. Kozy, R. W. Atchinson, L. A. Kingsley, J. I. Ward, D. P. Greenberg, and the Adult Pertussis Vaccine Efficacy Study Group. 2002. Evaluation of Chlamydia pneumoniae and Mycoplasma pneumoniae as etiologic agents of persistent cough in adolescents and adults. J. Clin. Microbiol. 40:637–640. Wadowsky, R. M., S. Laus, T. Libert, S. J. States, and G. D. Ehrlich. 1994. Inhibition of a PCR-based assay for Bordetella pertussis by using calcium alginate fiber and aluminum shaft components of a nasopharyngeal swab. J. Clin. Microbiol. 32:1054–1057. Waring, A. L., T. A. Halse, C. K. Csiza, C. J. Carlyn, K. Arruda Musser, and R. J. Limberger. 2001. Development of a genomics-based PCR assay for detection of Mycoplasma pneumoniae in a large outbreak in New York State. J. Clin. Microbiol. 39:1385–1390. Waris, M. E., P. Toikka, T. Saarinen, S. Nikkari, O. Meurman, R. Vainionpa ¨a ¨, J. Mertsola, and O. Ruuskanen. 1998. Diagnosis of Mycoplasma pneumoniae pneumonia in children. J. Clin. Microbiol. 36:3155–3159. Welti, M., K. Jaton, M. Altwegg, R. Sahli, A. Wenger, and J. Bille. 2003. Development of a multiplex real-time quantitative PCR assay to detect Chlamydia pneumoniae, Legionella pneumophila and Mycoplasma pneumoniae in respiratory tract specimens. Diagn. Microbiol. Infect. Dis. 45:85– 95. Williamson, J., B. P. Marmion, D. A. Worswick, T. W. Kok, G. Tannock, R. Herd, and R. J. Harris. 1992. Laboratory diagnosis of Mycoplasma pneumoniae infection. 4. Antigen capture and PCR gene amplification for detection of the Mycoplasma: problems of clinical correlation. Epidemiol. Infect. 109:519–537. Witt, D. J., D. E. Craven, and W. R. McCabe. 1987. Bacterial infections in adult patients with the acquired immunodeficiency syndrome (AIDS) and AIDS-related complex. Am. J. Med. 82:900–906. Wubbel, L., L. Muniz, A. Ahmed, M. Trujillo, C. Carubelli, C. McCoig, T. Abramo, M. Leinonen, and G. H. McCracken, Jr. 1999. Etiology and treatment of community acquired pneumonia in ambulatory children. Pediatr. Infect. Dis. J. 18:98–104. Zingangirova, N. A., O. V. Popova, C. V. Solovjeva, A. L. Gintzburg, and S. V. Prozorovsky. 1993. Development of a PCR-based method for diagnosing Mycoplasma pneumoniae infection. Lett. Appl. Microbiol. 16:106–109.
