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Standardizing Chlamydia pneumoniae Assays: Recommendations from the Centers for Disease Control and Prevention (USA) and the Laboratory Centre for Disease Control (Canada)

  1. Scott F. Dowell1,
  2. Rosanna W. Peeling5,
  3. Jens Boman6,
  4. George M. Carlone1,
  5. Barry S. Fields1,
  6. Jeannette Guarner1,
  7. Margaret R. Hammerschlag4,
  8. Lisa A. Jackson2,
  9. Cho-Chou Kuo3,
  10. Matthias Maass7,
  11. Trudy O. Messmer1,
  12. Deborah F. Talkington1,
  13. Maria Lucia Tondella1,
  14. Sherif R. Zaki1, and
  15. C. pneumoniae Workshop Participantsa
  1. 1National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta
  2. 2Department of Epidemiology, School of Public Health and Community Medicine, University of Washington, Seattle
  3. 3Department of Pathobiology, School of Public Health and Community Medicine, University of Washington, Seattle
  4. 4SUNY Health Science Center at Brooklyn, New York
  5. 5Laboratory Centre for Disease Control, Health Canada, Ottawa, Ontario, Canada
  6. 6Department of Clinical Virology, University Hospital of Umea, Umea, Sweden
  7. 7Institute of Medical Microbiology and Hygiene, Medical University of Lübeck, Lübeck, Germany
  1. Reprints or correspondence: Dr. Scott F. Dowell, Mailstop C-12, Centers for Disease Control and Prevention, 1600 Clifton Rd. NE, Atlanta, GA 30333 (sdowell{at}cdc.gov).
 
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Abstract

Chlamydia pneumoniae has been associated with atherosclerosis and several other chronic diseases, but reports from different laboratories are highly variable and “gold standards” are lacking, which has led to calls for more standardized approaches to diagnostic testing. Using leading researchers in the field, we reviewed the available approaches to serological testing, culture, DNA amplification, and tissue diagnostics to make specific recommendations. With regard to serological testing, only use of microimmunofluorescence is recommended, standardized definitions for “acute infection” and “past exposure” are proposed, and the use of single immunoglobulin (Ig) G titers for determining acute infection and IgA for determining chronic infection are discouraged. Confirmation of a positive culture result requires propagation of the isolate or confirmation by use of polymerase chain reaction (PCR). Four of 18 PCR assays described in published reports met the proposed validation criteria. More consistent use of control antibodies and tissues and improvement in skill at identifying staining artifacts are necessary to avoid false-positive results of immunohistochemical staining. These standards should be applied in future investigations and periodically modified as indicated.

Chlamydia pneumoniae is a fastidious bacterium that was first established as a cause of acute respiratory infection >15 years ago; more recently, it has been associated with certain chronic diseases, including atherosclerotic cardiovascular disease. The ever-expanding spectrum of diseases associated with C. pneumoniae infection has led a sizable influx of new investigators and laboratories to become involved in Chlamydia-related research. Test results are often contradictory and difficult to interpret; dramatic findings from one laboratory are unconfirmed by others. As a result, researchers in the field and external reviewers have called for validated and standardized diagnostic techniques to promote research applications and improve the recognition and care of patients infected with C. pneumoniae [1, 2].

A meeting was recently convened by the Centers for Disease Control and Prevention (Atlanta) and the Laboratory Centre for Disease Control (Ottawa, Ontario, Canada) to review current diagnostic tests for C. pneumoniae and provide recommendations for standardized approaches.

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Serological Testing

There are no wholly satisfactory serological methods for diagnosis of C. pneumoniae infection. Problems arise from the difficulty in obtaining appropriately paired serum samples, the high background of IgG antibody prevalence in certain adult populations, the lack of standardized testing methods, and a shortage of high-quality reagents.

For patients with acute C. pneumoniae infection, it is important to take into consideration the kinetics of the antibody response. In patients with primary infection, IgM antibody appears ∼2–3 weeks after the onset of illness and is generally undetectable after 2–6 months. IgG antibody may not reach high titer until 6–8 weeks after the onset of illness. C. pneumoniae infection does not induce good protective immunity, and reinfection may occur. In cases of reinfection, IgM antibody may not appear and the level of IgG antibody titer increases quickly, within 1–2 weeks. Serological testing most often provides only a retrospective diagnosis of acute infection, because a convalescent serum specimen is needed to show a 4-fold increase in titer, so it is not optimum for patient management. Nevertheless, serological testing is the most useful means of determining the cause of an outbreak or the prevalence of infection in epidemiologic studies.

Review of Currently Used Tests: Serological Testing

Complement fixation (CF), whole-inclusion fluorescence, and EIA cannot currently be endorsed. CF has objective end points and can detect increases in antibody levels in specimens obtained at intervals as close as 1 week apart, but the test cross-reacts with other Chlamydia species and other enteric bacteria, the sensitivity for detecting reinfection is low, and reagents are not readily available [3-5]. Whole-inclusion fluorescence tests are available as commercial kits, but they also are not species specific and have not been widely evaluated [4]. The EIA holds the most promise, and several kits are commercially available, although none has been approved by the US Food and Drug Administration (FDA) for use in the United States. Advantages of the EIA include high throughput, objective end points, technical accessibility, and an electronic record of the results. However, the limited published evaluations of these kits that have appeared to date have included reports of problems with both sensitivity and specificity [6]. Therefore, no currently available assay can be recommended because of a lack of peer-reviewed evaluations to document that the specificity is adequate when compared with that of microimmunofluorescence (MIF). In addition, use of EIA as a screening method is not endorsed because the high sensitivity required by this approach has not been demonstrated.

The MIF test is the serological testing method of choice for diagnosis of acute C. pneumoniae infection. It was the use of the MIF that led to the identification of C. pneumoniae as a distinct species of Chlamydia [7]. It is the only species-specific antibody test available that can measure isotype-specific antibody titers to all Chlamydia species simultaneously. The specificity of the MIF test can be attributed to the use of purified elementary bodies of all 3 species of Chlamydia rather than reticulate bodies that express predominately genus-specific epitopes.

The test format uses purified formalinized elementary bodies from C. pneumoniae, Chlamydia trachomatis, and Chlamydia psittaci that have been fixed onto glass slides as distinct dots of antigen. Dilutions of sera are placed over the antigen dots and incubated. However, the assay is technically complex, interpretation is subjective, and neither reagents nor diagnostic criteria have been standardized [8]. Kits based on the MIF format are commercially available. Their performance characteristics require further evaluation and peer-reviewed publication before endorsement.

Specific Recommendations for Serological Testing

A standardized approach to performing and interpreting the MIF assay is presented in table 1. Quality assurance procedures are particularly important to emphasize because of the subjective nature of the interpretation; they are included in table 1.

Figure 1
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Figure 1

Positive results of staining of smooth muscle cells for Chlamydia pneumoniae in an atheroma (arrow) by use of the TT-401 antibody, an avidin-biotin complex (ABC) method, and horseradish peroxidase. Reprinted with permission from [63].

Figure 2
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Figure 2

A, False-positive Chlamydia staining of smooth muscle cell in an atheroma (arrow) by use of the avidin-biotin complex (ABC) method, horseradish peroxidase, and a monoclonal antibody specific for Chlamydia (CF-2). B, The same tissue block showing similar staining with a nonspecific monoclonal antibody of the same isotype (IgG2a antibody to Lassa fever virus). C, False-positive Chlamydia staining of inflammatory cells (macrophages) by use of ABC, horseradish peroxidase, and a monoclonal antibody specific for Chlamydia (CF-2). D, The same tissue block showing similar staining with a nonspecific monoclonal antibody of the same isotype (IgG2a antibody to Lassa fever virus).

Table 1
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Table 1

Recommendations for use of the microimmunofluorescence test.

Several caveats regarding MIF serological testing are important to emphasize. Diagnosis of acute infection based on a single IgG titer cannot be routinely recommended; if single IgG titers are reported, they should be interpreted with caution. Serum samples obtained from elderly patients and from patients with chronic obstructive pulmonary disease have had persistently high IgG titers in the absence of clinically apparent disease. Failure to adsorb serum before IgM testing may lead to false-positive IgM results due to the presence of rheumatoid factor in the sera. The quality of the IgA conjugates has been found to vary, and their use requires careful evaluation and further standardization. Finally, the absence of MIF antibodies in persons with culture-confirmed infection has been reported. This is rare in adults but may be more common in young children [9-11].

One of the most challenging aspects of C. pneumoniae testing is the identification of persons with persistent or chronic infection by means of serological testing. Persistently elevated IgG or the presence of IgA antibodies have been frequently used [12-14]. There is no reference test for validating persistent infection, and studies that have attempted to correlate single-sample IgG or IgA antibody levels with disease status have produced equivocal results, owing in part to the different methodologies and titer cutoff points that were used in different studies [15, 16]. It has been proposed that high IgA titers may be a better marker of chronic C. pneumoniae infection than are IgG titers because serum IgA has a half-life of 5–7 days, whereas IgG has a half-life of weeks to months. However, there is at present no validated serologic marker of persistent or chronic infection, and the use of serological testing to define patients as “persistently infected” must await further validation.

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Culture

C. pneumoniae is an obligate intracellular bacterium and must be cultivated within a eukaryotic host cell. The specificity of culture is dependent on the ability of the laboratory worker to distinguish true Chlamydia inclusions from artifacts on microscopic examination of the cell monolayer after fluorescent antibody staining has been performed. Tests that are not based on culture, such as PCR, have become widely used for detection of Chlamydia infection, in part because of the technical complexity and low yield of culture protocols. Problems with low yields may also be related to the contamination of culture with Mycoplasma species [17, 18]. However, culture remains essential to document the viability of the organism, to provide isolates of the organism for biological characterization and antimicrobial susceptibility testing, and to assess microbiologic efficacy in treatment trials.

Review of Currently Published Protocols for Culture

All currently accepted culture procedures involve inoculation of a specimen onto a human cell line via centrifugation. The inoculated cells are incubated and are later stained with a fluorescent-labeled antibody specific to Chlamydia to visualize the bacteria that are multiplying within the host cells.

There are a number of modifications to these procedures that remain controversial. These include simultaneous centrifuging of the cell line and inoculum onto the culture vessel [19], the use of serum-free cell culture medium [20], prolongation of culture times, and increases in the number of times the cell cultures are centrifuged after inoculation [21, 22]. Host-cell monolayers have been pretreated with polyethylene glycol, trypsin, and diethylaminoethyl dextran to improve the recovery of isolates of C. pneumoniae from either true or mock specimens [22, 23]. Other researchers, however, have reported that diethylaminoethyl dextran pretreatment actually decreased the size of inclusions [24] or failed to document improved recovery by use of either technique [21]. None of these modifications has been sufficiently tested to warrant their routine recommendation.

There is a good deal of controversy regarding the number of times a culture should be passaged before the results are determined [25-27]. Most laboratories agree that ⩾2 passages after the primary culture step is performed are needed to maximize the recovery of C. pneumoniae isolates from respiratory specimens. Other reports have successfully used greater numbers of passages, particularly for isolation from tissue specimens, although no systematic comparison of passage numbers has been attempted. Increased passages may result in a concentration of cell debris that may contribute to nonspecific staining of the monolayer.

We recommend that respiratory specimens should be cultured by means of primary isolation procedures plus 2 additional passages. Tissue specimens should be cultured by means of primary isolation procedures plus 4–6 additional passages.

Specific Recommendations for Culture

Specimen types. Specimens obtained for detection of C. pneumoniae respiratory tract infection by use of culture include swabs of the nasopharynx or oropharynx, sputum specimens, bronchoalveolar lavage specimens, and tissue biopsy specimens. C. pneumoniae has not been successfully cultured from blood samples, although the DNA can be detected in samples of peripheral blood mononuclear cells, and the organism has been recovered from a limited number of vascular tissue specimens.

Specimen collection. Swab specimens should be collected only on swabs with a Dacron tip and an aluminum or plastic shaft. Swabs with calcium alginate or cotton tips and wooden shafts may inhibit the growth of the organism, depending on the adhesive used, and are unacceptable. Swabs should be placed in 2SP transport medium (sucrose, 68.4 g; potassium phosphate dibasic, 2.01 g; potassium phosphate monobasic, 1.01 g; gentamicin, 10 µg/mL; amphotericin B, 100 µg/mL; vancomycin, 25 µg/mL; and 10%–20% fetal calf serum, made up to a final volume of 1000 mL, pH 7.2) and not removed before transport. Bronchoalveolar lavage, sputum, and pleural fluid samples should be collected in 2SP at a ratio of specimen to medium of 1:2.

Specimen transport. All specimens that can be processed in the laboratory within 24 h should be held and shipped at 4°C (on wet ice). Samples that cannot be processed within 24 h should be frozen and held at −70°C.

Specimen processing. Swabs should be vortexed in the transport medium for 15–20 s and pressed against the side of the tube to extract all of the liquid. One hundred to 200 µL of this fluid is used as the inoculum. The specimen is then centrifuged at 8000–10,000g, resuspended in cell culture medium, and homogenized. It should be noted that sputum frequently inhibits cell growth and may be toxic to the monolayer. Calcified areas of vascular tissue specimens should be removed and the tissue resuspended in cell culture medium before homogenization.

Assay procedures and quality control. A standardized approach to culture procedures and quality assurance testing is presented in table 2. The use of methanol as a fixative for the monolayers before staining should be avoided or evaluated in advance, because there is some evidence that the antigenicity of certain proteins may be destroyed [28].

Table 2
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Table 2

Recommendations for use of culture for Chlamydia pneumoniae.

Whether a few inclusions that fail to propagate in subsequent passages should be considered a “true-positive” result of culture has been an area of debate. For all results of culture for C. pneumoniae, we recommend that the detection of an average of ⩾1 inclusions per well or tube be considered a “presumptive positive.” Only if the strain can be propagated by means of subsequent passage or confirmed by use of an additional test, such as PCR, should it be reported as a “confirmed positive.” It should also be recognized that C. pneumoniae has been cultured from specimens of the upper respiratory tract obtained from asymptomatic persons.

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PCR

The description of the clinical spectrum of C. pneumoniae disease has been hampered by the lack of sufficiently sensitive diagnostic methods. Nucleic acid-based amplification techniques, such as PCR, have identified C. pneumoniae in clinical samples ranging from respiratory specimens [29-31] to samples of vascular tissue [32-35], serum [36], and peripheral blood mononuclear cells [37]. Despite significant improvements in the development of molecular methods for the detection of C. pneumoniae, some laboratories report consistent detection of the organism in specimens of vascular tissue [32, 33, 38, 39], whereas others do not [40-42]. This variation may be related to differences in means of specimen collection and processing, primer design, nucleic acid extraction, amplification product detection, or prevention and identification of false-positive and false-negative results.

Review of Current Tests: PCR

Table 3 summarizes 18 reports regarding PCR assays, published as of 1 May 2000, with the target regions, product sizes, and methods for detection of the products. Although many in-house PCR methods are available for detection of C. pneumoniae, the sensitivity and specificity of the majority of these tests remain unknown. More studies need to be conducted using proper controls and a large number of clinical specimens obtained from patients to compare and evaluate more adequately the usefulness of different PCR tests for the diagnosis of C. pneumoniae infection. In addition, comparison of PCR results with those of a sensitive culture system and at least 1 other validated PCR assay that targets a different gene or a different sequence of the same gene is necessary to validate any newly proposed PCR assay.

Table 3
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Table 3

PCR assays for detection of Chlamydia pneumoniae in clinical specimens.

Among the assays described in table 3, only 4 widely used protocols, which are highlighted at the top of the table, satisfy the optimal criteria for a validated assay. First, they have been validated for sensitivity and specificity in ⩾2 outside laboratories using both calibrated artificial specimens and true clinical specimens. Second, sensitivity has been documented to a level of detection of <1 inclusion-forming unit (IFU). Finally, for each of these assays, specificity has been documented against other Chlamydia species as well as a wide range of other prokaryotic and eukaryotic DNA. We emphasize that all of these assays are research tools and none have been commercially standardized or cleared by the FDA.

Each PCR type has advantages and disadvantages that should be carefully considered before new assays are designed or evaluated. Seven PCR approaches and the advantages and disadvantages of each approach are briefly reviewed.

Nested PCR is, in general, more sensitive than is single-step PCR because of the 2-step amplification and the use of 2 sets of primers. The disadvantages of nested PCR are the increased risk of contamination and reamplification of the products, which makes the assay more time-consuming and expensive.

A multiplex PCR amplifies >1 target sequence in the same assay. For the Chlamydia genus, this has been used to discriminate among species. Multiplex reactions decrease sensitivity and specificity if the annealing temperatures for the individual primers are not identical. In general, these assays are not as sensitive as single-target PCR.

Internal or amplification controls allow the monitoring of PCR assay inhibition, which may be caused by a number of factors. These controls can have the disadvantage of competing for primers when identical primers are used for both target genes and internal controls. If a different set of primers is used to detect the internal control, differences in amplification conditions may decrease sensitivity.

Hot-start PCRs increase specificity by preventing nonspecific primer binding at temperatures that are less than the optimum temperature. Such assays are not recommended unless special hot-start Taq polymerases are used, because for all others, the need to open the tube to add more polymerase increases the potential for contamination.

Touchdown PCRs increase specificity by allowing the initial primer-template hybridization events to occur at annealing temperatures that are greater than the optimum annealing temperature. However, this type of PCR requires more cycles, which increases the duration of the PCR run.

Hybridization probe methods should routinely be used to ensure the specificity of the PCR product. They may also increase sensitivity, compared with standard visualization of PCR products in agarose gel after electrophoresis and ethidium bromide staining. These methods have the disadvantage of increased cost of reagents and are, in general, more time-consuming than are nonprobe methods.

The fluorescent probe-based assays that are currently being developed have the advantages of a closed system that avoids contamination by PCR product carryover. They can have real-time or end point readings, depending on the system that is used. These assays may be more sensitive and are inherently more specific than is single-step PCR because of dual primer and probe binding. However, the equipment that is required is very expensive.

Specific Recommendations for PCR

Specimen collection and processing. Specific recommendations regarding collection, transport, and processing of clinical specimens are similar to those described in the culture section. One aliquot (1 mL) of the transport medium should be centrifuged at ∼18,000g for 15 min and the pellet should be processed for DNA extraction. Commercially available cell-preparation tubes facilitate separation of mononuclear cells from whole blood. Tissue samples should be cut into small pieces (∼25 mg) and processed for DNA extraction. Specimens should be formed into aliquots to avoid >1 freeze-thaw cycle for optimal yield.

Specimens, controls, and PCR mixture reagents should be handled with dedicated pipettes in physically separated areas to avoid contamination. Aerosol barrier pipette tips, dedicated laboratory coats, and gloves are strongly recommended. Benchtop and equipment should be monitored routinely for DNA contamination by conducting swipe tests.

Assay procedures and quality control: DNA extraction. C. pneumoniae DNA should be extracted from clinical samples by use of a highly efficient and reliable protocol. Any new extraction method should be validated before routine use and should be assessed for problems with inhibitors of DNA polymerases.

Assay procedures and quality control: positive and negative controls. These should be included in all runs, in parallel with the clinical samples throughout the extraction and detection procedures. The positive controls should consist of small and very small amounts of DNA (titrations down to <10 ng of DNA) from a culture with <10 IFU. Each laboratory should prepare titrations of their C. pneumoniae stocks and form a large number of low and very low positive controls into aliquots. At least 1 negative control, consisting of water in place of the clinical specimen, should be run every fifth DNA extraction.

Assay procedures and quality control: amplification controls. The use of spiked nonrelated DNA adds validity to results by identifying potential inhibitors in the samples or in the PCR reaction itself. Such controls should be included in newly developed assays. Several different internal controls have been used, such as λ phage DNA [56], MIMICS or competitive primers [59, 60], and cloned fragments into the pUC19 vector [48]. Because of the potential for competition of the control with the target sequence, a low copy number of control DNA is important.

Development and quality control of new PCR assays. There is a critical need for commercially standardized, FDA-cleared assays. In the meantime, researchers should design primers and probes that are based on a highly conserved gene sequence by using sequence databases. The targeted DNA sequences should be searched by use of Basic Local Alignment Sequence Tool (available at http://www.ncbi.nlm.nih.gov/BLAST/) to check for specificity. Any newly developed primers or probes should be validated for analytic sensitivity and specificity.

Sensitivity should be determined by performing titrations of designated isolates to determine the lowest level of detection of the target gene. At a minimum, both C. pneumoniae type strain TW-183 (ATCC VR-2282) and CM-1 (ATCC VR-1360) should be titrated with each newly designed assay and the number of IFUs quantified as described in the culture section above. Ideally, all available C. pneumoniae strains should be tested for contamination with Mycoplasma species by use of a genus-specific PCR and with stock cultures that are similarly titrated. The specificity of new primers and probes should be tested with a bank of DNA preparations from C. psittaci, C. trachomatis, and other bacteria and viruses commonly found in the respiratory tract, with human DNA, and with at least 1 of the 4 recommended PCRs in table 3 by the laboratory developing the assay and by an independent laboratory.

Interpretation and reporting of results. For increased specificity, positive samples are often re-extracted and reanalyzed, but such selective repeat testing of only positive specimens introduces deliberate bias toward decreased sensitivity while increasing specificity. Whenever feasible, those persons who conduct the assays and interpret the results for research studies should be blinded to the patient status (case patient or control) and results of other testing (antibody status, culture or tissue diagnostic results). The resulting publications should specify how blinding was ensured or why it was not done.

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Tissue Diagnostics (Immunohistochemistry)

C. pneumoniae has been detected in tissue specimens by use of a variety of methods [15, 61]. Of those methods, immunofluorescence, immunohistochemistry (IHC), and in situ hybridization offer the advantage of preserving tissue morphology and permitting localization of the infectious agent to specific areas and cells. Cell types reported to be susceptible to infection include macrophages, endothelium, and smooth muscle [61]. Of the tissue diagnostic methods, IHC has been the most frequently used in studies of C. pneumoniae, and we will focus on this method. For IHC, careful interpretation is the critical challenge, because true-positive results of staining (figure 1) and false-positive results of staining (figure 2) can be very difficult to distinguish.

As of 1 May 2000, >20 publications have reported the detection of C. pneumoniae by use of IHC in atheromatous plaques obtained from diverse sites in human [32, 35, 38, 61-73] and animal subjects [74-76]. Detection rates in human atheromata have varied widely (21%–71%) between various laboratories [15, 38, 61, 70]. Studies that have reported detection of C. pneumoniae in the same atheromatous plaque by use of both IHC and other methods, such as culture or PCR, have shown poor correlation between the different methods [40, 61, 70]. Typically, detection rates that are determined by use of IHC are higher than those determined by use of PCR. This is attributed to several factors, including faster degradation of DNA (compared with antigens), difficulty in extraction of DNA from atheromas (mostly due to calcifications), and the presence of PCR inhibitors [61, 70, 77]. It is possible that these discrepancies may be due, in part, to false-positive and false-negative results of IHC. However, this has been difficult to evaluate in the absence of an accepted and standardized approach for both the methodology and interpretation of IHC results.

Review of Currently Used Tests and Recommendations for IHC

Procedure. The most widely used IHC assay is the avidin-biotinylated immune-complex method [78]. However, there are interlaboratory variations at different steps in the method, including pretreatment of the tissues, tyramide signal amplification, and colorimetric detection. Because the effect of variation in the aforementioned procedural steps has not been evaluated, we cannot yet recommend a standardized approach. An IHC study that compares these methodological issues in control tissues is recommended.

Antibodies. The majority of published studies have used CF-2, a monoclonal antibody directed against the lipopolysaccharide of all Chlamydia species [71, 75, 77, 79], or RR-402 and TT-401, 2 antibodies that are specific for C. pneumoniae [61, 80, 81]. There is some evidence of differences in reactivity of the various antibodies [82]. Use of CF-2 is a reasonable compromise for initial screening because no other Chlamydia species has been found in atheromas, but patients positively identified by use of this antibody should be further tested with specific antibodies for C. pneumoniae and C. trachomatis [34, 62, 63, 68, 72, 73].

Negative control antibodies must be used for every specimen to assess background staining of the tissue. Most reported studies have used the antibodies in mouse ascitic fluid as the only negative control. However, the best negative antibody control should be an antibody of the same isotype as the Chlamydia antibody being used in the assay. For example, if CF-2, which is an IgG2a antibody, is used, the negative control antibody should be an IgG2a antibody directed against a different infectious agent that does not cross-react with Chlamydia species. Therefore, we recommend the use of 2 negative control antibodies for each tissue block. The first should be either normal mouse ascitic fluid or hyperimmune serum, and the second should be a non-Chlamydia antibody of the same isotype.

Tissues and controls. Both fresh and formalin-fixed tissues have been used for IHC Chlamydia testing. One positive and one negative tissue control should be carefully selected and used consistently with each experimental run. Theoretically, infected human specimens should be the best positive tissue control. However, IHC staining for C. pneumoniae in human atheromas has been described as very focal and scanty, and it has not been possible to identify tissue blocks that consistently yield a positive result. Other positive control tissues that are currently used include infected tissue culture cells and tissue from experimentally infected animals. Negative tissue controls included in each run should either be uninfected cells of the same type as the Chlamydia-infected cells, or they should be specimens of normal artery, brain, lung, or other tissues (as appropriate for the experimental tissue). In addition, at least 2 sections from the same paraffin block as the case being studied should be incubated with negative antibody controls and included in each run. In summary, each staining run should include 1 positive and 1 negative tissue control incubated with the 2 positive and 2 negative antibodies that are used on the specimen of interest.

Interpretation of results. Establishing the difference between signal and background is the crucial issue. Interpretation of IHC for infectious diseases tends to rely more heavily on correlations between signal morphology (granular staining, intact bacteria) and histopathological context (type of cell associated with signal). Accurate interpretation requires specialized training and, at a minimum, the ability to consistently identify and distinguish the major inflammatory cells (polymorphonuclear cells, mast cells, and plasma cells) and pigments (lipofuscin, hemosiderin) in tissue sections. Only intracytoplasmic IHC staining of macrophages, endothelial cells, and smooth muscle cells should be considered a positive result [61]. An intracytoplasmic granular staining pattern in the correct location and in the absence of artifact (as assessed by means of control slides) constitutes a true-positive result. There is disagreement regarding whether a homogenous staining pattern in the proper context can be considered a true-positive result or whether such staining should always be regarded as artifact.

The key conclusions and recommendations that resulted from this meeting are summarized in table 4 and should provide immediate guidance to those working with current C. pneumoniae assays. In addition, meeting participants emphasized the need for concentrated efforts on several future research priorities, all of which should benefit from a standardized approach to assay use, interpretation, and quality control.

Table 4
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Table 4

Recommendations for standardizing Chlamydia pneumoniae diagnostic assays.

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Study Group Members

The following were participants in the meeting on Chlamydia pneumoniae diagnostic assay standardization (Atlanta, 27–28 April 2000): Dr. Claudiu Bandea, Dr. Carolyn Black, Dr. George M. Carlone, Dr. Scott Dowell, Dr. Barry Fields, Dr. Jeannette Guarner, Dr. Trudy Messmer, Dr. Siobhán O'Connor, Dr. John Papp, Ms. Mindy J. Perilla, Dr. Anne Schuchat, Ms. Valerie Stevens, Dr. Deborah Talkington, Dr. M. Lucia Tondella, Dr. Chris A. Van Beneden, Dr. Sherif Zaki, and Ms. Elizabeth R. Zell (National Center for Infectious Diseases, Centers for Disease Control and Prevention, Atlanta); Dr. Cynthia Cohen (Immunohistochemistry and Image Cytometry, Anatomic Pathology, Emory University Hospital, Atlanta); Dr. Lee Ann Campbell, Dr. Cho Chou Kuo, Dr. San Pin Wang, (Department of Pathobiology), Dr. J. Thomas Grayston (Department of Epidemiology), and Dr. Lisa A. Jackson (Department of Epidemiology, School of Public Health and Community Medicine, University of Washington, Seattle); Dr. Carolyn D. Deal (Center for Biologics Evaluation and Research, FDA Division of Bacterial Products, Washington DC); Dr. Charlotte Gaydos (Infectious Diseases Division, Johns Hopkins Hospital, Baltimore); Dr. Margaret Hammerschlag (SUNY Health Science Center at Brooklyn, New York); Ms. Laura Schindler (Infectious Diseases Laboratory, University of Louisville, Kentucky); Dr. Christopher E. Taylor (National Institute of Allergy and Infectious Disease, Washington DC); Dr. Jim Mahony (Regional Virology and Chlamydiology Laboratory, St. Joseph's Hospital, McMaster University, Canada); Dr. Rosanna W. Peeling (Laboratory Centre for Disease Control, Health Canada, Ottawa, Canada); Dr. Ignatius William Fong (Division of Infectious Diseases, University of Toronto, Canada); Dr. Maija Leinonen (National Public Health Institute, Oulu) and Dr. Pekka Saikku (Department of Medical Microbiology, University of Oulu, Oulu, Finland); Dr. Matthias Maass (Institute of Medical Microbiology and Hygiene, Medical University of Lubeck, Germany); Dr. Jacobus M. Ossewaarde (Research Laboratory for Infectious Diseases National Institute of Public Health and the Environment, The Netherlands); Dr. Kenneth Persson (Department of Clinical Microbiology, Malmö General Hospital, Malmö, Sweden); Dr. Jens Boman (Department of Clinical Virology, University Hospital of Umea, Umea, Sweden); and Dr. Petra Apfalter (Department of Clinical Microbiology, Hygiene Institute, University of Vienna, Austria).

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Footnotes

  • ↵a Members of the study group are listed after the text.

  • Received November 2, 2000.
  • Accepted December 28, 2000.
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