0
Conferences |

The Laboratory Evaluation of Opportunistic Pulmonary Infections FREE

James H. Shelhamer, MD; Vee J. Gill, PhD; Thomas C. Quinn, MD; Stephen W. Crawford, MD; Joseph A. Kovacs, MD; Henry Masur, MD; and Frederick P. Ognibene, MD
[+] Article and Author Information

An updated, edited summary of a Combined Clinical Staff Conference held on 29 September 1994 at the National Institutes of Health, Bethesda, Maryland. Authors who wish to cite a section of the conference and specifically indicate its author may use this example for the form of reference: Gill VJ. Laboratory evaluation of specimens, pp 586-588. In: Shelhamer JH, moderator. The laboratory evaluation of opportunistic pulmonary infections. Ann Intern Med. 1996; 124:585-599. Requests for Reprints: James H. Shelhamer, MD, Building 10, Room 7-D-43, National Institute of Health, Bethesda, MD 20892 Current Author Addresses: Drs. Shelhamer, Kovacs, Masur, and Ognibene: Critical Care Medicine Department, Clinical Center, Building 10, Room 7-D-43, National Institutes of Health, Bethesda, MD 20892.


Copyright ©2004 by the American College of Physicians


Ann Intern Med. 1996;124(6):585-599. doi:10.7326/0003-4819-124-6-199603150-00008
Text Size: A A A

The patient population at risk for opportunistic pulmonary infections has increased during the last decade.The spectrum of organisms causing opportunistic infections has also grown. With an ever broader list of potential therapeutic options and a growing differential diagnosis, a specific diagnosis of the cause of pulmonary disease becomes more important. Recent microbiologic advances have helped to facilitate the laboratory diagnosis of some of these agents. Immunoassays are available for the detection of antigen in nasopharyngeal secretions (respiratory syncytial virus, influenza), in serum (Cryptococcus species), and in urine (Legionella or Histoplasma species). Rapid-culture techniques are available for the culture and detection of various viruses, including cytomegalovirus. Molecular probes can now assist in the rapid identification of Mycobacterium tuberculosis and some fungi. In the near future, polymerase chain reaction-based techniques may assist in the detection of Pneumocystis carinii and Legionella, Chlamydia, Mycoplasma, and mycobacteria species. An expeditious evaluation of pulmonary disease requires an understanding of the differential diagnosis of likely causes of pulmonary disease in specific immunosuppressed patient populations, an understanding of the most appropriate specimens to process for these diagnoses, and an understanding of the limitations (sensitivity and specificity) of these diagnostic tests. An understanding of the most appropriate specimens and tests in a given institution should allow for early, relatively specific treatment of many potentially life-threatening infections.

Dr. James H. Shelhamer (Critical Care Medicine Department, Clinical Center, National Institutes of Health [NIH], Bethesda, Maryland): Because the spectrum of processes that cause pulmonary disease in immunosuppressed patients is so broad, eminently logical reasons exist for obtaining specific information about the causative process—for example, to make certain that the patient receives appropriate therapy and that inappropriate, potentially toxic, and expensive therapies are avoided. However, these arguments for specific diagnosis are countered by valid concerns that the most useful diagnostic tests will be invasive and may ultimately be nondefinitive, thus incurring expense and morbidity without altering the clinician's initial management plan.

During the past several years, the ability to diagnose infectious causes of pulmonary disease has increased considerably. Sputum, tracheal secretion, bronchoalveolar lavage, blood, and even urine samples can be examined directly. The organism can be visualized by a tinctorial or fluorescent stain, or evidence of an organism may be detected by tests for specific antigens or nucleic acid. In some instances, with the use of newer methods, extremely sensitive assays can detect the presence of a pathogen within 24 or 48 hours of sample submission. Rapid-culture techniques can provide additional information within days about the presence of viruses, mycobacteria, or fungi as well as bacteria.

Major issues for clinicians who manage immunosuppressed patients have emerged. These issues include 1) the availability of assays for specific pathogens; 2) the sensitivity and specificity of each assay; 3) whether determining the presence of organism by culture, antigen detection, or detection of its nucleic acid proves that the organism is the cause of pulmonary dysfunction; 4) whether a particular test result is accurate and reliable enough to use to determine therapy and to make decisions about epidemiologic issues, including isolation precautions; and 5) whether the morbidity that some procedures entail, the staff time that is required for the collection and processing of the samples, and the expense that the procedures incur are truly warranted. In this Combined Staff Conference, we review some of the major advances that have been made in rapid identification of microbial pathogens, and we assess the role of these assays in clinical practice.

Dr. Vee J. Gill (Clinical Pathology Department, Clinical Center, NIH): As the range of pathogens that a microbiology laboratory can detect increases, clinicians need to recognize the optimal type of specimen to submit (Table 1) and the types of tests that can be done (Table 2). Although Gram stain and culture remain the mainstay of traditional microbiologic tests, the implementation of new methods in the clinical laboratory has led to more sensitive and more rapid detection techniques. These, in turn, can benefit patient care.

Table Jump PlaceholderTable 1.  Specimens for Optimal Diagnosis of Pulmonary Infections*
Table Jump PlaceholderTable 2.  Direct Stains*, Direct Tests, and Culture Available for Common Pulmonary Pathogens †

For routine bacteria, Gram stains of lower respiratory tract secretion or tissue samples are still important for providing rapid initial guidance on the morphologic characteristics of the bacteria. Although Gram stain of sputum has fallen into disfavor with some clinicians, an abundance of organisms (10 per high-power field) in an adequate specimen with neutrophils (25 per high-power field) is still considered highly suggestive of significance. However, both Gram stains and cultures are admittedly difficult to interpret in neutropenic patients or in those who have already received extensive treatment with antimicrobial agents. Some clinicians are enthusiastic about specimens obtained by protected brushes or bronchoalveolar lavage, both of which reduce upper airway contamination of specimens. Such specimens can then be cultured quantitatively [12]. Quantitative culturing can add sensitivity and specificity to the diagnosis of bacterial pneumonia, but it is time consuming. Many clinicians and laboratory directors question whether the additional information that is gained justifies the added expense.

In subsequent sections, we present the current state of the art as well as new methods for the diagnosis of viruses, Legionella species, Mycoplasma pneumoniae, Chlamydia pneumoniae, Pneumocystis carinii, mycobacteria species, and fungi. For each of these major groups of pathogens, specific technical advances are ready to be incorporated into clinical laboratory procedures, and newer methods are being developed for use in the near future.

Overview of New Laboratory Tests for the Diagnosis of Pulmonary Infections
Viruses

Rapid isolation and identification of respiratory viruses within 1 to 2 days can now be done using shell vial cultures that are stained with virus-specific fluorescent monoclonal antibodies. With the shell vial culture, the specimen is centrifuged onto a tissue culture monolayer contained within a screw-capped vial. The monolayer, which is grown on a coverslip within the vial, can be stained using various respiratory virus pools or virus-specific fluorescent monoclonal antibodies. By setting up multiple vials, one can stain a coverslip at 1-day or 2-day (or other) intervals as desired. This method is shown schematically in Figure 1. The technical simplicity of shell vial cultures has made it possible for them to be done in-house at many laboratories, providing useful information in a timely fashion.

Grahic Jump Location
Figure 1.
Schematic of the shell vial virus isolation technique.
Grahic Jump Location
Legionella Species

For the laboratory diagnosis of Legionella pneumonia, bronchoalveolar lavage fluid samples and lung biopsy specimens remain the most reliable specimens and are preferred to expectorated sputum samples. Currently available methods include direct fluorescent monoclonal antibody staining and cultures using supplemented media for Legionella species. A urinary antigen radioimmunoassay for L. pneumophila serotype 1 was introduced several years ago but has not been readily available because the radioimmunoassay format has not been practical for clinical microbiology laboratories to set up. A commercially available urinary antigen assay that uses an enzyme-linked immunoassay now enables more laboratories to use this test. However, because this test is highly specific for L. pneumophila serotype 1, infection with a different serotype or species would go undetected. Successful use of the polymerase chain reaction on respiratory specimens for the diagnosis of Legionella infection has recently been reported.

Mycoplasma and Chlamydia Species

Laboratory diagnosis of infection caused by M. pneumoniae and C. pneumoniae relies heavily on the detection of serologic conversion. Both agents can be cultivated, but with difficulty, so that this service is not routinely offered by clinical laboratories. In the future, polymerase chain reaction may be used to establish an early diagnosis for these pathogens.

In addition to shell vial culturing, fluorescent antibody staining of smears made directly from specimens such as bronchoalveolar lavage fluid samples can yield quick results. Reagents that can be used for direct staining of specimens are available for respiratory syncytial virus, cytomegalovirus, herpes simplex virus, and varicella-zoster virus. Commercially available enzyme immunoassays have become more diverse and offer technically easy yet rapid testing; such tests are already available for respiratory syncytial virus and influenza A virus. Methods that provide results on the same day that the specimen is submitted, either by direct staining or by enzyme immunoassay, are attractive to clinical laboratories, but each assay must be assessed to ensure that its sensitivity and specificity are adequate.

Pneumocystis carinii

Advances in the detection of P. carinii are attributable to improvement in rapid direct staining of induced sputum, bronchoalveolar lavage fluid, and lung biopsy specimens. The ability of microbiology laboratories to provide same-day staining with stains such as monoclonal fluorescent antibody stains has resulted in more sensitive and timely detection of pneumocystis pneumonia. Polymerase chain reaction has also been shown to increase detection of P. carinii, particularly when done on induced sputum specimens.

Mycobacteria Species

Mycobacterial isolation and identification have been significantly improved by the use of more rapid-culture techniques such as the radiometric BACTEC (Becton-Dickinson, Sparks, Maryland) system. Used in conjunction with new commercially available DNA probes (Accuprobe, Gen-Probe, San Diego, California), the time required for the isolation and identification of the significant mycobacterial pathogens has been greatly reduced. Figure 2 describes this innovative probe technology, which detects specific ribosomal RNA targets, allowing identification of an isolate within 1 day by a simple chemiluminescent assay.

Grahic Jump Location
Figure 2.
Simplified schematic of the Accuprobe hybridization assay (Gen-Probe).

*The selection reagent selectively splits the acridinium ester from the single-stranded DNA probe but not from the double-stranded hybrid.

Grahic Jump Location

Because direct detection of Mycobacterium tuberculosis in sputum specimens using molecular techniques such as polymerase chain reaction may yield even more sensitive and rapid diagnosis, continued progress in the use of these methods is especially warranted.

Fungi

Advances in fungal diagnostics have been limited. Direct detection by smears, culture, and specific antigen assays (for Cryptococcus and Histoplasma species) are the methods on which clinicians currently rely. One improvement that has increased the utility of direct smears for fungi is the use of the calcofluor white stain. This is a rapid, easy-to-read stain in which fungal elements brightly fluoresce, but it requires the use of a fluorescent microscope. Other recent developments include commercial DNA probes (Gen-Probe) [28] that are available for the identification of H. capsulatum, Coccidioides immitis, and Blastomyces dermatitidis. Use of these probes greatly reduces the time and increases the accuracy of identification of these pathogens.

Polymerase Chain Reaction as a Diagnostic Tool

As we have mentioned, polymerase chain reaction or similar amplification techniques are currently under development for the detection of many infectious agents. In most instances in which polymerase chain reaction has been applied, the results suggest that amplification-based detection will greatly increase our ability to make specific diagnoses. The basic polymerase chain reaction is a powerful tool that may eventually permit more definitive and rapid identification of difficult-to-detect or slow-growing organisms as well as of organisms that cannot be cultured. Polymerase chain reaction results in a doubling of copies of the specified target DNA with each round of amplification, eventually resulting in million-fold levels of amplification. Figure 3 shows how this amplification is achieved. The product of amplification must then be detected; this can be done in various ways, including agarose gel electrophoresis followed by staining with ethidium bromide. Using a sequence-specific probe to confirm the specificity of the amplified DNA product also increases sensitivity, but traditionally this probing has required the use of radioisotopes. Alternative nonradioisotopic methods [such as chemiluminescent or enzyme-coupled probes] to detect specific amplification products are preferred for use in the clinical laboratory setting and will help speed the incorporation of polymerase chain reaction as a diagnostic tool. The immediate availability of polymerase chain reaction testing in the clinical laboratory has been hindered by 1) the complexity and time-consuming nature of the assays, 2) the need for personnel trained in molecular methods to do the testing, and 3) the need to define an optimal protocol for each organism and then to document the level of sensitivity and specificity of that protocol. Other important problems, such as contamination leading to false-positive reactions, the presence of inhibitors in clinical specimens, and assessing the clinical relevance of positive polymerase chain reaction assays, remain to be resolved. Polymerase chain reaction assays to improve the accuracy of the diagnosis of cytomegalovirus, L. pneumophila, Mycoplasma pneumoniae, C. pneumoniae, P. carinii, and Mycobacterium tuberculosis infections have been worked on by many investigators; these assays are discussed in the following sections.

Grahic Jump Location
Figure 3.
Schematic of the polymerase chain reaction.
Grahic Jump Location

Dr. Stephen W. Crawford [Fred Hutchinson Cancer Research Center, Seattle, Washington]: The diagnosis of viral pneumonia requires 1) clinical evidence of a pneumonia, 2) the presence of virus in the lung or blood, and 3) a causal link between the isolated virus and the pulmonary disease.

Requirements for Diagnosis

Viruses associated with pneumonia in immunosuppressed patients include double-stranded DNA viruses such as herpesviruses (cytomegalovirus, herpes simplex virus, varicella-zoster virus, Epstein-Barr virus, and human herpesvirus 6) and adenovirus as well as single-stranded RNA viruses such as influenza A and B, paramyxoviruses (respiratory syncytial virus and parainfluenza viruses 1, 2, and 3), measles, and picornaviruses. These are obligate intracellular parasites. Replication requires host cellular machinery, and release of completed versions may be associated with cell lysis. Thus, unlike bacteria and fungi, viral presence indicates infection, not colonization. The degree to which the infection is accompanied by an inflammatory response and by organ dysfunction determines whether the viral infection causes “disease.” Excretion of herpes viruses, such as cytomegalovirus, can occur without pneumonia [35]. Thus, culture of virus from respiratory specimens indicates infection, but not necessarily disease, and isolation of virus from sites remote to the lung does not necessarily confirm the cause of the pneumonia.

Sites of Viral Isolation

Isolation of respiratory viruses from either respiratory tissues or secretions, or, in some instances, from the blood, are of potential clinical importance. Detection of virus in lung tissue is the most convincing evidence of a viral cause of a pneumonia. The significance of the detection of viruses in the upper respiratory tree varies with the type of virus. Respiratory viruses that contain RNA genomes, such as myxoviruses and paramyxoviruses (influenza, respiratory syncytial virus, parainfluenza), do not establish persistent infection (latency), and, with these viruses, isolation almost always indicates active viral disease. These viruses are rarely found in the absence of symptoms of upper respiratory infection [6], and their presence is of limited duration. Although nasopharyngeal viral excretion strongly suggests that a respiratory virus is the cause of pneumonia, the true specificity of such isolation in determining the cause of pneumonia is unclear; this isolation may represent an upper respiratory pathogen unrelated to the process causing pneumonia.

Herpesvirus and adenovirus that contain DNA may result in lifelong persistence of infection and are capable of reactivation. Excretion from extrapulmonary sites, including the throat and blood, suggests disease but is not diagnostic for a pneumonia and may occur in asymptomatic patients [46].

Methods of Virus Detection

Respiratory viruses can be detected by many techniques, including culture (conventional tube or shell vial cultures), detection of viral antigens by monoclonal antibody staining or by enzyme immunoassays, detection of viral nucleic acids by in situ hybridization or polymerase chain reaction, and serologic studies. Standard “tube” culture using cell types that are sensitive to viral infection and that show typical cytopathic changes when infected is the “gold standard” for respiratory virus detection. Tube culture may require weeks of incubation in some cases (for example, cytomegalovirus). Rapid culture of most respiratory viruses has been achieved with centrifugation (shell vial) cultures, in which the presence of viral replication in the tissue culture is confirmed with fluorescent monoclonal antibodies within 1 to 2 days of inoculation [78]. The sensitivity and specificity of centrifugation culture of bronchoalveolar lavage fluid for the detection of cytomegalovirus (and presumably of other respiratory viruses) approximates that of culture of lung tissue [9]. Culture is sensitive for cytomegalovirus and herpes simplex virus, whereas respiratory syncytial virus and parainfluenza are successfully cultured less often. False-negative results due to technical factors (such as factors in blood and respiratory secretions that are cytotoxic for the tissue culture monolayer) are seen, and specialized laboratories are required to maintain appropriate cell cultures for viral incubation.

Serologic conversion provides evidence of exposure. However, it lacks sensitivity and specificity for a given episode of pneumonia, and documentation takes several weeks. Serologic evidence is often detected after the clinical need to establish a diagnosis. Serologic studies are of value in identifying immunosuppressed patients at risk for reactivation of latent viruses, such as cytomegalovirus and herpes simplex virus [10].

Characteristic histologic and cytologic abnormalities can be seen in cytologic and histopathologic specimens in some cases of viral pneumonia. “Owl's eye” Cowdry type A intranuclear inclusions are relatively specific for cytomegalovirus pneumonia, “smudge” cells for adenovirus, and the respiratory syncytial cell for respiratory syncytial virus infection, but these are insensitive markers. Such cytologic changes are specific when seen in cytospin preparations for bronchoalveolar lavage, but they are of low sensitivity [9, 11].

Specific and sensitive detection of viral antigens within infected cells can be achieved from respiratory secretion (including bronchoalveolar lavage fluid) and blood samples using labeled monoclonal antibodies by most virology and pathology laboratories within hours of sample collection [9, 1113]. Direct fluorescent antibody staining of bronchoalveolar lavage or of nasopharyngeal secretions correlates closely with culture results, and the sensitivity probably is increased with the inclusion of pooled monoclonal antibodies to multiple antigenic sites [11].

Similarly, peroxidase-labeled antibody staining of the peripheral blood buffy coat is highly specific in some immunosuppressed hosts (such as patients receiving transplants) for tissue-invasive cytomegalovirus diseases, with the presence of antigenemia correlating with disease. The sensitivity of peroxidase-labeled monoclonal antibodies to the cytomegalovirus-encoded immediate-early antigen, pp65, is similar to that of culture in most patients and tends to be detected earlier than culture [1421]. Commercially available enzyme immunoassay kits permit the detection of viral antigens in respiratory secretions. These assays appear to be most useful in the detection of respiratory syncytial virus, parainfluenza virus, and possibly influenza viruses in nasopharyngeal specimens [2223].

Detection of viral DNA is possible from tissue with in situ hybridization and from respiratory specimens and blood samples with polymerase chain reaction [2425]. Polymerase chain reaction of blood leukocytes for cytomegalovirus is more sensitive than either culture or assay for antigenemia [2630]. However, because of its high sensitivity, it may detect a small copy number of virus genome with a resulting low positive but high negative predictive value for disease [3132]. Additionally, prompt processing (within hours) of clinical specimens is required. Thus, the clinical utility of peripheral blood polymerase chain reaction is probably limited to excluding cytomegalovirus as a potential cause of an episode of pneumonia. As research tools, polymerase chain reaction and in situ hybridization will help to clarify the biology of viral pneumonia.

Virus as a Cause of Pneumonia

It is sometimes difficult to determine with certainty that a virus isolated from the lung is the cause of a pneumonic process (Table 3). The strongest evidence is both detection of typical histopathologic changes (such as intranuclear inclusion bodies typical of cytomegalovirus infection) and isolation of the virus in culture in the absence of other identifiable causes of pneumonia. However, such manifestations of infection are not uniformly present [33], and lung tissue is not always available for pathologic evaluation. Often the diagnosis is presumed, on the basis of viral detection from appropriate fluids, in a patient at risk who has characteristic clinical signs.

Table Jump PlaceholderTable 3.  Significance of Respiratory Virus Detection in the Presence of Pneumonia in Patients Receiving Bone Marrow and Solid Organ Transplant*

Bone marrow transplant recipients with diffuse pneumonia have a high mortality rate if cytomegalovirus is cultured from the lung, regardless of the presence of typical cytomegalovirus infection histopathology [10, 29]. Also, isolation of cytomegalovirus from bronchoalveolar lavage fluid specimens or blood samples is associated with a high (> 50%) incidence of subsequent pneumonia among asymptomatic bone marrow transplant recipients [4, 6]. In addition, quantitation of cytomegalovirus in bronchoalveolar lavage fluid specimens does not correlate with disease [34]. Therefore, the operational definition of cytomegalovirus pneumonia after marrow transplantation is detection of cytomegalovirus from lungs in the presence of pneumonia. The diagnosis of viral pneumonia among recipients of solid organ transplantation may be similarly viewed. However, many centers restrict the diagnosis of definite viral pneumonia to those cases with typical cytologic changes in which no other pathogens have been identified [12]. This approach may under-represent the true incidence of viral pneumonia. Cytomegalovirus rarely causes pneumonia in patients with the acquired immunodeficiency syndrome (AIDS) despite the frequent isolation of the virus in these patients. The presence of cytomegalovirus does not appear to influence mortality, regardless of treatment [3536].

Approach to Diagnosis of Viral Pneumonia

All immunosuppressed patients with diffuse pneumonia should be evaluated for viral pneumonia. Among patients seropositive for cytomegalovirus, an antigenemia test with negative results probably excludes cytomegalovirus pneumonia. Positive results from nasopharyngeal swab by enzyme immunoassay, direct fluorescent antibody, or culture may increase suspicion of respiratory viral infection (respiratory syncytial virus, influenza, parainfluenza) among patients with concomitant symptoms of upper respiratory infection, but they do not confirm the cause. Minimal evaluation should include bronchoscopy with bronchoalveolar lavage. The degree to which lung biopsy helps in the diagnosis of viral infection is unclear. All bronchoalveolar lavage fluid specimens should be examined for viruses by routine cytologic studies and, ideally, for cytomegalovirus and herpes simplex virus by centrifugation culture. During endemic seasons, cultures for respiratory syncytial virus, influenza, and parainfluenza should be done. If no viral culture facilities are available, monoclonal fluorescent antibody staining with probes specific for these viruses should detect most cases.

Dr. Thomas C. Quinn (National Institute of Allergy and Infectious Diseases, NIH): Legionella species, Mycoplasma pneumoniae, and C. pneumoniae are now well-recognized pathogens that warrant immediate therapy. None of the organisms grow on standard bacteriologic media; none are identified on Gram stain; and none respond to the standard empiric choice of penicillin or cephalosporin antibiotic therapy frequently used to treat pneumonia.

Legionella Species

Fifteen serogroups of L. pneumophila based on surface antigen analysis and 33 other Legionella species have now been identified, many of which have been shown to be responsible for pneumonia [3739]. Serogroup 1 is responsible for about 80% of disease caused by L. pneumophila, with serogroups 4 and 6 responsible for most other cases [38]. Infections caused by Legionella species other than L. pneumophila are uncommon, constituting less than 20% of infections. Of these pathogens, L. micdadei, L. longbeachae, L. dumoffii, and L. bozemanii appear to be the most common.

Five methods are currently used for the laboratory diagnosis of Legionella infections (Table 4). These include isolation of the organism on selective culture medium, determination of antibody level, detection of the bacterium in tissue or body fluid specimens using direct fluorescent antibody tests, detection of antigenuria, and detection of DNA by the polymerase chain reaction.

Table Jump PlaceholderTable 4.  Laboratory Diagnosis of Legionella Pneumonia*

Legionella species can be isolated from clinical specimens on selective medium using supplemented charcoal yeast extract medium (BCYE α) [38, 40]. The organism has been successfully isolated from sputum, transtracheal aspirates, endotracheal suction specimens, blood, biopsied lung tissue, pleural fluid, bronchial lavage fluid, pericardial fluid, and peritoneal fluid and from the respiratory sinuses. The disadvantage of culture is that it takes 5 to 10 days for colonies to appear, and the specimens must be processed in a precise manner on the abovementioned selective media for optimal isolation.

Acute and convalescent serologic studies are of diagnostic value, but it takes 6 to 8 weeks for the antibody titers to increase, thereby limiting the diagnostic value of these studies early in the course of the infection [41]. In immunocompromised patients, the antibody response may be severely impaired. Most laboratories use the indirect immunofluorescent antibody technique to determine antibody concentrations. About 75% of immunologically normal patients with culture-proven legionnaire's disease caused by L. pneumophila serogroup 1 develop a fourfold increase in titer by 8 weeks after onset of illness. Because up to 30% of healthy populations sampled have L. pneumophila serogroup 1 antibody titers of 1:128 or greater, only a fourfold increase in titer can be considered significant [4142]. Cross-reactions have also been reported in patients with other causes of pneumonia, and specificity has been estimated to be 90% for a fourfold titer increase.

The direct fluorescence antibody test is a rapid-detection test that uses a monoclonal antibody to all serogroups of L. pneumophila[38, 43]. This technique has been used successfully with expectorated sputum, endotracheal suction aspirates, biopsied lung tissue, and transtracheal aspirates. Use of secretions or biopsied tissue obtained by bronchoscopy has not resulted in high yield. The true sensitivity of the direct fluorescence antibody test is unknown, although 25% to 70% of patients with culture-proven L. pneumophila infection have positive results from direct fluorescent antibody tests [44]. The test specificity is greater than 90%, but cross-reactivity to Bordetella pertussis occurs. Results of the direct fluorescence antibody tests of sputum remain positive for 2 to 4 days after initiation of specific antibiotic therapy for L. pneumophila.

L. pneumophila serogroup 1 antigenuria can be detected using a commercial radioimmunoassay [45]. Cross-reactions between serogroups are uncommon, thereby limiting the usefulness of this test for the diagnosis of other Legionella infections. The advantage of this urine test is its high sensitivity, estimated to be 95% in culture-proven cases and 80% in patients with serologically proven disease. Specificity is also very high (estimated to be 99%). An enzyme-linked immunoassay is now commercially available. Test results can be obtained the day of testing, although it should be noted that they may remain positive for weeks to months after recovery from pneumonia.

Although not commercially available yet, amplification of Legionella nucleic acids by polymerase chain reaction has the potential to offer rapid results and to increase the sensitivity of current detection methods used on respiratory samples within a 24-hour period [46]. Polymerase chain reaction has been proven effective in the detection of L. pneumophila in bronchoalveolar lavage fluid specimens, nasopharyngeal swabs, and sputum samples. In a recent study of 40 immunocompromised patients with pneumonia at the NIH, we were able to use bronchoalveolar lavage fluid samples to detect L. pneumophila in 4 patients (10%) [47]. Polymerase chain reaction is also currently used for the detection of Legionella species in environmental samples [48]. Because primers are used for the detection of the 5s rRNA gene of Legionella species, polymerase chain reaction is capable of detecting Legionella species other than L. pneumophila.

Mycoplasma pneumoniae

The laboratory diagnosis of Mycoplasma pneumoniae infection can be made either by culture on selective media or by detection of an appropriate increase in specific antibody titer (Table 5). Mycoplasma pneumoniae can be isolated from throat washings, sputum, or throat swabs 7 to 10 days after inoculation in broth media [49]. A presumptive identification of Mycoplasma pneumoniae can be made if colonies show heme absorption of red blood cells.

Table Jump PlaceholderTable 5.  Diagnostic Tests for Mycoplasma pneumoniae

Serologic diagnosis depends on a fourfold increase in complement fixation titer for acute and convalescent sera [50]. IgM antibody first appears 1 week after infection, peaks at 4 to 6 weeks, and does not start to decrease until 4 to 6 months later. One third to three fourths of patients with Mycoplasma pneumoniae develop a fourfold or greater increase in titers of cold hemagglutinin [51], but these IgM antibodies are not diagnostic of Mycoplasma pneumoniae infection because they can be associated with various other infections as well as with connective tissue and neoplastic disorders. Polymerase chain reaction for Mycoplasma pneumoniae infection has also recently become available [5253]. In a recent study of 34 patients with respiratory illness and evidence of pneumonia, evidence of Mycoplasma pneumoniae infection was obtained in 10 patients (29%); in 8 of these 10 patients, results of both polymerase chain reaction and serologic studies were positive. However, it should be noted that the detection of Mycoplasma pneumoniae in the respiratory tract does not necessarily correlate with respiratory disease, at least not in an immunocompetent patient. Consequently, serologic tests should be used in addition to polymerase chain reaction to distinguish between acute and persistent infections.

All three Chlamydia species—C. psittaci, C. trachomatis, and C. pneumoniae—have been associated with pulmonary infection [54]. Chlamydia psittaci is responsible for psittacosis, a disease of birds, of which man is an accidental host. Chlamydia trachomatis is a common cause of pneumonitis in infants born to women with genital infection. Of the three species, C. pneumoniae, formerly known as TWAR, is the most common cause of acute respiratory disease, responsible for 5% to 15% of cases of community-acquired pneumonia [55], and according to one survey, for 10% of cases of pneumonia in immunocompromised patients [56].

Laboratory diagnosis usually depends on isolation of the organism, an increase in serologic titer, or polymerase chain reaction (Table 6). Because C. pneumoniae is a fastidious organism, it is difficult to isolate and requires in vitro tissue culture. It has been isolated in either HL cells or HEp-2 cells [5758]. Nasopharyngeal swabs provide the best specimen, but they must be transported in specialized transport media and inoculated immediately in the cell culture. Culture results are not usually positive for 72 hours or longer.

Table Jump PlaceholderTable 6.  Diagnostic Tests for Chlamydia pneumoniae

Most investigators have relied on serologic diagnosis using the microimmunofluorescence test. Grayston and colleagues [55] proposed criteria for serologic diagnosis of C. pneumoniae infection that have been used by many laboratories and clinicians. Patients with acute infection have a fourfold increase in the IgG titer, a single IgM titer of 1:16 or greater, or a single IgG titer of 1:512 or greater. Patients with past or preexisting infection have an IgG titer of at least 1:16 and less than 1:512. Because the microimmunofluorescence assay is not widely available, the complement fixation test has been used as an alternative. This test is genus-specific and should be used primarily for the diagnosis of lymphogranuloma venereum and psittacosis. Grayston and colleagues [55] found that about one third of patients with C. pneumoniae had detectable complement fixation antibody.

Because of the limitations of both culture and serologic studies for the establishment of early diagnosis, polymerase chain reaction has recently been developed, providing a sensitive and specific assay for early detection [56, 59]. Using primers for either the major outer membrane protein or the 16s rRNA gene of C. pneumoniae, polymerase chain reaction has been shown to be highly sensitive and specific. In one study at the Clinical Center, National Institutes of health, we screened 132 culture-negative bronchoalveolar lavage fluid specimens from 108 immunocompromised patients (34% of whom tested positive for human immunodeficiency virus [HIV] infection) and 7 healthy volunteers [56]. Thirteen specimens (9.8%) from 12 immunocompromised patients (11.1%) yielded positive results. No healthy volunteer had a positive polymerase chain reaction in a bronchoalveolar lavage specimen. Only 2 of these 12 infected patients had other microbiologic agents, including respiratory syncytial virus and P. carinii, implicated in the pneumonia. Only 4 of the 13 patients had diagnostic titers of antibody to C. pneumoniae. The inability of patients to respond to specific antigenic stimuli promptly or at all may have been caused by their immunocompromised state. Thus, more rapid tests such as polymerase chain reaction may be a useful addition to the more conventional methods.

Dr. Joseph A. Kovacs (Critical Care Medicine Department, Clinical Center, NIH): Despite the extensive use of prophylaxis, Pneumocystis carinii remains one of the most common causes of opportunistic pneumonia in immunosuppressed patients, especially patients with HIV infection. Human P. carinii cannot be cultured in vitro at present; therefore, unambiguous diagnosis of P. carinii pneumonia requires direct detection of the organism in an appropriate clinical specimen. Furthermore, because available data suggest that P. carinii is not a colonizing pathogen, detection of P. carinii in pulmonary samples from patients who have not received treatment, at least by standard tinctorial or immunofluorescent assays, is currently equivalent to diagnosis of P. carinii pneumonia.

Bronchoscopy using either bronchoscopic biopsy or bronchoalveolar lavage has a very high diagnostic yield. The major advantage of bronchoscopic biopsy is that multiple small specimens can be obtained and histopathologically examined for pathogens other than P. carinii. For P. carinii pneumonia, the sensitivity of transbronchial biopsy or bronchoalveolar lavage individually is more than 90% [6062].

Sputum induction using hypertonic saline has replaced bronchoscopy as the initial diagnostic procedure at many centers. This noninvasive procedure has a diagnostic yield of 50% to 90%. Increased experience in both obtaining and examining the sample has resulted in yields of more than 80% at many institutions [6364]. It is a rapidly done technique with relatively few risks, although it may not detect other potential causes of pneumonia. Negative results may require follow-up bronchoscopy if clinical circumstances warrant definition of the causative process.

For many years, the standard method for detecting P. carinii in clinical samples has been tinctorial staining [65]. Tinctorial stains include the cyst wall stains, such as toluidine blue-O and methenamine silver stains, which will stain only the cysts of P. carinii and not the more numerous trophozoite form of the organism. Cyst wall stains also stain fungi. The second category of tinctorial stains, Giemsa and Diff-Quik, stain not only the intracystic sporozoite, but also the more numerous trophozoite form; unfortunately, they will also stain other organisms, such as yeast, as well as cells and subcellular debris, and this makes them more complex to interpret. Recently, monoclonal antibodies to human P. carinii, which react with both cysts and trophozoites, have been found to be very useful diagnostically [64]. Prospective blinded studies have shown that immunofluorescent assays have a higher sensitivity than tinctorial stains, primarily in the examination of induced sputum samples (69% to 92% compared with 28% to 80%, respectively) [64, 6668]. These monoclonal antibodies are not known to cross-react with any other microbial forms likely to be found in human specimens and thus are highly specific.

Detection of organisms using the polymerase chain reaction has recently been applied to P. carinii[6970]. The most commonly used primers are based on the mitochondrial ribosomal RNA [69]. The major advantage of polymerase chain reaction is an increased sensitivity compared with tinctorial or immunofluorescent stains, but this comes at a cost of decreased specificity, which may be due to low numbers of organisms that are not causing the disease or to polymerase chain reaction-related technical problems. In a recent study comparing immunofluorescence to an enzyme-based polymerase chain reaction assay for examination of both sputum and bronchoalveolar lavage fluid specimens, the sensitivity in sputum specimens increased from 78% with immunofluorescence to 100% with the polymerase chain reaction assay with virtually no loss of specificity [71]. For bronchoalveolar lavage fluid samples, the two techniques were equally sensitive at 100% and virtually identical in terms of specificity. Based on these and other data, it appears that in bronchoalveolar lavage fluid examination, polymerase chain reaction will have no role in diagnosis of P. carinii infection, but it may have a diagnostic role in sputum examination. This test may also have a role in the typing of strains as well as in epidemiologic and environmental studies.

A final issue in the diagnosis of P. carinii pneumonia is the effect that aerosol pentamidine prophylaxis has on the detection of P. carinii. The use of aerosol pentamidine prophylaxis has been reported to be associated with an increase in upper lobe disease and a concomitant decrease in the sensitivity of both routine bronchoalveolar lavage and induced sputum examinations [7273]. Sensitivity in this situation can be improved to more than 90% by multilobe lavage including the upper and the middle lobe [7475]. The rare cases of extrapulmonary disease, which also appear to be more common in patients receiving aerosol pentamidine, can be diagnosed either clinically (for example, choroiditis) or histopathologically [7677].

Dr. Henry Masur [Critical Care Medicine Department, Clinical Center, NIH]: To diagnose mycobacterial disease in immunosuppressed patients, three basic issues must be addressed: 1) whether mycobacterial disease is present, 2) whether the mycobacterial isolate is Mycobacterium tuberculosis, and 3) whether the mycobacterium is clinically important. Mycobacterium tuberculosis receives special consideration not only because of its pathogenicity, since other mycobacteria can also cause life-threatening disease, but because of its potential for person-to-person transmission and the epidemiologic and economic need to determine if respiratory isolation for the patient is or is not necessary.

To determine if mycobacterial disease is present, most laboratories still rely on direct microscopy to detect organisms in smears from appropriate specimens [7879]. Indirect tests relying on detection of IgG or IgM antibodies in serum, on detection of enzymes such as adenosine deaminase or antigens in various types of clinical specimens, or on detection of cell wall lipids by chromatographic techniques have evoked considerable interest, but none of these tests shows a combination of sensitivity, specificity, and practicality adequate to warrant its routine use [78]. Direct microscopy is most often done on respiratory secretions, although it can obviously be applied to any clinical specimen. For most adults, expectorated or induced sputum specimens are screened initially, with bronchoalveolar lavage or lung biopsy necessary only in difficult diagnostic situations [78, 80]. With adequate encouragement, at least 80% of patients should be able to produce an expectorated or induced sputum sample. Gastric lavages are not commonly used, except in young children who cannot produce sputum; these specimens are useful for culture and for examination by direct techniques, despite earlier concerns about the frequent occurrence of nonpathogenic mycobacterial saprophytes in the stomach [81]. The sensitivity of direct microscopy depends on the stain used, the quality of the specimen, and the time and expertise involved in examining it; generally, at least 105 organisms/mL of specimen must be present, although some laboratories appear to be capable of detecting 102 to 103 organisms/mL [78]. This technique is very specific for identifying the presence of mycobacteria, but not for species identification. Laboratories are encouraged to use a fluorochrome stain such as auramine rhodamine for screening rather than a tinctorial stain such as the Ziehl-Neelson stain because the former is faster for screening specimens, requires less expertise to be read accurately, and is believed by many to be more sensitive.

Nucleic acid probes for detecting mycobacteria directly in clinical specimens initially elicited considerable interest but did not prove to be sufficiently sensitive to detect culture positive specimens reliably. Nucleic acid amplification techniques such as polymerase chain reaction have a greater potential for detecting fewer organisms and have been shown to be quite useful in detecting Mycobacterium tuberculosis; polymerase chain reaction based on the insertion element 156110 [8183] and the rRNA amplification system developed by Gen-Probe, the Amplified Mycobacterium Tuberculosis Direct Test [82, 84], have shown very high sensitivity for smear-positive specimens, and have detected Mycobacterium tuberculosis in a substantial fraction of smear-negative samples. These techniques are specific for Mycobacterium tuberculosis and do not give a positive signal when other mycobacteria are present, but they may give a positive signal in some patients with previously treated Mycobacterium tuberculosis disease or who are purified protein derivative (PPD)-positive without active disease, which complicates the interpretation of a positive result [82]. A negative amplification result is useful for determining that acid-fast organisms, if present, are not Mycobacterium tuberculosis and that Mycobacterium tuberculosis is unlikely to be present, which can be epidemiologically important. A positive result may indicate an active, inactive, or latent process. The use of polymerase chain reaction and other molecular methods is an exciting prospect for the rapid, accurate diagnosis of mycobacterial disease. Experience thus far has been limited to a few laboratories that have the expertise to develop their own methods. As yet, unfortunately, no simple test is available commercially in the United States to enable laboratories and physicians to benefit from the potentially increased sensitivity and speedy detection that these methods permit.

Culture of a specimen is the most sensitive technique for establishing the presence of active mycobacterial disease. Specimens can be plated on solid agar such as Lowenstein-Jensen culture medium, or in liquid medium such as Middlebrook 7H/11. The time until growth is detected depends on the size of the inoculum, the specific mycobacterial species and strain, and the presence of inhibiting drugs. Liquid systems are generally faster than solid medium systems for detecting growth [7879]. Radiometric culture systems that operate on the principle of a radiolabeled substrate being metabolized into a detectable gas are considerably faster and are being used by an increasing number of large laboratories; results are generally available 7 to 10 days earlier than with conventional solid media [79].

Respiratory specimens are most often the material cultured, but any potentially infected body fluid or tissue should be considered. In some instances, certain mycobacteria other than Mycobacterium tuberculosis may grow, but this may represent colonization or even contamination of the specimen. Mycobacterium avium complex can often be the former, and Mycobacterium gordonae, the latter. Pleural fluid and blood should not be overlooked as specimens for culture. In patients with human immunodeficiency virus infection, for example, blood cultures for Mycobacterium avium complex can be positive in many patients, especially those with low CD4+ lymphocyte counts [85]. If the BACTEC radiometric culture system is used, medium 13A should be used for blood samples rather than 12B, which is used for all other specimens.

Although nucleic acid probes are not used for direct testing of specimens, they have become a major advance in the laboratory's ability to identify specific mycobacterial groups or species accurately, once there is adequate growth of mycobacteria in a solid or liquid medium. Commercially available probes (such as Accuprobe) can identify Mycobacterium tuberculosis complex, Mycobacterium avium complex, Mycobacterium kansasii, and Mycobacterium gordonae within hours [7879]. This novel molecular approach (Figure 2), using DNA probes directed against ribosomal sequences, has dramatically improved identification of mycobacteria, reducing identification time from several weeks to 1 day once sufficient growth is present.

Is the mycobacterium identified from a specimen always important? Although the presence of Mycobacterium tuberculosis always represents disease that must be treated, and Mycobacterium kansasii usually represents disease requiring treatment as well, there are no universal rules for determining when other mycobacteria are pathogens, colonizers, or contaminants. The quantity of mycobacteria, the species, the host, and the clinical situation must be assessed. Some generalizations are helpful. As noted above, Mycobacterium gordonae has been identified as a true pathogen only rarely: It is most often a waterborne contaminant. Mycobacterium avium complex rarely causes localized or disseminated disease in any population other than patients with HIV infection. (However, a few cases of Mycobacterium avium complex causing parenchymal lung disease in persons with no underlying processes do occur.) In patients with HIV infection, blood isolates almost always represent disseminated disease, yet sputum, urine, or stool isolates often represent colonization and have not been shown to predict efficiently the future development of dissemination in studies done to date.

Dr. Frederick P. Ognibene (Critical Care Medicine Department, Clinical Center, NIH): The most common fungal pathogens responsible for pulmonary infections in immunocompromised hosts and the tests that may be helpful in the diagnosis of pneumonia caused by these organisms are shown in Table 7. A major frustration in pulmonary diagnostics has been the inability to develop useful tests for the fungi of most concern, Candida species and Aspergillus species.

Table Jump PlaceholderTable 7.  Utility of Diagnostic Tests for Fungal Pneumonia*

The isolation of Aspergillus species from respiratory secretions had been regarded as limited in usefulness in the antemortem diagnosis of invasive pulmonary aspergillosis. The detection of tissue invasion by fungal hyphae was considered the “gold standard” for diagnosis. However, Yu and colleagues [86] published data that indicated that in all of their patients with leukemia or neutropenia or both who ultimately had documented invasive aspergillosis, either A. fumigatus or A. flavus was isolated from cultures of the respiratory tract. They concluded that the isolation of Aspergillus organisms from sputum or lavage culture was highly predictive of invasive pulmonary infection, and many physicians follow their approach despite uncertainty about the specificity of this finding [8687]. Histopathologic evidence of fungi with the characteristic 45-degree angle branching and transverse septae confirm the diagnosis. No serologic studies for Aspergillus species are adequately sensitive or specific [8889].

Zygomycetes genera, including Mucor and Rhizopus, may also cause serious pulmonary disease in severely immunocompromised patients. A sputum or bronchoalveolar lavage specimen with a positive wet mount showing characteristic morphologic features or a positive culture strongly suggests that these agents are the cause of the pulmonary disease. Histopathologic specimens of the lung reveal broad, nonseptate hyphae with irregular right-angle branching. No reliable serologic tests exist for diagnosis.

Infection with Fusarium species may mimic the presentation of aspergillosis. Patients may have associated skin lesions or orbitofacial involvement. Blood cultures are frequently positive and provide a definitive diagnosis. No useful serologic studies exist for this pathogen [90].

For cryptococcosis, a sputum or bronchoalveolar lavage specimen that is either smear- or culture-positive for Cryptococcus neoformans is considered diagnostic because this encapsulated organism does not occur as a commensal. Although serologic tests may yield positive results for cryptococcal antigen in as many as 90% of patients with cryptococcal meningitis, results are positive in only 50% of patients with pneumonia alone [91]. A histologic specimen is often necessary to establish the diagnosis of cryptococcal pulmonary disease, because sputum and pleural fluid cultures may be negative in half of patients.

The diagnosis of disseminated histoplasmosis depends on a positive culture, typically from blood. For a pulmonary infection, a positive bronchoalveolar lavage smear or culture is diagnostic. However, it is important to note that a 2- to 4-week period may be required for Histoplasma capsulatum to grow in cultures. The histopathologic detection of intracellular organisms in viable pulmonary tissue is diagnostic evidence of disseminated disease. In nonimmunosuppressed patients, serologic studies are highly sensitive and provide confirmatory diagnostic evidence of Histoplasma infection [92]. Complement fixation has been used to detect antibody, and a titer of 1:32 or greater or a fourfold increase in titer is evidence of active or recent infection. Immunodiffusion studies are more specific but less sensitive than complement fixation. The presence of both immunoprecipitation bands (called M and H) is the most specific serodiagnostic finding, but it is also rare. A positive M band alone is a fairly specific finding. These serologic responses may occur in less than 50% of immunosuppressed patients. Detection of H. capsulatum antigen in specimens of blood and urine is done at only a few centers, but detection of antigen is believed to be useful for the diagnosis of disseminated histoplasmosis [93].

Coccidioides immitis can cause an acute, progressive pneumonia that may lead to dissemination to extrapulmonary foci. A positive culture of sputum or bronchoalveolar lavage is diagnostic. Wet mount staining is often helpful when the characteristic spherules with endospores are detected. Elevated complement fixing antibody studies are a hallmark of disseminated disease; titers of at least 1:32 indicate disseminated disease in most patients [92].

For the accurate diagnosis of Candida pneumonia, routine sputum and bronchoalveolar lavage cultures are not useful. Candida pneumonia is probably an unusual cause of pulmonary dysfunction, although the lung may be extensively involved with microemboli in patients with disseminated candidiasis. Histologic evidence of inflammation in the presence of Candida organisms is believed to be the most convincing evidence for Candida pneumonia. Blood cultures are not always positive in invasive disease caused by Candida species. In many patients with disseminated disease (not just pneumonia), blood cultures may be negative [94]. No reliable serologic studies exist for this pathogen [89].

Dr. Shelhamer: For an increasing number of community-acquired and opportunistic pathogens, new diagnostic tests that permit rapid detection of organisms in clinical specimens are becoming available. These test include direct detection systems that allow identification of organisms in clinical specimens within 24 hours by microscopy using tinctorial or immunologic staining techniques, by detection of antigen, or by detection of nucleic acid. Culture systems are also becoming more sensitive and more rapid, often providing results within a few days.

Should every reference diagnostic laboratory do all of these tests? In many cases, these tests are expensive and require considerable expertise. Some laboratories may not have the volume or the appropriate number of positive specimens during the course of a year to warrant doing each test. Some tests may not provide accurate, sensitive, or specific enough information to alter clinical management and thus are not practical for many cost-conscious facilities.

Even if these tests are done, clinicians need considerable expertise to interpret them appropriately. Each test has very specific meaning; unfortunately, these tests may signify different clinical information in different patient populations, and thus no easy, universally appropriate algorithm can be devised for their interpretation.

It is important for clinicians to be cognizant of the availability of these evolving tests. Some of the tests can be extremely valuable in certain well-defined situations, even if all of them cannot be routinely done. For instance, any center that has a large population of allogeneic bone marrow transplant recipients might consider doing rapid cytomegalovirus tests, such as some combination of shell vial cultures, buffy-coat antigen tests, or even polymerase chain reaction. A hospital with a high rate of Legionella infection, tuberculosis, or respiratory syncytial virus might consider instituting the appropriate rapid diagnostic test to ensure appropriate therapy in the case of legionellosis and to prevent nosocomial transmission as well as ensure appropriate therapy in the cases of tuberculosis and respiratory syncytial virus infection. Thus, institutions will have to pick and choose from this increasingly complex but useful menu, and develop tests that are appropriate for their patient population, their epidemiologic trends, and their budget.

Dr. Gill: Clinical Pathology Department, Clinical Center, Building 10, Room 2-C-385, National Institutes of Health, Bethesda, MD 20892.

Dr. Quinn: 1159 Ross Research Building, 720 Rutland Avenue, Baltimore, MD 21205.

Dr. Crawford: Pulmonary and Critical Care Medicine, Fred Hutchinson Cancer Research Center, 1124 Columbia Street, Seattle, WA 98104.

Thorpe JE, Baughman RP, Frame PT, Wesseler TA, Staneck JL.  Bronchoalveolar lavage for diagnosing acute bacterial pneumonia. J Infect Dis. 1987; 155:855-61.
 
Xaubet A, Torres A, Marco F, Puig-De la Bellacasa J, Faus R, Agustividal A.  Pulmonary infiltrates in immunocompromised patients. Diagnostic value of telescoping plugged catheter and bronchoalveolar lavage. Chest. 1989; 95:130-5.
 
Ruutu P, Ruutu T, Volin L, Tukiainen P, Ukkonen P, Hovi T.  Cytomegalovirus is frequently isolated in bronchoalveolar lavage fluid of bone marrow transplant recipients without pneumonia. Ann Intern Med. 1990; 112:913-6.
 
Schmidt GM, Horak DA, Niland JC, Duncan SR, Forman SJ, Zaia JA.  A randomized, controlled trial of prophylactic ganciclovir for cytomegalovirus pulmonary infection in recipients of allogeneic bone marrow transplants; The City of Hope-Stanford-Syntex CMV Study Group. N Engl J Med. 1991; 324; 1005-11.
 
Goodrich JM, Mori M, Gleaves CA, Du Mond C, Cays M, Ebeling DF, et al.  Early treatment with ganciclovir to prevent cytomegalovirus disease after allogeneic bone marrow transplantation. N Engl J Med. 1991; 325:1601-7.
 
Ljungman P, Gleaves CA, Meyers JD.  Respiratory virus infections in immunocompromised patients. Bone Marrow Transplant. 1989; 4:35-40.
 
Gleaves CA, Smith TF, Shuster EA, Pearson GR.  Comparison of standard tube and shell vial cell culture techniques for the detection of cytomegalovirus in clinical specimens. J Clin Microbiol. 1985; 21:217-21.
 
Rabalais GP, Stout GG, Ladd KL, Cost KM.  Rapid diagnosis of respiratory viral infections by using a shell vial assay and monoclonal antibody pool. J Clin Microbiol. 1992; 30:1505-8.
 
Crawford SW, Bowden RA, Hackman RC, Gleaves CA, Meyers JD, Clark JG.  Rapid detection of cytomegalovirus pulmonary infection by bronchoalveolar lavage and centrifugation culture. Ann Intern Med. 1988; 108:180-5.
 
Meyers JD, Flournoy N, Thomas ED.  Nonbacterial pneumonia after allogeneic marrow transplantation: a review of ten years' experience. Rev Infect Dis. 1982; 4:1119-32.
 
Emanuel D, Peppard J, Stover D, Gold J, Armstrong D, Hammerling U.  Rapid immunodiagnosis of cytomegalovirus pneumonia by bronchoalveolar lavage using human and murine monoclonal antibodies. Ann Intern Med. 1986; 104:476-81.
 
Paradis IL, Grgurich WF, Dummer JS, Dekker A, Dauber JH.  Rapid detection of cytomegalovirus pneumonia from lung lavage cells. Am Rev Respir Dis. 1988; 138:697-702.
 
Martin WJ 2d, Smith TF. Rapid detection of cytomegalovirus in bronchoalveolar lavage specimens by a monoclonal antibody method. J Clin Microbiol. 1986; 23:1006-8.
 
van der Bij W, Schirm J, Torensma R, van Son WJ, Tegzess AM, The TH.  Comparison between viremia and antigenemia for detection of cytomegalovirus blood. J Clin Microbiol. 1988; 26:2531-5.
 
Boeckh M, Bowden RA, Goodrich JM, Pettinger M, Meyers JD.  Cytomegalovirus antigen detection in peripheral blood leukocytes after allogeneic marrow transplantation. Blood. 1992; 80:1358-64.
 
de Gast GC, Boland GJ, Vlieger AM, de Weger RA, Verdonck LF, Zwaan FE, et al.  Abortive human cytomegalovirus infection in patients after allogeneic bone marrow transplantation. Bone Marrow Transplant. 1992; 9:221-5.
 
Erice A, Holm MA, Gill PC, Henry S, Dirksen CL, Dunn DL, et al.  Cytomegalovirus (CMV) antigenemia assay is more sensitive than shell vial cultures for rapid detection of CMV in polymorphonuclear blood leukocytes. J Clin Microbiol. 1992; 30:2822-5.
 
Mazzulli T, Rubin RH, Ferraro MJ, D'Aquila RT, Doveikis SA, Smith BR, et al.  Cytomegalovirus antigenemia: clinical correlations in transplant recipients and in persons with AIDS. J Clin Microbiol. 1993; 31:2824-7.
 
Bein G, Bitsch A, Hoyer J, Steinhoff J, Fricke L, Machnik H, et al.  A longitudinal prospective study of cytomegalovirus pp65 antigenemia in renal transplant recipients. Transpl Int. 1993; 6:185-90.
 
Halwachs G, Zach R, Pogglitsch H, Holzer H, Tiran A, Iberer F, et al.  A rapid immunocytochemical assay for CMV detection in peripheral blood of organ-transplanted patients in clinical practice. Transplantation. 1993; 56:338-42.
 
van den Berg AP, van Son WJ, Haagsma EB, Klompmaker IJ, Tegzess AM, Schirm J, et al.  Prediction of recurrent cytomegalovirus disease after treatment with ganciclovir in solid-organ transplant recipients. Transplantation. 1993; 55:847-51.
 
Michaels MG, Serdy C, Barbadora K, Green M, Apalsch A, Wald ER.  Respiratory syncytial virus: a comparison of diagnostic modalities. Pediatr Infect Dis J. 1992; 11:613-6.
 
Thomas EE, Book LE.  Comparison of two rapid methods for detection of respiratory syncytial virus (RSV) (TestPack RSV and ortho RSV ELISA) with direct immonufluorescence and virus isolation for the diagnosis of pediatric RSV infection. J Clin Microbiol. 1991; 29:632-5.
 
Hackman RC, Myerson D, Meyers JD, Shulman HM, Sale GE, Goldstein LC, et al.  Rapid diagnosis of cytomegaloviral pneumonia by tissue immunofluorescence with a murine monoclonal antibody. J Infect Dis. 1985; 151:325-9.
 
Schmidt CA, Oettle H, Wilborn F, Jessen J, Timm H, Schwerdtfeger R, et al.  Demonstration of cytomegalovirus after bone marrow transplantation by polymerase chain reaction, virus culture and antigen ejection in buffy coat leukocytes. Bone Marrow Transplant. 1994; 13:71-5.
 
van Dorp WT, Vlieger A, Jiwa NM, van Es LA, van der Ploeg M, van Saase JL, et al.  The polymerase chain reaction, a sensitive and rapid technique for detecting cytomegalovirus infection after renal transplantation. Transplantation. 1992; 54:661-4.
 
The TH, van der Ploeg M, van den Berg AP, Vlieger AM, van der Giessen M, van Son WJ.  Direct detection of cytomegalovirus in peripheral blood leukocytes-a review of the antigenemia assay and polymerase chain reaction. Transplantation. 1992; 54:193-8.
 
Jiwa NM, Van Gemert GW, Raap AK, Van de Rijke FM, Mulder A, Lens PF, et al.  Rapid detection of human cytomegalovirus DNA in peripheral blood leukocytes of viremic transplant recipients by the polymerase chain reaction. Transplantation. 1989; 48:72-6.
 
Schmidt CA, Oettle H, Wilborn F, Jessen J, Timm H, Schwerdtfeger R, et al.  Demonstration of cytomegalovirus after bone marrow transplantation by polymerase chain reaction, virus culture and antigen ejection in buffy coat leukocytes. Bone Marrow Transplant. 1994; 13:71-5.
 
Einsele H, Ehninger G, Steidle M, Fischer I, Bihler S, Gerneth F, et al.  Lymphocytopenia as an unfavorable prognostic factor in patients with cytomegalovirus infection after bone marrow transplantation. Blood. 1993; 82:1672-8.
 
Bitsch A, Kirchner H, Dennin R, Hoyer J, Fricke L, Steinhoff J, et al.  The long persistence of CMV DNA in the blood of renal transplant patients after recovery from CMV infection. Transplantation. 1993; 56:108-13.
 
Einsele H, Steidle M, Vallbracht A, Saal JG, Ehninger G, Mller CA.  Early occurrence of human cytomegalovirus infection after bone marrow transplantation as demonstrated by the polymerase chain reaction technique. Blood. 1991; 77:1104-10.
 
Myerson D, Hackman RC, Nelson JA, Ward DC, McDougall JK.  Widespread presence of histologically occult cytomegalovirus. Hum Pathol. 1984; 15:430-9.
 
Slavin MA, Gleaves CA, Schoch HG, Bowden RA.  Quantification of cytomegalovirus in bronchoalveolar lavage fluid after allogeneic marrow transplantation by centrifugation culture. J Clin Microbiol. 1992; 30:2776-9.
 
Miles PR, Baughman RP, Linnemann CC Jr.  Cytomegalovirus in the bronchoalveolar lavage fluid of patients with AIDS. Chest. 1990; 97:1072-6.
 
Millar AB, Patou G, Miller RF, Grundy JE, Katz DR, Weller IV, et al.  Cytomegalovirus in the lungs of patients with AIDS. Respiratory pathogen or passenger? Am Rev Respir Dis. 1990; 141:1474-7.
 
Balows A, Fraser DW.  International Symposium on Legionnaires' Disease. Ann Intern Med. 1979; 90:489-703.
 
Edelstein PH, Meyer RD. Legionella pneumonias. In: Pennington JE, ed. Respiratory Infections: Diagnosis and Management. 3d ed. New York: Raven; 1994:455-84.
 
England AC 3d, Fraser DW, Plikaytis BD, Tsai TF, Storch G, Broome CV. Sporadic legionellosis in the United States: the first thousand cases. Ann Intern Med. 1981; 94:164-70.
 
Kirby BD, Snyder KM, Meyer RD, Finegold SM.  Legionnaires' disease: report of sixty-five nosocomially acquired cases of review of the literature. Medicine (Baltimore). 1980; 59:188-205.
 
Edelstein PH.  Detection of antibodies to Legionella. In: Rose NR, de Macario EC, Fahey JL, Friedman H, Penn GM, eds. Manual of Clinical Laboratory Immunology. 4th ed. Washington, DC: American Soc Microbiology; 1992; 10:685-9.
 
Helms CM, Massanari RM, Zeitler R, Streed S, Gilchrist MJ, Hall N, et al.  Legionnaires' disease associated with a hospital water system: a cluster of 24 nosocomial cases. Ann Intern Med. 1983; 99:172-8.
 
Fanta CH, Pennington JE.  Pneumonia in the immunocompromised host. In: Pennington JE, ed. Respiratory Infections: Diagnosis and Management. 3rd ed. New York: Raven; 1994:275-94.
 
Edelstein PH, Beer KB, Sturge JC, Watson AJ, Goldstein LC.  Clinical utility of a monoclonal direct fluorescent reagent specific for Legionella pneumophila: comparative study with other reagents. J Clin Microbiol. 1985; 22:419-21.
 
Aguero-Rosenfeld ME, Edelstein PH.  Retrospective evaluation of the Du Pont radioimmunoassay kit for detection of Legionella pneumophila serogroup 1 antigenuria in humans. J Clin Microbiol. 1988; 26:1775-8.
 
Jaulhac B, Nowicki M, Bornstein N, Meunier O, Prevost G, Piemont Y, et al.  Detection of Legionella spp. in bronchoalveolar lavage fluids by DNA amplification. J Clin Microbiol. 1992; 30:920-4.
 
Summersgill J, Gaydos C, Quinn T, Fowler C, Ramirez J.  Detection of Legionella in an immunocompromised population with pneumonia using the polymerase chain reaction [Abstract]. Interscience Conference on Antimicrobial Agents and Chemotherapy. Orlando, Florida; 1994:165.
 
Bej AK, Mahbubani MH, Atlas RM.  Detection of viable Legionella pneumophila in water by polymerase chain reaction and gene probe methods. Appl Environ Microbiol. 1991; 57:597-600.
 
Kenny GE.  Mycoplasmas. In: Balows A, ed. Manual of Clinical Microbiology. 5th ed. Washington, DC: American Soc Microbiology; 1991:478-82.
 
Kenny GE, Kaiser GG, Cooney MK, Foy HM.  Diagnosis of Mycoplasma pneumoniae pneumonia: sensitivities and specificities of serology with lipid antigen and isolation of the organism on soy peptone medium for identification of infections. J Clin Microbiol. 1990; 28:2087-93.
 
Kenny GE.  Serology of mycoplasmal infections. In: Rose NR, Friedman H, Fahey JL, eds. Manual of Clinical Laboratory Immunology. Washington, DC: American Soc Microbiology; 1986:440-5.
 
van Kuppeveld FJ, Johansson KE, Galama JM, Kissing J, Bolske G, Hjelm E, et al.  16s rRNA based polymerase chain reaction compared with culture and serological methods for diagnosis of Mycoplasma pneumoniae infection. Eur J Clin Microbiol Infect Dis. 1994; 13:401-5.
 
Buck GE, O'Hara LC, Summersgill JT.  Rapid, sensitive detection of Mycoplasma pneumoniae in simulated clinical specimens by DNA amplification. J Clin Microbiol. 1992; 125:680-4.
 
Quinn TC. Chlamydia pneumonia. In: Shelhamer J, Pizzo PA, Parrillo JE, Masur H, eds. Respiratory Disease in the Immunosuppressed Host. Philadelphia: JB Lippincott; 1991:298-311.
 
Grayston JT, Campbell LA, Kuo CC, Mordhorst CH, Saikku P, Thom DH, et al.  A new respiratory tract pathogen: Chlamydia pneumoniae strain TWAR. J Infect Dis. 1990; 161:618-25.
 
Gaydos CA, Fowler CL, Gill VJ, Eiden JJ, Quinn TC.  Detection of Chlamydia pneumoniae by polymerase chain reaction-enzyme immunoassay in an immunocompromised population. Clin Infect Dis. 1993; 17:718-23.
 
Roblin PM, Dumornay W, Hammerschlag MR.  Use of HEp-2 cells for improved isolation and passage of Chlamydia pneumoniae. J Clin Microbiol. 1992; 30:168-71.
 
Cles LD, Stamm WE.  Use of HL cells for improved isolation and passage of Chlamydia pneumoniae. J Clin Microbiol. 1990; 28:934-40.
 
Gaydos CA, Quinn TC, Eiden JJ.  Identification of Chlamydia pneumoniae by DNA amplification of the 16s rRNA gene. J Clin Microbiol. 1992; 30:796-800.
 
Mones JM, Saldana MJ, Oldham SA.  Diagnosis of Pneumocystis carinii pneumonia. Roentgenographic-pathologic correlates based on fiberoptic bronchoscopy specimens from patients with the acquired immunodeficiency syndrome. Chest. 1986; 89:522-6.
 
Stover DE, White DA, Romano PA, Gellene RA.  Diagnosis of pulmonary disease in acquired immune deficiency syndrome (AIDS). Role of bronchoscopy and bronchoalveolar lavage. Am Rev Respir Dis. 1984; 130:659-62.
 
Ognibene FP, Shelhamer J, Gill V, Macher AM, Loew D, Parker MM, et al.  The diagnosis of Pneumocystis carinii pneumonia in patients with the acquired immunodeficiency syndrome using subsegmental bronchoalveolar lavage. Am Rev Respir Dis. 1984; 129:929-32.
 
Bigby TD, Margolskee D, Curtis JL, Michael PF, Sheppard D, Hadley WK, et al.  The usefulness of induced sputum in the diagnosis of Pneumocystis carinii pneumonia in patients with the acquired immunodeficiency syndrome. Am Rev Respir Dis. 1986; 133:515-8.
 
Kovacs JA, Ng VL, Masur H, Leoung G, Hadley WK, Evans G, et al.  Diagnosis of Pneumocystis carinii pneumonia: improved detection in sputum with use of monoclonal antibodies. N Engl J Med. 1988; 318:589-93.
 
Smith JW, Bartlett MS.  Laboratory diagnosis of Pneumocystis carinii infection. Clin Lab Med. 1982; 2:393-406.
 
Midgley J, Parsons P, Leigh TR, Collins JV, Shanson DC, Husain OA, et al.  Increased sensitivity of immunofluorescence for detection of Pneumocystis carinii [Letter]. Lancet. 1989; 2:1523.
 
Ng VL, Virani NA, Chaisson RE, Yajko DM, Sphar HT, Cabrian K, et al.  Rapid detection of Pneumocystis carinii using a direct fluorescent monoclonal antibody stain. J Clin Microbiol. 1990; 28:2228-33.
 
Fortun J, Navas E, Marti-Belda P, Montilla P, Hermida JM, Perez-Elias MJ, et al. Pneumocystis carinii pneumonia in HIV-infected patients: diagnostic yield of induced sputum and immunofluorescent stain with monoclonal antibodies. Eur Respir J. 1992; 5:665-9.
 
Wakefield AE, Pixley FJ, Banerji S, Sinclair K, Miller RF, Moxon ER, et al.  Detection of Pneumocystis carinii with DNA amplification. Lancet. 1990; 336:451-3.
 
Lipschik GY, Gill VJ, Lundgren JD, Andrawis VA, Nelson NA, Nielsen JD, et al.  Improved diagnosis of Pneumocystis carinii infection by polymerase chain reaction on induced sputum and blood. Lancet. 1992; 340:203-6.
 
Cartwright CP, Nelson NA, Gill VJ.  Development and evaluation of a rapid and simple procedure for detection of Pneumocystis carinii by PCR. J Clin Microbiol. 1994; 32:1634-8.
 
Levine SJ, Kennedy D, Shelhamer JH, Kovacs A, Feuerstein IM, Gill VJ, et al.  Diagnosis of Pneumocystis carinii pneumonia by multiple lobe, sitedirected bronchoalveolar lavage with immunofluorescent monoclonal antibody staining in human immunodeficiency virus-infected patients receiving aerosolized pentamidine chemoprophylaxis. Am Rev Respir Dis. 1992; 146:838-43.
 
Jules-Elysee KM, Stover DE, Zaman MB, Bernard EM, White DA.  Aerosolized pentamidine: effect on diagnosis and presentation of Pneumocystis carinii pneumonia. Ann Intern Med. 1990; 112:750-7.
 
Levine SJ, Masur H, Gill VJ, Feuerstein I, Suffredini A, Brown D, et al.  Effect of aerosolized pentamidine prophylaxis on the diagnosis of Pneumocystis carinii pneumonia by induced sputum examination in patients infected with the human immunodeficiency virus. Am Rev Respir Dis. 1991; 144:760-4.
 
Yung RC, Weinacker AB, Steiger DJ, Miller TR, Stern EJ, Salmon CJ, et al.  Upper and middle lobe bronchoalveolar lavage to diagnose Pneumocystis carinii pneumonia. Am Rev Respir Dis. 1993; 148:1563-6.
 
Raviglione MC.  Extrapulmonary pneumocystosis: the first 50 cases. Rev Infect Dis. 1990; 12:1127-38.
 
Shami MJ, Freeman W, Friedberg D, Siderides E, Listhaus A, Ai E.  A multicenter study of Pneumocystis choroidopathy. Am J Ophthalmol. 1991; 112:15-22.
 
Wolinsky E.  Conventional diagnostic methods for tuberculosis. Clin Infect Dis. 1994; 19:396-401.
 
Woods GL, Witebsky FG.  Current status of mycobacterial testing in clinical laboratories. Results of a questionnaire completed by participants in the College of American Pathologists Mycobacteriology E survey. Arch Pathol Lab Med. 1993; 117:876-84.
 
Fishman JA, Roth RS, Zanzot E, Enos EJ, Ferraro MJ.  Use of induced sputum specimens for microbiologic diagnosis of infections due to organisms other than Pneumocystis carinii. J Clin Microbiol. 1994; 32:131-4.
 
Klotz SA, Penn RL.  Acid-fast staining of urine and gastric contents is an excellent indicator of mycobacterial disease. Am Rev Respir Dis. 1987; 136:1197-8.
 
Schluger NW, Kinney D, Harkin TJ, Rom WN.  Clinical utility of the polymerase chain reaction in the diagnosis of infections due to Mycobacterium tuberculosis. Chest. 1994; 105:4:1116-21.
 
Abe C, Hirano K, Wada M, Kazumi Y, Takahashi M, Fukasawa Y, et al.  Detection of Mycobacterium tuberculosis in clinical specimens by polymerase chain reaction and Gen-Probe Amplified Mycobacterium Tuberculosis Direct Test. J Clin Microbiol. 1993; 31:3270-4.
 
Noordhoek GT, Kolk AH, Bjune G, Catty D, Dale JW, Fine PE, et al.  Sensitivity and specificity of PCR for detection of Mycobacterium tuberculosis: a blind comparison study among seven laboratories. J Clin Microbiol. 1994; 32:277-84.
 
Jones BE, Young SM, Antoniskis D, Davidson PT, Kramer F, Barnes PF.  Relationship of the manifestations of tuberculosis to CD4 cell counts in patients with human immunodeficiency virus infection. Am Rev Respir Dis. 1993; 148:1292-7.
 
Yu VL, Muder RR, Poorsattar A.  Significance of isolation of Aspergillus from the respiratory tract in diagnosis of invasive pulmonary aspergillosis. Results from a three-year prospective study. Am J Med. 1986; 81:249-54.
 
Weinberger M, Elattar I, Marshall D, Steinberg SM, Redner RL, Young NS, et al.  Patterns of infection in patients with aplastic anemia and the emergence of Aspergillus as a major cause of death. Medicine (Baltimore). 1992; 71:24-43.
 
Ruchel R.  Diagnosis of invasive mycoses in severely immunosuppresed patients. Ann Hematol. 1993; 67:1-11.
 
Bennett JE.  Rapid diagnosis of candidiasis and aspergillosis. Rev Infect Dis. 1987; 9:398-402.
 
Richardson SE, Bannatyne RM, Summerbell RC, Milliken J, Gold R, Weitzman SS.  Disseminated fusarial infection in the immunocompromised host. Rev Infect Dis. 1988; 10:1171-81.
 
Chmel H.  Fungal infections in the immunocompromised host. Clinical syndromes and diagnosis. In: Murphy JW, Friedman H, Bendinelli M, eds. Fungal Infections and Immune Responses. New York: Plenum; 1993:405-24.
 
Wheat LJ.  The role of the serologic diagnostic laboratory and the diagnosis of fungal disease. In: Sarosi GA, Davies SF, eds. Fungal Diseases of the Lung. 2d ed. New York: Raven; 1993:29-38.
 
Wheat LJ, Kohler RB, Tewari RP.  Diagnosis of disseminated histoplasmosis by detection of Histoplasma capsulatum antigen in serum and urine specimens. N Engl J Med. 1986; 314:83-8.
 
Anaissie E.  Opportunistic mycoses in the immunocompromised host: experience at a cancer center and review. Clin Infect Dis. 1992; 14(Suppl 1):S43-53.
 

Figures

Grahic Jump Location
Figure 1.
Schematic of the shell vial virus isolation technique.
Grahic Jump Location
Grahic Jump Location
Figure 2.
Simplified schematic of the Accuprobe hybridization assay (Gen-Probe).

*The selection reagent selectively splits the acridinium ester from the single-stranded DNA probe but not from the double-stranded hybrid.

Grahic Jump Location
Grahic Jump Location
Figure 3.
Schematic of the polymerase chain reaction.
Grahic Jump Location

Tables

Table Jump PlaceholderTable 1.  Specimens for Optimal Diagnosis of Pulmonary Infections*
Table Jump PlaceholderTable 2.  Direct Stains*, Direct Tests, and Culture Available for Common Pulmonary Pathogens †
Table Jump PlaceholderTable 3.  Significance of Respiratory Virus Detection in the Presence of Pneumonia in Patients Receiving Bone Marrow and Solid Organ Transplant*
Table Jump PlaceholderTable 4.  Laboratory Diagnosis of Legionella Pneumonia*
Table Jump PlaceholderTable 5.  Diagnostic Tests for Mycoplasma pneumoniae
Table Jump PlaceholderTable 6.  Diagnostic Tests for Chlamydia pneumoniae
Table Jump PlaceholderTable 7.  Utility of Diagnostic Tests for Fungal Pneumonia*

References

Thorpe JE, Baughman RP, Frame PT, Wesseler TA, Staneck JL.  Bronchoalveolar lavage for diagnosing acute bacterial pneumonia. J Infect Dis. 1987; 155:855-61.
 
Xaubet A, Torres A, Marco F, Puig-De la Bellacasa J, Faus R, Agustividal A.  Pulmonary infiltrates in immunocompromised patients. Diagnostic value of telescoping plugged catheter and bronchoalveolar lavage. Chest. 1989; 95:130-5.
 
Ruutu P, Ruutu T, Volin L, Tukiainen P, Ukkonen P, Hovi T.  Cytomegalovirus is frequently isolated in bronchoalveolar lavage fluid of bone marrow transplant recipients without pneumonia. Ann Intern Med. 1990; 112:913-6.
 
Schmidt GM, Horak DA, Niland JC, Duncan SR, Forman SJ, Zaia JA.  A randomized, controlled trial of prophylactic ganciclovir for cytomegalovirus pulmonary infection in recipients of allogeneic bone marrow transplants; The City of Hope-Stanford-Syntex CMV Study Group. N Engl J Med. 1991; 324; 1005-11.
 
Goodrich JM, Mori M, Gleaves CA, Du Mond C, Cays M, Ebeling DF, et al.  Early treatment with ganciclovir to prevent cytomegalovirus disease after allogeneic bone marrow transplantation. N Engl J Med. 1991; 325:1601-7.
 
Ljungman P, Gleaves CA, Meyers JD.  Respiratory virus infections in immunocompromised patients. Bone Marrow Transplant. 1989; 4:35-40.
 
Gleaves CA, Smith TF, Shuster EA, Pearson GR.  Comparison of standard tube and shell vial cell culture techniques for the detection of cytomegalovirus in clinical specimens. J Clin Microbiol. 1985; 21:217-21.
 
Rabalais GP, Stout GG, Ladd KL, Cost KM.  Rapid diagnosis of respiratory viral infections by using a shell vial assay and monoclonal antibody pool. J Clin Microbiol. 1992; 30:1505-8.
 
Crawford SW, Bowden RA, Hackman RC, Gleaves CA, Meyers JD, Clark JG.  Rapid detection of cytomegalovirus pulmonary infection by bronchoalveolar lavage and centrifugation culture. Ann Intern Med. 1988; 108:180-5.
 
Meyers JD, Flournoy N, Thomas ED.  Nonbacterial pneumonia after allogeneic marrow transplantation: a review of ten years' experience. Rev Infect Dis. 1982; 4:1119-32.
 
Emanuel D, Peppard J, Stover D, Gold J, Armstrong D, Hammerling U.  Rapid immunodiagnosis of cytomegalovirus pneumonia by bronchoalveolar lavage using human and murine monoclonal antibodies. Ann Intern Med. 1986; 104:476-81.
 
Paradis IL, Grgurich WF, Dummer JS, Dekker A, Dauber JH.  Rapid detection of cytomegalovirus pneumonia from lung lavage cells. Am Rev Respir Dis. 1988; 138:697-702.
 
Martin WJ 2d, Smith TF. Rapid detection of cytomegalovirus in bronchoalveolar lavage specimens by a monoclonal antibody method. J Clin Microbiol. 1986; 23:1006-8.
 
van der Bij W, Schirm J, Torensma R, van Son WJ, Tegzess AM, The TH.  Comparison between viremia and antigenemia for detection of cytomegalovirus blood. J Clin Microbiol. 1988; 26:2531-5.
 
Boeckh M, Bowden RA, Goodrich JM, Pettinger M, Meyers JD.  Cytomegalovirus antigen detection in peripheral blood leukocytes after allogeneic marrow transplantation. Blood. 1992; 80:1358-64.
 
de Gast GC, Boland GJ, Vlieger AM, de Weger RA, Verdonck LF, Zwaan FE, et al.  Abortive human cytomegalovirus infection in patients after allogeneic bone marrow transplantation. Bone Marrow Transplant. 1992; 9:221-5.
 
Erice A, Holm MA, Gill PC, Henry S, Dirksen CL, Dunn DL, et al.  Cytomegalovirus (CMV) antigenemia assay is more sensitive than shell vial cultures for rapid detection of CMV in polymorphonuclear blood leukocytes. J Clin Microbiol. 1992; 30:2822-5.
 
Mazzulli T, Rubin RH, Ferraro MJ, D'Aquila RT, Doveikis SA, Smith BR, et al.  Cytomegalovirus antigenemia: clinical correlations in transplant recipients and in persons with AIDS. J Clin Microbiol. 1993; 31:2824-7.
 
Bein G, Bitsch A, Hoyer J, Steinhoff J, Fricke L, Machnik H, et al.  A longitudinal prospective study of cytomegalovirus pp65 antigenemia in renal transplant recipients. Transpl Int. 1993; 6:185-90.
 
Halwachs G, Zach R, Pogglitsch H, Holzer H, Tiran A, Iberer F, et al.  A rapid immunocytochemical assay for CMV detection in peripheral blood of organ-transplanted patients in clinical practice. Transplantation. 1993; 56:338-42.
 
van den Berg AP, van Son WJ, Haagsma EB, Klompmaker IJ, Tegzess AM, Schirm J, et al.  Prediction of recurrent cytomegalovirus disease after treatment with ganciclovir in solid-organ transplant recipients. Transplantation. 1993; 55:847-51.
 
Michaels MG, Serdy C, Barbadora K, Green M, Apalsch A, Wald ER.  Respiratory syncytial virus: a comparison of diagnostic modalities. Pediatr Infect Dis J. 1992; 11:613-6.
 
Thomas EE, Book LE.  Comparison of two rapid methods for detection of respiratory syncytial virus (RSV) (TestPack RSV and ortho RSV ELISA) with direct immonufluorescence and virus isolation for the diagnosis of pediatric RSV infection. J Clin Microbiol. 1991; 29:632-5.
 
Hackman RC, Myerson D, Meyers JD, Shulman HM, Sale GE, Goldstein LC, et al.  Rapid diagnosis of cytomegaloviral pneumonia by tissue immunofluorescence with a murine monoclonal antibody. J Infect Dis. 1985; 151:325-9.
 
Schmidt CA, Oettle H, Wilborn F, Jessen J, Timm H, Schwerdtfeger R, et al.  Demonstration of cytomegalovirus after bone marrow transplantation by polymerase chain reaction, virus culture and antigen ejection in buffy coat leukocytes. Bone Marrow Transplant. 1994; 13:71-5.
 
van Dorp WT, Vlieger A, Jiwa NM, van Es LA, van der Ploeg M, van Saase JL, et al.  The polymerase chain reaction, a sensitive and rapid technique for detecting cytomegalovirus infection after renal transplantation. Transplantation. 1992; 54:661-4.
 
The TH, van der Ploeg M, van den Berg AP, Vlieger AM, van der Giessen M, van Son WJ.  Direct detection of cytomegalovirus in peripheral blood leukocytes-a review of the antigenemia assay and polymerase chain reaction. Transplantation. 1992; 54:193-8.
 
Jiwa NM, Van Gemert GW, Raap AK, Van de Rijke FM, Mulder A, Lens PF, et al.  Rapid detection of human cytomegalovirus DNA in peripheral blood leukocytes of viremic transplant recipients by the polymerase chain reaction. Transplantation. 1989; 48:72-6.
 
Schmidt CA, Oettle H, Wilborn F, Jessen J, Timm H, Schwerdtfeger R, et al.  Demonstration of cytomegalovirus after bone marrow transplantation by polymerase chain reaction, virus culture and antigen ejection in buffy coat leukocytes. Bone Marrow Transplant. 1994; 13:71-5.
 
Einsele H, Ehninger G, Steidle M, Fischer I, Bihler S, Gerneth F, et al.  Lymphocytopenia as an unfavorable prognostic factor in patients with cytomegalovirus infection after bone marrow transplantation. Blood. 1993; 82:1672-8.
 
Bitsch A, Kirchner H, Dennin R, Hoyer J, Fricke L, Steinhoff J, et al.  The long persistence of CMV DNA in the blood of renal transplant patients after recovery from CMV infection. Transplantation. 1993; 56:108-13.
 
Einsele H, Steidle M, Vallbracht A, Saal JG, Ehninger G, Mller CA.  Early occurrence of human cytomegalovirus infection after bone marrow transplantation as demonstrated by the polymerase chain reaction technique. Blood. 1991; 77:1104-10.
 
Myerson D, Hackman RC, Nelson JA, Ward DC, McDougall JK.  Widespread presence of histologically occult cytomegalovirus. Hum Pathol. 1984; 15:430-9.
 
Slavin MA, Gleaves CA, Schoch HG, Bowden RA.  Quantification of cytomegalovirus in bronchoalveolar lavage fluid after allogeneic marrow transplantation by centrifugation culture. J Clin Microbiol. 1992; 30:2776-9.
 
Miles PR, Baughman RP, Linnemann CC Jr.  Cytomegalovirus in the bronchoalveolar lavage fluid of patients with AIDS. Chest. 1990; 97:1072-6.
 
Millar AB, Patou G, Miller RF, Grundy JE, Katz DR, Weller IV, et al.  Cytomegalovirus in the lungs of patients with AIDS. Respiratory pathogen or passenger? Am Rev Respir Dis. 1990; 141:1474-7.
 
Balows A, Fraser DW.  International Symposium on Legionnaires' Disease. Ann Intern Med. 1979; 90:489-703.
 
Edelstein PH, Meyer RD. Legionella pneumonias. In: Pennington JE, ed. Respiratory Infections: Diagnosis and Management. 3d ed. New York: Raven; 1994:455-84.
 
England AC 3d, Fraser DW, Plikaytis BD, Tsai TF, Storch G, Broome CV. Sporadic legionellosis in the United States: the first thousand cases. Ann Intern Med. 1981; 94:164-70.
 
Kirby BD, Snyder KM, Meyer RD, Finegold SM.  Legionnaires' disease: report of sixty-five nosocomially acquired cases of review of the literature. Medicine (Baltimore). 1980; 59:188-205.
 
Edelstein PH.  Detection of antibodies to Legionella. In: Rose NR, de Macario EC, Fahey JL, Friedman H, Penn GM, eds. Manual of Clinical Laboratory Immunology. 4th ed. Washington, DC: American Soc Microbiology; 1992; 10:685-9.
 
Helms CM, Massanari RM, Zeitler R, Streed S, Gilchrist MJ, Hall N, et al.  Legionnaires' disease associated with a hospital water system: a cluster of 24 nosocomial cases. Ann Intern Med. 1983; 99:172-8.
 
Fanta CH, Pennington JE.  Pneumonia in the immunocompromised host. In: Pennington JE, ed. Respiratory Infections: Diagnosis and Management. 3rd ed. New York: Raven; 1994:275-94.
 
Edelstein PH, Beer KB, Sturge JC, Watson AJ, Goldstein LC.  Clinical utility of a monoclonal direct fluorescent reagent specific for Legionella pneumophila: comparative study with other reagents. J Clin Microbiol. 1985; 22:419-21.
 
Aguero-Rosenfeld ME, Edelstein PH.  Retrospective evaluation of the Du Pont radioimmunoassay kit for detection of Legionella pneumophila serogroup 1 antigenuria in humans. J Clin Microbiol. 1988; 26:1775-8.
 
Jaulhac B, Nowicki M, Bornstein N, Meunier O, Prevost G, Piemont Y, et al.  Detection of Legionella spp. in bronchoalveolar lavage fluids by DNA amplification. J Clin Microbiol. 1992; 30:920-4.
 
Summersgill J, Gaydos C, Quinn T, Fowler C, Ramirez J.  Detection of Legionella in an immunocompromised population with pneumonia using the polymerase chain reaction [Abstract]. Interscience Conference on Antimicrobial Agents and Chemotherapy. Orlando, Florida; 1994:165.
 
Bej AK, Mahbubani MH, Atlas RM.  Detection of viable Legionella pneumophila in water by polymerase chain reaction and gene probe methods. Appl Environ Microbiol. 1991; 57:597-600.
 
Kenny GE.  Mycoplasmas. In: Balows A, ed. Manual of Clinical Microbiology. 5th ed. Washington, DC: American Soc Microbiology; 1991:478-82.
 
Kenny GE, Kaiser GG, Cooney MK, Foy HM.  Diagnosis of Mycoplasma pneumoniae pneumonia: sensitivities and specificities of serology with lipid antigen and isolation of the organism on soy peptone medium for identification of infections. J Clin Microbiol. 1990; 28:2087-93.
 
Kenny GE.  Serology of mycoplasmal infections. In: Rose NR, Friedman H, Fahey JL, eds. Manual of Clinical Laboratory Immunology. Washington, DC: American Soc Microbiology; 1986:440-5.
 
van Kuppeveld FJ, Johansson KE, Galama JM, Kissing J, Bolske G, Hjelm E, et al.  16s rRNA based polymerase chain reaction compared with culture and serological methods for diagnosis of Mycoplasma pneumoniae infection. Eur J Clin Microbiol Infect Dis. 1994; 13:401-5.
 
Buck GE, O'Hara LC, Summersgill JT.  Rapid, sensitive detection of Mycoplasma pneumoniae in simulated clinical specimens by DNA amplification. J Clin Microbiol. 1992; 125:680-4.
 
Quinn TC. Chlamydia pneumonia. In: Shelhamer J, Pizzo PA, Parrillo JE, Masur H, eds. Respiratory Disease in the Immunosuppressed Host. Philadelphia: JB Lippincott; 1991:298-311.
 
Grayston JT, Campbell LA, Kuo CC, Mordhorst CH, Saikku P, Thom DH, et al.  A new respiratory tract pathogen: Chlamydia pneumoniae strain TWAR. J Infect Dis. 1990; 161:618-25.
 
Gaydos CA, Fowler CL, Gill VJ, Eiden JJ, Quinn TC.  Detection of Chlamydia pneumoniae by polymerase chain reaction-enzyme immunoassay in an immunocompromised population. Clin Infect Dis. 1993; 17:718-23.
 
Roblin PM, Dumornay W, Hammerschlag MR.  Use of HEp-2 cells for improved isolation and passage of Chlamydia pneumoniae. J Clin Microbiol. 1992; 30:168-71.
 
Cles LD, Stamm WE.  Use of HL cells for improved isolation and passage of Chlamydia pneumoniae. J Clin Microbiol. 1990; 28:934-40.
 
Gaydos CA, Quinn TC, Eiden JJ.  Identification of Chlamydia pneumoniae by DNA amplification of the 16s rRNA gene. J Clin Microbiol. 1992; 30:796-800.
 
Mones JM, Saldana MJ, Oldham SA.  Diagnosis of Pneumocystis carinii pneumonia. Roentgenographic-pathologic correlates based on fiberoptic bronchoscopy specimens from patients with the acquired immunodeficiency syndrome. Chest. 1986; 89:522-6.
 
Stover DE, White DA, Romano PA, Gellene RA.  Diagnosis of pulmonary disease in acquired immune deficiency syndrome (AIDS). Role of bronchoscopy and bronchoalveolar lavage. Am Rev Respir Dis. 1984; 130:659-62.
 
Ognibene FP, Shelhamer J, Gill V, Macher AM, Loew D, Parker MM, et al.  The diagnosis of Pneumocystis carinii pneumonia in patients with the acquired immunodeficiency syndrome using subsegmental bronchoalveolar lavage. Am Rev Respir Dis. 1984; 129:929-32.
 
Bigby TD, Margolskee D, Curtis JL, Michael PF, Sheppard D, Hadley WK, et al.  The usefulness of induced sputum in the diagnosis of Pneumocystis carinii pneumonia in patients with the acquired immunodeficiency syndrome. Am Rev Respir Dis. 1986; 133:515-8.
 
Kovacs JA, Ng VL, Masur H, Leoung G, Hadley WK, Evans G, et al.  Diagnosis of Pneumocystis carinii pneumonia: improved detection in sputum with use of monoclonal antibodies. N Engl J Med. 1988; 318:589-93.
 
Smith JW, Bartlett MS.  Laboratory diagnosis of Pneumocystis carinii infection. Clin Lab Med. 1982; 2:393-406.
 
Midgley J, Parsons P, Leigh TR, Collins JV, Shanson DC, Husain OA, et al.  Increased sensitivity of immunofluorescence for detection of Pneumocystis carinii [Letter]. Lancet. 1989; 2:1523.
 
Ng VL, Virani NA, Chaisson RE, Yajko DM, Sphar HT, Cabrian K, et al.  Rapid detection of Pneumocystis carinii using a direct fluorescent monoclonal antibody stain. J Clin Microbiol. 1990; 28:2228-33.
 
Fortun J, Navas E, Marti-Belda P, Montilla P, Hermida JM, Perez-Elias MJ, et al. Pneumocystis carinii pneumonia in HIV-infected patients: diagnostic yield of induced sputum and immunofluorescent stain with monoclonal antibodies. Eur Respir J. 1992; 5:665-9.
 
Wakefield AE, Pixley FJ, Banerji S, Sinclair K, Miller RF, Moxon ER, et al.  Detection of Pneumocystis carinii with DNA amplification. Lancet. 1990; 336:451-3.
 
Lipschik GY, Gill VJ, Lundgren JD, Andrawis VA, Nelson NA, Nielsen JD, et al.  Improved diagnosis of Pneumocystis carinii infection by polymerase chain reaction on induced sputum and blood. Lancet. 1992; 340:203-6.
 
Cartwright CP, Nelson NA, Gill VJ.  Development and evaluation of a rapid and simple procedure for detection of Pneumocystis carinii by PCR. J Clin Microbiol. 1994; 32:1634-8.
 
Levine SJ, Kennedy D, Shelhamer JH, Kovacs A, Feuerstein IM, Gill VJ, et al.  Diagnosis of Pneumocystis carinii pneumonia by multiple lobe, sitedirected bronchoalveolar lavage with immunofluorescent monoclonal antibody staining in human immunodeficiency virus-infected patients receiving aerosolized pentamidine chemoprophylaxis. Am Rev Respir Dis. 1992; 146:838-43.
 
Jules-Elysee KM, Stover DE, Zaman MB, Bernard EM, White DA.  Aerosolized pentamidine: effect on diagnosis and presentation of Pneumocystis carinii pneumonia. Ann Intern Med. 1990; 112:750-7.
 
Levine SJ, Masur H, Gill VJ, Feuerstein I, Suffredini A, Brown D, et al.  Effect of aerosolized pentamidine prophylaxis on the diagnosis of Pneumocystis carinii pneumonia by induced sputum examination in patients infected with the human immunodeficiency virus. Am Rev Respir Dis. 1991; 144:760-4.
 
Yung RC, Weinacker AB, Steiger DJ, Miller TR, Stern EJ, Salmon CJ, et al.  Upper and middle lobe bronchoalveolar lavage to diagnose Pneumocystis carinii pneumonia. Am Rev Respir Dis. 1993; 148:1563-6.
 
Raviglione MC.  Extrapulmonary pneumocystosis: the first 50 cases. Rev Infect Dis. 1990; 12:1127-38.
 
Shami MJ, Freeman W, Friedberg D, Siderides E, Listhaus A, Ai E.  A multicenter study of Pneumocystis choroidopathy. Am J Ophthalmol. 1991; 112:15-22.
 
Wolinsky E.  Conventional diagnostic methods for tuberculosis. Clin Infect Dis. 1994; 19:396-401.
 
Woods GL, Witebsky FG.  Current status of mycobacterial testing in clinical laboratories. Results of a questionnaire completed by participants in the College of American Pathologists Mycobacteriology E survey. Arch Pathol Lab Med. 1993; 117:876-84.
 
Fishman JA, Roth RS, Zanzot E, Enos EJ, Ferraro MJ.  Use of induced sputum specimens for microbiologic diagnosis of infections due to organisms other than Pneumocystis carinii. J Clin Microbiol. 1994; 32:131-4.
 
Klotz SA, Penn RL.  Acid-fast staining of urine and gastric contents is an excellent indicator of mycobacterial disease. Am Rev Respir Dis. 1987; 136:1197-8.
 
Schluger NW, Kinney D, Harkin TJ, Rom WN.  Clinical utility of the polymerase chain reaction in the diagnosis of infections due to Mycobacterium tuberculosis. Chest. 1994; 105:4:1116-21.
 
Abe C, Hirano K, Wada M, Kazumi Y, Takahashi M, Fukasawa Y, et al.  Detection of Mycobacterium tuberculosis in clinical specimens by polymerase chain reaction and Gen-Probe Amplified Mycobacterium Tuberculosis Direct Test. J Clin Microbiol. 1993; 31:3270-4.
 
Noordhoek GT, Kolk AH, Bjune G, Catty D, Dale JW, Fine PE, et al.  Sensitivity and specificity of PCR for detection of Mycobacterium tuberculosis: a blind comparison study among seven laboratories. J Clin Microbiol. 1994; 32:277-84.
 
Jones BE, Young SM, Antoniskis D, Davidson PT, Kramer F, Barnes PF.  Relationship of the manifestations of tuberculosis to CD4 cell counts in patients with human immunodeficiency virus infection. Am Rev Respir Dis. 1993; 148:1292-7.
 
Yu VL, Muder RR, Poorsattar A.  Significance of isolation of Aspergillus from the respiratory tract in diagnosis of invasive pulmonary aspergillosis. Results from a three-year prospective study. Am J Med. 1986; 81:249-54.
 
Weinberger M, Elattar I, Marshall D, Steinberg SM, Redner RL, Young NS, et al.  Patterns of infection in patients with aplastic anemia and the emergence of Aspergillus as a major cause of death. Medicine (Baltimore). 1992; 71:24-43.
 
Ruchel R.  Diagnosis of invasive mycoses in severely immunosuppresed patients. Ann Hematol. 1993; 67:1-11.
 
Bennett JE.  Rapid diagnosis of candidiasis and aspergillosis. Rev Infect Dis. 1987; 9:398-402.
 
Richardson SE, Bannatyne RM, Summerbell RC, Milliken J, Gold R, Weitzman SS.  Disseminated fusarial infection in the immunocompromised host. Rev Infect Dis. 1988; 10:1171-81.
 
Chmel H.  Fungal infections in the immunocompromised host. Clinical syndromes and diagnosis. In: Murphy JW, Friedman H, Bendinelli M, eds. Fungal Infections and Immune Responses. New York: Plenum; 1993:405-24.
 
Wheat LJ.  The role of the serologic diagnostic laboratory and the diagnosis of fungal disease. In: Sarosi GA, Davies SF, eds. Fungal Diseases of the Lung. 2d ed. New York: Raven; 1993:29-38.
 
Wheat LJ, Kohler RB, Tewari RP.  Diagnosis of disseminated histoplasmosis by detection of Histoplasma capsulatum antigen in serum and urine specimens. N Engl J Med. 1986; 314:83-8.
 
Anaissie E.  Opportunistic mycoses in the immunocompromised host: experience at a cancer center and review. Clin Infect Dis. 1992; 14(Suppl 1):S43-53.
 

Letters

NOTE:
Citing articles are presented as examples only. In non-demo SCM6 implementation, integration with CrossRef’s "Cited By" API will populate this tab (http://www.crossref.org/citedby.html).

Comments

Submit a Comment
Submit a Comment

Summary for Patients

Clinical Slide Sets

Terms of Use

The In the Clinic® slide sets are owned and copyrighted by the American College of Physicians (ACP). All text, graphics, trademarks, and other intellectual property incorporated into the slide sets remain the sole and exclusive property of the ACP. The slide sets may be used only by the person who downloads or purchases them and only for the purpose of presenting them during not-for-profit educational activities. Users may incorporate the entire slide set or selected individual slides into their own teaching presentations but may not alter the content of the slides in any way or remove the ACP copyright notice. Users may make print copies for use as hand-outs for the audience the user is personally addressing but may not otherwise reproduce or distribute the slides by any means or media, including but not limited to sending them as e-mail attachments, posting them on Internet or Intranet sites, publishing them in meeting proceedings, or making them available for sale or distribution in any unauthorized form, without the express written permission of the ACP. Unauthorized use of the In the Clinic slide sets will constitute copyright infringement.

Toolkit

Want to Subscribe?

Learn more about subscription options

Advertisement
Forgot your password?
Enter your username and email address. We'll send you a reminder to the email address on record.
(Required)
(Required)