Standard precautions apply. Ensure that material from the site of infection is collected, and contamination by normal flora is avoided.
Decontaminate skin or mucous membranes that must be crossed to obtain the specimen.
Use appropriate sterile supplies to collect the specimen. Place the specimen in a sterile, leakproof container for transport. Ensure that the lid is firmly tightened, but avoid overtightening. Use specific transport medium and/or procedures as required for suspected pathogens (described below) or if transport to the lab will be prolonged (>2 hours). Apply a label to the specimen with information to identify the patient and type of specimen.
Transport the specimen to the lab as quickly as possible, avoiding extremes of temperature.
Note that collection protocols for some types of specimens require specific training and/or certification of the health care professional performing the collection. Examples include collection of bone marrow and cerebrospinal fluid (CSF) specimens.
Smears of patient specimens are stained and examined for the presence of mycobacteria. They may provide early evidence of TB or other mycobacterial diseases. Certain dyes bind to the thick, mycolic acid-rich cell walls of mycobacteria. The lipids of the cell wall make the cells resistant to decolorization with strong acid-alcohol solutions. AFB staining should be undertaken for most specimens that are submitted for mycobacterial culture.
▼ There are two types of AFB stains: chromogenic (carbol-fuchsin stains [hot, Ziehl-Neelsen; cold, Kinyoun]) and fluorogenic (auramine O + rhodamine). After staining, the smear is destained with an acid-alcohol solution, typically HCl in ethanol. Mycobacteria retain the stain.
In chromogenic methods, slides are examined using a 100× oil immersion objective with light microscopy. Nonmycobacterial cells are counterstained with methylene blue. Mycobacteria appear red, whereas other bacteria are stained blue.
In fluorogenic methods, auramine-stained slides are examined by fluorescence microscopy using a 25× or 40× objective. Mycobacteria are yellow-orange against a dark background. The improved signal-to-noise ratio of auramine O fluorochrome staining, allowing scanning with lower power objective, results in examination of a greater area of the slide in a given time and therefore greater sensitivity. Any detected organisms should be confirmed by examination for typical morphology using the 100× objective. Some laboratories confirm positive fluorochrome smears with a carbol-fuchsin-based stain.
Specimens should be collected and transported to the laboratory according to recommendations for mycobacterial cultures.
Turnaround time: <24 hours.
Negative results: Negative. Detection of mycobacteria requires 10,000 or more organisms per milliliter or gram of sample for consistent detection. Sensitivity may be improved by concentration of specimen, such as by centrifugation, and by examination of multiple specimens. Rapidly growing
mycobacteria, such as Mycobacterium fortuitum, have relatively thin layers of cell wall mycolic acid and may decolorize with acid-alcohol decolorizing solutions. These organisms may be stained using a modified acid-fast stain.
Positive results: Positive specimens are very likely (>90%) to yield growth of mycobacteria in culture. In a minority of patients, usually with cavitary or extensive tuberculosis, sputum AFB stains may remain persistently positive for weeks after patients have converted to negative cultures. Nonviable organisms may be detected by AFB stains.
Standardized protocols, such as those published by the American Thoracic Society, should be followed carefully to ensure sensitive detection and accurate interpretation of smears.
Common pitfalls: Care must be taken to avoid contaminating slides with acid-fast organisms. Common causes of slide contamination are use of tap water for solution preparation, carryover between slides with immersion oil, and use of common staining chambers.
This stain may be used for detection of Nocardia in patient specimens or culture isolates when nocardiosis is suspected on the basis of clinical presentation or because of typical morphology in culture isolates. Rapidly growing mycobacteria, such as M. fortuitum, may be negative by routine acid-fast staining, but positive by modified acid-fast staining.
The modified acid-fast stain is also used to confirm nocardioform organisms detected by Gram stain. The modified acid-fast stain is useful for differentiating Nocardia (positive) from Streptomyces (negative), especially in culture isolates.
The modified acid-fast stain is similar to the carbol-fuchsin-based acidfast stains (Ziehl-Neelsen or Kinyoun stains) except that a less active decolorizer is used (1% H2SO4 or 3% HCl in aqueous solution).
Specimens should be collected and transported as appropriate for routine bacterial cultures for the specimen type.
Turnaround time: 24-72 hours.
Expected results: Negative. Negative stains do not rule out nocardiosis.
Positive results: The morphology of positive staining organisms should be correlated with clinical signs and symptoms, and differential diagnosis. Delicate, branching filaments that retain the carbol-fuchsin stain are consistent with nocardiosis.
Nocardia may stain poorly in direct specimen staining. Other species of aerobic actinomycetes, such as Rhodococcus equi and occasionally coryneform bacteria, may be modified acid-fast stain positive.
Real-time PCR-based qualitative test for the detection of human adenovirus (HAdV) DNA using nasopharyngeal swabs (NPSs) specimens. There are seven species of HAdV, A to G. The molecular tests detect but do not differentiate HAdV species A, B, C, D, E, and F. Respiratory infections are caused mainly by species B and C. Adenovirus is also a target tested on the respiratory viral pathogen panel.
For testing individuals exhibiting signs and symptoms of acute respiratory infection
Positive results: Indicates the presence of adenovirus nucleic acid
Negative results: Does not rule out the presence of adenoviruses because viruses may be present at levels below the detection limits of this assay
The results of the test should be used as an aid in diagnosis and should not be used as the sole basis for treatment or other patient management decisions.
Adenovirus DNA may be detected in asymptomatic individuals in certain settings.
This assay tests only for presence of adenovirus; additional testing for other respiratory viruses or other pathogens is needed in case of negative results.
Aerobic cultures are indicated for the detection of common aerobic bacterial pathogens in patient specimens taken from sites with signs and symptoms of bacterial infection (e.g., swelling, redness, heat, pus, or exudate). The objective of culture is to identify potential pathogens and to provide isolates for antibiotic susceptibility or other relevant testing.
Specimens may be inoculated on several types of aerobic culture plates and broth media and may include selective and enriched media. Typical media for aerobic cultures include the following:
▼ Supportive media to isolate nonfastidious organisms, like sheep blood agar (SBA).
▼ Enriched media to isolate organisms with special nutritional requirements, like chocolate agar.
▼ Selective media to suppress the growth of specific types of bacteria in specimens likely to be contaminated with normal flora, like feces. Selective media are often formulated so that colonies of different types of organisms have different appearances. MacConkey agar is an
example. Only non-fastidious gram-negative bacilli are able to grow (Selective). Lactose fermenters are distinguished from lactose non-fermenters (Differential).
▼ Solid versus broth media:
Solid media (culture plates) are inoculated with a small amount of specimen. Mixed cultures are recognized by differences in colony morphology. The amount of each type of organism (and relative proportions in mixed cultures) can be estimated (e.g., rare, light, moderate, or heavy). Pyogenic infections are usually associated with growth of a single (or predominant) pathogen in moderate or heavy amounts.
Broth media can be inoculated with a larger volume of specimen than agar plates, which may improve detection of infections with low concentrations of pathogens, like bacteremia, but the amount of bacteria in the specimen cannot be estimated from broth cultures. Broth cultures have been associated with an increased rate of contamination.
Site-specific bacterial cultures (e.g., sputum culture, genital culture) are recommended, if available.
Some fastidious bacterial pathogens, like Legionella and Francisella, have specific nutritional or incubation requirements for efficient isolation. Specific cultures targeting these organisms should be requested when clinically indicated.
Expected results: No pathogen isolated.
Turnaround time: 48-72 hours.
▼ In positive cultures, additional time is required for isolation, identification, susceptibility testing, and further characterization, as appropriate.
The value of positive cultures critically depends on the quality of specimen collection: collection from the infected site and avoidance of contaminating flora.
Anaerobic culture is recommended for infections at sites likely to be infected by anaerobic pathogens. Examples include pelvic infections, intra-abdominal infections, abscesses, and traumatic and surgical wounds. Certain aerobic pathogens, such as Legionella species, require special processing or culture techniques for detection.
▼ Specimens may be collected from sites that are not the primary site of active infection (even though there may be signs of inflammation at the site).
▼ Inadequate site preparation may result in false-positive cultures due to specimen contamination with endogenous flora. Contaminated specimens may also mask the recognition of slow-growing or fastidious pathogens in the culture.
Some bacterial pathogens lack enzymes, like peroxidase, catalase, and superoxide dismutase, that efficiently inactivate reactive oxygen species. Strict anaerobes only grow in atmospheres with 0% O2, while aerotolerant organisms may tolerate up to 5% O2.
Anaerobic cultures are indicated for evaluation of patient specimens taken from sites with signs and symptoms of bacterial infection (e.g., swelling, redness, heat, pus, or exudate). Infections associated with anaerobic pathogens include surgical and traumatic wounds, sinusitis and pararespiratory infections, pelvic and intra-abdominal infections, osteomyelitis, myositis, gangrene, and necrotic wounds, abscesses, and actinomycosis and infections associated with fistula formation.
See Introduction for a discussion of general details related to bacterial cultures. Provide the laboratory with relevant clinical details to ensure special cultures procedures are not required. Specimens are inoculated on several types of anaerobic culture media. Media should be fresh and prereduced. Typical media for anaerobic cultures include the following:
▼ Supportive agar media, like Schaedler agar or CDC anaerobic blood agar
▼ Selective/differential agar media:
Phenylethyl alcohol agar or CNA for anaerobic gram-positive pathogens
Kanamycin-vancomycin-laked blood agar, for anaerobic gram-negative bacilli
Bacteroides bile-esculin agar, for Bacteroides fragilis group
Egg yolk agar, for characterization of Clostridioides (Clostridium) species
Cycloserine-cefoxitin-egg yolk-fructose agar (CCFA), for Clostridioides (Clostridium)
Broths, like enriched thioglycolate medium or chopped meat broth, are used to isolate pathogens present in low concentrations.
Successful anaerobic culture depends on maintaining strict anaerobic conditions through the whole diagnostic process, from specimen collection through final culture methods.
Because of the anaerobic endogenous flora, specimens from the following sites should not be submitted for anaerobic culture: sputum or other lower respiratory specimens; swabs from skin or mucosal surfaces; specimens from the GI tract (including fistulae, stoma surfaces, and so on); superficial ulcers or eschars, including decubitus ulcers; vaginal or cervical swabs; or urine (except suprapubic aspirate urine).
When submitting samples, ensure that sufficient specimen is collected for all of the diagnostic testing required (e.g., aerobic, fungal, and/or mycobacterial cultures and stains). Minimize exposure to atmospheric oxygen and transport in an anaerobic transport system. Do not refrigerate or
freeze specimens for anaerobic culture. Note the following: Specimens collected and transported for anaerobic culture are also acceptable for aerobic bacterial, fungal, or mycobacterial culture, provided a sufficient volume of specimen is provided.
Turnaround time: Incubation for 5-7 days.
▼ Additional time is required for positive cultures for additional testing required for isolation, confirmation as anaerobic (aerotolerance testing), identification, susceptibility testing, and further characterization, as needed. Anaerobic infections are frequently polymicrobial; final results may require several weeks for full laboratory evaluation, if needed.
Expected results: No anaerobic pathogen isolated
Anaerobic infections are frequently polymicrobial. Initial isolation and aerotolerance testing may require repeated subculture of the primary culture media. Many anaerobic pathogens are slow growing and biochemically indolent, making identification, susceptibility testing, and further characterization of isolates in the laboratory much slower than most aerobic bacterial pathogens. Therefore, patient care decisions must often be made before results of testing are available, limiting the clinical utility of extensive workup of mixed anaerobic cultures.
▼ Anaerobic culture may be significantly compromised by collection and transport conditions that are not strictly anaerobic or because of refrigeration during transport. Inadequate site preparation may result in false-positive cultures due to specimen contamination with endogenous flora. Contaminated cultures may also mask the recognition of slow-growing or fastidious anaerobic pathogens in the culture.
Antimicrobial susceptibility testing is performed to determine if an isolate is capable of expressing resistance to an antimicrobial agent. Panels of antibiotics typically used for treatment are inoculated for different groups of pathogens, like staphylococci and enteric GNBs. Routine test methods determine growth or inhibition of the test organism in vitro at different antibiotic concentrations. Quantitative test results are translated into interpretive categories.
Methods that detect specific resistance mechanisms, like β-lactamase, PBP2a, or molecular testing, may be used as adjunctive testing. Additional testing, like the D-Test, the oxacillin disk screen, and carbapenemase confirmation testing, may also performed as needed.
The Clinical and Laboratory Standards Institute (CLSI) has produced widely accepted standards for the performance, interpretation, and limitations of antimicrobial susceptibility testing for a wide range of pathogens. Standardized methods include broth dilution, agar dilution, and disk diffusion (Kirby-Bauer) testing. Other susceptibility methods, like automated platforms and gradient diffusion methods, have been validated against CLSI methods.
A number of factors must be carefully standardized in order to achieve reproducible results, including bacterial suspension preparation, specific media and additives, incubation conditions, etc. Also, the antibiotics against which an organism or group of organisms can be tested is clearly defined, as well as the range of antibiotic concentrations tested.
Standardized broth dilution methods determine the lowest concentration of antibiotic that inhibits growth of the challenge isolate, the minimum inhibitory concentration (MIC). The MIC values are compared to antibiotic concentrations achievable in vivo to generate interpretations regarding the likely utility of a specific antibiotic treating infection with the patient’s isolate.
▼ Susceptible (S): The organism does not show resistance to the antibiotic.
▼ Intermediate (I): The organism is not fully susceptible. However, therapy may be successful for treating infection at sites where the agent is concentrated, like urine. Also, therapy may be successful if high antibiotic dosage, frequency or delivery route (IV) are used.
▼ Susceptible-dose dependent (SDD): This category has recently been added for specific intermediate organism/antibiotic combinations for which clinical data confirm clinical effectiveness when specific dosing regimens are used.
▼ Resistant (R): The organism exhibits resistance mechanisms that indicate the agent is unlikely to cure infection or improve patient outcome.
▼ Nonsusceptible (NS): Reported for isolates for which only the S category has been defined because of the rarity of resistant clinical isolates. Because of this, isolates reported as NS should have repeat testing for identification and susceptibility.
Poorly collected specimens may yield growth of irrelevant organisms in culture, which may mistakenly be implicated in the infectious process.
Standardized methods are not defined or locally available for all types of organisms, like some fastidious organisms. Also, test methods or interpretive criteria have not been defined for all pathogen/antibiotic combinations. The use of nonstandardized testing or interpretations may lead to therapeutic errors.
This test is intended for the rapid initial detection of Streptococcus pneumoniae, Haemophilus influenzae type b, group B β-hemolytic Streptococcus (GBS), or Neisseria meningitidis in CSF. Published reports have demonstrated limited sensitivity for the detection of the target pathogens in patients with meningitis. Test results rarely change the management or therapy of patients. There may be some utility in patients who have been treated with antibiotics prior to CSF collection. There is some evidence that the performance for initial detection of GBS meningitis in neonates is acceptable.
Latex particles are coated with antibodies directed against specific antigens of the pathogens noted above. Agglutination should occur if the antigen is present in CSF. Specimens are collected and transported according to directions for CSF culture.
Turnaround time: <4 hours.
The sensitivity and specificity are too low to be recommended for routine use. Results must be carefully interpreted in the context of the patient’s clinical signs and symptoms.
The most common cause of fungemia is Candida albicans, which typically grows well in routine blood cultures. Other Candida species and Cryptococcus neoformans are usually efficiently isolated by routine blood cultures.
Fungal blood cultures are used for detection of bloodstream infection (BSI) caused by fungi, especially when dimorphic species and uncommon pathogens are suspected. Fungal blood cultures may be useful in septic patients in whom routing blood cultures are negative. The culture is indicated primarily for patients with increased risk of invasive fungal infection (IFI), including cancer, extensive therapy with broad-spectrum antibiotics, and HIV and other immunocompromising conditions.
Turnaround time: 4 weeks.
Inoculate blood culture system according to the manufacturer’s recommendations. Alert the laboratory if infection due to Malassezia furfur is suspected. Special culture processing is needed for isolation of this lipophilic yeast. Transport to the laboratory at room temperature.
Expected results: No growth
Most commonly isolated pathogens in positive cultures:
▼ Yeasts: C. albicans, non-albicans Candida species, and C. neoformans
▼ Dimorphic fungus: Histoplasma capsulatum
▼ Mold: Fusarium and Scedosporium species
Aspergillus species are rarely isolated by blood culture even in the presence of acute systemic infection.
The mycobacterial blood culture is used for the detection of BSI due to Mycobacterium species. Mycobacteremia is most commonly seen in patients with AIDS, although it may occur in other congenital and acquired immunocompromising conditions, including patients taking chronic corticosteroid therapy and malignancies. Growth of mycobacteria in culture requires the use of specialized, supplemented media with prolonged incubation time. Lysis of blood cells improves detection, by releasing phagocytized organisms, and is used in most methods.
Turnaround time: 4-8 weeks.
Ideally, inoculate blood directly into mycobacterial blood culture vials, or designated transport tube, like a lysis centrifugation tube, according to manufacturer’s instructions. Blood inoculated into routine blood culture media is unacceptable. If a specific vial is not available, collect 5-10 mL of blood anticoagulated with SPS or heparin. Transport to the laboratory at room temperature.
Expected results: No growth.
Positive results: Mycobacterium avium complex (MAC) is the most commonly isolated pathogen. Mycobacterium tuberculosis may be isolated at the time of hematogenous spread associated with severe primary or reactivation disease. Rapidly growing mycobacteria, such as M. fortuitum, have been associated with chronic indwelling vascular catheters and other prosthetic material.
Some mycobacterial infections are rarely associated with mycobacteremia. EDTA-anticoagulated blood should not be used for mycobacterial blood culture inoculum.
The routine blood culture is used for detection of BSIs due to common aerobic and anaerobic bacterial and yeast pathogens. Potentially pathogenic isolates are identified, and susceptibility testing is performed, as appropriate. Special testing is required for the detection of mycobacteria, parasites, viruses, and fungal pathogens.
▼ Sepsis syndrome, fever, chills, malaise, hypotension, poor perfusion, toxicity, tachycardia, and hyperventilation.
▼ Evaluation of serious localized infections, such as pneumonia, urinary tract infections (UTIs), and meningitis. Classic signs and symptoms may be absent in infants, the elderly, and patients with certain medical or surgical conditions.
▼ Most commercially available blood culture systems recommend inoculation of blood into two broth media: one aerobic and one anaerobic. Lysis centrifugation methods may be used for routine detection of BSIs due to bacteria or yeast but are more typically used for detection of mycobacteremia or fungemia.
Turnaround time: Generally, incubation for 5-7 days. Most true-positive blood cultures become positive within 24-48 hours after inoculation.
Decontamination of the collection site is the most important factor in preventing false-positive (contaminated) blood cultures. Inoculate media according to the manufacturer’s instructions. Usually, 8-10 mL of blood is inoculated into each blood culture bottle. A smaller inoculum volume, based on weight or age, is recommended for small children. Submission of two or three independently drawn (different venipuncture sites) blood cultures is recommended for the initial evaluation of patients with suspected BSI. Transport blood cultures to the laboratory at room temperature.
Expected results: No growth.
Positive results: Bacteremia or fungemia present. Positive blood cultures must be carefully evaluated to assess the possibility of false-positive culture, usually due to specimen contamination at the time of collection. Interpret all positive blood cultures in the context of number of positive blood cultures, organism identified, and clinical and laboratory signs and
symptoms. In patients with clinically relevant BSIs (true positive), the pathogen is typically isolated from a majority of cultures/bottles collected. In patients with contaminated blood cultures (false positive), a common contaminant is typically isolated in a single culture or bottle, whereas other cultures that are drawn during evaluation remain negative.
Negative results: Bacteremia and fungemia at the time of specimen collection are unlikely. False-negative results may be seen in patients with prior antimicrobial therapy. False-negative results may be caused by inoculation of blood culture bottles with less than the recommended volume of blood. Because clinically significant bacteremia may be intermittent, collection of two or three blood cultures is recommended to rule out bacteremia.
The significance of positive blood cultures must be evaluated in terms of several factors, including patient signs and symptoms, the intrinsic pathogenicity of the blood culture isolate, number of positive cultures, number of isolates in culture (mixed cultures typically represent contamination), and cultures positive at other infected sites for the blood culture isolate.
Routine blood cultures are optimized for detection of the pathogens most frequently associated with BSIs. Clinically relevant BSIs may be associated with pathogens for which special blood cultures are required (e.g., mycobacteria, dimorphic fungi, fastidious bacteria).
▼ Decreased sensitivity because of such factors as a low volume of blood inoculated into blood culture media. Decreased specificity because of contamination due to poor preparation of collection site.
This test is used to detect parasites circulating in peripheral blood. It should be ordered in patients when infection caused by Plasmodium species (malaria), Babesia species (babesiosis), Trypanosoma species (sleeping sickness, Chagas disease), or certain microfilaria species or systemic infection with Leishmania species is suspected. Thin and thick blood smears are prepared from free-flowing capillary blood or EDTA-anticoagulated blood. Smears are inspected after staining with Giemsa, Wright, or Wright-Giemsa stain. For positive smears, the level of parasitemia should be determined for each specimen.
▼ Preliminary examination should be performed “STAT” if malaria is suspected (ideal turnaround time <4 hours). Final report for initial positive smears: <24 hours.
EDTA-anticoagulated blood is most commonly collected. For microfilariae, the diurnal circulation of some species must be taken into account in timing specimen collection (Loa loa, 10 am to 2 pm; Wuchereria or Brugia species, 8 pm to 4 am). Transport specimens to the laboratory and prepare smears as soon as possible. If microfilaremia is suspected, special concentration techniques (Knott) may be requested.
In general, collect specimens on each of 3 successive days. Collect specimens every 6-12 hours (or until positive) for optimal detection in suspected cases. Blood should be examined in treated patients after 24, 48, and 72 hours to determine effectiveness of therapy.
Negative results: Blood parasites not detected. However, a single negative specimen cannot rule out parasitemia.
Positive results: Disease caused by specific parasite confirmed.
Low level of parasitemia may require the examination of multiple specimens for detection. Examination of smears prepared from buffy coat preparations may improve the sensitivity of detection for some parasites, like microfilaria and trypanosomes. The efficient detection of microfilaria requires specimen collection during the specific hours when circulation of the parasite is expected. Polymerase chain reaction (PCR) may be requested for patients with a high index of suspicion of malaria or babesia if initial blood parasite examinations are negative.
In effectively treated patients, the level of parasitemia should drop very quickly. In patients with drug-resistant parasites, the level may remain stable or even increase.
Sterile fluid-filled spaces are present at a number of anatomic sites and are subject to infection. Examples of sterile fluids include peritoneal, pleural, pericardial, and synovial/joint fluid. Infections associated with CSF are life threatening and associated with a different etiology of bacterial
pathogens, so these cultures are typically processed differently than other sterile fluids. Urine is also processed with different culture techniques because of its connection with the external environment, via the urethra, and pathogenesis of infection. A broad etiology of bacterial pathogens may cause infections of sterile sites, and culture methods are optimized for recovery of organisms present in low concentrations.
Sterile fluid cultures are used to diagnose serious infections in normally fluid-filled spaces in the body. Collect sterile fluid cultures from sites associated with signs and symptoms of inflammation, including redness, swelling, pain, heat, fluid accumulation, and pus formation.
Method: Supportive, enriched, and selective/differential solid agar and broth media are typically inoculated. Anaerobic culture should be requested if there is a significant possibility of anaerobic pathogens. If infection with an uncommon, fastidious pathogen is suspected, the laboratory should be informed so that special cultures can be inoculated.
Turnaround time: Cultures are incubated for up to 7 days. Additional time is required for isolation, identification, susceptibility testing, and further characterization, as needed.
Fluid aspiration is performed after preparation of the puncture site in a manner consistent with a preparation of a surgical site. Specimens from drainage devices should not be submitted because of the high incidence of contamination with endogenous flora.
Because many infections are associated with a low concentration of organisms, collection of larger volumes of fluid and repeated sampling improve culture sensitivity. Transport fluid to the lab in a sterile transport container. In addition, always submit 1-10 mL of fluid in an anaerobic transport vial for Gram stain and anaerobic culture, if indicated. Ensure that sufficient fluid is collected for all microbiology tests required. Sensitivity may be improved by inoculation of up to 10 mL of fluid into blood culture media at the patient bedside. However, if polymicrobial infection is suspected, like traumatic peritonitis, inoculation of blood culture media is not recommended.
If volume is limited, prioritize testing for the sample, and then collect additional material if needed.
Swabs should not be used for fluid collection, like intraoperative cultures.
Specimens transported in anaerobic transport vials are acceptable for inoculation of cultures for aerobic bacterial, mycobacterial, and fungal cultures.
Submission of several specimens prior to antibiotic therapy may significantly improve sensitivity of culture detection.
The use of anticoagulants is discouraged because of possible inhibition of some pathogens. If anticoagulation is required, heparin or SPS is recommended.
Transport specimens at room temperature; do not refrigerate or freeze specimens.
Negative results: Infection is less likely, but not excluded by a negative culture, especially after initiation antibiotic therapy. Uncommon, fastidious pathogens may not be isolated in culture without inoculation of special media.
Positive results: Indicates infection of the sterile site, but cultures that may be contaminated with endogenous flora must be interpreted with caution in the context of quantity of bacterial growth, purity of culture, Gram stain findings, and clinical signs and symptoms. Infected peritoneal fluid may yield numerous aerobic and anaerobic pathogens. Extensive identification and susceptibility testing are usually not clinically useful: final results are often not available until well into therapy, and empirical treatment is usually effective.
This test is used to detect acute infection caused by the slow-growing, fastidious pathogen B. pertussis, the cause of pertussis or whooping cough. Nasopharyngeal specimens should be submitted for B. pertussis culture. Aspirates into a trap, using an infant-sized feeding tube, are preferred. Alternatively, a small-tipped NPS is passed along the floor of the nasal cavity into the posterior nasopharynx and allowed to sit for 30 seconds prior to removal. Anterior nares or throat specimens are unacceptable. Regan-Lowe transport medium is recommended. Specimens are inoculated onto an enriched, selective agar, usually Regan-Lowe media.
Turnaround time: Most cultures are positive in 7-10 days, although some cultures are incubated for up to 14 days. Additional time is required for final identification and further characterization.
Negative results: A negative culture does not exclude the diagnosis of pertussis, especially when a specimen is collected after the early, acute phase of infection.
Positive results: Confirms the diagnosis of pertussis.
The sensitivity of culture for B. pertussis falls significantly after the first 7-14 days after onset of symptoms. Poor specimen collection, submission of specimens other than nasopharyngeal specimens, and submission of specimens during the chronic phase of disease are associated with poor sensitivity of culture.
PCR and serologic methods are discussed below.
Pertussis, a respiratory tract infection also known as “whooping cough,” is highly contagious. It is caused by the gram-negative coccobacillus B. pertussis.
It is characterized clinically by a severe and prolonged cough. A clinical diagnosis will form the basis of most pertussis diagnosis and treatment decisions.
Serodiagnosis may be used to confirm the clinical diagnosis of pertussis.
Expected result: Negative.
Aids in the detection of B. pertussis infection in patients. Optimal time point is ≥2-8 weeks after onset of consistent symptoms.
Natural infection with B. pertussis results in production of immunoglobulin IgA, IgG, and IgM antibodies to a variety of antigens; primary immunization induces mainly IgG and IgM antibodies.
Serologic testing typically involves comparing levels of pertussis antibodies IgA or IgG to pertussis toxin [PT], filamentous hemagglutinin [FHA], pertactin [PRN], fimbriae, or sonicated whole organism.
The most reliable serologic approach to diagnosis of pertussis is with simultaneous testing of paired acute and convalescent sera. A significant increase (fourfold or greater) in IgG or IgA antibody titers to PT or FHA, comparing convalescent to acute sample, suggests recent B. pertussis infection in patients with a clinical illness compatible with pertussis.
Paired sera, however, are not practical in most clinical settings. Single-sample serology tests for antipertussis toxin IgG must be collected at least 2 weeks after symptom onset. A high antibody titer >2 years following vaccination supports the diagnosis of pertussis.
Positive results: IgG antibody to B. pertussis detected, which may indicate a current or past exposure/immunization to B. pertussis
Laboratory diagnosis of pertussis is complicated by the limitation of available tests and lack of standardization.
Interference of previous vaccinations or previous infections with serodiagnosis, cross-reactivity with other Bordetella species, and the variable response to B. pertussis antigens limits interpretation.
CDC-recommended first-line tests for pertussis are B. pertussis/B. parapertussis by PCR and/or B. pertussis culture.
The IgG serology test results are not interpretable in children younger than 11 years of age because of interference due to persistent antibody formed by childhood vaccination. The test also cannot be interpreted in older patients who have received the Tdap vaccination in the previous 3 years.
For diagnosis of individuals suspected of a respiratory tract infection caused by B. pertussis or B. parapertussis
Negative results: DNA of B. pertussis or B. parapertussis was not detected; this result does not preclude B. pertussis or B. parapertussis infection.
Positive results: DNA of B. pertussis or B. parapertussis was detected in the patient’s sample; this result does not rule out coinfections with other respiratory pathogens.
New nucleotide variants in the primer/probe regions may lead to false-negative molecular results.
This assay test only for presence of B. pertussis or B. parapertussis, additional testing for other respiratory pathogens is needed in case of negative results.
Lyme disease is the most common tick-borne disease in the United States, Canada, and Europe. It is a bacterial infection caused by six species in the spirochete family Borreliaceae.
In North America, infection is caused primarily by B. burgdorferi and, less commonly, in a region of the upper mid-West, by Borrelia mayonii. In Europe and Asia, infection is caused primarily by either Borrelia afzelii or Borrelia garinii, less commonly by B. burgdorferi, and rarely by Borrelia spielmanii or Borrelia bavariensis. There is a broad spectrum of manifestations, and severity of disease is due, in part, to differences in the infecting species.
A sensitive serum serology screening test for the detection of IgG and/or IgM antibodies to B. burgdorferi is essential for the diagnosis.
In the United States, most laboratories use a whole-cell sonicate preparation of B. burgdorferi as antigen for initial serologic (enzyme immunoassay [EIA]) assays. This test approach has high sensitivity because of multiple antigens in the whole-cell sonicate preparation. However, because some of these antigens are cross-reactive with antigens from the host or other
pathogens, specificity of the EIA alone is not optimal. Additional FDA-cleared EIAs that use as few as 1 to several antigens, which results in a higher specificity and similar sensitivity than that for whole-cell sonicate EIAs, have recently become commercially available. There are several FDA-cleared assays that target the cell surface variable major protein-like sequence expressed (VlsE) lipoprotein and its sixth invariable region, the C6 peptide. These Borrelia antigens are highly conserved and immunogenic among all Lyme borreliosis species and strains and cause an early antibody response useful for diagnostic testing.
Aids in the diagnosis of Lyme disease in suspected at-risk patients.
Expected results: Negative.
In serologic testing for anti-B. burgdorferi antibodies, a two-tier conditional strategy is recommended to support the diagnosis. The traditional two-tiered testing algorithm uses a sensitive enzyme immunoassay followed by a more specific Western blot test. Separate IgM and IgG blots are typically performed.
Several modified algorithms using two enzyme immunoassays have been developed to improve the sensitivity of diagnosis for early Lyme disease and reduce the need for Western blot testing. One strategy uses a whole-cell-based ELISA followed by a C6 ELISA. However, clinical laboratories are not generally using this approach because the two enzyme immunoassays have been cleared by the U.S. Food and Drug Administration (FDA) only for use as first-tier assays and have not been FDA cleared for sequential use in replacing Western blots.
Should not be used to screen general population. False-negative results can occur if the patient is tested too early; repeat testing in 2-4 weeks. The IgG antibody response is usually not detectable until 4-6 weeks after infection; the IgM antibody response usually not detected during the first 2 weeks of infection, peaking 3-6 weeks following infection.
False-positive results may occur from other spirochetal diseases, autoimmune diseases, or other infections (Epstein-Barr virus [EBV], HIV, syphilis, infectious mononucleosis, etc.). IgG antibodies can be detected as early as 2 weeks, and both IgM and IgG antibodies can remain detectable for years. Diagnosis depends on clinical features, combined with available laboratory tests.
A positive IgM result alone should not be used to support the diagnosis of Lyme disease in patients who have had symptoms of early infection for >6-8 weeks without treatment. If a patient has a positive IgM ELISA and IgM Western blot after that time, but a negative IgG blot, the IgM test most likely represents a false-positive result, or possibly evidence of past treated infection.
Borrelia miyamotoi should be considered in the evaluation of patients who present with an acute nonspecific febrile illness if they are from an area where B. miyamotoi has been previously identified. In patients suspected of having B. miyamotoi based upon their clinical presentation, the diagnosis depends upon the availability of laboratory testing by PCR in blood or CSF. Serologic assays are less useful in diagnosing acute infection since assays that detect antibodies to B. miyamotoi are not always positive early in the course of disease.
The Western blot assay for antibodies to B. burgdorferi, the primary cause of Lyme disease, is a qualitative method of categorizing specific immunoreactivities in serum or plasma to B. burgdorferi proteins that have been formatted according to molecular weight into discrete bands on nitrocellulose strips.
The Western blot assay for B. burgdorferi is used as a second-tier test to characterize the specificity of an individual’s immune response to the component proteins of B. burgdorferi by identifying the presence, relative level, and pattern of reactivities to the complete set of the bacterial antigens. The assay is used routinely to provide supportive serologic evidence of infection following a more sensitive but less specific screening test (such as EIA) for general reactivity to B. burgdorferi. Both IgM and IgG reactivities to the bacterial proteins are assayed to provide information on the evolution of the immune response relative to the stage of infection (i.e., early localized, early disseminated, or late disseminated). A caveat to this use is that IgM testing is not recommended in patients with symptoms present for >1-2 months. For such patients, IgG testing alone should be performed.
Reactivity scores: Specimen reactions with protein bands are first scored in terms of relative reaction intensity versus a cutoff control or minimally positive (“+”) band reaction intensity by a positive control specimen.
Test interpretation (IgM class reactivities)
▼ Positive results: At the early stage of the disease (1-2 weeks after the onset of erythema migrans), reactivity scores of “+” or greater for at least two of three clinically significant proteins: 41, 39, ospC (24) kDa
▼ Negative results: Absence of any band reactivity on the test strip or reactivity for only one of the three clinically significant proteins
Test interpretation (IgG class reactivities)
▼ Positive results: At the later stages of the disease (2-6 or more weeks after the onset of erythema migrans or initial symptoms), reactivity
scores of “+” or greater for at least 5 of 10 clinically significant proteins: 93, 66, 58, 45, 41, 39, 30, 28, 23, 18 kDa
▼ Negative results: Absence of any band reactivity on the test strip or reactivity for <5 of the 10 clinically significant proteins
Minimum specimen volume is 40 µL (20 µL each for the IgM and IgG tests).
Like any second-tier test, the positive predictive value for a Western blot assay is a function of the a priori likelihood of the disease based on clinical and epidemiologic criteria, whereas the negative predictive value is not as well defined because of the variability of the immune response among infected individuals. Cross-reactive diseases are most frequently evidenced by reactivity to the 41-kDa flagellar protein and at much lower frequency to the 66-kDa heat shock protein. Specimens from patients diagnosed with Ehrlichia or Babesia infections can show other Borrelia-specific bands.
Quantitative bacterial cultures of specimens collected bronchoscopically (bronchoalveolar lavage [BAL] or protected brush) are submitted for the evaluation for ventilator-associated pneumonia (VAP). The diagnosis of VAP is challenging, requiring a combination of clinical, imaging, and laboratory studies. Cultures are assessed in comparison to predetermined thresholds.
Protected brush and BAL specimens are collected using standard procedures.
Brush: After collection, the brush end should be removed, using sterile technique, and placed in a small volume (1 mL) of nonbacteriostatic saline for transport.
BAL: Undiluted samples should be transported to the laboratory as quickly as possible.
▼ A Gram stain is performed on all specimens.
▼ Known volumes of the specimen (or specimen dilutions) are quantitatively inoculated onto solid agar media, including SBA and chocolate and MacConkey agar (and other media as required for uncommon pathogens, such as Legionella), and spread so that individual colonies can be counted; quantitative results are calculated on the basis of the number of colonies isolated and dilution factor.
Protected brush: The brush is vigorously agitated in the saline transport fluid to release trapped microorganisms. The saline is then used to prepare dilutions for media inoculation.
BAL: A measured aliquot of BAL fluid is used to prepare dilutions for media inoculation.
▼ After incubation, the concentration of each respiratory pathogen is calculated using the colony count on the solid media, volume inoculated onto the solid media, and the dilution of the original specimen. Common contaminants, like coagulase-negative staphylococci or diphtheroids, are not quantified. Cultures are interpreted on the basis of the isolate identification, quantity of isolate in culture, and the presence of other flora, especially endogenous flora of low pathogenicity.
Turnaround time: Incubation for 48 hours. Additional time is required for pathogen isolation, identification, susceptibility testing, and further characterization, if needed.
Expected results: A low quantity of endogenous upper respiratory flora is often seen.
Gram stain results: VAP is supported by many polymorphonuclear neutrophil (PMN) and a predominant bacterial morphotype, like GNB or GPCs in clusters. VAP is less likely when few PMN (<50% of nucleated cells) and mixed bacterial morphotypes without predominant form are seen.
Positive culture results: In patients with clinical VAP:
▼ Bronchial brush: Growth of >1,000 cfu/mL of a respiratory pathogen is significant.
▼ BAL: Growth of >10,000 cfu/mL of a respiratory pathogen is significant.
Negative culture results: False-negative cultures may be caused by prior antimicrobial therapy. Detection of pneumonia caused by certain fastidious pathogens may require inoculation of special media. Heavy contamination of the specimen with endogenous flora may mask the growth of the causative pathogen.
Human infection may be caused by several species of the genus Brucella. These organisms are fastidious, slow-growing gram-negative bacilli capable of producing severe localized and systemic infection. Infections have typically been acquired by zoonotic transmission, primarily related to livestock and dairy industries. There is great concern regarding the use of this organism for a bioterror-related attack. It is critical that the laboratory be informed whenever brucellosis is suspected to minimize potential for laboratory-acquired infection.
This culture is used to isolate Brucella species from clinical specimens. Because of the risk of laboratory-acquired infection and because isolation of Brucella species may represent a sentinel event in a bioterror attack, most clinical microbiology laboratories limit the workup to screen suspected isolates. Isolates that fail to “rule out” are to their local public health laboratory for identification and further characterization.
Method: Specimens are inoculated onto a blood agar, chocolate agar, MacConkey agar, and Thayer-Martin agar (if contamination with endogenous flora is suspected).
Turnaround time: Isolation and preliminary identification are usually available in 3-7 days. Additional time is required for transfer to the local public health laboratory, confirmation of identification, and further testing.
Bone marrow and blood are the specimens of choice for patient evaluation. Specimens from other infected tissue or sites should also be submitted for culture. Serologic testing is recommended for diagnosis in patients with suspected brucellosis.
Expected results: Negative.
Positive results: Isolation of Brucella in culture is diagnostic for brucellosis.
Brucella may be difficult to detect by Gram stain in primary specimens.
Common pitfalls: Because brucellosis may present after a prolonged incubation period, or present with nonspecific symptoms and an indolent onset, the diagnosis may not be considered until progression into the chronic phase of illness. Clinicians may fail to request specific cultures for brucellosis, or alert the laboratory of their clinical suspicion.
Brucellosis is a reportable disease. Patients with a clinical suspicion or laboratory diagnosis of brucellosis must be reported to the local department of health.
CSF culture is used for specific diagnosis of bacterial meningitis. Patients commonly present with severe headache, fever, neck stiffness, and meningeal signs, mental status changes, and signs of systemic toxicity.
▼ CSF is inoculated onto sheep blood and chocolate agar, incubated aerobically. Broth media may be inoculated. Special media or culture
conditions may be used for non-community-acquired meningitis, such as infections associated with trauma and prosthetic implants.
Turnaround time: Routine cultures are incubated for 96 hours. Additional time is required for isolate identification, susceptibility testing, and further characterization, as needed.
CSF is collected by needle aspiration after preparation of the puncture site in a manner consistent with a surgical site preparation.
Fluid is transported in a sterile container or tube with a tight-fitting lid.
CSF should be transported at room temperature; do not refrigerate or freeze for transport.
CSF specimens submitted for bacterial culture are also acceptable for fungal or mycobacterial stains and culture, antigen testing, and Venereal Disease Research Laboratory (VDRL), if sufficient volume of fluid is submitted.
Expected results: No growth. False-negative cultures may be caused by low pathogen concentration in CSF, especially when low-volume samples are submitted, or prior antibiotic therapy.
Positive results: Positive CSF culture supports a specific diagnosis of meningitis. False-positive cultures may be caused by contamination with endogenous skin flora. For most bacterial pathogens, CSF samples in patients with acute bacterial meningitis usually show increased WBCs (PMNs predominate), increased protein, and decreased glucose.
A broad etiology, which may require a number of different tests for diagnosis, may be considered for patients presenting with signs and symptoms of meningitis. The volume of CSF submitted is often insufficient for optimal sensitivity for the range of tests requested.
This culture method is used to diagnose infections caused by C. trachomatis, an obligate intracellular pathogen. Tests based on nucleic acid amplification have emerged as the most sensitive methods for diagnosis of Chlamydia genital infections, but Chlamydia culture is still required for specimen types for which molecular diagnostic tests have not been validated. Chlamydia cultures should also be performed in cases that may have legal implications, such as rape and child abuse.
▼ Infected cells from patient specimens are inoculated onto cultured eukaryotic cells, most commonly McCoy cells.
▼ Positive cultures are now most commonly detected by staining fixed monolayers with specific anti-C. trachomatis antibodies; positive cultures show staining of intracellular inclusions. The sensitivity of cultures for C. trachomatis detection may be improved by blind subculture of a primary culture after the initial incubation.
Turnaround time: Cultures are incubated for 72 hours. An additional 48-72 hours are required if primary cultures are subcultured prior to final examination.
It is critical to collect infected epithelial cells from infected sites using toxicity-tested swabs or other device. Swabs may be premoistened with sterile nonbacteriostatic saline before specimen collection. Scrapings or biopsy specimens may be submitted for some specimen types.
Place specimens into a Chlamydia transport medium, such as 2-SP, and transport to the laboratory at 4°C. Deliver to the laboratory as quickly as possible.
Specimens commonly submitted for Chlamydia culture come from the following sites:
▼ Cervix: Remove excess mucus from the exocervix. Insert a new swab approximately 1 cm into the cervical canal and gently rotate for 10-15 seconds.
▼ Urethra: Clean the distal urethra and meatus with a swab. Insert a new thin-shafted swab 2-4 cm into the urethra and gently rotate for 10-15 seconds.
▼ Conjunctiva: Remove excess purulent discharge with a swab. With a new swab, gently rotate over the affected conjunctival surface.
▼ Anus: Insert a premoistened swab into the anorectal juncture and rotate gently. The swab should not be heavily stained with feces.
▼ Fallopian tube or epididymis: Place aspirate into an equal volume of Chlamydia transport media.
▼ Respiratory tract (neonate): Place aspirate or wash into an equal volume of Chlamydia transport media.
Expected results: No growth.
Positive results: Chlamydia culture is very specific for infection caused by C. trachomatis.
Negative results: Chlamydia infection is not ruled out by a negative culture. Repeat testing, using a nucleic acid amplification test if appropriate for the site, is recommended for patients with a high suspicion for chlamydial infection.
Chlamydophila species, C. psittaci and C. pneumoniae, are not isolated by C. trachomatis culture. The following specimens are not recommended for
Chlamydia culture: peritoneal fluid, urethral discharge, urine, cul-de-sac fluid, vagina, or throat.
▼ Poor specimen collection (sample selection or collection technique) or loss of viability during transport. Swabs may be toxic for C. trachomatis. Specific types and lots of swabs should be tested for toxicity before releasing before clinical use. Urethral specimens should not be collected within 1 hour after the patient has urinated.
Laboratory tests for C. difficile are not used as the primary criteria for diagnosis of C. difficile infection (CDI). Rather, they are performed to support a clinical diagnosis of CDI.
Severe diarrhea without other cause is present in the vast majority of patients with CDI. Most patients also have fever, elevated WBCs, and abdominal pain. Severe diarrhea is defined as three or more unformed stools in 24 hours. Unformed stool will flow and take the shape of the transport container.
Note: Laxatives and stool softeners may cause significant diarrhea. It is recommended that laxatives be discontinued, C. difficile testing deferred, and the patient reevaluated for possible CDI 48-72 hours later.
C. difficile is a major cause of antibiotic-associated diarrhea and pseudomembranous colitis, and it represents a significant and serious agent of nosocomial infection. CDI may be mild and self-limited after discontinuation of antibiotics, but a significant number of patients have persistent and/or severe diarrheal illness that may progress to pseudomembranous colitis or toxic megacolon.
C. difficile is an anaerobic, spore-forming, gram-positive bacillus. It forms several toxins (toxins A and B) that have been used as the basis for detection. C. difficile detection is recommended for patients in whom diarrheal illness develops after antibiotic therapy or during hospitalization, especially when colitis (increased fecal leukocytes) is a prominent feature.
Liquid stool specimens collected in clean containers with tight-fitting lids should be transported to the laboratory at room temperature within 2 hours. If transport will be prolonged, the specimen should be held at refrigerator temperature. Do not freeze.
▼ Cytotoxicity: These assays detect C. difficile toxin B in feces. Stool, passed through a membrane filter to remove bacteria, is inoculated on cultured eukaryotic cells. The cell monolayer is examined for 48 hours for evidence of typical “actinomorphic” cytotoxicity. In order to exclude nonspecific cytotoxicity that may be seen with stool filtrates, neutralization of the cytotoxic effect, using anti-C. difficile antiserum, must be demonstrated.
▼ Toxigenic culture: These cultures isolate C. difficile organisms from feces. C. difficile may be isolated from stool using a spore selection technique (by heat shock or alcohol pretreatment of a stool suspension prior to medium inoculation) and selective media. Because not all C. difficile isolates produce the toxins associated with disease, culture supernatants must be tested for toxin production to support a diagnosis of C. difficile-associated disease.
▼ Toxin detection using immunodiagnostic procedures: Toxin immunoassays demonstrate the presence of C. difficile toxin A and/or B in feces. A number of commercially available latex agglutination or EIA kits have been developed for the detection of C. difficile toxin A and/or toxin B in stool samples. These assays provide rapid turnaround time.
▼ C. difficile glutamate dehydrogenase antigen (GDH) detection: These assays demonstrate the presence in feces of an antigen present in all C. difficile strains. Because glutamate dehydrogenase is not specific for toxigenic C. difficile, or for the presence of a significant amount of toxin in stool, positive specimens must be tested for toxin to support the diagnosis of C. difficile-associated disease.
▼ Molecular diagnostic methods: These methods detect the presence of toxigenic C. difficile in feces, but they do not provide information about the amount of toxin present in stool. Molecular diagnostic methods for detection of the C. difficile are commercially available and provide the most sensitive assay for CDI. The test is very specific when used on patients with typical clinical signs and symptoms of C. difficile disease. It is important to make sure before sending sample for molecular testing that patient has diarrhea and multiple stools per day and is not on stool softeners or laxatives. Sensitivity of the molecular assays may cause clinically false positives if asymptomatic C. difficile carriers are tested. Molecular assays are detecting toxin B (tcdB) gene, rarely toxin A (tcdA) gene, which are parts of the pathogenicity locus (PaLoc). Some assays test also for binary toxin (CDT) locus containing two genes (cdtA and cdtB) and specifically the single base pair deletion at nucleotide 117 in the tcdC gene. This deletion disrupts the action of negative regulation of toxin production in “hypervirulent” strains 027/NAP1/BI, which exhibit increased toxin production.
▼ Molecular diagnostic, immunodiagnostic assays, and the glutamate dehydrogenase assays: 24 hours
▼ Cytotoxicity assays: 24-72 hours
▼ Culture: 96+ hours
Expected results: Negative.
▼ Tests that detect the presence of significant quantities of toxin in feces (cytotoxicity and toxin EIA tests) are the most specific for supporting a diagnosis of CDI, but they may be less sensitive in symptomatic disease associated with low concentration of toxin.
▼ Tests that detect the presence of C. difficile in feces (GDH EIA, toxigenic culture, NAAT) represent the most sensitive detection of C. difficile; however, these methods do not determine the amount of toxin actually present in feces. They may be, therefore, less specific when stool is submitted from patients without a clinical diagnosis of CDI.
The available assays vary somewhat in sensitivity and specificity for diagnosis of C. difficile disease. The choice of diagnostic methods must take cost, assay performance, turnaround time, and other factors into consideration.
Positive C. difficile test results must be interpreted with caution in infants; toxin may be detected in the stool of healthy infants without signs of diarrheal illness or colitis.
This culture is used to detect C. diphtheriae in clinical specimens. It should be considered in patients who present with signs and symptoms consistent with diphtheria, either localized pharyngeal or cutaneous infection or systemic disease, suggesting the action of diphtheria toxin on the heart, central or peripheral nervous system, liver, or kidney. Diphtheria is now uncommon in countries that have implemented widespread vaccination programs against this pathogen.
▼ Agar-enriched media containing cysteine and tellurite, such as cysteine-tellurite blood agar or modified Tinsdale agar, are used for C. diphtheriae detection. Plates are incubated at 35-37°C for at least 48 hours in room air.
▼ On cysteine-tellurite agar, C. diphtheriae produces black or gray colonies; on modified Tinsdale agar, black colonies are surrounded by a dark brown halo.
▼ In isolates with characteristic appearance on modified Tinsdale agar, a negative urease reaction distinguishes C. diphtheriae from other Corynebacterium species. Definitive identification of suspect colonies must be confirmed by biochemical, molecular, or other techniques. C. diphtheriae isolates should be referred to test for toxin production.
Turnaround time: 48-72 hours for initial isolation. Additional time is required for confirmation of the identity, toxin testing, and further characterization of suspicious isolates.
The laboratory should be alerted before specimen submission to ensure that appropriate medium is available for culture inoculation.
Swab specimens are collected from multiple inflamed sites of the pharynx or other respiratory mucosal surfaces. Collection of specimens from near or under any diphtheritic membrane is recommended.
Aspirate, swab, or tissue samples for detection of cutaneous diphtheria are collected for skin lesions.
Routine transport media can be used for transport.
Expected results: No growth.
Positive results: Isolation of toxigenic strains of C. diphtheriae from upper respiratory or cutaneous lesions is diagnostic of diphtheria.
Negative results: Submission of multiple specimens may be required for C. diphtheriae isolation.
This test may be ordered for the early diagnosis of infections caused by C. neoformans. It is recommended for immunocompromised patients presenting with clinical signs of meningitis. Testing is most sensitive when testing CSF for cryptococcal meningitis. Testing serum may be submitted for detection of infection at other sites.
▼ There are several formats for commercially available cryptococcal antigen tests, including latex agglutination, EIA, and lateral flow assays. Determination of antigen titer is recommended for positive CSF specimens to monitor response to treatment.
▼ Titers of 1:8 or greater indicates active disease. Approximately 95% of patients with cryptococcal meningitis are detectable by cryptococcal antigen testing of the CSF.
▼ The sensitivity for CSF is 93-100%, and for serum, it is 83-97%. Specificity for both specimen types is typically >95%.
Turnaround time: <24 hours.
Expected results: Negative.
Positive results: Cryptococcal infection very likely. Positive results should be confirmed by culture.
Negative results: Cryptococcal infection unlikely. Use fungal culture to definitively rule out cryptococcal infection.
False-negative reactions may occur, especially due to a prozone effect in latex agglutination assays. Prozone effect may be minimized by testing serially diluted specimens. Some isolates from profoundly immunocompromised patients may produce very little polysaccharide capsular material, resulting in false-negative tests.
There are several sources of false-positive reactions. Positive reactions caused by rheumatoid factor (RF) may be reduced by pretreatment of the specimen with pronase, EDTA, or reducing agents. The syneresis
fluid from agar media can cause false-positive results; an aliquot of the specimen for cryptococcal antigen testing should be removed before medium inoculation. Finally, several uncommon pathogens, including Trichosporon asahii, Rothia, and Capnocytophaga species, can cause false-positive cryptococcal agglutination reactions.
▼ Positive cryptococcal antigen titers should be confirmed by culture to document active infection and rule out false-positive reactions. Some infected patients may have very low antigen titers. All specimens submitted for cryptococcal antigen testing should be accompanied by cultures of spinal fluid, blood, or other potentially infected material for fungal isolation.
Antigen titers are usually higher in patients with AIDS compared to those seen in HIV-negative patients with cryptococcal infection. In patients with AIDS, baseline CSF antigen titers <1:2,048 are associated with improved prognosis. Antigen titers should fall with effective antifungal therapy. Steady or increasing cryptococcal antigen titers, even with sterilization of cultures, are an indication of likely treatment failure and recurrence of infection.
This test is used to evaluate diarrheal disease in patients at risk for cryptosporidiosis, specifically for the identification of Cryptosporidium parvum in stool specimens.
▼ Enzyme immunoassays are used. EIAs have very high sensitivity (near 100%) and specificity (near 100%) compared with a series of stool O&P examinations. For information about microscopic Cryptosporidium detection, see the test Ova and Parasite Examination, Stool.
▼ Different EIA assays for fecal parasite detection have different specimen requirements (fresh vs. preserved) and transport conditions. Laboratories should provide assay-specific information for their providers.
Turnaround time: 24-48 hours.
Expected results: Negative
Examination of several specimens improves detection in patients with light infection. A series of O&P examinations is recommended in patients with repeatedly negative immunoassays in whom parasitic infection is still suspected.
CMV is a ubiquitous viral pathogen. Most infections in immunocompetent patients are asymptomatic or mildly symptomatic, including a self-limited mononucleosis syndrome. In immunocompromised patient populations, including neonates, patients with AIDS, and transplant patients, serious localized (e.g., retinitis, colitis, polyradiculopathy, encephalopathy) or systemic infection may occur.
▼ Specimens for CMV culture are usually inoculated onto monolayers of human fibroblasts (e.g., foreskin, fetal lung). Shell vial cultures provide a more rapid turnaround time than tube cultures but are somewhat less sensitive for detection.
Turnaround time: Specimens with high viral loads, such as urine, may give positive results within several days, but negative cultures may require incubation for up to 4 weeks before signing out as negative. Shell vial cultures are processed for growth at 48-72 hours after inoculation.
Specimens should be collected according to general recommendations for virus culture of the specimen type.
Specimens should be collected early in acute infection.
Urine is most often recommended for evaluation of neonates with suspected CMV infection. For evaluation of patients with suspected viremia, heparinized whole blood or isolated buffy coat cells are used to inoculate cultures.
CMV is a fastidious virus and should be delivered to the laboratory as quickly as possible. Most specimens should be placed in a viral transport medium and transported at 4°C; do not freeze.
Expected results: Negative.
Negative results: Negative cultures do not rule out CMV infection; they may be due to loss of viability after collection or low viral load in the specimen submitted.
Positive results: Positive cultures usually indicate active CMV infection. Occasionally, positive cultures represent asymptomatic shedding of virus not associated with disease.
Positive cultures may be due to asymptomatic shedding during latent infection; correlation with histopathology and other clinical signs and symptoms may be needed to ensure specific diagnosis.
The CMV quantitative assay uses real-time PCR to quantify CMV DNA extracted from plasma, serum, CSF, saliva, or urine. The first WHO International Standard for human cytomegalovirus (HCMV), NIBSC code 09/162, was accepted by all laboratories in the United States allowing for standardization of nucleic acid amplification technique (NAAT) assays. The test quantifies CMV DNA and the results expressed in IU/mL (international units/mL). The qualitative assay reports only if virus was detected or not detected in the sample.
Normal values: Not detected (the result is below the level of detection of the assay).
Not detected: CMV DNA is not detected.
Number in IU/mL: Calculated viral load (IU/mL) is within the linear range.
>ULoQ (upper limit of quantitation): Result is above the validated linear range; if client desired a viral load, the original specimen should be diluted with CMV-negative human EDTA plasma/serum, and test should be repeated; reported result is calculated by multiplication of obtained result by the dilution factor.
Although rare, mutations of the viral genome may result in underquantification or failure to detect the presence of the virus in these cases (false-negative result).
Human CMV is a herpes virus. It is ubiquitous, species specific, and spread by close human contact. Primary infection may be acquired through different transmission routes and in different periods of life (e.g., congenital, perinatal, and postnatal infections). Serologic diagnosis of CMV infection
relies on the detection of IgG and IgM antibodies. CMV IgM appears within 2-4 weeks and persists for several weeks. In addition, CMV IgM may reappear during secondary CMV infection. CMV IgG can be detected typically after 4 weeks and persists for years to life. Unequivocal diagnosis of CMV primary infection is achieved by documenting a CMV IgG seroconversion on acute-convalescent pairs of serum samples.
Aids in the diagnosis of mononucleosis-like illness in immunocompetent patients.
Normal range: Negative.
Serology reporting is done as qualitative (negative, equivocal, or positive) based on the cutoff values established by manufacturer-specific clinical trials. A negative result, however, does not always rule out acute CMV infection.
A diagnosis of recent or acute CMV is considered probable (though not definite) in the following circumstances:
▼ The detection of CMV-specific IgM antibodies (suggesting recent seroconversion)
▼ A fourfold or greater increase in CMV-specific IgG titers in paired specimens obtained at least 2-4 weeks apart
Screening of the general population should not be performed. The positive predictive value depends on the likelihood of the virus being present. Testing should only be performed on patients with clinical symptoms or when exposure is suspected. Diseases such as Epstein-Barr viral syndrome, toxoplasmosis, and hepatitis may cause symptoms similar to CMV infection and must be excluded before confirmation of diagnosis.
The IgM response may not be detectable in the very early stage of the infection or if the patient is immunocompromised. If clinical exposure to CMV is suspected despite a negative finding, a second sample should be collected and tested after 1 or 2 weeks.
Poliovirus, coxsackie viruses (A and B), and echoviruses are enteroviruses (EVs). As the name implies, EVs most commonly replicate in the GI tract, and fecal-oral transmission is typical. Most clinical manifestations of EV infection are outside the GI tract. EV infection is most commonly considered in children who present with signs and symptoms of aseptic
meningitis in summer months. EVs also cause a severe sepsis syndrome in neonates (<2 weeks of age), pleurodynia, myocarditis and cardiomyopathy, and respiratory and oral mucosal diseases. Except for neonatal sepsis syndrome and endemic or vaccine-related poliomyelitis, EV infection is usually followed by complete recovery.
This test is used to detect viral infections caused by EVs. A number of different cell lines are susceptible to EV infection. Different EVs show differing infectivity for specific cell lines, so a number of different lines are typically inoculated for EV isolation.
Tube cultures may be incubated for up to 4 weeks before signing out as negative.
CSF cultures are usually positive within 7 days.
Stool cultures, or other specimen types with higher concentrations of virus, are often positive within several days.
Specimens should be collected within the 1st week after onset of symptoms.
Specimens should be collected according to general recommendations for virus culture of the specimen type. For patients with aseptic meningitis, CSF should be transported to the laboratory on wet ice (4°C). Submission of stool for viral culture may improve the detection of EV central nervous system (CNS) infection.
Expected results: Negative.
Positive and negative results should be interpreted in the context of clinical signs and symptoms.
Positive results: Supports infection by enterovirus.
Negative results: Enteroviral infection is less likely, but cannot be confidently ruled out. Submission of multiple specimens and sampling different patient sources may improve detection.
Submission of specimens >7 days after onset of acute infection is associated with decreased sensitivity. Cell culture is negative in 25% or more of patients presenting with typical EV infection. EVs may grow slowly in culture. Coxsackie A isolates grow poorly in culture; sensitivity for detection is fairly low. Commercially available RT-PCR methods have emerged as the most sensitive and specific tests for the detection of EV aseptic meningitis.
The enterovirus molecular assay amplifies nucleic acid by nucleic acid sequence-based amplification (NASBA) or reverse transcription real-time PCR (rtRT-PCR) for the qualitative detection of enterovirus RNA in
cerebral spinal fluid, respiratory samples, or other specimens. There is only one FDA-approved assay testing CSF samples. Other tests and specimen types have to be validated by clinical laboratories. The enterovirus is part of the larger respiratory viral/pathogen panel.
CSF is tested for patients with signs and symptoms of meningitis or encephalitis.
Respiratory specimens are tested for patients with respiratory symptoms of enterovirus such as nasal discharge, congestion and stuffiness, cough, sore throat, mild fever, mild body aches, and flu-like symptoms. Particularly, in 2014, enterovirus D68 (EV-D68) caused a nationwide outbreak of severe respiratory illness, causing severe symptoms (difficulty breathing, chest pain, wheezing, blue lips) in some children, particularly those with asthma or other lung conditions.
Stool sample is tested in case of enterovirus causing vomiting, diarrhea, and abdominal pain.
Negative results: Enterovirus RNA is not detected in the specimen; negative result in CSF should be confirmed by cell culture.
Positive results: Enterovirus RNA was detected in the specimen. This positive result does not rule out other caused of meningitis, including bacteria, mycobacteria, other viruses, and fungi.
The result should be used only in conjunction to clinical observations and other clinical information.
Common molecular assays don’t differentiate rhinovirus from enterovirus. Rhinoviruses (RVs) and enteroviruses (EVs) are different species of the Enterovirus genus, but phylogenetically closely related.
EBV quantitative and qualitative PCR assays detect the presence of EBV DNA in clinical specimens, most commonly plasma or serum but also CSF, sterile body fluids, and respiratory specimens. Normal adults usually do not have detectable EBV DNA in their plasma/serum, although normal adults previously infected with EBV will have low levels of EBV DNA in their lymphocytes.
Monitoring the level of viral reactivation and/or disease activity, particularly in the posttransplant and chemotherapy patients.
In diagnosis, prognosis, prediction, and prevention of diseases as mononucleosis, lymphoma, sarcoma, and carcinoma.
Monitoring EBV viral load in patients with infectious mononucleosis, allogeneic transplant, and nasopharyngeal carcinoma.
The EBV DNA level in healthy carriers is low and restricted to the cellular compartment of the blood. The high level is characteristic of EBV-related disease.
Patients with active infection or EBV-related cancer tend to have high levels of EBV DNA in the plasma or serum.
Positive result in asymptomatic patients may be due to recent infection or viral shedding.
▼ Detected: Above the upper limit of detection.
▼ Detected not quantified: Result above limit of detection but below limit of quantitation.
▼ Detected quantified: Expressed in copies/mL and/or log10copies/mL.
Currently, there is no international standard available for calibration of this assay. Therefore, caution should be taken when interpreting results obtained by different laboratories or assay methodologies.
EBV is the major cause of infectious mononucleosis (IM) and is a widely disseminated herpesvirus that is spread by intimate contact between susceptible persons and EBV shedders. EBV spreads primarily via passage of saliva. The virus can persist in the oropharynx of patients with IM for up to 18 months following clinical recovery. EBV has also been isolated in both cervical epithelial cells and male seminal fluid, suggesting that transmission may also occur sexually. This test comprises four serologic markers: EBV-NA (nuclear antigen IgG), EBV-VCA (viral capsid antigen) IgG and IgM, infectious mononucleosis antibody, and EBV-EA IgG (early antigen IgG).
Normal range: Negative
Tests for EBV:
▼ IgG-VCA: Indicates past infection and immunity. May be present early in illness, usually before clinical symptoms are present. Detected at onset in 100% of cases; only 20% show a fourfold increase in titer after visiting a physician. Decreases during convalescence but detectable for many years after illness; therefore, not helpful as the sole test for establishing diagnosis of IM.
▼ IgM-VCA: Detected at onset in 100% of cases; high titers present in serum 1-6 weeks after onset of illness, start to fall by the 3rd week, and usually disappear in 1-6 months. Sera are often taken too late to be detected. It is almost always present in active EBV infection and, thus, most sensitive and specific to confirm acute IM. May be positive in other herpesvirus infections (especially CMV); therefore, confirmation with IgG and EBV-NA assays is recommended.
▼ Early antigen: IgG antibodies to early antigen are present at the onset of clinical illness. There are two subsets of EA IgG: anti-D and anti-R. The presence of anti-D antibodies is consistent with recent infection, since titers disappear after recovery; however, their absence does not exclude acute illness because the antibodies are not expressed in a significant number of patients. Anti-R antibodies are only occasionally present in IM. Early antigen anti-D titers rise later (3-4 weeks after onset; is transient) in course of IM than AB-VCA and disappear with recovery; combined with IgG-VCA suggests recent EBV infection; only found in 70% of patients with IM due to EBV. High titers are found in nasopharyngeal carcinoma due to EBV. Early antigen anti-R antibodies occur rarely in primary EBV infection, 2 weeks to months after onset, and may persist for a year; more often in atypical or protracted cases. No clinical significance; high titers are found in chronic active EBV infection or Burkitt lymphoma.
▼ Epstein-Barr nuclear antigen: Last antibodies to appear and are rare in the acute phase; develops 4-6 weeks after onset of clinical illness and rises during convalescence (3-12 months) and persists for many years after illness. Absence when IgM-VCA and anti-D are present implies recent infection. Appearance early in illness excludes primary EBV infection. Appearance after previous negative test evidences recent EBV infection.
Diagnosing IM in patients with suspected IM and a negative heterophile test. Also aids in diagnosis of other suspected illnesses associated with EBV infection.
See Table 3-1.
EBV serology testing should not be performed as a screening procedure for the general population. The predictive value of a positive or negative result depends on the prevalence of analyte in a given patient population.
Testing should only be done when clinical evidence suggests the diagnosis of EBV-associated infectious mononucleosis.
TABLE 3-1. Interpretation of Epstein-Barr Virus (EBV) Serologic Status
Specific Antibody Response
Antibodies to EBV have been demonstrated in all population groups with a worldwide distribution; approximately 90-95% of adults are eventually EBV seropositive. EBV acquired during childhood years is often subclinical; <10% of children develop clinical infection despite the high rates of exposure.
The false-negative rates are highest during the beginning of clinical symptoms (25% in the 1st week; 5-10% in the 2nd week, 5% in the 3rd week).
Approximately 10% of mononucleosis-like cases are not caused by EBV. Other agents that produce a similar clinical syndrome include CMV, HIV, toxoplasmosis, HHV-6, hepatitis B, and possibly HHV-7.
Escherichia coli strains associated with enterohemorrhagic GI infections produce Shiga toxin and are most commonly, but not exclusively, associated with E. coli O157:H7 strains. Enterohemorrhagic E. coli O157:H7 gastroenteritis commonly presents with abdominal pain with vomiting and diarrhea. Stool may become bloody with signs of colitis. In most patients, the symptoms resolve within a week. Rare patients, usually the elderly or very young patients, develop hemolytic uraemic syndrome (HUS) with onset commonly occurring 7 days or more after onset of diarrheal symptoms.
This culture is used to diagnose GI infection caused by STEC. (Stool may be tested directly for the presence of Shiga toxin as an alternative to culture isolation). Special medium (sorbitol-MacConkey agar) is used to screen stool. Suspicious isolates are confirmed by serotyping and/or Shiga toxin production. E. coli O157:H7 strains are almost all sorbitol negative.
Turnaround time: 24-48 hours. Additional time is needed for positive cultures to confirm final identification.
Specimens are collected and transported according to recommendation for routine stool culture.
Expected results: Negative.
Negative results: Infection is unlikely, but a single negative culture does not rule out infection by enterohemorrhagic E. coli.
Positive results: A positive test indicates infection by E. coli O157:H7 in patients with a compatible clinical presentation.
Cultures are usually positive only in acute, early infection. The use of stool culture for evaluation of patients with HUS is limited. The use of sorbitol-MacConkey agar is not sensitive for the detection of non-O157 Shiga toxin-producing strains of E. coli; alternative testing methods should be used in areas where toxigenic non-O157 strains are prevalent or during outbreaks caused by non-O157 strains. Antibiotic therapy of E. coli O157:H7 infection is not routinely recommended; treatment may induce Shiga toxin production and increase disease severity.
Assays for Shiga toxin genes, stx1 and stx2, and E. coli O157 serotype using stool samples are part of the high multiplexed gastrointestinal molecular panels (see below).
In addition, a low-plexed assay that detects Shiga toxin-producing E. coli directly from a patient’s stool sample, the Shiga Toxin Direct Test is a sample-to-result, automated assay that detects stx1 and stx2 genes and identifies the hypervirulent serotype O157.
Patients with symptoms of gastroenteritis: severe stomach cramps, diarrhea (often bloody), and vomiting. Accurate, rapid identification of STEC,
particularly of E. coli O157:H7, is critical for patient management and disease control.
Positive for Shiga toxin 1 gene (stx1), Shiga toxin 2 gene (stx2), or both: genes for Shiga toxin 1, Shiga toxin 2, or both were detected by PCR, indicating the likely presence of a Shiga toxin-producing E. coli such as O157:H7.
Positive for E. coli O157:H7: E. coli O157:H7 DNA was detected.
Negative for Shiga toxin genes: Shiga toxin genes not detected by PCR, suggesting that a Shiga toxin-producing E. coli, such as O157, is not present.
Negative for E. coli O157:H7: E. coli O157:H7 DNA was not detected.
If positive for Shiga toxin genes but negative for E. coli O157:H7: Positive for non-O157 STEC.
Clinical laboratories should submit all STEC isolates and Shiga toxin-positive broths to a public health laboratory for additional testing.
Laboratories that use direct molecular methods for detection and identification of Shiga toxin-producing E. coli and E. coli O157:H7 might not have capability to grow the organism to provide isolates for further testing.
New nucleotide variants in the primer/probe regions may lead to false-negative molecular results.
Collaboration with microbiology laboratories and state laboratories is necessary.
The presence of fecal leukocytes is an indication of an inflammatory process of the colon, including colitis caused by invasive enteric pathogens. A number of GI infections are typically associated with the presence of fecal leukocytes: infections caused by Shigella spp., Salmonella spp., Campylobacter spp., Yersinia spp., enteroinvasive E. coli, and C. difficile and amebic dysentery.
This test is used to detect leukocytes in stool. A fecal leukocyte examination may be indicated for patients with a clinical diarrheal syndrome and signs of colitis. A fixed smear or wet mount of diarrheal stool is stained with methylene blue and examined for the presence of PMNs using a high-power objective. Fecal lactoferrin may be ordered as a surrogate marker of activated fecal granulocytes.
Turnaround time: <24 hours.
Stool is collected according to recommendations for stool culture and transported to the laboratory within 2 hours.
Expected results: Negative.
Negative fecal leukocyte examination does not rule out significant bacterial enteric infection.
Positive results support a diagnosis of invasive gastrointestinal infection or noninfectious colitis. Enteroinvasive GI infections are usually associated with 3+ to 4+ fecal leukocytes (1-4 PMN/HPF or >5 PMN/HPF) with sensitivity >50% for specimens with results of 3+ or greater. The positive predictive value increases with increasing numbers of PMN/HPF.
Significant infections caused by a number of enteric pathogens, including Vibrio spp., enterohemorrhagic E. coli, and viral agents, do not show an increase in fecal leukocytes. An increase in fecal leukocytes is not specific for infection and may be caused by other conditions, such as inflammatory bowel disease.
Francisella tularensis is a slow-growing, fastidious, gram-negative bacillus capable of producing severe infection, including localized and systemic disease. Infections have typically been acquired by zoonotic tick-borne transmission or direct contact. The common reservoir for organisms includes rabbits, rodents, deer, squirrels, and other wild mammals. Domestic animals may harbor the organism. This organism is easily transmissible, so it is critical that the laboratory be informed whenever tularemia is suspected. Typical disease syndromes include glandular, oculoglandular, and ulceroglandular tularemia; oropharyngeal tularemia; typhoidal tularemia; and pneumonic tularemia. There is great concern regarding the use of this organism for a bioterror-related attack.
This culture is used to isolate F. tularensis from clinical specimens.
▼ Specimens for F. tularensis isolation should be inoculated onto cysteine-enriched agar media. Blood-cysteine-glucose agar is recommended; most clinical isolates will grow on chocolate, Thayer-Martin, and nonselective buffered charcoal-yeast extract (BCYE) agar. Enriched broth media, such as thioglycollate broth, should also be inoculated. Blood agar and MacConkey agar are typically inoculated for clinical specimens for isolation of other possible pathogens.
▼ Because of the risk of laboratory-acquired infection and because isolation of F. tularensis may represent a sentinel event in a bioterror attack, most clinical microbiology laboratories limit the workup of suspected isolates to simple tests to rule out suspicious colonies, referring isolates that fail to “rule out” to their local public health laboratory for
identification and further characterization. Final results for testing, therefore, may be delayed compared to common bacterial isolates.
Turnaround time: Isolation and a preliminary identification are usually available within 3-6 days. Additional time is required for transfer to the local public health laboratory, confirmation of identification, and further testing.
Lymph node aspirate, ulcerative lesions, sputum, BAL, or other localized specimens, depending on the clinical presentation, are usually submitted for diagnosis.
Culture of multiple specimens from different infected tissues may improve detection.
Blood cultures are recommended for evaluation of patients with suspected tularemia.
Serologic testing is recommended for diagnosis in patients with suspected tularemia.
Expected results: Negative. After the acute phase of infection, cultures may become negative. Tularemia cannot be confidently ruled out by negative cultures.
Positive results: Isolation of F. tularensis is diagnostic of tularemia. Tularemia is a reportable disease; positive cultures must be reported to the local department of health.
Because F. tularensis organisms are tiny and faintly staining, direct detection by Gram stain of clinical specimens is uncommon. Cultures late in infection may be negative. Serologic diagnosis may be helpful in patients with disease consistent with tularemia, but negative cultures.
▼ The diagnosis of tularemia may not be entertained until after the most acute phase of illness, a time when cultures are less likely to be positive.
▼ Clinicians may fail to request specific cultures for tularemia or alert the laboratory of their clinical suspicion.
(1-3)-β-D-glucan (BG) is a cell wall component of most fungi, except Zygomycetes and Cryptococcus species. BG has been used as a biomarker for IFIs, including candidemia and Pneumocystis pneumonia; an FDA-approved test for quantitative BG detection is available. In patients with IFI or Pneumocystis pneumonia, significant levels of BG may be detected in the serum significantly earlier than clinical signs and symptoms or detection of infection by laboratory or imaging studies. Decreasing BG levels have been associated with treatment success.
Blood samples are collected and transported according to standard protocol. Samples are allowed to clot and serum separated for testing (minimum 0.5 mL).
BG testing may be submitted for early evaluation of patients at risk for IFI or Pneumocystis pneumonia or to monitor the effectiveness of therapy.
Expected results: Not detected.
▼ For IFI in neutropenic patients, BG levels ≥80 pg/mL are consistent with emerging or active infection (sensitivity approximately 65%; specificity approximately 95%).
▼ For Pneumocystis pneumonia, a higher cutoff value (≥100 pg/mL) has been recommended, yielding sensitivity approximately 95% with approximately 99% specificity.
Indeterminate results: Detectable BD levels below cutoff values do not establish or reject a diagnosis. Repeat testing is recommended.
Negative results: Fungal infection or Pneumocystis pneumonia is unlikely, but infection with Cryptococcus species or Zygomycetes is not ruled out.
The BG assay is not specific for any specific fungal pathogen. Additional testing is required for identification of the infecting species. Evaluation of patients at risk for IFI or Pneumocystis pneumonia should not be evaluated by BD as the sole diagnostic testing. Fungal culture, wet mount, imaging studies, histopathology, and other relevant diagnostic evaluations should be performed as relevant.
Galactomannan is an Aspergillus antigen that may be detected in the serum of patients with invasive aspergillosis (IA). The most sensitive galactomannan detection uses monoclonal specific antibodies in an EIA format. Detection of Aspergillus galactomannan has been shown to provide good sensitivity and specificity for IA. Galactomannan testing may improve patient management of patients at risk for IA because culture and histopathologic methods have limited sensitivity for specific detection and Aspergillus culture isolates may represent contamination or patient colonization.
Serum is most commonly collected for testing; standard laboratory procedures for collection and transport are used. Other specimens may be acceptable and are collected and transported according to instructions of the assay manufacturer.
Specimens may be collected from patients at risk for invasive aspergillosis. Sequential testing may improve detection in patients with emerging AI. Testing is not recommended for patients receiving antifungal prophylaxis because this treatment markedly decreases sensitivity of the assay.
Expected result: Not detected.
Positive results: Positive results support a diagnosis of invasive aspergillosis, but two consecutive positive results are required for a positive patient evaluation. A second specimen should be collected, before initiation of antifungal treatment, from patients to confirm initially positive galactomannan results. False-positive results may be caused by cross-reactive antigens from nonaspergillus species. Positive reactions, even persistently positive reactions, should be interpreted in the context of clinical presentation and other signs and symptoms.
Negative results: The probability of invasive Aspergillus is reduced, but AI is not excluded. Repeat testing in high-risk patients may improve detection of early AI. False-negative reactions may be caused by collection of specimens after initiation of antifungal treatment, low fungal load in serum (e.g., localized infection), high galactomannan antibody titers, or other factors.
Sensitivity of assays may be limited early in active infection. False-positive results may be seen in up 18% of patients without AI. Attempts to confirm positive results by culture or evaluation of patients at risk for invasive aspergillosis should not be evaluated with galactomannan as the sole diagnostic testing. Fungal culture, wet mount, imaging studies, and histopathology should be attempted to minimize the possibility of false-positive diagnosis. Histopathology and other relevant diagnostic evaluations should be performed as relevant.
Fungal cultures are indicated when clinically significant fungal infection is suspected. Symptomatic fungal infections commonly can be characterized as follows:
▼ Superficial (skin/nail/hair)
▼ Subcutaneous (chromoblastomycosis, mycetoma, phaeohyphomycotic cyst, sporotrichosis)
▼ Systemic mycosis (e.g., coccidioidomycosis)
▼ Opportunistic mycosis (e.g., aspergillosis)
Fungal cultures are used as the most sensitive routine laboratory method for detection of fungal infections.
▼ The media inoculated vary, depending on the specimen submitted and type of pathogen suspected.
▼ Direct examination, such as wet mount or calcofluor white staining, should be performed for most specimen types; see Fungal Wet Mount. Specimens for routine fungal culture are inoculated onto nonselective media, such as BHI or Sabouraud dextrose-BHI agar. For specimens likely to be contaminated, selective media, such as inhibitory mold agar, are inoculated. An enriched medium, such as BHI blood agar, is inoculated to improve recovery of dimorphic fungal pathogens.
▼ Special media may be inoculated for some types of specimens or suspected pathogen, like Bird (niger) seed agar for C. neoformans, chromogenic agar for differentiation of Candida isolates, or dermatophyte test medium for dermatophytes. If M. furfur is suspected, medium supplemented with a source of long-chain fatty acids (like olive oil) is inoculated.
▼ Inoculated media are typically incubated at 25-30°C in room air for up to 4 weeks. Cultures for isolation of systemic, dimorphic pathogens may be incubated at 35-37°C. Cultures for fastidious pathogens are incubated for up to 8 weeks.
▼ Media inoculated for aerobic bacterial culture will support the growth of the common yeast pathogens, Candida species, so specific culture for yeast is not usually required.
See Blood Culture, Fungal for information regarding the detection of fungemia.
▼ Cultures for yeast are incubated for 7 days. Routine fungal cultures are incubated for up to 4 weeks. Cultures for systemic dimorphic pathogens are incubated for up to 8 weeks. Additional time is needed for isolation and identification of isolates.
Specimens are collected using sterile technique and transported in a sterile container within 2 hours. Store specimens at 4°C if transport will be delayed.
Most specimens for fungal culture are collected following standard specimen collection instructions. Special collection:
▼ Collection of specimens using swabs is not recommended, except for samples from mucous membranes for the diagnosis of candidiasis.
▼ Pluck multiple hairs (10 or more) and scrape scalp from involved areas.
▼ Wipe affected nail with 70% alcohol. Submit nail clippings and scrapings from beneath nails in a clean container.
▼ Wipe affected skin lesions with 70% alcohol. Scrape the advancing margin to remove superficial cells and keratinized material; submit in a clean container.
Expected results: No growth.
Positive results: Positive cultures must be carefully interpreted to ensure that endogenous fungal flora and culture contaminants are recognized.
The results of fungal cultures may not be available when decisions regarding therapy of acute infection are required. Empiric therapy may be required.
Common pitfalls: Isolation of endogenous Candida species or environmental mold contaminants may result in unneeded treatment.
Clinical information, such as travel history, immune status, and animal exposure, should be included on the requisitions for fungal cultures.
Histopathologic testing and immunologic testing are important methods for diagnosis of IFIs.
A diagnosis of vaginal and oral candidiasis (thrush) can be made reliably by direct microscopic examination (Gram stain or wet mount) of scrapings of mucosal surfaces without need for fungal culture.
Detection of antigen or fungal products, such as cryptococcal antigen, histoplasma antigen, β-D-glucan, or galactomannan, may be useful for diagnosis.
The India ink wet mount is less sensitive than cryptococcal antigen testing for meningitis caused by C. neoformans.
A direct examination for fungal elements may provide a rapid detection of fungal infection and is recommended for most types of specimens submitted for fungal culture.
This test is used for the direct detection of fungal forms in patient specimens. The specimen is processed to form a liquid suspension of the patient sample.
▼ Solid specimens, such as tissues, should be minced to facilitate suspension.
▼ The specimen may be suspended in saline or a 10% KOH solution. KOH may improve liquefaction of the specimen and also lyses host cells and keratin; fungal cells are resistant to KOH digestion.
▼ A cover slip is added for examination with regular or phase-contrast light microscopy.
▼ Calcofluor white, a fluorogenic dye that binds to specific polysaccharide bonds found in fungal cell walls, may be added to the KOH solution to improve microscopic visualization of fungi.
Turnaround time: 24 hours.
Specimens should be collected and transported according to guidelines for fungal culture of the specimen type.
Expected results: Negative.
Positive results: Provides evidence of fungal infection. Fungal elements may be characterized on the basis of morphology (e.g., budding yeast, aseptate hyphae, conidia-forming structures consistent with Aspergillus species).
Negative results: Fungal infection is not ruled out by a negative wet mount examination.
The morphology of objects must be examined carefully to exclude artifacts or nonspecific absorption of calcofluor dye to nonfungal objects, such as capillaries.