After completing this chapter, the reader should be able to
Describe the basic methods that may be used in the diagnosis of invasive fungal infections
Discuss the laboratory tests that are commonly used in the diagnosis of common viral infections such as influenza, herpes simplex virus, cytomegalovirus, and respiratory syncytial virus
Discuss the laboratory tests that are commonly used in the diagnosis of human immunodeficiency virus; describe the laboratory tests that are commonly used in the assessment and monitoring of patients with human immunodeficiency virus infection
Discuss the laboratory tests that are commonly used in the diagnosis of infections due to Mycobacterium tuberculosis and nontuberculous mycobacteria
The assessment, diagnosis, and treatment of a patient with a fungal, viral, or mycobacterial infection can be challenging tasks for most clinicians. This may be partly due to the nonspecific presentation and clinical recognition of infectious processes; the continuously changing taxonomy and diagnostic procedures of infecting organisms; the emergence of multidrug resistant pathogens such as Candida auris and Aspergillus tanneri; the understanding of drug resistance mechanisms and interpretation of susceptibility characteristics; and the recognition of the potential role of older and newer therapeutic agents in the prevention and/or treatment of these infections.1–8 It is important to note that diagnostic tests for many infectious diseases, particularly the diagnosis of COVID-19 caused by a coronavirus (ie, SARS-CoV-2) and human immunodeficiency virus (HIV) infection, are continuously evolving to reflect technological advances in laboratory procedures.8,9 Nucleic acid amplification test (NAAT), sequencing-based identification methods, and/or proteomics (eg, matrix-assisted laser desorption/ionization-time-of-flight mass spectroscopy [MALDI-TOF MS]) are transforming diagnostic microbiology and becoming rapidly accepted detection techniques of fungal, viral, and mycobacterial pathogens.10–13
This chapter describes the laboratory tests commonly used in the diagnosis of common infections due to fungi, mycobacteria, and viruses. Laboratory tests used in the diagnosis of viral hepatitis are addressed in Chapter 15.
Fungi are classified as one of the six kingdoms of life. There are approximately 500 named species of fungi that are known to cause infection in humans and other vertebrate animals.14 Approximately 50 fungal species are associated with infections in healthy subjects and most fungal infections occur in immunocompromised or debilitated patients by organisms that are part of the normal human flora. However, an increasing number of serious and life-threatening opportunistic infections are being caused by ubiquitous environmental molds.
Some of the most challenging and frustrating aspects of diagnostic medical mycology are the terminology, taxonomy, classification, and nomenclature of fungi.14,15 For example, the correct name for a species of fungi is that which was published earliest and met the requirements in the International Code of Botanical Nomenclature for algae, fungi, and plants (http://www.iapt-taxon.org/nomen/main.php). Since January 1, 2013, the concept of “One Fungus/One Name” has been applied, eliminating the use of dual names (eg, anamorph and teleomorph names of fungal species). All subsequent names are considered synonyms; however, exceptions do exist, particularly when a later name is more commonly used than the earlier name or if research requires a species to be transferred to a different genus. Changes have occurred within the kingdom fungi (eg, the phylum Zygomycota is no longer recommended because of polyphyletic characteristics). Because of these issues, the reader is referred to the latest editions of standard microbiology textbooks and reference manuals (eg, Manual of Clinical Microbiology, American Society for Microbiology Press) and the following links (http://www.mycobank.org/; https://mycology.adelaide.edu.au/; http://www.indexfungorum.org/names/names.asp; https://www.fungaltaxonomy.org/; http://www.clinicalfungi.org) for more detailed information and updates on taxonomy and classifications of fungi.14,15 In addition, a glossary of common mycological terms is often included.
Fungi are eukaryotic and can be either unicellular or multicellular organisms. Fungi have cell walls composed mainly of chitins, glucan, and mannam with a membrane-bound cell nucleus with chromosomes. The dominant sterol in the cytoplasmic membrane of fungi is ergosterol, compared with cholesterol in mammalian cells. These organisms are heterotropic (eg, require exogenous energy sources) and can reproduce by either asexual (involving mitosis) or sexual (involving meiosis) cell division. Fungi may exist in a morphologic form that results from sexual reproduction (teleomorph, or perfect state) and a form that results from asexual reproduction (anamorph, or imperfect state), in which each of the forms has its own name (eg, the sexual form of Scedosporium apiospermum complex is Pseudallescheria boydii). The use of separate anamorph and teleomorph species names ended in January 2013, and all legitimate names for the species can be used. It will likely take a significant period (eg, a decade or more) before these nomenclatural changes of fungal organisms achieve stability and acceptance.
Fungi have traditionally been categorized into mold, yeast, or dimorphic fungi based on morphologic and structural features (Table 19-1).14–18Molds (or moulds) are long, cylindrical, and threadlike (filamentous) fungi that form multicellular mycelium or thallus, an intertwined mass of branching hyphae (tube-like extensions or filament-like cells), with septa (having cross walls; being septate) or pauciseptate. An asexual spore (conidium) is produced on conidiophores, a specialized hyphal structure that serves as a stalk, and macroconidia and microconidia may be present. Thermally, monomorphic molds can be divided into four groups: (1) Mucormycetes, (2) dematiaceous fungi, (3) dermatophytes, and (4) hyaline hyphomycetes. Mucormycetes have broad hyphae that are almost nonseptate, with asexual spores (sporangiospores) formed by cleavage in a saclike structure (sporangium). The most common Mucormycetes observed in the clinical laboratory are from the order Mucorales, which are associated with severe fungal infections referred to as mucormycosis. Two genera from the order Entomophthorales, Basidiobolus and Conidiobolus, are less commonly observed Mucormycetes but are responsible for subcutaneous infections in otherwise healthy individuals. Dematiaceous fungi produce dark-colored colonies of olive, brown, gray, or black due to melanin pigment in the cell walls. Some of the common infections associated with dematiaceous fungi include chromoblastomycosis, phaeohyphomycosis, and mycetoma. Dematiaceous fungi also cause tinea nigra and black piedra. Dermatophytes are most often associated with superficial fungal infections (tinea or ringworm) of the skin, hair, and nails. These filamentous fungi colonize the outermost layer of the skin and digest keratin as a source of nutrients. The three genera (Microsporum, Trichophyton, and Epidermophyton) are differentiated by their conidium formation (macroconidia or microconidia). Hyaline hyphomycetes are colorless, septate hyphae molds that produce conidia that may be either colorless or pigmented. Coccidioides immitis and Coccidioides posadasii are known pathogens from this group whereas most other organisms cause opportunistic infections in immunocompromised patients.
Categorization of Selected Medically Important Fungia
aRefer to reference 15 for further details and other organisms not listed.
bCoccidioides immitis is often placed with dimorphic fungi (listed in this table under hyaline hyphomycetes because it does not produce yeast-like colonies or cells at 35°C to 37°C on routine mycology agar).
cSeveral Candida spp. have had a change in nomenclature to names of teleomorphs (in parenthesis): C lusitaniae (Clavispora lusitaniae); C krusei (Pichia kudriavzevii); C kefyr (Kluveromyces marxianus); C guilliermondii (Meyerozyma guilliermondii); C lipolytica (Yarrowia lipolytica). We have kept the original Candida spp. names because most laboratories and medical providers are more familiar with this nomenclature of fungi.
dThe phylum Glomerulomycota (Mucormycetes) has replaced the former term Zygomycetes. Mucormycetes is divided into two subphylums, Mucormycotina and Entomophthoromycotina. The subphylum Mucormycotina contains the order Mucorales, which includes the genera Rhizopus, Mucor, Rhizomucor, and Lichtheimia (formerly Absidia). The subphylum Entomophthoromycotina contains the order Entomophthorales, which includes genera Basidiobolus and Conidiobolus.
eSporothrix schenckii grows as a dematiaceous mold when incubated at 25°C to 30°C but is yeast like at 35°C to 37°C. It is commonly categorized among dimorphic fungi but is often considered a dematiaceous mold.
Yeasts (and yeast-like organisms) appear as round or oval cells that are unicellular and generally reproduce at their surface by budding (blastoconidia). Some produce pseudohyphae (an elongated chain of cells, like a chain of sausages, resembling hyphae; however, borders between cells are delineated by marked constrictions), whereas others have true hyphae (tend to be straighter and without constrictions at the septa), which may be septate or without septa (aseptate). Ascospores, a sexual spore in a saclike structure (ascus), are produced by only some yeast. Yeasts are the most frequently encountered fungi in the clinical microbiology laboratory and are considered opportunistic pathogens. Candida spp. and Cryptococcus spp. are among the most common yeasts causing fungal infections. Yeasts are not considered a formal taxonomic group but a growth form of unrelated fungi (members of the phyla Basidiomycota and Ascomycota).
Thermally, dimorphic fungi have two distinct morphologic forms in which their growth forms can change from a multicellular mold (in their natural environment or when cultured at 25°C to 30°C) to budding, unicellular yeasts (during tissue invasion or when cultured at 35°C to 37°C). Medically important dimorphic fungi include Histoplasma capsulatum, Blastomyces dermatitidis, and Paracoccidioides brasiliensis. Each of these fungi is considered pathogenic and must be handled with caution in the clinical laboratory.
The Identification of Fungi
The following section provides a summary of the common methods currently used in diagnostic testing of medically important fungi.15–18 Fungal identification has traditionally been based on morphologic characteristics, such as the color of the colonies, the size and shape of cells, the presence of a capsule, and the production of hyphae, pseudohyphae, or chlamydospores. Culture remains the “gold standard” in most clinical microbiology laboratories and is the only method that allows subsequent susceptibility testing. NAAT, molecular characteristics, and/or proteomics (MALDI-TOF MS) are rapidly gaining a larger routine role in fungal identification, particularly when morphology-based identification is atypical, confusing, or not helpful (eg, organisms that fail to sporulate) and in cases in which precise identification is required (eg, epidemiologic studies).18–22 Because no one test is perfect, it is often necessary to perform several diagnostic tests (both morphologic and genotypic methodologies) to maximize the accuracy of fungal identification. Laboratory diagnosis of fungal infections includes direct microscopic examination, morphologic identification, isolation in culture, and use of non–culture-based methods, such as antigen and/or antibody detection, 1,3-β-d-glucan detection, molecular and nonmolecular diagnostic testing, and MALDI-TOF MS.10,14,18–27 As with all types of infections, appropriate biological specimens need to be selected, collected, and transported to the laboratory for immediate processing.16 Because different fungi are capable of causing infection at a number of anatomic sites, specimens from the site of infection and peripheral blood specimens should be considered and submitted for culture and microscopic examination. Communication with the laboratory regarding the clinical infection and suspected fungi is important and may be useful for determining how best to process specimens safely and efficiently, including pretreatment and staining procedures, selection and incubation of media, and choice of additional diagnostic testing. Early identification of the infecting fungal pathogen may have direct diagnostic, epidemiologic, prognostic, and therapeutic implications.
The first step is usually identifying yeast-like fungi (pasty, opaque colonies) from molds (large, filamentous colonies that vary in texture, color, and topography). Drawings, color plates, and brief descriptions found in standard textbooks can serve as guides and assist in the preliminary identification of fungi seen on direct microscopic examination of clinical specimens.14–17 Microscopic examination of the clinical specimen can delineate morphologic features (Table 19-2) and often provides preliminary identification of many fungi (eg, Aspergillus spp., Mucormycetes, dematiaceous molds). Microscopic morphology can often provide definitive identification of a mold whereas the addition of biochemical tests, serology, nucleic acid-based molecular testing, and/or MALDI-TOF MS are usually needed for identifying the genus and species of most yeast and yeast-like fungi. Direct microscopic examination of properly stained clinical specimens and tissue sections is usually the most rapid (within a few minutes or hours) and cost-effective method for a preliminary diagnosis of fungal infection. In addition, microscopic detection of fungi can assist the laboratory in the selection of media and interpretation of culture results.
Laboratory Testing and Characteristic Features Used in the Diagnosis of Selected Opportunistic and Pathogenic Fungi
Round to oval budding yeasts (3–6 µm in diameter), singly, in chains, or in small loose clusters; true hyphae (no or slight constrictions at the septa) and pseudohyphae (5–10 µm in diameter; chains of elongated blastoconidia) when invading tissues; blastoconidia develop along the sides of either type of hyphae
Candida glabrata slightly smaller (2–5 µm in diameter) than other species and does not produce any hyphal forms
Variable morphology; colonies usually pasty, white to tan, and opaque; may have smooth or wrinkled morphology
Clusters of blastoconidia, pseudohyphae and terminal chlamydospores in some species
Serum (1,3)-β-d-glucan (Fungitell Assay Reagent) indicated for presumptive diagnosis of invasive fungal infection and should be used in conjugation with other diagnostic procedures
EIA test for detection of Candida mannan antigen and antimannan antibody; both the antigen and antibody test should be performed together to maximize early diagnosis
Morphology on CHROMagar, corn meal agar, rapid trehalose test
Germ tube production by Candida albicans, Candida dubliniensis, and Candida stellatoidea
T2 Candida Panel
Most commonly in CSF and blood; bone marrow; catheter sites; respiratory sites; skin, mucous membrane; urine; multiple systemic sites
Spherical (football shaped) or round, budding yeasts of variable size (2–15 µm) with thin dark walls; thick capsule may or may not be present; no hyphae or pseudohyphae
Colonies are shiny, mucoid, dome-shaped, and cream to tan in color
Budding spherical cells of varying size; capsule present; no pseudohyphae; cells may have multiple narrow-based buds
LFD, LA, or EIA test for polysaccharide antigen
Tests for urease (+), phenoloxidase (+), and nitrate reductase (−)
Mucicarmine and melanin stains in tissue
Differentiation from C gattii with CGB (l-canavanine, glycine, bromthymol blue) agar India ink can demonstrate capsule in ~50% of cases on smeared specimens; this low sensitivity has resulted in antigen testing or MALDI-TOF MS as primary laboratory techniques for identification
Septate with uniform diameter (3–6 µm); dichotomously branched hyphae at 45° angles; tend to grow in radial fashion (like spokes on a wheel)
Varies with species; Aspergillus fumigates: blue-green to gray; Aspergillus flavus: yellow to green; Aspergillus niger: black with white margins and yellow surface mycelium
Varies with species; conidiophores with enlarged vesicles covered with flask-shaped metulae or phialides; hyphae are hyaline and septate
Galactomannan detection (eg, ELISA assay: Platelia Aspergillus GM antigen kit) is useful tool in the diagnosis of invasive aspergillosis, especially in hematologic malignancy and stem cell transplantation patients
Serum (1,3)-β- d-glucan (Fungitell Assay Reagent) indicated for presumptive diagnosis of invasive fungal infection and should be used in conjugation with other diagnostic procedures (eg, PCR)
Useful for chronic and allergic aspergillosis
Identification based on colony characteristics and microscopic morphology features to identify Aspergillus
Culture-based identification remains important
Molecular techniques (eg, sequencing-based and non-sequence-based PCR methods) are attractive as alternative methods (eg, Luminex xMAP technology with Aspergillus species-specific assays)
MALDI-TOF MS attractive as alternative method but requires adequate sample preparation procedure and supplementation with reference spectra
Broad, thin-walled, pauciseptate hyphae (6–25 µm) with nonparallel sides and branching irregularly, nondichotomous, and at various angles; hyphae stain poorly with GMS stain and often stain well with H&E stain
Colonies are rapid growing, wooly, or fluffy (cotton candylike), and gray-black in color
Differentiation of various genera based on presence and location (or absence) of rhizoids, nature of sporangiophores, shape of columella, appearance of an apophysis, and size and shape of the sporangia
Identification based on microscopic morphologic features
Culture results need to be evaluated along with the clinical presentation, examination of morphologic features, and/or histopathology
Round, thick-walled spherules that vary in size (20–200 µm in length); mature spherules contain small (2–15 µm in diameter) endospores; septate hyphae, barrel-shaped arthroconidia may be seen in cavitary and necrotic lesions
Great variation in morphology; at 25°C or 37°C, colonies initially appear moist and glabrous, rapidly develops a white, cottony aerial mycelium, which becomes gray-white to a tan or brownish
Hyaline hyphae with rectangular (barrel-shaped) arthroconidia separated by empty disjunctor cells
EIA can be used with urine, serum, plasma, BAL, CSF, and other body fluid samples (uses antibodies against Coccidioides galactomannan)
Various methods (ID and CF most reliable) for initial antibody screening; EIA assays also available and more often used for screening with confirmation of positive results by ID and CF
Identification by direct microscopic visualization and histopathologic examination
AccuProbe may be useful for confirmation of unknown isolates as Coccidioides species (but does not distinguish between two species of Coccidioides)
Serum (1,3)-β- d-glucan (Fungitell Assay Reagent) indicated for presumptive diagnosis of invasive fungal infection but has a limited role because of low sensitivity (44%)
Exoantigen and nucleic acid probe tests
PCR-based assay (Gene STAT.MDx Coccidioides test) for detection from clinical samples was recently approved by FDA
Small (2–6 µm in diameter) yeast-like cells of varying sizes and shapes (oval or round or cigar shaped); single or multiple elongated “pipe stem;” bud is on a narrow base
At 25°C–30°C, colonies are initially small, smooth, moist and white to pale orange to orange-gray with no cottony aerial hyphae; later, colonies become moist, wrinkled, leathery, or velvety and darken to brown or black; at 35°C—37°C, colonies are white to tan, dry, smooth, and yeast-like
At 25°C–30°C, thin or narrow, septate, and branching, with slender, tapering conidiophores rising at right angles; conidia borne in rosette-shaped clusters at the end of the conidiophores; at 35°C–37°C, variable-sized round, oval, and fusiform budding yeasts (cigar bodies)
LA test is commercially available (rarely used but could be helpful for disseminated cases, and potentially CNS sporotrichosis when culture-based diagnosis had failed)
Oval (2.5–5 µm in length) or elongated or cylindrical, curved yeast-like cells; found within histiocytes (intracellular); has visible septa and budding does not occur (reproduces by fission [arthroconidium-like])
Colonies are flat, powdery to velvet, and tan, and then produce diffusible red-yellow pigment at 25°C–30°C; at 35°C–37°C, colony is soft, white to tan, dry, yeast-like
At 25°C–30°C, smooth conidiophores with four to five terminal metulae bearing phialides; chains of short, narrow extensions connect the round to oval conidia in a “paint brush” distribution; at 35°C–37°C, arthroconidial yeast cells divide by fission and may elongate
Demonstration of thermal dimorphism
NAAT (eg, single-step or nested PCR) are attractive as alternative methods but are mainly in-house systems without standardization being available
MALDI-TOF MS attractive as alternative method requiring supplementation with reference spectra
Respiratory sites; multiple systemic sites
Cysts are round, ovoid, or collapsed crescent shaped (4–7 µm in diameter); stains should be used to visualize cyst forms for diagnosis; trophozoites are small (1–4 µm in diameter), pleomorphic forms
Serum (1,3)-β-d-glucan (Fungitell Assay Reagent) indicated for presumptive diagnosis of invasive fungal infection and should be used in conjugation with other diagnostic procedures
Useful for epidemiology studies but not for diagnosis
Direct microscopic detection using stains: methenamine silver (Gomori/Grocott), Giemsa and rapid Giemsa-like, toluidine blue, direct and indirect immunofluorescence, calcofluor white, Gram-Weigert, and papanicolaou
Several nucleic acid detection techniques (eg, real-time PCR) have been successful and commercially available outside the United States
EIA = enzyme immunoassay; MALDI-TOF MS = matrix-assisted laser desorption/ionization-time-of-flight mass spectroscopy; CSF = cerebrospinal fluid; LFD = lateral-flow device; LA = latex agglutination; ELISA = enzyme-linked immunosorbent assay; PCR = polymerase chain reaction; NAAT = nucleic acid amplification technology; ID = immunodiffusion; CF = complement fixation; BAL = bronchoalveolar lavage; CNS = central nervous system; FDA = Food and Drug Administration.
The Gram stain that is typically used for bacterial processing may also allow the detection of most fungi, especially Candida spp., because the size of the smallest fungi is similar in size to large bacteria; the presence of budding cells can also be observed. A wide range of stains is available (Table 19-3) to assist in the rapid detection of fungal elements.10,14,17,18 A common approach to wet preparations of specimens or smeared dried material is to use a 10% solution of potassium hydroxide (KOH) with or without fluorescent calcofluor white stain. The strong alkaline KOH solution digests tissue elements to allow better visualization of the fungi, while the calcofluor white stain binds to chitin and polysaccharides in the fungal cell wall, allowing it to appear white under ultraviolet light. Specific staining techniques are often used to outline morphologic features that are diagnostic and distinctive of the suspected fungal organism (eg, India ink stain for detection of a polysaccharide capsule of Cryptococcus neoformans). In suspected cases of histoplasmosis, the Giemsa or Wright stain is useful for detecting intracellular yeast cells within macrophages from blood or bone marrow specimens.
Stains Used to Enhance the Direct Microscopic Detection of Fungi
Polysaccharide capsule produced by Cryptococcus neoformans (in CSF)
Mucopolysaccharide stain showing clear halos against black background; often negative in many cases of meningitis
Calcofluor white (CFW)
Most fungi, including Pneumocystis jirovecii (cysts)
Binds to chitin in fungal cell wall and fluoresces bluish white against dark background; requires fluorescent microscope; mixed with KOH for easier and rapid observation of fungi
Dematiaceous fungi, Cryptococcus neoformans, and Cryptococcus gattii; may also be useful for Aspergillus fumigates, Aspergillus flavus, Trichosporon spp., and some Mucormycetes
Stains fungi brown to black against reddish background; demonstration of melanin or melanin-like substances in the lightly pigmented agents of phaeohyphomycosis
Giemsa or Wright
Visualization of intracellular Histoplasma capsulatum; trophic forms of Pneumocystis jirovecii; fission yeast cells of Talaromyces marneffei
Stains blue-purple (fungi and bacteria); examination of bone marrow or peripheral blood smears for disseminated disease
Gomori methenamine silver (GMS)
Most fungi in histopathologic sections; Pneumocystis jirovecii (respiratory specimens)
Detects fungi elements; however, requires specialized staining method; stains hyphae and yeast forms gray to black against a pale green or yellow background
Yeast and pseudohyphae appear gram-positive and hyphae (septate and aseptate) appear gram-negative
Commonly performed on clinical specimens; some fungi stain poorly (eg, Cryptococcus spp., Nocardia)
Gridley fungus (GF)
Most fungi in histopathologic sections
Fungi stain purplish red; filaments of Actinomycetes are not stained
Hematoxylin and eosin (H&E)
General purpose histopathologic stain; best method for visualizing host tissue reactions to infecting fungus
Stains some fungal elements violet to bluish purple in contrast to lighter background; Aspergillus spp. and Zygomycetes stain well; some fungi difficult to differentiate from background
Commercial antibodies used in the immunohistochemical diagnosis of fungal infections, especially to distinguish fungal elements on atypical appearing tissue sections
India ink, nigrosin
Cryptococcus neoformans (in CSF)
Sensitivity is <50% of meningitis cases
Nocardia (filaments are partially acid-fast and stain pink) and some isolates of Blastomyces dermatitidis
Actinomyces and other actinomycetes are negative
Cryptococcus neoformans (capsular material) and Cryptococcus gattii; cell walls of Blastomyces dermatitidis and Rhinosporidium seeberi
Histopathologic stain for mucin; capsular material stains deep rose to red; tissue elements stain yellow
Histopathologic stain for fungi, especially yeast cells and hyphae in tissue; commonly used stain by dermatopathologists
Fungal elements stain bright pink–magenta or purple against orange background (picric acid counterstain) or green background (if light green used); hyphae of molds and yeast can be readily distinguished; demonstrates double-contoured refractile wall of Blastomyces dermatitidis; Nocardia do not stain well
Potassium hydroxide (KOH)
Most fungi (more readily visible); hyaline molds and yeast appear transparent; dematiaceous molds display golden brown hyphae
Used to dissolve tissue material, allowing more visible fungal elements; can be combined with calcofluor white for fluorescence detection
Pneumocystis jirovecii (respiratory specimens: biopsy or BAL)
Stains cysts of Pneumocystis jirovecii reddish blue or dark purple against light blue background
Histopathologic stains are extremely valuable for identifying fungal elements in tissues and host tissue reactions to fungal infection.24,25 Histology laboratories commonly use stains such as hematoxylin and eosin for these general purposes. Periodic acid-Schiff and Gridley fungus stains can also assist in visualization of fungal elements, especially if debris is present in the tissue background. Special stains (eg, Gomori methenamine silver, mucicarmine, and Fontana-Masson) are useful for enhancing the detection of specific fungal elements (Table 19-3) for a histopathological diagnosis of fungal infections.15,17,18
Culture remains the gold standard for isolation and identification of fungi suspected of causing infection. Petri plates are preferred over screw-cap tubes because of the larger surface area and dilution of inhibitory substances in the specimens. However, for laboratory safety reasons, most thermally dimorphic fungi (eg, Histoplasma, Blastomyces, Paracoccidioides, T marneffei) and Coccidioides spp. are pathogenic and should be grown on slants (ie, avoid the use of Petri plates and slide culture). A variety of media are available for the isolation and cultivation of yeasts and molds (Table 19-4).15–17 Sabouraud dextrose, brain heart infusion, and inhibitory mold agars are enriched media commonly recommended to permit the growth and isolation of yeasts and molds. Several media, with (selective) and without (nonselective) inhibitory agents, should be used because no one medium is adequate for all the different types of specimens or organisms. Antibiotics such as chloramphenicol or gentamicin are included as inhibitory substances of most bacterial contaminants, whereas cycloheximide is used to inhibit saprobes and prevent the overgrowth of contaminating molds. Nonselective media (without inhibitory agents) should be used with specimens from sterile sites, and when suspected, fungi are likely to be inhibited by cycloheximide (eg, Aspergillus fumigatus, C neoformans/gattii, Lomentospora prolificans, T [Penicillium] marneffei, some Candida spp., most Mucormycetes) or by antibiotics (eg, Nocardia or other filamentous bacteria). Direct microbiological examination (outlined previously) of clinical specimens can assist in the selection of media based on specimen type and suspected pathogen. In addition, the choice of media will be influenced by the patient population, local endemic pathogens, cost, availability, and laboratory preferences.
Examples of Various Media Used for the Recovery of Fungi from Clinical Specimens
COMMENTS AND USES
Primary Media Without Antibacterials or Antifungals
Brain heart infusion (BHI) agar
Enriched media used for cultivation and isolation of all fungi; designed to enhance the recovery of fastidious dimorphic fungi more than SDA does
Sabouraud dextrose agar (SDA)
General-purpose medium that supports primary growth or sporulation and provides classic pigment and morphology
SDA, Emmons modification
Compared with SDA, Emmons Modification contains 2% (versus 4%) glucose and has a pH of 6.9–7.0 (versus pH 5.6 [slightly acidic])
Enriched media using combined ingredients of BHI and SDA; supports growth of all fungi; designed for better recovery of fastidious dimorphic fungi than does SDA
Primary Media with Antibacterials or Antifungals
Any of the above media
Usually with chloramphenicol (inhibits gram-negative and gram-positive bacteria) with or without gentamicin (inhibits gram-negative bacteria); penicillin and streptomycin have also been used; cycloheximide added to inhibit sensitive fast-growing saprophytic fungi
Inhibitory mold agar (IMA)
A selective and enriched media providing better recovery of fastidious fungi than SDA; usually contains chloramphenicol; some formulations contain gentamicin
Littman Oxgall agar
General purpose selective medium for isolation of all fungi. Oxgall restricts the spreading of fungal colonies; contains crystal violet and streptomycin to inhibit bacteria growth
Mycosel or mycobiotic agar
Selective medium containing chloramphenicol and cycloheximide primarily used for isolation of dermatophytes; can also be used for isolation of other pathogenic fungi from contaminant specimens
Dermatophyte test medium (DTM)
Screening medium for the recovery, selection, and differentiation of dermatophytes (eg, Microsporum, Trichophyton, Epidermophyton) from hair, skin, and nail (keratinous) specimens; contains chloramphenicol, gentamicin, and cycloheximide; other saprophytic fungi and Aspergillus spp. can grow on this medium (thus, it is recommended only as a screening medium)
Yeast extract phosphate agar with ammonia (Smith’s medium)
Used for isolation and sporulation of slowly growing dimorphic fungi (ie, Histoplasma capsulatum and Blastomyces dermatitidis) from contaminated specimens; contains chloramphenicol and ammonium hydroxide to suppress the growth of bacteria, molds, and yeasts
Chromogenic media used for direct and rapid differentiation of many clinically important yeast spp; contains chloramphenicol to inhibit bacteria and is available with or without fluconazole (selection of fluconazole-resistant Candida krusei); CHROMagar differentiates more Candida spp. than CAN2; useful in identifying mixed cultures of yeasts
Chromogenic media used for direct and rapid identification of Candida albicans versus other species of yeasts; useful in identifying mixed cultures of yeasts
Canavanine glycine bromothymol blue (CGB) agar
Solid medium recommended for use in qualitative procedures for selective and differential isolation of Cryptococcus gattii (ie, Cryptococcus serotypes B and C) from Cryptococcus neoformans
Cornmeal agar (CMA)
CMA with 1% dextrose used for the cultivation of fungi and differentiation of Trichophyton mentagrophytes from Trichophyton rubrum (based on pigment production); CMA with Tween 80 used for the cultivation and differentiation of Candida spp. (based on mycelial characteristics); Tween 80 promotes growth and production of red pigment by Trichophyton rubrum
Potato dextrose agar (PDA) or potato flake agar (PFA)
PDA is used to stimulate conidium production by fungi and enhance pigment production by some dermatophytes; PDA is most commonly used with slide culture technique to view morphologic characteristics; PFA used for the simulation of conidia of fungi; PFA can be made selective by including cycloheximide and chloramphenicol
Rapid sporulation agar (RSA)
Used for isolation and identification of dermatophytes; contains chloramphenicol and chlortetracycline to inhibit bacteria and cycloheximide to inhibit saprobic fungi; similar to DTM but can increase the production of conidia and improve color visualization of the isolate (bromothymol blue in the formulation)
Niger seed or bird seed agar and esculin base medium (EBM)
Selective and differential medium for isolation of Cryptococcus spp., especially C neoformans and C gattii; chloramphenicol as a selective agent; creatinine to enhance melanization of some strains of C. neoformans
aSeveral commercial sources of yeast chromogenic agar media are available.17 CHROMagar Candida and chromID Candida agar are examples of selected chromogenic agar products that are approved by the FDA for use in U.S. laboratories. Candida spp. identified with chromogenic agar differs by medium and manufacturer. Further testing is required for final identification.
Proper temperature and adequate time for incubation are necessary to optimize the recovery of medically important fungi from clinical specimens. Inoculated media should be incubated aerobically at 30°C. If an incubator at that temperature is not available, then 25°C (room temperature) can be considered. Other temperatures (eg, 35°C to 37°C for thermally dimorphic organisms) should be reserved for selected fungi that prefer a higher temperature. In general, yeasts are detected within 5 days or less, dermatophytes within 1 week, and dematiaceous and dimorphic fungi between 2 and 4 weeks. Cultures should be regularly reviewed (eg, every day the first week, every 2 to 3 days the second week, twice during the third week, once weekly thereafter) to account for the growth rates and identification of fungi. It is generally recommended that fungal cultures be incubated for 4 weeks before being considered negative (no growth for fungus). Several factors influence the length of incubation including the choice of media (eg, yeasts on chromogenic [48 hours] versus routine media [5 to 7 days]) and type of fungus suspected (eg, slow-growing dimorphic systemic fungi may need 8 weeks).
Once the organism has been cultured and isolated, the following approach has usually been conducted: (1) determine the morphology of the unknown fungus and determine if it is consistent with any of the groups listed in Table 19-1 or filamentous bacterium (some of the aerobic actinomycetes [eg, Nocardia] resemble fungi and must be ruled out) and (2) note the rate of growth, colony, and microscopic morphologies of the possible organism(s) (Table 19-2) and refer to necessary textbooks to compare descriptions, drawings, color plates, discussions of characteristics, and other test results to assist in differentiating the likely organism.15,18 In the case of yeasts and yeast-like organisms, additional testing, such as the germ tube test, biochemical testing using commercially available systems, or the urease test, may allow species identification of isolates from various body sites. Both NAAT and MALDI-TOF MS are increasingly being used to modernize clinical microbiology laboratories. These rapid, inexpensive, and accurate methods for identification of fungal organisms allow less dependency on performing time-consuming biochemical procedures and/or needing visual expertise for detection of microscopic and colonial morphology.10,18–24,26
Cell wall components of various invasive fungi have been used as diagnostic markers for antigen testing. Galactomannan is a polysaccharide component of the cell wall of Aspergillus and it is released by growing hyphae. A commercial enzyme-linked immunosorbent assay (ELISA) (Platelia Aspergillus Galactomannan Test [Bio-Rad Laboratories, Marens-La-Coquette, France]) is available to detect circulating galactomannan antigen in serum or bronchoalveolar lavage (BAL) fluid and has been shown to be an earlier diagnostic marker for invasive aspergillosis in neutropenic patients with hematologic malignancies and hematopoietic stem cell transplantation.18,27 The monitoring of antigen titers has also been shown to correlate with the response to antifungal therapy, patient survival, and autopsy findings in neutropenic patients. Other genera of fungi, including Histoplasma, Penicillium, Alternaria, Geotrichum, and Paecilomyces, have shown reactivity to the assay kit. Several causes of false-positive results have also been reported, including patients receiving specific antibiotics (eg, piperacillin-tazobactam, amoxicillin-clavulanic acid), certain foods (eg, pasta, vegetables, milk), and Plasma-Lyte A. The detection of galactomannan is also reduced in patients receiving antifungal agents active against molds and patients with chronic granulomatous disease. A nongalactomannan antigen method has recently been developed as a point-of-care (POC) test for rapid detection of invasive pulmonary aspergillosis. This rapid detection method uses a monoclonal antibody (JF5) and a lateral-flow device (LFD) (OLM Diagnostics, Newcastle upon Tyne, UK) to detect an extracellular glycoprotein antigen produced by A. fumigatus. This test has a high negative predictive value and good sensitivity and specificity, especially with BAL fluid. U.S. Food and Drug (FDA) approval of this test for diagnostic use is still pending.
Antigen testing is considered the primary diagnostic test in screening cerebrospinal fluid (CSF) for suspected cases of cryptococcal meningitis because the India ink procedure has a low sensitivity. The combination of antigen detection test and an India ink stain of the CSF are recommended for the primary evaluations of suspected cases of cryptococcal meningitis. Several commercial kits are available for the detection of cryptococcal antigen in serum and CSF.18 Galactoxylomannan is a polysaccharide capsular component of Cryptococcus and is the antigen detected for infections caused by serotypes of C neoformans, C gattii, and C deneoformans. Latex agglutination (LA) and enzyme immunoassay (EIA) methods are sensitive (93% to 100%) and specific (93% to 100%) diagnostic tests for the detection and quantitation of circulating cryptococcal antigen in serum and CSF. The reported titer determinations of the two testing methods (eg, LA versus EIA testing) or from different commercial latex kits are not numerically similar. Thus, the same testing method and latex kit should be used to monitor serial samples for a patient. Numerous causes have been responsible for false-positive results, with both testing methods including rheumatoid factor, soaps, disinfectants, hydroxyethyl starch, malignancy, and infections associated caused by bacteria, Trichosporon, Capnocytophage, Rothia, or Geotrichum beigelii. Both low and high antigen titers can result in false-negative results. Recently, LFD device has become available for measuring cryptococcal antigen from serum and CSF samples. The advantages of LFD include similar specificity and increased sensitivity as LA and EIA for testing serum samples, ease of use, lower costs compared with other test kits, and the similar accuracy between whole blood samples and blood obtained from finger pricks. False-positive results with LFD have been reported at low titers in patients without a history of cryptococcal infection.
Enzyme immunoassay can be used to detect a polysaccharide antigen from H capsulatum in body fluids (eg, serum or plasma, urine, CSF, or BAL fluid). It has been recommended that the antigen screening test be validated by antibody testing (eg, immunodiffusion [ID] and complement fixation [CF]).18 The diagnosis of histoplasmosis should be based on a combination of diagnostic test results because antigen testing is associated with cross-reactivity to other fungal infections (eg, blastomycosis, coccidioidomycosis, paracoccidioidomycosis) and false-positive results (eg, rheumatoid factor, rabbit antithymocyte globulin). The test sensitivity varies with disease presentation (eg, 77% for acute pulmonary histoplasmosis, 34% for subacute pulmonary histoplasmosis, 21% for chronic pulmonary histoplasmosis, 92% for progressively disseminated histoplasmosis), patient groups (eg, HIV infection, immunocompromised, disseminated diseases), and specimen type (60% to 86% in serum, 80% to 95% in urine, 25% to 50% in CSF, 93.5% in BAL). A monoclonal antibody ELISA has also been developed and has improved sensitivity (98%) and specificity (97%).
Antigen detection tests for H capsulatum, B dermatitidis, and Coccidioides species are performed by the clinical reference laboratory, MiraVista Diagnostics (Indianapolis, IN), on a fee-for-service basis. Antigen detection is generally not used as a diagnostic tool and has a limited role for blastomycosis and coccidioidomycosis because of low levels of detection in antigenemia and antigenuria, false-positive reactions, and/or cross-reactions are common in patients with other mycoses.
Mannin, a major component of the Candida cell wall, has been the main diagnostic marker used in antigen detection tests of Candida species.18 For serology testing, antimannin antibodies can be monitored because mannin can induce a strong antibody response toward oligomannose epitopes. A wide range in assay sensitivity and specificity has been reported when either antigen or antibody detection tests are used alone for the diagnosis of candidemia and disseminated candidiasis. The combined detection of circulating mannan and antimannan antibodies in serum or plasma by immunoenzymatic assays (Platelia Candida Ag Plus and Platelia Candida Ab Plus, Bio-Rad Laboratories, Marens-La-Coquette, France) has been recommended to maximize the early diagnosis of invasive candidiasis. Concomitant detection of mannan and antimannan antibodies for Candida species has a median sensitivity of >80%, particularly for C albicans, C glabrata, and C tropicalis.
Several different serology methodologies (eg, ID, countercurrent immunoelectrophoresis, ELISA, CF tests, fluorescent-enzyme immunoassay) have been investigated for the detection of specific fungal pathogens.18,24 Interpretation of results for most fungal infections requires knowledge of the laboratory technique used to perform the antibody testing. Serologic assays are most useful as diagnostic testing of fungal infections in the immunocompetent host because a poor antibody response is common in immunosuppressed patients, resulting in a false-negative result.
Serologic tests (ie, ID and CF) play an important role in the clinical diagnosis of infections caused by H capsulatum.18,24,27 These tests have been the most useful in patients with chronic pulmonary or disseminated histoplasmosis. The ID test is more specific than CF test and can serve as a useful screening procedure with and without the CF test. The CF test is more sensitive than ID but has shown cross-reactivity and positive results in patients with various other types of infections, including bacterial, viral, mycobacterial, and fungal (eg, aspergillosis, blastomycosis, candidiasis, coccidioidomycosis, paracoccidioidomycosis). CF test can also have false-negative results in the presence of rheumatoid factor or cold agglutinins. Other serologic assays (eg, LA, EIA, ELISA) have also been evaluated and may be useful for the diagnosis of specific types of H capsulatum infections. The potential of cross-reactivity or lack of commercial availability limits the current use of these assays.
The ID and CF tests are reliable serologic methods for the diagnosis of invasive infections of coccidioidomycosis and paracoccidioidomycosis.18,27 The ID test mainly detects immunoglobulin M (IgM) antibodies (heated coccidioidin) and is useful in the diagnosis of active disease. The ID has replaced the historical use of a tube precipitin test. The CF detects IgG antibodies (unheated coccidioidin) and is useful in diagnosing acute or chronic diseases and predictive for monitoring treatment response and a poor prognosis. LA and a highly sensitive EIA are also available; however, false-positive results have been noted. These tests should only be used as a screening tool, in which a positive result must be confirmed by another method.
Finally, serology testing methods (ie, immunoelectrophoresis, ELISA, and fluorescent-enzyme immunoassay) for Aspergillus-specific antibodies are useful for the diagnosis of noninvasive diseases such as allergic bronchopulmonary aspergillosis, aspergilloma, and chronic cavitary aspergillosis.18 Low sensitivity and/or specificity currently limit the use of serology testing as definitive diagnosis of invasive fungal infections caused by species of Blastomyces, Candida, and Cryptococcus.
Commercial assays for the detection of (1,3)-β-d-glucan, a polysaccharide present in cell wall of common pathogenic yeasts, have been used as a panfungal diagnostic tool for invasive fungal infections such as aspergillosis, Fusarium infection, trichosporonosis, and candidiasis.18,27 This assay has also been used to detect (1,3)-β-d-glucan from P jirovecii, in both HIV-positive and HIV-negative patients. This diagnostic assay is not useful for mucoraceous molds (eg, Zygomycetes such as Rhizopus, Mucor, and Absidia), which do not produce (1,3)-β-d-glucan, or Cryptococcus species and B dermatitidis, because they produce only low levels of (1,3)-β-d-glucan. Limited evaluations have assessed this diagnostic assay for the detection of histoplasmosis and coccidioidomycosis.
In the United States, Fungitell assay (Associates of Cape Cod Inc., East Falmouth, MA) is widely available and the only FDA-approved screening test for detecting (1,3)-β-d-glucan in serum. The manufacturer’s recommended guidelines for a positive serum (1,3)-β-d-glucan value is ≥80 pg/mL and a negative value <60 pg/mL; values between 60 and 79 are considered indeterminate (http://www.acciusa.com). Repeat testing (eg, twice weekly) is recommended to improve the predictive value and specificity of the test. False-positive results have been observed in patients receiving hemodialysis (with cellulose membranes), treated with certain blood products (eg, albumin, immunoglobulins), having bacterial bloodstream infections, mucositis, or graft-versus-host disease, and/or exposed to glucan-containing materials (eg, gauze or swabs). Concurrent β-lactam therapy, such as piperacillin–tazobactam or amoxicillin–clavulanate, and antitumoral polysaccharides have also been associated with false-positive results. Because of these potential risks of a false-positive result, the test may be more useful in excluding a diagnosis of invasive fungal infection when results are reported negative. Because this assay is nonspecific with varying levels of sensitivity and specificity, its use should be in conjunction with clinical examination of the patient and other diagnostic tests and procedures to make a conclusive diagnosis of invasive fungal infection.
Molecular diagnostic tests have a significant and increasing role in the detection and identification of fungi.15,18–24 The advantages of these techniques include organisms being observed microscopically but not grown on culture; a more rapid and objective identification of molds with unrecognizable or unproductive structures or yeasts not included in commercial databases; the ability to differentiate fungi with similar characteristics; and the precise genotyping for epidemiology studies and updates to taxonomy, classification, and nomenclature of fungi.18 The reader is referred to a glossary of common molecular terms for comprehending information in this rapidly evolving field.15,18
The ribosomal targets and internal transcribed spacer regions have been the main target used for molecular identification of fungi. Procedural steps that are commonly involved with molecular identification techniques include extraction of DNA, amplification of DNA segment of interest, and DNA analysis. Amplification is most often performed by polymerase chain reaction (PCR) with non–sequencing-based or sequencing-based identification methods. A wide variety of methodologies are available, including local and in-house laboratory-developed PCR tests. Clinicians need to contact their laboratory to determine which tests are available and which molecular diagnostic tests may need to be sent to a reference laboratory. In addition, selection of appropriate primers and/or probes for molecular testing often relies on the initial impressions and/or characteristics of the isolate, particularly for less robust molecular identification methods. In many situations, a combination of morphologic and molecular testing methods is best used for species identification.
Many evaluations have been ongoing for different PCR assays for invasive Candida spp. and Aspergillus infections.15,19,27 Limited and variable sensitivity and specificity have been some of the main issues restricting the routine use of this method. Additional issues that need to be addressed include specimen type, sample volume, best method of DNA extraction, target range, and definitions of positive results. However, further evaluation with standardized methodology and decreased inconsistencies between tests should allow PCR to become a promising method for detection of Candida and Aspergillus spp. Like other fungal infections, nucleic acid detection has been extensively used as a research-based tool with a slower progression to routine use by clinical microbiology laboratories and/or commercial availability.27
Several molecular-based diagnostic assays have been cleared by the FDA and are commercially available for clinical use in the United States.15,18–22,27 Even more devices have been marked Conformitè Europèene In Vitro Diagnostic (CE-IVD) and marketed for use in Europe.19 These devices have incorporated detection technologies such as DNA amplification followed by magnetic resonance, peptide nucleic acid fluorescent in situ hybridization (PNA-FISH), chemiluminescent labeled with single-stranded DNA probes, nested multiplex PCR or real-time PCR with DNA melt-curve analysis, and a system platform involving PCR amplification, flow cytometry, and dual-lasers detection. Most of the available commercial devices have focused on the detection and identification of fungi species commonly associated in infections, such as Aspergillus, Cryptococcus, and several Candida spp. (ie, C albicans, C glabrata, C krusei, C parapsilosis, and C tropicalis). The following brief discussion highlights a few examples of molecular devices that have FDA clearance for clinical use in the United States.
The T2Candida Panel and automated T2Dx Instrument (T2Biosystems, Lexington, MA) uses novel technologies to allow rapid (eg, 3 to 5 hours) and accurate diagnosis of invasive candidiasis directly from whole blood samples (no need for blood culture and isolation of Candida spp.).15,18–22,27 The instrument is a fully automated as a clinical multiplex benchtop diagnostic system using PCR and miniaturized magnetic resonance technology. The T2Candida panel can identify C albicans, C tropicalis, C parapsilosis, C glabrata, and P kudriavzevii. Other panels are in development, including T2Cauris panel (research use only) for the detection of the emerging multidrug-resistant Candida auris. A T2Bacteria panel has FDA clearance and is available for use on the same instrument.
PNA-FISH (Yeast Traffic Light PNA FISH, OpGen, Gaithersburg, MD) provides rapid identification of up to five Candida spp. directly from yeast-positive blood cultures.15,18–22 This device has a fast turnaround time (eg, approximately 90 minutes) with high sensitivity (92% to 100%) and specificity (94% to 100%). The rapid identification methodology used is hybridization of fluorescent PNA probes to organism-specific rRNA, with detection via fluorescent microscopy. After the Gram stain and the hybridization process are completed, C albicans and C parapsilosis are identified microscopically as bright green fluorescing cells, while Candida tropicalis fluoresces bright yellow, and C glabrata and C krusei fluoresce bright red. Other yeasts do not fluoresce. The colors of the light probes also provide an indication about the potential use of fluconazole in these patients because C albicans and C parapsilosis are generally susceptible to fluconazole (green light for go), C glabrata can be resistant to fluconazole, and C krusei is intrinsically resistant to fluconazole (red light for stop). The yellow signal produced by C tropicalis indicates that caution should be used because fluconazole susceptibility is variable for this organism. This method has a significant impact over traditional identification methods, which could take up to 3 or more days for identification of Candida spp., as well as guiding the most effective antifungal drug therapy. The PNA-FISH methodology is also used for rapid identification of bacteria, including gram-positive (ie, Staphylococcus aureus, coagulase-negative staphylococci, enterococci) and gram-negative (ie, Escherichia coli, Klebsiella pneumoniae, Pseudomonas aeruginosa) organisms. A single automated system using FISH technology (Accelerate Pheno BC kit, Accelerate Diagnostics, Inc., Tucson, AZ) has also been developed and can identify C albicans, C glabrata, and gram-positive and gram-negative organisms.
Probe-based assays for culture identification of fungi have become commercially available.15,18–22 AccuProbe (Hologic, Inc., Mississauga, ON, Canada) uses luminometer to detect hybridization of a chemiluminescent labeled, single-stranded DNA probe to target rRNA present in a fungal culture. Three separate probes have FDA clearance and are available for the culture identification of dimorphic fungi, including Blastomyces dermatitidis, Coccidioides immitis, and Histoplasma capsulatum. Performance data have demonstrated high sensitivity (>98%) and specificity (>99%) for each pathogen. The B dermatitidis probe has the potential to cross-react with other fungi, including Emmonsia species, Paracoccidiodes brasiliensis, and Gymnascella spp. In addition, the Coccidioides probe is unable to distinguish between species, namely Coccidioides immitis and Coccidioides posadasii. AccuProbe tests are also available for culture identification of bacteria (ie, Neisseria gonorrheae, S aureus, Listeria monocytogenes, Streptococcus pneumoniae) and mycobacteria.
BioFire FilmArray (BioFire, Salt Lake City, UT) is an automated in vitro diagnostic device that detects multiple nucleic acid targets by using nested multiplex PCR with DNA melting curve analysis.15,18–22 Six different identification panels are currently available, with yeast being included on two of these panels. The Blood Culture ID (BCID) Panel can test for 43 targets associated with bloodstream infections, including gram-positive and gram-negative bacteria, yeast, and 10 antimicrobial resistance genes. The yeast detected by the BCID Panel includes six Candida spp. (ie, C albicans, C auris, C glabrata, C krusei, C parapsilosis, and C tropicalis) and Cryptococcus neoformans/gatti. This panel requires a positive blood culture sample. The overall sensitivity and specificity for the BCID Panel are 99% and 99.8%, respectively. Cryptococcus neoformans/gatti, along with 13 common bacterial and viral pathogens, are included on the Meningitis/Encephalitis (ME) Panel. The ME Panel can directly detect pathogens from a 0.2 mL sample of CSF, and has also demonstrated a high sensitivity (94.2%) and specificity (99.8%).
Luminex (xMAP and xTAG, Luminex Molecular Diagnostics, Inc., Austin, TX) is a commercially available multianalyte profiling platform that can provide detection and identification of clinically important pathogens directly from positive blood culture bottles and other types of clinical samples.15,18–22 The platform has combined PCR amplification, flow cytometry, and dual-laser detection system to provide multiplexed assay capabilities. The xMAP (x = analyte or unknown; MAP = Multi-Analyte Profiling) hybridization technology can detect an antigen (target) by using a capture antibody attached to the surface of a color-coded microbeads (microsphere) and a detection antibody that incorporates a fluorescent label. Luminex-based technology has allowed rapid and reliable identification of clinically important fungal pathogens, including up to 10 genus- and 29 species-specific diagnoses. xTAG (the TAG name is derived from three nucleic acid bases being used: T, A, G) consists of the MagPlex-TAG microsphere (that are precoupled with anti-TAG sequence) and the user designator primer and TAG sequence that complements the x-TAG sequence on the bead. The xTAG analyte-specific reagents can be combined with xMAP instruments for amplification and detection. Evaluations of the xTAG Fungal ASR assay have demonstrated that multiple yeast species could be identified with 100% sensitivity, 99% specificity, and 99% positive and 100% negative predictive values when compared with traditional fungal culture results.
MALDI-TOF Mass Spectrometry
Matrix-assisted laser desorption-ionization time-of-flight (MALDI-TOF) mass spectrometry (MS) is ideal for genus and species identification and has the potential for accurate strain typing and identification for fungi, bacteria, and mycobacterium.5,10,15,18–20,26 This technology is a rapid and accurate method for identifying yeasts and molds recovered on culture media and using sample preparation for MALDI-TOF MS. Reports on the use of MALDI-TOF MS for routine rapid identification have focused on clinically important yeasts (eg, Candida spp., including Candida auris, C neoformans, and C gattii spp.) and dermatophyte species (eg, Neoscytalidium spp., Trichophyton, Microsporum, Epidermophyton, and Arthroderma). Identification of dimorphic and filamentous fungi as well as molds (eg, Aspergillus spp., Fusarium spp., Pseudallescheria–Scedosporium complex, Penicillium spp., Lichtheimia spp.) have been more challenging because of different developmental forms on agar media and the influences of the phenotype. MALDI-TOF MS is becoming the primary diagnostic method for rapid identification of fungus isolates in the clinical microbiology laboratory. However, its initial use for fungal identification has moved at a slower pace than the current use of MALDI-TOF MS for bacterial identification.
The advantages of the MALDI-TOF MS for fungal identification are low cost of materials (a few cents) for each organism identification, ease of performance, and rapid, accurate results (approximately 11 minutes if just one isolate is tested; 2.5 minutes per isolate in a batch of 96 isolates, with the average time per isolate in published reports being 4 to 6 minutes). The simplicity of MALDI-TOF MS removes the specific skills and ability to visually identify fungi macroscopic and microscopic characteristics. Current limitations include the initial costs of instrumentation for the system, lack of sample preparation techniques, and inadequate fungal spectra in commercial database and software of manufacturers (ie, two commercial systems currently exist in the United States: Bruker, Billerica, MA and Vitek MS, bioMérideux, Inc, Durham, NC). The expansion of database libraries and developments in sample preparation are rapidly evolving to establish validated and routine procedures for large numbers of clinically important fungal strains and species. MALDI-TOF MS is also becoming a reliable and reproducible method for antifungal susceptibility testing (eg, caspofungin for isolates of Candida spp.).20,28 Finally, MALDI-TOF MS is also being investigated for epidemiologic testing of fungal isolates for outbreak investigations and as an early surveillance test and/or screening tool for antifungal drug resistance.20
Antifungal Susceptibility Testing
The importance of antifungal susceptibility testing has become increasingly recognized as a useful component in the treatment optimization of invasive infections caused by Candida spp. because of the increasing number of available antifungal agents, emerging resistance issues to standard therapy, and the changing epidemiology of invasive fungal disease. Obtaining antifungal susceptibility testing is particularly important when azole-resistant isolate is suspected, when failure to respond to antifungal therapy has occurred, and in species in which acquired resistance often exists (ie, Candida glabrata, Candida auris, and Aspergillus fumigatus).
The CLSI and European Committee on Antimicrobial Susceptibility Testing (EUCAST) has developed standardized reference methods for macrodilution and microdilution susceptibility testing of yeasts and molds and broth microdilution method for dermatophyte.28–30 Agar-based alternative methodologies, including agar dilution, disk diffusion, E-test methods, and semisolid agar, have also been applied to susceptibility tests of yeasts and molds. The commercial availability of simplified and/or automated testing methods (eg, E-test and other gradient strip testing; Vitek 2; Sensititre YeastOne) consistent with CLSI reference methods is allowing an increasing number of clinical laboratories to routinely perform antifungal susceptibility testing. Molecular testing methods for the detection of resistance have also been expanding.28–30
Interpretive MIC breakpoints based on CLSI- and EUCAST-recommended in vitro susceptibility testing methods have been recommended for Candida spp.28–30 Comprehensive reviews regarding the microbiological, molecular, pharmacokinetic-pharmacodynamic, and clinical antifungal data for Candida spp. provide species-specific interpretive clinical breakpoints for azole agents and the echinocandins (Table 19-5).28 These data have been used to establish epidemiologic cutoff values, detect emerging resistance among Candida spp., and harmonize antifungal susceptibility testing standards by CLSI and EUCAST.28–30 In addition, tentative breakpoints for the multidrug-resistant pathogen Candida auris have been proposed (https://www.cdc.gov/fungal/candida-auris/c-auris-antifungal.html).
Species-Specific Breakpoints for In Vitro Susceptibility Testing of Candida spp. According to CLSI and EUCAS
CLSI = Clinical and Laboratory Standards Institute (revised breakpoints from M27-S4 document); EUCAST = European Committee for Antimicrobial Susceptibility Testing; S = susceptible; R = resistant.
aThe wild-type populations of C parapsilosis to anidulafungin and micafungin and C glabrata to fluconazole are classified as intermediate (I) category (values between S and R) to accommodate use of these agents in some clinical situations.
bThe wild-type population of C glabrata is classified by CLSI as susceptible dose-dependent (SDD) to fluconazole to accommodate use of fluconazole at higher doses in some clinical situations.
Notes: Clinical breakpoint of ≤1 mg/L for amphotericin has been set by EUCAST for Candida albicans. EUCAST breakpoints for caspofungin have not been established due to an unacceptable variation in MIC ranges.
Source: Adapted with permission from Johnson EM, Cavling-Arendrup M. Susceptibility test methods: yeast and filamentous fungi. In: Carroll KC, Pfaller MA, Landry ML, et al, eds. Manual of Clinical Microbiology. 12th ed. Washington, DC: American Society for Microbiology Press; 2019:2351–2375.
Interpretive breakpoint criteria for amphotericin B and some of the triazole agents against selected Aspergillus spp. have been reported; other fungal pathogens remain to be standardized.28–30 The recommended EUCAST clinical breakpoints for Aspergillus fumigatus include ≤1 mg/L (susceptible) and >2 mg/L (resistant) for amphotericin, itraconazole, and voriconazole, and ≤0.125 mg/L (susceptible) and >0.25 mg/L (resistant) for posaconazole (provided sufficient drug concentrations can be achieved).
More than 650 viruses are known to cause infection in humans and other vertebrate animals.31 The three major properties that classify viruses into families include (1) the nucleic acid (NA) core (either DNA or RNA but not both); (2) whether the viral NA is single-stranded or double-stranded (https://viralzone.expasy.org/656); and (3) the presence or absence of a lipoprotein envelope (Tables 19-6 and 19-7).13,31,32 Viruses also differ based on their genome topology (eg, linear, circular, single versus multiple segments). Virus families can be further categorized based on morphology (eg, size, shape, and substructure), mode of replication, and molecular and genomic characteristics. The most recent information on the rapidly changing classification and taxonomy of viruses can be obtained from the website database (http://ictv.global/report/) that has been established by The International Committee on Taxonomy of Viruses (ICTV). The 2019 ICTV report now recognizes five hierarchical ranks consisting of 55 orders, 168 families, 103 subfamilies, 1421 genera, and 6,590 species of viruses; however, a larger number of viruses remain unclassified.
Characteristics and Laboratory Diagnosis of Selected DNA Viruses of Medical Importance to Humans
aCommonly used methods in clinical laboratories: electron microscopy (EM); immunoassays (IA) including immunofluorescence assay (IFA) and NAATs.
bThe isolation of some pathogens (eg, smallpox) requires biosafety level (BSL) 3 or 4 facilities, usually only in specialized centers collaborating with World Health Organization. The isolation of Vaccinia virus requires BSL-2 (grows readily in cell culture).
The ability to detect and accurately identify viruses in the clinical laboratory has increased during the last 30 years as a result of wider applicability of diagnostic laboratory techniques with increased sensitivity and decreased turnaround time, the availability of newer reagents and rapid commercial diagnostic kits, and the addition of new antiviral drugs for specific viral infections.1,13,32–37 In addition, several NA amplification tests (NAATs), specifically PCR and real-time PCR, are allowing routine clinical laboratories to provide virology services for the increasing frequency of infectious diseases that depend on rapid viral diagnosis.1,13,32
It is important to note that all diagnostic tests for the identification of viruses are not available at each institution, and the clinician must establish a relationship with the laboratory that will be performing viral testing. In certain clinical situations, samples may need to be sent out for diagnostic testing at either large reference or public health laboratories because they are able to provide the necessary methods that are difficult or impossible to routinely perform in the clinical virology laboratory. In addition, certain viruses (eg, arboviruses, arenaviruses, filoviruses, Variola virus, and rabies virus) require testing at biosafety level (BSL) 3 or 4 facilities and are often sent to the Centers for Disease Control and Prevention (CDC) or the CDC’s Division of Vector-Borne Infectious Diseases.13,32,33
The ability to accurately diagnose a viral infection is highly dependent on appropriate selection, timing, collection, and handling of biological specimens.32–34 In general, the highest titers of viruses are present early in the course of illness and decrease as the duration of illness increases. Therefore, it is important to collect specimens for the detection of viruses in the early course of an infection. In most cases, identification of viruses is a specimen-driven process. Because collection procedures are highly dependent on viruses being suspected, attention needs to be taken regarding collection containers and devices, and transport systems (eg, whether a viral transport medium is needed). The different types of clinical specimens that can be collected for viral culture and antigen detection include respiratory specimens (eg, nasopharyngeal swabs, aspirates, and washes; throat swabs; BAL and bronchial washes), blood, bone marrow, CSF, stool, biopsy tissue, urine, ocular specimens, vesicles and other skin lesions, and amniotic fluid. In addition, specimens for molecular diagnostic testing (eg, PCR and other nucleic amplification techniques) must be obtained following specific guidelines so that the stability and amplifiability of the NAs are ensured.
Once the sample is collected, it should be promptly transported to the laboratory in a sterile, leak-proof container using the appropriate viral transport media to maximize viral recovery. Every effort should be made to prevent delay between the time of specimen collection and its arrival to the laboratory. When delays are expected, viral samples should be refrigerated at 4°C or frozen at –70°C. Subsequently, the laboratory will need to follow specific processing procedures for each specimen and the different diagnostic viral test methodologies.
The laboratory techniques used in the diagnosis of viral infections include cell culture, cytology and histology, electron microscopy (EM), antigen detection, NAATs, and serologic testing.1,13,32–38 The choice of test(s) varies depending on the clinical syndrome or disease, virus(es) involved, patient characteristics, collection site, purposes of the test (eg, screening, diagnosis, confirmation or monitoring), time to result, laboratory capabilities/staff expertise, and cost. The following section, as well as Tables 19-6 and 19-7, provides a brief summary of the common methods currently used in diagnostic testing of common viruses.1,13,31,32 For more detailed information, the reader is referred to current published literature, standard reference books, and the latest edition of reference manuals (eg, Manual of Clinical Microbiology). A list of virology services offered by the CDC can be found on the CDC website (https://www.cdc.gov/laboratory/specimen-submission/list.html).
The use of cell culture rapidly expanded the knowledge about the epidemiology, clinical characteristics, and diagnosis of common viral infections in the 1950s and 1960s. Subsequently, the use of cell cultures to isolate a virus became the gold standard method for the diagnosis of viral infections in most clinical virology laboratories for the next 50 years.13,32–36 During that time, other technologies for viral detection were slowly introduced in the clinical laboratories, including enzyme immunoassays, IgM class capture assays, rapid centrifugation cultures, direct viral antigen detection from clinical specimens by immunofluorescence, and monoclonal antibodies for identification. However, rapid and accurate serology and molecular methodologies have become cornerstones for virus detection and identification in clinical laboratories during the past decade, resulting in a decline in the use and prominence of cell cultures in larger academic medical centers and tertiary-care facilities. Despite these changes in routine diagnostic virology, viral cultures play an important role in the discovery of new or unknown viruses, identification of variants of known viruses, detection of drug-resistant viruses, typing of serologic strains, detection of viruses in special patient populations (eg, immunocompromised patients), research and development of antiviral drugs and vaccines, and performance of viral susceptibilities.34–36
The advantages of cell culture include good specificity and sensitivity, the capability of detecting multiple viruses if present, and the cultivation of the virus for further laboratory testing (eg, susceptibility testing, serologic strain typing), if needed.32,34–36 Cell cultures can be useful when combined with highly specific monoclonal antibodies or engineered cell lines (eg, to produce virus-induced enzymes), especially if the cost of other testing methods is greater than cell cultures or when the clinical laboratory does not have the ability and equipment to perform molecular detection methods. The disadvantages of cell culture include the long time needed for the detection of viruses using conventional cell culture (eg, days to weeks), the need for cell culture facilities, the expense of performing cell culture, and the fact that the methodology is not applicable to all viruses (eg, viruses that have not grown in conventional cell cultures [ie, Group C rhinovirus]). This greater demand for technical laboratory experience with cell cultures, need for comprehensive quality control program, and strict procedures for handling biohazardous materials in the clinical virology laboratory are being replaced by rapid and sensitive antigen screening assays and NAATs.34
Several different types of cell culture are available to grow clinically important viruses.32,34–36 Each virus requires a predefined cell line, which is established once a cell culture has been subcultured in vitro (the reader is referred to a comprehensive list of available cell lines and virus susceptibility profiles34). The different types of cell lines can be divided into three categories: primary, diploid (also called low passage cell lines), and heteroploid. Primary cell lines (eg, rhesus monkey kidney [RhMK] cells or human amnion cells) are prepared from animal or human tissues and can withstand only one or two passages until the cells die. Diploid cell lines are usually derived from fetal or newborn cells (eg, human embryonic lung fibroblast lines such as WI-38 or MRC-5) and can undergo 20 to 50 passages before cells are unable to survive. Continuous cell lines can undergo an indefinite number of passages without reducing the sensitivity to virus infection. Heteroploid cell lines are characteristically derived from human or animal cancers (eg, human epidermoid lung carcinoma [HEp-2, HeLa]) or are cells transformed in vitro (eg, LLC-MK2). Heteroploid cell lines can also include genetically engineered cells (eg, ELVIS cell mixture for the detection of herpes simplex virus [HSV] types 1 and 2). Most specimens are inoculated onto two or more cell lines (eg, RhMK, MRC-5, HEp-2) based on the most likely viruses associated with the type of clinical specimen that was submitted.
The growth of a virus from a clinical specimen provides direct evidence that the patient was infected with a virus. The main method for detecting growth from the cell culture method is by microscopic examination of the unstained cell cultured monolayers for morphologic changes or cytopathic effect (CPE).32,34–36 The characteristics of the CPE (eg, which cell culture types were affected; what is the resultant shape of the cells; whether the effect is focal or diffuse; the time of its appearance and progression) can be used for primary and definitive identification of the virus. Subsequently, direct and indirect fluorescent antibody (DFA and IFA, respectively) staining of cells with virus-specific monoclonal antibodies harvested from the culture is often used to confirm the identification of the virus (the reader is referred to a comprehensive list of available DFA and IFA reagents and target virus to detect34). Molecular or ancillary traditional testing can alternatively be used for viral identification.
Some viruses, such as influenza, parainfluenza, and mumps virus, grow in cell cultures without producing CPE so that other methods are used to identify and detect these viruses, including hemadsorption and interference.32,34–36 Hemadsorption involves the removal of the culture medium from the inoculated cell culture, adding a suspension of erythrocytes, and examining for hemadsorption with a low-power microscope as manifested by adherence of the red cells to the cell culture monolayer due to the presence of a hemadsorbing virus. Hemadsorption is used to detect these viruses, which can grow rapidly and reach high titers in cell cultures without producing CPE. Used to detect viruses such as rubella, interference involves growing a virus that yields a cell culture resistant to other viruses (to which it is normally susceptible). The viruses that produce hemadsorption or interference subsequently can be identified by staining with virus-specific monoclonal antibodies or antiserum.
Shell vial cultures with centrifugation and pre-CPE detection are used to decrease the amount of time required to grow a virus by conventional cell cultures.32,34–36 This technique makes use of cells grown on microscope coverslips that are placed within shell vials and covered with culture media. After cultures are incubated for 1 to 3 days, FA staining is performed on the cells on the coverslips to recognize an antigen in the nucleus of infected cells. Shell vial cultures have been commonly applied for the detection of cytomegalovirus (CMV), HSV, varicella-zoster virus (VZV), enteroviruses, and the human respiratory viruses. Centrifugation-enhanced rapid cell cultures can also be used with cocultivated cells (eg, mixture of two cell lines together) or genetically engineered cells (eg, ELVIS [enzyme-linked virus-inducible system], BGMK-hDAF [buffalo green monkey kidney cell line]) for the rapid identification (eg, 16 to 72 hours) and blind staining of multiple viruses from a single shell vial or tray well.32,34–36
The detection of specific viral NAs by molecular diagnostic techniques is revolutionizing the field of diagnostic virology.1,13,32,34,38 NAATs have become the “gold standard” in clinical virology laboratories and are replacing older techniques such as cell cultures for detecting clinically significant viruses. Many different techniques are used in viral NA detection, including direct hybridization assays, target (template) amplification (eg, PCR, self-sustained sequence replication method, strand displacement amplification), and signal amplification (eg, branched-chain DNA [bDNA] assay and hybrid capture assay). Among these, PCR has been the most important technique in diagnostic virology because of its versatility in detecting DNA or RNA and being able to provide qualitative and quantitative information on specific viral NAs.
The use of NA detection has become the standard of care (eg, hepatitis C virus [HCV] and HIV) or the test of choice for routine diagnosis of many viral infections (eg, bocaviruses, HSV central nervous system [CNS] infections, human HVS 6 and 7, human metapneumovirus, human papillomavirus [HPV]).1,13,32 The FDA has cleared or approved commercial molecular detection assays; several viruses, including hepatitis B and C viruses (HBV, HCV); HIV; HSV; CMV; adenovirus; avian flu; enteroviruses; influenza; and HPV. An FDA-approved simple multiplex PCR test (eg, xTAG Respiratory Viral Panel) is also available for rapidly screening common respiratory viruses (eg, respiratory syncytial virus [RSV], influenza A and B, adenovirus) or subtypes.1,13,38–41 In addition, FDA-approved, high-throughput, syndromic viral tests are also available for detection of GI and CSF pathogens (eg, BioFire FilmArray GI and Meningitis/Encephalitis panels).1,39–43 An up-to-date listing of cleared or approved nucleic acid diagnostics tests is available at the FDA website (https://www.fda.gov/medical-devices/vitro-diagnostics/nucleic-acid-based-tests).
The advantages of viral NA detection methods include the rapidity of results (eg, hours for real-time PCR and one to several days for other methods), maximal sensitivity for virus-specific detection and identification, adequate to excellent specificity, dramatic increase in availability of commercial assays, the ability to detect viruses that are difficult to culture, and the ability to detect NAs without viable virus present in the clinical specimen. Historically, equipment and reagents costs, service contracts, and technical expertise have been the major barriers to implementing molecular testing. However, NAAT has rapidly evolved and allows viruses such as influenza virus and RSV to be detected within 20 to 30 minutes at near POC and with sensitivity similar to other laboratory testing methodologies.1,13,39–41,44 Several devices (eg, FilmArray Respiratory Panel EZ assay [Biofilm, Salt Lake City, UT]; ID NOW RSV [Abbott Diagnostics, Scarborough, ME]; the cobas Liat Influenza A/B & RSV [Roche Diagnostics, Indianapolis, IN]) have been waived by the Clinical Laboratory Improvement Amendments (CLIA).13,40,41,44 Molecular assays have become the standard of care for diagnosing viral infections and monitoring antiviral therapy and patient outcomes.1,13,32
Cytology and Histology
Cytopathologic effects (CPEs) on cells are produced by many viruses. Cytologic examination can be performed on smears prepared from samples that are applied to a microscope slide or “touch preps” of unfixed tissues.32,34–36 Cytologic findings are suggestive of a viral infection and provide identification of cell morphologies (eg, “owl’s eye” nuclear inclusions consistent with CMV), cell lysis, or other cell changes (eg, vacuolation, syncytia, inclusion bodies). The specific virus cannot be identified unless virus-specific immunostaining techniques are used. Applications of cytology to viral diagnosis include the Tzanck smear with Giemsa reagent for demonstrating the presence of HSV or VZV infection, Papanicolaou staining of cells obtained from the uterine cervix (Pap smear) for providing evidence of HPV infection, and cytologic staining of urinary sediments for screening the presence of either CMV or polyomaviruses JCV and BKV.
Similar to cytology, histologic examination of tissue provides evidence to suggest a group of viruses that may be causing infection, but it does not identify a specific virus.32,34 Despite this shortcoming, histopathology has been useful in differentiating between asymptomatic viral shedding and clinically important infections of CMV and has been used for the diagnosis of CMV infections in tissue samples obtained from biopsy or at autopsy. In addition, detection of specific viral antigens by immunohistochemistry and detection of specific viral NAs by in situ hybridization (ISH) or PCR has allowed specific viruses to be identified by histopathology.
Viruses are the smallest infectious pathogens that range in diameter from 18 to 300 nm.32,37 Direct visualization of a virus with a light microscope can be performed only on pathogens with a diameter >200 nm. Electron microscopy (EM) allows visualization of characteristic viral morphology and, unlike direct detection or molecular methodologies, is capable of detecting the distinctive appearances of multiple viruses, if present.32,37 EM is considered the most useful routine test for poxviruses.66 Diagnostic virology laboratories also commonly use EM for detection of viruses that are not detected with cell cultures or other methods (eg, gastroenteritis viruses such as noroviruses, coronaviruses, astroviruses, enteric adenovirus, and calicivirus).13,32,34,37,43
Several techniques have been incorporated to allow the visualization of viruses with EM from various types of clinical specimens. Negative staining is a technique for identification of viruses in fluid samples, stool samples, and blister fluid. Thin sectioning can be performed on tissue samples that have been fixed with specific fixatives for EM study, and it can be used to visualize herpesviruses, respiratory viruses, and rabies virus.
More sensitive methods are replacing the routine use of EM for detecting clinically significant viruses.13,32,37 An advantage of EM is its economical, quick (eg, same day), adaptable, and straightforward approach for detecting viruses. The major disadvantages of EM include poor sensitivity, initial equipment expenses, and need for highly skilled laboratory staff.
Direct Antigen Detection
Antigen detection methods involve the use of virus-specific antibodies directed toward viral antigens in a clinical specimen.1,13,32,38 Examples of viruses that can be identified by direct antigen detection include RSV, influenza virus, parainfluenza virus, adenovirus, HSV, VZV, CMV, rotavirus, HBV, and measles virus.1,13,32,39–41 The advantages of direct antigen detection include the rapidity of diagnosis (eg, several hours to 1 day), usefulness for the identification of viruses that are difficult to culture, and detection of viral specific antigens even if viable virus is not present in the clinical specimen. The disadvantages include the potential for false-positive and false-negative results, difficulty of performing batch testing, and lack of sensitivity necessary for diagnostic applications for all viruses (eg, not applicable for rhinoviruses because there are >90 serotypes and cross-reacting antibodies).
The techniques commonly used for antigen detection include immunofluorescence assay (IFA; direct and indirect), EIA (including ELISA), chemiluminescent and fluorescence-based immunoassay, and particle agglutination assays. Several membrane immunochromatographic assays (dipstick tests) are available as influenza diagnostic tests (eg, Directigen Flu A or A+B Test, QuickVue influenza).13,32,39,40 These viral antigen tests have become simple to use, are low cost, and allow rapid detection (≤30 minutes) of specific antigens from a single specimen at POC (eg, outpatient facilities, physician offices, patient bedside) and in the clinical laboratory. Many of these RIDT kits are CLIA-waived because the methodologies are simple to use and accurate, and the likelihood of erroneous results is negligible.13,39,45 Rapid influenza diagnostic tests (RIDTs) have lower specificity but variable sensitivity (higher in children and for detecting influenza A). The need for improved sensitivity of RIDTs was also observed during the 2009 pandemic of H1N1 influenza. These issues, in part, resulted in a medical device reclassification by the FDA in 2017 and additional compliance requirements of RIDTs for influenza.13,39,45 Further details on RIDTs and NA detection-based tests for influenza virus can be found at the CDC website (https://www.cdc.gov/flu/professionals/diagnosis/rapidlab.htm).
Serologic tests are designed to detect an antibody response in serum samples after exposure to viral antigens has occurred.32 The major uses of serology for the detection of viral infections include the demonstration of immunity or exposure to a virus, the diagnosis of postinfectious sequelae, and the screening of blood products. In several clinical situations, serologic testing remains the primary means for the laboratory diagnosis of viruses that are difficult to culture or detect by direct methods (eg, rubella virus, Epstein-Barr virus, hepatitis viruses, HIV, arboviruses).1,13,32 Serologic testing may also serve as a supportive or adjunctive role in clinical situations in which viral cultures or direct detection methods are available.
For viral infections, serologic testing can identify the virus, distinguish the strain or serotype, differentiate between primary infection and reinfection, and determine if the infection is in an acute or convalescent phase. Virus-specific immunoglobulin antibodies (eg, IgM or IgG) are produced during the time course of a viral infection. In general, virus-specific IgM is detected in serum sooner than virus-specific IgG. The results measure the relative concentration of antibody in the body as a titer, with the titer representing the lowest antibody concentration (or inverse of the greatest dilution; a dilution of 1:128 is expressed as a titer of 128) that demonstrates activity in a patient’s serum. The exact value for a titer varies with each testing method, the specific virus involved, the timing of specimen collection, and the presence of active disease.
For most viral infections, virus-specific IgM can be detected as soon as 3 to 7 days after the onset of infection. The presence of virus-specific IgM in a single serum sample shortly after the onset of symptoms (acute phase) is usually indicative of a recent or current primary infection. Titers of virus-specific IgM usually decline to near undetectable amounts within 1 to 4 months after the onset of infection. Virus-specific IgG can be detected during the acute phase of infection (eg, 1 to 2 weeks) and continues to increase for several months before reaching a maximal titer. Thereafter, the IgG titer declines, but it usually remains detectable in serum for the remainder of a person’s life. Seroconversion has occurred when at least a 4-fold increase in IgG titer has occurred between serum samples collected in the acute and convalescent (two to four weeks afterward) phases. The presence of virus-specific IgG is also indicative of a past infection.
Serologic tests are also used to assess the immunity or exposure to a virus. The presence of antibody can detect which patients have been previously infected by or vaccinated for a specific virus. For example, a positive result (presence of antibody) for rubella in a woman of childbearing age implies that congenital infection will not occur during subsequent pregnancies. A negative result (absence of antibody) implies susceptibility to infection, and the woman should receive rubella vaccination as a preventative measure if she is not pregnant. Some other examples of viruses for which serologic determination of immune status is useful include hepatitis A and B (HAV, HBV), measles, mumps, parvovirus B19, and VZV.1,13,32
The techniques commonly used for serologic assays include CF, EIA, IFA, anticomplement immunofluorescence, and western immunoblotting. In the diagnosis of certain viral syndromes (eg, CNS infections), a serology panel may be helpful so that a battery of antigens is tested for antibody to several viruses. The advantages of viral serology include the assessment of immunity or response of a virus isolated from a nonsterile site, serum specimens are easy to obtain and store, and it can be used to identify viruses that are difficult to culture or detect by immunoassay. The disadvantages include the time to results (eg, few days to weeks), the potential for cross-reactions between different viruses, and the need for both acute and convalescent specimens.
Antiviral Susceptibility Testing
The emergence of drug-resistant strains of viruses to antiviral agents is an increasing problem, especially in immunocompromised hosts. Unlike antibiotics, in vitro susceptibility testing of viruses has not been routinely available. The major variables that have limited the standardization of antiviral susceptibility testing include cell lines, inoculums titer, incubation period, testing range of antiviral drug concentrations, reference strains, assay methodology, and criteria, calculation, and interpretation of end points.46 However, emerging molecular technology and the phasing out of virus culture-based methods have permitted phenotypic and genotypic antiviral susceptibility testing vividly advance.
The FDA-cleared assays for viral susceptibility testing for the past decade have mainly been limited to phenotypic and genotypic assays for HIV. Antiviral resistance and cases of drug failure has led to increased interest in susceptibility testing of HSV, VZV, CMV, and influenza viruses.46 Thus far, most susceptibility testing for these pathogens has been limited to research use only or laboratory user-developed tests. The CLSI has published an approved standard for phenotypic susceptibility testing of HSV.47 This standard outlined the use of a plaque reduction assay and denotes resistance to acyclovir and foscarnet when inhibitory concentration 50% (IC50) values are ≥ 2 mcg/mL and ≥ 100 mcg/mL, respectively. Proposed guidelines for antiviral susceptibility results of HSV, CMV, VZV, and influenza A and B viruses for various other phenotypic testing methods (ie, DNA hybridization, EIA, neuraminidase inhibition assay, late antigen reduction assay) and antiviral agents (ie, famciclovir, vidarabine, cidofovir, ganciclovir, neuraminidase inhibitors) have also been outlined.46 Interpretation of these values must be carefully made in conjunction with the clinical response of the individual patient. Additional consensus documents and further standardization of phenotypic and genotypic assays for antiviral susceptibility testing are needed.
HUMAN IMMUNODEFICIENCY VIRUS
Human immunodeficiency virus (HIV) is the causative agent of AIDS. The HIV virus is an enveloped, positively stranded RNA virus that belongs to the Retroviridae (retrovirus) family and Lentivirus genus.48 The mature virus measures approximately 100 nm in diameter and has a characteristic conical core containing proteins, enzymes, and two identical copies of single-stranded RNA. Viral proteins within the core and the lipid envelope play a significant role in the detection, diagnosis, and treatment of HIV.49,50 The replication process of HIV involves transcription of viral RNA into proviral DNA using the reverse transcriptase (RT) enzyme. The proviral DNA is then integrated into the host’s genome using the integrase enzyme, resulting in lifelong latent infection. The virus is transmitted to humans by the exchange of blood or other body fluids containing the virus through sexual contact; exposure to contaminated blood; transfusion of contaminated blood and blood products; or via contaminated needles (eg, intravenous drug users or accidental needle sticks). In addition, infants can acquire HIV from an infected mother in utero, during labor or delivery, or during breastfeeding.49,51
There are two distinct serotypes of HIV, namely HIV-1 and HIV-2; while HIV-1 is the most prevalent serotype of HIV infections worldwide, HIV-2 infection is most commonly distributed in Western Africa and other limited geographic locations.49–51 Routine diagnostic testing of HIV-2 is not recommended in the United States because its prevalence is extremely low. Thus, the following discussion focuses mainly on laboratory tests used for the diagnosis and management of HIV-1 infection. However, HIV-2 testing may be indicated in persons at risk for HIV-2 infection or for those who have symptoms suggestive of HIV infection with negative or indeterminate test results for HIV-1. In addition, all blood donations in the United States are tested for both HIV-1 and HIV-2.49–51
Laboratory Tests for Human Immunodeficiency Virus-1 Infection
Several laboratory tests are available for the diagnosis and monitoring of patients with HIV-1 infection. The most common virologic testing methods include HIV-1 antibody assays, HIV-1 p24 antigen assays, DNA-PCR, plasma HIV-1 RNA (viral load) assays, and viral phenotypic and genotypic assays. In addition, the absolute number of CD4+ lymphocytes and the ratio of helper (CD4+) to suppressor (CD8+) lymphocytes (CD4+:CD8+ ratios) are routinely measured to evaluate the patient’s immune status and response to antiretroviral therapy, because HIV primarily infects and depletes CD4+ T helper lymphocytes. Viral cultures for HIV are not typically performed beyond clinical research studies due to the labor-intensive nature of the testing methods as well as the extensive time required to obtain results.33–35
Laboratory tests for HIV-1 infection are clinically used for diagnosing HIV-1 infection, monitoring progression of HIV infection and the response to antiretroviral therapy, and screening blood donors. The selection of these tests is highly dependent on the clinical situation, the patient population, and the specified purpose for the testing, as described in Table 19-8.8,51–54The following section briefly reviews each of the specific tests, but more comprehensive descriptions of the various commercial assays and their clinical applications can be found elsewhere.49,51,52,55
Recommended Laboratory Tests for the Diagnosis, Monitoring, and Blood Donor Screening for HIV
Diagnosis of HIV infection, including acute infection (excluding infants)
Risk of disease progression greater with HIV RNA >100,000 copies/mL
Response to antiretroviral therapy
Plasma HIV RNA viral load and CD4+ T cell count
Decision to start therapy should be based on laboratory results as well as clinical findings, patient interests, adherence issues, and risks of toxicity and drug interactions
Antiretroviral drug resistance testing
Phenotypic and genotypic resistance assays
Recommended for acute and chronic HIV infection on entry into care, treatment naïve patients, pregnant patients, and cases of virologic failure (testing recommended within 4 weeks of treatment discontinuation)
Not recommended for patients with HIV RNA <1,000 copies/mL
Infection with HIV affects both humoral and cell-mediated immune function. The humoral immune response results in the production of antibodies directed against HIV-specific proteins and glycoproteins. For most patients, antibodies to HIV-1 can be detected in the blood by 4 to 8 weeks after exposure to the virus. However, it may take up to 6 to 12 months in some patients. There are several tests currently available for the detection of HIV antibody in infected patients.
The methodology of EIA (commonly referred to as ELISA) is widely used as the initial screening test to detect HIV-specific antibodies.49,51,52,56 Like all immunoassays, ELISA is based on the concept of antigen and antibody reaction to form a measurable precipitate. The ability of ELISA to detect HIV antibodies during earlier infection has improved over recent years. Although less specific, first-generation ELISA tests introduced in 1985 were capable of detecting HIV-1 antibodies as soon as 40 days after exposure. Second-generation ELISA, introduced in 1987, incorporated recombinant antigens that increased specificity, sensitivity, and the ability to detect antibodies as soon as 34 days after infection. With the introduction of antigens from HIV-2, and the addition of antigens from HIV-1 groups M, N, and O and group M subtypes in the 1990s, specificity and sensitivity were improved. Third-generation ELISA, introduced in the mid-1990s, was redesigned as an antigen-antibody-antigen format, which dramatically improved sensitivity and specificity, and able to detect IgM and non-IgG antibodies as soon as 22 days after infection.
Testing for Human Immunodeficiency Virus While on Pre-Exposure Prophylaxis
Brian W. is an HIV-negative partner in a serodiscordant relationship who is currently taking tenofovir/emtricitabine for HIV pre-exposure prophylaxis (PrEP). He reports taking PrEP regularly without missed doses and does note occasional unprotected intercourse with his HIV-positive partner. He routinely gets tested for HIV every 3 months but is wondering if he needs to wait that long to get tested if he has an HIV sexual exposure from his partner. He has heard that it takes several months before HIV can be detected after exposure, and he would rather know as soon as possible.
QUESTION: How soon after exposure can current HIV tests detect infection?
DISCUSSION: The current tests for diagnosing HIV infection include the fourth-generation ELISA (or EIA) and a supplemental antibody differentiation immunoassay. Because fourth-generation assays combine the detection of HIV-1 and HIV-2 antibodies as well as HIV-1 p24 antigen, they are able to detect both acute and established HIV infection. The detection of HIV-1 p24 antigen by third-generation and fourth-generation immune assays is estimated to be between 14 and 20 days after infection. Thus, this patient does not need to wait until his scheduled routine HIV test and could seek earlier testing for reassurance. Additionally, he should be counseled on the importance of condom use while taking PrEP and on the decreased likelihood of transmission if his partner has an undetectable HIV viral load.
Speciation of Mycobacteria
Eva J. is a 42-year-old woman with HIV/AIDS, cerebral toxoplasmosis, and disseminated MAC. She was admitted to the hospital 10 days ago with abdominal pain and found to have a psoas abscess, which was drained. A preliminary culture result of this abscess stained positive for AFB with pending speciation. She has been taking tenofovir alafenamide/emtricitabine/bictegravir, ethambutol and azithromycin, and atovaquone for the past 4 months since her diagnosis of HIV/AIDS, toxoplasmosis, and disseminated MAC, respectively. Although she is taking medications with activity against mycobacteria, it would be best to tailor her antimycobacterial regimen to address the organism causing her psoas abscess.
QUESTION: What laboratory could be used to determine the species of this AFB?
DISCUSSION: The staining methods can detect the presence of mycobacteria in a clinical specimen but cannot differentiate between species of mycobacteria. Molecular techniques that use NA amplification (PCR) to detect M tuberculosis complex in acid-fast smear positive or negative respiratory specimens have been commercially developed to augment identification. Because of their high specificity and faster results, the CDC recommends performing NA amplification tests on respiratory specimens of patients suspected of having pulmonary TB. However, none of these assays are currently FDA-approved for the detection of Mycobacterium from nonrespiratory specimens such as in our patient case. For optimal cultivation and identification of mycobacteria, a combination of culture media, including at least one solid growth medium and one liquid growth medium, should be used during specimen processing to facilitate growth and optimize pigment production of the organism. Cultures for mycobacteria typically require prolonged incubation periods, sometimes up to 8 weeks, because most of the more common pathogens grow rather slowly. Rapidly growing mycobacteria such as M fortuitum, M chelonae, and M abscessus typically grow within 7 days on solid media, while slow growing mycobacteria such as M tuberculosis complex, MAC, M kansasii, and M marinum require 7 days to 7 weeks for growth. Hence, culture tubes or plates are examined weekly during the incubation period for the presence of mycobacterial growth. Colonies grown in culture are examined microscopically for characteristic colonial morphologic features, pigmentation, and growth rate; are subjected to biochemical tests; and are evaluated using rapid molecular detection methods, such as PCR methods mentioned earlier, DNA hybridization using DNA probes, and chromatographic methods, such as high-performance liquid chromatography or gas liquid chromatography, to detect mycobacterial lipids for definitive identification. The DNA probes can be used only with mycobacteria grown in culture (not directly on patient specimens), are highly sensitive and specific, and are commercially available for the rapid identification of M tuberculosis complex, M gordonae, M kansasii, and MAC. The molecular methods have replaced the use of biochemical tests in many laboratories because they provide more accurate identification in a significantly shorter time frame, within 14 to 21 days of specimen receipt as compared with several weeks or months using traditional identification methods. Recently, the use of MALDI-TOF MS for the identification of mycobacteria and other organisms was approved by the FDA. Although more expensive and not widely available, MALDI-TOF can provide faster time to speciation. However, the speciation of certain mycobacteria such as MTBC remains mostly unavailable with this method.
Fourth-generation ELISA detects the presence of both HIV-1 and HIV-2 antibodies and HIV-1 p24 antigen, which has reduced the detection period to approximately 15 days after infection, similar to the period of detection of p24 antigen.51 The improved sensitivity of fourth-generation assays has led to current recommendations by the CDC for their use as initial test in the diagnosis of HIV (Figure 19-1).51 Commercial fourth-generation ELISA kits used by most diagnostic laboratories can detect both HIV-1 and HIV-2 and have a sensitivity and specificity of >99%; however, false-positive and false-negative results can occur, particularly with previous generation assays. False-positive results have been reported with improper specimen handling (eg, heating) and in patients with autoimmune diseases, recent influenza vaccination, acute viral infection, alcoholic liver disease, chronic renal failure requiring hemodialysis, lymphoma, hematologic malignancies, and positive rapid plasma reagin (RPR) tests due to reactivity of the antibodies used for testing. Several causes have been identified for false-negative ELISA results and include the concomitant use of immunosuppressive therapy, the presence of severe hypogammaglobulinemia, and testing for HIV infection shortly after infection (acute HIV infection) or too late in the course of HIV infection.49,51
The results of a fourth-generation assay are reported as reactive (positive) or nonreactive (negative). If the initial result is nonreactive, no further testing for HIV antibodies is performed and the person is considered uninfected unless they are suspected of having acute HIV infection. When the initial result of a fourth-generation test is reported as reactive, a supplemental assay that differentiates between HIV-1 and HIV-2 antibodies must be performed. If the supplemental antibody differentiation test result is nonreactive or indeterminate, an HIV-1 nucleic acid test (NAT) must be performed to confirm the diagnosis of HIV-1 infection.51 Currently, laboratories can routinely use only the APTIMA HIV-1 RNA Qualitative Assay (Gen-Probe, Inc., San Diego, CA) as an aid in diagnosing HIV-1 (including acute or primary infection), and quantitative (viral load) HIV-1 RNA assays require a written order by a physician.57 As of this writing, no NAT has been approved for HIV-2 by the FDA, and the CDC recommends consultation with an expert in suspected cases of HIV-2 infection.
The western blot (WB) was the confirmatory test of choice for detecting HIV-specific antibody for many years.49,51,56,58 WB is a protein electrophoretic immunoblot technique that detects specific antibodies to HIV protein and glycoprotein antigens. The proteins and glycoproteins are detected by WB as “bands” and can be divided into Env (envelope) glycoproteins (gp41, gp120, gp160), Gag or nuclear proteins (p17, p24/25, p55), and Pol or endonuclease-polymerase proteins (p34, p40, p52, p68). Criteria for the interpretation of WB results have been published by different healthcare organizations and may vary depending on the issuing body. WB traditionally was considered to have a low level of sensitivity and high specificity, but newer data showed the WB misinterpreted most HIV-2 infections as being HIV-1.51 The WB test is technically difficult to perform, expensive, and associated with a relatively high rate of indeterminate results (4% to 20%) and has a long turnaround time (results are available in 1 to 2 weeks).49,51,52 Because of these limitations and the availability of more reliable assays, WB is no longer recommended for the diagnosis of HIV in the United States.51
Compared with WB, the use of an HIV-1/HIV-2 differentiation assay detects HIV-1 antibodies earlier, reduces indeterminate results, identifies HIV-2 infections, has a shorter turnaround for test results, and an overall lower cost.51 To date, only the Geenius HIV 1/2 Supplemental Assay (Bio-Rad Laboratories, Redmond, WA) differentiation ELISA remains available as an FDA-approved supplemental test following a reactive fourth-generation HIV-1 and HIV-2 antibody and HIV-1 p24 antigen initial assay.59 Differentiation assays are not without flaws, and an increase in the number of HIV-1–positive individuals with false-positive HIV-2 results (more than the total number of confirmed and probable HIV-2 cases) has called for the revision of the CDC-recommended HIV laboratory testing algorithm (Figure 19-1) by many experts.60
An assay that does not require supplemental HIV-1/HIV-2 antibody differentiation was approved by the FDA in July 2015. The Bio-Rad BioPlex 220 HIV Ag-Ab Assay (Bio-Rad Laboratories) simultaneously detects HIV antibodies and HIV-1 p24 antigen and provides separate antibody results for HIV-1 and HIV-2.59 Considered a fifth-generation assay, specimens reactive only to HIV-1 p24 antigen do not require antibody confirmation, and specimens reactive only to HIV antibodies do not require antigen confirmation.61 Because it eliminates supplemental antibody differentiation and antibody/antigen confirmatory procedures, broad implementation of this assay would require a change in the CDC HIV laboratory testing algorithm (Figure 19-1). As of June 2020, no changes in the testing algorithm have been announced or published by the CDC with regards to a fifth-generation assay.
Rapid human immunodeficiency virus screening tests
Technological advances have allowed for the development of rapid (eg, 30 minute) screening tests for the detection of HIV antibodies. The methodology of the rapid HIV antibody assays involves typically either membrane EIA or immunocytochemical assays. Currently, only seven FDA-approved rapid HIV-1 screening tests remain available on the market in the United States: OraQuick ADVANCE Rapid HIV-1/2 Antibody Test (OraSure Technologies, Bethlehem, PA); Reveal Rapid HIV-1 Antibody Test (MedMira, Halifax, Nova Scotia); INSTI HIV-1/HIV-2 Antibody Test (bioLytical Laboratories, Richmond, BC); Alere Determine HIV-1/2 Ag/Ab Combo (Alere Scarborough, Scarborough, ME); (Clearview Complete) SURE CHECK HIV 1/2 Assay, (Clearview) HIV 1/2 STAT-PAK Assay, and Chembio DPP HIV 1/2 Assay (Chembio Diagnostic Systems, Medford, NY).49,59 All currently approved tests display specificity and sensitivity similar to ELISA, require only basic training in test performance and interpretation, can be stored at room temperature, and require minimal specialized laboratory equipment.49,55 All assays, except for the Reveal Rapid HIV-1 Antibody Test, have received waivers under CLIA, which allow POC testing where dedicated laboratories are not available. POC tests are less complex to perform than traditional diagnostic tests and have minimal chance for error.49,55
Rapid HIV testing has proven useful for the detection of HIV infection in a number of clinical settings, such as during labor and delivery of pregnant women at high risk for HIV infection with unknown serostatus for the purposes of preventing perinatal transmission, facilities where return rates for HIV test results are low, and following occupational exposure to potentially HIV-infected body fluids (eg, through a needlestick) in which immediate decisions regarding postexposure prophylaxis are needed.49,55,58 It is important to note that rapid HIV tests are not recommended by the CDC for the diagnosis of HIV infection due to insufficient data, and a positive result from a rapid HIV test must be confirmed with a fourth-generation ELISA before the final diagnosis of HIV infection can be established. If the results of a rapid HIV test are negative, a person is considered uninfected. However, retesting should be considered in persons with possible exposure to HIV within the previous 3 months because the testing may have been performed too early in the infection to detect antibodies to the virus.49,53,55
Noninvasive human immunodeficiency virus-1 tests
Several tests have been developed for testing HIV antibodies from oral fluid or urine.49,51,58,59 The OraSure Western Blot Kit (OraSure Technologies, Bethlehem, PA) is a supplemental confirmatory test for the OraSure HIV-1 Oral Specimen Collection Device, which uses a cotton fiber pad that is placed between the cheek and lower gum for 2 minutes to collect oral mucosal transudate containing IgG. The OraSure HIV-1 oral device is FDA-approved and has a specificity of 99.4% (similar to EIA). The DPP HIV 1/2 Assay (Chembio Diagnostic Systems) detects antibodies against HIV-1 and HIV-2 by swabbing a flat pad against the upper and lower outer gums once and then inserting the sample collection pad into the developer solution vial. After 25 to 40 minutes, the results window indicates whether HIV antibodies have been detected. Two urine-based HIV-1 tests (ELISA and WB) have also been FDA-approved for use but are associated with lower sensitivity and specificity than the oral fluid testing. As with ELISA testing of blood samples, confirmatory testing is required for both types of tests if the initial results are positive. Noninvasive HIV-1 testing should be considered in persons who are unable to access healthcare facilities, have poor venous access, or are reluctant to have their blood drawn. The advantages of these tests include avoidance of blood drawing for sample collection, ease of use, low cost, and stability of samples for up to 3 weeks at room temperature.49,51,57
Home sample collection tests
As of June 2020, the OraQuick In-Home HIV Test (OraSure Technologies) remains the only FDA-approved assay available over-the-counter for at-home use.62 The test requires swabbing upper and lower gums with a collection stick that is inserted into a test solution for 20 minutes, after which time the results can be read. A line on the control and test areas indicates a positive result. The advantages of home sample collection tests include ready access to HIV-1 testing, convenience, lower costs, anonymity, and privacy.51,57
p24 Antigen Tests
A main structural core protein of HIV is p24, with levels of p24 antigen being elevated during the early stages of HIV infection. Testing for p24 antigen has diagnostic use during early infection when low levels of HIV antibody are present.49,52 The direct detection of HIV-1 p24 antigen can be performed using a plasma or serum EIA assay.51,58 Similar to antibody testing, positive results of the initial testing for HIV-1 p24 antigen must be retested in duplicate using the same EIA method. In addition, these results need to be confirmed with a neutralization assay due to the potential for p24 antigen testing to produce false-positive reactions resulting from interfering substances.49,51
The p24 antigen test may be used for screening blood donors, detecting growth in viral cultures, and serving as an alternative diagnostic test for HIV-1 in patients suspected of having acute HIV infection or infants <18 months of age born to HIV-infected mothers (Table 19-8).49,51,58 The advantages of the p24 antigen test include earlier detection of HIV infection compared with antibody testing (16 days versus 22 days), ease of performance, low cost, and specificity that approaches 100%. However, the DNA PCR assays and plasma HIV-RNA concentrations demonstrate greater sensitivity and have replaced the p24 antigen tests in many clinical situations.
Human Immunodeficiency Virus DNA Polymerase Chain Reaction
HIV DNA PCR is used for early detection of proviral HIV-1 DNA in a patient’s peripheral blood mononuclear cells. HIV DNA PCR is currently recommended for the diagnosis of HIV infection in infants (<18 months of age) born to HIV-infected mothers, and any clinical situations in which antibody tests are inconclusive or undetectable (Table 19-8).49,53 Antibody tests are not useful for diagnosing HIV infection in infants because maternal HIV antibodies can persist in the infant for up to 18 months after birth. Therefore, infants should be tested with the HIV DNA PCR or HIV RNA viral load test initially between 14 and 21 days of age, then at 1 to 2 months, and age 3 to 6 months.53 Negative tests at birth can be repeated at 14 days of life because the assay sensitivity is increased by 2 weeks of life. To confirm the diagnosis of HIV infection, a positive result at any sampling time needs to be confirmed by a second HIV virologic test. HIV infection may be excluded in infants with two or more negative HIV virologic results when initial testing occurred at age ≥1 month and the second testing occurred at age ≥4 months.53
The advantages of the HIV DNA PCR test include a high level of sensitivity and specificity (96% and 99% at ~1 month of age, respectively), the requirement for only a small volume of blood (eg, 200 µL), and the rapid turnaround time.49 The disadvantages include the expense, the high level of interlaboratory variability, and the availability of only one commercial assay (COBAS AmpliPrep/COBAS TaqMan HIV-1 Test; Roche Diagnostics, Indianapolis, IN), which is not currently FDA-approved for proviral DNA quantitation.49,53
Human Immunodeficiency Virus-RNA Concentration (Human Immunodeficiency Virus RNA Viral Load)
The accurate measurement of plasma HIV-RNA concentrations (also known as the HIV viral load) in conjunction with CD4+ T lymphocyte count has become an essential component in the management of patients with HIV-1 infection.8,49,53 These two laboratory tests provide the clinician with information regarding a patient’s virologic and immunologic status, which is needed to make decisions regarding the initiation or changing of antiretroviral therapy and to predict the risk of disease progression from HIV infection to AIDS. In addition, plasma HIV-RNA concentrations can assist in the diagnosis of HIV infection in selected clinical situations (Table 19-8).8,53
Methods that measure the amount of HIV-RNA in plasma include coupling reverse transcription to a DNA polymerase chain reaction (RT-PCR), identification of HIV-RNA with signal amplification by bDNA, and NA sequence-based amplification. Currently, there are five commercial assays approved by the FDA for clinical use: Amplicor HIV-1 Monitor version 1.5 (Roche Molecular Systems, Pleasanton, CA) in standard and ultrasensitive versions; Versant HIV-1 RNA 3.0 Assay (bDNA) (Siemens Healthcare Diagnostics, Tarrytown, NY); COBAS AmpliPrep/COBAS TaqMan HIV-1 versions 1 and 2 (Roche Diagnostics); and RealTime HIV-1 (Abbott Molecular, Des Plaines, IL).8,49,59 The results of these tests are expressed as the number of HIV copies/mL. Higher HIV-RNA levels (eg, >100,000 copies/mL) represent a substantial risk for disease progression. These assays differ in their dynamic ranges and lower limits of detection of HIV viral copies/milliliter of plasma. For example, the Versant assay has a lower limit of detection of <75 copies/mL whereas the COBAS AmpliPrep/COBAS TaqMan version 2 assay detection limit is <20 copies/mL. Some of these assays have different versions according to the degree of automation and simplicity. For instance, the Amplicor HIV-1 Monitor version 1.5 is a manual test; the COBAS Amplicor HIV-1 Monitor is semiautomated, and the COBAS AmpliPrep/COBAS Amplicor HIV-1 Monitor is automated. Additionally, the Amplicor HIV-1 Monitor and its variants exist as two FDA-approved assays—standard and ultrasensitive—due to limited dynamic range. The standard assay has a lower limit of detection of 400 copies/mL compared with the ultrasensitive assay, which has a limit of 50 copies/mL. It is recommended to use both assays concurrently when testing samples that fall outside of the dynamic range. Therefore, if a viral load is reported as “undetectable,” it signifies that the plasma HIV-RNA concentrations are below the lower limits of detection of the assay used.8,49 When performing plasma HIV-RNA viral load levels for each patient, the same laboratory and method should be used to minimize variation.
Once the diagnosis of HIV-1 infection has been confirmed, a plasma HIV-RNA level should be measured to assist in assessing antiretroviral treatment efficacy. Ideally, a plasma HIV-RNA level (and CD4+ T lymphocyte count) is measured on two separate occasions as the baseline measurement. Although antiretroviral therapy is recommended for everyone regardless of HIV-RNA levels and CD4+ T lymphocyte count, these values along with clinical findings and symptoms, the willingness of the patient to adhere to therapy, and the potential complications associated with therapy help decide whether treatment can be temporarily deferred and for how long when patients cannot initiate therapy right away. When initiating antiretroviral therapy, current guidelines recommend monitoring plasma HIV-RNA levels at the following time intervals: immediately before treatment initiation and 2 to 8 weeks after starting or changing antiretroviral drug therapy; 3 to 4 months following therapy initiation and every 3 to 6 months while on therapy; and any time when clinically indicated, including for patients who experience a significant decline in CD4+ T lymphocyte count.8 Testing HIV-RNA is not recommended during the period of an acute illness (eg, bacterial or P jirovecii pneumonia) or in patients who have been recently vaccinated because these circumstances may increase the viral load for 2 to 4 weeks. For patients receiving antiretroviral therapy, the goals of therapy include specific reductions in the HIV viral load measured in log reductions over a given time frame as well as achieving a viral load “below the limits of detection.” Changes in the amount of plasma HIV-RNA are often reported in log base 10 values. For example, a change from 10,000 to 1,000 copies/mL in a patient on antiretroviral therapy would be considered a 1-log decrease in viral load. With optimal therapy, plasma HIV-RNA levels should decrease by ≥1 log10 during the first 2 to 8 weeks after initiation of therapy and should continue to decline over subsequent weeks, with the ultimate goal of achieving an undetectable viral load (eg, <50 copies/mL) 16 to 24 weeks after initiation of therapy.8 However, every patient responds differently. The reader is encouraged to review the most recent HIV diagnostic and treatment guidelines because recommendations for different patient populations are continuously being modified as newer data become available.8,53,54
CD4+T Lymphocyte Count
The cell-mediated immune function effects of HIV infection are demonstrated by reductions in the CD4+T lymphocyte count. Flow cytometry can be used to identify the various subsets of lymphocytes by their cluster of differentiation (CD) of specific monoclonal antibodies to surface antigens. The CD4+ T lymphocytes are the helper-inducer T cells, whereas the CD8+ T lymphocytes are the cytotoxic-suppressor T cells. HIV infection causes a decrease in the total number of lymphocytes (particularly the CD4+ T lymphocytes) as well as changes in the ratios of the different types of lymphocytes. CD4+ T lymphocyte counts of <200 cells/mm3 (normal count 800 to 1,100 cells/mm3) or a CD4+ T lymphocyte percentage of <14% of the total lymphocyte count (normal 40% of total lymphocytes) are indicative of severe immunosuppression, placing the patient at risk for development of opportunistic infections.8 A CD4+/CD8+ ratio <1 is considered a hallmark of HIV infection, and lower ratios are indicative of immune senescence and associated with serious non-AIDS events and death.63 This ratio is not commonly used in clinical practice as it may reflect immune activation and not immune reconstitution but is often used in research in trying to characterize immune activation or senescence. Because of its limited clinical utility, measurement of the CD4+/CD8+ ratio is not routinely recommended.8
Current HIV treatment guidelines recommend initiation of antiretroviral therapy in all HIV-infected patients regardless of the plasma HIV RNA viral load or CD4+ T lymphocyte count.8 As stated earlier, the CD4+ T lymphocyte count is used in conjunction with the plasma HIV-RNA level to provide essential information regarding a patient’s virologic and immunologic status and risk of disease progression from HIV infection to AIDS and to determine whether to modify antiretroviral therapy or initiate chemoprophylaxis against opportunistic infections.8,53,54 Because of the significant impact on disease progression and survival, most guidelines recommend monitoring CD4+ T lymphocyte counts at baseline and every 3 to 6 months to assess the immunologic response to treatment in patients starting or modifying antiretroviral treatment, or even yearly or longer in people on stable therapy.8
Phenotypic and Genotypic Assays for Antiretroviral Drug Resistance
Resistance of HIV-1 to antiretroviral drugs is an important cause for treatment failure. Genotypic assays use gene sequencing or probes to detect resistance mutations in genes of circulating RNA known to confer drug resistance in the RT, protease (PR), envelope, and integrase genes of HIV-1. Two genotypic assays have been approved by the FDA: TruGene (Siemens Healthcare Diagnostics, Tarrytown, NY) and ViroSeq (Celera Diagnostics, Alameda, CA).50,59Phenotypic assays measure the quantity of viral replication in the presence of various concentrations of antiretroviral agents. Sequences from the RT, PR, envelope, and integrase genes of the patient’s HIV virus are inserted into a wild-type virus in the laboratory. The concentration of the drug needed to inhibit 50% of viral replication (IC50) is reported. The ratio of IC values for the test and reference viruses is calculated and used to report the quantitative fold-increase in resistance of each antiretroviral agent. The interpretation of results from both assays is complex and requires expert knowledge and consultation.8,49
The current guidelines recommend drug resistance testing for patients with acute and chronic HIV infection when they enter into care, regardless of the decision to initiate antiretroviral therapy; pregnant women prior to antiretroviral therapy initiation; women contemplating or entering pregnancy with a detectable HIV-RNA level during treatment; patients with virologic failure while receiving antiretroviral therapy; and patients with suboptimal suppression of plasma HIV-RNA level after the initiation of antiretroviral therapy.8,54
Genotypic assays are typically recommended for resistance testing in the treatment of naïve patients and pregnant women. In the setting of virologic failure, current guidelines recommend performing resistance testing immediately after or within 4 weeks of discontinuation of antiretroviral therapy for optimal results. Resistance testing is discouraged in patients with plasma HIV-RNA levels <1,000 copies/mL or patients who have been off antiretroviral therapy for extended periods of time because the population of mutant virus may not be sufficient for detection. A newer generation of genotypic assays that analyze proviral DNA and do not depend on detectable HIV-RNA can be considered in patients with undetectable HIV-RNA.8
The advantages and disadvantages of genotypic and phenotypic assays for the detection of HIV-1 resistance have been previously described.51 The genotypic assay may be preferred over the phenotypic assay because of availability, clinical utility, faster turnaround time (1 to 2 weeks versus ≥2 weeks), and lower cost.50 Because they are labor-intensive and more expensive than genotypic assays, phenotypic assays are generally reserved for cases in which antiretroviral resistance cannot be predicted based on known genetic mutations alone. However, both assays are complex, technically demanding, and expensive and are not routinely performed in most clinical laboratories. Proviral DNA genotypic assays may be useful in people with a history of multiple treatments, treatment failures, or no available genotypic resistance test results who are virally suppressed on a complex antiretroviral regimen, which they need to modify due to drug–drug interactions, pill fatigue, or toxicity.8 Proviral DNA genotypic assays may not be useful in people on their first or second antiretroviral regimen without prior failures or with available prior genotypic testing results.
Additional Laboratory Testing for Patients with Human Immunodeficiency Virus
Coreceptor tropism assays
After attachment of HIV to the CD4+ T lymphocytes, fusion of the virus and CD4+ cell membranes involves binding to a coreceptor molecule. The two coreceptors used by HIV are chemokine coreceptor 5 (CCR5) and CXC coreceptor (CXCR4). The recently approved antiretroviral agent maraviroc is a CCR5-coreceptor antagonist that prevents the entry of HIV into the CD4+ cell by binding to the CCR5 receptor. Most acutely or recently infected patients harbor the CCR5-tropic virus, while untreated patients with advanced disease and those with disease progression shift from CCR5-tropic to CXCR4-tropic or both (dual-tropic or mixed-tropic). Treatment-experienced patients with high levels of drug resistance are more likely to harbor dual-tropic or mixed-tropic virus. Current HIV treatment guidelines recommend performing a coreceptor tropism assay when considering the use of a CCR5-coreceptor antagonist or in the event of virologic failure during maraviroc therapy.8 Currently, a phenotypic assay, Trofile (Monogram Biosciences, South San Francisco, CA), and a genotypic tropism assay, HIV-1 Coreceptor Tropism (Quest Diagnostics, Madison, NJ), are available in the United States.84
Abacavir, a nucleoside reverse transcriptase inhibitor, is associated with a potentially life-threatening hypersensitivity reaction reported in 5% to 8% of patients in clinical trials. The hypersensitivity reaction appears to occur more frequently in white patients (5% to 8%) than black patients (2% to 3%) and is associated with the presence of MHC class I allele HLA-B*5701. Treatment guidelines recommend screening patients for the presence of HLA-B*5701 prior to the initiation of abacavir-containing regimens in areas in which the screening test is available, and patients with a positive result should not receive abacavir.8 However, the initiation of abacavir therapy can be reasonably considered using clinical judgment with appropriate monitoring and extensive patient education about the signs and symptoms of the hypersensitivity reaction in settings in which the HLA-B*5701 screening test may not be available.
Mycobacteria are nonmotile, nonspore-forming, aerobic bacilli that continue to cause infection as well as significant morbidity and mortality, especially in developing countries.64–72 Currently, >100 species of mycobacteria have been identified, with only a number of species causing infection in humans, including M tuberculosis, M leprae, M avium complex, M kansasii, M fortuitum, M chelonae, M abscessus, and M marinum.11,64–66 Depending on the species, mycobacteria can be nonpathogenic, pathogenic, or opportunistic and, therefore, may cause infection in both immunocompetent and immunocompromised hosts. Table 19-9 lists the most common pathogenic mycobacteria species with their typical associated infections and environmental sources.11,64–69
Mycobacteria are generally divided into two groups based on epidemiology and spectrum of disease: (1) the M tuberculosis complex (MTBC), including the species M tuberculosis, M bovis, M bovis bacille Calmette-Guerin (BCG), M africanum, and M microti; and (2) nontuberculous mycobacteria (NTM; also referred to as mycobacteria other than tuberculosis), which include all other species of mycobacteria.11,64,66M tuberculosis is the most clinically significant mycobacteria and is the causative organism of tuberculosis (TB). The incidence of TB in the United States declined between 1953 (when it became a notifiable disease) and 1985 as a result of improved diagnostic methods, enhanced public health efforts to isolate patients infected with TB, and the introduction of effective antimycobacterial agents.65,66,70 This decline in TB cases led experts to predict the elimination of the disease by 2010. However, between 1986 and 1992, an increase in the incidence of TB was observed in the United States due to deterioration of the TB public health programs, emergence of the HIV epidemic, increase in immigration to the United States, and emergence of MDR-TB.65,70 Since 1992, the number of cases of TB in the United States has steadily declined due to improved public health control strategies.71 Despite advances in medical care and treatment, TB continues to be one of the most common infectious diseases worldwide. The World Health Organization estimated that approximately 10 million persons were afflicted with TB worldwide in 2018, and there were more than 1.4 million deaths that year.72 Because TB can be transmitted from person to person, rapid diagnosis is necessary to decrease the spread of infection.70
Identification of Mycobacteria
Mycobacteria possess several unique characteristics that contribute to the difficulties with the growth, identification, and treatment of these organisms. The cell wall of mycobacteria is complex and composed of peptidoglycan, polypeptides, and a lipid-rich hydrophobic layer.11,66 This cell wall structure confers a number of distinguishing properties in the mycobacteria, including (1) resistance to disinfectants and detergents, (2) the inability to be stained by many common laboratory identification stains, (3) the inability of mycobacteria to be decolorized by acid solutions (a characteristic that has given them the name of acid-fast bacilli [AFB]), (4) the ability of mycobacteria to grow slowly, and (5) resistance to common anti-infective agents.11,64,66 These characteristics have led to the continuous modification and improvement of laboratory practices used in the identification and diagnosis of mycobacterial infections. Many of these laboratory practices involve specialized staining techniques, growth media, identification techniques, environmental conditions (BSL-3 facilities for M tuberculosis), and susceptibility testing methods that may be unavailable in some clinical laboratories.66,68
The ability to accurately cultivate mycobacteria is highly dependent on the appropriate selection and collection of biologic specimens for staining and culture.11,64–66 Because different mycobacteria are capable of causing a number of infections (Table 19-9), the following biologic specimens may be submitted for mycobacterial culture based on the site of infection: respiratory tract secretions (eg, expectorated or induced-sputum and bronchial washings) or gastric lavage specimens for the diagnosis of pulmonary TB; CSF for the diagnosis of TB meningitis; blood for the diagnosis of disseminated M avium complex (MAC) infection; stool for the diagnosis of disseminated MAC infection; and urine, tissue, exudate, or wound drainage, bone marrow, sterile body fluids, lymph node tissue, and skin specimens for infection due to any mycobacteria.65–68 For the diagnosis of pulmonary TB, several early morning expectorated or induced-sputum specimens are recommended to enhance diagnostic accuracy. Biologic specimens for mycobacterial culture should be immediately processed according to specified guidelines to prevent the overgrowth of bacteria that may also be present in the specimen and should be concentrated to enhance diagnostic capability.65,66,68 Similar to the processing of specimens for bacterial culture, biologic specimens submitted for mycobacterial culture should be stained for microscopic examination and plated for culture. The staining techniques and culture media, however, are somewhat different because mycobacteria are poorly visualized in the Gram stain (they do not reliably take up the dyes and are referred to as acid-fast) and take longer to grow than conventional bacteria.
Staining with subsequent microscopic examination for mycobacteria is a rapid diagnostic test that involves the use of stains that are taken up by the lipid and mycolic acid components in the mycobacterial cell wall.11,66 Several acid-fast stains are available for the microscopic examination of mycobacteria, including carbolfuchsin-based stains that are viewed using light microscopy (Ziehl-Neelsen or Kinyoun method) and fluorochrome stains (auramine-rhodamine) examined under fluorescence microscopy that are thought to be more sensitive tests, especially on direct specimens.11,64–67,73 The sensitivity of the staining method is highly dependent on the type of clinical specimen, the species of mycobacteria present, the technique used in specimen processing, the thickness of the smear, and the experience of the laboratory technologist.11,65,66 The staining methods can detect the presence of mycobacteria in a clinical specimen but cannot differentiate between species of mycobacteria. Therefore, several molecular techniques have been commercially developed to augment identification, which use NA amplification (PCR) to detect M tuberculosis complex in acid-fast smear–positive respiratory specimens (Amplicor Mycobacterium tuberculosis Test by Roche Diagnostics, and the Amplified Mycobacterium tuberculosis Direct Test by Gen-Probe) or acid-fast smear–negative respiratory samples (Amplified M tuberculosis Direct Test by Gen-Probe).11,64,11,66,68,73 Both of these tests display high sensitivity in detecting the presence of M tuberculosis complex in smear-positive respiratory specimens (>97%).11,66,73 Because of their high specificity and the rapid availability of results, the CDC recommends performing NA amplification tests on respiratory specimens of patients suspected of having pulmonary TB.12,74 However, none of these assays are currently FDA-approved for the detection of Mycobacterium from nonrespiratory specimens.12,74,75
For optimal cultivation and identification of mycobacteria, a combination of culture media, including at least one solid growth medium and one liquid growth medium, should be used during specimen processing to facilitate growth and optimize pigment production of the organism.11,66,67,69,70,73 The preferred commercially available solid growth media for the cultivation of mycobacteria include an agar-based medium such as Middlebrook 7H10 or an egg-based medium such as Lowenstein-Jensen. Several liquid growth media systems are available for culture of mycobacteria, some of which use continuous automated monitoring systems for the detection of mycobacterial growth.11,66,67,69 The liquid growth media systems often provide more rapid isolation of AFB compared with conventional solid media, with results within 10 days as compared with 17 days or longer using solid growth media.65–68,73 The most commonly used semiautomated systems with liquid growth media include the MB Redox (Heipha Diagnostika Biotest, Heidelberg, Germany), BACTEC 460 TB system (BD), the Septi-Chek AFB System (BD), and the Mycobacteria Growth Indicator Tube (BD).11,66,69,73 The most commonly used automated, continuous monitoring systems with liquid growth media include the ESP Culture System II (Trek Diagnostics, Westlake, OH), the BACTEC 9000 MB System (BD), the MB BacT/Alert System (bioMérieux, Marcy-l’Étoile, France), and the BACTEC MGIT 960 (BD Biosciences, Sparks, MD).11,66,69,73 Once growth is detected in the liquid media systems, an acid-fast stain is performed on the specimen to confirm the presence of mycobacteria, with subsequent subculture onto solid media.
The optimal growth conditions of mycobacteria depend on the species; therefore, the clinical laboratory should follow a standardized procedure outlining the process that should be used to enhance cultivation of the suspected Mycobacterium from the submitted clinical specimen based on the suspected site of infection. The optimal conditions for incubation of mycobacterial cultures are 28°C to 37°C in 5% to 10% CO2 for 6 to 8 weeks, depending on the organism.11,66,68,69 Cultures for mycobacteria typically require prolonged incubation periods, sometimes up to 8 weeks, because most of the more common pathogens grow rather slowly. Rapidly growing mycobacteria such as M fortuitum, M chelonae, and M abscessus typically grow within seven days on solid media, while slow-growing mycobacteria such as M tuberculosis complex, MAC, M kansasii, and M marinum require 7 days to 7 weeks for growth.11,67,69 Therefore, culture tubes or plates are examined weekly during the incubation period for the presence of mycobacterial growth.
Colonies grown in culture are examined microscopically for characteristic colonial morphologic features, pigmentation, and growth rate, are subjected to biochemical tests, and are evaluated using rapid molecular detection methods, such as PCR methods mentioned earlier, DNA hybridization using DNA probes, and chromatographic methods, such as high-performance liquid chromatography or gas liquid chromatography, to detect mycobacterial lipids for definitive identification.11,66–69 The DNA probes can be used only with mycobacteria grown in culture (not directly on patient specimens), are highly sensitive and specific, and are commercially available for the rapid identification of M tuberculosis complex, M gordonae, M kansasii, and MAC.11,66–69 The molecular methods have replaced the use of biochemical tests in many laboratories because they provide more accurate identification in a significantly shorter time frame, within 14 to 21 days of specimen receipt, as compared with several weeks or months using traditional identification methods.11,65 For example, the Xpert MTB/RIF Assay (Cepheid, Sunnyvale, CA) was cleared by the FDA in 2013 for the identification of MTBC and rifampin resistance mutations via DNA RT-PCR from respiratory specimens, and was adopted by the CDC and WHO as part of the initial diagnostic tools.76,77 Later in 2017, the FDA cleared the use of MALDI-TOF MS for the identification of mycobacteria and other organisms with the Vitek MS system (bioMérieux, Inc, Marcy-l’Étoile, France).78 Although this method of identification is rapid, there are still significant disadvantages with respect to cumbersome extraction protocols, inability to identify MTBC species, and high initial set up and operation costs.79
Susceptibility Testing of Mycobacteria
The choice of antibiotic or antimycobacterial agent to use in the treatment of mycobacterial infection depends on the species of mycobacteria involved. It is important for the clinician to have an understanding of the typical susceptibility patterns of specific mycobacterial species, the current treatment guidelines outlining which and how many drugs to use, and the methodology available for drug susceptibility testing for the particular mycobacterial species being treated.
Standardized guidelines have been published for the susceptibility testing of M tuberculosis complex.80 Susceptibility testing is currently recommended on the initial isolate of all patients with M tuberculosis complex infection, on isolates from patients who remain culture positive after 3 months of appropriate therapy, and on isolates from patients who are not clinically responding to therapy.11,66,70,80 Susceptibility testing of M tuberculosis complex is initially performed with primary antituberculous agents such as isoniazid (using two concentrations, 0.2 and 1 mcg/mL), rifampin, ethambutol, and pyrazinamide. However, if resistance to any of these first-line drugs is detected, susceptibility testing should be subsequently performed using second-line drugs, including streptomycin, a higher concentration of ethambutol (10 mcg/mL), ethionamide, capreomycin, ciprofloxacin, ofloxacin or levofloxacin, kanamycin, p-aminosalicylic acid, and rifabutin.66,68,80
Susceptibility testing of M tuberculosis complex can be performed directly using mycobacteria from a smear-positive specimen (direct method) or using mycobacteria isolated from culture (indirect method).11,66,68 The direct method of mycobacterial susceptibility testing produces faster results but is less standardized so that susceptibility testing is usually performed using isolates grown in culture.66,68 Four conventional methods are used worldwide for determining the susceptibility of M tuberculosis isolates to antituberculous agents, including the absolute concentration method, the resistance ratio method, the agar proportion method, and the agar proportion method using liquid medium (commercial radiometric, nonradiometric, or broth systems, including the BACTEC 460TB System [BD]; BACTEC MGIT 960 [BD]; ESP Culture System II [Trek Diagnostics, Westlake, OH]; and MB/BacT-Alert 3D [bioMérieux]). The agar proportion method and the commercial broth systems are the methods commonly used for mycobacterial susceptibility testing in the United States.
The agar proportion method is a modified agar dilution test that evaluates the extent of growth of a standardized inoculum of M tuberculosis in control and drug-containing agar medium. The organism is considered resistant if growth is >1% on the agar plate containing critical concentrations of the antituberculous drug.80 The critical concentration for each drug represents the lowest concentration of the drug that inhibits 95% of wild-type M tuberculosis strains that have never been exposed to the drug.66,80 Susceptibility results using the agar proportion method are typically available 21 days after the plates have been inoculated.
The commercial susceptibility testing systems use liquid growth medium in which the growth of the organism is measured in the presence and absence of antituberculous drugs.65–69,73,80 These systems provide rapid susceptibility results for the primary antituberculous agents but cannot be used for susceptibility testing of second-line agents. Commercial susceptibility tests are recommended over agar proportion methods because the results are often available within 5 to 7 days after inoculation and can help guide appropriate therapy without the unnecessary delay of the agar proportion method.65,68,69,80
Several other susceptibility testing methods are currently being evaluated for drug susceptibility testing of M tuberculosis. Many antimycobacterial drugs are now available in E-test strips, including streptomycin, ethambutol, isoniazid, and ethionamide.11,73,80 Several studies have evaluated their performance as compared with the commercial methods; however, further studies are needed to validate the use of the E-test as a suitable, alternative susceptibility testing method. In addition, newer molecular methods for drug susceptibility testing of M tuberculosis are currently being evaluated that are easier to perform and produce more reliable results in a shorter period.11,66–69 These methods include PCR amplification, DNA sequencing line probe assays, and reverse hybridization-based probe assays for the detection of specific drug-resistance mutations. As stated earlier, the use of the Xpert MTB/RIF Assay (Cepheid) for the identification of MTBC and the rifampin resistance mutations in the rpoB gene is recommended as part of initial diagnosis of MTBC but does not invalidate the need for additional molecular and traditional growth methods of susceptibility testing against other first- and second-line agents. Further study, however, is warranted before newer susceptibility testing methods can replace conventional methods.
Guidelines for susceptibility testing of NTM have recently been published by the CLSI, in which testing is recommended on the initial isolate for clinically significant isolates (blood, tissue) that display variability in susceptibility to antituberculous drugs or for organisms that may be associated with acquired resistance.80 The guidelines contain protocols for the susceptibility testing of rapidly growing NTM (M fortuitum, M chelonae, and M abscessus) and slow-growing NTM (MAC, M kansasii, and M marinum), including the recommended methodology and drugs to be tested for each organism.11,66,68,80 Standard broth microdilution should be used to evaluate the susceptibility of any clinically significant, rapidly growing NTM.67,80 Drugs that may be considered for susceptibility testing include amikacin, cefoxitin, ciprofloxacin, clarithromycin, doxycycline, imipenem, sulfamethoxazole, trimethoprim–sulfamethoxazole, and tobramycin (for M chelonae only).67,80 Susceptibility testing of MAC is recommended for clarithromycin only using a broth-based method (macrodilution or microdilution) for initial blood or tissue isolates in patients with disseminated infection, for clinically significant isolates from patients receiving previous or current macrolide therapy, for isolates from patients who develop bacteremia while receiving macrolide prophylaxis, and for isolates from patients who relapse while receiving macrolide therapy.67,80 Regarding the other slow-growing NTMs, susceptibility testing of M kansasii should be routinely performed only on rifampin using the commercial radiometric systems, broth microdilution, or the modified proportion method, while routine susceptibility testing of M marinum is not recommended.67,80 The integration of molecular and culture-based methods for both MTBC and NTM for better estimation of drug efficacy is also addressed in CLSI guidelines.
The Mantoux test or tuberculin skin test (TST) is one test available for the detection of latent TB (LTBI) and involves the intradermal injection of a purified protein derivative (PPD) of the tubercle bacilli, which is obtained from a culture filtrate derived by protein precipitation.68,81 Injection of the PPD into individuals previously exposed to TB elicits a delayed hypersensitivity reaction involving T cells that migrate to the area of intradermal injection (usually the dorsal aspect of the forearm), inducing the release of lymphokines that produce induration and edema within 48 to 72 hours after injection. The diameter of induration is measured between 48 and 72 hours after injection by a healthcare professional.68,81 Published guidelines are available for interpretation of the TST reaction based on the size of the induration and clinical and demographic characteristics of the patient. An induration of ≥5 mm is considered positive in persons at high risk of developing TB disease, including HIV infected patients; patients receiving immunosuppressive therapy including tumor necrosis factor blocking agents; patients who have been recently exposed to a person with TB; and patients with an abnormal chest radiographic consistent with prior TB.64,68,81 An induration of ≥10 mm is considered positive in patients who are not immunocompromised but who are at high risk for having LTBI or for progressing from LTBI to TB, such as persons born in high TB burden countries; injection drug users; residents and employees of high-risk settings (eg, prisons, healthcare facilities, and mycobacteria laboratory personnel); persons with chronic medical conditions of high risk (eg, diabetes, silicosis, and chronic renal failure), and children younger than 5 years of age. An induration of ≥15 mm is considered positive in persons with no risk factors or at low risk for developing active infection with TB.
A two-step TST is recommended by the CDC in certain populations (initial skin testing of newly hired healthcare workers without a documented negative TST within the past 12 months and persons expected to undergo serial screening for TB, such as residents and staff of long-term care facilities) to identify individuals with past TB infection whose delayed-type hypersensitivity to tuberculin has diminished over time.82,83 The first TST is administered as described previously, with a second TST administered following the same procedure 1 to 3 weeks later in persons with a negative initial test result.82 The premise behind the administration of two TSTs in these settings is to delineate between past TB infection or BCG vaccination from recent conversion/infection. That is, the first injection stimulates (boost) the delayed hypersensitivity response in a patient with previous TB infection or BCG vaccination and the second TST then elicits a positive reaction.82,83
Blood Assay for Mycobacterium Tuberculosis
Two in vitro diagnostic tests using whole blood have become recently available in the United States for the detection of latent M tuberculosis infection: (1) a blood assay for M tuberculosis (BAMT), the QuantiFERON-TB Gold In-Tube test (Cellestis, Chadstone, Victoria, Australia) and (2) T-SPOT.TB (Oxford Immunotec, Oxford, UK).65,68 These tests use ELISA to measure the amount of interferon-γ released from sensitized lymphocytes from prior exposure to M tuberculosis following overnight incubation with PPD from M tuberculosis and control antigens.68,84 Because the tests use peptide antigens from M tuberculosis, they have more specificity than the TST for the diagnosis of latent M tuberculosis infection and do not produce false-positive results in patients with previous BCG vaccination or infection due to NTM.82,84 The results of BAMT testing are stratified according to risk for TB infection (like the TST) but are not influenced by the subjectivity of reader bias or error, as may be seen with the TST. Because these tests are unable to distinguish between active or latent infection, the exact role of the BAMT and the TST in the diagnosis of latent TB infection are unclear and currently under investigation. However, the BAMT test can be used to assist in the diagnosis of latent TB infection in high-risk populations, such as recent immigrants from high-prevalence countries, injection drug users, inmates, prison employees, and healthcare workers at high risk for exposure to TB.82,84 The BAMT test can also be considered the initial and serial screening test for latent TB infection in healthcare workers and military personnel.82,84 It is important to note that the FDA has approved these tests as aids in the diagnosis of M tuberculosis infection and are intended to be used in conjunction with other diagnostic techiques.84
New testing methodologies and diagnostic guidelines continue to become available for the recovery, identification, and susceptibility testing of fungi, viruses, and mycobacteria. These processes created new opportunities and challenges for clinical microbiology laboratories with regard to what diagnostic tests offer and for which pathogens. Introduction and rapid acceptance of MALDI-TOF MS has allowed proteomics to be applied for the rapid identification of fungi and mycobacterial pathogens. Rapid antigen tests, NAATs, and serology have revolutionized diagnostic microbiology and impacted the need for traditional staining and culturing procedures, particularly in viral infections. A variety of molecular diagnostic assays allowed laboratory testing of SARS-CoV-2 to expand and become rapidly available during the recent global pandemic. Accurate SARS-CoV-2 NAATs have been critical for viral detection, public health quarantine decisions for preventing transmission, infection control measures, and patient management and treatment guidelines. Multiplex NAAT tests have become commercially available with syndromic panels that include fungal, viral, and bacterial pathogens commonly associated with respiratory tract infections, gastrointestinal pathogen detection, and central nervous system infections. Rapid POC testing for viral infections such as RSV, influenza, and HIV are allowing the latest diagnostic technology to move from the clinical laboratory setting to outpatient clinics, emergency departments, and healthcare offices. Genotypic and phenotypic resistance testing is regularly performed for HIV infections. Susceptibility and resistance testing for mycobacterial, fungal, and other viral pathogens continue to expand. Not all laboratories have broad testing capabilities with the latest technologies. It is essential to consult your local laboratory and appreciate the current diagnostic methods being offered, which tests are referred to external reference and/or public health laboratories, and the likely turnaround times in reporting results.
1. Matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF MS) has become a proteomic methodology for the identification of bacterial, fungal, and mycobacteria colonies on a variety of laboratory media. How does MALDI-TOF MS work for identification of fungi? What advantages does MALDI-TOF MS offer compared with traditional identification methods of fungi? What are the limitations of using MALDI-TOF MS for identification of fungi?
ANSWER: A pure colony of fungus is needed to produce a mass spectrum fingerprint characteristic of the organism. A colony of fungal cells is spread across the conductive metallic target plate (“spot”) of the MALDI-TOF MS. The cells may or may not be treated (eg, with formic acid) on the target plate, and the spot is overlaid with a matrix and air dried. This plate is loaded into the ionization chamber of the mass spectrometer and ultraviolet laser pulses hit the target spot, causing the sample to be vaporized (forming ions). A cloud of desorbed and ionized molecules is released to be accelerated by a high-voltage electric charge and fly through the time-of-flight (TOF) tube (or field-free drift region). The ions strike the detector at the other end of the flight tube (smaller ions travel faster than larger ions) and masses of the ions are determined from the time it took them to travel the length of the flight tube. A mass spectral signature is determined by mass-to-charge ratio (essentially molecular weight given MALDI predominantly produces singly charged ions) and the number of ions with a particular size striking the detector (intensity). Mass spectrometry identifies the protein fingerprints and a direct comparison of the spectral pattern of the organism in question is made with commercial or supplemental database patterns of different microorganisms. Depending on the database, it may be possible to identify the genus, species, and/or strain levels of the detected microbe.
The MALDI-TOF MS approach offers multiple advantages, including fast results, affordability, ease of use, and a highly accurate identification of fungal colonies to allow earlier decision-making of optimal antifungal therapy. Turnaround time is approximately 3 to 5 minutes for batch runs (compared with 1.5 days for traditional methods). This method is technically simple to perform, has minimal waste disposition, and has low reagent cost per test (although acquisition of the initial instrumentation system is rather costly). The method is considered robust and reliable because of an extremely low level of false identification of yeast and several filamentous fungi. It can be particularly helpful when morphology-based identification is not possible (ie, atypical morphology, lack of sporulate, lengthy identification processes) or if the phenotypic result is confusing. MALDI-TOF MS can extend its use beyond organism identification to include the detection of drug resistance and biomarkers. It is also a reliable and reproducible method for growth-based antifungal susceptibility testing of yeast (eg, fluconazole and echinocandin susceptibility testing of Candida) although it is labor-intensive and time-consuming.
Some of the major limitations of MALDI-TOF MS for identification of fungi center around reference mass spectral databases, specimen requirement, possibilities of errors, and the updates of taxonomy that are regularly needed. Most databases for MALDI-TOF MS are proprietary (not publicly available) and differences exist between manufacturers, including identification algorithms and instrumentation. In addition, limited commercial databases, variability in the spectra because of structural features of mold colonies, and lack of standardized processes has constrained the routine use of mold identification (except Aspergillus spp.) in some clinical laboratories. Widely published user-developed or in-house fungal databases can be used; however, applying and/or creating such supplemental databases may be difficult for some clinical laboratories. Another disadvantage is the requirement of organism recovery on culture media before sample preparation can begin for MALDI-TOF MS. Ongoing evaluations have had inconsistent results for organism detection directly from clinical samples. Finally, there is a learning curve for users of this system, including colony testing (eg, homogeneity of the smear, amount of organism and placement on target plate location, minimizing the use impure colonies, cleanup procedures of reusable plates) and minimizing sources of errors.
2. In vitro testing for circulating galactomannan can be used as a diagnostic tool for invasive aspergillosis. What is galactomannan? Which fungal pathogen is it most useful as a diagnostic aid? What type of assay is used to detect galactomannan in the United States? Outline the precautions to consider when interpreting the results from galactomannan testing.
ANSWER: Galactomannan is a polysaccharide present as a major cell wall component of the mold Aspergillus. Concentrations of galactomannan are released during the growth phase of the infection, with the highest concentrations occurring during the terminal phases of aspergillosis.
The Platelia Aspergillus Ag is an EIA sandwich microplate assay cleared by the FDA for the detection of Aspergillus galactomannan antigen in serum and BAL fluid. The assay uses a rat monoclonal antibody (EBA-2) directed to bind Aspergillus galactomannan antigen. The antibody binds to the antigen and serves as the detector for the antigen in the conjugate reagent (peroxidase-linked monoclonal antibody). The performance of the galactomannan assay has been similar for adults and children.
Serum galactomannan can often be detected 7 to 14 days before other diagnostic tests become apparent. The test should be used in conjunction with other diagnostic procedures (eg, microbiological culture, histologic examination of biopsy samples, radiographic evidence) as an aid in the diagnosis of invasive aspergillosis. The combined use of galactomannan antigen testing and NAAT (ie, PCR) has been shown as a successful approach in the earlier diagnosis of invasive aspergillosis. A negative result does not rule out the diagnosis of invasive aspergillosis. Repeat testing is recommended if the result is negative but invasive aspergillosis is suspected. Patients at risk for invasive aspergillosis should have a baseline serum tested and should be monitored twice a week for increasing galactomannan antigen levels. Galactomannan antigen concentrations may be useful in the assessment of therapeutic response (eg, antigen concentrations decline in response to antifungal therapy).
Other genera of fungi such as Penicillium and Paecilomyces have shown reactivity with the monoclonal antibody used in the assay. Specimens containing Histoplasma antigen may cross-react in the Aspergillus galactomannan assay. The specificity of the assay for Aspergillus species cannot exclude the involvement of other fungal pathogens (eg, Fusarium, Alternaria, and Mucorales). The assay may exhibit reduced detection of galactomannan in patients with chronic granulomatous disease and Job syndrome.
Certain foods (eg, pasta, cereals, rice, cow’s milk) contain galactomannan. It is thought that damage to the gut wall by cytotoxic therapy, irradiation, or graft-versus-host disease enables translocation of the galactomannan from the gut lumen into the blood, which may result in the high false-positive rate of this assay. Antibiotics such as piperacillin–tazobactam, amoxicillin, and/or amoxicillin–clavulanate (eg, drugs derived from the genus Penicillium) have been demonstrated to cross-react with monoclonal antibody in the assay. False-positive galactomannan results have occurred in patients receiving Plasma-Lyte, either for intravenous hydration or BAL. The concomitant use of antifungal therapy in some patients with invasive aspergillosis may result in reduced sensitivity of the assay or false-negative results.
3. In vitro testing for circulating (1,3)-β-d-glucan can be used as a diagnostic tool for several invasive fungal pathogens. What is (1,3)-β-d-glucan? Briefly explain the type of assay used to detect (1,3)-β-d-glucan in the United States. Outline the pathogens this test can detect and what precautions need to be considered when interpreting the results from (1,3)-β-d-glucan testing.
ANSWER: The unique composition of the fungal cell wall makes it particularly well suited to be a nonmolecular biomarker test. A broad range of pathogenic molds and yeasts have the polysaccharide (1,3)-β-d-glucan present in their cell wall. Concentrations of (1,3)-β-d-glucan are released into the blood of patients with several types of invasive fungal infections.
In the United States, Fungitell assay (Associates of Cape Cod Inc., East Falmouth, MA) is widely available and is the only FDA-approved screening test for detecting (1,3)-β-d-glucan in serum. There are several commercially available assays outside the United States. This test uses the coagulation pathway derived from the Limulus horseshoe crab as the detector method given (1,3)-β-d-glucan can activate factor G in this coagulation cascade. Optimal sensitivity is usually obtained by testing patients twice weekly. False-positive results have been associated with the administration of intravenous immunoglobulins or albumin, hemodialysis using cellulose membranes, the use of glucan-containing gauzes or swabs, bacteremia, mucositis or graft-versus-host disease, and the use of selected antibiotics and antitumoral polysaccharides.
(1,3)-β-d-glucan is associated with Aspergillus, Candida, and Fusarium. In addition, the (1,3)-β-d-glucan test has been shown to be a sensitive and specific diagnostic marker for Pneumocystis jirovecii in patients with and without HIV. Cryptococci and the mucoraceous molds (eg, Zygomycetes such as Rhizopus, Mucor, and Absidia) have little to no measurable amounts of (1,3)-β-d-glucan, so this test will not be positive in patients with these fungal infections. The test has also performed poorly in patients with infections caused by Blastomyces spp. The major limitation in a positive test result is the lack of evidence on the specific fungal etiology.
Guidelines from the Infectious Diseases Society of America (IDSA), the European Organization for Research and Treatment of Cancer. and the Mycosis Study Group Education and Research Consortium support the use of (1,3)-β-d-glucan to aid in the diagnosis of invasive aspergillosis. Guidelines from the European Society of Clinical Microbiology and Infectious Diseases (ESCMID) have recommended its use for the diagnosis of candidemia, invasive candidiasis, and chronic disseminated candidiasis. The results of (1,3)-β-d-glucan testing have been shown to be more sensitive for detecting invasive aspergillosis versus invasive candidiasis. Most studies support the use of (1,3)-β-d-glucan in hematology patients because a limited number of studies have been conducted in patients without cancer and concerns regarding test specificity have occurred in solid-organ transplant recipients.
4. What laboratory tests are used in the diagnosis of HIV infection? What surrogate laboratory markers are used to assess the immunocompetence of patients infected with HIV? What laboratories are available to detect HIV resistance to available antiretrovirals?
ANSWER: Current recommendations for the diagnosis of HIV infection include the initial use of a fourth-generation antibody/antigen combination ELISA (or EIA) followed by a supplemental HIV-1/HIV-2 antibody differentiation immunoassay when the initial test is reactive (positive), and the use of NA tests when the antibody differentiation immunoassay is nonreactive or indeterminate (Figure 19-1). The CD4 cell count and the plasma HIV viral load are the two surrogate laboratory markers that are routinely used throughout the course of HIV infection. A CD4+ T-cell count is used to assess the immunocompetence of patients infected with HIV. An absolute CD4+ T-cell value of <200 cells/mm3 or <14% is associated with significant immunocompromise and risk for different opportunistic infections. The CD4+/CD8+ T-cell ratio is sometimes used to assess immune senescence or activation in the research context, but it is not clinically useful, generally, and therefore not routinely recommended. Plasma HIV viral load is a marker of antiretroviral treatment efficacy. HIV genotypic assays sequence or probe genes in specific areas of the virus in which different classes of antiretrovirals exert their action. This type of assay is recommended after HIV diagnosis, before starting antiretroviral therapy, and when HIV resistance is suspected based on unresponsiveness of the viral load despite good adherence to therapy. Genotypic assays most commonly are performed on RNA when HIV-RNA viral load is detectable, ideally ≥1,000 copies/mL, but can be obtained from proviral DNA when HIV-RNA is undetectable. HIV phenotypic assays measure the quantity of viral replication in the presence of various concentration of antiretroviral agents. Because they are labor-intensive and more expensive than genotypic assays, phenotypic assays are generally reserved for cases in which antiretroviral resistance cannot be predicted based on known genetic mutations alone.
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