Chapter 4: Principles of Laboratory Diagnosis of Infectious Diseases

The diagnosis of a microbial infection begins with an assessment of clinical and epidemiologic features, leading to the formulation of a diagnostic hypothesis. Anatomic localization of the infection with the aid of physical and radiologic findings (eg, right lower lobe pneumonia, subphrenic abscess) is usually included. This clinical diagnosis suggests a number of possible etiologic agents based on knowledge of infectious syndromes and their courses. The specific cause or etiologic diagnosis is then established by the application of methods described in this chapter. A combination of science and art on the part of both the clinician and laboratory worker is required: The clinician must select the appropriate tests and specimens to be processed and, where appropriate, suggest the suspected etiologic agents to the laboratory. The laboratory scientist must use the methods that will demonstrate the probable agents, and be prepared to explore other possibilities suggested by the clinical situation or by the findings of the laboratory examinations. The best results are obtained when communication between the clinician and laboratory is maximal.

The general approaches to laboratory diagnosis vary with different microorganisms and infectious diseases. However, the types of methods are usually some combination of direct microscopic examinations, culture, antigen detection, and antibody detection (serology). Nucleic acid amplification (NAA) assays that allow direct detection of genomic components of pathogens are now being used in many clinical microbiology laboratories. Particularly promising are newer multiplexed polymerase chain reaction (PCR) platforms that enable rapid detection of multiple potential pathogens in a single test reaction. Panels are available for respiratory, gastrointestinal, and central nervous system pathogens directly on appropriate specimens and identification of positive blood cultures. Despite such progress, however, traditional methods remain important and complementary, since isolation of microorganisms by culture is needed for most antimicrobial susceptibility testing. Not all pathogens are detected in these panels, and only known pathogens are sought. Therefore, this chapter considers the principles of infectious disease laboratory diagnosis and the methods available. Details about particular agents are discussed in the relevant chapters and in the section about infectious disease syndromes and etiologies at the back of the book. All diagnostic approaches begin with some kind of specimen collected from the patient.

The primary connection between the clinical encounter and the diagnostic laboratory is the specimen submitted for processing. If it is not appropriately chosen and/or collected, no degree of laboratory skill can rectify the error. Failure at the level of specimen collection is the most common reason for failing to establish an etiologic diagnosis, or worse, for suggesting a wrong diagnosis. In the case of bacterial infections, the primary problem lies in distinguishing resident or contaminating normal floral organisms from those causing the infection. The three specimen categories illustrated in Figure 4–1A-C are discussed in the text that follows.

Direct Tissue or Fluid Samples

Direct specimens (Figure 4–1A) are collected from normally sterile tissues (lung, liver) and body fluids (cerebrospinal fluid, blood). The methods range from needle aspiration of an abscess to surgical biopsy. In general, such collections require the direct involvement of a physician and may carry some risk for the patient. The results are always useful because positive findings are diagnostic and negative findings can exclude infection at the suspected site.

Indirect Samples

Indirect samples (Figure 4–1B) are specimens of inflammatory exudates (expectorated sputum, voided urine) that have passed through sites known to be colonized with the resident microbiota. The site of origin is usually sterile in healthy persons; however, some assessment of the probability of contamination with normal flora during collection is necessary in interpretation of the results. This assessment requires knowledge of the potential contaminating flora as well as the probable pathogens to be sought. Indirect samples are usually more convenient for both physician and patient, but carry a higher risk of misinterpretation. For some specimens, such as expectorated sputum, guidelines to assess specimen quality have been developed by correlation of clinical and microbiologic findings.

Samples from Microbiota Sites

Frequently, the primary site of infection is in an area known to be colonized with many organisms (pharynx and large intestine) (Figure 4–1C). This is primarily an issue with bacterial diagnosis because they dominate the makeup of the microbiota. In such instances, examinations are selectively made for organisms known to cause infection that are not normally found at the infected site. For example, the enteric pathogens Salmonella, Shigella, and Campylobacter may be selectively sought in a stool specimen or only β-hemolytic streptococci in a throat culture. In these instances, selective media that inhibit growth of the other bacteria are used or, if growing, they are simply ignored. Molecular assays that target the specific pathogens in these specimens are becoming more widely used in place of the selective cultures.

The selection of specimens for viral diagnosis is easier because there is usually little resident viral flora to confuse interpretation. This allows selection guided by knowledge of which sites are most likely to yield the suspected etiologic agent. For example, enteroviruses are the most common viruses involved in acute infection of the central nervous system.

Specimens that might be expected to yield these agents on culture or in molecular assays include nasopharyngeal or throat swabs, stool, and cerebrospinal fluid.

Specimen Collection and Transport

The sterile swab is the most convenient and most commonly used tool for specimen collection; however, it provides the poorest conditions for survival, can only absorb a small volume of inflammatory exudate, and is easily contaminated with adjacent microbiota. The worst possible specimen is a dried-out swab; the best is a collection of 5 to 10 mL or more of the infected fluid or tissue. The volume is important because infecting organisms that are present in small numbers may not be detected in a small sample.

Specimens should be transported to the laboratory as soon after collection as possible because some microorganisms survive only briefly outside the body. For example, unless special transport media are used, isolation rates of the organism that causes gonorrhea (Neisseria gonorrhoeae) are decreased when processing is delayed beyond a few minutes. Likewise, many respiratory viruses survive poorly outside the body. In contrast, some bacteria survive well and may even multiply after the specimen is collected. The growth of enteric gram-negative rods in specimens awaiting culture may, in fact, compromise specimen interpretation and interfere with the isolation of more fastidious organisms. Significant changes are associated with delays of more than 3 to 4 hours.

Various transport media have been developed to minimize the effects of the delay between specimen collection and laboratory processing. In general, they are buffered fluid or semisolid media containing minimal nutrients and are designed to prevent drying, maintain a neutral pH, and minimize growth of bacterial contaminants. Other features may be required to meet special requirements, such as an oxygen-free atmosphere for obligate anaerobes or specific (validated) collection-transport systems for molecular assays.

Of the infectious agents discussed in this book, only some of the parasites are large enough to be seen with the naked eye. Bacteria and fungi can be seen clearly with the light microscope when appropriate methods are used. Individual viruses can be seen only with the electron microscope, although aggregates of viral particles in cells (viral inclusions) may be seen by light microscopy. Various stains are used to visualize and differentiate microorganisms in smears and histologic sections.

Light Microscopy

Direct examination of stained or unstained preparations by light (bright-field) microscopy (Figure 4–2A) is particularly useful for detection of bacteria, fungi, and parasites. Even the smallest bacteria (1-2 μm wide) can be visualized, although all require staining and some require special lighting techniques. As the resolution limit of the light microscope is near 0.2 μm, the optics must be ideal if small organisms are to be seen clearly by direct microscopy.

Bacteria may be stained by a variety of dyes, including methylene blue, crystal violet, carbol-fuchsin (red), and safranin (red). The two most important methods, the Gram and acid-fast techniques, use staining, decolorization, and counterstaining in a manner that helps to classify as well as stain the organism.

The Gram Stain

The differential staining procedure described in 1884 by the Danish physician Hans Christian Gram has proved one of the most useful in microbiology and medicine. The procedure (Figure 4–3A) involves the application of a solution of iodine in potassium iodide to cells previously stained with an acridine dye such as crystal violet. This treatment produces a mordanting action in which purple insoluble complexes are formed with ribonuclear protein in the cell. The difference between gram-positive and gram-negative bacteria is in the permeability of the cell wall to these complexes on treatment with mixtures of acetone and alcohol solvents. This extracts the purple iodine-dye complexes from gram-negative cells, whereas gram-positive bacteria retain them. An intact cell wall is necessary for a positive reaction, and gram-positive bacteria may fail to retain the stain if the organisms are old, dead, or damaged by antimicrobial agents. Rarely, a gram-negative organism (eg, Acinetobacter) will appear gram positive. The stain is completed by the addition of red counter-stain such as safranin, which is taken up by bacteria that have been decolorized. Thus, cells stained purple are gram positive, and those stained red are gram negative. As indicated in Chapter 21, gram positivity and negativity correspond to major structural differences in the cell wall.

In many bacterial infections, the etiologic agents are readily seen on stained Gram smears of pus or fluids. The purple or red bacteria are seen against a gram-negative (red) background of leukocytes, exudate, and debris (Figures 4–3A and 4–4C). This information, combined with the clinical findings, may guide the management of infection before culture results are available. Interpretation requires considerable experience and knowledge of probable causes, of their morphology and Gram reaction, and of any organisms normally present in health at the infected site.

The Acid-fast Stain

Acid fastness is a property of the mycobacteria (eg, Mycobacterium tuberculosis) and related organisms. Acid-fast organisms generally stain very poorly with dyes, including those used in the Gram stain. However, they can be stained by prolonged application of more concentrated dyes, by penetrating agents, or by heat treatment. Their unique feature is that when stained, acid-fast bacteria resist decolorization by concentrations of mineral acids and ethanol that remove the same dyes from other bacteria. This combination of weak initial staining and strong retention once stained is related to the high lipid content of the mycobacterial cell wall. Acid-fast stains are completed with a counterstain to provide a contrasting background for viewing the stained bacteria (Figure 4–3B).

In the acid-fast procedure, the slide is flooded with carbol-fuchsin (red) and decolorized with hydrochloric acid in alcohol. When counterstained with methylene blue, acid-fast organisms appear red against a blue background (Figure 4–4D). A variant is the fluorochrome stain, which uses a fluorescent dye (auramine, or an auramine–rhodamine mixture), followed by decolorization with acid–alcohol. Acid-fast organisms retain the fluorescent stain, which allows their visualization by fluorescence microscopy. The fluorochrome stain is more sensitive and allows rapid screening and, therefore, has become the method of choice in most laboratories performing testing for acid-fast organisms.

Fungal and Parasitic Stains

The smallest fungi are the size of large bacteria, and all parasitic forms are larger. This allows detection in simple wet mount preparations, often without staining. Fungi in sputum or body fluids can be seen by mixing the specimen with a potassium hydroxide solution (to dissolve debris) and viewing with a medium power lens. The use of simple stains or the fluorescent calcofluor white improves the sensitivity of detection. Another technique is to mix the specimen with India ink, which outlines the fungal cells (Figure 4–4F). Detection of the cysts and eggs of parasites requires a concentration procedure if the specimen is stool, but once done they can be visualized with a simple iodine stain (Figure 4–5).

Dark-field and Fluorescence Microscopy

Some bacteria, such as Treponema pallidum, the cause of syphilis, are too thin to be visualized with the usual bright-field illumination. They can be seen by use of the dark-field technique. With this method, a condenser focuses light diagonally on the specimen in such a way that only light reflected from particulate matter, such as bacteria, reaches the eyepiece (Figure 4–2B). The angles of incident and reflected light are such that the organisms are surrounded by a bright halo against a black background. This type of illumination is also used in other microscopic techniques, in which a high-light contrast is desired, and for observation of fluorescence. Fluorescent compounds, when excited by incident light of one wavelength, emit light of a longer wavelength and thus a different color. When the fluorescent compound is conjugated with an antibody as a probe for detection of a specific antigen, the technique is called immunofluorescence, or fluorescent antibody microscopy (Figure 4–6). The appearance is the same as in dark-field microscopy except that the halo is the emitted color of the fluorescent compound (Figures 4–2C and 4–6C and D). Immunofluorescence stains are the most commonly used stains for detection of viruses though these are being replaced by the more sensitive molecular assays. For improved safety, most modern fluorescence microscopy systems direct the incident light through the objective from above (epifluorescence).

Electron Microscopy

Electron microscopy shows structures by transmission of an electron beam and has 10 to 1000 times the resolving power of light microscopic methods. For practical reasons, its diagnostic application is limited to virology, where, because of the resolution possible at high magnification, it offers results not possible by any other method. Using negative staining techniques for direct examination of fluids and tissues from affected body sites enables visualization of viral particles. In some instances, electron microscopy has been the primary means of discovery of viruses that do not grow in the usual cell culture systems. Molecular assays are replacing electron microscopy for detection of many viruses.

Growth and identification of the infecting agent in vitro is usually the most sensitive and specific means of diagnosis and is, thus, the method most commonly used. Theoretically, the presence of a single live organism in the specimen can yield a positive result. Most bacteria and fungi can be grown in a variety of artificial media, but strictly intracellular microorganisms (eg, Chlamydia, Rickettsia, and viruses) can be isolated only in cultures of living eukaryotic cells. The culture of some parasites is possible, but used only in highly specialized laboratories.

Isolation and Identification of Bacteria and Fungi

Almost all medically important bacteria can be cultivated outside the host in artificial culture media. A single bacterium placed in the proper culture conditions multiplies to quantities sufficient to be seen by the naked eye. Bacteriologic media are soup-like recipes prepared from digests of animal or vegetable protein supplemented with nutrients such as glucose, yeast extract, serum, or blood to meet the metabolic requirements of the organism. Their chemical composition is complex, and their success depends on matching the nutritional requirements of most heterotrophic living things. The same approaches as well as some of the same culture media used for bacteria are also used for fungi.

Growth in media prepared in the fluid state (broths) is apparent when bacterial numbers are sufficient to produce turbidity or macroscopic clumps. Turbidity results from reflection of transmitted light by the bacteria; depending on the size of the organism, more than 10⁶ bacteria per milliliter of broth are required. The addition of a gelling agent to a broth medium allows its preparation in solid form as plates in Petri dishes. The universal gelling agent for diagnostic bacteriology is agar—a polysaccharide extracted from seaweed. Agar has the convenient property of becoming liquid at approximately 95°C but not returning to the solid gel state until cooled to less than 50°C. This allows the addition of a heat-labile substance such as blood to the medium before it sets. At temperatures used in the diagnostic laboratory (37°C or lower), broth–agar exists as a smooth, solid, nutrient gel. This medium, usually termed “agar,” may be qualified with a description of any supplement (eg, blood agar).

A useful feature of agar plates is that the bacteria can be separated by spreading a small sample of the specimen over the surface. Bacterial cells that are well separated from others grow as isolated colonies, often reaching 2 to 3 mm in diameter after overnight incubation. This allows isolation of bacteria in pure culture because the colony is assumed to arise from a single organism (Figure 4–7). Colonies vary greatly in size, shape, texture, color, and other features called colonial morphology. Colonies from different species or genera often differ substantially, whereas those derived from the same strain are usually consistent. Differences in colonial morphology are very useful for separating bacteria in mixtures and as clues to their identity. Some examples of colonial morphology are shown in Figure 4–8.

New methods that do not depend on visual changes in the growth medium or colony formation are also used to detect bacterial growth in culture. These techniques include optical, chemical, and electrical changes in the medium, produced by the growing numbers of bacterial cells or their metabolic products. Many of these methods are more sensitive than classic techniques and, thus, can detect growth hours, or even days, earlier than traditional methods. Some have also been engineered for instrumentation and automation. For example, one fully automated system that detects bacterial metabolism fluorometrically can complete a bacterial identification and antimicrobial susceptibility test in 2 to 4 hours.

Culture Media

Over the last 100 years, countless media have been developed by microbiologists to aid in the isolation and identification of medically important bacteria and fungi. Only a few have found their way into routine use in clinical laboratories. These may be classified as nutrient, selective, or indicator media.

Nutrient Media

The nutrient component of a medium is designed to satisfy the growth requirements of the organism to permit isolation and propagation. For medical purposes, the ideal medium would allow rapid growth of all agents. No such medium exists; however, several suffice for good growth of most medically important bacteria and fungi. These media are prepared with enzymatic or acid digests of animal or plant products such as muscle, milk, or soybeans. The digest reduces the native protein to a mixture of polypeptides and amino acids that also includes trace metals, coenzymes, and various undefined growth factors. For example, one common broth contains a digest of casein (milk curd) and a digest of soybean meal. To this nutrient base, salts, vitamins, or body fluids such as serum may be added to provide pathogens with the conditions needed for optimum growth.

Selective Media

Selective media are used when specific pathogenic organisms are sought in sites with an extensive microbiota (eg, Campylobacter species in fecal specimens). In these cases, other bacteria may overgrow the suspected etiologic species in simple nutrient media, either because the pathogen grows more slowly or because it is present in much smaller numbers. Selective media usually contain dyes, other chemical additives, or antimicrobial agents at concentrations designed to inhibit contaminating flora but not the suspected pathogen.

Indicator Media

Indicator media contain substances designed to demonstrate biochemical or other features characteristic of specific pathogens or organism groups. The addition to the medium of one or more carbohydrates and a pH indicator is frequently used. A color change in a colony indicates the presence of acid products and thus of fermentation or oxidation of the carbohydrate by the organism. The addition of red blood cells (RBCs) to plates allows the hemolysis produced by some organisms to be used as a differential feature. In practice, nutrient, selective, and indicator properties are often combined to various degrees in the same medium. It is possible to include an indicator system in a highly nutrient medium and also make it selective by adding appropriate antimicrobials. Some examples of culture media commonly used in diagnostic microbiology are listed in Appendix 4–1, and more details of their constitution and application are provided in Appendix 4–2.

Atmospheric Conditions

Aerobic

After inoculation, cultures of most aerobic bacteria are placed in an incubator with temperature maintained at 35°C to 37°C. Slightly higher or lower temperatures are used occasionally to selectively favor a certain organism or organism group. Most bacteria that are not obligate anaerobes grow in air; however, CO₂ is required by some and enhances the growth of others. Incubators that maintain a 2% to 5% concentration of CO₂ in air are frequently used for primary isolation, because this level is not harmful to any bacteria and improves isolation of some. A simpler method is the candle jar, in which a lighted candle is allowed to burn to extinction in a sealed jar containing plates. This method adds 1% to 2% CO₂ to the atmosphere. Some bacteria (eg, Campylobacter) require a microaerophilic atmosphere with reduced oxygen (5%) and increased CO₂ (10%) levels to grow. This can be achieved by using a commercially available packet that is placed in a jar which is then sealed similar to the anaerobic system described further.

Anaerobic

Strictly anaerobic bacteria do not grow under the conditions just described, and many die when exposed to atmospheric oxygen or high oxidation–reduction potentials. Most medically important anaerobes grow in the depths of liquid or semisolid media containing any of a variety of reducing agents, such as cysteine, thioglycollate, ascorbic acid, or even iron filings. An anaerobic environment for incubation of plates can be achieved by replacing air with a gas mixture containing hydrogen, CO₂, and nitrogen and allowing the hydrogen to react with residual oxygen on a catalyst to form water. A convenient commercial system accomplishes this chemically in a packet that is added before the jar is sealed. Specimens suspected to contain significant anaerobes should be processed under conditions designed to minimize exposure to atmospheric oxygen at all stages.

Clinical Microbiology Procedures

Routine laboratory procedures for processing specimens from various sites are needed because no single medium or atmosphere is ideal for all bacteria. Combinations of broth and solid-plated media and aerobic, CO₂, and anaerobic incubation must be matched to the organisms expected at any particular site or clinical circumstance. Examples of such routines are shown in Table 4–1. In general, it is not practical to routinely include specialized media for isolation of rare organisms such as Corynebacterium diphtheriae or Legionella pneumophila. For detection of these and other uncommon organisms, the laboratory must be specifically informed of their possible presence by the physician. Appropriate media and special procedures can then be included.

Identification

When growth is detected in any medium, the process of identification begins. Identification involves methods for obtaining pure cultures from single colonies, followed by tests designed to characterize and identify the isolate. The exact tests and their sequences vary with different groups of organisms, and the taxonomic level (genus, species, subspecies, etc) of identification needed varies according to the medical usefulness of the information. In some cases, only a general description or the exclusion of particular organisms is important. For example, a report of “mixed oral flora” in a sputum specimen or “No Salmonella, Shigella, or Campylobacter isolated” in a fecal specimen may provide all the information needed. MALDI-TOF (matrix-assisted laser desorption ionization-time of flight) mass spectroscopy has become major new tool used for the rapid identification of microorganisms already isolated in pure culture; it complements but does not replace fully the need for traditional methods.

Features Used to Classify Bacteria and Fungi

Cultural Characteristics

Cultural characteristics include the demonstration of properties such as unique nutritional requirements, pigment production, and the ability to grow in the presence of certain substances (sodium chloride, bile) or on certain media (MacConkey, nutrient agar). Demonstration of the ability to grow at a particular temperature or to cause hemolysis on blood agar plates is also used. For fungi, growth as a yeast colony or a mold is the primary separator. For molds, the morphology of the mold structures (hyphae, conidia, etc) are the primary means of identification.

Biochemical Characteristics

The ability to attack various substrates or to produce particular metabolic products has broad application to the identification of bacteria and yeast. The most common properties examined are listed in Appendix 4–3. Biochemical and cultural tests for bacterial identification are analyzed by reference to tables that show the reaction patterns characteristic of individual species. In fact, advances in computer analysis have now been applied to identification of many bacterial and fungal groups. These systems use the same biochemical principles together with computerized databases to determine the most probable identification from the observed test pattern.

Toxin Production and Pathogenicity

Direct evidence of virulence in laboratory animals is rarely needed to confirm a clinical diagnosis. In some diseases caused by production of a specific toxin, the toxin may be detected in vitro through cell cultures or immunologic methods. Neutralization of the toxic effect with specific antitoxin is the usual approach to identify the toxin. Toxin testing is conducted only in specialized laboratories. Molecular assays have been developed for some toxins.

Antigenic Structure

Viruses, bacteria, fungi, and parasites possess many antigens, such as capsular polysaccharides, surface proteins, and cell wall components. Serology involves the use of antibodies of known specificity to detect antigens present on whole organisms or free in extracts (soluble antigens). The methods used for demonstrating antigen-antibody reactions are discussed in “Antibody Detection (Serology).”

Genomic Structure

Nucleic acid–sequence relatedness as determined by homology and direct sequence comparisons have become a primary determinant of taxonomic decisions. They are discussed later in the section on nucleic acid methods.

Isolation and Identification of Viruses

Cell and Organ Culture

Living cell cultures that can support their replication are the primary means of isolating pathogenic viruses. The cells are derived from a tissue source by outgrowth of cells from a tissue fragment (explant) or by dispersal with proteolytic agents such as trypsin. They are allowed to grow in nutrient media on a glass or plastic surface until a confluent layer one cell thick (monolayer) is achieved. In some circumstances, a tissue fragment with a specialized function (eg, fetal trachea with ciliated epithelial cells) is cultivated in vitro and used for viral detection. This procedure is known as organ culture.

Three basic types of cell culture monolayers are used in diagnostic virology. The primary cell culture, in which all cells have a normal chromosome count (diploid), is derived from the initial growth of cells from a tissue source. Redispersal and regrowth produce a secondary cell culture, which usually retains characteristics similar to those of the primary culture (diploid chromosome count and virus susceptibility). Monkey and human embryonic kidney cell cultures are examples of commonly used primary and secondary cell cultures.

Further dispersal and regrowth of secondary cell cultures usually lead to one of two outcomes: the cells eventually die, or they undergo spontaneous transformation, in which the growth characteristics change, the chromosome count varies (haploid or heteroploid), and the susceptibility to virus infection differs from that of the original. These cell cultures have characteristics of “immortality”; that is, they can be redispersed and regrown many times (serial cell culture passage). They can also be derived from cancerous tissue cells or produced by exposure to mutagenic agents in vitro. Such cultures are commonly called cell lines. A common cell line in diagnostic use is the Hep-2, derived from a human epithelial carcinoma. A third type of culture is often termed a cell strain. This culture consists of diploid cells, commonly fibroblastic, that can be redispersed and regrown a finite number of times; usually, 30 to 40 cell culture passages can be made before the strain dies out or spontaneously transforms. Human embryonic tonsil and lung fibroblasts are common cell strains used in clinical virology laboratories that are continuing to perform cultures. Molecular assays that are faster, more sensitive, and more cost-effective are replacing the standard viral culture techniques in many laboratories.

Shell vial techniques using coverslips with monolayers of cell lines have been developed for some viruses (eg, cytomegalovirus, respiratory viruses) to provide a more rapid culture method. Virus is amplified in the cell culture vials after low-speed centrifugation. Then fluorescent staining techniques using monoclonal antibodies for the specific virus are used to detect early viral antigens prior to the development of cytopathic effect (CPE).

Cells from Cancerous Tissue May Grow Continuously

Detection of Viral Growth

Viral growth in susceptible cell cultures can be detected in several ways. The most common effect is seen with lytic or cytopathic viruses; as they replicate in cells, they produce alterations in cellular morphology (or cell death), which can be observed directly by light microscopy under low magnification (×30 or ×100). This CPE varies with different viruses in different cell cultures. For example, enteroviruses often produce cell rounding, pleomorphism, and eventual cell death in various culture systems, whereas measles and respiratory syncytial viruses cause fusion of cells to produce multinucleated giant cells (syncytia). The microscopic appearance of some normal cell cultures and the CPE produced in them by different viruses are illustrated in Figure 4–9.

Other viruses may be detected in cell culture by their ability to produce hemagglutinins. These hemagglutinins may be present on the infected cell membranes, as well as in the culture media, as a result of release of free, hemagglutinating virions from the cells. Addition of erythrocytes to the infected cell culture results in their adherence to the cell surfaces—a phenomenon known as hemadsorption. Another method of viral detection in cell culture is by interference. In this situation, the virus that infects the susceptible cell culture produces no CPE or hemagglutinin, but can be detected by “challenging” the cell culture with a different virus that normally produces a characteristic CPE. The second, or challenge, virus fails to infect the cell culture because of interference by the first virus, which is thus detected. This method is obviously cumbersome, but has been applied to the detection of rubella virus in certain cell cultures.

For some agents, such as Epstein-Barr virus (EBV) or human immunodeficiency virus (HIV), even more novel approaches may be applied. Both EBV and HIV can replicate in vitro in suspension cultures of normal human lymphocytes such as those derived from neonatal cord blood. Their presence may be determined in several ways; for example, EBV-infected B lymphocytes and HIV-infected T lymphocytes express virus-specified antigens and viral DNA or RNA, which can be detected with immunologic or genomic probes. In addition, HIV reverse transcriptase can be detected in cell culture by specific assay methods. In addition, immunologic and nucleic acid probes (see further text) can also be used to detect virus in clinical specimens or in situations in which only incomplete, noninfective virus replication has occurred in vivo or in vitro. An example is the use of in-situ cytohybridization, whereby specific labeled nucleic acid probes are used to detect and localize papillomavirus genomes in tissues where neither infectious virus nor its antigens can be detected.

In-Vivo Isolation Methods

In-vivo methods for isolation, although carried out only in highly specialized laboratories, are also sometimes necessary. The embryonated hen’s egg is still often used for the initial isolation and propagation of influenza A virus. Virus-containing material is inoculated on the appropriate egg membrane, and the egg is incubated to permit viral replication and recognition. Animal inoculation is now only occasionally used by highly specialized laboratories for detecting some viruses. The usual animal host for viral isolation is the mouse; suckling mice in the first 48 hours of life are especially susceptible to many viruses. Evidence of viral replication is based on the development of illness, manifested by such signs as paralysis, convulsions, poor feeding, or death. The nature of the infecting virus can be further elucidated by histologic and immunofluorescent examination of tissues or by detection of specific antibody responses. Many arboviruses and rabies virus can be detected in this system.

Viral isolation from a suspect case involves a number of steps. First, the viruses that are believed to be most likely involved in the illness are considered, and appropriate specimens are collected. Centrifugation or filtration and addition of antimicrobials are frequently required with respiratory or fecal specimens to remove organic matter, cellular debris, bacteria, and fungi, which can interfere with viral isolation. The specimens are then inoculated into the appropriate cell culture systems. The time between inoculation and initial detection of viral effects varies; however, for most viruses positive cultures are usually apparent within 5 days of collection. With proper collection methods and application of the diagnostic tools, as discussed further, many infections can even be detected within hours. In contrast, some viruses may require culture for a month or more before they can be detected.

Viral Identification

On isolation, a virus can usually be tentatively identified to the family or genus level by its cultural characteristics (eg, the type of CPE produced). Confirmation and further identification may require enhancement of viral growth to produce adequate quantities for testing. This result may be achieved by inoculation of the original isolate into fresh culture systems (viral passage) to amplify replication of the virus, as well as improve its adaptation to growth in the in vitro system.

Neutralization and Serologic Detection

Of the several ways of identifying the isolate, the most common is to neutralize its infectivity by mixing it with specific antibody to known viruses before inoculation into cultures. The inhibition of the expected viral effects on the cell culture such as CPE or hemagglutination is then evidence of that virus. As in bacteriology, demonstration of specific viral antigens is a useful way to identify many agents. Immunofluorescence and enzyme immunoassay (EIA) are the most common methods.

Cytology and Histology

In some instances, viruses produce specific cytologic changes in infected host tissues that aid in diagnosis. Examples include specific intranuclear inclusions seen in neuronal infections due to herpes simplex (Cowdry type A bodies) and due to intracytoplasmic inclusions in rabies (Negri bodies), and cell fusion, which results in multinucleated epithelial giant cells (eg, measles and varicella zoster). Although such findings are useful when seen, their overall diagnostic sensitivity and specificity are usually considerably less than those of the other methods discussed.

Electron Microscopy

When virions are present in sufficient numbers, they may be further characterized by specific agglutination of viral particles on mixture with type-specific antiserum. This technique, immune electron microscopy, can be used to identify viral antigens specifically or to detect antibody in serum using viral particles of known antigenicity.

Some viruses (eg, human rotaviruses and hepatitis A and B viruses) grow poorly or not at all in the laboratory culture systems currently available. However, they can be efficiently detected by immunologic or molecular methods (described later in this chapter).

Diagnostic microbiology makes great use of the specificity of the binding between antigen and antibody. Antisera of known specificity are used to detect their homologous antigen in cultures, or more recently, directly in body fluids. Conversely, known antigen preparations are used to detect circulating antibodies as evidence of a current or previous infection with that agent. Many methods are in use to demonstrate the antigen–antibody binding. The greatly improved specificity of monoclonal antibodies has had a major impact on the quality of methods where they have been applied. Before discussing their application to diagnosis, the principles involved in the most important methods are discussed.

Methods for Detecting an Antigen–Antibody Reaction

Precipitation

When antigen and antibody combine in the proper proportions, a visible precipitate is formed. Optimum antigen–antibody ratios can be produced by allowing one to diffuse into the other, most commonly through an agar matrix (immunodiffusion). In the immunodiffusion procedure, wells are cut in the agar and filled with antigen and antibody. One or more precipitin lines may be formed between the antigen and antibody wells, depending on the number of different antigen–antibody reactions occurring. Counterimmunoelectrophoresis (CIE) is immunodiffusion carried out in an electrophoretic field. The net effect is that antigen and antibody are rapidly brought together in the space between the wells to form a precipitin line.

The amount of antigen or antibody necessary to produce a visible immunologic reaction can be reduced if either is on the surface of a relatively large particle. This condition can be produced by fixing soluble antigens or antibody onto the surface of RBCs or microscopic latex particles suspended in a test tube or microtiter plate well (Figure 4–10). Whole bacteria are large enough to serve as the particle if the antigen is present on the microbial surface. The relative proportions of antigen and antibody thus become less critical, and antigen–antibody reactions are detectable by agglutination when immune serum and particulate antigen, or particle-associated antibody and soluble antigen, are mixed on a slide. The process is termed slide agglutination, hemagglutination, or latex agglutination depending on the nature of the sensitized particle.

Neutralization

Neutralization takes some observable function of the agent, such as CPE of viruses or the action of a bacterial toxin, and neutralizes it. This is usually done by first reacting the agent with antibody and then placing the antigen–antibody mixture into the test system. The steps involved are illustrated in Figure 4–11. In viral neutralization, a single antibody molecule can bind to surface components of the extracellular virus and interfere with one of the initial events of the viral multiplication cycle (adsorption, penetration, or uncoating). Some bacterial and viral agents directly bind to RBCs (hemagglutination). Neutralization of this reaction by antibody blocking of the receptor is called hemagglutination inhibition (Figure 4–12).

Complement Fixation

Complement fixation assays depend on two properties of complement. The first is fixation (inactivation) of complement on formation of antigen–antibody complexes. The second is the ability of bound complement to cause hemolysis of sheep [RBCs coated with anti-sheep RBC antibody (sensitized RBCs)]. Complement fixation assays are conducted in two stages: The test system reacts the antigen and antibody in the presence of complement; the indicator system, which contains the sensitized RBCs, detects residual complement. Hemolysis indicates that complement was present in the indicator system and, therefore, that antigen–antibody complexes were not formed in the test system. Primarily used to detect and quantitate antibody, complement fixation has been largely replaced by simpler methods that can be readily automated.

Labeling Methods

Detection of antigen–antibody binding may be enhanced by attaching a label to one (usually the antibody) and detecting the label after removal of unbound reagents. The label may be a fluorescent dye (immunofluorescence), a radioisotope (radioimmunoassay, or RIA), or an enzyme (enzyme immunoassay, or EIA). The presence or quantitation of antigen– antibody binding is measured by fluorescence, radioactivity, or the chemical reaction catalyzed by the enzyme.

Immunofluorescence

The most common labeling method in diagnostic microbiology is immunofluorescence (Figure 4–6), in which antibody labeled with a fluorescent dye, usually fluorescein isothiocyanate (FITC), is applied to a slide of material that may contain the antigen sought. Under fluorescence microscopy, binding of the labeled antibody can be detected as a bright green halo surrounding bacterium or, in the case of viruses, as a fluorescent clump in or on an infected cell. The method is called “direct” if the FITC is conjugated directly to the antibody with the desired specificity. In “indirect” immunofluorescence, the specific antibody is not labeled, but its binding to an antigen is detected in an additional step using an FITC-labeled anti-immunoglobulin antibody that binds to the specific antibody. Choice between the two approaches involves purely technical considerations.

RIA and EIA

The labels used in RIA and EIA are more suitable for liquid phase assays and are used particularly in virology. They are also used in direct and indirect methods and many other ingenious variations such as the “sandwich” methods, so called because the antigen of interest is “trapped” between two antibodies (Figure 4–13). These extremely sensitive techniques are discussed further with regard to antibody detection.

Serologic Classification

For most important antigens of diagnostic significance, antisera are commercially available. The most common test methods for bacteria are agglutination and immunofluorescence, and, for viruses, neutralization. In most cases, these methods subclassify organisms below the species level and, thus, are primarily of value for epidemiologic and research purposes. The terms “serotype” and “serogroup” are used together with numbers, letters, or Roman numerals with no apparent logic other than historical precedent. For a few genera, the most fundamental taxonomic differentiation is serologic. This is the case with the streptococci, in which an existing classification based on biochemical and cultural characteristics was superseded because a serologic classification scheme developed by Rebecca Lancefield correlated better with disease.

Before these techniques can be applied to the diagnosis of specific infectious diseases, considerable study of the causative agent(s) is required. Antigen–antibody systems may vary in complexity from a single epitope to scores of epitopes on several macromolecular antigens, whose chemical nature may or may not be known. The cause of the original 1976 outbreak of Legionnaires disease (caused by L pneumophila) was proved through the development of immune reagents that detected the bacteria in tissue and antibodies directed against the bacteria in the serum of patients. Now, more than 40 years later, there are more than a dozen serotypes and many additional species, each requiring specific immunologic reagents for antigen or antibody detection for diagnosis.

Antibody Detection (Serology)

During infection—viral, bacterial, fungal, or parasitic—the host usually responds with the formation of antibodies, which can be detected by modification of any of the methods used for antigen detection. The formation of antibodies and their time course depend on the antigenic stimulation provided by the infection. The precise patterns vary depending on the antigens used, the classes of antibody detected, and the method. An example of temporal patterns of development and increase and decline in specific antiviral antibodies measured by different tests is illustrated in Figure 4–14. These responses can be used to detect evidence of recent or past infection. The test methods do not inherently indicate immunoglobulin class, but can be modified to do so, usually by pretreatment of the serum to remove IgG to differentiate the IgM and IgG responses. Several basic principles must be emphasized:

1. In an acute infection, the antibodies usually appear early in the illness, and then rise sharply over the next 10 to 21 days. Thus, a serum sample collected shortly after the onset of illness (acute serum), and another collected 2 to 3 weeks later (convalescent serum) can be compared quantitatively for changes in specific antibody content.

2. Antibodies can be quantitated by several means. The most common method is to dilute the serum serially in appropriate media and determine the maximal dilution that will still yield detectable antibody in the test system (eg, serum dilutions of 1:4, 1:8, and 1:16). The highest dilution that retains specific activity is called the antibody titer.

3. The interpretation of significant antibody responses (evidence of specific, recent infection) is most reliable when definite evidence of seroconversion is demonstrated; that is, detectable specific antibody is absent from the acute serum but present in the convalescent serum. Alternatively, a fourfold or greater increase in antibody titer supports a diagnosis of recent infection; for example, an acute serum titer of 1:4 or less and a convalescent serum titer of 1:16 or greater would be considered significant.

4. In instances in which the average antibody titers of a population to a specific agent are known, a single convalescent antibody titer significantly greater than the expected mean may be used as a supportive or presumptive evidence of recent infection. However, this finding is considerably less valuable than those obtained by comparing responses of acute and convalescent serum samples. An alternative and somewhat more complex method of serodiagnosis is to determine which major immunoglobulin subclass constitutes the major proportion of the specific antibodies. In primary infections, the IgM-specific response is often dominant during the first days or weeks after onset, but is replaced progressively by IgG-specific antibodies; thus, by 1 to 6 months after infection, the predominant antibodies belong to the IgG subclass. Consequently, serum containing a high titer of antibodies of the IgM subclass would suggest a recent, primary infection.

The immunologic methods used to identify bacterial or viral antigens are applied to serologic diagnosis by simply reversing the detection system: that is, using a known antigen to detect the presence of an antibody. The methods of serologic diagnosis to be used are selected on the basis of their convenience and applicability to the antigen in question. As shown in Figure 4–14, the temporal relationships of antibody response to infection vary according to the method used. Of the methods for measuring antigen–antibody interaction discussed previously, those now used most frequently for serologic diagnosis are agglutination, RIA, and EIA.

Western Blot

The Western blot immunoassay is another technique that is now commonly used to detect and confirm the specificity of antibodies to a variety of epitopes. Its greatest use has been in the diagnosis of HIV infections (see Chapter 18), in which virions are electrophoresed in a polyacrylamide gel to separate the protein and glycoprotein components and then transferred onto nitrocellulose. This is then incubated with patient serum, and antibody to the different viral components is detected by using an antihuman globulin IgG antibody conjugated with an enzyme label. Newer EIA assays that detect the P24 antigen as well as antibodies to HIV obviate the need for the Western Blot assay in those laboratories using these assays (Figure 4–15).

Antigen Detection

Theoretically, any of the methods described for detecting antigen–antibody interactions can be applied directly to clinical specimens. The most common of these is immunofluorescence, in which antigen is detected on the surface of the organism or in cells present in the infected secretion. The greatest success with this approach has been in respiratory infections in which a nasopharyngeal, throat washing, sputum, or bronchoalveolar lavage specimen may contain bacteria or viral aggregates in sufficient amount to be seen microscopically. Although the fluorescent tag makes it easier to find organisms, these methods are generally not as sensitive as culture. With some genera and species, the immunofluorescent detection of antigens in clinical material provides the most rapid means of diagnosis, as with Legionella and respiratory syncytial virus.

Another approach to detecting antigens is to detect free antigen released by the organism into body fluids. This offers the possibility of bypassing direct examination, culture, and identification tests to achieve a diagnosis. Success requires a highly specific antibody, a sensitive detection method, and the presence of the homologous antigen in an accessible body fluid. The latter is an important limitation, because not all organisms release free antigen in the course of infection. At present, diagnosis by antigen detection is limited to some bacteria and fungi with polysaccharide capsules (eg, Haemophilus influenzae), to Chlamydia, and to certain viruses. The techniques of agglutination with antibody bound to latex particles, CIE, RIA, and EIA are used to detect free antigen in serum, urine, cerebrospinal fluid, and joint fluid. Live organisms are not required for antigen detection, and these tests may still be positive when the causative organism has been eliminated by antimicrobial therapy. The procedures can yield results within 1 or 2 hours, sometimes within a few minutes. This feature is attractive for office practice because it allows diagnostic decisions to be made during the patient’s visit. A number of commercial products detect group A streptococci in sore throats with over 90% sensitivity; however, because these tests are less sensitive than culture, negative results must still be confirmed by culture.

As with the human genome, the genome sequence of the major human pathogens has or soon will be determined. These data are placed in widely available computer databases and have already been used for applications ranging from taxonomy to detection of antimicrobial resistance genes. Some of the methods and applications relevant to the study of infectious diseases are briefly summarized in the following discussion. The student is referred to textbooks of molecular biology for more complete coverage.

Methods of Nucleic Acid Analysis

DNA Hybridization and Probes

If the DNA double helix is opened, leaving single-stranded (denatured) DNA, the nucleotide bases are exposed and, thus, are available to interact with other single-stranded nucleic acid molecules. If complementary sequences of a second DNA molecule are brought into physical contact with the first, they hybridize to it, forming a new double-stranded molecule in that area. A probe is a cloned DNA fragment that has been labeled so that it can be detected if it hybridizes to complementary sequences in such a test system (Figure 4–16). The probe may be derived from the gene for a known protein of the pathogen or be empirically derived just for diagnostic purposes. The methods that allow the hybridization to take place include those that immobilize the single-stranded target DNA on a membrane, as in the Figure, or liquid-phase assays, which can be rapid and automated. A variant in which the DNA is separated by agarose gel electrophoresis before binding to the membrane is called Southern hybridization.

Agarose Gel Electrophoresis

Nucleic acids may be separated in an electrophoretic field in an agarose (highly purified agar) gel. The speed of migration depends on size, with the smaller molecules moving faster and appearing at the bottom (end) of the gel. This method is able to separate DNA fragments in the range of 0.1 to 50 kilobases, which is far below the size of bacterial genomes but includes some naturally occurring genetic elements such as bacterial plasmids. This analysis can be refined by the use of restriction endonucleases, which are enzymes derived from bacteria that recognize specific nucleotide sequences in DNA molecules and digest (cut) them at all sites where the sequence appears. Thus, plasmids of the same size may be differentiated by the size of fragments generated by endonuclease digestion of DNA as shown in Figure 4–17A-C. Agarose gel electrophoresis, endonuclease digestion, and the specificity of a probe may all be combined in searches for the source specific genes as shown in Figure 4–18A-E.

Nucleic Acid Amplification

Nucleic acid amplification (NAA) methods such as the PCR allow the detection and selective replication of a targeted portion of the genome. The basic PCR technique uses synthetic oligonucleotide primers and special DNA polymerases in a way which allows repeated cycles of synthesis of only a segment of a targeted DNA molecule that may be as large as an entire genome. The specificity is provided by the sequence of approximately 20 nucleotides in each primer pair, which are crafted to flank the desired segment of the genome. The DNA polymerases used are ones that operate at unusually high temperatures. This allows the use of temperature to control shifts between separation of the complementary DNA strands (so primers can bind) and replication of the DNA sequence that lies between the two primers. Because each strand generates a new fragment, the increase is exponential. In a machine called a thermocycler, the targeted DNA can be amplified 1 million to 1 billion times in 20 to 30 cycles (Figure 4–19A-D). Other NAA methods use the same principles.

Application of Nucleic Acid Methods to Infectious Diseases

Bacterial and Viral Genomes

The only intact genetic elements of infectious agents that are small enough to be directly detected and sized by agarose gel electrophoresis are bacterial plasmids. Not all bacterial species typically harbor plasmids, but those that do may carry one or a number of plasmids ranging in size from less than 1 to more than 50 kilobases. This diversity makes the presence or absence, number, and sizes of plasmids of considerable value in differentiating strains for epidemiologic purposes. Because plasmids are not stable components of the bacterial genome, plasmid analysis also has the element of a timely “snapshot” of the circumstances of a disease outbreak. The specificity of these results can be improved by digesting the plasmids with restriction endonucleases before electrophoresis. Two plasmids of the same size from different strains may not be the same, but if an identical pattern of fragments is generated from the digestion, they almost certainly are. These principles are illustrated in Figures 4–16 and 4–17.

Because of their larger size, the chromosomes of bacteria must be digested with endonucleases to resolve them on gels. For viruses, the outcome is much like that with plasmids, depending on the genomic size and the endonuclease used. Digested bacterial chromosomes can be compared in this manner, but the number of fragments is very large and the patterns complex. The combined use of endonucleases, which make infrequent cuts, and electrophoretic methods able to resolve large fragments can produce a comparison comparable to that possible with plasmids. This approach is also used for analysis of the multiple chromosomes of fungi and parasites.

DNA Probes

Probes may be recovered from NAA procedures or more commonly synthesized as a single chain of nucleotides (oligonucleotide probe) from known sequence data. They may contain a gene of known function or simply sequences empirically found to be useful for the application in question. When labeled with a radioisotope or other marker and used in hybridization reactions, they can detect the homologous sequences in unknown specimens (Figure 4–16) or further refine gel electrophoresis findings (Figure 4–18).

The diagnostic use of DNA probes is to detect or identify microorganisms by hybridization of the probe to homologous sequences in DNA extracted from the entire organism. A number of probes have been developed that can quickly and reliably identify organisms already isolated in culture. The application of probes for detection of infectious agents directly in clinical specimens such as blood, urine, and sputum is more difficult because only a small number of organisms may be present. This problem of sensitively can be overcome by combining probes with NAA methods (see further text). This approach offers the potential for rapid diagnosis and the detection of characteristics not possible by routine methods. For example, a bacterial toxin gene probe can demonstrate both the presence of the related organism and its toxigenicity without the need for culture.

Applications of Polymerase Chain Reaction

The amplification power of the PCR offers a solution for the sensitivity problems inherent in the direct application of probes in clinical specimens. The nucleic acid segment amplified by PCR can be detected by direct hybridization with the probe (Figure 4–19D2) or for greater specificity after electrophoresis and Southern transfer (Figure 4–19D3,4). This approach has been successful for a wide range of infectious agents and awaits only further resolution of practical problems for wider use.

Another creative use of PCR has been in the study of infectious agents seen in tissue but not grown in culture. PCR primers derived from sequences known to be highly conserved among bacteria, such as ribosomal RNA, have been applied to tissue specimens. The amplification produces enough DNA to clone and sequence. This sequence can then be compared with sequences published for other organisms using computers. Thus, taxonomic relationships can be inferred for an organism that has never been isolated in culture.

Ribotyping

Ribotyping also makes use of the conserved nature of bacterial ribosomal RNA and of the ability of RNA to hybridize to DNA under certain conditions. Labeled ribosomal RNA of one organism can be hybridized with restriction endonuclease–digested chromosomal DNA of another. In this case, ribosomal RNA is being used as a massive probe of restriction fragments separated by electrophoresis. Hybridization to multiple fragments is common, but if the organisms are genetically different, the restriction fragments, which contain the ribosomal RNA sequences, will vary in size. The pattern of bands produced by epidemiologically related strains can then be compared side by side.

The application of some combination of the principles described in this chapter is appropriate to the diagnosis of any infectious disease. The usefulness of any individual method differs among infectious agents as a result of biologic variation and uneven study. In general, for agents that can be grown in vitro, culture remains the “gold standard” as both the most sensitive and specific method. Molecular methods have the potential to replace culture and have in some areas, especially virology. Aside from cost, their broader application in infectious disease diagnosis must deal with their highly specific nature. Depending on the clinical situation, the specimen introduced at the beginning of this chapter could be directed at a very narrow or very broad question. If the question is only the diagnosis of a short list of diseases (AIDS, gonorrhea, tuberculosis, malaria), a DNA probe approach can be rapid, sensitive, and practical. Very often, however, the question is “almost anything” or at least a wide range of possibilities. Multiplex PCR platforms can extend the utility of NAA methods; however, culture is difficult to replace totally because it offers broader detection of the common together with a reasonable chance of catching the uncommon and even the rare infection.