Production and Labeling of Probe Nucleic Acid.

In keeping with the requirement of complementation for hybridization, the probe design (i.e., probe length and the sequence of nucleic acid bases) depends on the sequence of the intended target nucleic acid. Therefore, the selection and design of a probe depends on the intended use. For example, if a probe is to be used to recognize only gram-positive bacteria, its nucleic acid sequence must be specifically complementary to a nucleic acid sequence common only to gram-positive bacteria and not to gram-negative bacteria. Even more specific probes can be designed to identify a particular bacterial genus or species, virulence, or an antibiotic-resistance gene present in certain strains in a given species.

In the past, probes were produced through a labor-intensive process involving recombinant DNA and cloning techniques with the nucleic acid sequence of interest. More recently, probes have been chemically synthesized using instrumentation, a service that is commercially available. The base sequence of potential target genes, sequence patterns, or gene fragments for probe design is easily accessed using computer on-line services for nucleic acid sequence information (e.g., GENBANK, National Center for Biological Information). In short, the design and production of nucleic acid probes is now relatively easy. Although probes may be hundreds to thousands of bases long, oligonucleotide probes (i.e., those 20 to 50 bases long) usually are sufficient for detection of most clinically relevant targets.

All hybridization tests must have a means to detect or measure the hybridization reaction. This is accomplished with the use of a “reporter” molecule attached to the single-stranded nucleic acid probe. Probes may be labeled with a variety of molecules, but most commonly, radioactive (e.g., 32P, 3H, 125I, or 35S), biotin-avidin, digoxigenin, fluorescent, or chemiluminescent labels are used (Figure 8-2).

Radioactive labels are directly incorporated through chemical modification into the probe molecule. With the use of radioactively labeled probes, hybridization is detected by the emission of radioactivity from the probe-target complex (see Figure 8-2, A). Quantification of the complexes may be achieved through scintillation counting or densitometry. Although this is a highly sensitive method for detecting hybridization, the requirements for radioactive training, monitoring, licensing, and disposal of radioactive waste have limited the use of radioactive labeling in the diagnostic setting.

Biotinylation is a nonradioactive alternative for labeling nucleic acid probes that involves the chemical incorporation of biotin. Biotin labels are classified as indirect, based on the need for a secondary complex formation. Biotin-labeled probe-target nucleic acid duplexes are detected using avidin, a biotin-binding protein conjugated with an enzyme, such as horseradish peroxidase. When a chromogenic substrate is added, the peroxidase produces a colored product that can be detected visually or spectrophotometrically (see Figure 8-2, B).

Other nonradioactive labels are based on principles similar to those of biotinylation. For example, with digoxigenin-labeled probes, hybridization is detected using antidigoxigenin antibodies conjugated with an enzyme. Successful duplex formation means the enzyme is present; therefore, with the addition of a chromogenic substrate, color production, resulting in color formation, is interpreted as positive hybridization. Alternatively, the antibody may be conjugated with fluorescent dyes that can be directly detected without a secondary enzymatic reaction to produce a colored or fluorescent end product.

Chemiluminescent reporter molecules can be chemically linked directly to the nucleic acid probe without using a conjugated antibody. These molecules (e.g., acridinium or isoluminol) emit light during hybridization between the chemiluminescent-labeled probe and target nucleic acid. The light is detected using a luminometer (see Figure 8-2, C).

Fluorescent labels and fluorimetric reporter groups (e.g., fluorescein and rhodamine) are also considered direct nucleic acid probes. In addition to direct detection probes, fluorimetric reporter groups may be complexed with avidin, digoxigenin, or secondary antibodies, creating a secondary labeling process with additional fluorophores.

Preparation of Target Nucleic Acid.

Because hybridization is driven by complementary binding of a homologous nucleic acid sequence between probe and target, the target nucleic acid must have a single strand and the base sequence integrity must be maintained. Failure to meet these requirements results in negative hybridization reactions as a result of factors such as target degradation, insufficient target yield, and the presence of interfering substances such as organic chemicals (i.e., false-negative results).

Because the relatively rigorous procedures for releasing nucleic acid from the target microorganism can be deleterious to the molecule’s structure, obtaining target nucleic acid and maintaining its appropriate conformation and sequence can be difficult. The steps in target preparation vary, depending on the organism source of the nucleic acid and the nature of the environment from which the target organism is being prepared (i.e., laboratory culture media; fresh clinical material, such as fluid, tissue, or stool; and fixed or preserved clinical material). Generally, target preparation steps involve enzymatic and/or chemical destruction of the microbial envelope to release target nucleic acid, the removal of contaminating molecules such as cellular components (protein), stabilization of target nucleic acid to preserve structural integrity and, if the target is DNA, denaturation to a single strand, which is necessary for binding to complementary probe nucleic acid. Nucleic acid extraction procedures are optimized to ensure a high degree of purity, integrity, and yield of the desired nucleic acid.

Nucleic acid extractions may be classified as organic or nonorganic extractions. Organic extractions use phenol, chloroform, or isoamyl alcohol to disrupt the cellular membranes and denature and remove proteins. After chemical treatment with the organic solution, the mixture is centrifuged, which results in the separation or phasing of the cellular material layered over the top of the organic molecules and waste along the bottom of the tube. The aqueous phase, containing the desired nucleic acid, is then extracted from the organic phase, and the resulting nucleic acid is precipitated using a buffered solution. Nonorganic extractions rely on protein precipitations and nucleic acid precipitations without the use of organic chemicals. Cell membranes and proteins are denatured with a detergent, and the proteins are precipitated with a salt solution. Nonorganic extractions are fast, easy, and do not require the disposal of hazardous organic materials.

DNA isolation is not as technically demanding as RNA extraction methods. RNA may be degraded rapidly by RNAse enzymes. RNAse enzymes are very stable, ubiquitous in the environment, and elevated in certain tissues, such as the placenta, liver, and some tumors. Guanidinium isothiocyanate may be used to denature and inactivate RNAse to preserve the nucleic acid sample before analysis.

Two primary physical methods are available for nucleic acid extraction: liquid-phase extraction, which requires a large sample volume, and solid-phase extraction, which requires a smaller sample volume. Solid-phase extractions are typically simpler than liquid-phase extractions, providing for ease of operation, processing of large batches, high reproducibility, and adaptability to automation. Solid-phase extractions use solid support columns constructed of fibrous or silica matrices, magnetic beads, or chelating agents to bind the nucleic acids.

Mixture and Hybridization of Target and Probe.

Designs for mixing target and probe nucleic acids are discussed later, but some general concepts regarding the hybridization reaction require consideration.

The ability of the probe to bind the correct target depends on the extent of base sequence homology between the two nucleic acid strands and the environment in which probe and target are brought together. Environmental conditions set the stringency for a hybridization reaction, and the degree of stringency can determine the outcome of the reaction. Hybridization stringency is most affected by:

With greater stringency, a higher degree of base-pair complementarity is required between probe and target to obtain successful hybridization (i.e., less tolerance for deviations in base sequence). Under less stringent conditions, strands with less base-pair complementarity (i.e., strands having a higher number of mismatched base pairs within the sequence) may still hybridize. Therefore, as stringency increases, the specificity of hybridization increases and as stringency decreases, specificity decreases. For example, under high stringency a probe specific for a target sequence in Streptococcus pneumoniae may only bind to target prepared from this species (high specificity), but under low stringency the same probe may bind to targets from various streptococcal species (lower specificity). Therefore, to ensure accuracy in hybridization, reaction conditions must be carefully controlled.

Detection of Hybridization.

The method of detecting hybridization depends on the reporter molecule used for labeling the probe nucleic acid and on the hybridization format (see Figure 8-2). Hybridization using radioactively labeled probes is visualized after the reaction mixture is exposed to radiographic film (i.e., autoradiography). Hybridization with nonradioactively labeled probes is detected using colorimetry, fluorescence, or chemiluminescence, and detection can be somewhat automated using spectrophotometers, fluorometers, or luminometers, respectively. The more commonly used nonradioactive detection systems (e.g., digoxigenin, chemiluminescence, fluorescence) are able to detect approximately 10⁴ target nucleic acid sequences per hybridization reaction.

Solution Format.

In the solution format, probe and target nucleotide strands are placed in a liquid reaction mixture that facilitates duplex formation; hybridization occurs substantially faster than with a solid support format. However, before duplex formation can be detected, the hybridized, labeled probes must be separated from the nonhybridized, labeled probes (i.e., “background noise”). Separation methods include enzymatic digestion (e.g., S1 nuclease) of single-stranded probes and precipitation of hybridized duplexes, use of hydroxyapatite or charged magnetic microparticles that preferentially bind duplexes, or chemical destruction of the reporter molecule (e.g., acridinium dye) attached to unhybridized probe nucleic acid. After the duplexes have been “purified” from the reaction mixture and the background noise minimized, hybridization detection can proceed by the method appropriate for the type of reporter molecule used to label the probe (Figure 8-3).

Solid Support Format.

Either probe or target nucleic acids may be attached to a solid support matrix and still be capable of forming duplexes with complementary strands. Various solid support materials and common solid formats exist, including filter hybridizations, southern or northern hybridizations, sandwich hybridizations, and in situ hybridizations.

Filter (membrane) hybridization has several variations. Filter hybridizations are often referred to as “dot blots.” The target sample, which can be previously purified DNA, the microorganism containing the target DNA, or the clinical specimen containing the microorganism of interest, is affixed to a membrane (e.g., nitrocellulose or nylon fiber filters). To identify specimens, samples are usually oriented on the membrane using a template or grid. The membrane is chemically treated, causing release of the target DNA from the microorganism and denaturing the nucleic acid to single strands. The membrane is then submerged in a solution containing labeled nucleic acid probe and incubated, allowing hybridization to occur. After a series of incubations and washings to remove unbound probe, the membrane is processed for detection of duplexes (Figure 8-4, A). An advantage of this method is that a single membrane can hold several samples for exposure to the same probe.

Southern hybridization is another method that uses membranes as the solid support. In this instance, the nucleic acid target is purified from the organisms and digested with specific enzymes to produce several fragments of various sizes (Figure 8-4, B) (also see Enzymatic Digestion and Electrophoresis of Nucleic Acids later in this chapter). The nucleic acid fragments, which carry a net negative charge, are subjected to an electrical field, forcing them to migrate through an agarose gel matrix (i.e., gel electrophoresis). Because fragments of different sizes migrate through the porous agarose at different rates, they can be separated by molecular size. When electrophoresis is complete, the nucleic acid fragments are stained with the fluorescent dye ethidium bromide so that fragment “banding patterns” can be visualized on exposure of the gel to ultraviolet (UV) light. For southern hybridization, the target nucleic acid bands are transferred to a membrane that is submerged in solution, allowing for hybridization of the nucleic acid probe. After hybridization, the southern hybridization membrane is used to detect the specific target nucleic acid fragment carrying the base sequence by using radiolabeled, fluorescent, or substrate-labeled detection. The complexity, time, and labor intensity of the procedure precludes its common use in most diagnostic settings.

With sandwich hybridizations two probes are used. One probe is attached to the solid support, is not labeled, and via hybridization “captures” the target nucleic acid from the sample to be tested. The presence of this duplex is then detected using a labeled second probe that is specific for another portion of the target sequence (Figure 8-4, C). Sandwiching the target between two probes decreases nonspecific reactions but requires a greater number of processing and washing steps. For such formats, plastic microtiter wells coated with probes have replaced filters as the solid support material, thereby facilitating the use of these multiple-step procedures for testing a relatively large number of specimens.

In Situ Hybridization.

In situ hybridization allows a pathogen to be identified in the context of the pathologic lesion being produced. This method uses patient cells or tissues as the solid support phase. Tissue specimens thought to be infected with a particular pathogen are processed in a manner that maintains the structural integrity of the tissue and cells, yet allows the nucleic acid of the pathogen to be released and denatured to a single strand with the base sequence intact. Although the processing steps required to obtain quality results can be technically demanding, this method is extremely useful, because it combines the power of molecular diagnostics with the additional information provided through histopathologic examination.

Peptide Nucleic Acid (PNA) Probes.

PNA probes are synthetic pieces of DNA that have unique chemical characteristics in which the negatively charged sugar-phosphate backbone of DNA is replaced by a neutral polyamide backbone of repetitive units (Figure 8-5). Individual nucleotide bases can be attached to this neutral backbone, which then allows the PNA probe to hybridize to complementary nucleic acid targets. Because of the synthetic structure of the backbone, these probes have improved hybridization characteristics, providing faster and more specific results than traditional DNA probes. In addition, because these probes are not degraded by ubiquitous enzymes, such as nucleases and proteases, they provide a longer shelf-life in diagnostic applications. PNA FISH is a novel fluorescent in situ hybridization (FISH) technique that uses PNA probes to target species-specific ribosomal RNA (rRNA) sequences. Upon penetration of the microbial cell wall, the fluorescent-labeled PNA probes hybridize to multicopy rRNA sequences within the microorganisms, resulting in fluorescent cells. Recently, AdvanDx (Woburn, Massachusetts) introduced in vitro diagnostic kits (using PNA FISH), which have been approved by the U.S. Food and Drug Administration (FDA). These kits can be used to directly identify S. aureus and C. albicans and to differentiate Enterococcus faecalis from other enterococci in blood cultures. In brief, a drop from a positive blood culture bottle is added to a slide containing a drop of fixative solution. After fixation, the fluorescent-labeled PNA probe is added and allowed to hybridize; slides are washed and air dried. After the addition of a mounting medium and a coverslip, the slides are examined under a fluorescent microscope using a special filter set. Identification is based on the presence of bright green, fluorescent-staining organisms (Figure 8-6, A and B). For negative results, only slightly red-stained background material is observed (Figure 8-6, C and D). Multiple studies have been done to evaluate the efficacy of the PNA FISH kits for identifying S. aureus and C. albicans in positive blood cultures. The kits have demonstrated high sensitivity and specificity.

Hybridization with Signal Amplification.

To increase the sensitivity of hybridization assays, methods have been developed in which detection of the binding of the probe to its specific target is enhanced. For example, one commercially available kit uses genotype-specific RNA probes in either a high-risk or low-risk cocktail to detect the human papillomavirus (HPV) DNA in clinical specimens (see Chapter 66). Essentially, sensitivity of HPV detection by hybridization is increased by multimeric layering of reporter molecules, increasing their number on an antibody directed toward DNA-RNA hybrids using chemiluminescence; thus, sensitivity of detection is enhanced by virtue of greater signal produced (i.e., chemiluminescence) for each antibody bound to target.

Extraction and Denaturation of Target Nucleic Acid.

For PCR, nucleic acid is first extracted (released) from the organism or a clinical sample potentially containing the target organism by heat, chemical, or enzymatic methods. Numerous manual methods are available to accomplish this task, including a variety of commercially available kits that extract either RNA or DNA, depending on the specific target of interest. Other commercially available kits are designed to extract nucleic acids from specific types of clinical specimens, such as blood or tissues. Most recently, automated instruments (e.g., the Roche MagNaPure) have been introduced to extract nucleic acid from various sources, such as bacteria, viruses, tissue, and blood (Figure 8-7).

Once extracted, target nucleic acid is added to the reaction mix containing all the necessary components for PCR (primers, nucleotides, covalent ions, buffer, and enzyme) and placed into a thermal cycler to undergo amplification (Figure 8-8). For PCR to begin, target nucleic acid must be in the single-stranded conformation for the second reaction, primer annealing, to occur. Denaturation to a single strand, which is not necessary for RNA targets, is accomplished by heating to 94°C (Figure 8-9). Of note, for many PCR procedures, especially those involving commonly encountered bacterial pathogens, disruption of the organism to release DNA is done in one step by heating the sample to 94°C.

Primer Annealing.

Primers are short, single-stranded sequences of nucleic acid (i.e., oligonucleotides usually 20 to 30 nucleotides long) selected to specifically hybridize (anneal) to a particular nucleic acid target, essentially functioning like probes. As noted for hybridization tests, the abundance of available gene sequence data allows for the design of primers specific for a number of microbial pathogens and their virulence or antibiotic resistance genes. Thus, primer nucleotide sequence design depends on the intended target, such as unique nucleotide sequences, genus-specific genes, species-specific genes, virulence genes, or antibiotic-resistance genes.

Primers are designed in pairs that flank the target sequence of interest (see Figure 8-9). When the primer pair is mixed with the denatured target DNA, one primer anneals to a specific site at one end of the target sequence of one target strand, and the other primer anneals to a specific site at the opposite end of the other, complementary target strand. Usually primers are designed to amplify an internal target nucleic acid sequence of 50 to 1000 base pairs. The annealing process is conducted at 50° to 58°C or higher. Annealing or hybridization of primers is optimized according to the nucleic acid sequence. The nucleic acid sequence of the primer determines the optimal annealing melting temperature (Tm) for the primers. The melting temperature is defined as the temperature at which 50% of the primers are hybridized to the appropriate complementary sequence. Because of the complementary binding of nucleotides, the melting temperature may be determined for a known nucleotide sequence. The melting temperature is calculated according to a simple formula:

2X(A+T)+4 X(G+C)

Primer pairs should be optimally designed to anneal within 1 to 2 degrees of each other to maintain the specificity of the amplification reaction. Once the duplexes have been formed, the last step in the cycle (amplification), which mimics the DNA replication process, begins.

Extension of Primer-Target Duplex.

Annealing of primers to target sequences provides the necessary template format that allows the DNA polymerase to add nucleotides to the 3’ terminus (end) of each primer and extend sequence complementary to the target template (see Figure 8-9). Taq polymerase is the enzyme commonly used for primer extension, which occurs at 72°C. This enzyme is used because of its ability to function efficiently at elevated temperatures and to withstand the denaturing temperature of 94°C through several cycles. The ability to allow primer annealing and extension to occur at elevated temperatures without detriment to the polymerase increases the stringency of the reaction, thus decreasing the chance for amplification of nontarget nucleic acid (i.e., nonspecific amplification).

The three reaction steps in PCR occur in the same tube containing the mixture of target nucleic acid, primers, components to optimize polymerase activity (i.e., buffer, cation [MgCl2], salt), and deoxynucleotides. To minimize the time lag required to alter the reaction temperature between denaturation, annealing, and extension over several cycles, automated programmable thermal cyclers are used. These cyclers hold the reaction vessel and carry the PCR mixture through each reaction step at the precise temperature and for the optimal duration.

As shown in Figure 8-9, for each target sequence originally present in the PCR mixture, two double-stranded fragments containing the target sequence are produced after one cycle. At the beginning of the second cycle of PCR, denaturation produces four templates to which the primers will anneal. After extension at the end of the second cycle, there will be four double-stranded fragments containing target nucleic acid. Therefore, with completion of each cycle, there is a doubling or logarithmic increase of target nucleic acid, and after the completion of 30 to 40 cycles, 10⁷ to 10⁸ target copies will be present in the reaction mixture.

Although it is possible to detect one copy of a pathogen’s gene in a sample or patient specimen by PCR technology, detection is dependent on the ability of the primers to locate and anneal to the single target copy and on optimum PCR conditions. Nonetheless, PCR has proved to be a powerful amplification tool to enhance the sensitivity of molecular diagnostic techniques.

Detection of PCR Products.

The specific PCR amplification product containing the target nucleic acid of interest is referred to as the amplicon. Because PCR produces an amplicon in substantial quantities, any of the basic methods previously described for detecting hybridization can be adopted for detecting specific amplicons. Detection involves using a labeled probe specific for the target sequence in the amplicon. Therefore, solution or solid-phase formats may be used with reporter molecules that generate radioactive, colorimetric, fluorometric, or chemiluminescent signals. Probe-based detection of amplicons serves two purposes: it allows visualization of the PCR product, and it provides specificity by ensuring that the amplicon is the target sequence of interest and not the result of nonspecific amplification.

When the reliability of PCR for a particular amplicon has been well established, hybridization-based detection may not be necessary; confirming the presence of the correct-size amplicon may be sufficient. This is commonly accomplished by subjecting a portion of the PCR mixture, after amplification, to gel electrophoresis. After electrophoresis, the gel is stained with ethidium bromide to visualize the amplicon and, using molecular weight–size markers, the presence of amplicons of appropriate size (the size of the target sequence amplified depends on the primers selected for PCR) is confirmed (Figure 8-10).

Derivations of the PCR Method.

The powerful amplification capacity of PCR has prompted the development of several modifications that enhance the utility of this methodology, particularly in the diagnostic setting. Specific examples include multiplex PCR, nested PCR, quantitative PCR, RT-PCR, arbitrary primed PCR, and PCR for nucleotide sequencing.

Multiplex PCR is a method by which more than one primer pair is included in the PCR mixture. This approach offers a couple of notable advantages. First, strategies including internal controls for PCR have been developed. For example, one primer pair can be directed at sequences present in all clinically relevant bacteria (i.e., the control or universal primers), and the second primer pair can be directed at a sequence specific for the particular gene of interest (i.e., the test primers). The control amplicon should always be detectable after PCR; absence of the internal control indicates that PCR conditions were not met, and the test must be repeated. When the control amplicon is detected, absence of the test amplicon can be more confidently interpreted to indicate the absence of target nucleic acid in the specimen rather than a failure of the PCR assay (Figure 8-11).

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