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Clinical Microbiology Reviews, January 2006, p. 165-256, Vol. 19, No. 1
0893-8512/06/$08.00+0     doi:10.1128/CMR.19.1.165-256.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Real-Time PCR in Clinical Microbiology: Applications for Routine Laboratory Testing

M. J. Espy,* J. R. Uhl, L. M. Sloan, S. P. Buckwalter, M. F. Jones, E. A. Vetter, J. D. C. Yao, N. L. Wengenack, J. E. Rosenblatt, F. R. Cockerill III, and T. F. Smith

Division of Clinical Microbiology, Department of Laboratory Medicine and Pathology, Mayo Clinic, Rochester, Minnesota

SUMMARY
INTRODUCTION
REAL-TIME PCR INSTRUMENTS
REAL-TIME PCR PROBE TECHNOLOGIES
    5' Nuclease (TaqMan) Probes
    Molecular Beacons
    FRET Hybridization Probes
NUCLEIC ACID EXTRACTION
    Manual Extraction
    Automated Extraction
    Auxiliary Procedures To Enhance Extraction
REAL-TIME PCR ASSAY DEVELOPMENT
    Target Nucleic Acid Selection
    PCR Primer and Probe Design
    Assay Optimization
BIOSAFETY CONSIDERATIONS
QUALITY CONTROL AND QUALITY ASSURANCE
    Verification and Validation
    Positive and Negative Controls
    Internal and Inhibition Controls
    Reagents
    Quality Assurance
    Contamination
IMPLEMENTATION OF REAL-TIME PCR TESTING IN THE CLINICAL MICROBIOLOGY LABORATORY
    Facilities Requirements
    Personnel Requirements
    Work Flow Design
        Example of work flow design:
        Example of work flow design:
COSTS
    Royalties
    Reagents and Instrumentation
    Personnel
    Cost Savings at the Bedside
    Coding and Reimbursement
APPLICATION OF REAL-TIME PCR FOR CLINICAL MICROBIOLOGY TESTING
BACTERIA OTHER THAN MYCOBACTERIA SPP.
    General Bacteria
    Slow-Growing or Poorly Culturable Bacteria
    Agents of Community-Acquired Pneumonia
    Agents of Meningitis
    Potential Agents of Bioterrorism
    Bacterial Antibiotic Resistance Genes
MYCOBACTERIA
VIRUSES
    Qualitative Viral Assays
        Herpes simplex virus.
        Herpes simplex virus CNS disease.
        Herpes simplex virus dermal and genital disease.
        Varicella-zoster virus dermal disease.
        Varicella-zoster virus CNS disease.
        Cytomegalovirus CNS disease.
        Epstein-Barr virus CNS lymphoproliferative disease.
        Enterovirus CNS disease.
        Polyomaviruses.
        JCV CNS disease.
        Parvovirus.
        West Nile virus.
    Respiratory Viruses
        Influenza viruses.
        Rous sarcoma virus.
        Adenovirus.
        Metapneumovirus.
        Parainfluenza virus.
        Severe acute respiratory syndrome coronavirus.
    Poxviruses
QUANTITATIVE VIRAL ASSAYS
    Cytomegalovirus
    Epstein-Barr Virus
    BK Virus
    Viral Hepatitis Agents
    Human Immunodeficiency Virus
FUNGI
    Aspergillus Species
    Candida Species
    Pneumocystis jiroveci
PARASITES
    Plasmodium spp.
    Babesia spp.
    Trypanosoma spp.
    Leishmania spp.
    Toxoplasma spp.
    Trichomonas spp.
    Cryptosporidium, Entamoeba, and Giardia spp.
ACKNOWLEDGMENTS
REFERENCES

   SUMMARY
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Real-time PCR has revolutionized the way clinical microbiology laboratories diagnose many human microbial infections. This testing method combines PCR chemistry with fluorescent probe detection of amplified product in the same reaction vessel. In general, both PCR and amplified product detection are completed in an hour or less, which is considerably faster than conventional PCR detection methods. Real-time PCR assays provide sensitivity and specificity equivalent to that of conventional PCR combined with Southern blot analysis, and since amplification and detection steps are performed in the same closed vessel, the risk of releasing amplified nucleic acids into the environment is negligible. The combination of excellent sensitivity and specificity, low contamination risk, and speed has made real-time PCR technology an appealing alternative to culture- or immunoassay-based testing methods for diagnosing many infectious diseases. This review focuses on the application of real-time PCR in the clinical microbiology laboratory.


   INTRODUCTION
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Real-time PCR has revolutionized the way clinical microbiology laboratories diagnose human pathogens (25, 71, 73, 294, 456). This testing method combines PCR chemistry with fluorescent probe detection of amplified product in the same reaction vessel. In general, both PCR and amplified product detection are completed in an hour or less, which is considerably faster than conventional PCR and detection methods. Hence, for some time this technology was referred to as rapid-cycle real-time PCR. Other descriptions of real-time PCR in the early literature included homogeneous PCR and kinetic PCR.

Real-time PCR testing platforms provide equivalent sensitivity and specificity as conventional PCR combined with Southern blot analysis. Since the nucleic acid amplification and detection steps are performed in the same closed vessel, the risk for release of amplified nucleic acids into the environment, and contamination of subsequent analyses, is negligent compared with conventional PCR methods. Real-time PCR instrumentation requires considerably less hands-on time and testing is much simpler to perform than conventional PCR methods. Additionally, accelerated PCR thermocycling and detection of amplified product permits the provision of a test result much sooner for real-time PCR than for conventional PCR. The combination of excellent sensitivity and specificity, low contamination risk, ease of performance and speed, has made real-time PCR technology an appealing alternative to conventional culture-based or immunoassay-based testing methods used in the clinical microbiology for diagnosing many infectious diseases. This review focuses on the application of real-time PCR in the clinical microbiology laboratory.


   REAL-TIME PCR INSTRUMENTS
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Specifications for commercially available real-time PCR instruments, including the nucleic acid probe formats supported, excitation and detection wavelengths, maximum number of samples per run, reaction volumes, and relative thermocycling times are presented in Table 1. The large capacity (≥96-microwell format) instruments, which include the ABI Prism series (7000, 7300, and 7500), the MyiQ and iCycler, Mx4000, MX3000p, Chromo4, Opticon and Opticon 2, and SynChron, may be particularly useful in laboratories with large numbers of specimens. However, thermocycling on these instruments is slower than on other lower capacity instruments, including the LightCycler 1.0, LightCycler 2.0, and SmartCycler II. This is due to the use of a solid-phase material for heat conductance (heating block principle). The large-capacity instruments support high-volume testing while the rapid, lower capacity instruments permit the work flow flexibility that may be especially useful for laboratories that test fewer samples. In summary, work load and work flow issues may dictate which system is best for different-sized laboratories and test volumes.


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  		TABLE 1. Instruments available

 
The Rotor-Gene instrument uses inexpensive standard plastic tubes for the PCR vessel and air for heat transfer with 72 reactions per run. This instrument is intermediate in speed because time is needed for heat conductance to the center of the tubes. The SmartCycler and LightCycler use specialized vessels for rapid heat transfer and can complete a PCR in 30 to 40 min. An additional few minutes are required for the melting curve analysis on the LightCycler.

All the instruments support all or some of the dyes used for TaqMan probes and molecular beacons. Currently, only the LightCycler supports fluorescence resonance energy transfer (FRET) hybridization probe detection with melting curve analysis. Quantitation of target nucleic acid is possible with any of the instruments and supported detection formats.

Recently, analyte-specific reagents (ASRs) and Food and Drug Administration (FDA)-approved kits have become available in the United States for testing on several real-time PCR instruments. The commercial availability of these reagents now make it considerably easier for many clinical microbiology laboratories to adapt real-time PCR testing platforms into their work flow. Because laboratory-developed (also referred to as in-house developed or home brew) real-time PCR tests required considerable expertise to develop and validate, they are generally limited to larger laboratories, especially referral laboratories. The availability of ASRs and kits will also facilitate the development of common testing protocols and standards so that proper comparative clinical studies can be performed and ultimately reliable test results can be ensured for the patient.

In addition to the usual considerations for new instrument purchase (physical space requirement, cost of instrument, disposables, and reagents, instrument maintenance and service, reliability, upgrades, etc.), selection of a real-time PCR instrument and real-time detection format requires consideration of test volume, probe detection requirements, turnaround time for results, personnel requirements, and software. Also, sample preparation requirements must be considered as this can add to the hands-on time per sample, turnaround time, and expense. Several manufacturers have developed semiautomated nucleic acid extraction instruments for use in tandem with real-time PCR instruments. These include the MagNA Pure LC and MagNA Pure Compact for use with the LightCycler instrument, the GeneExpert for use with the SmartCycler II, and the ABI Prism 6700 for use with the ABI Prism series instruments.


   REAL-TIME PCR PROBE TECHNOLOGIES
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One detection method for nucleic acid detection with real-time PCR uses SYBR Green to detect the accumulation of any double-stranded DNA product. SYBR Green provides sensitive detection but is not specific. The use of SYBR Green with instruments that can perform a melting curve analysis to determine the melting temperature, Tm, permits detection of different amplification products based upon the %G+C content and length of the amplification product. This is similar but not equivalent to agarose gel electrophoresis where the separation is based primarily on length. Because SYBR Green assays are not specific, they are often used for screening assays where further analysis of specimens is performed to confirm the results.

Sensitive and specific detection is possible with real-time PCR by using novel fluorescent probe technology probes. Three types of nucleic acid detection methods have been used most frequently with real-time PCR testing platforms in clinical microbiology: 5' nuclease (TaqMan probes), molecular beacons, and FRET hybridization probes (Fig. 1). These detection methods all rely on the transfer of light energy between two adjacent dye molecules, a process referred to as fluorescence resonance energy transfer (500). Collectively, these three types of probes are frequently referred to as FRET probes and this general term has been used in some sections of this review. However, when specifically referring to each of these three types of probes, only FRET appears in the name of one, i.e., FRET hybridization probes. Because FRET hybridization probes consist of two separate probes, the term dual FRET hybridization probes has also been used to describe this specific type of nucleic acid detection method.


Figure 1
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  		FIG. 1. Real-time probe technologies. (A) 5' nuclease (TaqMan) probe. (B) Molecular beacon. (C) FRET hybridization probes. (Reprinted from reference 73 with kind permission of Springer Science and Business Media.)

 
For all types of FRET probes, as the distance between adjacent dye molecules increases, FRET decreases. For TaqMan probes or molecular beacons, the two dye molecules are attached to a single probe. In contrast, for FRET hybridization probes, dyes are attached separately to two probes that align in a head-to-tail configuration on target nucleic acid DNA. (For purposes of discussion in this review, the nucleic acid which is targeted for an assay is henceforth referred to as target nucleic acid or simply the target.) The first dye is a fluorescent dye, and the second can be either a quencher dye or another fluorescent dye which can absorb fluorescent light transferred from the first dye and reemit light at a different wavelength. The proximity of the two dyes in the probe(s) is determined by the nucleic acid architecture of the probe(s). However, the mechanisms to achieve a fluorescent signal with the TaqMan, molecular beacon, or FRET hybridization probe format are different.

5' Nuclease (TaqMan) Probes

The first real-time fluorescent probes developed were 5' nuclease probes, which are commonly referred to by their proprietary name, TaqMan probes (Fig. 1A). A TaqMan probe is a short oligonucleotide (DNA) that contains a 5' fluorescent dye and 3' quenching dye. To generate a light signal (i.e., remove the effects of the quenching dye on the fluorescent dye), two events must occur. First, the probe must bind to a complementary strand of DNA at 60°C. Second, at this temperature, Taq polymerase, the same enzyme used for the PCR, must cleave the 5' end of the TaqMan probe (5' nuclease activity), separating the fluorescent dye from the quenching dye.

A single TaqMan probe can be used for detection of amplified target DNA. If the intent of the assay is to differentiate a single nucleotide polymorphism from a wild type sequence in the target DNA, then a second probe with the complementary nucleotide(s) to the polymorphism and a fluorescent dye with a different emission spectrum is utilized. Thus, TaqMan probes can be used to detect a specific, predefined polymorphism under the probe in the PCR amplification product. For this application, two reaction vessels are required, one with a complementary probe to detect wild-type target DNA and another for detection of a specific nucleic acid sequence of a mutant strain. Because TaqMan probes require 60°C for efficient 5' nuclease activity, the PCR is usually cycled between 95 and 60°C for amplification. In addition, the cleaved (free) fluorescent dye accumulates after each PCR temperature cycle, and therefore can be measured at any time during the PCR cycling, including the hybridization step. This is in contrast to molecular beacons and FRET hybridization probes, for which fluorescence can only be measured during the hybridization step.

Molecular Beacons

Molecular beacons are similar to TaqMan probes but are not designed to be cleaved by the 5' nuclease activity of Taq polymerase (Fig. 1B). These probes have a fluorescent dye on the 5' end and a quencher dye on the 3' end of the oligonucleotide probe. A region at each end of the molecular beacon probe is designed to be complementary to itself, so at low temperatures, the ends anneal, creating a hairpin structure. This integral annealing property positions the two dyes in close proximity, quenching the fluorescence from the reporter dye. The central region of the probe is designed to be complementary to a region of the PCR amplification product. At high temperatures, both the PCR amplification product and probe are single stranded. As the temperature of the PCR is lowered, the central region of the molecular beacon probe binds to the PCR product and forces the separation of the fluorescent reporter dye from the quenching dye. The effects of the quencher dye are obviated and a light signal from the reporter dye can be detected. If no PCR amplification product is available for binding, the probe reanneals to itself, forcing the reporter dye and quencher dye together, preventing fluorescent signal.

Typically, a single molecular beacon is used for detection of a PCR amplification product and multiple beacon probes with different reporter dyes are used for single nucleotide polymorphism detection. By selection of appropriate PCR temperatures and/or extension of the probe length, molecular beacons will bind to the target PCR product when an unknown nucleotide polymorphism is present but at a slight cost of reduced specificity. There is not a specific temperature thermocycling requirement for molecular beacons, so temperature optimization of the PCR is simplified.

FRET Hybridization Probes

FRET hybridization probes, also referred to as LightCycler probes, represent a third type of probe detection format commonly used with real-time PCR testing platforms (Fig. 1C). FRET hybridization probes are two DNA probes designed to anneal next to each other in a head-to-tail configuration on the PCR product. The upstream probe has a fluorescent dye on the 3' end and the downstream probe has an acceptor dye on the 5' end. If both probes anneal to the target PCR product, fluorescence from the 3' dye is absorbed by the adjacent acceptor dye on the 5' end of the second probe. The second dye is excited and emits light at a third wavelength and this third wavelength is detected. If the two dyes do not align together because there is no specific DNA for them to bind, then FRET does not occur between the two dyes because the distances between the dyes are too great. A design detail of FRET hybridization probes is the 3' end of the second (downstream) probe is phosphorylated to prevent it from being used as a primer by Taq during PCR amplification. The two probes encompass a region of 40 to 50 DNA base pairs, providing exquisite specificity.

FRET hybridization probe technology permits melting curve analysis of the amplification product. If the temperature is slowly raised, eventually the probes will no longer be able to anneal to the target PCR product and the FRET signal will be lost. The temperature at which half the FRET signal is lost is referred to as the melting temperature of the probe system. The Tm depends on the guanine plus cytosine content and oligonucleotide length. In contrast to TaqMan probes, a single nucleotide polymorphism in the target DNA under a hybridization FRET probe will still generate a signal, but the melting curve will display a lower Tm. The lowered Tm can be characteristic for a specific polymorphism underneath the probes; however, a lowered Tm can also be the result of any sequence difference under the probes. The target PCR product is detected and the altered Tm informs the user there is a difference in the sequence being detected. Generally, more than three base pair differences under a FRET hybridization probe prevent hybridization at typical annealing temperatures and are not detected.

This trait of FRET hybridization probes is advantageous in cases where the genome of the organism is known to mutate at a high frequency, such as with viruses. When a single or limited number (<3) of known polymorphisms occur between two similar targets, FRET hybridization probes can also be used for discriminating strains of organisms. An example of this application is the identification of herpes simplex virus type 1 (HSV-1) and HSV-2 (Fig. 2). Like molecular beacons, there is not a specific thermocycling temperature requirement for FRET hybridization probes. Molecular beacons and FRET hybridization probes, unlike TaqMan probes, are both recycled (conserved) in each round of PCR temperature cycle. Also, for Molecular beacons and FRET hybridization probes, unlike TaqMan probes, fluorescent signal does not accumulate as PCR product accumulates after each PCR cycle.


Figure 2
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  		FIG. 2. Melting curves obtained after PCR amplification of HSV DNA.

 

   NUCLEIC ACID EXTRACTION
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A critical preanalytical step for real-time PCR assays, as well as any assay in which nucleic acid is analyzed, is nucleic acid extraction. Extraction methods that work for one pathogen in a particular specimen type may not work for another pathogen in another specimen type. For example, herpes simplex virus DNA can be extracted relatively easily from genital swabs (115, 118), whereas extraction of DNA from vancomycin-resistant enterococci in stool samples may be considerably more challenging (451).

A few general comments about extraction of nucleic acid from microorganisms can be made. The thick cell wall of gram-positive bacteria is more difficult to disrupt than the relatively thinner cell wall of gram-negative bacteria. Substances that may inhibit amplification such as heme in blood or bile in stool must be removed. The released nucleic acids should be maintained in an aqueous solution to protect them from degradation. Nucleic acids should be eluted into a small volume in order to maximize detection.

Extraction of clinical specimens can be accomplished either by manual or automated methods. A survey of the literature demonstrates the ability of various commercially available methods to successfully extract a wide variety of specimens for bacterial, viral, and fungal targets (Tables 2 and 3).


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  		TABLE 2. Manual methods of nucleic acid extraction and purification for rapid real-time PCR assays

 

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  		TABLE 3. Automated systems of nucleic acid extraction and purification for rapid real-time PCR assays

 
Manual Extraction

Phenol-chloroform has been used successfully for the extraction of nucleic acids (290, 396). However, phenol is a caustic and corrosive agent, and its use should be considered a safety hazard by clinical microbiology laboratory. A number of commercial manufacturers have developed manual extraction kits for use by clinical laboratories. Some of the most frequently used manual kits as reported in peer reviewed publications are presented in Table 2. These kits vary as to the method, cost, and time required for extraction (Table 2). This variability permits the flexibility in choosing the kit that best suits the needs of a specific laboratory. Manual extraction kits typically use noncorrosive agents making them safe to use by laboratory personnel. While these kits are generally inexpensive and easy to use, they have several drawbacks.

Processing of samples by manual methods requires multiple manipulations. As the number of samples to be extracted increases, so does the potential for contamination due to increased manipulation. In the United States, Clinical Laboratory Improvement Amendments of 1988 (CLIA) regulations (http://www.cms.hhs.gov/clia/) consider manual extraction high-complexity testing, and therefore this type of testing can only be performed by laboratory personnel with appropriate academic credentials. In order to ensure reproducible results, extensive training is necessary to achieve consistency among laboratory personnel performing manual extraction. Some manual kits use ethanol to precipitate the nucleic acids. If not properly removed, excess ethanol residues can inhibit the PCR (502). Finally, manual extraction is a laborious, time-consuming process which requires the undivided attention of the technologist performing this technique in order to ensure optimal results.

Automated Extraction

Automated extraction instruments are manufactured by a number of different companies, and like manual methods vary in method, cost, and time requirements for extraction. Additionally, these instruments vary as to specimen capacity per run and size (footprint) (Table 3). While these systems have not been as widely used as manual methods, a number of studies have reported their utility for extraction of a variety of specimen types (Table 3). Studies which compared manual and automated extraction methods have reported automated extraction to be equivalent and in some instances superior to manual methods (116, 139, 143, 226, 446).

Automated extraction systems have certain inherent advantages over manual methods. Recovery of nucleic acids from automated instruments is consistent and reproducible. Automated extraction systems keep sample manipulation to a minimum, reducing the risk for cross contamination of samples. Many of the instruments are closed systems, further reducing the risk for contamination. Automated systems are typically walk-away systems, and do not require constant attention, which permits personnel to perform other duties. The procedures associated with these instruments could potentially be classified as moderate complexity based on the the Clinical Laboratory Improvement Amendments of 1988 (13). Therefore, laboratory assistants may be able to perform sample extraction with these instruments. Finally once these systems have been validated and proper maintenance procedures are in place, quality control monitoring is less intensive than that required for manual extraction (137).

While the benefits of automated extraction are considerable, there are potential drawbacks. It is most economical when instruments are fully loaded; therefore, a significant number of samples (50 to 100/day) should be processed in order to justify the capital investment that is required for these instruments. The footprints of automated extraction instrumentation may require space that is not currently available in the laboratory. In addition to the cost for equipment, costs for disposables also need to be considered. Some vendors are now manufacturing smaller versions of earlier models of their instruments (Table 3). While these instruments extract significantly fewer samples at a time, they are less expensive and have a smaller footprint than the parent instrument. These smaller versions may be viable options for smaller laboratories which process lower numbers of specimens.

Auxiliary Procedures To Enhance Extraction

Recently, new products have been developed to facilitate the extraction of nucleic acid from clinical samples. Stool transport and recovery buffer (S.T.A.R.; Roche Diagnostics Corporation, Indianapolis, IN) has been used successfully for the extraction of historically challenging specimens such as stool (451). S.T.A.R. buffer has three important properties: infectious organisms are inactivated, degradation of nucleic acids is minimized, and the binding of the nucleic acids to magnetic beads, as is used in the extraction process of MagNA Pure (Roche Diagnostics Corporation), is enhanced.

The swab extraction tube system (S.E.T.S.; Roche Diagnostics Corporation) kit, shown in Fig. 3, is a simple method for rapidly and efficiently recovering specimen attached to and absorbed into the fibers of a collection swab. For some organisms, studies have demonstrated that specimen which is retrieved in a microcentrifuge tube by the S.E.T.S. method, can be directly, or after a quick lysis step (boiling), analyzed by a LightCycler real-time PCR instrument (195, 499). Alternatively the centrifuged material can be extracted by the MagNA Pure instrument to obtain a cleaner preparation of nucleic acids.


Figure 3
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  		FIG. 3. S.E.T.S. The inner tube will hold standard-sized swabs (approximately 5 to 8 mm in diameter, 10 of 25 mm in length) during spinning procedures. The collection tube is used for collection and storage of liquid from the swab. The screw cap is for tightly closing the collection tube.

 
IsoCode Stix (Schleicher and Schuell, Keene, NH), a method for stabilizing blood samples to be transported long distances, can be used to preserve samples for later testing by real-time PCR. This specimen transport device has been coupled with real-time PCR assays for the detection of blood-borne parasites such as malaria (561). This method is not recommended for use with RNA assays.


   REAL-TIME PCR ASSAY DEVELOPMENT
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Target Nucleic Acid Selection

The target primer sequences must be unique in order to identify a specific organism or an organism group, (e.g., group A streptococcus or Mycobacterium genus screen), quantitate a microbe (e.g., cytomegalovirus), or identify unique virulence genes (e.g., verotoxin genes) or genes or mutations associated with antimicrobial resistance (e.g., mecA gene or mutations in rpoB gene associated with rifampin resistance) which can occur across strains or species. Moreover, the PCR primer must be able to identify with high efficiency and specificity the target primer sequences in the specimen of interest (e.g., stool or perianal swab specimens for vancomycin-resistant enterococci). A search for the intended primer sequence in a DNA database such as the National Center for Biotechnology Information (NCBI) database (http://www.ncbi.nlm.nih.gov/BLAST/) may reveal cross-reactivity. However, since the databases currently available represent only a small portion of the nucleic acid sequences for microbes in complex specimen matrices such as stool, specimens and related organisms must also be tested to confirm the lack of cross-reactivity. The target nucleic acid sequence should also be conserved in the organism to be identified or quantitated. If sequence data of the intended target area shows a significant frequency of polymorphisms a more conserved site should be chosen.

PCR Primer and Probe Design

PCR primers provide the first level of specificity for the PCR assay, and primers that only amplify one product will provide the best assay sensitivity. Since real-time PCR also incorporates highly specific homogeneous probe detection, the annealing temperature for probes can be several degrees below the melting temperature of the primers. PCR primers should have a low potential to form secondary structures, including self and crosshybridization with other oligonucleotides in the PCR. This becomes increasingly more difficult as more oligonucleotides are added to the reaction. Details for design of primers and probes are beyond the scope of this review and have been described extensively in two recent publications (191, 500).

Assay Optimization

Optimization of assay conditions can be more challenging for conventional PCR. Due to the numerous manual steps and time requirements for conventional PCR, the assessment of different testing parameters is a painstaking process. For example, several days were frequently required to evaluate the effects of changing a single parameter (e.g., optimal magnesium concentration). Because real-time PCR is more automated and has a shorter test turnaround time, optimization experiments can be performed within hours instead of days.

For real-time PCR a few key components should be optimized in order to achieve maximum results (17, 35, 228, 483). These factors include magnesium concentration, which allows the polymerase enzyme to function at an optimal level; primer and probe concentrations, which affect the sensitivity and specificity of the assay respectively; and the use of additives such as dimethyl sulfoxide, which can aid in the denaturation of nucleic acids with high G+C. The type of polymerase enzyme utilized can also play a significant role, polymerases which permit hot-start PCR are preferrable. These enzymes do not function until a critical maximum temperature is reached, which reduces the generation of nonspecific sequence fragments.


   BIOSAFETY CONSIDERATIONS
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Clinical microbiology laboratories receive and process a wide variety of specimens, including urine, stool, whole blood, plasma, sputum, and swab materials. These specimens may contain a number of transmissible infectious agents including hepatitis viruses and human immunodeficiency virus (HIV). As molecular testing becomes more commonplace, the question of at what point during the extraction process are specimens rendered noninfectious arises. Many extraction kits contain guanidinium salts as one of their compoenents. Studies have shown that guanidinium salts will disrupt cellular integrity and neutralize inhibitory substances (66). However, there are no published studies that demonstrate treatment with guanidinium will ensure that specimens are not infectious.

The MagNA Pure mixes a guanidinium isothiocyanate-containing lysis solution with the sample and incubates it at room temperature for 2 min. We have found (unpublished observations) that this treatment renders 108 Staphylococcus aureus/ml nonviable. However, further studies are required to determine if guanidinium has the same effect on other infectious agents. Until these studies are completed individuals using real-time PCR in clinical laboratories should practice universal precautions, i.e., treating all specimens as if they were infectious (10).

In the past, infectious agents, such as anthrax, have been weaponized for use in biological warfare. The intentional release of anthrax spores in the U.S. mail system in the fall of 2001 emphasized the urgent need for rapid and safe laboratory techniques for detecting Bacillus anthracis in suspicious powders as well as human specimens (179, 289, 350). The Centers for Disease Control and Prevention (CDC) has issued guidelines for the processing and testing of specimens obtained from a suspected outbreak of bioterrorism, in order to protect first-line workers (direct healthcare providers and laboratory workers) (10). In the case of a smallpox outbreak, rapid and accurate laboratory detection is critical in order to quickly contain the infection, however, this may be difficult as smallpox is a level 4 organism, and as such, must be tested at institutions with specialized biosafety level 4 containment facilities (i.e., CDC or United States Army Medical Research Institute of Infectious Diseases).

Autoclaving has been shown to be an effective way to inactivate potential agents of bioterrorism, while permitting the nucleic acid to remain intact for analysis by PCR assays (119, 125, 179, 289, 350). The authors of these publications demonstrated that autoclaving anthrax spores and vaccinia virus, a close relative of smallpox virus, destroyed their ability to be infectious, while not affecting the integrity of their nucleic acid so it could be detected by PCR techniques.

As indicated in the preceding discussion, S.T.A.R. buffer (Roche Diagnostics Corporation) not only stabilizes nucleic acid during transport at room temperatures, but can inactivate pathogens. We have observed that S.T.A.R. buffer can inactivate many bacteria, including such pathogens as Mycobacterium tuberculosis and Escherichia coli OH157:H7 isolated from culture, or present in complex matrices such as respiratory and stool specimens without damaging the integrity of the DNA (unpublished data). The pathogen-inactivating properties of S.T.A.R. buffer provides laboratories an added level of safety for processing and transporting pathogens for nucleic acid analysis.


   QUALITY CONTROL AND QUALITY ASSURANCE
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Verification and Validation

Clinical relevance, cost, instrumentation, and ease of performance should be considered when evaluating a new test procedure (109), but of primary importance is the verification and validation of test performance. The ability of a laboratory test to consistently produce accurate and precise results is not only essential, it is the core of quality assurance programs for clinical laboratories (302). A detailed protocol for the verification of new test methods should be established by the laboratory prior to the verification procedure. Table 4 provides guidelines for verification of new test methods (333). Additionally, documentation of validation is necessary to demonstrate that a verified test continues to perform according to the laboratory's requirements. These procedures help ensure the consistency of the results and that laboratory personnel remain competent to perform tests and report results.


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  		TABLE 4. Verification guidelines

 
Guidelines developed by regulatory agencies are not current for real-time PCR applications in clinical microbiology. The Clinical Laboratory Standards Institute (formerly the National Committee for Clinical Laboratory Standards) published a set of guidelines for molecular diagnostic methods in infectious disease testing in 1995; however, these guidelines were provided before the introduction of real-time PCR technology (332). These are considered guidelines, not standards, for infectious disease testing, and currently are undergoing revision.

The most recent document addressing quality control standards for molecular test systems is the revised CLIA 1988 document published in the Federal Register, 24 January 2003 (4). This document addresses requirements for certain quality control provisions and personnel qualifications. It combines and reorganizes requirements for test management, quality control and quality assurance, and also changes the requisite consensus for grading proficiency testing challenges. The CLIA 88 document stipulates that prior to test implementation, clinical laboratories verify the manufacturer's performance specifications and confirm they can be replicated by laboratory personnel when following the procedure. For laboratory developed tests or modification of test systems, laboratories are required to establish their own performance specifications prior to implementation of the new or modified test.

Because nucleic acid test methods are changing and evolving so rapidly, existing guidelines have been difficult to apply. The challenges to clinical laboratories include determining the type of verification experiments required for a real-time PCR assay and an acceptable number and type of specimens to evaluate. Providing a single set of guidelines for real-time PCR which envelops all the necessary verification and validation by all accreditation agencies would be of great benefit to laboratories acquiring this new technology. Along with the need for a well defined quality control program for real-time PCR qualitative assays, there is need for guidelines for quantitative real-time PCR assays. To date such guidelines only exist for a select number of blood-borne viruses (341).

Quality control allows the laboratory to minimize the reporting of inaccurate results, to report results with a high degree of confidence and to decrease costs by detecting errors prior to reporting results (137). One goal of the laboratory quality control program is to reduce the number of controls needed for reporting acceptable results. The following information relates to specific controls used during testing as well as the quality control of reagents used for testing. This discussion is not intended to be all-inclusive nor definitive, and is based to some extent on experience at Mayo Clinic with real-time PCR and our interpretation of published guidelines for generic molecular testing.

Positive and Negative Controls

Ideally, patient specimens containing the target nucleic acid are used as the positive control, but this is often not practical or feasible. An acceptable positive control is pooled negative specimens spiked with whole organisms or if that is not available, a representative sample of the nucleic acid to be detected. The positive control should be at a concentration near the lower limit of detection of the assay to challenge the detection system yet at a high enough level to provide consistent positive results.

A blank control such as water or buffer is often used as a negative control. However, an optimal negative control is a sample containing nontarget nucleic acid to demonstrate that nonspecific PCR amplification and detection of amplified product is not occurring. In addition, the negative control is used to demonstrate that the reagents are not contaminated with target nucleic acid and can be used to compensate for background signal generated by the reagents. The recommendation for a negative control every fifth tube to monitor PCR contamination (332) may be excessive with real-time PCR assays. The closed system for amplification and detection used with real-time PCR virtually eliminates the amplicon contamination caused by the opening and closing of reaction vessels which is problematic with conventional PCR and detection methods. Even with the closed system of real-time PCR, the laboratory may still choose to add uracil-N-glycosylase to the PCR mix for another level of amplicon contamination control.

The 2003 revisions to CLIA 88 rule does not specifically address real-time PCR assays but recommendations from the molecular testing sections can be applied to real-time PCR assays. Table 5 summarizes the 2003 revised CLIA 88 document (4) and CLSI (332) quality control recommendations. In both of these documents it is recommended that each molecular amplification run of samples contain positive and negative controls. Additionally, in the CLIA document it is indicated that a test system which includes nucleic extraction also include a positive and negative control, with the positive control capable of detecting errors in the nucleic acid extraction procedure. For a quantitative assay, two positive controls representing two different concentrations of target nucleic acid are recommended in both the CLIA 88 and CLSI documents. Laboratories should establish the number and frequency of controls based on manufacturers criteria and agency recommendations.


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  		TABLE 5. Positive and negative control recommendations for molecular testing

 
Internal and Inhibition Controls

An acceptable specimen should be free of inhibitory substances that could produce a false-negative result. Some clinical samples may contain substances which are not always removed by the extraction process and which may inhibit the PCR amplification. Inhibition of amplification can be detected by the introduction of an internal control, also referred to as a recovery template.

Based on the requirements of regulatory or accrediting agencies, individual laboratories should determine when an internal control is required in an assay. For example, the 2003 revision to CLIA 88 document does not address internal controls nor have a requirement for assessment of inhibition of PCR chemistry. In contrast, the College of American Pathologist (CAP) molecular pathology inspection checklist indicates that the laboratory must determine the likelihood of the generated result being a false-negative result due to inhibition when there is no amplification of product (76). If the test is performed according to manufacturer instructions, published data containing the inhibition rate may be used for documentation. Internal controls for laboratory developed assays or modified FDA assays should be determined on a case-by-case basis taking into account the probability that the specimen source contains inhibitory substances. Specimen types such as stool or sputum are generally more inhibitory to PCR chemistry than serum or plasma specimens. Also, the assay performance characteristics (sensitivity, specificity, accuracy, etc.), the implications of a false-negative result and the degree to which a clinical diagnosis depends on laboratory results, require consideration. Internal controls may be naturally present in the original specimen, added to the specimen prior to extraction, or added to the PCR reagent mix before amplification. The simplest way to establish inhibition is to add target nucleic acid to a portion of the sample and perform the test to show that if target nucleic acid were present, the PCR would have been positive. Unfortunately, this approach increases the cost of the assay.

Examples of the different types of internal controls that have been used for real-time PCR assays are shown in Table 6. Homologous and heterologous internal controls are those which do not naturally occur within the specimen source. These have also been referred to as exogenous controls as they must be added to the specimen. Homologous controls are coamplified with target DNA using the same PCR primers. However, the internal sequence of the homologous control DNA internal to the PCR primer sites is genetically engineered to be different from the target DNA such that a different product signal occurs with FRET detection.


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  		TABLE 6. Examples of internal controls used in real-time PCR assays

 
An example of a homologous internal control is shown in Fig. 4. Heterologous controls consist of separate amplifiable targets. Since these do not contain the target sequence, a separate set of PCR primers and probes are required for amplification and detection respectively. Housekeeping genes occur naturally within the specimen being tested and therefore are referred to as endogenous controls (341). The housekeeping genes occur in all human nucleated cell types and therefore these types of controls are commonly used in human genetic studies. There is no single housekeeping gene that is suitable for all experimental conditions and articles have been published on the variability of certain housekeeping genes in different systems (326, 498).


Figure 4
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  		FIG. 4. Recovery template. The recovery template (internal control) has the same sequence as the PCR product except the probe region has been replaced with a sequence complementary to recovery template probes. FRET detection of the target DNA is with a probe labeled with the Red 640 dye in channel 2 of the LightCycler while the recovery template is detected with a probe labeled with the Red 705 dye in channel 3. A small amount of recovery template is added to the PCR and is amplified along with the target DNA by the same primers. Thus, the two reactions compete for the primers. Normally, the recovery template is amplified in all samples, including the negative control. If neither the recovery template nor target DNA is amplified, then it is assumed that inhibition of the PCR has occurred and the test for that sample is not valid. However, if target DNA is amplified but the recovery DNA template is not amplified, then it is assumed that the target DNA is present in a proportionally greater amount. In this situation, partial inhibition of the PCR may be present but the target DNA is successfully amplified or the recovery template may not be able to compete for primers and the recovery template signal may be weak or not present. When this occurs, the positive result is valid because the recovery template amplification result is unnecessary.

 
Real-time PCR assays used in microbiology require optimal sensitivity and the use of internal controls should not decrease the sensitivity of the assay. Performing competitive assays by amplifying serial dilutions of the target DNA with and without the internal control should reveal if the sensitivity of the assay is affected (501). Generally, procedures for synthesis of homolgous internal controls are too complex for the clinical microbiology laboratory (55). A number of manufacturers of real-time PCR ASRs and kits are including homologous internal controls for their assays, which obviates this cumbersome task for the novice user of real-time PCR.

Reagents

The quality control of reagents is extremely important to ensure the success of real-time assays (56). Frequently, commercially available master mix components that contain standardized concentrations of reagents are available, but these do not always include PCR primers and FRET probes.

Whether PCR primers are purchased from vendors or laboratory developed, some method of chromatographic purification should be applied. Purification recovers oligonucleotides of the correct length. Truncated oligonucleotides can affect a PCR by consuming reaction reagents and forming nonspecific amplification products (159). The presence of these irregular oligonucleotides can also falsely elevate the final concentration of the working primers thus affecting the performance of the assay. The CLSI MM3 guideline recommends that laboratories obtain a certificate of analysis from PCR primer vendors. These certificates may contain sequence data, base composition, molecular weight of the sequence, concentration and method of purification (332). Vendors may provide PCR primer concentration, but these concentrations should be verified in the laboratory. Laboratories synthesizing their own oligonucleotide PCR primers should perform chromatographic purification and determine also PCR primer concentration. In compliance with the CAP Molecular Pathology inspection checklist, each new lot of reagent should be tested in parallel with the old reagent lot using both positive and negative patient samples ensuring the same results are obtained with both reagent lots (76).

Purification of FRET probes is especially important to separate both dye and oligonucleotides that have not coupled to form the FRET probes and to remove oligonucleotides with an incorrect length (554). While the quality of probe synthesis has improved greatly over the past few years, quality control is required to avoid probe lots with reduced performance. The CLSI MM3 guideline for PCR primers discussed above should also apply to FRET probes. Some vendors provide a tracing (chromatogram, polyacrylamide gel electrophoresis analysis, etc.) of the purified FRET probe as part of their quality control documentation which may augment quality control. Another quality control service provided at a nominal charge by some vendors is to determine if a particular FRET hybridization probe set is capable of producing FRET. The company will design and synthesize an oligonucleotide complementary to both probes and a melting curve analysis is performed. The production of a melting peak at the predicted Tm will confirm that the FRET hybridization probe set is capable of producing FRET and is acceptable to use. This probe validation process can also be completed in the laboratory by following the method provided on the Idaho Technology website (http://www.idahotech.com) under probe classroom.

Proper storage of reagents can result in an increase in shelf life. FRET probes may arrive in a lyophilized form and the recommendation is to store them at room temperature until resuspended. Some manufacturers state that the hydrated probes should be stored at 4°C for daily use or aliquoted into smaller volumes and stored at –20°C. Numerous freeze-thaw cycles can be detrimental to the FRET probes. The PCR primers can be stored in a similar manner as the probes. The completed master mix (containing primers and probes) can be stored at –20°C for extended periods of time, without degradation of the mix (452). We have also found this to be true with most of our real-time PCR assay master mix reagents used at the Mayo Clinic. The complete mix is stored at either 4°C or –20°C (assay dependent) for 1 to 6 months without loss of activity. However, we have observed that the length and temperature of storage are assay dependent and conditions of storage require validation for each assay. The advantage of freezing the master mix is assay reproducibility, time savings in setting up assays, and reduced reagent contamination (452).

Quality Assurance

After implementation of the real-time PCR test it is necessary to continue to monitor performance of the assay, equipment, reagents, and personnel. For example, technologists monitor patient specimen positivity rates for all real-time PCR assays used in our institution on a weekly or monthly basis. If a sudden increase in positivity rate occurs, this could reflect seasonal variances of disease frequency (e.g., influenza virus or group A streptococcus), disease outbreak, or specimen-to-specimen or amplification product contamination. Daily quality control of reagents including positive and negative controls and/or extraction controls should be performed. In compliance with accrediting and regulatory agencies, comparable performance of new reagent lots compared with old reagent lots should be verified. Instrument performance should be assessed biannually when multiple instruments are used interchangeably, also as required by accrediting and regulatory agencies. Competency of personnel performing tests must also be evaluated. Examples of competency assessment are included under the Personnel Requirements section below.

Contamination

The risk of contamination is considerably less with real-time PCR compared to conventional PCR, but still can occur (341). Since real-time PCR amplification is performed in a closed system, there is no need for individual air-controlled rooms as is recommended for conventional PCR. In our experience with real-time PCR, specimen to specimen contamination has become a greater challenge than amplified product contamination. The most obvious situation where specimen-to-specimen contamination can occur is with the transfer of specimen to the PCR vessel or to the DNA extraction tube. Care must be taken to avoid contamination of the pipette device with specimen and to avoid the creation of an aerosol by blowing out the specimen from the tip.

Certain types of sample sources are known to contain a high concentration of organisms that may lead to specimen-to-specimen contamination, namely, viral agents. The inclusion of negative controls and continual trend analysis of the assay are used to recognize a contamination event. Additionally, unidirectional work flow should be followed. Separation of procedural steps will require separate work spaces in the laboratory, as detailed below under the ssection on facilities requirements. As with all methods performed in the laboratory, good laboratory practice is critical for accurate results.


   IMPLEMENTATION OF REAL-TIME PCR TESTING IN THE CLINICAL MICROBIOLOGY LABORATORY
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Implementation of real-time PCR testing platforms in the clinical microbiology laboratory requires careful consideration of facility requirements, personnel requirements, and work flow design. These considerations are similar to those required for implementation of any new type of testing method. A review of these requirements related to our experience at Mayo Clinic with implementing LightCycler technology into the clinical microbiology laboratory is provided in the following discussion. At the Mayo Clinic, some of our real-time PCR assays have been used routinely in the clinical laboratory since early 2000.

Facilities Requirements

As previously mentioned, a physical separation of processes, equipment, and reagents is recommended, to minimize the risk of specimen-to-specimen contamination. Four different work areas are suggested, including a reagent preparation area to prepare PCR master mix, a sample processing area where procedures, including nucleic acid extraction, occurs, a target loading area where the specimen is added to the PCR master mix in the reaction vessel, and an amplification area where thermocycling and probe detection occurs.

The reagent preparation area should be kept free of all patient specimens and DNA extracts. Protocols for the sample preparation area should minimize the number of tubes that are simultaneously open. Each of the work areas should contain dedicated working materials, reagents, and pipetting devices. Reagents should be prepared and aliquoted into single use or small volumes. This ensures ease of use and less chance for contamination.

All working surfaces should be cleaned before and after use, preferably with a reagent that destroys nucleic acid such as a 5% bleach solution. The manufacturer's recommendations should be followed for cleaning of instrument components (e.g., carousels with the LightCycler), processing blocks, and other instrument surfaces and parts.

Gloves should be changed frequently, at least before beginning each of the separate tasks required in a dedicated work area and should always be changed if moving from one work area to another work area. The use of aerosol-resistant pipette tips and pipette tips long enough to prevent specmien contact with the pipetter aids in the prevention of specimen contamination (502). Enzyme contamination control systems such as uracil-N-glycosylase can be incorporated into the real-time PCR master mix as an added safeguard to sterilize amplified product that may be carried over to subsequent batches of tests.

Personnel Requirements

Personnel should be trained in both the preanalytical (specimen processing and extraction) and the analytical procedures. Many current laboratory professionals do not have training or experience in molecular methods and also lack theoretical knowledge of molecular microbiology. Based on our experience at the Mayo Clinic, providing a variety of methods for attaining this knowledge is useful. Some vendors are willing to provide overview presentations on molecular biology as well as technical information on their specific testing platform. Appreciation of the fundamentals will help to avoid cookbook testing and will later allow more careful and focused troubleshooting.

Clinical microbiologists who have not had formal training in molecular microbiology still possess many of the critical skills necessary for success in performing real-time PCR testing. Especially important is meticulous attention to detail, strict adherence to standard operating procedures, and use of aseptic technique.

These skills are easily transferable from culture based conventional microbiologic testing to real-time PCR testing. At the Mayo Clinic we noted that providing training on the basics of accurate pipetting was fundamental, especially for those lacking experience with micropipetting.

Well-written training materials, including training checklists and detailed standard operating procedures for each real-time PCR test, should be available. The training checklist serves to standardize the training of all personnel. At the Mayo Clinic, we believe that identifying a technical expert to provide one-on-one training for real-time PCR is critical. Technologists are required to successfully complete a panel of unknown samples and perform the procedure under direct observation of the technical expert to ensure flawless manipulations throughout the procedure. They also are required to analyze a previous run of samples with a variety of unusual results. This allows them to perfect their skills manipulating the computer software associated with the real-time PCR instrument and ensures consistent analysis and reporting of results. Overall, our technologists have been very enthusiastic about implementation of real-time PCR and excited to learn the new technology.

The availability of resources for troubleshooting is a consideration when selecting a molecular platform for the clinical laboratory. Laboratory-developed tests require that the technical resources to resolve problems related to the assay are available within the laboratory. Use of ASRs and United States Food and Drug Administration-approved tests allow the use of technical support resources available from the manufacturer for troubleshooting problems related to the assay or instrumentation.

Work Flow Design

After selection and successful implementation of a real-time PCR testing platform into the clinical laboratory, efficiencies may be gained by the implementation of additional tests which use the same methodology. Different real-time PCR tests may have subtle variations (e.g., differences in nucleic acid extraction procedures), but overall the methodology is very similar. This attribute reduces the personnel resources required for training and implementation of subsequent tests.

Not unlike other microbiology tests, work flow and testing schedules for real-time PCR tests are determined by the arrival times of specimens into the laboratory, clinical urgency for the results, and laboratory hours of operation. Many of the real-time PCR platforms are most efficiently run in a batch mode. Some vendors provide identical thermocycling protocols for different ASRs for the same instrument. This allows testing for multiple analytes within the same run, which enhances the efficiency of the testing platform.

Example of work flow design: real-time PCR for detection of group A streptococci from throat swabs. At the Mayo Clinic in August 2002 we replaced a conventional testing method (rapid antigen screen with backup culture for rapid antigen negative results) with the LightCycler Strep-A assay (Roche Diagnostics Corporation, Indianapolis, IN) (456, 499). This real-time PCR test is as sensitive as the gold standard method, culture, and provides same-day conclusive results for all patients, whereas the antigen/backup culture method required up to 48 h for a conclusive result for the majority of patients. A simple lysis and extraction method of the swab sample is performed using the S.E.T.S. tube (Roche Diagnostics Corporation) before testing in the LightCycler (Fig. 5).


Figure 5
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  		FIG. 5. Work flow algorithm for processing specimens for the laboratory diagnosis of group A streptococcal infections by real-time PCR.

 
Prior to implementing this real-time PCR assay we worked extensively with our clinical colleagues to review the performance characteristics of this novel testing method, prepare educational materials for patients, determine an appropriate testing schedule, and clarify what health care providers and patients should expect. One misconception of our healthcare providers that we had to clarify was that a majority of patients had conclusive results using the conventional antigen screen. In fact, the sensitivity of the rapid antigen test in our hands was approximately 50% compared to culture. Therefore, during the height of the streptococcal pharyngitis season, when the incidence of true-positive results was {approx}30%, {approx}85% of patients had to wait up to 48 h for a conclusive result by culture. This was because with {approx}30% true-positive cases and {approx}70% true-negative cases, half of the true-positive cases, or 15%, were detected by rapid antigen and the other half, or 15% of true-positives, as well as the 70% true-negatives required detection by culture. Cultures are held for 48 h.

In discussions with our health care providers, it became clear that there were other considerations beyond education on test performance. These included, specimen transportation and arrival times, (especially for samples coming from more distant clinics) and issues related to the expeditious provision of antibiotics (i.e., closing times of local pharmacies). Based on these considerations, we implemented testing five times daily (4:00 a.m., 11:30 a.m., 3:00 p.m., 5:30 p.m., and 8:30 p.m.), 7 days/week, with additional batches set up as needed during the peak season.

Additionally, we have eliminated most of the follow-up procedures required by health care providers by using the following innovative processes with our clinical colleagues (456). Patients identify their preferred pharmacy at the time the throat swab is collected. A standardized prescription form (including the antibiotics prescribed and the patient's preferred pharmacy) is completed by the healthcare provider, which then accompanies the specimen to the laboratory. At the conclusion of the test run, results are entered into the laboratory information system and transmitted to the patient's electronic medical record. The results from the electronic medical record are delivered to a computerized message center, allowing patients to obtain their secure results by telephone and pick up the prescription prepared for them at their selected pharmacy. Prescriptions are faxed to numerous local and regional pharmacies by the clinical microbiology laboratory staff (Fig. 5).

In a comparison of the personnel required for performing rapid antigen test and back up culture of antigen negative specimens, we realized a savings of 2.1 full time equivalents. This is based on an annual testing volume of 26,000 tests with the rapid antigen test being performed at four satellite locations in Rochester, Minnesota. Performance of the Roche Strep-A ASR test allowed us to centralize testing in one location. This results in a more efficient process that saved personnel effort.

In summary, t