Antibiotics in Laboratory Medicine, 6 Ed.

Chapter 8. Applications, Significance of, and Methods for the Measurement of Antimicrobial Concentrations in Human Body Fluids

Shelly Latte, Emilia Mia Sordillo, and Stephen C. Edberg

The proliferation of antimicrobial agents in recent years presents clinical laboratories with a potentially large number of antibiotics to assay. Many of the methods described in the previous edition of this book (367) remain the testing standard, whether for routine clinical therapeutic drug monitoring (e.g. automated immunoassay based methods) or for monitoring during the initial evaluation of a new antimicrobial agent (e.g. high pressure liquid chromatography/HPLC). Method modifications that have been introduced to increase assay sensitivity include improved sample preparation methods prior to HPLC and utilization of mass spectrometry for analyte detection following HPLC.

Automated chemical methods for the determination of antibiotic concentrations in body fluids, most commonly serum or plasma, have become commonplace and can be performed with relative ease at reasonable costs. These automated methods have largely replaced bioassays, which are not routinely performed. Therapeutic drug monitoring of aminoglycosides is generally performed by enzyme immunoassay (EIA), particle-enhanced turbidimetric inhibition immunoassay (PETINIA), fluorescence polarization immunoassay (FPIA), cloned enzyme donor immunoassay (CEDIA), chemiluminescent microparticle immunoassay (CMIA), or similar labeled immunoassays because of their ease of performance, rapid turnaround times, and reasonable costs.

For example, Abbott Laboratories (Abbott Park, IL) produces CEDIAs, CMIAs, and PETINIAs for gentamicin, tobramycin, amikacin, and vancomycin which can be run on Abbott’s Architect, an automated chemistry analyzer. Other immunoassay-based tests are available from numerous manufacturers including Beckman-Coulter (Jersey City, NJ), Roche Diagnostics (Indianapolis, IN), Siemens Diagnostics (Tarrytown, NY), and Vitros (Ortho Clinical Diagnostics Inc, Rochester, NY.)

The PETINIA assays previously produced by Dade Behring, Inc, for gentamicin, tobramycin, amikacin, and vancomycin were acquired by Siemens AG in 2007, and can be run on the company’s Dimension and Dimension Vista instruments. These assays use latex particle antibiotic conjugates to compete for monoclonal antibody binding sites with the free, unconjugated antibiotic in the sample. The binding of unbound antibiotic to the antibody decreases the rate of aggregation of the particle-antibody complexes, which is inversely proportional to the concentration of antibiotic in the sample.

Roche Diagnostics’ (Indianapolis, IN) CEDIA gentamicin II and tobramycin assays run on several of the company’s automated chemistry analyzers. In the CEDIA assays, an inactive fragment of recombinant bacterial β-galactosidase that has been genetically engineered into two inactive fragments is conjugated to the study of antibiotic. Under assay conditions, the two enzyme fragments spontaneously reassociate to form a fully active enzyme that cleaves a chromogenic substrate. A color change is generated that is spectrophotometrically measured. Binding of a monoclonal antibody directed at the enzyme-drug conjugate interferes with enzyme reassociation. Free analyte in the sample competes with the conjugated drug for antibody sites. The quantity of enzyme that is formed and the resultant absorbance change are directly proportional to the amount of analyte that is present in the sample.

Vancomycin and chloramphenicol (CAM) levels are routinely obtained through the use of immunoassay or high-performance liquid chromatography (HPLC)–based methods. Determination of sulfonamide, trimethoprim, β-lactam, macrolide, quinolone, tetracycline, and other antimicrobial levels is largely confined to reference laboratories. Methods include HPLC, immunoassay, and bioassay, although this latter method has fallen into disfavor because of assay variability and potential interference when multiple drugs are administered. While radioimmunoassay (RIA) and gas chromatography techniques can also be used for antibiotic concentration analysis, these methods generally confer little advantage over existing immunoassays and are infrequently used.

Interestingly, despite the major advances in technology and the large increase in the number of antimicrobial drugs, the clinical indications for therapeutic monitoring of antibiotic concentrations are limited to a few select drugs and clinical settings. This is largely because of the improved toxicity profiles and large therapeutic windows of the majority of currently used antimicrobials. Most routine antimicrobial therapeutic drug monitoring involves aminoglycoside and glycopeptide (vancomycin) antibiotics and, to a lesser extent, CAM. Antituberculosis, antifungal, and antiretroviral therapies present additional applications in which therapeutic drug monitoring may prove to be of value.

Recently, concerns regarding increasing resistance among pathogenic organisms and the emergence of obesity as a major health issue have highlighted the importance of weight-based dosing paired with assessment of blood levels for many drugs. A notable example is the recommendation to maintain serum vancomycin through levels above 10 mg per L, and from 15 to 20 mg per L in serious infections, as outlined in the 2009 consensus guideline from the American Society of Health-System Pharmacists, the Infectious Diseases Society of America, and the Society of Infectious Diseases Pharmacists (373).

The assay of antibiotics or antibiotic material dates back to the demonstration of lysozyme in an agar diffusion assay system by Fleming (1) in 1922. Thus, the first use and many of the other initial uses of assay systems were for the detection of antibiotic activity that resulted from the in vivo production of antimicrobial substances. Assays of antibiotics in blood, urine, and other body fluids and tissues were primarily performed in conjunction with the determination of antibiotic pharmacology and pharmacokinetics.

In the late 1960s, accurate, rapid assays to measure blood levels of antimicrobial substances, particularly those with narrow toxic/therapeutic ratios, were sought (27). The value of such information for the optimal use of certain antibiotics eventually became apparent, but the popularization of rapid serum assays required most of the decade of the 1970s. Data from several laboratories demonstrated that the efficacy of some antibiotics correlated with their peak serum antibiotic levels (8,9; Williams DN et al., unpublished data, 1984). Primarily, as a consequence of this work, the measurement of aminoglycoside levels began to be performed in all seriously ill patients who received these drugs, irrespective of the patients’ renal status (10).

Toxicity was also shown to be related to both peak and trough serum levels. Although the study by Line et al. (11) showed a correlation between streptomycin toxicity and trough levels, the concept that trough levels correlated better with toxicity than peak levels for aminoglycoside drugs remained in dispute for some time. Indeed, although it was quite clear from retrospective analyses that toxicity was associated with high serum levels of these antibiotics, it was difficult to prove that toxicity occurred at particular blood levels for a given drug. Table 8.1 demonstrates the toxic/therapeutic ratios of some commonly used aminoglycosides (12).

The measurement of antibiotic concentrations in various fluids has been a prominent aspect of the evaluation of new antibiotics and the quality control of their manufacture. With the availability of rapid, accurate assays, the measurement of antibiotic material in serum and other body fluids is feasible, desirable, and widely practiced for these purposes. Clinically, such assays have been used primarily for the determination of aminoglycoside levels in serum, for which peak levels primarily establish adequacy of therapy, while trough levels reflect potential drug accumulation and toxicity.

Although the most common reason for performing rapid serum antibiotic assays in hospitals is to regulate therapy with aminoglycoside antibiotics and vancomycin, there are other indications. For patients who have organ dysfunction, such as hepatic or renal failure, it is desirable to know whether or not antibiotics that are metabolized or eliminated (e.g., CAM) by damaged or imperfectly functioning organs accumulate. In some patients who are taking oral antibiotics, it may be important to know the extent to which they are absorbing the drugs from their gastrointestinal tracts (1316). Therapeutic monitoring of β-lactam, macrolide, tetracycline, and other antibiotics with wide therapeutic windows occurs infrequently, but may be performed to assess compliance, and/or in patients with renal failure.

Aminoglycoside levels are usually ordered to establish the adequacy of therapy and to prevent toxicity. Retrospectively, there is a correlation between both peak and trough levels and toxicity. In one study, Black et al. (17) found a significant correlation between ototoxicity of amikacin and both peak and trough levels, with the P value being slightly lower for the peak than the trough value.

Another indication for the measurement of antibiotics from human body fluids has been the establishment of therapeutic levels of drugs in various body fluids. The field of the pharmacokinetics of antibiotics has developed rapidly, along with the number of drugs that are currently available to treat serious infections. The concept of the “class” or “type” antibiotic that is representative of all related members is no longer valid. For example, we can no longer predict the tissue distribution of all cephalosporins based on the activity of cephalothin. Many of the third-generation cephalosporins appear in extravascular body fluid compartments, while cephalothin does not. Therefore, the ability to assay each cephalosporin level in cerebrospinal fluid is desirable. As pharmaceutical chemists modify currently available antibiotics, additional generations and individual drugs will continue to be developed.

In the 1960s and 1970s, most work concentrated on microbiologic assay systems for the major classes of antibiotics. The 1970s witnessed the evolution of reference methods to rapid procedures that became available in clinical laboratories. By the late 1970s, approximately 75% to 85% of all assays in clinical laboratories were microbiologic in nature, with most of the remainder being RIAs (18). Subsequently, commercially available, nonisotopic immunoassays largely replaced microbiologic assays. Methods for the assay of virtually all classes of antibiotics by HPLC have become available and can be performed in a broad spectrum of laboratories. Immunoassays for aminoglycosides and vancomycin have become routine “black box” procedures because of the widespread availability of commercial kits. Assays for antituberculous, antifungal, and antiviral medications can be found in their respective chapters.


The selection of an assay method depends on the clinical and research needs and on the capabilities and resources of the laboratory in which the assay is to be performed. For high-volume assays that require rapid turnaround, commercially sold immunoassays that use specific monoclonal antibodies and are performed on automated chemistry analyzers will be the methods of choice. Low-volume assays for which monoclonal antibodies, commercial kits, or automated chemistry analyzers are unavailable may be performed by HPLC or, in some instances, bioassay. As a rule of thumb, a laboratory that processes 10 or more specimens per day should select the most automated, least labor-intensive methods available, even if reagent and equipment costs are increased by doing so. Because the availability of assays to detect minimal deviations from the therapeutic range can be clinically important, sensitivity may take precedence over specificity (19). There remain circumstances in which microbiologic assays may still be useful with particular antibiotics. They have the advantage of being performed without costly or dedicated laboratory equipment. In addition, they determine the concentration of all active metabolites in one step.

There is an extremely broad array of microbiologic agar diffusion assays that are available for use. Each combination of agar, pH, and organism has been chosen to optimize the measurement of a given antibiotic, either alone or in combination with other antibiotics. For clinical purposes, however, the absolute sensitivity of the organism used is of less importance than such factors as the turnaround time of the test, the ability to store plates, and the resistance pattern of the organism. Rarely is it necessary for clinical assays to equal the sensitivity of research assays (20,21). When microbiologic assays in agar are well designed and measurements of zone sizes are made properly, these assays have shown excellent agreement with immunologic techniques (2225).

A study was undertaken in Great Britain in the mid-1970s in which salted serum specimens of gentamicin were sent to numerous clinical laboratories that performed microbiologic assays for measurement of the drug in body fluids (26). This study classified the performance of fewer than 20% of laboratories, on average, as “good.” Those laboratories with the least experience fared the worst. Radioenzyme assays and RIAs were studied according to similar protocols. Repeated testing of the same laboratories demonstrated marked improvement because of either increased motivation or reexamination of and improvement in their techniques. Pocket calculators can be programmed to provide linear regression analysis with such factors as slope, intercept, and coefficient of variation, and it is recommended that such records be maintained as an internal quality control check of a bioassay (27). Deviations beyond 2 standard deviations (SDs), or 1 SD in the same direction, an inordinate number of times should prompt reexamination of the method.

Proficiency testing samples with serum samples spiked with antibiotics such as amikacin, gentamicin, tobramycin, and vancomycin are available from the College of American Pathologists and other authorized organizations. Participation in Centers for Medicare and Medicaid Services–approved proficiency testing programs for clinically tested antibiotic analytes for which such programs are available is required under the Clinical Laboratory Improvement Amendments of 1988 and for laboratory accreditation by certifying entities, irrespective of the testing method used. For antibiotics for which no Centers for Medicare and Medicaid Services–approved proficiency testing samples are available, laboratories must document biannual verification of the accuracy of their procedures.a

Table 8.2 summarizes the general advantages and disadvantages of the agar diffusion assay method. Almost all of the equipment necessary to perform the microbiologic assay successfully is familiar to microbiology technicians and is readily available in the microbiology laboratory. The only factor that needs to be rigidly controlled is the preparation of the antibiotic standards (28). For those laboratories without sensitive electrical balances or a person experienced in preparing such standards, the hospital pharmacy can be extremely helpful and is often willing to prepare standards. Skilled technical personnel are not required to perform the assay, and it generally takes less than 2 hours to train a technician to successfully complete it. Once the standards are made and familiarity with the calculation of results and preparation of media is attained, an antibiotic blood level can be set up within 10 to 20 minutes of receipt of the specimen. With most Staphylococcus, Enterobacteriaceae, and Bacillus assays, results are available within 2 to 4 hours (2931). Finally, inexpensive equipment is used, limiting the cost of the assay and possible problems with instrument accessibility.

The major disadvantage of the microbiologic assay is the steep slope that is generated, especially for the assay of aminoglycoside antibiotics. As a consequence, a small difference in measurement can significantly alter the apparent concentration (32). Vernier calipers or automated instruments that are designed especially for this purpose, rather than millimeter rulers, should always be used to measure zone sizes. Rather than hand drawing a “best” straight line, x and y values should be interpolated from a regression curve. The regression curve can be generated with a pocket calculator. A second or third antibiotic that is present in the specimen and unknown to the laboratory can, of course, lead to erroneously high results. Because a large percentage of hospitalized patients receive two or more antibiotics, it is incumbent on the assayist to contact the ward or the pharmacy to inquire as to what antibiotics the patient is receiving, irrespective of the information that is provided on the requisition form (33,34).

It is probably convenient when choosing a microbiologic assay to choose an organism, such as Klebsiella, that is inherently resistant to many antibiotics. The technician should be sure that likely combinations of antibiotics will not act synergistically on the organism that is selected. Inordinately high results should be confirmed by repeating the assay with the specimen diluted 1:5 and 1:10 in normal human serum and by determining whether an unknown second antibiotic is present (35).

Immunoassays are the method of choice for aminocyclitol/aminoglycoside antibiotics. The major advantages of these assays include specificity, automated analysis, microprocessor-controlled calculations, and versatility of analytes that can be measured with the required instrumentation. Immunoassays for aminoglycosides and other antibiotics are invariably competitive assays, in which bound or conjugated drug competes with free drug in the specimen for antibody binding sites. Some assay formats that are commonly used in clinical laboratories to measure aminoglycoside concentrations are listed in Table 8.3. For the measurement of gentamicin and tobramycin concentrations, the Abbott AXSYM FPIA (Abbott Laboratories, Abbott Park, IL), the Dade Dimension turbidimetric inhibition immunoassay (Siemens Diagnostics, Tarrytown, NY), and the Beckman Synchron (Beckman Coulter, Inc, Brea, CA) reagent are among the most commonly used assay systems in clinical laboratories. For the analysis of amikacin concentrations, the Abbott TDX/TDX FLEX (Abbott Laboratories, Abbott Park, IL) is a very popular assay format.

Although the material costs per test are higher for immunoassays than for microbiologic assays, the labor costs are significantly less. The necessary instrumentation costs significantly less than RIA or radioenzymatic assay equipment, and, in most cases, capital expenditures for instruments can be transferred to the costs of individual tests as disposable items. Because many reagents have refrigerated lifetimes of up to 12 weeks after they are constituted, these methods are applicable to laboratories of any size. Moreover, specialized technologists are not needed to perform immunoassays. In deciding between various assay methods and formats, one often must balance the costs of labor versus the costs of supplies. For clinical purposes, the sensitivity of nonisotopic immunoassays is no different from that of RIAs (3643).

HPLC has become the method of choice for analysis of β-lactam concentrations in serum and other body fluids. Again, one must deal with the proviso that the equipment is expensive. However, one can process large numbers of specimens quite accurately in short periods of time (44,45). HPLC technology is the primary method of analysis of antibiotic concentrations in human body fluids in the research setting, especially when new drugs are being investigated. It is the method of choice when one wishes to analyze individual components of an antibiotic or its metabolites (46).

A particular advantage of HPLC is the ability to quantify closely related compounds in a mixture (4753). HPLC procedures have been developed for the analysis of almost all antibiotics used for the treatment of human diseases (5456). Unfortunately, the high cost of HPLC instruments makes it impractical for this technique to compete with immunoassays once a specific antibody has been produced for an antibiotic. In addition to the cost of the HPLC apparatus, the procedure generally requires a trained technologist and dedicated instrument to the analysis of particular antibiotics or classes of antibiotics for a defined period of time. Different antibiotic classes may require different columns and conditions. Unlike gas-liquid chromatography (GLC), however, the basic columns in HPLC technology (C8 and C18) may be used in the reverse-phase mode for all water-soluble antibiotics. The primary modifications that are required for the measurement of these drug concentrations are to the solvent systems. Because of the ability of HPLC to separate constituents in a mixture (e.g., the three gentamicin components), its analytical capabilities extend beyond those of RIAs (5760).


To achieve optimal results, specimens should be processed as soon as possible after they are obtained. Each antibiotic loses potency at its own rate. For example, aminoglycosides are much more stable than the penicillins.

Serum or ethylenediaminetetraacetic acid (EDTA)–treated plasma is the recommended specimen for measuring blood concentrations of aminoglycosides. Heparin in the concentrations present in heparin-containing blood collection tubes may inactivate aminoglycoside antibiotics through complex formation. This can cause underestimates of aminoglycoside concentrations. Heparin in therapeutic concentrations does not appear to have in vivo effects on aminoglycoside activity, but heparin-containing collection tubes should not be used unless their suitability has been verified by the manufacturer of an assay. Although sera of patients receiving gentamicin can be stored between 20°C and +25°C for up to 2 days without any significant effect, samples that are not tested within 2 hours should be stored at 0°C to 5°C to avoid possible inactivation by coadministered β-lactam antibiotics (61,62).

Acceptable samples for measurement of blood CAM concentrations include serum and EDTA or citrate anticoagulated plasma. These specimens should be kept protected from light and, if not analyzed immediately, stored frozen.

Serum or EDTA anticoagulated plasma are appropriate specimens for analysis of vancomycin concentrations. Heparin-containing samples are generally also considered acceptable. However, reports of vancomycin instability in the presence of heparin recommend exercising caution in testing such samples. Verification of specimen suitability is essential.

In general, if a specimen cannot be processed within 1 to 2 hours of its receipt, steps must be taken to ensure its potency. If it is to be processed the same day, it should be refrigerated at less than 4°C. If it cannot be processed the same day, it should be frozen. If the specimen can be processed within 3 to 4 days, freezing at −20°C should be sufficient. If storage is for a longer period, the specimen should be frozen at −70°C. If a tissue specimen cannot be processed within 2 to 3 hours, it should be frozen at −20°C for processing within 24 hours. If storage will be for more than 24 hours, the specimen should be frozen at −70°C (63).


The clinical assay of antibiotics from sites other than serum or plasma has generally not been standardized (64). The determination of antibiotic concentrations in fluids other than blood involves all the variability that accompanies the measurement of antibiotic levels in serum and much more (6568). These fluids can contain different types and amounts of proteins, chemical compounds, and cellular compounds and have different pH values as compared with serum. In order for an assay to be valid, standards must be prepared in the same milieu as the sample or in a matrix that has been demonstrated to be equivalent to it. In assaying a fluid, one must be sure that the specimen is not contaminated by blood. Although rather simplistic in practice, the avoidance of such contamination is difficult and sometimes not readily detectable (69).

No universal method is described that can be used for the assay of antibiotic concentrations in all fluids. The key to the assay of antibiotics in body fluids is that the standard antibiotic dilution curves should be prepared in the same media as the patient samples. For example, in assaying the level of an antibiotic in joint fluid, one should prepare standards in normal human joint fluid or in a solvent with a high protein content. In the determination of gentamicin concentrations in spinal fluid, it was found that if the standards were made in water, a 400% error could result. If the standards were made in 0.5% saline, the error was reduced to 50%. Finally, if the suspending medium was 150 mmol/L NaCl per 4.5 mmol/L CaCl2, the results were not significantly different from those for cerebrospinal fluid (70).

One may determine the antibiotic concentration in fluids such as cerebrospinal fluid, joint fluid, or any other nonviscous fluid by diluting the antibiotic standards in normal fluid and performing the microbiologic assay, as described in the section “Elements Influencing Microbiologic Assays,” for total biologic activity (71). When assaying an antibiotic that is not readily degraded (e.g., aminoglycosides and CAM), an immunologic or chemical assay is best.

It is important to establish the concentration of antibiotic that was actually present in the fluid under study from that obtained in the presence of contaminating blood. One should analyze the fluid specimen for blood by weighing the fluid and measuring the amount of blood inside it (72). This may be done by spectrophotometric analysis. The fluid should be centrifuged at 3,000 rpm for 15 minutes and the supernatant placed in a 1-cm spectrophotometric cell. After brief aeration, the absorbance at 576 nm is recorded and corrected for turbidity produced by cellular debris by recording the absorbances at 600 and 624 nm. At these latter wavelengths, the hemoglobin absorption is less than 10% of the maximum.

To the absorbance at 600 nm is added the difference between the absorbances at 600 and 624 nm. The resulting absorbance represents the contributions made by the turbidity to the absorbance at 576 nm. This correction is then subtracted from total absorbance at 576 nm, with the remainder providing a measure of the tissue hemoglobin content. The absorbance value so obtained can be standardized in terms of hemoglobin concentration by making similar measurements with a sample of the patient’s own blood, diluted 1:150.

Another potential problem in the assay of antibiotic concentrations in tissue is storage. When performing fluid assays, one tends to collect the fluid and freeze it until ready for use. In contrast, most clinical assays of antibiotic blood levels are performed soon after the specimen has been received.

The stability of an antibiotic depends on the medium in which it is suspended. An antibiotic that is stable in buffer for long periods of time at 20°C or at 4°C may be quite unstable at the same temperature in serum or tissue (73). One should store all body fluids at −70°C prior to assay. It is incumbent on the assayist, when storing antibiotics in a given tissue medium, to store a standard in parallel to assess any loss of activity (74). At this time, it is not possible to definitively predict how an antibiotic’s stability will be affected by the particular fluid or tissue in which it is found (73,74). Antibiotics have been assayed successfully from many body sites by employing the principles described. Each fluid and antibiotic must be treated as a unique combination. Procedures that are satisfactory for one fluid and antibiotic pair will not necessarily prove satisfactory for another pair, even if the two are closely related.

Urine should be buffered to the optimum pH of the antibiotic in question, and its protein and sugar content should be recorded. Normal pooled urine from patients who are not receiving antibiotics, buffered in the same way, should be used as controls. The Bacillus subtilis technique has been used successfully for the determination of antibiotic concentrations in urine. As described in the section “Elements Influencing Microbiologic Assays,” dilutions in a phosphate buffer provide good results. It has been reported that individuals may naturally excrete organic acids that can be antibacterial. Although this scenario is not common, the assayist should be on guard for spuriously high urine antibiotic levels due to the presence of such organic acids. One must filter-sterilize the urine if it contains microorganisms (75,76).


Microbiologic assays are relative rather than absolute (77). In one type of assay, an antibiotic concentration is determined from the microbiologic response of a strain of test organism to a series of standard antibiotic concentrations.

Agar Methods

Assays in agar fall into three broad categories: one-dimensional, two-dimensional, and three-dimensional. The one-dimensional assay employs a test tube or a capillary tube in which seeded agar has been poured and allowed to harden as the agar medium (Fig. 8.1). An antibiotic test suspension is pipetted onto the surface of the hardened agar, and zones of inhibition in one dimension are formed. The one-dimensional assay is especially appropriate for the assay of antibiotics under anaerobic conditions. This method never gained popularity in clinical laboratories in the United States, although it has been used frequently in Japan (78). The method does not lend itself to automation. Its major disadvantages are that it requires complex sample preparation and time-consuming dilution steps. The one-dimensional assay can be useful in determining the antibiotic fluid levels in a pediatric population or in other circumstances in which only a small amount of fluid is available for testing. A procedure for the one-dimensional tube assay is provided later in this chapter.

Clinical laboratories have most commonly used two-dimensional or three-dimensional assays. A two-dimensional assay is one in which the antibiotic diffuses directly against a wall of seeded bacteria. This typically involves one of two designs. In the first technique, wells are cut in agar and seeded throughout with the test organism. In the second method, the test organism is swabbed onto the surface of the agar. Antibiotic disks are then placed on the agar surface. The assay is considered two-dimensional because the concentration of antibiotic as it diffuses radially is equal at any given distance from its source. In a three-dimensional assay, the antibiotic diffuses vertically to the bottom of the Petri plate, in addition to migrating along the surface of the agar. At a distance x from the disk, therefore, the concentration of antibiotic may not be identical in all dimensions. As the agar thickness in a three-dimensional system decreases, the likelihood that the antibiotic concentrations are equal at any given distance x from the source increases because the three-dimensional system is physically moving toward a two-dimensional system.

For clinical assays, the differences between the three- and two-dimensional systems are of little significance. In research settings, both systems should be compared to establish that they yield equal results before a three-dimensional system is used. The two-dimensional assay is theoretically somewhat sounder than the three-dimensional assay (79). The three-dimensional assay is one in which cylinders, fish spines, or disks are placed on the surfaces of seeded agar plates. The well-type, two-dimensional assay and paper disk-type, three-dimensional assay have been extensively used clinically. One generally can determine the lower limits of antibiotic concentrations using well-type assays.

Elements Influencing Microbiologic Assays

In performing microbiologic assays, one must carefully account for the many conditions that affect the action of the antibiotic under study and the growth properties of the organism. Deviations from the use of rigid controls result in erroneous assay values (80). The most prominent factors that affect the performance of microbiologic assays follow.


Basic to the assay of antibiotic concentrations is a system design that determines the levels accurately. One must be certain that the zone of inhibition around a source of antibiotic is produced in direct proportion to the amount of antibiotic that is contained within that source. One must also be certain that the response is linear within the range that is normally encountered in clinical specimens. Because dose-response curves are sigmoidal in shape, the assayist must test a sufficient number of specimens to be assured that they fall on the dose-response curve. Environmental factors such as temperature and pH must not affect the assay in either the high or low ranges.


There are a wide variety of media from which to choose for the assay of antibiotic concentrations. Grove and Randall (81) in 1955 described 11 different media that were available for the assay of antibiotic concentrations in human specimens. Almost all subsequently published methods have used some variation of these 11 types of media. The choice of medium is related to the antibiotic under study, the assay design, and the test organism. Although a detailed discussion of the relative merits of each medium–antibiotic–organism combination is not within the scope of this chapter, the medium chosen should be optimized for the assayed antibiotic (79) (Table 8.4). A medium that is satisfactory for use with one antibiotic–organism combination may not be acceptable for the measurement of the same antibiotic concentration in an assay that uses a different organism (82).

The pH of the agar may vary with the mixture of ingredients and the method of preparation. Although the agars are made according to the same formulation, they may not be identical. This variability is the result of the undefined nature of many of the ingredients. For example, yeast extract and animal infusions are in no way standardized. Table 8.4 presents the effects that different agar preparations and pH values have on the results of an assay for the measurement of gentamicin concentrations. Although most researchers have not thought it necessary to use indicators in media, several have found it useful to add fermentable sugars that enable the determination of zones of inhibition by the accompanying pH changes (83). The spraying of plates with tetrazolium blue allows for the ascertainment of zone size more rapidly than by visible inspection alone (84). Agars differ in their susceptibilities to pH changes, crispness of zones, absolute sensitivities, and slopes of the standard curves that they produce (85). Because antibiotic standards and patients’ specimens are included on the same plates in most clinical assays, such variations are internally controlled (Fig. 8.2).

Antibiotic Standards

The production of antibiotic standards is the most critical factor in these assays. Because microbiologic assays are relative assays, any inconsistency in the antibiotic standards results in erroneous concentration measurements. Each antibiotic is differentially active at different pH values (78,86). Because it is impractical to make antibiotic standards from powder each time an assay is to be performed, one may produce and store working concentrations of the antibiotic in small aliquots. Practically, the antibiotics are dissolved in appropriate buffers (Table 8.5) at high concentrations. These 1-mL aliquots can be stored at −20°C but should not be kept longer than 3 months. The aminoglycosides are extremely stable and probably can be held longer than the penicillins, which are more labile (79,87,88). Subsequent dilutions from these buffers are made in appropriate body fluids for assay use.

Antibiotics must be weighed using an analytical balance. The potency of the antibiotic should be calculated based on active micrograms per milligram of powder. In addition, the identical antibiotic must be present in the standard and the sample. For example, gentamicin is a compound composed of three molecular elements. They should be present in the same proportion in the assay standards as they are in the pharmacy.

It is also important that there not be a second antibacterial substance present in the antibiotic standard. Because pharmacy materials may contain preservatives, they should not be used as standards except under emergency circumstances. It is necessary to make standards as close in composition to the patient’s specimen as possible.

For the determination of antibiotic levels in blood, the specimen should be diluted in plasma or serum (89). As discussed subsequently, this principle also applies to the determination of antibiotic levels from other tissue fluids. Considerable work has been performed to establish the types of sera that may be used for dilution. Horse serum, bovine serum, normal human serum, normal human serum inactivated at 56°C for 30 minutes, bovine serum albumin (BSA), and fetal calf serum have been the most extensively studied (61,9092). It was found that, when antibiotics were suspended in different sera than that used in the test samples, errors of 80% to 367% in measured antibiotic concentrations resulted (6). In assaying antibiotics from cerebrospinal fluid, 150 mmol/L NaCl per 4.5 mmol/L CaCl2 should be added to a phosphate buffer or inaccuracies occur (70). Apparently, it is not necessary to physically buffer the diluent serum (90). It also appears that cations present in sera at physiologic concentrations do not appreciably affect clinical assays (90).

Two factors that are present in sera may, however, exert some effect on microbiologic assays. The gentamicin recovery rate in normal serum has been reported to be between 80% and 90%. However, in uremic serum (blood urea nitrogen levels >50 mg/100 mL), the recovery rate of gentamicin has been shown to be only 50% to 69%. A more drastic decrease in the tobramycin level in the setting of uremia has been demonstrated. There has been some controversy about whether or not high levels of bilirubin in sera can interfere with the results of microbiologic assays of antibiotic concentrations. Blood bilirubin levels of greater than 23 mg/100 mL were shown to cause erroneous determinations of antibiotic concentrations (93). Bilirubin levels of 8 mg/100 mL did not affect the assay under study. Most workers have found that only assays in which bilirubin levels are above 20 mg/100 mL are adversely affected (6,78). These levels are exceedingly rare in clinical samples. Suspending the antibiotic in a matrix equivalent medium that is similar to that from which it came corrects for several factors, the most important of which is protein binding.

Physical Factors

Disks must be known to be effective for assaying antibiotics. Schleicher & Schuell BioScience, Inc (Keene, NH) 740-E disks have been used extensively. Although individual investigators have used disks of different diameters, the smallest disk that produces good zone sizes in the range of the anticipated antibiotic levels should be used. If one is using the recommended 740-E disk, 20 µL of sample should be used in the assay. The maximum volume that this disk accurately holds is 25 µL. The filter paper must lie flat on the surface of the agar or irregular zone sizes will be produced. Supersaturation can lead to surface distortions.

The well technique is approximately five to six times more sensitive than techniques that use paper disks (93). Wells can be conveniently punched in agar with a metal cylinder that is attached to a suction device (94). It is necessary, however, to allow the agar to harden for at least 15 minutes so that cracking does not occur around the wells. The wells can be filled using capillary pipettes because slight overfilling does not produce significant errors in the results obtained (94). Wells should be filled while a low-watt light bulb is maintained at an angle of approximately 30 degrees so that the wells in an assay plate contain uniform amounts of samples or standards. Because the agar depth affects the sensitivity of the test, it is best to add as little agar as possible to the container when a plate is poured.

Choice of Organism

One can use a microbiologic assay to measure the concentration of almost any drug for which a sufficiently susceptible test organism can be isolated (84). It is quite simple to choose an organism for the assay of a sample in which only one antibiotic is present. However, one must exercise considerable caution in organism selection for an assay in which the specimen contains antibiotics in addition to the test drug.

The usual method is to use, as the test organism, a microorganism that is very sensitive to one of the antibiotics and insensitive to the other (81). This dictum appears simple but is often not reliable in practice (61). One must take into account synergy or antagonism, even though the test organism may appear to be resistant to one of the antibiotics. Thresholds at which one antibiotic interferes with another are published (95). The optimum way to assay antibiotics in a mixture is to chemically separate them by electrophoresis or chromatography prior to their measurement (96). This is probably not practical in clinical laboratories. Many test strains have been isolated that adequately take into account the physiologic levels of antibiotic combinations found in humans.

Organisms used for the clinical measurement of antibiotic blood levels most commonly include Bacillus (84,97100), Staphylococcus aureus (82,93,101,102), Sarcina lutea (103,104), Streptococcus(102,105), Clostridium(99,106,107), KlebsiellaProvidencia (108), and others (Fig. 8.3). Fungi and bacteria have been used for the assay of chemotherapeutic drugs (109,110). Assays that use bioluminescent bacteria (111,112) have also been employed. The actual choice of organism depends on the antibiotic to be measured, its susceptibility pattern, its ability to produce clear and crisp zones on the given agar medium, and its ability to provide results within 4 hours. The organism that is selected impacts upon the minimum level of antibiotic that one can detect. Table 8.6 demonstrates the lower levels of detectability for some common antibiotics with assays that use Kirby-Bauer control strains Staphylococcus aureus, American Type Culture Collection (ATCC) strain 25923, and Escherichia coli ATCC strain 25922 as test organisms.

For the rapid determination of antibiotic levels, vegetative organisms must not be more than 24 hours old. The organism can be inoculated the night before the test is run or, more simply, swabbed off a plate, including a Kirby-Bauer sensitivity plate, that is no more than 1 day old (3). An organism with a particularly unusual sensitivity pattern may be seen infrequently. A 4- to 6-hour growth of bacteria can be diluted 1:1 in either fetal calf serum or 7% BSA and stored at −20°C to −70°C for up to 3 weeks. These frozen cultures can be thawed and used directly for rapid antibiotic assays (113,114). Although varying the size of the inoculum changes the sensitivity of the test, this also affects the test time, with a large inoculum reducing both test time and sensitivity.

It is not possible in this chapter to detail all conditions for the assay of the large number of antibiotics that are used in humans. Table 8.7 provides references for assays that are based on the previously mentioned microbiologic principles for some commonly used antibiotics. Table 8.7 also presents the conditions that are required for the determination of concentrations by diffusion assay for some of the major classes of antimicrobials. The choices of test organism, test agar, and buffer diluent shown are optimal for the class of antibiotic. One needs only substitute the required test organism for Klebsiella, use the proper agar and diluent described in Table 8.7, and choose the proper dilution series for the antibiotic standard, as described in Table 8.8.

As with other microbiologic assays, when distinct zone sizes appear around the wells or disks, they may be measured. The concentration of antibiotic is calculated in the same way as described for the B. subtilis assay of aminoglycoside concentrations. If one substitutes another test organism, buffer diluent, or test agar for those described here, this would not necessarily invalidate the assay but would likely decrease its sensitivity. As with all assays, one must obtain a straight line for the standards when plotting zone diameters versus logarithms of the antibiotic concentrations. The B. subtilis assay described subsequently can determine the clinically applicable levels of almost all medically used antibiotics (115).

The basic approach used in bioassays is exemplified by the method of Lund et al. (116). This method is typical of those that use organisms that are resistant to all but specific antibiotics for the rapid assay of antibiotic concentrations in clinical material (4,114,117). Although any organism that possesses the appropriate sensitivity and meets the criteria described here can be used, the method of Lund et al. (116) provides a good model. The multiresistant Klebsiella strain they described is available from the ATCC. The method has been successfully field-tested. Prior to the substitution of another organism in this method, users should ensure that the new organism meets the criteria previously described. This strain of Klebsiella is resistant to most commonly used antibiotics, except gentamicin, tobramycin, and amikacin.

Presence of Aminoglycosides

Because the aminoglycoside antibiotics have broad activity, the assay of other classes of antibiotics that are present in combination with them has proved difficult. There are few bacteria that are resistant to aminoglycosides but sensitive to other classes of antibiotics. A group B Streptococcus has been used that allows the determination of clindamycin (CLD) concentrations in the presence of aminoglycosides (118). Care should be exerted when using any streptococcus if a patient is receiving penicillin-type antibiotics because of possible synergy between penicillin and aminoglycosides with this genus. Because aminoglycoside antibiotics are not active under anaerobic conditions, it is possible to determine the level of CLD in the presence of gentamicin using Clostridium perfringens (99). The addition of calcium and other divalent cations to the medium has proved effective (119). However, high levels of calcium may affect bacterial growth and render the medium somewhat turbid. Based on the same principle, cellulose phosphate powder can be incubated with the serum specimen to remove gentamicin (120).

Aminoglycoside antibiotics are inactivated by the polyanionic detergent sodium polyanetholesulfonate in a stoichiometric precipitation reaction (121). The addition of this material to media allows for the rapid determination of the levels of all other classes of antibiotics in the presence of all aminoglycosides (122). Using either nutrient agar or plate count agar, sterile 5% sodium polyanetholesulfonate solution (Grobax; Hoffmann-LaRoche, Nutley, NJ) is added to establish a final concentration of 0.8%. Using the Kirby-Bauer S. aureus ATCC strain 25923, or E. coli ATCC strain 25922, the standard well-type bioassay is run. For penicillin, cephalosporins, erythromycin, tetracycline, CAM, and vancomycin, S. aureus is the better choice. For antibiotics that are more active against gram-negative bacteria, such as ampicillin or the polymyxins, E. coli is preferred. The procedure described for the Klebsiella assay can be followed with either of these strains.

One-Dimensional Assays

As previously described, one-dimensional assays can be useful for the determination of antibiotic concentrations under anaerobic conditions and for assays for which only small amounts of specimen are available.


Nutrient agar is diluted with an equal amount of 1% peptone in water and brought to pH 7.8 (the pH varies depending on the type of antibiotic to be assayed). It is dispensed for storage in 19-mL amounts.


The bacterial test strain (S. aureus 658P) is grown overnight in trypticase soy broth. It is diluted 1:100 and mixed well by shaking to break up clumps. Before use, the agar is melted and cooled to 48°C in a temperature-controlled water bath. Then 1 mL of the bacterial suspension is added to 19 mL of the test agar. The bacteria and the test agar are mixed well. The test agar is pipetted into conical test tubes that have an internal diameter of approximately 3 mm (K tubes used for determination of complement fixation in the Kolmer test in syphilis serology are satisfactory). The agar is added to a depth of 3 mm from the bottom of the tube. The agar is allowed to harden (at least 5 minutes). Standards for many commonly used antibiotics are described in Table 8.5. Each standard is pipetted into a different tube. Each standard should be repeated three times. The patient’s serum is treated identically. The antibiotic standards and patient’s serum should be added until they are 1.5 mm above the agar layers. Small deviations from this level do not affect the accuracy of the method. The tubes are incubated at 37°C overnight.


Typical assay results are presented in Figure 8.1. To determine inhibition of growth, each tube is laid on its side. Using an eyepiece with the ability to measure distances, the distance in millimeters from the point where the patient’s specimen and the test agar meet (the meniscus of the test agar) to the point where growth of the test strain begins (often it is the place where large colonies are seen) is recorded. This measurement is repeated for each standard and the patient’s specimen. The results are calculated as for any dose-response curve. On the ordinate (y axis), the logarithm of the antibiotic concentration is plotted. The depth of inhibition is plotted on the abscissa (x axis). The concentration of the patient’s specimen is found by drawing a vertical line from the depth of inhibition on the x axis to the standard line and drawing a horizontal line from the point of intersection to the y axis, where the concentration of the antibiotic is read directly.

Amoxicillin and Clavulanic Acid

β-Lactamase inhibitors protect β-lactam antibiotics from destruction by these enzymes. Clavulanic acid, sulbactam, and tazobactam are those currently used. Because β-lactamase inhibitors are combined with β-lactam antibiotics in the same pharmaceutical preparations, a procedure for the assay of amoxicillin and clavulanic acids is presented (123).

Augmentin consists of amoxicillin trihydrate and the potassium salt of clavulanic acid as the anhydrous free acids in the ratio of two parts amoxicillin to one part clavulanic acid. Amoxicillin may be assayed in the presence of clavulanic acid. A conventional microbiologic assay technique with S. lutea as the assay organism can be used to measure amoxicillin concentrations in body fluids after the administration of Augmentin. This is not the case with clavulanic acid because its very low level of antibacterial activity precludes the use of a microbiologic assay technique that depends on measurement of antibacterial activity. Instead, clavulanic acid concentrations in body fluids can be assayed by a microbiologic agar diffusion method involving measurement of the inhibition of β-lactamase activity of a strain of Enterobacter aerogenes. In the method subsequently described, clavulanic acid does not interfere with the microbiologic assay of amoxicillin in clinical specimens, and amoxicillin does not influence the measurement of clavulanic acid.

This method uses a large-plate microbiologic assay technique in which samples are added to wells punched in agar. Modifications may be made as required; for example, Petri dishes may be used instead of large plates, and the specimens may be applied to the plates by assay cylinders, paper disks’ fish spines, beads, and others. Moreover, the bacteriologic media used are not critical and individual laboratories may find their own modifications to be more suitable for their purposes than those described.

In principle, specimens should be assayed as soon as possible after collection and should not be stored for more than a few days before assays are performed. As a rule, β-lactam compounds are relatively unstable in aqueous solutions and in body fluids, and clavulanic acid is no exception. Consequently, care must be taken in the handling and storage of clinical specimens that contain amoxicillin and clavulanic acid. Serum specimens may be stored for up to 2 days in the refrigerator (4°C) or for up to 3 days in a freezer (−20°C). Clavulanic acid is relatively unstable in undiluted urine, particularly at alkaline pH, and is less stable (in urine) at −20°C than at 4°C. Accordingly, urine specimens for assay should be diluted 10-fold in citrate buffer, pH 6.5, as soon as possible after collection and stored at 4°C. Under these conditions, specimens of urine containing amoxicillin and clavulanic acid may be kept for up to 5 days at 4°C before assay.

The stability of clavulanic acid is influenced by the concentration, pH, and composition of the buffer solution that is used as a diluent. Solutions of clavulanic acid are most stable at pH 6.0 to 7.0, and preparations in citrate buffers or distilled water are more stable than those in phosphate buffers. Also, aqueous solutions that contain amoxicillin and clavulanic acid are more stable at 4°C than at −20°C. Consequently, aqueous solutions of Augmentin, clavulanic acid, or amoxicillin for microbiologic tests should be prepared in 0.1 mol/L of a citrate buffer, pH 6.5. Solutions that are prepared in this medium and contain up to 0.1 mol/L of amoxicillin or clavulanic acid may be stored at 4°C for up to 4 weeks.


Apparatus.The following equipment are required:

 1.  Large glass assay plates

 2.  Pasteur pipettes (nominally 30 drops/mL) or standard dropping pipettes (0.02 mL per drop)

 3.  Punches for cutting holes (7- to 8-mm diameter)

 4.  Lancets or broad needles

 5.  Test tubes, flasks, and pipettes as required

 6.  Leveling tripods or a level surface

 7.  Water bath at 50°C

 8.  Incubator at 30°C or 37°C

 9.  Needle-point calipers

Antibiotic medium 2 (81) is obtainable commercially in dehydrated form from Chesapeake Biological Laboratories (Baltimore, MD), Oxoid, Inc (Ogdensburg, NY), and Difco Laboratories (Sparks, MD). It is prepared by dissolving beef extract (1.5 g), yeast extract (3.0 g), peptone (6.0 g), and agar (15.0 g) in 1,000 mL of distilled water and adjusting the pH to 6.5 or 6.6. Volumes of 300 mL are distributed in screw-capped bottles and sterilized at 15 psi for 15 minutes.

The required amount of medium is melted and maintained at 50°C in a water bath until the plates are ready to be poured. The assay plates are placed on a level surface or on leveling tripods and adjusted until they are level. The plates are sterilized by swabbing them with alcohol, followed by flaming. The swabbing and flaming procedure is performed twice. The inoculum is added to each bottle of agar and mixed thoroughly. The surface of the plate is flamed and the inoculated medium is poured evenly over the plate. The surface of the agar is again flamed to eliminate air bubbles, and the agar is allowed to solidify with the lid slightly open. The plates may be kept in a refrigerator at 4°C for up to 24 hours.

Buffer.Sorensen’s buffer (0.1 mol/L citrate buffer, pH 6.5) is used. Solution A is 0.1 mol/L disodium citrate (21.0 g of citrate acid in water, dissolved in 200 mL of 1 N NaOH [4.0 g/1,000 mL] and diluted to 1,000 mL). Solution B is 0.1 N NaOH (4.0 g/1,000 mL). Fifty-four milliliters of solution A is added to 46 mL of solution B. The pH of final solution is checked and, if necessary, adjusted by adding the required amounts of solution A or B.

Standard Solutions.Normal pooled human serum is sterilized by filtration and tested for the absence of antibacterial activity. The quantity of laboratory reference standard amoxicillin trihydrate that is equivalent to the required amount of amoxicillin pure, free acid is accurately weighed and dissolved in 0.1 mol/L citrate buffer, pH 6.5. Dilutions are made in pooled human serum for the assay of serum specimens or in 0.1 mol/L of a citrate buffer, pH 6.5, for the assay of other specimens or urine. Serial dilutions are prepared in the requisite diluent to give the necessary range of standard solutions. Each specimen is diluted to give a concentration that is estimated to fall within the range of the standard line.

Serum specimens are diluted in pooled human serum. Specimens of urine are diluted in 0.1 mol/L citrate buffer, pH 6.5. The required dilution depends on the dose of the drug that was administered and on the time at which the specimen was taken. The mean serum concentrations of amoxicillin and clavulanic acid in fasting volunteer subjects 1 hour after the administration of single oral 375 mg doses of Augmentin are 5.6 µg/mL for amoxicillin and 3.7 µg/mL for clavulanic acid. The mean urine concentration of clavulanic acid is 545 µg/mL.

Bacteria.A suspension of S. lutea (National Collection of Type Cultures strain 8340, ATCC strain 9341) is prepared by using a loop to inoculate 100 mL of nutrient broth in a 500 mL Erlenmeyer flask with organisms from a stock culture that has been grown on a nutrient agar slant and stored in the refrigerator at 4°C. The flask is incubated for 48 hours at 37°C and stored at 4°C. Stock cultures on agar slants may be kept for up to 4 months at 4°C. Broth suspensions may be held for 4 or 5 weeks at 4°C. An inoculum of 0.8 mL of the broth suspension into 100 mL of agar usually provides satisfactory growth on the large assay plates.

Procedure.Seeded plates are taken from the refrigerator and the agar surface is blotted dry with filter paper. Holes 8 mm in diameter are cut with a punch. The agar plugs are removed with a lancet or broad needle. The holes are filled with the specimens and standard solutions using a Pasteur or dropping pipette. The pipette is rinsed three times in buffer solution between each sample loading. The plates are incubated overnight at 30°C or 37°C.

Interpretation.The diameters of the inhibition zones are measured and the standard and sample responses are averaged. The mean inhibition zone diameters of the standard solutions are plotted against the logarithm of the antibiotic concentrations using semilogarithmic paper. The best fitting straight line connecting the points is constructed. The concentration of each dilution of specimen is determined by extrapolation from the straight line.

Clavulanic Acid

Clavulanic acid is a weak antibacterial agent. However, it is a potent inhibitor of certain β-lactamases. This latter property is the basis of a microbiologic assay for clavulanic acid in clinical specimens. In brief, a subinhibitory concentration (60 µg/mL) of benzylpenicillin is added to a nutrient agar that is inoculated with the β-lactamase–producing organism E. aerogenes BRL strain 1. Plates are poured and wells are cut in the agar in the usual fashion. At the concentrations tested, clavulanic acid has no inhibitory effect on the growth of the assay organism, but it does inhibit the β-lactamase activity of the enterobacterium, thereby preventing destruction of the benzylpenicillin incorporated in the agar. As a consequence, inhibition zones are produced by the penicillin, the diameters of which are proportional to the concentration of clavulanic acid in the test sample.

The assay for the determination of clavulanic acid is identical to that for amoxicillin, with the following exceptions. The assay medium is adjusted to pH 7.4. The amount of laboratory standard material of potassium clavulanate that is equivalent to the required amount of the pure, free clavulanic acid is accurately weighed and dissolved in 0.1 mol/L of a citrate buffer, pH 6.5. Nutrient broth inoculated with a wire loop from a nutrient agar slant of E. aerogenes BRL strain 1 (or Klebsiella pneumoniae ATCC strain 29665) and incubated overnight at 37°C is used to inoculate the assay agar. A fresh culture should be used for each assay. An inoculum of 3.0 mL of overnight broth culture in 100 mL of agar produces satisfactory growth on large assay plates.

The inoculum is added to the agar and benzylpenicillin is added to the inoculated agar to give a final concentration of 6 µg of benzylpenicillin per milliliter of agar. The agar is poured into the plates, as described for the assay of amoxicillin. When the agar has set, the plates are stored in the refrigerator at 4°C and are used as soon as possible on the same day. This is essential because the assay organism is able to inactivate the penicillin incorporated in agar plates if they are allowed to stand at room temperature or overnight at 4°C. The results are interpreted as for the amoxicillin assay as previously described.

Three-Dimensional Assays

Antibiotic medium 11 (Difco Laboratories, Detroit, MI) is adjusted to pH 7.9. This agar may be stored in 25-mL aliquots in the refrigerator for up to 1 month. Prior to performing the assay, an appropriate number of tubes are melted and brought to 50°C; 0.4 mL of an overnight growth of Klebsiella in trypticase soy broth is added. The suspension is thoroughly mixed and 9 mL is poured into two 100 × 15-mm plastic Petri plates. Alternatively, 0.4 mL of a heavily inoculated 5-hour broth culture can be added to the agar.


For each assay, 0.02 mL of the patient’s serum is pipetted with sterile disposable capillary pipettes onto Schleicher & Schuell 740-E disks (Schleicher & Schuell BioScience, Inc, Keene, NH). Alternatively, wells can be cut in the agar. The method described by Sabath et al. (6,93) is followed using the same number of standards and samples of the patient’s serum.

Calculation of Results

Generally for a given antibiotic, results are obtained from a plot of the logarithm of the antibiotic concentration versus the zone diameter. This is quite adequate for calculations involving limited ranges of antibiotic concentrations. However, if the concentration range is greater than fourfold, it is better to plot the logarithm of the antibiotic concentration versus the square of the zone diameter. Lines that visually appear straight on plots of the logarithm of the antibiotic concentration versus zone diameter have frequently been shown by computer analysis to have low coefficients of variation (97). This caveat is particularly important if the slope of a line is steep. When using logarithmic paper to calculate antibiotic concentrations in this way, very small differences in values can produce large changes in the apparent concentrations, especially in the range of 10 to 100 µg/mL. It is best to use an assay system that provides the flattest possible line. Pocket calculators or handheld computers can provide “best fit” straight lines, coefficients of variation, SDs, and direct interpolations. Keeping permanent records of these calculations provides a strong internal quality control. One may, for example, detect the decay of antibiotic standards by changes in line slopes or differences in coefficients of variation.

When used to perform clinical assays, the microbiologic method produces results in 4 hours or less. Figures 8.4 and 8.5 illustrate that longer incubations affect the zone size only slightly. The size of the zone is, in effect, established approximately 2 or 3 hours after incubation. Small, uniform increases in zone size do not affect the slope of the line or the final calculation of antibiotic concentration.

Electrophoretic Separation of Antibiotics

Methods have been developed to separate antibiotics in a mixture by gel electrophoresis and to assay these separated compounds by covering the gel with an indicator bacterium in agar. After incubation, zones of inhibition are seen in the covering agar slab. These methods were especially useful before HPLC became available and may still have some applicability if one needs to assay for disparate classes of antibiotics in a mixture (124,125).

Broth Methods

Turbidimetric Assays

For completeness, the basic parameters of the assay of antibiotics by turbidimetric means are mentioned. These techniques have largely been relegated to the assay of antibiotic-containing materials in situations in which one wants to screen large numbers of samples in a short period of time. Turbidity is used here to mean any technique in which the growth of bacteria in a liquid medium is used to quantify the amount of antibiotic in a solution. Included under this heading is a consideration of techniques in which the change in the number of bacteria is directly measured photometrically (as numbers or mass of organisms) and, as an extension, those in which a product such as a change in pH or the release of CO2 is measured.

An examination of the theory of optical methods was explored by Kavanaugh (79). Only the pertinent factors involved in the assay of antibiotics by turbidimetric means are discussed here. The assay of antibiotics by photometry depends on the direct relationship between growth of bacteria in culture and the amount of antibiotic present. Generally, this relationship is displayed in a plot of growth per unit time versus percent transmission or the optical density of the bacterial suspension. It should be noted that absorbance measures mass or volume rather than the concentration of bacteria (79). This relationship can be stated as a form of Beer’s Law or Beer-Lambert Law in which Io is the amount of light leaving a suspension, I is the amount of light entering a suspension, OD is the optical density, N is the mass of bacteria, E is the extinction coefficient of the particles, a is the optically effective area of the particle, and b is the thickness of the suspension. Early work on the assay of antibiotics utilized modifications of this equation from commercial fermentation processes for the assay of antibiotics. Several investigators have attempted to modify it for use in clinical assays (105).

log(Io/I) = OD = (N)[Eab/2.3]

Irrespective of the instrument or technique used, procedures must be employed to straighten the generally curved calibration lines for turbidimetric microbiologic assays. Although it is beyond the scope of this chapter to treat the subject fully, the following relationships should be useful for the applied assay. A theoretical dose-response line for an antibiotic that decreases the growth rate of the test organism is

Nt = No exp(ko + f(v)kM – kaC)t,

where Nt is the concentration of the test inoculum after an incubation period of t, No is the concentration of the test inoculum at the beginning of the assay, ko is the generation rate constant in the absence of antibiotic, C is the concentration of antibiotic with an inhibitory coefficient of ka, kM is the effect of the medium, and the function f(v) is a function of the volume of the sample added to the assay. This is a theoretical expression because the major characteristics of the equation, although interrelated, are not absolutely known.

In an individual assay, No, ko, f(v), kM, ka, and t (time) are constant. Therefore, the variables of the equation may be related as follows:

log N = G + BC,

where G and B are constants that are intrinsic to the assay. Because most spectrophotometers use absorbance (A), this equation may be translated into

log A = E + FC.

To straighten the line, this equation may be modified as

log (A + M) = O + PC.

The algebraic sign M is used to straighten the line over the particular concentration range in question. The constant M compensates for two sources of curvature: the nonlinear relationship between log N and C, and the inherent nonlinear responsiveness of photometers.

Because of this complexity, pure photometry has proven successful only in situations in which all conditions except the quantity of a single antibiotic in solution could be rigidly controlled. The Abbott Laboratories MS-2 instrument (Abbott Park, IL), a device that continually monitors bacterial growth in optical density units, has been used to determine antibiotic blood levels under clinical conditions using a growth curve analysis (126). The principle of this technique is that, for any dose-response curve, the response is the turbidity of bacteria in solution. The MS-2 instrument determines turbidity using red light–emitting diodes. The test culture is inoculated into the cuvette. After the culture reaches logarithmic growth phase, the culture is pulled down into compartments, each of which contains a disk. The disks can contain either a patient’s serum or a standard. Generally, the patient’s serum is tested in two compartments and each standard is repeated twice. After approximately 3 hours of incubation, the optical density (after minor corrections that are required because of the physical nature of the instrument) is plotted on the y axis and the logarithm of the antibiotic concentration is plotted on the x axis. The instrument has a built-in computer that enables it to continuously monitor the growth of the test cultures. All calculations are automated. In the early 1980s, we used this instrument successfully with S. aureus as a test organism to determine the concentration of aminoglycosides in blood (126).

Broth Dilution Bioassay for Polymyxins

Broth dilution bioassay is rarely performed. However, this technique may prove useful for the measurement of polymyxin-class antibiotics. It was found that in normal agar, but not agarose, assays of colistin concentration yielded false-low blood and urine values (127). This was probably occurring because a highly diffusible precursor was being converted into a poorly diffusible active compound. The turbidimetric technique more accurately reflected what was actually occurring in the body than did agar techniques. Levels of the polymyxin group of antibiotics can be determined by a modification of the standard turbidimetric technique in which the number of viable organisms is counted.

Procedure.A strain of Pseudomonas aeruginosa that is sensitive to the polymyxins is employed as the test organism. The inoculum that is used for each antibiotic concentration determination is 0.05 mL of a 1- to 8-hour culture (grown in trypticase soy broth) that has been diluted at a 1:10,000 ratio in normal human serum. The patient’s specimen is diluted in the same fluid that is to be assayed (serum specimens are diluted in serum, cerebrospinal fluid specimens are diluted in cerebrospinal fluid, and so on) to yield final percentage concentrations of 90%, 80%, 70%, 60%, 50%, 40%, 30%, 20%, 10%, and 0%. The final volume of each of these dilutions is 0.5 mL after the addition of 0.05 mL of the inoculum. Standards should be incubated with normal human serum for 18 hours at 37°C. This preincubation is necessary for activation of the polymyxin-type drugs.

Immediately after the addition of the inoculum, all tubes are incubated at 37°C for 30 minutes. After incubation, the tubes are plunged into an ice bath (0°C). The number of bacteria in each tube is determined by performing serial 10-fold dilutions with distilled water and plating 0.1 mL of each dilution on the surface of trypticase soy agar plates. The total number of viable bacteria per milliliter of inoculum is calculated from these colony counts after 18 hours of incubation at 37°C.

The end point for each titration of the polymyxins in each patient’s serum (or other fluid) is the smallest amount of patient’s serum that results in a reduction of inoculum to 10% of the original viable count. To increase the precision of the end points, the concentration of the subject’s serum or the concentration of known antibiotic that causes a reduction of 90% of the viable organisms is read from a curve that is drawn on semilogarithmic paper by plotting the number of organisms on the logarithmic axis versus the concentration on the arithmetic axis. Performing the assay with normal human serum alone accounts for any reduction in bacteria that is caused by nonspecific factors.

Interpretation.The antibacterial activity of the polymyxin standards is compared with the antibacterial activity of the patient’s specimen. The calculations are as set forth in the “Turbidimetric Assays” section. The titer of the patient’s serum is multiplied by the minimum inhibitory concentration of the organism for the individual polymyxin to yield the amount of polymyxin-type antibiotic in the patient’s serum.

pH Change Assays

Antibiotics act on bacteria by halting their growth and/or metabolism. Antibiotic-induced changes bacterial metabolic activity can be used to determine antibiotic concentrations in body fluids and tissues. The methods used are based on the principle that the larger the quantity of antibiotic that is in contact with a bacterial population, the greater will be the modification of the population’s metabolic pathways. As with all dose-response assays, one must obtain a standard dose-response curve and calculate from only those points that fit the equation for a straight line.

The most popular method of this type employs a change in the pH of the growth medium as a measure of the amount of antibiotic that is present. If no antibiotic is present, the test strain grows and produces a change in the pH of the growth medium by metabolizing a constituent (e.g., a sugar, which would lower the pH, or urea, which would increase the pH). If an antibiotic is added to the system and the test organism is affected by it, the change in pH is lessened. The urease-based bioassay is representative of this group of assays and can be used by laboratories without special equipment.

Adenosine Triphosphate Measurement Assays

A potentially automatable assay that is a combination of a bioluminescence procedure and a chemical assay uses rates of endogenous adenosine triphosphate (ATP) production by Klebsiella edwardsii to determine antibiotic concentrations. The amount of ATP that is present in a broth reflects bacterial growth, which is inversely related to the amount of antibiotic present. Standards are inoculated with the test strain in parallel with the patient’s serum. After 2 hours of incubation at 37°C, the tubes are extracted with an ethylenediaminetetraacetate/H2SO4 solution. The amount of bacterial ATP that is released is determined by spectrophotometry (128). A plot of the antibiotic concentration versus the relative amount of ATP released is used to calculate the antibiotic blood level. One should heat the serum prior to performing the assay in order to destroy human adenosine triphosphatase. Instruments that enumerate bacteria in urine and other body sites based on this principle are available from a number of vendors, including Celsis International (Newmarket, Suffolk, United Kingdom), Coral Biotechnology (San Diego, CA), and New Horizons Diagnostics Corporation (Columbia, MD).


Methods that involve the use of luminescent bacteria have been proposed to determine the activity of antibiotics in serum (111,112). One type of assay called the induced test is based on the ability of some antibiotics to inhibit luciferase synthesis by luminescent bacteria (129). Ulitzur’s (129) method, described in the following section, is sensitive, rapid, and potentially automatable.

Procedure.The serum is heated at 56°C for 30 minutes to eliminate bacterial activity. To 0.8 mL of serum, 0.2 mL of 10% NaCl containing 0.1 mol/L 3-(N-morpholino) propanesulfonic acid buffer and 1.5% glycerol is added. The final pH value of the mixture should be 7.9 for all antibiotics except CAM and tetracycline, which are tested at pH 6. A Photobacterium leiognathi 8SD18 cell suspension (200 µL, 3 × 108 cells/mL) is added to 0.8 mL of the serum, as well as to 1 mL of antibiotic-free pooled serum. After 10 minutes of preincubation at 30°C, proflavin is added to give a final concentration of 1.5 µg/mL for pH 7.9 or 25 µg/mL for pH 6.0. The vials are then incubated with gentle shaking at 30°C. Proflavin is a DNA-intercalating agent that induces the luminescence system of dark mutant luminescent bacteria. The results are recorded after 40 to 60 minutes of incubation. Luminescence is measured with a photometer/photomultiplier and is universely proportional to the concentration of the antibiotic in the specimen.

The bioluminescence test specifically determines the activity of the tested antibiotic as a de novo protein synthesis inhibitor. Antibiotics that act on DNA or cell wall synthesis are not detected by this test. The bioluminescence test is more sensitive than most available bioassays. This high sensitivity may be attributable to the greater susceptibility of newly synthesized proteins to the inhibitory actions of many antibiotics.

Serum Inhibitory Concentration and Serum Bactericidal Concentration

Broth dilution tests that employ patient serum as the antimicrobial milieu date to the work of Schlichter and McLean (130), who reported on the serum inhibitory concentrations (SIC) in 10 patients with streptococcal endocarditis. They obtained serum from these 10 patients, geometrically diluted each sample in broth, and inoculated each dilution with the patient’s own organism (131). They obtained a titer of each patient’s serum that reflected the serum’s ability to inhibit macroscopic growth of the microbe. Fisher (132) subsequently extended this procedure by subculturing each tube that failed to exhibit macroscopic evidence of growth after overnight incubation. The highest dilution of a patient’s serum, or titer, that was able to kill the microbial inoculum was called the serum bactericidal concentration(SBC). SIC/SBC tests have since been applied to specimens from patients receiving therapy for intravascular and other closed-space infections (133).

Procedure.The patient’s infecting microbe is adjusted to yield between 105 and 106 colony-forming units (CFU)/mL in each tube (134). It is important that the actual CFU per milliliter value be determined. To do this, 0.1 mL each of a 1:100 and a 1:1,000 dilution of the inoculum is spread over the surface of the appropriate agar medium. After overnight incubation, the inoculum size is calculated from the number of colonies that are present on the plate that has between 20 and 200 colonies on it. For example, if there are 40 colonies on the plate that was inoculated with a 1:1,000 dilution, the original inoculum contained 4 × 105 CFU/mL.

The serum is diluted geometrically and the test is performed in a manner that is analogous to the minimum inhibitory concentration/minimum bactericidal concentration procedures. For most organisms, the broth can be Mueller-Hinton broth, with brain-heart or Levinthal medium used for fastidious microbes. To each of 12 tubes, 1 mL of broth is added. To the first tube is added 1 mL of serum, followed by thorough mixing. One milliliter from tube 1 is added to tube 2 and is thoroughly mixed. One milliliter from tube 2 is added to tube 3, and so forth. This process is continued through tube 12. However, 2 mL of broth is added to tube 12, rather than 1 mL as in the other tubes. One milliliter is removed from tube 12 and is transferred to a sterile test tube as a sterility control. A geometric dilution of the patient’s serum has, therefore, been made from 21 to 212. One should also have a 14th tube that contains only the patient’s undiluted serum. To each tube is added 1 mL of the patient’s own microbe, diluted to yield a final concentration of 105 or 106 CFU/mL. All transfers should be made with sterile pipettes. All tubes are incubated in ambient air at 35°C for 18 to 24 hours. The lowest concentration (dilution) of the patient’s serum that completely inhibits visible growth is the SIC. Concentration or dilution is converted to titer by the formula: titer = 1 per dilution. As the titer increases, the amount of serum in the tube decreases.

The SBC is determined by spreading 0.1 mL from each tube that does not show turbidity and the first tube that does show turbidity over the entire surface of a 100-mm diameter agar plate that contains the appropriate growth medium. A different pipette should be used for each transfer from a tube. The plates are incubated for 24 hours and preliminary colony counts are performed and recorded. The plates should be incubated for 48 hours when examining staphylococci and for up to 72 hours when examining other microbes. This extended incubation allows the organism to grow on the surface of agar and obviates any effects that transferred antibiotic may exert. The SBC is the lowest concentration (i.e., highest titer) of serum that produces 99.9% killing. For example, if the inoculum in each tube was 2.0 × 105CFU/mL, killing would be defined as an agar plate that demonstrates no more than 20 colonies from a 0.1 mL subculture.

Interpretation.Much controversy surrounds the use of SIC/SBC tests (135,136). Importantly, few groups have performed the procedure in the same manner. Variations in protocols have involved inoculum size, type of broth, bactericidal end point, time of incubation, timing of the blood sample, volume of the broth sample, and the definition of the bactericidal end point. For these reasons, this test is no longer performed at Yale.


Although considerable effort has gone into the determination of antibiotic concentrations in body fluids by direct chemical analysis, there are few instances in the clinical laboratory when one can use these methods. Because these assays are based on reactions that involve specific chemical groups on antibiotic molecules, the concentrations of antibiotics determined by many chemical assays include active drugs as well as metabolic breakdown products that retain the reacting moiety. Consequently, a chemical assay is most satisfactory in a pharmaceutical fermentation process in which one is dealing with a pure substance.

In vivo, from 0% to more than 95% of an administered drug may not exist in its native state. Because a clinical assay is intended to determine the amount of active material circulating, chemical assays may yield false-high values. In addition, for many antibiotics, all their breakdown products may not be known. When chemically assaying the amount of active drug from biologic sources for such antibiotics, it is dangerous to use estimates of average metabolically active fractions. In addition, secondary substances (either other chemotherapeutic agents or normal body constituents) may interfere with these assays. Because it is very difficult to exclude all possible agents in a given assay, falsely elevated values can unpredictably result.

Most chemical assays require extraction of the antibiotic prior to analysis. This extraction leads both to an inordinate number of technical procedures and to a greater possibility of error. Also, extracted material often requires specialized environmental safety conditions. Although these obstacles are not insurmountable in industry, they may be unwieldy for clinical laboratories.

The advantages of chemical assays for clinical laboratories are, at present, more theoretical than practical. Chemical assays can potentially be automated and should provide rapid turnaround times. Moreover, chemical analysis yields an absolute quantity, as opposed to a relative response, as in the microbiologic assay. This makes the standardization and implementation of controls easier. Only those assays that are useful in the routine and/or clinical research laboratory are discussed in the following paragraphs.

Colorimetric Assays


The thiobarbituric assay of streptomycin has been modified to allow it to determine clinically important levels of many deoxy sugars (137). Although the chromatogen that is produced when streptomycin and thiobarbituric acid react is quite stable, biologic homogenates that contain glycoproteins, sugars, and plasma proteins have been found to interfere with the assay. Changing the temperature of incubation from 100°C to 37°C eliminates all but plasma protein interference. Automated dialysis and computerized result analysis can potentially be used to design automated assays based on the method.

A crude solution of the enzyme is used in the reaction. The acetylated product, rather than being adsorbed to phosphocellulose paper, is not used at all. Thiol coenzyme A that is produced in the reaction is allowed to react with 5,5′-dithiobis(2-nitrobenzoic acid). The product of this reaction is thionitrobenzoic acid, a compound that has a maximum absorbance at 412 nm. The amount of thionitrobenzoic acid produced is a measure of the aminoglycoside concentration. For successful application of this assay, protein impurities must be removed. One should use a more pure enzyme suspension than that required for radioenzymatic assay procedures. With this assay system, gentamicin blood levels can be obtained within 15 minutes (138). Efforts to increase the sensitivity of the assay and to establish the optimal reaction conditions are underway. It should be noted that vancomycin, which is often used in conjunction with gentamicin and streptomycin, does not interfere in the chemical assays described in this chapter.

β-Lactam Antibiotics

Concentrations of the penicillins and related antibiotics in human specimens have been spectrophotometrically determined by a variety of methods. Most methods rely on a β-lactamase to hydrolyze the β-lactam ring of the drug and involve the measurement of the end products of the reaction (139,140). Historically, these methods have suffered from an inability to distinguish one form of penicillin from another. A relationship can be established between the consumption of iodine by a penicillin and the quantity of the penicillin in solution.

In one useful method, interfering protein is easily removed. Isopropyl alcohol is added to an aliquot of serum to remove the protein. The protein is precipitated by centrifugation (16,000 rpm in a Sorvall RC-2 centrifuge [Thermo Fischer Scientific, Inc., Waltham, MA, Norwalk, CT]). Two equal volume aliquots of the supernatant are taken. One aliquot is used as the sample, and the other is used as a blank. Iodine solution (0.01 N), buffered at pH 6.5, is added to each aliquot. A measured amount of aqueous penicillinase (Rikker penicillinase, 1,000 units/mL) is added to the sample while an equal amount of distilled water is added to the blank. After 25 minutes, the excess iodine in each tube is quantitated using a thiosulfate reagent and a starch indicator. The penicillin concentration can be calculated from the difference in iodine uptake between the specimen and the blank. The amounts of base and penicillinase that are required, as well as the incubation times, vary with the penicillins that are being measured. As a general formula, the amount of penicillin-type antibiotic (although susceptibility to penicillinase varies) can be calculated by the general formula:

µg/mL penicillin = (V2−V1)(MW/N)/sample weight in mg

where V2 is the volume of 0.01 N iodine consumed after inactivation with alkali, V1 is the iodine consumption before inactivation, MW is the molecular weight of the individual penicillin, and N is the number of iodine equivalents (141).

Another technique is to assay for a specific constituent of the penicillin molecule. This approach proved successful in the determination of 6-amino-penicillinoic acid with a glucosamine reagent (142). An additional method of analysis relies on the differential absorption of light in a given wavelength by different penicillin molecules to determine antibiotic levels when more than one drug is present. Ampicillin has a higher absorbance at 268 nm in a solution of pH 5 than in one at pH 9. This property has been used to measure concentrations of ampicillin in the presence of cloxacillin (143).

To avoid the difficulties inherent in the alkalized starch-iodine method, a procedure was developed that substituted chloroplatinic acid for base in the assay. Chloroplatinic acid degrades penicillin to penicillinoic acid, which can be measured colorimetrically. The color generated is more stable than that produced with the alkaline method. In the assay, 0.13 mL of 0.2% chloroplatinic acid is added to the penicillin-containing specimen and brought to a final volume of 4 mL with distilled water. The mixture is kept at room temperature for 30 minutes, after which time 1.5 mL of starch-iodine color reagent is added. The starch-iodine color reagent is prepared by adding equal volumes of water-soluble starch solution (0.8%) and 480 mol/L of a 4.8 mmol/L potassium iodine solution. After incubation for 5 minutes at room temperature, the amount of penicillin may be calculated from the absorbance measurement at 260 nm (101).

Active Sulfonamides

The body fluid is extracted into ethyl acetate from a nondeproteinized sample, yielding active unchanged sulfonamide and an acetylated inactive component. The acetylated component does not react in this assay.

Reagents.The following reagents are used:

 1.  McIlvain buffer, pH 5.5, which is made by mixing 8.6 volumes of a 0.2 mol/L aqueous solution of citric acid with 11.4 volumes of a 0.4 mol/L aqueous solution of disodium phosphate

 2.  Ethyl acetate

 3.  2 N solution of HCl in acetone/water. The required quantity of this reagent must be freshly prepared immediately before each series of analyses by mixing one volume of 8 N HCl with three volumes of acetone. This product should not be kept for more than a few hours and should be discarded as soon as a brown color develops.

 4.  0.1% sodium nitrate in a mixture of 3:1 acetone/distilled water

 5.  5% solution of sulfaminic acid in 3:1 acetone/distilled water

 6.  0.1% solution of α-naphthylethylenediamine dihydrochloride in a 3:1 mixture of acetone/distilled water

 7.  Methanol

Procedure.For plasma samples, 1 mL of McIlvain buffer is pipetted into a 10- to 15-mL shaking tube that is sealed by either an ether-tight glass or polyethylene stopper. Plasma (0.1 mL) and ethyl acetate (5 mL) are added. The tube is mixed by shaking for 10 minutes for extraction and simultaneous partial deproteinization and centrifuged 5 minutes at 3,000 rpm. The proteins settle between the two liquid phases as a fine precipitate.

Three milliliters of the supernatant is transferred to a test tube and 0.5 mL of HCl is added and mixed. Then 0.5 mL of 0.1% sodium nitrite solution is added, mixed, and allowed to stand for 6 minutes. One-half milliliter of sulfaminic acid solution is added and mixed well by shaking until there is no further liberation of gas bubbles. After 3 minutes, 0.5 mL of α-naphthylethylenediamine dihydrochloride solution is added and mixed. One-half milliliter of absolute methanol is then added. The tube is mixed until a homogeneous liquid phase is achieved. The tube is closed with a polyethylene stopper. In 20 minutes to 1 hour, one can determine the concentration of the product photometrically, as described in the following texts.

Total Sulfonamides

Reagents.The following reagents are used:

 1.  20% trichloroacetic acid

 2.  3 N aqueous HCl

 3.  0.1% aqueous sodium nitrite

 4.  0.5% aqueous sulfaminic acid

 5.  0.1% aqueous solution of α-naphthylethylenediamine dihydrochloride (Note that all reagents can be stored at −20°C for up to 1 year. Some reagents may freeze and should be well mixed after defrosting.)

Procedure.Four milliliters of distilled water is pipetted into a shaking tube that holds 10 to 15 mL and can be closed with either an ether-tight glass or polyethylene stopper. Then 0.2 mL of plasma is added and the tube is mixed well. The tube is placed in a boiling water bath for 4 minutes. One milliliter of 20% trichloroacetic acid is immediately added, followed by thorough mixing of the tube. The tube is centrifuged for 10 minutes at 3,000 rpm. Three milliliters of the supernatant is transferred to a test tube and 0.5 mL of 3 N HCl is added and mixed. The tubes are sealed and placed in a boiling water bath. After 1 hour, hydrolysis is complete. The tubes are allowed to cool and any condensation drops that may have formed along the walls are washed down. One-half milliliter of 0.1% sodium nitrite is added, mixed, and allowed to stand for 6 minutes. Sulfaminic acid (0.5%) is added to the tube, which is mixed by shaking until no gas bubbles are released. One-half milliliter of 0.1% aqueous α-naphthylethylenediamine dihydrochloride is added and mixed. The solution can be measured spectrophotometrically, as described later, in approximately 20 to 60 minutes.

For urine samples, the procedure is the same as that described for plasma, except that in the assay for active sulfonamides, 0.15 mL of urine is mixed with 5 mL of McIlvain buffer, and 1 mL of this mixture is added to ethyl acetate. In the assay for total sulfonamides, 0.5 mL of urine is diluted with 20 to 50 mL of distilled water (depending on the amount of sulfonamide present). Standards in a similar solvent should be prepared to cover the range of sulfonamide concentrations that are expected in the sample and run in parallel with the patient samples. In addition, distilled water and a sample of the same type of specimen (e.g., plasma or urine) from a patient who has not received sulfonamides are also run as negative controls. All tubes are assayed at 554 nm. The value for each tube is determined by subtracting from the reading for each patient sample the reading obtained from the body fluid that does not contain sulfonamides. A standard Beer’s Law or Beer-Lambert Law type of curve is constructed. One can interchange standard curves among the sulfonamides.

Occasionally, there may be so much sulfonamide present in a sample that it may not fall on the straight portion of the line. In these instances, samples should be diluted in the same body fluid from which they were collected and the assay should be repeated. If the identical body fluid is not available, distilled water can be used as a diluent with little error for all body fluids except bile. These methods are satisfactory for all sulfonamides.


Trimethoprim (TMP) can be extracted from body fluids with chloroform at a basic pH, extracted back into dilute H2SO4, and oxidized with KMnO4 in an alkaline milieu to yield the fluorescent trimethoxybenzoic acid (TMBA) (144).

Reagents.The following reagents are used:

 1.  0.1 N sodium carbonate solution

 2.  analytical-grade chloroform

 3.  0.01 N H2SO4

 4.  0.1 mol/L KMnO4 in 0.1 N NaOH

 5.  35% formaldehyde

 6.  1 N H2SO4

Procedure.For the production of standard solutions of TMP, (a) 29.24 mg of TMBA is weighed in a 100-mL volumetric flask, which is then filled with chloroform; and (b) 5 mL of the solution from step (a) is added to a 100-mL volumetric flask, which is then filled with chloroform (this yields 20 µg/mL TMP). From this working standard solution, dilutions can be made in chloroform to cover the range of TMP that is encountered in biologic material.

To extract the antibiotic, 8 mL of Na2CO3, 10 mL of chloroform, and 1 or 2 mL of biologic fluid are added to a 25-mL shaking tube. The tube is stoppered and, to avoid emulsion, inverted gently head-over-tail for 4 minutes. The tube is centrifuged for 10 minutes at 3,000 rpm. As much of the aqueous phase as possible is aspirated into a new shaking tube and 4 mL of 0.01 N H2SO4 and 4 mL of the chloroform extract from the previous step are added. The tube is mixed by shaking for 10 minutes and centrifuged as described.

For oxidation of TMP to TMBA, 3 mL of the H2SO4 extract and 2 mL of the alkaline KMnO4 solution are added to a shaking tube. The tube is mixed and placed in a 60°C water bath for 20 minutes. Then 0.3 mL of formaldehyde is added and the tube is mixed. One milliliter of 1 N H2SO4 is added, and the tube is placed in a 60°C water bath for 20 minutes. The tube is mixed and cooled to room temperature.

For extraction of TMBA, 2 mL of chloroform is added and the tube is mixed by vigorous shaking for 10 minutes, followed by centrifugation at 3,000 rpm. The clear chloroform phase is transferred into a quartz cuvette, and fluorescence is measured at an activation wavelength of 375 nm and a fluorescence wavelength of 360 nm. The amount of TMP is determined by

C = (Ma)(Cs)(F)/Ms,

where C is the concentration of TMP in the sample, Cs is the concentration of drug in the standard, Ma is the fluorescence of the TMP-containing sample after blank subtraction, Ms is the fluorescence reading of the standard after subtraction of the chloroform bank, and F is a constant that takes into account the conversion yield of TMP in TMBA (for 1 mL of body fluid, it is 5.442 and for 2 mL, it is 2.721).

Plotting the amount of TMBA and TMP (obtained through the conversion factor) versus fluorescent intensity should yield straight lines. The method is useful for the determination of TMP in samples from all body fluids. It has a sensitivity of 20 ng of TMP/mL of plasma. Other drugs, including the sulfonamides, do not react in this procedure. The method determines the concentration of unmetabolized, active TMP.


CAM is used for the therapy of acute bacterial meningitis, rickettsial infections, typhoid fever, and brain abscesses. However, CAM may cause hematologic toxicity. Rarely, aplastic anemia with a high fatality rate occurs. In addition, CAM can cause bone marrow suppression, which is reversible and dose-related. Toxicity occurs in relation to dose when plasma levels exceed 80 mol/L (25 µg/mL). Therapeutic monitoring of serum CAM concentrations may be helpful in evaluating and maintaining effective levels of this potentially toxic antibiotic. This may be particularly important for patients with compromised liver status. In premature neonates, metabolism of the drug is unpredictable. High serum levels in these infants can produce the fatal gray baby syndrome.

In one useful chemical method for the determination of CAM concentrations, the drug is extracted from clinical specimens in isoamyl acetate. After extraction, the CAM concentration can be determined by analysis of the yellow color that develops when the extract reacts with isonicotinic acid hydrazine and sodium hydroxide (145). It appears that many of the biologic breakdown products of CAM are not extractable by this procedure.

Duplicate standards are prepared at 10, 20, and 30 µg/mL. Two milliliters of a phosphate buffer (0.1 mol/L; 2.21 µg of NaH2PO4 · H2O and 7.61 g of Na2HPO4 · H2O in 1 L of distilled water) is added to all tubes. Then 0.5 to 1.0 mL of serum or a standard solution is added. Distilled water is added to an additional tube as a negative control. Three milliliters of isoamyl acetate (Fisher Scientific Co, Hampton, NH) is pipetted into each tube. Each tube is tightly stoppered, mixed well by shaking for approximately 10 minutes, and centrifuged (16,000 rpm in a Sorvall RC-2 centrifuge [Thermo Fischer Scientific, Inc., Waltham, MA, Norwalk, CT] is optimal). To each tube that contains 2 mL of the supernatant solvent, 1.0 mL each of 1.5 N NaOH and 3% isonicotinic acid hydrazide (Eastman Kodak Co, Rochester, NY) are added. The tubes are then stoppered and incubated in a water bath at approximately 30°C for 45 minutes. The tubes are agitated periodically to ensure good mixing. After the incubation step has been completed, the yellow underlayer is aspirated with a Pasteur pipette and the absorbance is measured with a spectrophotometer at 430 nm. The blank is read as the negative control. The standard dose-response curve is then constructed by plotting the absorbance at 430 nm (the reading obtained from the test less the reading obtained with the blank) on the x axisversus the logarithm of antibiotic concentration on the y axis. Another colorimetric assay for CAM is described next.

Reagents.The enzyme reagent includes 200 mmol/L glycylglycine, pH 8, 1 mmol/L magnesium chloride, 7 mmol/L oxamic acid, 0.09 mmol/L acetyl coenzyme A, 60 U/L CAM acetyltransferase (CAT), 90.5 mmol/L nicotinamide adenine dinucleotide (NAD), 0.2 mmol/L thiamine pyrophosphate (TPP), 0.6 mmol/L 2-oxoglutarate, 20 U/L 2-oxoglutarate dehydrogenase (2-OGDH), and 0.01% GAFAC RE-610 (GAF Corp, Wayne, NJ). This reagent must be used within 2 hours of preparation. However, if the acetyl coenzyme A is omitted, the reagent can be stored at 4°C for 24 hours without a significant loss in activity of either enzyme. The color reagent, a solution of 0.2 mmol/L 2-(2-benzothiazolyl)-5-styryl-3-(4-phthalhydrazidyl) tetrazolium chloride in 12 mmol/L citric acid, that contains 0.02 mmol/L 1-methoxy-phenazine methosulfate, 0.04% Nonidet P-40, and 0.1% sodium azide, is stored in a dark bottle at 4°C. Citric acid provides maximum reagent stability.

Procedure.The enzymatic reactions are individually optimized with respect to buffer type, pH, and substrate and cofactor concentrations. To facilitate a rapid reaction, acetyl coenzyme A is required by CAT at a concentration that is in excess of the sample CAM concentration. The 2-OGDH reaction requires the substrate 2-oxoglutarate and the cofactors NAD, TPP, and magnesium ions. Oxamic acid is included as an inhibitor of endogenous serum lactate dehydrogenase activity. When the two enzymatic reactions are combined, glycylglycine buffer (200 mmol/L, pH 8) facilitates rapid reactions and gives maximum enzyme stability.

Dehydrogenase activity is detected by using reduction of the tetrazolium salt 2-(2-benzothiazolyl)-5-styryl-3-(4-phthalhydrazidyl) tetrazolium chloride (146) to a formazan dye, with the highest molar extinction coefficient under the prevailing assay conditions. CAT activity is determined by measuring the increase in absorbance at 412 nm of an assay mixture containing 100 mmol/L Tris-HCl, pH 8.0, 0.1 mmol/L acetyl coenzyme A, 0.1 mmol/L CAM, and 1 mmol/L 5,5-dithiobis (2-nitrobenzoic acid) (Ellman’s reagent or DTNB). The reaction is initiated by adding 25 µL of CAT to 1 mL of assay mixture in a semimicrocuvette (path length of 1 cm).

2-OGDH (EC activity is determined by measuring the increase in absorbance at 340 nm at 30°C of an assay mixture (1 mL) that contains 50 mmol/L of potassium phosphate buffer, pH 8.0, 1 mmol/L MgCl2, 2.5 mmol/L NAD, 0.2 mmol/L TPP, 0.1 mmol/L coenzyme A, 2.5 mmol/L cysteine, and 2 mmol/L 2-oxoglutarate. The reaction is initiated with 25 µL of 2-OGDH.

The serum sample or CAM standard (0.1 mmol/L, 32 mg/L, 0.1 mL) is added to the enzyme reagent (0.5 mL) in a semimicrocuvette (path length of 1 cm), mixed well, and incubated at room temperature for 4 minutes. Color reagent (0.5 mL) is added and, after incubation at room temperature for exactly 2 minutes, the absorbance of the reaction mixture is measured at 575 nm. After 2 minutes, the reaction mixture exhibits a gradual increase in absorbance. A sample blank is prepared and its absorbance is measured by following the same procedure, except that glycylglycine buffer (200 mmol/L, pH 8) is substituted for the enzyme reagent.

The method is based on the reduction of a pale tetrazolium salt to a strongly colored formazan dye by NADH. The thiol groups of 2-OGDH also act as reducing agents, causing formazan production independent of the NADH reaction. Therefore, a reagent blank is required in addition to the sample blank, the absorbance of which is added to the sample blank value. The absorbance of the reagent blank is constant.

Performance.The assay is linear over the CAM range of 5 to 200 mol/L. The intra- and interbatch coefficients of variation (precision) are 1.4% to 4.9% and 4.3% to 6.3%, respectively. Mean recoveries (accuracy) from CAM-spiked (0.1 mmol/L and 0.025 mmol/L) serum samples from normal individuals and patients with renal failure were 98.4% and 105.6% for serum from normal individuals and 100.9% and 106.8% for serum from patients with renal failure. The method does not detect the inactive prodrug forms of CAM, that is, CAM succinate (intravenous preparations) and CAM palmitate (oral suspensions). Of the metabolites tested (CAM base, reduced base, and glycolic acid), only glycolic acid is recognized by CAT, with a cross-reactivity of 81%. However, this is a minor metabolite (<3%) and its detection is not considered important to the clinical utility of the assay.

Previous work indicates that CAT does not recognize CAM glucuronide (147). The method correlates well with reversed-phase high-performance liquid chromatography (RP-HPLC), which is specific for microbiologically active CAM. The assay also detects thiamphenicol, a CAM analog, with similar sensitivity and with a linear response up to a serum concentration of 100 µM. The endogenous colored compounds bilirubin and hemoglobin interfere with the color reaction when they are present in serum at concentrations above 200 µM and 0.2 mg/dL, respectively, because of a shift in the optimum wavelength of the final color. Thus, because of colorimetric interference, this assay is not recommended for use with grossly hemolyzed samples or when bilirubin concentrations exceed 200 µM.

A main advantage of this procedure is the speed with which an accurate CAM measurement can be obtained. The method requires no pretreatment of the sample such as heating or solvent extractions, as is required for HPLC, the more commonly employed assay for CAM. Moreover, a result is available within 6 minutes. The precision of the assay described here, when performed manually, is similar to that of the automated enzyme-multiplied immunoassay technique (EMIT) (148). However, it has the advantages of producing a linear response (allowing a single-point calibration) and requiring only a simple spectrophotometer to measure absorbance. Because it is a two-reagent system, the assay may also be adapted for a wide range of discrete analyzers. The method is specific and shows no significant interference by high concentrations of urea, creatinine, or phenolic compounds, which may be present in the serum of patients with renal failure (147).


Bacteria are often resistant to antibiotics because they produce inactivating enzymes. Benveniste and Davies (149) and Davies et al. (150) provided the basis for radioenzymatic techniques in their description of a method by which they could determine the types of enzymes that inactivated certain antibiotics. Their basic method is used for the theoretical study of bacterial resistance.

Radioenzyme assays have been largely replaced. They were originally a by-product of the study of how enzymes destroy aminoglycosides and CAM. Table 8.2 presents the general advantages and disadvantages of radioenzymatic assays. Radioactive ATP in the presence of adenylating enzyme transfers radioactive 14C to the aminoglycoside. Aminoglycosides, which are positively charged, stick to negatively charged phosphocellulose papers. By enumerating the radioactive counts on these phosphocellulose papers, the extent to which adenylation occurred can be measured. Acetylating enzymes, which transfer acetyl groups from radioactive acetyl coenzyme A to aminoglycosides, can be similarly employed, with the measured counts on phosphocellulose paper reflecting the amount of acetylation that took place (149,150).

These adenylation and acetylation reactions provide the basis for radioenzymatic assays. It should be noted that the reactions are stoichiometric in that the amount of transfer in both adenylation and acetylation reactions is directly related to the quantity of antibiotic in the solution. The reaction takes place in several steps.

Many authors have used a method by which periplasmic enzymes are released from bacteria due to changes in osmotic pressure (151,152). By this method, less than 4% of the intracellular bacterial contents are released. However, the method is time-consuming and technically involved. Sonication has been investigated as a faster and easier method of obtaining the enzyme (153). The enzymes obtained by sonication appear to be as effective as those produced by osmotic shock. However, enzymes obtained by sonication may be somewhat more contaminated and unstable because of the release of proteolytic enzymes inside the bacteria (154). This could limit the length of time that the enzyme can be stored, a critical factor in the long-term usage of radioenzymatic techniques (155).

One may prepare the sonicate as follows. A 16-hour culture of E. coli RS/W677 is centrifuged at 14,000 × g for 5 minutes. The sediment is washed twice in 30 mmol/L NaCl plus 10 mmol/L Tris-HCl, pH 7.8. After the second wash, the pellet is suspended in 0.5 mmol/L MgCl2 at 4°C (5 mL of 0.5 mmol/L MgCl2 per 100 mL of original culture volume). The suspension is then sonicated for 20 seconds using a Dawe-type 3057A Soniprobe (Dawe Instruments Ltd, London, United Kingdom) set to give a 4 amp current. The cellular debris is removed by centrifugation at 25,000 × g for 20 minutes. The supernatant is divided into 0.1-mL aliquots and stored at 20°C (94).

Because it is undesirable to have to prepare the enzyme frequently, its stability has been studied under various conditions (154). Several methods have been developed to decrease enzyme lability. Keeping the enzyme frozen and in an ice bath are effective techniques (36,156). The enzyme has been shown to be stable for 24 hours at 4°C and 30 days at −20°C with BSA (157). Storing partially purified enzyme in reducing agents appears to increase its life span (158). Freezing the enzyme at low temperatures, such as −70°C, in quantities that will be used in a day’s run, appears to be the most efficient and effective technique (159,160). It increases the storage life of the enzyme and significantly decreases the amount of technician time that is required to set up the assays.


A molecule is fluorescent when it can receive light at one wavelength and emit it at another wavelength. The wavelengths at which a given molecule receives and emits light are often specific to the molecule, a reactive group on a molecule, or a class of molecules. One major disadvantage of these assays is that other material present in the specimen, particularly radiologic fluorescein dyes and certain proteins, can fluoresce and interfere with the test. However, fluorescent assays typically are much more sensitive than chemical assays and can quantify compounds in the nanogram and often picogram per milliliter range.

Fluorescence can be determined in one of two ways. First, one may use the inherent fluorescent properties of the molecule. Second, for antibiotics that are either weakly fluorescent or nonfluorescent, one may covalently link a strongly fluorescent moiety to the drug in question. Because of the need for an extraction step, the possible need for coupling steps, the need for rather specialized equipment, the lack of standardized techniques, and the frequent inability to distinguish between active and inactive antibiotics, fluorescence methods have not been widely used in clinical laboratories, although they have been used in industry and U.S. Public Health Service laboratories. The reader is referred to the book by Undenfriend (161) for a general view of fluorescent analysis.

Although penicillins and cephalosporins are not generally inherently fluorescent, many produce fluorescent compounds under hydrolysis in the presence of acid (162). Attempts to simplify the extraction procedures and the number of technical manipulations that are required to generate such reactions have led to methods for measurement that can be clinically useful (141,163). For example, ampicillin concentrations can be determined in the absence of ampicillinoic acid by extraction. Standards should be prepared to cover the expected range of concentrations and serum blanks should be run to obtain measurements of background fluorescence.


Different tetracyclines require different fluorescent reagents to enhance their light-emitting properties (164). Each tetracycline should be tested individually to optimize these assays. Hall (165) has expanded the technique to measure the concentrations of tetracycline mixtures in plasma by using acid hydrolysis or alkaline degradation to convert the tetracycline into a fluorescent form. In these methods, aluminum salts are used to enhance the fluorescence of the end products. This fluorescence is measured with a spectrofluorometer. Each tetracycline exhibits a different structural arrangement of the chemical groups that surround the fluorescent nucleus of the anhydrous salts, creating individual fluorescent characteristics (165). A general method for the fluorometric determination of tetracyclines follows (166).


An Aminco-Bowman spectrophotofluorometer (American Instrument Company, Silver Spring, MD) fitted with a xenon arc lamp and an R 136 photomultiplier, or its equivalent, should be employed. Mirrors and 1-mm slits are placed in the cell housing.


For most tetracyclines, one can use 0.5 mol/L magnesium acetate tetrahydrate plus 0.3 mol/L sodium barbitone in ethanedial. Minocycline, and other 7-aminotetracyclines, can be measured with a mixture of 0.2 mol/L magnesium acetate and 0.2 mol/L citric acid in ethanedial.


A 0.2-mL aliquot of the sample (serum or other fluid) is mixed with 0.4 mL of a phosphate buffer (3 mol/L NaH2PO4 plus 1 mol/L Na2SO3) and thoroughly extracted with 2.5 mL of amyl acetate. The phases are separated by allowing them to settle or centrifuging the tube (approximately 500 × g). Two milliliters of the organic (top) phase are transferred to a Brown fluorometer cuvette. A suitable fluorescence reagent (0.6 mL) is added. (Fluorescence reagents are described later.) The tubes are mixed by shaking for 5 minutes. The turquoise fluorescence in the lower phase is read 20 minutes or more after shaking. If the lower phase is cloudy, the tubes are centrifuged (500 × g for 2 or 3 minutes) before reading. Standards (0.2 mL of a 10 mmol/L solution of the appropriate tetracycline) and blanks (0.2 mL of water) are also made.

For the determination of minocycline, a pH 6.5 buffer (0.5 mol/L NaH2PO4 plus 0.5 mol/L Na2HPO4) should be used instead of the phosphate sulfite buffer (137). Fluorescence is measured with excitation at 405 nm and emission at 490 nm. The fluorescence of 7-aminotetracyclines is read with excitation at 380 nm and emission at 480 nm (166).


Immunologic assays came into use in the mid-1970s. These assays were based largely on existing equipment and merely exploited procedures available for the assay of hormones. However, for the first time, the assay of antibiotics was removed from the realm of the specialist. These assays could be performed in a central location, with the instrument playing the primary role. The development of the means to elicit specific, high-titered antibodies to hapten antibiotics allowed RIA techniques to deliver a specificity that was impossible with biologic agents. As with any nonbiologic assay, however, one always had to be particularly careful not to measure a nonactive metabolite.

Because RIA equipment was expensive, the measurement of antibiotics was often mixed in with the assay of other drugs, resulting in significant delays in processing. The requirement to maintain a stock of highly active radioisotopes (often with short half-lives) gave impetus to the development of nonisotopic immunoassays. Unlike hormones, which are present in extremely small amounts and require highly sensitive methods of detection, antibiotics are generally present in levels above 0.5 µg/mL. Because other small molecules, such as antiepileptics and drugs of abuse, are also present in these levels, a technology was developed to measure small molecules by somewhat less sensitive, nonisotopic means. Unlike the stimulatory role played by existing RIA equipment in the development of RIA, the impetus for the development of the new nonisotopic immunoassays was largely the need to assay the aminoglycoside class of antibiotics.

Before the application of this technique to the measurement of antibiotic concentrations in 1975, RIA had been used for a number of years for the quantitation of hormonal substances (167). All RIAs of antibiotics employ three broad reaction steps. The first step involves three components: radiolabeled antigen (antibiotic); high-titer, high-avidity antibody to that antigen; and unlabeled antigen (antibiotic obtained from the patient’s serum). The reaction with either labeled or unlabeled antigen produces antibody combined with labeled antigen or antibody combined with unlabeled antigen. The more unlabeled antigen that is present in the reaction mixture, the less radiolabeled antigen combines with antibody.

After equilibrium is reached, one must quantify the amount of bound antibody in the mixture. This step involves either the removal of the bound antigen-antibody complex from solution by a precipitating agent or the removal of the unbound antigen from solution by chemical means. Because antigen/antibody reactions are stoichiometric in nature, the quantification of either the bound radiolabeled or the free radiolabeled antigen is directly proportional to the antibiotic level in a sample (168170).

Although there are individual modifications, the test procedures follow a basic course (167). First, one incubates a known quantity of tritiated antibiotic with antibody (of known potency) to that antibiotic. A given amount of patient’s serum is added and the mixture is allowed to come to equilibrium. These assays are heterogeneous and produce sigmoidal coprecipitation curves. Early investigators were hampered because only the relatively linear part of the sigmoidal curve could be used for the calculation of antibiotic concentrations. Robard et al. (171) devised a method in which the sigmoidal curve was converted to a straight line, thereby permitting the calculation of results over a much wider range of values. They found that the plot of y = 100 (B/B°) may be linearized by the equation representing a straight line in which logit (y) = A + B log x, where x is the amount of unlabeled or patient drug, and yrepresents the percentage of antibiotic that is bound.

This logit conversion not only allowed a much wider range of antibiotic concentrations to be measured but also permitted the development of standard curves. As a result, one did not have to repeat all the controls each time a specimen was run. An advantage of immunologic assays is their within-class specificity. Although there may be cross-reactivity among members of the same drug class, there is no cross-reactivity among members of different classes (e.g., gentamicin and vancomycin).

Nonisotopic Immunoassays

The development of EIAs by Engvall and Perlmann (172,173) laid the foundation for design of nonradioisotopic immunoassays. In these immunoassays, an enzyme or fluorometric substance is coupled to the antigen or antibody in such a way that the activity of the parent compound is not appreciably affected. The significant difference between nonisotopic immunoassays and RIAs is the means of counting the label. In nonisotopic assays, the quantity label present is estimated from enzyme or fluorescence activity, which changes under the assay conditions.

Generally, when an enzyme–substrate reaction is used in an immunoassay, enzyme activity is measured as reaction velocity. In this situation, the reaction velocity must be proportional to the number of enzyme molecules that catalyze the reaction (174). In fluorescence immunoassays, a change in the nature of the interaction of light with the substrate is measured (175).

In many nonisotopic immunoassays, the label, whether fluorescent or nonfluorescent, is an enzyme. In order to distinguish RIAs from nonisotopic immunoassays, we must consider the characteristics of an enzyme label. First, it must be recognized easily as the label. The enzyme must be attached to the substrate in such a way that it maintains activity yet performs satisfactorily under the assay conditions. Second, the label must be stable. The label must not disintegrate when the assay is in process and it must be stable during storage. Invariably, the enzyme is covalently bound to the labeled molecule. Third, the enzyme must be quantitatively measurable. In nonisotopic immunoassays, one measures a secondary reaction product (i.e., produced by a substrate acting on the enzyme), rather than direct release of a radioisotope (176,177). Therefore, we do not measure the enzyme molecules themselves but instead measure the catalyzed reaction processes. With any enzymatic label, the product that is produced must be proportional to the amount of enzyme that catalyzes the reaction.

In considering the theory of enzyme activity and its relationship to the clinical assay, we must remember the Michaelis-Menten equation:

v = V [S/(S + Km)],

where v is the measured reaction velocity, S is the concentration of substrate, and Km is the calculated constant. In practice, one attempts to design the assay conditions so that S is much greater than Km. In this case, we may assume that v is approximately equal to V and that

v = (k)E2,

where E2 is the total amount of enzyme that is present in the reaction mixture. It should be noted that this equation implies that the reaction velocity is independent of time.

The equations just described require that the reactions be followed continuously as they occur. One must measure the accumulation of product or the change in fluorescence as it occurs over time. There have been three primary means of quantitatively following enzyme reactions for the assay of haptens, including antibiotics, under these conditions. First is the spectrophotometric assay. The spectrophotometer is useful when the product of the reaction can be measured between 190 and 800 nm. Most workers have attempted to measure the release of the product directly. However, the product may first have to be transformed into a colored product. The advantage of using a spectrophotometer is that this instrument is readily available in clinical laboratories.

Much work has involved the study of fluorometric methods, which is the second approach. The outstanding feature of fluorometry is its high degree of sensitivity. Fluorometic methods are generally 100 to 1,000 times more sensitive than spectrophotometric methods. This sensitivity is especially manifested either when the initial substrate is nonfluorescent and becomes so during the reaction or when there is a change in fluorescence polarization as the reaction proceeds.

The third method, which employs electrodes, offers several advantages. Hydrogen ion concentrations and automatic titration apparatus may be employed to continuously measure a reaction as it proceeds. Considerable work has been performed on an oxygen electrode, which is a type of polarography (54).

Fundamental to nonisotopic immunoassays is the preparation of hapten–protein conjugates, in which haptens (antibiotic) and proteins (generally an enzyme) both function naturally. Most of the described methods have attached the hapten to the protein by free amino, hydroxyl, or carboxyl groups. Haptens with hydroxyl groups may be conjugated to a carrier protein by activation of the carboxyl group, followed by acylation of amino groups in the protein. The mixed-anhydride procedure has been commonly used for this purpose. Figure 8.6 demonstrates the principles of the reaction. The procedure is performed directly with the hapten and the conjugate.

Alternatively, one may use the carbodiimide procedure, in which uridine-5-carboxylic acid is coupled directly to poly-DL-alanyl-poly-L-lysine with dicyclohexylcarbodiimide in 95% dimethylformamide. The carrier or enzyme, excess hapten, and reagent are simply stirred together in water for 30 minutes each day for several days.

Haptens with amino groups, which include the aminocyclitol/aminoglycoside class of antibiotics, may be coupled to proteins. The reactions that are used to conjugate these haptens depend on whether the groups are aromatic or aliphatic amines. If the amino group is an aromatic amine, the hapten may be conjugated to proteins by the classic diazotization procedure. For example, CAM may be conjugated by this means (178). With this method, one must reduce the nitro group to an amino group before the conjugation reaction can occur. If the hapten contains an aliphatic amine, it can be reacted with carboxyl groups by the carbodiimide reagent. The conditions of this reaction are straightforward and similar to those employed for the conjugation of carboxyl groups. Optimally, amino groups of the hapten molecule can be acetylated by the hetero-bifunctional reagent N-(m-maleimidebenzoyloxy) succinimide to introduce a maleimide residue. As shown in Figure 8.7, the maleimide residue may be coupled with thiol groups, which are converted from the disulfide bonds of cystine residues by reductive cleavage.

Few enzymes have been used to label antibiotics. Peroxidase was coupled to gentamicin by a modified Nakne’s method (8). One of the first enzymes used, which is still commonly employed in the EMIT system, is glucose-6-phosphate dehydrogenase. The means of covalently linking this enzyme to antibiotics is, however, a proprietary secret. β-D-Galactosidase has also been commonly employed. This reaction has been made possible through the use of hetero-bifunctional reagents such as N-(3-maleimidopropionylglycyloxy) succinimide (MPGS). A number of these reagents are equally useful. The general method is shown in Figure 8.8.

The bonding of ampicillin serves as a useful model of this type of antibiotic labeling. Fifty micromoles of ampicillin are dissolved in 0.05 mol/L of a phosphate buffer, pH 7.0. This is incubated with MPGS for 50 minutes at 30°C. After lyophilization, the powder is washed three times with 10 mL of ether/methylene chloride solution (2:1) to remove excess MPGS. Following desiccation, the powder is dissolved in 0.5 mol/L of a phosphate buffer, pH 6.0. Approximately 30% to 40% of the ampicillin is acetylated by MPGS. The acetylated ampicillin is coupled to the β-D-galactosidase enzyme by dissolving 1 mol in 1 mL of 0.05 mol/L phosphate buffer, pH 6.0, with 93 pmol of β-D-galactosidase and incubating the mixture overnight at room temperature. Thus, the conjugate is separated chromatographically using a Sepharose 6B column (1.8 × 30 cm) with 0.02 mol/L phosphate-buffered saline (PBS), pH 7.0, that contains 0.1% NaN3, as the eluent. One unit of enzyme activity may be defined as the amount of enzyme that hydrolyzes 1 µmol of 7-β-D-galactopyanosyloxy-4-methylcoumarin per minute (178).

Nonisotopic immunoassays invariably use competition between labeled and unlabeled drug for antibody binding sites to derive the antimicrobial concentration in a specimen. These assays may be divided into two types: heterogeneous and homogeneous. The heterogeneous type was the first to be developed and is best exemplified by the enzyme-linked immunosorbent assay. Heterogeneous assays require wash steps to separate the bound and free ligands. Most commonly, these assays have been used to measure and detect large molecules. Homogeneous assays do not require the removal of unbound ligand from the reaction mixture and, therefore, may be performed as single-step assays.

If an antibody is bound to the antigen, the substrate is unable to gain access to the catalytic site of the enzyme. Accordingly, the enzyme’s activity is inhibited. When unlabeled antigen is added, it competes with the enzyme-labeled antigen for binding to the antibody and the free form of the enzyme-labeled antigen is increased. As a result, enzyme activity is increased. Consequently, enzyme activity is proportional to the concentration of unlabeled antigen and can be used to quantitatively assay small hapten molecules.

Most commonly, the inhibition of enzyme activity on binding to antibody is caused by steric modification of the enzyme. Homogeneous assays are inherently more easily automated than are heterogeneous assays. These assays have been extensively developed for the measurement of a wide variety of molecules. The elimination in homogeneous assay of the need to remove unbound ligand from the reaction has made possible the generation of machinery that requires minimal technical input from the clinical laboratory. Automated instruments can measure enzyme activity through detection of reaction products and, with internally programmed computer analysis, derive the concentrations of the antibiotics in question.

Heterogeneous Nonisotopic Immunoassays

An assay for ampicillin is presented as a representative heterogeneous nonisotopic immunoassay. A mixture of 18 units of enzyme-labeled ampicillin (the production of which is described in the previous section) and 50 µL of 104-fold diluted rabbit ampicillin antiserum is incubated. A 0.1-mL sample or standard is added to 0.1 mL of the diluted ampicillin antiserum. To the sample or standard plus antiserum is added 0.5 mL of ampicillin–β-D-galactosidase conjugate and 0.05 mL of a 0.05 mol/L phosphate buffer, pH 6.0. This final volume of reactants is incubated for 6 to 8 hours at 4°C. Following incubation, 0.05 mL of normal rabbit serum, which has been diluted 1:500, is added to 0.05 mL of goat antirabbit immunoglobulin G (IgG) serum, which has been diluted 1:5 and incubated with the reactant solution for 8 to 16 hours at 4°C. After this incubation period, 1 mL of 0.05 mol/L phosphate buffer, pH 6.0, is added and the entire mixture is centrifuged at 2,500 rpm for 20 minutes. The centrifugation step is repeated twice. To the reactant phase, 0.15 mL of substrate solution is added, with incubation at 30°C for 60 minutes. The amount of ampicillin present is calculated in a fluorometer, with excitation wavelength of 365 nm and emission wavelength of 448 nm (Fig. 8.9) (178180).

To avoid the requirement for a centrifugation step, a method was developed that used antibody that was covalently linked to magnetizable particles (59). The sample is incubated with fluorescein-labeled gentamicin and antigentamicin serum to which has been attached magnetic particles. The magnetic particles are rapidly sedimented from the reaction mixture through contact with a polarized surface.

Receptor Antibody Sandwich Assay for Teicoplanin

Teicoplanin, a glycopeptide antibiotic with activity similar to that of vancomycin, is currently used in Europe, Japan, and other countries to treat severe infections caused by gram-positive bacteria. Traditional methods for measuring concentrations of teicoplanin include microbiologic assays, HPLC, and solid-phase enzyme receptor assays (181). These methods have several limitations. First, the microbiologic assay is not specific for teicoplanin in the presence of other antibiotics and is of low accuracy when it is performed on biologic fluids. HPLC requires specialized equipment and laborious extraction procedures for sample preparation. Finally, the solid-phase enzyme receptor assay does not always yield accurate results in complex specimens such as bronchial expectorates or skin homogenates. The following method is a receptor antibody sandwich assay that is able to quantify teicoplanin in complex matrices. The method is based on bioselective adsorption of teicoplanin onto microtiter plates coated with BSA-ε-aminocaproyl-D-alanyl-D-alanine, a synthetic analog of the antibiotic’s biologic target, followed by reaction with antiteicoplanin antibodies. The sandwich complexes are detected by incubation with peroxidase-labeled goat antibodies to rabbit IgGs and a chromogenic reaction with o-phenylenediamine.


Polyvinylchloride microtiter plates (96-well, Falcon Micro Test III flexible assay plates; Becton-Dickinson, Oxnard, CA) are coated with BSA-e-aminocaproyl-D-alanyl-D-alanine. The wells of the last vertical row of each plate are coated with BSA alone at a concentration of 10 mg/L and serve as blank wells. Each well in the microtiter plate contains fixed volumes (0.1 mL) of a standard solution of teicoplanin in PBS (0.15 mol/L NaCl and 0.05 mol/L sodium phosphate buffer, pH 7.3). The plates are incubated for 2 hours at 30°C in a covered, humidified box, washed eight times with PBS that contains 0.5 mL of Tween 201, and dried by shaking.

The wells are then filled with 0.1 mL of rabbit antiteicoplanin antiserum (diluted 250-fold with PBS/Tween 201 containing 3 g/L BSA) and allowed to react for 1 hour at room temperature. Microtiter plates are again washed with PBS/Tween 201. Each well is filled with 0.1 mL of peroxidase-conjugated goat antirabbit antibodies (diluted 1,500-fold with PBS/Tween 201/BSA). After reaction for 1 hour at room temperature and a wash step with PBS/Tween 201, 0.15 mL of chromogenic peroxidase substrate solution (1 g/L o-phenylenediamine and 3.5 mmol/L hydrogen peroxide in 0.1 mol/L sodium citrate buffer, pH 5) is added to each microtiter well. After 30 minutes of color development at 30°C, the reaction is stopped by adding 50 µL of 4.5 mol/L sulfuric acid to each well. Ten minutes later, the absorbance at 492 nm is measured in a Titertek Multiskan photometer (Titertek, Huntsville, AL). The binding curves are obtained by plotting, on semilogarithmic paper, the absorbances at 492 nm as a function of the teicoplanin concentration.


This assay has been used to detect teicoplanin in serum, ascitic fluid, skin homogenates, bronchial expectorates, pleural fluid, and prostate homogenates. Wells coated with only BSA, without BSA-ε-aminocaproyl-D-alanyl-D-alanine, have shown that nonspecific binding is negligible. A dose-response curve that is linear in the teicoplanin range of 0.004 to 0.15 mg/L indicates that the sandwich complex is formed despite the low molecular mass of the antibiotic.

The interaction of teicoplanin with BSA-ε-aminocaproyl-D-alanyl-D-alanine and the antiteicoplanin antibodies is highly specific for teicoplanin because the receptor antibody sandwich assay does not give a response when other glycopeptide antibiotics are present in the concentration range of 0.001 to 100 mg/L. However, when teicoplanin solutions are assayed in the presence of vancomycin, recoveries of teicoplanin are lower than expected. Because similar effects are likely with other antibiotics of the same class, receptor antibody sandwich assays should not be used to determine teicoplanin concentrations in the presence of other glycopeptide antibiotics without performing studies that investigate possible interferences. The mean analytical recovery of teicoplanin with this assay is 99.5%. The interassay coefficient of variation is 5.13% for all samples. The detection limit is 0.03 mg/L. Receptor antibody sandwich assays of this type detect the total amount of immunoreactive material that is present in the sample, irrespective of biologic activity (179).

Homogeneous Nonisotopic Immunoassays

Fluorescence Quenching

Fluorescence quenching was the first widely available, homogeneous, nonisotopic immunoassay procedure for the assay of antibiotic concentrations. The method is based on the decrease in fluorescence that results from the combination of antibody with fluorescein-labeled antibiotic. This technique is rapid, sensitive, does not require an extraction step, and is automatable (182184). The procedure is based on the same competitive binding principle that governs RIA.


In a fluorescence quenching assay for gentamicin, a drug that has been made fluorescent through a reaction with fluorescein thiocarbamyl (FTC), is mixed with a serum specimen that contains both gentamicin and antigentamicin antiserum. Antigentamicin antiserum combines with FTC-gentamicin and stoichiometrically decreases the fluorescence of the conjugated antibiotic. As the amount of unconjugated gentamicin in the serum sample increases, the amount of antigentamicin antiserum available for binding to the FTC–gentamicin conjugate decreases, causing a proportional rise in the emitted fluorescence (183,185).


Fluorometric measurements are made with a xenon arc lamp in standard 1 × 1 cm glass cells.


FTC-gentamicin is produced by the reaction of 0.425 g/L gentamicin free base with 0.50 g/L fluorescein isothiocyanate isomer (Sigma Biochemicals, Sigma-Aldrich, St. Louis, MO) in 50 nmol/L sodium carbonate–bicarbonate buffer, pH 9.0. The mixture is incubated for 2 hours at room temperature. Two milliliters of the reaction mixture is chromatographed on a 1 × 97-cm Sephadex G-15 column eluted at 1.8 mL/minute with a carbonate–bicarbonate buffer. The purity of the eluate can be determined by electrophoresing the products on Whatman no. 1 paper in 20 nmol/L of sodium carbonate–bicarbonate buffer, pH 9.0, at 10 V/cm for 2 hours. The band is visualized under shortwave ultraviolet (UV) light. Antigentamicin antiserum is prepared in rabbits, as for the RIA method (186).


Excitation is at 495 nm and emission at 540 nm. In constructing a standard curve, the fluorescence value of a serum sample without gentamicin should be subtracted from both the patient serum and standard readings.

Antigentamicin antibody dilution curves are determined by making doubling dilutions in buffer of antigentamicin antiserum and pipetting 1-mL aliquots into two series of tubes. A 0.5-mL aliquot of a 1:200 dilution of stock FTC-gentamicin solution in buffer is added to each tube of one series and mixed. To each tube of the other series, 0.5 mL of the buffer is added. The tubes of the latter series serve as blanks that allow estimation of the fluorescence produced by the antiserum itself. Incubation is at room temperature for 5 minutes. The fluorescence of the test mixture and blanks is then measured. The corresponding blank signal is subtracted from the total signal of each test mixture. Figure 8.10 presents a typical antibody dilution curve.

The quenching fluorescence assay method is performed by diluting 50 µL of serum specimen 1:50 in buffer. To prepare standards, gentamicin concentrations are made in a geometric series from 0.25 to 32 µg/mL and diluted 1:50 in buffer. Aliquots of 0.5 mL of the diluted samples or standards are pipetted in duplicate into two series of small tubes. To each tube of one series, 0.5 mL of the 1:200 dilution in buffer of the stock FTC-gentamicin solution is added and mixed. Then 0.5-mL aliquots of a 1:80 dilution in buffer or antigentamicin serum are added to each tube and the contents are immediately mixed. To each tube of the other series, 1 mL of buffer is added and mixed. This latter series serves for estimation of the fluorescence blank signals contributed by the intrinsic fluorescence of the serum samples or standards themselves.

The assay mixtures and blanks are incubated at room temperature for at least 5 minutes. All tubes are read in the fluorometer. The corresponding blank signal is subtracted from the total signal of each assay mixture tube. The relationship between the amount of gentamicin present in the sample and the fluorescent intensity is presented in Figure 8.11.


The fluorescence quenching method correlates with the RIA procedure at a correlation coefficient of 0.98. FTC-gentamicin is stable for more than 1 year. Once the antibody has been standardized, minimal technical manipulation is required for the method. Results are available within 15 minutes of receipt of a specimen. Also, the procedure is amenable to automation using continuous flow-type chemical analysis (183,185).


In an effort to develop a rapid and simple assay, the cost of which approaches that of the microbiologic assay, a hemagglutination inhibition assay for gentamicin was developed (112). The technique applies to gentamicin that is covalently linked to BSA, which is covalently linked to sheep erythrocytes and fixed in formaldehyde. This preparation is stable for up to 8 months in a refrigerator. The test is performed in Linbro microtiter plates (MP Biomedicals, Santa Ana, CA) with V wells. In principle, one drop of the patient’s diluted serum is mixed with one drop of antiserum and, after 5 minutes at room temperature, one drop of gentamicin-bound sheep erythrocytes is added. The mixture is then incubated for 60 to 90 minutes at room temperature. Each of the reagents is standardized. If the patient’s serum contains more gentamicin than is neutralized by the antiserum, agglutination of the gentamicin-coated sheep erythrocytes occurs. The lowest dilution of a patient’s serum that causes agglutination is recorded.

A gentamicin standard containing 10 ng/mL is diluted to produce final concentrations of 0.31, 1.86, 2.97, 4.96, 6.51, 9.61, and 18.9 µg/mL in each microwell. Appropriate controls are added to adjacent wells. A given amount of reconstituted antiserum is added to all wells except the control well and the tray is mixed by gentle rotation. After 5 to 10 minutes, 50 µL of reconstituted cells is added to each well and the tray is mixed again by shaking. After 1 hour, the well number showing a pellet of cells equal to that of the control well is recorded. In effect, a titer, similar to cross-over broth dilution assays, is obtained. Although one can obtain only discontinuous values (values for which standard individual control wells are available), the method may prove effective if produced cheaply enough for clinical purposes. Based on the principle stated here, latex particles as carriers have also been marketed for the rapid semiquantitation of the major aminoglycoside/aminocyclitol antibiotics (187).


The EMIT technique was developed from a free radical assay method that employed a spin label and detected drugs by electron spin resonance spectrometry. Electron spin resonance spectrometry requires expensive and very specialized equipment. EMIT was developed to bring the advantages of EIAs to clinical laboratories, without the requirement for large capital expenditures.

The heart of the EMIT is the attachment of an enzyme to the hapten. Commonly, lysozyme from egg whites, glucose-6-phosphate dehydrogenase from the bacterium Leuconostoc mesenteroides, malate dehydrogenase from pig heart mitochondria, and β-D-galactosidase from E. coli have been used as enzymes in EMIT assays. All these enzymes maintain significant activity after conjugation to the assayed drug.

Binding of antibody to the drug–enzyme conjugate inhibits the activity of the enzyme. The large protein antibody sterically hinders the association of the substrate with the active site of the enzyme, resulting in a reduction in the quantity of the product that is produced. The EMIT assay employs the stoichiometric competition between the antibiotic in the serum sample and the enzyme-conjugated drug for antibody binding sites to derive the serum antibiotic concentration. The activity of the conjugated enzyme is decreased on antibody binding and the amount of enzyme activity is directly proportional to the concentrations of free and bound enzymes present in the assay mixture. Therefore, one can derive the concentration of antibiotic in a sample from quantitative measurements of the reaction products by relating them to a standard curve. The major steps in immunoassay by EMIT are as follows:

 1.  Drug of unknown concentration + antibody → antibody–drug

 2.  Drug–enzyme conjugate + antibody → drug-enzyme-antibody

 3.  Substrate + antibiotic–enzyme → product

No separation step is required and the product is measured directly. Table 8.9 presents the major characteristics of the EMIT immunoassay. Table 8.10 lists the major advantages and disadvantages of the EMIT system.

EMIT procedures have been modified for a wide variety of automated instruments. For example, Dade Behring’s Syva EMIT Gentamicin 2000 (Siemens Diagnostics, Tarrytown, NY) homogeneous immunoassays are designed for used with most chemistry analyzers, including the company’s Dimension (Siemens Diagnostics, Tarrytown, NY) combined chemistry/immunochemistry workstations. Many automated instruments automatically analyze the rate of change in absorbance of the sample or standard, correct for the absorbance from the drug-free control, and fit the data to a log-logit curve. In most cases, the coefficients of variation of the instruments range from 2% to 3%, but generally, 5% should be expected. It must be noted that, because the standard curves are not linear, small analytical errors may cause relatively large concentration errors at their limits. Thus, accurate timing and pipetting are crucial. For this reason, automated systems, even though they require larger capital investments and cost more per test, are less labor-intensive and require fewer repeat assays than manual procedures (188,189).

As with immunoassays in general, there are few limitations on the use of EMIT or interferences with its performance. The most common interference is the presence of the unconjugated enzyme in the specimen. For example, lysozyme and malate dehydrogenase may be endogenous in urine specimens. Lipemia, hemolysis, and hyperbilirubinemia do not interfere significantly with EMIT assays. When using EMIT to measure antibiotic concentrations in urine, changes in the pH or ionic strength may introduce errors into the system. This can be particularly pronounced when urease-splitting microbes in the urine increase its pH. It had been noted with the EMIT, as well as with other nonisotopic immunoassay systems, that the reproducibility of standard curves diminished when most of the reagent in a bottle had been used. Since this phenomenon appeared to have been caused by reagent evaporation, many manufacturers began packaging their reagents in smaller vessels (190).

The labor cost per test in the EMIT system is relatively small compared with the microbiologic assay. Approximately 20% of EMIT and 60% of biologic assay costs are for labor. In general, for the assay of antibiotics, the rapid availability of the assays overcomes the requirement for single-drug analysis (191,192).

The basic apparatus for EMIT includes a UV/visible light spectrophotometer with a temperature-controlled cuvette, a timer/printer, and a pipette/diluter. In general, serum or urine samples are diluted with buffer before the antibody–substrate and enzyme–antibiotic reagents are added. The mixture is aspirated into a spectrophotometer and absorbance is measured at two time points.


The assay of amikacin by EMIT is presented as a model for the assay of aminoglycoside/aminocyclitol antibiotics by this method. The principles, apparatus, and procedures are identical for other antibiotics. In the EMIT procedure, serum or plasma is mixed with reagent antibiotic that is coupled to glucose-6-phosphate dehydrogenase (reagent A). After incubation, antibodies to the particular drug are added. Glucose-6-phosphate serves as the substrate for the enzyme, and NAD is used as a cofactor.

The antidrug antibody competitively binds to both the free antibiotic and the enzyme-labeled drug. Antibody binding to the antibiotic–enzyme conjugate inactivates the enzyme. Consequently, as the antibiotic concentration in the specimen increases, the activity of glucose-6-phosphate dehydrogenase proportionally increases. Enzyme activity is reflected in the conversion of NAD to NADH. This reduction reaction produces a color change that is spectrophotometrically measured. Since NAD serves as a cofactor with bacterial, but not human glucose-6-phosphate dehydrogenase, interferences due to the presence of the human form of the enzyme are avoided by using bacterial glucose-6-phosphate dehydrogenase (from L. mesenteroides) in the assay.


A spectrophotometer that is capable of measurement at 340 nm at a constant temperature of 30°C must be used. A data-handling device must be attached to the spectrophotometer to analyze and print absorbance readings. In the past, recommended spectrophotometers have included the Syva S-111 (Dade Behring, Inc, Deerfield, IL) and Gilford Stasar 111 (Gilford Instrument Laboratories, Inc, Oberlin, OH). Each of these spectrophotometers should be set in the absorbance mode, with distilled water set to an optical density of 1.000. The switch mode control is set to concentration, and the set display is placed at 2.667 with the concentration calibrator knob. The display is zeroed with the zero control knob. Data handling has been performed with a Syva CP-5000 clinical processor (Siemens Diagnostics, Tarrytown, NY), Syva CP-1000 timer/printer (Siemens Diagnostics, Tarrytown, NY), or Syva timer/printer model 2400 (Siemens Diagnostics, Tarrytown, NY). Automatic sample handling has been accomplished with the Syva pipette/diluter model 1500 (Siemens Diagnostics, Tarrytown, NY). Any semiautomatic pipette/diluter that is capable of sampling 50 µL and delivering this sample along with 250 mL of assay buffer with sufficient force to ensure that there is adequate mixing of the reactants is satisfactory.


Reagent A is amikacin to which has been covalently coupled glucose-6-phosphate dehydrogenase (the means of coupling is proprietary). This reagent, when reconstituted in buffer, is standardized to work with reagent B. Reagent A is reconstituted from the lyophilized form by adding 6.0 mL of distilled water. Following reconstitution, the reagent may be stored in the refrigerator overnight, but it must remain at room temperature for at least 2 hours before use. Reagent A should always be stored at 2°C to 8°C; it has a shelf life of 12 weeks.

Reagent B contains sheep antiamikacin antibody. This antibody–substrate reagent contains a standardized preparation of sheep γ-globulin, enzyme substrate glucose-6-phosphate, the coenzyme NAD, and preservatives in Tris buffer, pH 5.2. The lyophilized reagent is reconstituted with 6 mL of distilled water and must be allowed to remain at room temperature for at least 2 hours before use. Reagent B may be reconstituted and allowed to remain at 28°C overnight but must come to room temperature before use. It must be stored at 2°C to 8°C and is stable for 12 weeks.

The standard buffer solution used for dilution in the EMIT assay is 0.055 mol/L Tris-HCl buffer, pH 8.0, with a small amount of surfactant. The buffer solution is stable at room temperature for up to 12 weeks. Six amikacin calibrators and a control must be used. The calibrators are reconstituted with 1.0 mL of distilled water and the controls are reconstituted with 3.0 mL of distilled water. After reconstitution, the calibrators and controls must remain at room temperature for at least 2 hours before use. The calibrators and controls may be reconstituted and refrigerated overnight before use but must be at room temperature when the assays are run. The calibrators and controls must be stored at 2°C to 8°C and are stable for 12 weeks. The calibrators contain amikacin concentrations of 0, 2.5, 10, 20, and 50 µg/mL. The controls contain 15 µg/mL amikacin.


Preparations for the test are made as follows: (a) All reagents are prepared as previously described; (b) all reagents must be well mixed and brought to room temperature; (c) all instruments must be properly calibrated; (d) the spectrophotometer must be zeroed with distilled water to 0.000; (e) the pipetter/diluter should be primed and flushed with buffer to ensure that there are no air bubbles in the lines; and (f) there must be a sufficient number of beakers present in the work rack. For calibrating, and for assaying unknowns, measurements should be made in duplicate and the results averaged. Duplicate readings that differ by more than six absorbance units should be repeated. The pipette tips must be carefully wiped with laboratory tissues both before and after the delivery of each solution. Solutions should not be held in the tubing for more than 5 seconds before delivery. The samples are analyzed as follows:

 1.  The calibrator, control, or specimen is diluted by delivering 50 µL of the appropriate solution and 250 µL of buffer to a 2.0-mL disposable beaker.

 2.  The sample is diluted again by adding 50 µL of the diluted sample from no. 1 to 250 µL of buffer solution and delivering this mixture to a second 2.0-mL disposable beaker.

 3.  Fifty microliters of reagent A is mixed with 250 µL of buffer solution, with delivery to the second beaker.

 4.  The spectrophotometer flow cell is purged.

 5.  Fifty microliters of reagent B plus 250 µL of buffer solution is added to the second beaker.

 6.  Immediately after the addition of reagent B, the contents of the second beaker are aspirated into the spectrophotometer flow cell. The printer/recorder should be automatically activated.

 7.  For the remaining samples, controls, and calibrators, steps 1 through 6 are repeated.

 8.  The spectrophotometer flow cell must be cleaned with the cleaning solution supplied with the instrument and the pipette/dilution lines must be stored in distilled water. After a 15-second delay, absorbance readings are made for each sample. The change in absorbance over a 30-second measurement period is used to calculate results.

The difference between the average calibrator zero reading (A0) and the reading of each of the other calibrators (A), known as A – A0, must be determined to plot a standard curve and to calculate the concentrations of the unknown samples. The Syva CP-5000 (Siemens Diagnostics, Tarrytown, NY) clinical processor automatically calculates the concentration of amikacin. The Syva CP-1000 timer/printer (Siemens Diagnostics, Tarrytown, NY) and Syva timer/printer model 2400 (Siemens Diagnostics, Tarrytown, NY) have built-in memory functions that store the A0 reading so that the technologist may perform the necessary calculations on log-logit paper. The technologist derives the amikacin concentrations by preparing a standard curve and plotting A – A0 for each calibrator against the calibrator concentrations on the lot-specific graph paper that is supplied with each reagent kit. A best fit line is constructed. Each time a new bottle of reagent A, reagent B, or buffer is used, a new standard line must be prepared. Furthermore, new standard lines must be drawn whenever duplicate controls vary by more than 10% or if any calibrator point lies more than six absorbance units off the line.


The EMIT amikacin assay is designed to measure the concentration of this antibiotic in serum or plasma. The major form of nontechnical error is generated by cross-reactivity with other compounds. Kanamycin significantly cross-reacts with the amikacin assay. Table 8.11 lists the concentration of a number of antibiotics that are required to produce a 30% measurement error in a sample that contains 10 µg/mL kanamycin. The assay range of quantitation is between 2.5 and 50 µg/mL. The coefficient of variation between runs is typically approximately 10%.

The EMIT has proven accuracy in clinical settings with the following limitations: (a) When β-lactam antibiotics are present in addition to amikacin, specimens must be assayed immediately or stored frozen; (b) severely hemolytic, lipemic, or icteric samples may interfere with the assay; and (c) kanamycin shows significant cross-reactivity with the amikacin assay. Table 8.9 describes the major performance characteristics of the EMIT immunoassay system (193,194).


The recommended peak CAM concentration is from 10 to 20 mg/L (15 to 25 mg/L for meningitis). Because of CAM’s potential toxicity, therapeutic monitoring of blood levels may be desirable with its use. Bioassays involve tedious preparation and lengthy incubation times. HPLC uses expensive equipment, and highly trained personnel are needed to perform HPLC assays. The following method describes a Syva EMIT assay (Dade Behring, Inc, Deerfield, IL) that was developed for measurement of CAM in human serum (58).

The Syva EMIT kit may be purchased from Dade Behring, Inc. Calibrators and buffers are reconstituted according to the manufacturer’s instructions. The assay has been performed with a Cobas-Bio centrifugal analyzer (Roche Analytical Instruments, Nutley, NJ) that was equipped with Data Reduction and Nonlinear Standard Curves (DENS) program version 8326. A standard curve for CAM is stored in the Cobas-Bio analyzer (Roche Analytical Instruments, Nutley, NJ) and the results from the individual serum samples are compared with the standard curve.

The sensitivity of the assay, defined as the smallest amount of CAM that can be accurately measured, is 2.5 mg/L (7.7 mol/L) for EMIT. Lower concentrations of CAM can be detected but not accurately quantified. The within-day precision coefficient of variation for EMIT is 4.0% at 5.0 mg/L, and the between-day coefficient of variation is less than 5.5%. When compared with HPLC and bioassay methods, the EMIT is specific for CAM and correlates well by regression analysis. The EMIT measures only the biologically active base form of the drug, uses a small sample size (0.2 mL), and provides rapid results. However, the reagents are expensive, and personnel need special training to perform the assay.


The substrate-labeled immunofluorescent assay (SLIFA), like other immunoassays that are used to measure antibiotic concentrations, is based on competitive inhibition of the label of conjugated drug by drug that is present in the specimen. In the SLIFA procedure, the fluorescent moiety umbelliferone is generated from β-galactosylumbelliferone that is covalently bound to the antibiotic. In its usual state, the labeled antibiotic does not fluoresce. However, if the antibiotic–substrate reagent is cleaved by β-galactosidase, umbelliferone is released. When the β-galactosylumbelliferone–drug conjugate binds to the antidrug antibody, the conjugate is prevented from interacting with the enzyme. As in any competitive binding assay, the free antibiotic in serum sample competes with the conjugate for binding sites on the antibodies. The amount of conjugate available for the reaction is, therefore, directly related to the amount of free drug in the serum sample. In the SLIFA, the rate of increase in the intensity of fluorescence is proportional to the amount of antibiotic in the sample.

The reaction to label the aminoglycoside class of antibiotics with β-galactosylumbelliferone is a carbodiimide procedure. The reaction sequence proceeds in the same manner with all aminoglycoside/aminocyclitol antibiotics. For the aminoglycoside amikacin, the β-galactosylumbelliferone reaction is performed by adding 50 mg of the potassium salt of β-[7-(3-carboxycoumarinoxyl)]-β-galactoside to 171 mg of amikacin sulfate in 2 mL of water. The pH is adjusted to 3.8 and the mixture is cooled to 0°C (in an ice bath). Thirty milligrams of 1-ethyl-[3-(3-dimethylaminopropyl)] carbodiimide hydrochloride are added.

After 2 hours, the mixture is chromatographed at 25°C on a 2.5 × 50-cm column of CM-Sephadex C-25 (Sigma-Aldrich, St. Louis, MO). The effluent is monitored at 345 nm. The column is washed with 200 mL of 50 mmol/L ammonium formate to elute the unreacted β-galactosylumbelliferone-amikacin. A linear gradient is formed with 400 mL of 50 mmol/L and 400 mL of 1.8 mol/L ammonium formate solution and applied to the column. A peak of material is eluted at a concentration of approximately 1.4 mol/L ammonium formate. The column is washed with 600 mL of 1.8 mol/L ammonium formate. The carbodiimide reaction appears to lead to the formation of amide bonds between the carboxylic acid of 1β-[7-(3-carboxycoumarinoxyl)] galactoside and the primary amino groups of the aminoglycoside antibiotic.


In the past, SLIFA kits have been commercially available for the assay of aminoglycoside/aminocyclitol antibiotics, including gentamicin, tobramycin, and amikacin (195,196). In addition, noncommercial antibiotic SLIFA kits have been produced for other aminoglycoside/aminocyclitol antibiotics. The procedure for the assay of amikacin in human serum or plasma is identical to other aminoglycoside/aminocyclitol assays. Only the absolute amounts of some of the reactants differ.


The SLIFA may be performed with any fluorescence spectrophotometer. Most commonly, either an Aminco-Bowman spectrophotofluorometer (American Instrument Company, Silver Spring, MD) or an Ames fluorocolorimeter (Miles Laboratories, Elkhart, IN) has been employed. The Aminco instrument is set for excitation at 400 nm and emission at 450 nm. The Ames instrument requires a 450 nm, narrow bandpass, interference filter for excitation, and a glass 5–56 (blue) filter on top of a glass 3–73 (yellow) filter for emission of light. The described SLIFA is not completely hands off in the sense that there are no microprocessors and flow-through apparatus available to interpret the data. Accessory equipment includes (a) disposable fluorescence polystyrene cuvettes (Evergreen Scientific, Los Angeles, CA), (b) a pipetter/diluter equipped with a 250 µL reagent syringe and a 2.5 mL buffer syringe, (c) an accurate timer, and (d) 13 × 100-mm test tubes. It should be noted that aminoglycoside/aminocyclitol antibiotics may adsorb from dilute solutions onto glass. Therefore, plastic test tubes should always be used. (This physicochemical guideline applies to all assays, not just the SLIFA.)


The antibody–enzyme reagent is composed of 1.5 units of β-galactosidase and antiserum-to-amikacin in 50 mmol/L bicine/0.1% sodium azide buffer (Worthington Biochemical, Inc, Freehold, NJ). Bicine buffer ([NN-bis[2-hydroxyethyl]glysine) (grade A; Calbiochem, La Jolla, CA) is used at pH 8.5. The enzyme should be standardized at 25°C in 3-mL bicine buffer, pH 8.5, containing 3 mmol/L o-nitrophenyl-β-D-galactoside. The molar extinction coefficient for the product of this reaction, o-nitrophenyl, is 4.27 at 415 nm. One unit of enzyme activity hydrolyzes 1.0 mmol of substrate per minute. The antiserum has been commercially available (197). The antiserum must be of potency to inhibit the fluorescence to 10% to 20% of that in the absence of antiserum.

The antibiotic–drug conjugate consists of β-galacto-sylumbelliferyl-amikacin (0.007 absorbance units at 343 nm) in 5 mmol/L formate with 0.1% sodium azide buffer, pH 3.5. Each reaction mixture must contain 0.00035 absorbance units at 343 nm.

Standard amikacin concentrations should be made in normal human serum in the range of 0, 10, 20, 30, and 40 µg/mL amikacin. The range adjustment solution consists of a mixture of 7-hydroxycoumarin-3-(N-hydroxyethyl) carboxamide in 5 mmol/L formate, pH 3.5, with a final concentration of 0.1% sodium azide buffer. When 50 µL of the range adjustment solution is mixed with 1.5 mL of bicine buffer, the fluorescence intensity must be comparable to that absorbed with the high calibration samples at the end of a 20-minute incubation. The fluorescent antibiotic reagent must absorb maximally at 343 nm. However, on hydrolysis by β-galactosidase, the absorbance at 343 nm must decrease and a new maximum must appear at 405 nm. The absorbance of the enzyme–antibiotic conjugate at 405 nm after hydrolysis should be 1.6 times that at 343 nm before hydrolysis.


After reconstitution, all components can be stored at 2°C to 3°C for up to 10 weeks. All components must be at room temperature before the assay is performed. The assay proceeds as follows.

 1.  Using a pipette/diluter, 1:50 dilutions of the calibrators and samples are made by diluting 50 µL of each specimen with 2.5 mL of diluted bicine buffer.

 2.  Fifty microliters of the antibody–enzyme reagent and 500 µL of buffer are dispensed, using the pipette/diluter, into the reaction cuvettes that are required for duplicate determinations of the calibrators and samples.

 3.  Fifty microliters of each of the diluted calibrators and samples is mixed with 500 µL of buffer and added to the reaction cuvettes.

 4.  Starting in the same calibrator sequence, the reactions are initiated at timed intervals by dispensing 50 µL of the amikacin fluorogenic reactant and 500 µL of buffer to each cuvette. The reactants must be thoroughly mixed.

 5.  After approximately 19 minutes, the reactions are read in sequence by zeroing the instrument with buffer and reading the first specimen in the fluorometer after adjusting the fluorescence reading to 90% of full scale.

 6.  At 20 minutes, the fluorescence intensity of the highest calibrator is read and recorded. Then the fluorescence intensity of the remaining cuvettes is read and recorded in the same sequence as that in which the reactions were initiated.

 7.  The concentration of amikacin in the assay is determined by constructing a standard line by plotting the average fluorescence readings for duplicate calibrators versus the amikacin concentrations.

The SLIFA performs comparably with other radiometric and nonisotopic immunoassays. Table 8.12 presents the salient performance characteristics of the assay for amikacin. As demonstrated in Figure 8.12, kanamycin and tobramycin cross-react at 2.4% and 0.2%, respectively, changing the response to the maximum dose by 50%. Other aminoglycoside/aminocyclitol antibiotics, including gentamicin, netilmicin, sisomicin, and streptomycin, do not significantly interfere with the assay. Additional antibiotics that have been tested, including carbenicillin, cephalothin, CAM, erythromycin, methicillin, and tetracycline, do not appear to affect the assay (198).

The assay of amikacin by the SLIFA technique is accurate from 5 to 50 µg/mL. The accuracy of the SLIFA is affected by the same factors as other immunoassays of antibiotics (198).

Cycling Enzyme Assay

Cycling procedures are inherently appealing because they do not consume reagents during the course of reactions and can, therefore, use the same reagents for extended periods of time. Historically, the coenzymes NAD+ and flavin adenine dinucleotide have been most frequently used as reaction cofactors. The assays have traditionally suffered from a lack of suitable detection systems for biologic samples. Cycling procedures have been described that use alcohol dehydrogenase and malate dehydrogenase as reagents. These assays are of limited use with biologic specimens because significant interference is generally present during the enzymatic cycling steps and in the final fluorometric measurements. An enzymatic cycling procedure with two irreversible reactions that are catalyzed by NAD+ peroxidase and glucose-6-phosphate dehydrogenase has been devised. This cycling procedure is not adversely affected in an appreciable way by biologic fluids.

The reaction has been designed to proceed as follows:

 1.  Gentamicin + (NAD+-gentamicin) + antibody → gentamicin-antibody + (NAD+-gentamicin)-antibody.

 2.  (NAD+-gentamicin) + glucose-6-phosphate + glucose-6-phosphate dehydrogenase → (NADH-gentamicin) + H+ + 6-phosphogluconate (cycling step).

 3.  6-Phosphogluconate + NADP + glucose-6-phosphate dehydrogenase → NADPH + H+ + ribulose-5-phosphate + CO2 (indicator reaction).


A Perkin-Elmer model 555 UV/visible spectrophotometer (Perkin Elmer, Norwalk, CT) has been used for determining absorbance. This gentamicin assay has been performed with an Eppendorf model 1101M spectrophotometer (Eppendorf, Hamburg, Germany) that is equipped with a 334-nm filter.


NAD+ peroxidase (EC, glucose-6-phosphate dehydrogenase (EC 11.1.49), NAD phosphate (NADP+) 1-oxoreductase from L. mesenteroides, 6-phosphogluconate dehydrogenase (EC, NADP+ 2-oxoreductase, and β-D-glucose-6-phosphate crystallized monosodium salt were all obtained from Boehringer-Mannheim (Mannheim, Germany). Antigentamicin antibodies were obtained from Atlantic Antibodies (Scarborough, ME).


For synthesis of NAD+-labeled gentamicin, 5 g of gentamicin sulfate is dissolved in 15 mL of water. The pH of the solution is adjusted to 11.5 with 5 mol/L NaOH. The mixture is lyophilized and extracted in 2 L of boiling methanol. Free gentamicin base is dried in a vacuum. Gentamicin is conjugated to NAD+ after the coenzyme has been converted to N6-(2-carboxyethyl)-NAD+. One gram of gentamicin base is dissolved in 5 mL of water, with the pH adjusted to 7.0 using 6 mol/L HCl. One hundred milligrams of N6-(2-carboxyethyl)-NAD+ and 500 mg of 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide are dissolved in 2 mL of water. The pH is kept constant at 4.7 for a 3-hour incubation period. After incubation, the pH is adjusted to 7.0 and the mixture is applied to a Dowex 1-X2 column (Sigma-Aldrich, St. Louis, MO) (chloride form, 2.6 × 40 cm) and eluted with water (100 mL/hour). The UV-absorbing fraction is concentrated on a rotary evaporator. The mixture is desalted on a Sephadex G-15 column (Sigma-Aldrich, St. Louis, MO) (2.6 × 60 cm), with water as the eluent. The UV-absorbing material is applied to a BioRex 70 column (Bio-Rad Laboratories, Richmond, CA) (2.6 × 40 cm) that has been previously washed with 20 mmol/L ammonium acetate. The void volume is washed out at 100 mL/hour with 60 mmol/L ammonium acetate. Elution is accomplished with a linear gradient of ammonium acetate (20 to 50 mmol/L) over a period of 4 hours. The UV-absorbing fraction contains both NAD+ and gentamicin.

The fraction that contains both NAD+ and gentamicin is concentrated and fractionated on a Bio-Gel P4 column (Bio-Rad Laboratories, Richmond, CA) (2.6 × 60 cm) with water flowing at 40 mL/hour. After volume reduction to 10 mL, the two NADH and gentamicin peaks are stored at −30°C. These two peaks are analyzed for NAD+ content using ethanol and alcohol dehydrogenase. Peak 1 has a molar absorptivity at 265 nm of 24.1 × 103/mol/L/cm in the oxidized form and 20.7 × 103/mol/L/cm in the reduced form. Peak 2 has absorptivity of 2.9 × 103/mol/L/cm in the oxidized form and 19.2 × 103/mol/L/cm in the reduced form. Peak 1 has a gentamicin content by RIA of 1.45 µg/mL and peak 2 has a gentamicin content of 0.99 µg/mL.

Fifty microliters of sample is added to 50 µL of the ice-cooled cycling reagent that contains 0.2 mol/L Tris, 0.28 mol/L potassium acetate, 12 mmol/L β-D-glucose-6-phosphate, 12 mmol/L hydrogen peroxide, 200 µg/mL NAD+peroxidase, and 200 µg/mL glucose-6-phosphate dehydrogenase, pH 8.5. The solution is mixed for 2 hours at 30°C. After cooling, 0.5 mL of 20 mmol/L Tris, 30 mmol/L ammonium acetate, 0.1 mmol/L ethylenediaminetetraacetate, 0.2 g/L BSA, 0.6 mmol/L NADP+, and 10 µg/mL 6-phosphogluconate dehydrogenase, pH 7.7 (the spectrophotometric indicator reagent), is added. The absorbance at 334 nm is recorded after the mixture is incubated for 30 minutes at 25°C.


Figure 8.13 demonstrates the inhibition of NAD+-gentamicin peak 2 by antigentamicin antiserum at various gentamicin levels. A standard curve for gentamicin was calculated from these values and is displayed in Figure 8.14. The absorbance values for gentamicin standards and patient specimens are measured at 334 nm, and after subtracting blank absorbance values, related to the gentamicin concentrations.


The spectrophotometric enzyme cycling procedure is able to detect 5 ng of gentamicin (0.1 µg/mL). The method is equal to other nonisotopic immunoassays and RIA in the range of 1 to 10 µg/mL. The intraassay coefficient of variation is 6.1% with a mean at 7.4 µg/mL. The intraassay coefficient of variation is 3.4% at a concentration of 13.1 µg/mL. The specificity and performance of the spectrophotometric cycling enzyme assay are equal to those of other nonisotopic immunoassays and the standard RIA method. In addition, reagents are not rapidly consumed and measurements may be made on a simple spectrophotometer (199).


Like the other nonradioactive immunoassay techniques such as SLIFA and EMIT, the FPIA method was developed as a technology to assay small molecules. The theoretical basis for FPIA was described in the early 1970s by Dandliker et al. (200). Subsequently, Abbott Laboratories (Abbott Park, IL) accelerated the entry of this technology into clinical laboratories by developing an automated fluorescence polarization analyzer system (201).

A molecule naturally gravitates toward its lowest energy state but becomes excited after exposure to light. This excitation is the result of electrons moving from a lower energy shell to a higher energy shell. The leap to higher energy levels, governed by the laws of quantum mechanics, is temporary. The electron naturally returns to its original energy shell. The amount of energy gained during the first jump to a higher energy shell is greater than the amount of energy that the molecule loses when the electron returns to its normal orbit. The wavelength of light that is emitted during the return trip (emitted light) is longer than that which is absorbed during the primary jump (excitation light) and is measured as fluorescence. Ordinary (white) light contains a spectrum of wavelengths (white light). The electrical vectors of light waves that are produced by standard sources are randomly oriented. Excitation sources that are used in fluorometers can emit light at one, several, or a range of wavelengths of interest, depending on the applications for which the instrument is used. The intensity of the emitted fluorescent light is related to the excitation intensity of the initially absorbed light at the absorption wavelength.

Plane-polarized light is generated when light is passed through crystalline materials known as “polarizers.” A polarizer orients the electrical vectors of incoming light waves in a single plane. Fluorescence polarization occurs when fluorophores that have been excited by polarized light also emit polarized light (366). This occurs when the Brownian movement (rotational relaxation) of the fluorophores under analysis is slower than their fluorescence decay times. Small molecules rotate rapidly, such that their rotational relaxation times are much shorter than their fluorescence decay times. Consequently, small fluorophores emit depolarized fluorescent light while large fluorophores emit polarized fluorescent light (202).

In developing the FPIA technique, the fluorescence decay times and rotational relaxation times of the molecules under study needed to be considered. The fluorescence decay time of the molecule is the time interval from the moment it is struck by polarized light until it releases its emitted light (203). The rotational relaxation time describes the molecule’s Brownian motion and is the time an oriented molecule takes to leave alignment after an incoming burst of polarized light. To effectively determine analyte concentrations with an FPIA assay that uses routine equipment, the rotational relaxation time of the molecule under analysis must be 1 nanosecond or less. This limits the use of the technique to the analysis of small molecules. For example, immunoglobulins have rotational relaxation times of approximately 100 nanoseconds.

The fluorescent light intensity that is produced by the sample is measured both parallel to and perpendicular to the plane of the polarized excitation light. The fluorescence polarization is measured by the following formula:


where IPAR (parallel) is the intensity of fluorescence parallel to the plane of the excitation light and IPERP (perpendicular) is the intensity of fluorescence perpendicular to the plane of the excitation light.

In practice, the antibiotic is labeled with fluorescein. When the antibiotic binds to a specific antibody, the rotational relaxation time of the large antigen-antibody complex is increased so that it exceeds the fluorescence decay time of the label. The concentrations of antibody and fluorescent-labeled antigen are kept constant, with the only variable being the concentration of the antibiotic (unlabeled antigen) in the specimen. The higher the concentration of free antibiotic in the sample, the more this antigen limits binding of the labeled antibiotic to antibody, resulting in a proportional decrease in fluorescence polarization (204208). Conversely, as the concentration of unlabeled antibiotic decreases, the amount of fluorescence polarization proportionately increases.

Fluorescence Immunoassay Method


A specially designed fluorometer with microprocessor controls is available from Abbott Laboratories (Abbott Park, IL). The instrument and specimen flow pathways are described in the following texts.

The filament of a General Electric EFM 50W tungsten/halogen lamp (Fairfield, CT) is focused through a 3-mm diameter entrance aperture. After passing through heat-absorbing glass (BD-38; Corion Corp, Holliston, MA), the light is collimated. All lenses in the system are Melles Griot plano-convex lenses (Albuquerque, NM) that have a focal length of 18 mm and a diameter of 15 mm. The collimated light is directed through a 10-nm bandwidth, 485-nm center wavelength, excitation filter (Corion Corp, Holliston, MA). Light reflected from the transparent glass beam splitter (Corning coverglass no. 1, Corning, NY) is focused onto a UV 215 B reference silicon detector (EG & G, Salem, MA). The reference detector signal is used to monitor the intensity of the lamp. The light transmitted through the beam splitter passes through an HN38S polarizer/crystalloid transmission–type liquid crystal combination (Polaroid, Cambridge, MA). This beam splitter also serves to rotate the plane of polarization. The excitation light is directed onto the sample, which is contained in a 12 × 75-mm glass tube. The emitted light is collimated through a lens, passed through a 10-nm band with a 525-nm interference filter, and then directed through a vertical HN38S polarizer. The emitted light is focused onto a 3 × 8-mm aperture and R928 photomultiplier tube (Hamamatsu Corp, Middlesex, NJ). The excitation optics and the emission optics are perpendicular. The polarizer/liquid crystal combination is rotated in an electric field applied to the liquid crystal. When voltage is directed through the liquid crystal, no rotation occurs and horizontal light passes through the sample. When there is no voltage, the liquid crystal rotates by 90% and the sample is excited by vertical light (209,210).

Measurements are made as directed by the microprocessor through emission fluorescence intensity measured by the photomultiplier tube. The gain of the photomultiplier tube is controlled by a model PMT-20 A/N high-voltage power supply (Bertran Associates, Syosset, NY). Both the excitation and fluorescent polarization intensities are captured in the vertical and horizontal modes and converted from voltages to frequencies, which are measured by a counter timer. Intensity values are presented as the ratio of the frequency channel counts to the reference clock channel counts (211).


The fluorescein conjugate of amikacin is prepared by reacting the antibiotic with 5-[(4,6-dichlorotriazinyl) amino] fluorescein. The fluorescein compound is dissolved in methanol and added to amikacin in water at pH 9.0. The final concentration of the fluorescein reagent is 16 mol/L and that of amikacin is 160 mol/L. After 1 hour at room temperature, the mixture is applied to a diethylaminoethyl-cellulose chromatography column with 0.1 mol/L phosphate buffer, pH 8, as the eluent.

The amikacin–fluorescein conjugate is diluted in 0.1 mol/L Tris (hydroxymethyl) methylamine buffer, pH 7.5, that contains 1.212 g/L sodium dodecyl sulfate, 0.1 g/L bovine γ-globulin, and 0.1 g/L sodium azide. The concentration of the conjugate is approximately 100 nmol/L. Antiserum is diluted in a phosphate buffer, pH 7.5, that contains 0.1 g/L bovine γ-globulin, 0.1 g/L sodium azide, and 50 mg/L benzalkonium chloride. Standards and controls of amikacin are made as described for the SLIFA procedure.


Amikacin is assayed by transferring 20 µL of sample or control into 1 mL of buffer. This step is repeated. After 20 µL of the diluted sample is dispensed into a 12 × 75-mm disposable culture tube, 200 µL of dilution buffer is added. To this solution, 40 µL of conjugate is added, followed by 700 µL of buffer. Finally, 40 µL of antiserum solution is added and dispensed into the reaction vessel, followed by 1.0 mL of buffer.


Table 8.13 presents the salient characteristics of the FPIA for amikacin. The following antibiotics cross-react at a rate of less than 0.1% in the FPIA of amikacin: ampicillin, amphotericin, carbenicillin, cefamandole, cephalexin, cephaloglycin, cephaloridine, cephalothin, CAM, CLD, erythromycin, ethacrynic acid, 5-fluorocytosine, fortimicin A, fortimicin B, furosemide, fusidic acid, lincomycin, methicillin, methotrexate, oxytetracycline, penicillin, rifampin, sulfadiazine, sulfamethoxazole, tetracycline, ticarcillin, TMP, and vancomycin.

Table 8.14 presents the cross-reactivity of closely related compounds with the FPIAs of the aminoglycosides gentamicin, tobramycin, and amikacin. Figure 8.15 compares the FPIA results for amikacin with those of RIA, bioassay, and SLIFA. Discrepancies between the slopes for the different assays are most likely attributable to the sources of the standards or their preparation and storage. Coefficients of variation between the immunoassays were 6.96% or greater. Results seen here with amikacin are also applicable to the aminoglycosides gentamicin and tobramycin (205,212).

Micromethod Fluorescence Polarization Immunoassay of Aminoglycosides

Gentamicin and netilmicin are aminoglycoside antibiotics that are used to treat gram-negative bacterial infections. Monitoring the concentrations of these drugs, usually in venous blood samples, helps optimize care by limiting the incidence of toxicity and therapeutic failure. However, venous samples are often difficult to obtain from infants and children. Capillary blood spotted on filter paper has been used successfully for detection of inborn errors of metabolism and for monitoring drug concentrations. For example, this technique has been used with an HPLC assay that has been previously described in this chapter to detect netilmicin and sisomicin. The following method is an FPIA that determines the gentamicin and netilmicin concentrations from blood samples spotted on filter paper.


One hundred microliters of the standard aqueous solutions (0 to 20 µg/mL prepared in distilled water) and the gentamicin and netilmicin blood standards (0 to 20 µg/mL prepared with antibiotic-free pooled blood) are spotted onto filter paper. The spots are dried at 50°C for 10 minutes in an air-circulating oven or at room temperature for 5 hours. Using scissors, the blood-containing area is cut into five or six pieces and all are placed into one tube. Addition of 500 µL of warmed (35°C) 0.5 mol/L Na2HPO4 buffer to the tube is followed by incubation of the sample in an oven (35°C) for 60 minutes and centrifugation for 15 minutes (3,000 × g). The colorless clear filtrate is transferred to a well of a TDx cartridge (Abbott Laboratories, Abbott Park, IL) for measurement by FPIA.

The measurement of hemoglobin in extracts can be performed using a hemoglobin assay kit (Wako Pure Chemical Industries, Osaka, Japan). Twenty microliters of each extracted sample is mixed with 5 mL of cyanmethemoglobin reagent. After 5 minutes, the absorbance is measured at 540 nm and compared with that of the standard solution containing 3.58 of cyanmethemoglobin/5 mL.


Hemoglobin levels below 8.6 g/L do not affect the FPIA gentamicin assay. Ultrafiltration of a gentamicin-free sample with a hemoglobin level greater than 8.6 g/L gives a clear, colorless filtrate, demonstrating that the hemoglobin in the filtrate is almost completely removed by the ultrafiltration procedure. Maximum elution time from the spotted paper for both gentamicin and netilmicin is 60 to 90 minutes under the given extraction conditions. Recovery of gentamicin from dried blood-spotted samples in the concentration range of 1.5 to 20 µg/mL is from 92% to 115%. The lower limit of detection (LOD) of gentamicin or netilmicin with the FPIA reagent kits is 1 µg/mL each for samples spotted with 100 µL of whole blood.

The calibration curve for gentamicin or netilmicin in dried blood spots is linear over the concentration range of 1.0 to 20 µg/mL. The intraassay coefficients of variation are 5.6% and 6.3% for gentamicin concentrations of 5 and 10 µg/mL, and 4.6% and 1.6% for netilmicin concentrations of 10 and 20 µg/mL, respectively. The interassay coefficient of variation is 4.5% for a gentamicin concentration of 5 µg/mL and 9.7% for a netilmicin concentration of 10 µg/mL. Clinical samples from pediatric patients treated with netilmicin show excellent linear correlation when corrected for hematocrit levels. The applicability of the determination of gentamicin or netilmicin concentrations in dried blood spots by FPIA is limited because the quantification limit exceeds the effective range of the antibiotics (204).


Teicoplanin levels in serum can be determined by FPIA using fluorescein-labeled teicoplanin, which competes with unlabeled teicoplanin for antibody (213).


For the generation of the calibration curve, standards containing 0, 5.0, 10.0, 25.0, 50.0, and 100.0 µg/mL teicoplanin are prepared. The International Bioclinical reagent system is used to prepare standards and serum samples for measurement by the American Bioclinical FP analyzer (Upland, CA). This instrument calculates the millipolarization values and extrapolates the teicoplanin concentration for each sample by comparing the values to those of the calibration curve.


The lower LOD of teicoplanin is 0.5 µg/mL. Other antibiotics that interfere in this assay are shown in Table 8.15. The results of the FPIA agree well with those of bioassay, with a correlation coefficient of 0.901.


The general advantages and disadvantages of the RIA technique are shown in Table 8.2 (214,215) (Fig. 8.16). Because it is no longer in routine clinical use, it is not discussed further in this chapter.


Chromatography uses the ability of a compound in one type of medium to be selectively removed from that medium onto an adsorbent and to be quantitatively assayed. Most commonly, the compound is in a liquid and is adsorbed onto a solid (liquid chromatography) or is in the gas phase and is adsorbed onto a liquid (GLC). Inherent in any chromatographic method is the requirement for specialized adsorbent columns (which may be different for different antibiotics), the use of expensive equipment (especially in GLC), the use of various extraction procedures, and the requirement for a skilled technician to perform the assay.

The advantages of the procedure are that chromatography can separate even closely related compounds and analyze them separately. Generally, the method is quite sensitive, that is, able to quantitate between 1.0 and 0.5 µg/mL for most antibiotics and is quite rapid, requiring 30 to 60 minutes per specimen. The principles of chromatography have evolved from the early decades of the 20th century, with the most exciting applications occurring within the last 40 years.

Modern chromatography dates to 1941 when Martin and Synge (216) published their classic paper on liquid–liquid partition chromatography. They described individual rates of migration of substances that were a consequence of differing partition ratios between mobile and stationary liquid phases. Gas chromatography (GC) and thin-layer chromatography (TLC) appeared in the 1950s and were received enthusiastically. However, the time-consuming, crudely quantitative characteristics of TLC and the requirements of molecule volatility and thermostability for GC are disadvantages of these techniques. Building on chromatographic principles that evolved over five or more decades and experience gained from GC and TLC in particular, researchers developed HPLC in the late 1960s. The debut of HPLC was largely from the pioneering work of Horvath and Lipsky (217).

Although homogeneous and heterogeneous enzyme FPIA techniques may be viewed as competitive with HPLC, a more global view of the subject reminds us that each method has its own strengths and weaknesses and that the fields of therapeutic monitoring and pharmacokinetic assessment of antibiotics are strengthened by the appropriate application of each of these technologies. The advantages of HPLC are many. These will be amplified as improvements in instrumentation, packing materials, and methods are made. As with GC, mass spectroscopy (MS) is now being applied to HPLC (218). GC-MS will undoubtedly remain the method of choice for the most accurate identification and most sensitive quantitation of small, volatile, thermostable molecules. However, for many larger, nonvolatile, thermosensitive molecules, HPLC-MS has major potential as an analytic method. In this technique, HPLC is a preparatory step prior to MS, and allows the identification and quantification of molecules in the picogram range. Several papers have demonstrated the application of HPLC to MS and have presented instrument modifications to reduce MS contamination (219,220).


HPLC is a reference method, if not the major reference method, by which all new antimicrobial agents are studied, from discovery to clinical application. It plays a major role in drug preparation, pharmacologic assessment, and clinical monitoring (50,64,91,216,221). Thus, an understanding of the analysis of antimicrobial agents by HPLC is essential in clinical laboratory practice.

HPLC as a technology has assumed a central role in the analysis of all water-soluble molecules in clinical laboratories. This technique is the method of choice for the analysis of penicillin and cephalosporin antibiotics. HPLC is especially useful clinically in the measurement of third-generation cephalosporins in the nonblood body compartments (222). It is also being used more frequently to measure parent antibiotics and their metabolites. This section discusses this powerful tool and provides specific examples of how it is used to determine concentrations of most of the major classes of antibiotics in use today.

Many antibiotics have been analyzed successfully by HPLC, including aminoglycosides (223226), cephalosporins and penicillins (227230), tetracyclines (231,232), trimethoprim-sulfamethoxazole (371), and others (44,233,234). The interested reader is referred to reviews by Nilsson-Ehle (235) and Jehl et al. (236) for discussions of the general method and its application to the analysis of drugs.

A number of factors stimulated the application of HPLC, a technique that was already in common use in science and industry, to the quantitation of antibiotics in biologic fluids (237239). This technology was commonly used in modern laboratories to measure antiarrhythmic and anticonvulsant agents. The equipment and expertise were thus available. Several HPLC methods, using standard chromatographic equipment, have been developed that allow the simultaneous quantitation of multiple, closely related antibiotics (240243). Others permit simultaneous quantitation of parent drugs and some metabolites (9,48,71,244250). Simultaneous analysis of mixtures of antibiotics can be performed with small changes in operating conditions (251). Once the equipment has been obtained, the cost of quantifying antibiotics by HPLC is competitive with other methods (252256). Finally, the rapidity with which assays may be performed by HPLC is crucial for seriously ill patients who are receiving drugs with narrow toxic/therapeutic ratios or that have potentially severe side effects (257). HPLC methods can detect quantities of antibiotics as low as 500 ng/L, reflecting levels of sensitivity beyond what is needed in clinical practice (251).

Using the HPLC technique, highly reproducible, difficult separations are achieved rapidly and quantitatively (55,258260). The majority of HPLC separations are accomplished in less than 1 hour, with many requiring only a few minutes. A wide variety of molecular species may be separated and quantitated by HPLC, including both large and small molecules, ions, isomers, polymers, polar molecules, and nonpolar molecules (261263). Because HPLC is often performed at low or ambient temperatures, thermosensitive molecules are easily handled (264266). After the analyte molecules pass the instrument detector, they may be collected quantitatively in unaltered form for further purification or investigation.

Prior to reviewing the practical application of the liquid–liquid partition principle in HPLC, let us first briefly consider its theoretical basis. All HPLC methods use the same basic steps:

 1.  Extraction of the drug with a specific solvent

 2.  Separation of the drug on the solid phase by HPLC

 3.  Detection of the effluent from the solid phase by spectrometry

 4.  Quantitation of the amount of antibiotic present by peak height analysis or peak area analysis

The aim of chromatography in general is the resolution or separation of different molecular species. To understand resolution in the context of HPLC, we must consider a few explanatory equations. These equations and a more in-depth discussion may be found in an excellent concise paper by Guiochon (267) or in the work of Giddings (268). First, let us assume that the groups of molecules eluting from the chromatographic column or peaks on the chromatogram are Gaussian in distribution (not entirely true but adequate for mathematical interpretation). The resolution (R) of two Gaussian peaks is defined by the equation

R = 2(tR2 − tR1)/(W1 + W2),

where tR1 and tR2 are the retention times of the first and second peaks, respectively, and W1 and W2 are the widths at the base of the chromatographic peaks. If the two peaks are close, the resolution is defined by the equation

R = (sq rt. [N/4)[(α − 1)/α] [k′2/(1 + k′2)].

To understand this equation, let us look at its component factors. First, remember that the width at the base of a Gaussian distribution is equal to 4 SD (σ). N, or the plate number of the column (a standard measure of its efficiency), is defined as

N = (16tR/W)2 = (tRσt),

where σt is the SD of the peak in units of time and tR and W are the retention time and base width of the peak, respectively. The relative retention of the two compounds being considered, σ, is described by the equation

α = (tR2 − t0)/(tR1 − t0) = t′R2/t′R1 = k′2/k′1,

where t0 is the retention of an inert molecular species that theoretically is not retained by the column, t′R1 and t′R2 are the adjusted retention times, and k′ is the capacity factor of the column (how retentive it is in terms of these particular molecular species). The capacity factor, k, for each molecular species, may be redefined as

k = (tR − t0)/t0 = r′R/t0

This factor, k, is important because it is proportional to the equilibrium constant that describes the distribution of molecules between the chromatographic phases.

Applying these formulas to HPLC, the mobile solvent phase passes over the stationary phase at a constant rate. The two phases possess different chemical polarities. As the analyte molecules in the mobile phase pass over the stationary phase, those with polarity closer to that of the stationary phase are retained selectively for a time on the column. Conversely, the analyte molecules with polarity closer to that of the mobile phase tend to remain in the mobile phase, passing through the column faster. Passing through the instrument monitor sequentially, these groups of molecules give rise to peaks on the chromatogram (Fig. 8.17).

The term high-performance liquid chromatography encompasses several different types of chromatography. Systems that have a polar stationary phase with a nonpolar mobile phase are termed normal phaseor straight phase, while those having a nonpolar stationary phase with a polar mobile phase are termed reverse phase. A popular variant of RP-HPLC is ion-pair chromatography, where the polarity (and thus the retention) of the analyte is changed by adding a second ion of opposite charge (counter-ion). Ion-exchange chromatography is based on competition between the analyte ion and a second, similarly charged molecule for oppositely charged sites on the exchange resin. An early form of HPLC using a solid stationary phase that separates molecules on the basis of differing size and weight is termed size-exclusion or gel-permeation chromatography. Gel-permeation chromatography has wide industrial and research applications but is rarely used for antimicrobial determinations in clinical laboratories.

Much of the versatility of HPLC is because of the wide variety of instrument components and chromatographic conditions from which to choose. In its simplest form (Fig. 8.18), the HPLC system is a closed system composed of an injector, pump, chromatographic column, and detector, with a reservoir for the mobile phase liquid. The composition of the mobile phase selected depends on the physical characteristics of the analyte, the complexity of the sample mixture, and the column packing employed. Although initially the mobile phase was usually nonpolar (straight phase), most recent applications use a buffered aqueous mobile phase containing acetonitrile, methanol, or some other relatively polar organic (reverse-phase) compound. For an extensive review of solvent phase selection, the reader is referred to the work of Snyder and Kirkland (269).

The injector, either manual or automatic, is a simple but very important component that allows the sample to be placed into the mobile phase stream without disrupting the flow. The HPLC pump is critical in that it must present the mobile phase liquid to the chromatographic column at a constant pressure (usually <2,000 psi) to provide acceptable and reproducible analyte separations in a timely fashion. Pumps are available over a broad price range, with considerable variations in design that serve mainly to minimize pressure fluctuations.

Progress in the field of HPLC has been made through improvements in the chromatographic packing materials themselves and the consistency with which the columns are packed. High-quality, commercially prepared columns are available that offer improved batch-to-batch reliability and a wide selection of column lengths, efficiencies, types of support phases, particle sizes, shapes, pore sizes, pH ranges, and chemical specificities of the liquid stationary phase (270). The column sleeve is usually made of stainless steel, although the radial compression technology uses polyethylene (271).

In practice, most HPLC systems use a polar mobile phase and nonpolar stationary phase (reverse phase). In order to eliminate false peaks (bubbles) and to remove microparticulate matter, the mobile phase is degassed and filtered. The organic stationary phase is commonly composed of C8 or C18 aliphatic chains bonded chemically to microparticulate porous silica of 5 to 10 µm mean particle diameter, densely and uniformly packed into a stainless steel sleeve. A rule of thumb for column efficiency is that for an efficient column, a single theoretical plate should be two to five times the mean particle diameter of the support phase, with an overall efficiency of 3,000 to 5,000 theoretical plates. The internal diameter of the sleeve is usually 2 to 5 mm, with the column length varying from 3 cm to several meters, although typically, it is less than 30 cm. The rate of flow of the mobile phase is usually 1 to 3 mL per minute, requiring a pressure of up to about 2,000 psi to maintain the flow and to attenuate chromatographic time.

Most current methods are performed at ambient temperature or at up to approximately 50°C. Above this, silica-based packing materials begin to deteriorate. The pH of the mobile phase is kept below 7 to 7.5 for the same reason, although nonsiliceous packing materials are available that work well in the basic region.

The selection of the means by which the HPLC instrument “sees” the analyte molecules as they proceed in groups from the chromatographic column allows one to further tailor the instrument and the analysis for particular needs. Commonly used detectors include UV/visible, fluorescence, refractive index, electrochemical, infrared, and radiometric instruments, and mass spectrometry. UV detectors are most commonly used to monitor the effluent from the column, although fluorescence detectors, because of increased sensitivity and elimination of interfering peaks, are often preferred. State-of-the-art detectors tend to be microprocessor-controlled, with flow cell volumes down to 1 µL. However, less sophisticated detectors with flow cells of 10 µL are adequate for the majority of applications. Typically, 5 to 100 µL of the prepared sample is injected. The chromatographic time is usually less than 20 minutes. Current HPLC systems routinely work in the nanogram range. Between-day coefficients of variation are usually less than 5% to 7%. An internal standard is commonly used to minimize errors inherent in the system, particularly if some form of sample pretreatment is employed. For a concise review of the use of internal standards, the reader is referred to the work of Snyder and Kirkland (269).

Biologic fluids are typically complex mixtures that are composed of a wide variety of proteins, carbohydrates, lipids, and others. Beyond the deleterious effects these materials have on the injector, column packing material, and pump, their presence frequently interferes with the separation and quantitation of the analyte molecules under study. Consequently, some form of sample preparation is almost always required prior to the chromatography. Four major techniques are commonly employed to prepare samples for injection.

In most assays of biologic fluids, including antibiotic assays, it is essential to remove as much protein from the fluid under study as possible. Most directly, one can precipitate the protein, for example, with trichloroacetic acid. However, acid precipitation leaves many interfering materials in the sample and the resultant pH change may be deleterious to the analyte molecules, metabolites, packing material, or other equipment components.

Alternatively, one may remove the protein with an ultrafilter such as those available from Millipore Corporation (Billerica, MA) (272), but not all low molecular weight proteinaceous material can be removed. Other interfering materials may pass through the filter as well. Additionally, analyte is lost if there is significant protein binding. In spite of its limitations, this method has been gaining popularity (273).

Solvent extraction has been the most common preparatory technique. Organic solvents extract specific organic molecules selectively from the specimen by solvent partitioning. In this process, protein in the sample is usually denatured and left at the liquid–liquid interface. By appropriate solvent selection and manipulation of the specimen pH, a satisfactory separation of the analyte from most of the other materials in the specimen can frequently be achieved. For example, piperacillin can be extracted into chloroform/1-pentanol at an acidic pH (274). The organic solvent containing the drug can be removed by evaporation, or the drug can be extracted back into an aqueous solution with a pH of 7, where the drug is ionized and water-soluble. This latter step further cleans the specimen and may be used to concentrate the analyte if a reduced quantity of the aqueous phase is used.

Many compounds may be isolated by first adsorbing them either onto an ion-exchange resin (275) if they are charged or onto a bonded reverse-phase packing material if they are not charged (276), followed by elution. For example, the aminoglycoside/aminocyclitol antibiotics are very polar and difficult to extract with an organic solvent. However, they may be easily adsorbed onto Amberlite resin (The Dow Chemical Company, Midland, MI) (275) or silicic acid (12), eluted, and analyzed.

The advantages of HPLC—rapid availability of results, sensitivity, specificity, and the ability to measure several drugs and their metabolites simultaneously—allow for easy therapeutic monitoring of antibiotics. At the same time, they enhance antibiotic development and pharmacokinetic evaluation. Nearly all of the antibiotics in use today have been assessed by HPLC. Concurrent with improvements in HPLC equipment and packing materials, methods have been developed for the quantitation of certain antibiotics by EMIT, RIA, fluorescence assays, FPIA, and other nonisotopic immunoassays. When a choice of technique is possible, some factors that should be considered before a selection is made include the equipment and expertise that is available within the laboratory, the positive and negative attributes of the specific methods under consideration, the laboratory’s test volumes and costs, and the institution’s clinical requirements (56,221,258,277293).


At present, CAM is probably the antibiotic most commonly quantified by HPLC in clinical laboratories. Several relatively simple methods are available that give sensitive and reproducible results rapidly and require only small quantities of specimen. Additionally, CAM and its succinate or palmitate prodrug esters may be measured simultaneously. HPLC conditions for their measurement are similar to those employed for quantitation of theophylline, acetaminophen, and some anticonvulsants, thus requiring minimal method modification for the many laboratories already measuring these drugs by HPLC (294). Radioenzymatic assays (158,295) are available to measure CAM, and a modification of these has been compared with HPLC (296). Table 8.16 presents a summary of the HPLC methods available for measuring CAM. The authors have preferred the method of Velagapudi et al. (297) because the procedure employs ethyl acetate extraction, is performed at ambient temperature, is monitored at the absorption maximum of CAM (278 nm), and uses a closely related compound (thiamphenicol) as the internal standard (298). Table 8.16 presents recent useful HPLC methods (299).


Factors to consider when deciding whether or not to monitor blood concentrations of cephalosporin drugs include the potential for toxicity and concerns about patient compliance (for the oral preparations), possible failure to reach or sustain therapeutic concentrations for other reasons, or inappropriate use of expensive medications. Blood concentrations of these drugs may also be monitored for the purposes of pharmacokinetic studies (256). Table 8.16summarizes a number of available HPLC procedures for many of the cephalosporins (239,242,255,300314). Several clinically important cephalosporin antibiotics are reviewed in this chapter.

Oral cephalosporins (cefaclor, cefadroxil, cephalexin, and cephradine) are widely used antibiotics. Cefixime, a newer member of this class of drugs, offers broader pharmacokinetic properties than older cephalosporin drugs, with enhanced gram-negative activity. Cefixime can be used for the treatment of otitis media, urinary tract infections, and respiratory tract infections (370). Initial HPLC methods were limited to the detection of one or two cephalosporins with the same assay system, and in their ability to measure drug concentrations in clinically significant ranges. Method enhancements including modifications of the mobile phase acqueous/organic ratio enabled multiple cephalosporins to be measured with a single assay system (369). The method outlined below uses a single HPLC system to measure five oral cephalosporins in a clinically significant drug concentration range (311).


One Waters HPLC system (Millipore, Waters Division, Milford, MA) consists of a model 590 pump used to deliver the mobile phase and a model 481 variable wavelength detector set at 240 nm. Analysis may be performed using a 4.6-mm × 15-cm Altex Ultrasphere octyl C8 column (5 µm particles; Beckman Instruments, Berkeley, CA) with a silica RCSS Guard Pak precolumn (Waters, Milford, MA). Linear least squares regression analysis is performed using a laboratory automation system (model 3357; Hewlett-Packard, Paramus, NJ).

Chromatographic Conditions

The mobile phase consists of methanol/12.5 mmol/L monobasic sodium phosphate buffer (20:80 by volume), adjusted to pH 2.6 with concentrated phosphoric acid. The mobile phase is filtered, degassed before use, and delivered at ambient temperature at a flow rate of 2 mL/minute. The order of elution for the five cephalosporins is cefadroxil, cefaclor, cefixime, cephalexin, and cephradine, between the times of 3 and 20 minutes.

Five milligrams of cefixime or 5 mg of cephalexin is dissolved in 10 mL of methanol to yield 500 mg/L stock solutions. Each standard is then serially diluted 10-fold. Working standards for the other drugs are prepared similarly, dissolving 5 mg of each compound (cefadroxil, cefaclor, and cephradine) in 10 mL of methanol to give 500 mg/L stock solutions, which are diluted by 10-fold serial dilutions to yield 50 mg/L and 5 mg/L working standards.

Aliquots of the working standard solutions are added to polyethylene microcentrifuge tubes to give the desired drug concentrations. The solvent is then evaporated under a stream of nitrogen. The residue is reconstituted with 0.1 mL of serum, a known volume of internal standard (cefixime for the other four compounds, cephalexin for cefixime), and 0.1 mL of acetonitrile to precipitate serum proteins. Samples are vortex-mixed for 15 seconds and centrifuged at 14,000 × g for 2 minutes. The clear supernatant is evaporated under a stream of nitrogen. The residue is reconstituted with 0.1 mL of the mobile phase, and 50 to 80 µL is delivered by a WISP autoinjector (Waters, Milford, MA) into the chromatograph.


The peak height ratio is linear over the concentration ranges studied (0.1 to 10 mg/L for cefixime and 1 to 100 mg/L for the others). The range for the coefficients of variation for intra- and interday measurements is less than 15%. With the exception of cefaclor, the recovery of each cephalosporin is greater than 81.4% over these concentration ranges. The lower recovery of cefaclor is probably because of its limited solubility in the mobile phase. This becomes a problem only at concentrations of more than 50 mg/L. These drug levels are not normally observed clinically. Other than salicylic acid, which absorbs UV light at 240 nm, commonly administered drugs do not generally interfere with the assay. Moreover, salicylic acid is eluted at 22.1 minutes, not in the same time frame as any of the cephalosporins. The performance data for the cephalosporin assays are shown in Table 8.17.

By using a single system for all five cephalosporins, this HPLC method allows for monitoring without delays due to changes in equipment (311).

Measurement in Body Fluids Other Than Plasma

HPLC- based methods have also been used to determine achievable levels of a antibiotic in body fluids other than blood. For example, clinical trials of the oral cephalosporin cefditoren, which is currently approved in the United States for treatment of lower respiratory tract infections such as acute exacerbations of chronic bronchitis, community-acquired pneumonia, pharyngitis/tonsillitis, and uncomplicated skin-structure infections, included measurement of levels in urine (316) and lung epithelial lining fluid (315) in addition to plasma.

HPLC for Cefditoren Levels in Urine: Modifications to Equipment and Conditions

This HPLC method (316) specifies Nova-Pack C18, 15 cm × 0.39 mm (Waters Corporation, Milford, USA) as the stationary phase, a mobile phase of 75% 0.05 M disodium hydrogen phosphate dehydrated and 25% acetonitrile, adjusted to pH 7.0 by the addition of orthophosphoric acid, and a pump flow rate at 1.0 mL per minute. For cefditoren, the maximum ultraviolet (UV) absorbance is at a wavelength of 295 nm. The integrator was a Maxima 820 Waters data system.

Urine specimens are prepared by mixing 0.5 mL of each sample with 50 µL of an internal standard (aminopyrine, 0.1 mg per mL) plus 900 µL of water. A 25 µL aliquot of the prepared urine sample is injected onto HPLC for analysis. The retention times of ceftidoren and aminopyrine are 5.5 and 11.9 min, respectively. The reported intra- and interday variability (SD/mean × 100) is less than 9.15% for urine samples containing 10, 50 and 100 µg per mL cefditoren, with a limit of quantification of 10 mg per L for this assay method (316).

Liquid Chromatography-Tandem Mass Spectrometry LC-MS/MS for Cefditoren Levels in Lung Epithelial Lining Fluid (ELF)

This method (315) differs from the HPLC methods described above in that mass spectrometry rather than UV absorbance is used to detect the analyte after chromatographic separation, and in that all pipetting and centrifugation steps are carried out at 4°C.

As described earlier in this chapter, lung ELF samples collected by BAL must be centrifuged immediately after collection to remove cells, and the supernatants frozen and stored at −70°C until testing to maximize drug stability. The BAL samples must be concentrated, for example by freeze-drying, and reconstituted in a smaller volume of distilled water, before processing. For this method, the concentration of the internal marker urea is measured in the BAL sample [CBAL(urea)]and in a simultaneously collected serum or plasma sample [Cserum(urea)] to allow determination of a corrected cefditoren concentration in the lung ELF, according to the formula corrected cefditoren level = CBAL(cefditoren) X [Cserum(urea)/CBAL(urea)].


To test for cefditoren, one mL of each reconstituted sample is mixed with 100 µL methanol. A 20 µL aliquot of each sample is chromatographed on a reversed-phase column, eluted with an isocratic (constant concentration rather than a gradient) solvent system, and monitored by liquid chromatography-tandem mass spectrometry (LC-MS/MS) with a selected reaction monitoring (SRM) method, in positive mode, with precursor → product ion for cefditoren, m/z 507 → m/z 241. The precursor → product ion for the internal standard cefotaxime is m/z 456 → m/z 396. Under these conditions, cefditoren and the internal standard cefotaxime are eluted after approximately 1.2 min.

Calibration and inter-assay variation is assessed by testing samples in a range of concentrations (0.00200 mg per L, 0.0200 mg per L, 0.200 mg per L and 0.800 mg per L) prepared from a standard cefditoren solution in buffer in each run. An additional calibration standard such as cefotaxime prepared in a similar concentrations in buffer should be included. No interferences are noted between cefditoren and cefotaxime in this method. Calibration is performed by weighted (1/concentration) linear regression, and should show linearity between 0.00100 and 0.400 mg per L with quantification limits at the lowest calibration levels. The intra-day precision of the bronchoalveolar fluid assay is reported to ranges between 2.3 and 7.9%, and is 2.3% at 0.800 mg per L, 6.9% at 0.200 mg per L, 7.9% at 0.0200 mg per L, and 5.2% at 0.00200 mg per L. The analytical error of the bronchoalveolar fluid assay ranges from −5.0 to 4.7%, and is 4.7% at 0.800 mg per L, 1.0% at 0.200 mg per L, −3.4% at 0.0200 mg per L, and −5.0% at 0.00200 mg per L.

Calculation of the corrected cefditoren level in BAL requires that the urea concentration be determined. To determine the urea concentration in BAL samples by LC-MS/MS, 100 µl of an internal standard solution in Milli-Q-water and 1 ml gradient grade methanol are added to 1 ml of each human BAL sample, and a 10 µl aliquot of each sample is applied to a reversed-phase column, eluted with an isocratic solvent system, and monitored by LC-MS/MS with an SRM method in positive mode, with precursor to product ion for urea, m/z 157 3 m/z 114, and for the internal standard, m/z 160 3 m/z 115. Urea and the internal standard are eluted after approximately 2.3 min.

As for cefditoren, calibration and inter-assay variation is assessed by testing samples in a range of concentrations prepared from a standard urea solution in buffer and an additional calibration standard such as cefotaxime in each run. No interferences were observed for urea and the internal standard. Calibration by weighted (1/concentration) linear regression showed linearity of the urea calibration curves between 0.208 and 8.00 mg per L, and quantification limits at the lowest calibrator concentrations. The reported range for the inter-day precision and the analytical recovery of the spiked quality control standards is from 0.3 to 0.9%, and is 0.3% at 8.00 mg per L), 0.9% at 1.60 mg per L and 0.3% at 0.400 mg per L, with an analytical error of −1.4% at 8.00 mg per L), 4.2% at 1.60 mg per L, and 8.6% at 0.400 mg per L (315).

Penicillins and Aztreonam

Because new semisynthetic penicillins continue to be developed and require pharmaceutical, pharmacokinetic, and clinical assessment, HPLC has much to offer for this antibiotic group. Additionally, because of possible side effects, particularly those that are dose-related, selected therapeutic monitoring seems reasonable. Table 8.16 summarizes HPLC methods available for the penicillins. Note that some methods are designed for the assessment of pharmaceutical dosage forms but can probably be modified for use with biologic fluids (317331). Assays for some major clinically useful penicillin and penicillin derivative antibiotics are presented in this chapter.


Aztreonam is a totally synthetic, monocyclic, β-lactam antibiotic with low toxicity that exhibits specific activity against aerobic gram-negative bacteria and aminoglycoside and cephalosporin-resistant gram-negative organisms. Assays performed by microbiologic methods are lengthy and do not separate metabolites. The following assay describes an ion-pair HPLC method for the quantitative analysis of aztreonam and its metabolites in human and animal serum and urine.


Two HPLC systems may be used interchangeably and yield equivalent results. One consists of the following components: two M-6000A solvent delivery pumps, a model 660 solvent gradient programmer and a model U6K injector (all from Waters Associates, Milford, MA), a model LC75 variable-wavelength UV detector set at 293 nm, an autocontroller (Perkin-Elmer, Inc., Welleshey, MA), and a model 3390A printer/plotter/integrator (Hewlett-Packard, Palo Alto, CA). The other HPLC system is a Hewlett-Packard model 1084B HPLC, which is equipped with a built-in variable-wavelength UV detector also set at 293 nm, an autosampler, and a printer/integrator. With both systems, the analysis is performed using the same column system, that is, a Bondapak C18 column (inside diameter, 3.9 mm; length, 3.0 cm) and a guard column (inside diameter, 3.9 mm; length, 3.0 cm). The guard column is packed with Bondapak C18 on Corasil (Waters, Milford, MA).

Chromatographic Conditions

The mobile phase consists of 0.005 mol/L tetrabutylammonium hydrogen sulfate/0.005 mol/L (NH4)2SO4 and acetonitrile (80:20 by volume), adjusted to pH 3.0. The solution is filtered and degassed before HPLC use and is pumped at a flow rate of 2.0 mL/minute.


Serum is diluted with an equal volume of acetonitrile and centrifuged for 2 minutes at ambient temperature at 15,000 × g. Supernatants are removed, and 50 µL is used for HPLC analysis. Urine samples are diluted 10-fold with 0.005 mol/L tetrabutylammonium hydrogen sulfate, pH 3.0. Fifty microliters of sample is used for analysis.

Standards of aztreonam for serum and urine are dissolved in the mobile phase and prepared at concentrations of 1,000, 500, 200, 100, 50, 20, 10, 5, 1, and 0.5 µg/mL. A standard curve is constructed using peak area versus concentration. Serum standards are prepared over the same concentration ranges as the mobile-phase standards but are diluted with an equal volume of acetonitrile and centrifuged. Supernatants are then used for HPLC analysis. Aztreonam urine standards are made in the same concentration ranges as described. Urine samples are diluted 10-fold with 0.005 mol/L tetrabutylammonium hydrogen sulfate and are analyzed by HPLC under the same conditions as the standards that are dissolved in the mobile phase. Peak areas for aztreonam obtained from the serum standards are plotted on the previously constructed standard curve of aztreonam standards dissolved in the HPLC mobile phase.


It is possible to analyze urine and serum from all species for aztreonam without encountering interfering peaks. The results are linear for serum and urine over the concentration range of 0.1 mg/mL to 0.5 µg/mL. Recovery of samples is between 96% and 102%, and total analysis time for either specimen type is less than 10 minutes per sample. Use of tetrabutylammonium hydrogen sulfate with acetonitrile and (NH4)2SO4 to ion-pair the SO3 group of aztreonam to the reverse-phase column gives a good retention time, a symmetrical peak shape for aztreonam, and also allows the drug to be separated from biologic fluid components. HPLC agrees well with a microbiologic assay for both sample types. The HPLC method offers the advantages of speed and the ability to separate chemical entities. Only a small sample is required for HPLC analysis (330).


Mezlocillin, an acylureidopenicillin, is a semisynthetic penicillin with a broad spectrum of antimicrobial activity against gram-positive and gram-negative organisms, including Enterobacteriaceae, P. aeruginosa, and Bacteroides spp. Previous assay methods, including HPLC, that have been developed to measure the drug’s concentrations have not used internal standards. The following is a description of a reverse-phase, ion-pair, extraction HPLC assay that uses piperacillin as the internal standard.

Chromatographic Conditions

The mobile phase consists of 5 mmol/L of a phosphate buffer, pH 7.0, that contains 5 mmol/L tetrabutylammonium phosphate/acetonitrile (75:25). Fifteen microliters of the reconstituted eluate is injected onto a 250 × 4.5-mm Econosphere C18 column (5 µm particle size). Column effluent products are eluted at ambient temperature with the mobile phase at a flow rate of 1.0 mL/minute and are monitored at 220 nm.


Solid-phase preparation is used for samples in the following manner: 0.5 mL of serum sample that contains 100 µL of tetrabutylammonium phosphate and 15 µL of piperacillin internal standard (1.0 g/L) are applied to solid-phase extraction columns that have been previously activated by washing with methanol and 5 mmol/L tetrabutylammonium phosphate. The column is then washed with water and dried by aspiration. The drug is eluted with 600 µL of an equivolume solution of chloroform/acetone. The eluate is then evaporated and reconstituted with 100 µL of the mobile phase. The assay is calibrated with drug-free serum spiked with mezlocillin (10 to 300 mg/L).


The peak area ratio of mezlocillin to the internal standard is linearly related to the mezlocillin concentration. The run-to-run coefficient of variation is less than 5%. Analytical recovery is 67%. There are no known interferences with other antibiotics that have been tested. The use of a solid-phase preparation and an internal standard had made this method unique compared with other HPLC assays for mezlocillin (329).


Thienamycin, or (5R,6S)-2-aminomethyl-[(1R)-hydroxymethyl]-2-penem-3-carboxylic acid (Ciba-Geigy, Ardsley NY) belongs to a class of β-lactam antibiotics, the carbapenems, of which imipenem, meropenem, ertapenem, and doripenem are now used clinically in the United States. Carbapenems combine, in a single structure, the antimicrobial properties of penicillins and the cephalosporins, and are active against gram positive and gram negative organisms, although activity against Pseudomonas aeruginosa varies among member of the class. The following method is an HPLC assay for the determination of (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid in plasma and urine. It uses the closely related aminoethyl derivative of (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid as the internal standard.


An RP-HPLC system consisting of a Hewlett-Packard (Palo Alto, CA) model 1084B instrument equipped with a variable-volume injector and a variable-wavelength detector that is set at 320 nm may be used. Analysis is performed using a prepacked column (20 cm × 4.6 mm, internal diameter) filled with LiChrosorb RP-8 (10 µM Hewlett-Packard) and a precolumn (1 cm × 4.6 mm, internal diameter) filled with Nucleosil C18 (30 µm). For the plasma assay, both columns must be replaced after 120 to 150 injections to prevent a decrease in separation efficiency. Peak areas are given by the integrator/recorder (79 850 A LC terminal), while peak heights are measured manually.

Chromatographic Conditions

The mobile phase consists of a phosphate buffer (8 ×10−3 mol/L Na2HPO4/5.9 × 10−3 mol/L KH2PO4), pH 6, and is degassed and filtered before use. The mobile phase is delivered at a flow rate of 1 mL/minute at room temperature.


Plasma samples are prepared by adding 50 µL of internal standard solution (116.2 mol/L prepared in phosphate buffer, pH 6); 50 µL of phosphate buffer, pH 6; and 250 µL of a saturated solution of ammonium sulfate (53 µg of ammonium sulfate in 72 mL of water) to 250 µL of plasma in a glass tube. The solution is vortex-mixed for 30 seconds and centrifuged at 1,400 × g for 10 minutes. One hundred microliters of supernatant is used for analysis. For urine samples, a 1-mL volume of urine is diluted to 25 mL with phosphate buffer, pH 6. One milliliter of the diluted sample is added to 100 µL of the internal standard solution and 100 µL of phosphate buffer, pH 6, in a glass tube and vortex-mixed for 15 seconds. Thirty microliters of the mixture is used for analysis. For reasons of chemical stability, both samples needed to be injected as soon as possible after preparation.

Solutions of internal standards (116.2 µmol/L) as well as (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid stock solutions (2.05 mmol/L) are prepared in phosphate buffer, pH 6. Reference solutions are prepared by dilution with the same buffer. Plasma-calibrated samples are prepared by adding 50 µL of reference solution of (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid to 250 µL of drug-free plasma, and correspond to concentrations ranging from 1.64 to 410 µmol/L. Urine-calibrated samples are prepared by adding 100 µL of reference solutions of (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid to 1 mL of 25-fold diluted urine. Resulting concentrations range from 41 µmol/L to 1.025 mmol/L.


(5R,6S)-2-Aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid and the internal standard are well separated from plasma and urine components without any interferences. Calibration graphs for plasma and urine are obtained by plotting the (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid/internal standard peak area ratio against the concentration of (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid. Their equations are calculated using weighted linear regression. The coefficients of variation are 3% to 8.6% and recoveries are close to 100%. The method described is suitable for the determination of (5R,6S)-2-aminomethyl-6-[(1R)-hydroxyethyl]-2-penem-3-carboxylic acid in plasma and urine (324).

Measurement in Other Body Fluids, Urine, and Tissues

Reversed-phase HPLC with detection of separated drug by UV or by tandem mass spectrometry has also been used to evaluate the penetration of carbapenem antibiotics into body fluids other than plasma and into muscle and adipose tissue.

HPLC Measurement of Doripenem Penetration into Peritoneal Fluid

Doripenem is a parenteral carbapenem currently approved for treatment of intra-abdominal infections and complicated urinary tract infections including pyelonephritis. Ikawa et al. (333) used the HPLC method that follows to measure the total concentrations of doripenem in serum and peritoneal exudate to determine its peritoneal penetration. Samples were collected following intravenous administration of doripenem to uninfected patients who were undergoing abdominal surgery . To prepare serum or peritoneal exudate for HPLC, a sample (400 µL aliquot) was transferred to an ultrafiltration device (Nanosep 10K; Pall Corporation, Northborough, MA, USA) which was centrifuged at 12000 g for 10 min. A 20 µL aliquot of the filtrate was injected onto a chromatograph.

Chromatography was carried out using a reversed-phase column [XBridge C18 5µm (4.6 × 150 mm); Waters Corporation, Milford, MA, USA] and ultraviolet absorbance was detected at 300 nm. A mixture of 0.1 M sodium acetate buffer (pH 4.6) and acetonitrile (95:5, v/v) was used as a mobile phase at a flow rate of 1 mL per min. The column temperature was 40 C, and the retention time for doripenem was 5.9 min. The method was linear over a concentration range of 0.052100 mg per L and the lower limit of quantification was 0.05 mg per L in both serum and exudate. The precisions in intra- and inter-day assay (n = 6) were within 1.8% and 3.7%, respectively. The accuracies in the intra- and inter-day assay were 99.8% to 105.2% and 99.0% to 102.8%, respectively.

LC−MS/MS for Measurement of Carbapenems in Plasma, Urine and Tissue: Measurement of Doripenem and Its Metabolite, Doripenem-M-1, in Plasma and Urine

Cirillo et al. (332) described several methods based on reverse phase chromatography followed by tandem mass spectrometry (liquid chromatographic-triple quadrupole mass spectrometry [LC-MS/MS]) that were used in pharmacokinetic studies to detect doripenem and its inactive metabolite (doripenem-M-1) in human plasma and urine. Plasma and urine samples were processed at 4°C then stored frozen at −70°C until analyzed.

For the first method, plasma samples underwent solid phase extraction followed by separation with reverse phase chromatography. Chromatographic retention of doripenem was obtained on a Luna C18 column (150 × 4.6 mm; Phenomenex, Torrance, CA) with a mobile phase composed of 70:30 ammonium acetate:methanol. Column effluent was analyzed using the mass spectrometer (Sciex API 365, Concord, Ontario, Canada) equipped with Turbo ion spray in the positive ion mode. The lower limit of quantification for doripenem in plasma and urine was 0.200 µg per mL. For doripenem in plasma, mean accuracy for the quality control samples ranged from104% to 105%, and mean precision ranged from 5% to 6%. For doripenem in urine, mean accuracy for the quality control samples ranged from 103% to 107%, and mean precision ranged from 6% to 8%.

For the second method, 100 µL aliquots of the human plasma samples were prepared for HPLC by protein precipitation. Determination of levels of doripenem and its metabolite in plasma required the use of two different solid state and mobile phase systems. Chromatographic retention of doripenem and internal standards was obtained on an Atlantis HILIC Silica HPLC column (150 × 2.1 mm, 5-µm particle size; Waters, Milford, MA) under isocratic conditions with a mobile phase composed of acetonitrile and 20 mM ammonium formate containing 0.2% formic acid. Chromatographic retention of the metabolite doripenem-M-1 and internal standards was obtained on a Discovery HS F5, Supelco Analytical HPLC column (100 × 2.1 mm, 3-µm particle size; Sigma-Aldrich, St. Louis, MO) under isocratic conditions with a mobile phase composed of acetonitrile and water containing 0.05% formic acid.For both doripenem and doripenem-M-1, column effluent was analyzed by multiple reactions monitoring using the triple quadrupole mass spectrometer (Quattro Micro, Micromass, Waters, Milford, MA) equipped with electrospray in the positive ion mode. The validated linear range for doripenem was 0.100 to 20 µg per mL with the lower limit of quantitation at 0.100 µg per mL. The validated linear range for doripenem-M- 1 was 0.200 to 20 µg per mL with the lower limit of quantitation at 0.200 µg per mL.

For analysis in urine samples, chromatographic retention of doripenem, doripenem-M-1, and the internal standards was obtained on a Restek Allure PFP Propyl HPLC column (50 × 2.1 mm, 5-µm particle size; Bellefonte, Pennsylvania) under gradient conditions, with mobile phase A composed of 5:95 acetonitrile and water containing 0.1% formic acid and mobile phase B composed of acetonitrile containing 0.1% formic acid. Column effluent was analyzed using the mass spectrometer (Sciex API 5000) equipped with Turbo ion spray in the positive ion mode. The lower limit of quantification for doripenem and doripenem-M-1 in urine was 0.200 µg per mL.

Measurement of Doripenem in Skeletal Muscle and Subcutaneous Tissues

Burian et al (334, with permission) used ultra-HPLC followed by tandem mass spectrometry (UHPLC-MS/MS) to determine doripenem levels in saliva, plasma, and extracellular space fluid of skeletal muscle and subcutaneous adipose tissue. Unbound drug concentrations were determined using The UHPLC-MS/MS system consisted of a Dionex UltiMate 3000 Rapid Separation (Germering, Germany) and an Applied Biosystems MDS Sciex API 4000 (MDS Sciex, Concord, Ontario, Canada) triple quadruple mass spectrometer fitted with an electrospray ionization source. Chromatographic separation was employed using a Waters Acquity CSH C18, 2.1 by 50 mm, 1.7 µm UHPLC column (Dublin, Ireland). The mobile phase consisted of aqueous formic acid (A) (pH 2.80) and acetonitrile (B). Three percent B was increased to 10% within 1.75 min, followed by 95% B for 2.25 min and re-equilibration for 3.70 min. The total run time for each sample was 7.70 min. The flow rate was kept at 0.500 mL per min at a temperature of 23°C. The autosampler was set at 10°C. The retention time of doripenem was 1.49 min +/−2.7%. The MS detection was performed with multiple reaction monitoring (MRM; positive ion mode) for quantification, and settings were optimized for doripenem analysis: nebulizer gas, heater gas, and curtain gas were set at 30, 30, and 40 L per min. Turbo IonSpray voltage was 5,500 V at 550°C. The collision activated dissociation (CAD) flow was used at 4 L per min, and the declustering and entrance potentials were 51 V and 10 V, respectively. The optimized collision energy (CE) for the analyte varied from 21 eV for the quantifier to 23 eV for the qualifier. The dwell time was adjusted to 120 ms per transition. Quantification of doripenem was employed using MRM of the transition m/z 420.9¡274.3 (quantifier), while the qualifier transition was m/z 420.9¡342.1. The Analyst 1.5 software (MDS Sciex,Concord, Ontario, Canada) was used to control the UHPLC-MS/MS system and to perform the measurement. The limit of detection (LOD, signal-to-noise ratio = 3) was 0.005 mg per L, and the limit of quantification (LOQ, signal-to-noise ratio = 10) was 0.025 mg per L.

Measurement of Ertapenem in Plasma and Tissue

A similar method using LC/MS has been developed by Koal et al. for the measurement of ertapenem in human plasma (369). This method also has been used to measure pharmacokinetics of ertapenem in colorectal tissue (340).

Column-Switching Approach for Pretreatment Before HPLC

Incorporation of column-switching before HPLC, which simplifies pretreatment of samples and has the potential for automation and analysis of multiple samples without loss of assay sensitivity or specificity, has been used in methods developed to assay levels of multiple beta-lactam antibiotics (337). The column switching approach is particularly useful for preparation of samples containing antibiotics such as ertapenem that are susceptible to hydrolysis during exposure to acids and solvents, and that can be altered or destroyed during harsh sample pretreatment procedures. During pretreatment, the antibiotic being assayed is concentrated on the head of the extraction column, while endogenous constituents that must be removed from the sample are passed through the column to waste; the exposure to acid is relatively short.

Assays for measurement of ertapenem in plasma and urine have been described that employ column-switching for on-line extraction at a neutral pH prior to reverse-phase HPLC.

The lower limits of quantization for the plasma and urine assays are 0.125 and 2.5, respectively (336,335).For this assay, a plasma sample is centrifuged and then injected onto an extraction column using 25 mM phosphate buffer, pH 6.5. After 3 min, using a column-switching valve, the analyte is back-flushed with 10.5% methanol-phosphate buffer for 3 min onto a Hypersil 5 mm C BDS 10034.6 mm analytical column and then detected by its absorbance at 300 nm. The sample preparation and HPLC conditions for the urine assay are similar, except for a longer analytical column 15034.6 mm. The plasma assay is specific and linear from 0.125 to 50 mg per mL; the urine assay is linear from 1.25 to 100 mg per mL. This assay has been used with minimal modification for detection of ertapenem in blister fluid, to determine its tissue penetration (338)

Further modification of the online extraction using an acidic mobile phase (0.1% formic acid,pH3) and a large injection volume (≥150 µL) has been reported to increase assay sensitivity to a LLOQ of 0.025 µg per mL, enabling its use for determination of ertapenem in cerebrospinal fluid. (339) This assay method is described in detail below (with permission).


The HPLC autosampler (7l7plus) and loading pump for column-switching (pump 1, 600E) were from Waters (Milford, MA, USA). The elution pump (pump 2, Series 200 LC Micropump) and the UV-Vis detector (785A) were from Perkin-Elmer (Cupertino, CA, USA). Column-switching was performed by an Autochrom M10 column-switching valve (10-port) purchased from Valco Instruments Company (Houston, TX, USA). On-line extraction was performed using a Maxsil C18 (50 mm × 4.6 mm, 10m) column from Phenomenex (Torrance, CA, USA) and chromatography was performed using a BDS Hypersil C18 (100 mm × 4.6 mm, 5m) HPLC column from Keystone Scientific (Bellefonte, PA, USA). Data were collected, stored, and analyzed using Turbochrom Navigator Client/Server version 6.1 (Perkin-Elmer, Cupertino, CA, USA). An Eppendorf EDOS 5222 (Brinkmann Instruments, Westbury, NY, USA) pipettor was used for sample and standard aliquots.

Chromatorgraphic Conditions

Mobile phase 1 (MP1, 0.1% formic acid) and mobile phase 2 (MP2, ACN/0.1% formic acid, 15:85 (v/v)) were filtered and degassed through a 0.22m Magna R nylon filter (Whatman International, Maidstone, UK). The flow rates for pump 1 and pump 2 were 1.5 and 2.0 mL per min, respectively.


CSF samples were thawed and then vortexed. Sample aliquots of 200 µL were mixed with 50 µL of buffer (0.1 M MES buffer, pH 6.5) and transferred to autosampler vials, which were capped, vortexed and stored in a 5°C autosampler until use. Each 150 µL sample was injected onto the extraction column for 3.5 min at a flow rate of 1.5 mL per min with MP 1.After 3.5 min, the analyte was back-flushed off of the extraction column and directed to the analytical column using MP 2, at a flow rate of 2.0 mL per min. At 5 min after injection, the switching valve returned MP 1 through the extraction column for equilibration prior to the next injection. Ertapenem eluted in 7 minutes. The total run time was 10 min.


The intraday precision and accuracy of this assay for calibration standards (0.025−10 µg per mL) and quality control samples (0.1, 0.5 and 2.5 µg per mL) were < 6.2% coefficient of variation and 96.8-104.0% of nominal concentration, respectively).


Sulbactam, a penicillanic acid sulfone and a competitive and noncompetitive β-lactamase inhibitor, is administered with ampicillin and cefoperazone to expand the drug’s antimicrobial spectrum to include β-lactam–resistant organisms. Assays that were described before the following method showed interference from metabolic products and lacked the sensitivity to determine sulbactam concentrations in tissue (368). The assay next presented is an HPLC method that is able to determine sulbactam and cefoperazone concentrations in plasma, urine, and prostate tissue.


An RP-HPLC system that consists of a model 510B pump to deliver the mobile phase and a model 481 variable-wavelength detector (Waters Associates, Milford, MA) may be used. The detector settings include a wavelength of 313 nm, 0.05 absorbance units full-scale for plasma, and 0.01 absorbance units full-scale for tissues. The analysis is performed with a Beckman ODS-5 stainless steel column (Beckman Instruments, Fullerton, CA).

Chromatographic Conditions

The mobile phase consists of 89% of a 0.1 mol/L phosphate buffer, pH 6.1/11% acetonitrile. Tetrabutylammonium hydroxide (40%, 2 mL) is added to 1 L of buffer. The buffer is degassed and filtered before use. The mobile phase is delivered at a flow rate of 2.21 mL/minute. The retention time of the sulbactam imidazole derivative is 6.0 minutes.


Sample preparation involves a two-step procedure: derivatization and extraction. The derivatization procedure for all three specimen types consists of adding 0.5 mL of an imidazole reagent to 1.0 mL of each specimen and standard. The specimens and standards are then vortex-mixed and stored at 4°C to allow the derivatization process to be completed. Imidazole reagent is prepared by dissolving 8.5 g of imidazole in water, adding 5 N HCl to bring the solution to pH 6.8, and adjusting the volume to 40 mL with water. Specimens are then extracted by using an acetonitrile/dichloromethane procedure. The procedure includes the addition of 2.0 mL of acetonitrile to the specimen test tube to precipitate the plasma proteins, centrifugation at 3,000 × g for 5 minutes, and the decanting of the supernatant into 3.0 mL of dichloromethane. The test tube is mixed and centrifuged. A WISP automated sample processor is used to deliver 15 µL of the upper aqueous layer that contains the sulbactam imidazole derivative.

Prostate tissue is blotted, weighed, diluted with phosphate buffer, pH 6.1, and homogenized. A standard curve for the determination of sulbactam concentrations in pooled plasma is prepared with concentrations ranging from 0 to 100 µg/mL. A standard curve for determination of sulbactam concentrations in prostate tissue is prepared by the addition of 2 to 25 µg of sulbactam per gram to a tissue supernatant. A standard curve for the determination of sulbactam concentrations in normal urine is prepared with concentrations of 0 to 100 µg/mL. No internal standard is added to any sample.

Chromatograms do not contain interfering peaks from body metabolites in plasma, urine, or prostate tissue, or show interference with cefoperazone or other antimicrobial agents (175). The two-step procedure is linear from 100 to 1.0 µg/mL. Extraction efficiency of the sulbactam imidazole derivative is more than 90% for all specimen types. The coefficient of variation for inter- and intrabatch studies for all specimens is generally less than 11.4%. The recovery of sulbactam from urine is more than 50%.


A number of different methods are available for therapeutic monitoring of aminoglycoside levels. Most frequently, aminoglycoside levels in human specimens are obtained with nonisotopic immunoassays. Microbiologic assays, RIA, EMIT, HPLC, and several other assays have been compared for the monitoring of gentamicin concentrations (341,372). RIA, EMIT, FPIA, and HPLC are advantageous in that their results are available within a few hours, compared with 4 to 10 hours for microbiologic assays, depending on the method employed. However, interference from other aminoglycosides should be anticipated when using RIA or EMIT to quantify one of the members of this group. HPLC, although more complex and time-consuming than EMIT and, to a lesser extent, RIA, offers greater specificity than either method. Additionally, it is the only one of these methods that separates the components of gentamicin and other aminoglycosides. Table 8.16 provides information on several HPLC methods that are available to therapeutically monitor the aminoglycosides. If the decision is made to monitor an aminoglycoside by HPLC, the specific method selected will probably depend on the nature of the available equipment and the level of technical expertise within the laboratory (342,343). It has been the authors’ opinion that the HPLC method of Marples and Oates (275) deserves initial consideration.

Examples of the major clinically useful aminoglycoside/aminocyclitol antibiotics are presented here. Gentamicin was the first major aminoglycoside to be analyzed by HPLC. Its assay serves as a model for the assay of the other aminoglycosides.


A potassium borate buffer (0.4 mol/L) is prepared by the titration of 24.7 g of boric acid into 900 mL of distilled water. The pH is brought to 10.40 with concentrated potassium hydroxide solution. The solution is then diluted to 1 L.

o-Phthaldehyde (OPA) is prepared by dissolving 60 mg of OPA in 1 mL of methanol, followed by the addition of 2-mercaptoethanol (0.2 mL). The solution is gently stirred until complete decolorization occurs. Potassium borate buffer (100 mL) is added, and the mixture is vigorously stirred. The solution is placed in the supply vessel for OPA and flushed several times with nitrogen. It must be used within 2 days of preparation.

Stock gentamicin is prepared at a concentration of 1,000 µg/mL in 0.1 mol/L phosphate buffer, pH 8.0, and stored at −20°C until use. Appropriate dilutions of the stock solutions are made with water. A serum solution that contains 10 µg/mL gentamicin is prepared by the addition of gentamicin standard at 100 µg/mL to serum. Further dilutions from this serum standard are made in normal human serum.


A Tracor model 990 pump (Tracor Instruments, Austin, TX) is used to deliver mobile phase. A Schoeffel model 970 fluorometer (Schoeffel Instrument Corp, Westwood, NJ) is used to detect the fluorescence product that is formed by continuous flow, postcolumn derivation with OPA. Fluorescence excitation is at 340 nm and a KV418 filter is used for emission. The photomultiplier voltage is varied between 1,000 and 1,100 V, depending on the sensitivity required. OPA is supplied to a mixing tee from a pressurized glass vessel. A Cheminer fitting (CJ-303 1; Laboratory Data Control, Riviera Beach, FL) is used for the mixing tee, and a delay coil consisting of a Teflon tube (2.0 × 0.6 mm i.d.) is used between the mixing tee and detector. Analysis is performed by using a Bondapak C8 column (30 cm × 3.9 mm i.d.; Waters Associates, Milford, MA), with a precolumn (4.3 cm × 4.2 mm i.d.) packed with Micropart C18 phase-bonded silica gel (5 µm; Applied Science Laboratories, Deerfield, IL). The detector signal is processed and recorded by using a CDS 101 computing integrator (Varian) and a model 7123A recorder (Hewlett-Packard, Palo Alto, CA). Samples are injected using a Valco CV-6-VHPa-C20 injection valve with a 15-µL injection loop modified for variable-volume injection (Glenco Scientific, Houston, TX).

Chromatographic Conditions

Mobile phase compositions are expressed as the ratio of components, by volume, that are added to produce the final solution. No corrections need to be made for volume changes that occur as a result of mixing. Predominantly aqueous mobile phases are degassed before use by vacuum filtration through a Millipore type HA membrane filter (0.45 µm; Millipore Corp, Bedford, MA). Solutions that contain predominantly methanol are filtered through a solvent-resistant type of FH filter (0.5 µm). The mobile phase that is used for analysis contains 0.2 mol/L Na2SO4, 0.02 mol/L sodium pentasulfonate, and 0.1% (v/v) acetic acid in a water/methanol (97:3) mixture. The column flow rate is 2 mL/minute at 184 atm. The OPA flow rate is approximately 0.5 mL/minute.

Gentamicin is separated from interfering compounds in serum by ion-exchange gel chromatography. A 1.5-cm column with a bed volume of 1 mL is prepared with CM-Sephadex C-25 using 0.2 mol/L Na2SO4 as the initial buffer. Four hundred microliters of serum is applied to this column and eluted with 1 mL and 4 mL of the initial buffer in succession. The eluting buffer is changed to 0.01 mol/L NaOH in 0.2 mol/L Na2SO4, and 600 µL of this buffer is added to the column. After the column has drained completely, a second volume of alkaline buffer (400 µL) is added and the eluate is collected. Fifteen microliters of this fraction are injected for chromatographic analysis. Figure 8.19 presents the results of a typical assay. The chromatogram of a mixture of C1a, C2, and C1 gentamicin, all at 2.5 µg/mL, is pictured.



The liquid chromatography system is from Kontron with a Uvikon photometric detector (Kontron AG, Eching). The LiChrosorb columns (Sigma-Aldrich, St. Louis, MO) are connected by an eco-tube cartridge system.

Chromatographic Conditions

The mobile phase is methanol/water (5:95, v/v) containing 0.01 mol/L disodium n-heptanesulfonate adjusted to pH 14.0 with sodium acetate and acetic acid. The HPLC system is operated at a flow rate of 0.8 mL/minute, a pressure of 59 bar, and a temperature of 30°C.


A 1.0-mL plasma sample is extracted by dilution with n-heptanesulfonic acid and forced through a SAX Bond-Elut (Agilent Technologies, Inc, Santa Clara, CA) sample cleanup cartridge by vacuum. Plasma constituents are eluted from the cartridge with n-heptanesulfonic acid and teicoplanin is eluted with methanol. Detection is performed by measurement of the absorbance at 240 nm.


The calibration graph is linear in the range of 3 to 50 µg/mL. The detection limit is 0.2 to 0.4 µg/mL (44).


CLD, synthesized from microbially fermented lincomycin, is an antibiotic that is often used to treat infections with gram-positive and gram-negative anaerobes, as well as gram-positive aerobes. Other analytical procedures, including GC and microbiologic assays, exist for the detection of CLD in biologic fluids. However, the GC assay requires lengthy extraction and derivatization procedures, while the microbiologic assay lacks specificity and is less accurate than HPLC. Both of these assays are time-consuming and labor-intensive. Previously used HPLC assays have been applied only to the analysis of CLD concentrations in sterile fluids or water and have not been used with biologic fluids. The following assay is an excellent HPLC assay for the quantification of CLD in biologic fluids, serum, and plasma.


An RP-HPLC system is used that includes a model 510 pump (Waters Associates, Milford, MA) to deliver the mobile phase. A model 783 variable-wavelength UV detector (Spectros, Ramsey, NJ) with a detector wavelength of 198 nm and a filter response time of 0.1 second provides a reproducible, quantifiable peak that corresponds to 0.17 µg/mL. The monochromator of the detector is continuously purged with purified nitrogen to help decrease oxygen quenching and temperature-drift instability. Analysis is performed by using a Nova-Pak C18 octadecylsilane column (5 µm, 15 cm × 3.8 mm; Waters, Milford, MA) that has been wrapped in styrofoam to help stabilize the temperature. The detector signal is processed and recorded by using a Pro-840 computer printer (Waters, Milford, MA) for data collection, system operation, and data storage/retrieval.

Chromatographic Conditions

The mobile phase consists of acetonitrile/water/phosphoric acid/7.6 mmol/L trimethylammonium chloride (30:70:0.2:0.75), with a final pH of 6.7. The mobile phase is prepared in 2-L quantities in the following manner. A 600-mL volume of acetonitrile is transferred to a 2-L graduated cylinder. Nanopure (Millipore, Milford, MA) water is then poured into the cylinder until the fluid level reaches the 1,900-mL mark. Four milliliters of phosphoric acid is added along with 15 mL of a previously prepared 10% trimethylammonium chloride solution. Water is added to the 2-L mark and the contents are filtered under vacuum with a 0.45-µm membrane filter. The pH is adjusted to 6.7 with 1 mol/L sodium hydroxide and pumped through the column at a flow rate of 1.0 mL/minute and a pressure of 130 bar. A constant slow bubbling of helium through the mobile phase during operation is required to minimize the amount of oxygen in the system. Retention times for CLD and triazolam average 8 and 11.8 minutes, respectively.


Heparinized plasma and serum samples (200 µL) are prepared by protein precipitation with 0.5 mL of acetonitrile that contains the internal standard triazolam (44 ng/100 mL). Samples are vortex-mixed for 20 seconds and centrifuged at 3,000 × g for 10 minutes. The resulting supernatant is poured off the protein pellet and evaporated under nitrogen to a volume of 250 µL. A WISP autoinjector is used to deliver 15 to 30 µL of the concentrated supernatant into the HPLC system. Standards are human plasma samples with the addition of the internal standard triazolam and CLD at 1.5 and 12 µg/mL, respectively.


Standard curves are constructed using an unweighted least squares (y = mx + b) method with the origin as a data point and a floating intercept. All chromatograms must be recalculated with forced baselines, as provided in a special chromatographic software package (Waters Associates, Milford, MA). The use of peak height provides better linearity than peak area when applied over the entire standard curve. Replicate standard curves run over a 36-hour period show no loss of integrity. The ranges for the coefficients of variation for intraday and interday measurement are 2.4% to 5.7%. The recovery range is 96% to 118%, and there is no degradation of CLD in human plasma stored at −20°C for up to 56 days.

The use of a low-wavelength setting combined with the simple removal of oxygen from critical areas of the system allows for precise measurement of CLD concentrations in human plasma. This HPLC method also has the ability to measure CLD in other biologic fluids. It is fast, simple, and requires no extraction (288).


Roxithromycin is a macrolide antibiotic with an antibacterial spectrum of activity similar to that of erythromycin that exhibits enhanced, clinically desirable pharmacokinetic properties. Specifically, plasma levels are more sustained and are higher than for erythromycin, allowing for lower doses and less frequent administration. Previous assays for the macrolides have included microbiologic assays and HPLC using UV absorption fluorescence and electrochemical detection methods. Bioassays are lengthy and are not selective in the presence of active metabolites. HPLC that uses UV detection is limited because of weak macrolide absorbance in the low–UV wavelength range (<235 nm). Thus, large volumes of patient specimens (2 to 3 mL) are required to attain sufficient sensitivity and the weak macrolide absorbance results in increased background noise and large interferences from coextracted samples. The following HPLC assay uses electrochemical detection of roxithromycin in plasma and urine.


An RP-HPLC system is used and consists of a solvent delivery pump (type 364.000; Knauer, Berlin, Germany), a manual injector (model U6K; Waters Associates, Milford, MA), and a dual-electrode electrochemical detector (ESA model 5100A; Coulochem Environmental Sciences, Bedford, MA). The electrochemical detector is equipped with a guard cell that has been placed in-line before the injector to electrolyze components of the mobile phase. The model 5010 dual-electrode cell is operated in the oxidative screen mode. The applied cell potential of the screen electrode E1 is set at + 0.7 V and the sample electrode E2 at + 0.9 V. This allows irreversible oxidation of many compounds in plasma and urine at the first electrode (E1) without a decrease in the response for roxithromycin and the internal standard. Analysis is performed using a Bondapak C18 reverse-phase column (30 cm × 3.9 mm, internal diameter; 10-µm particle size; Waters Associates, Milford, MA). The detector signal is recorded by an Omniscribe recorder (Houston Instruments, Houston, TX).

Chromatographic Conditions

The mobile phase consists of acetonitrile/83 mmol/L ammonium acetate/methanol (55:23:22 by volume), with the pH adjusted to 7.5 with acetic acid. The pH affects the retention times and oxidation of the macrolides. A pH of 7.5 is optimal. The low ionic strength of the buffer also helps provide adequate conductivity and minimizes background current. Before use, the mobile phase is filtered and pumped at ambient temperature through the column at a flow rate of 1.0 mL/minute. The retention times of roxithromycin and erythromycin base are 9.8 and 7.0 minutes, respectively.


Two hundred–microliter aliquots of plasma or diluted urine (diluted 1:2 with isotonic NaCl solution), 100 µL of internal standard (10 µg/mL erythromycin), 600 µL of phosphate buffer, pH 9, and 3 mL of dichloromethane are pipetted into 10-mL glass extraction tubes, which are stoppered and mixed by shaking for 10 minutes. Following centrifugation at 2,000 × g for 5 minutes, the upper layer is discarded and the organic phase (2.5 mL) is evaporated at ambient temperature under a stream of nitrogen. The residue is reconstituted with 50 µL of methanol and vortex-mixed for 10 seconds. A 15-mL aliquot is injected into the HPLC system. Standards are human plasma and urine samples with roxithromycin at concentrations of 1 to 20 µg/mL. The internal standard erythromycin concentration is 10 µg/mL in doubly distilled water.

Chromatograms are recorded at a chart speed of 0.25 cm/minute and peaks are well resolved from endogenous plasma or urine compounds. Peak height ratios of roxithromycin to erythromycin are measured.


There is a linear relationship between concentration and response up to 25 µg/mL in plasma samples and diluted urine samples. Instrument precision as determined by repeated injection of 500 mg of roxithromycin is 2.3%. The ranges for the coefficients of variation for intraday and interday variation are 1.6% to 5.5% and 2.2% to 7.0%, respectively. The recovery ranges for roxithromycin and erythromycin are 6.82 ± 1.0% and 78.8 ± 2.4%, respectively, with no interfering peaks. Roxithromycin does not degrade upon storage at 37°C, 4°C, or −20°C. The use of dual coulometric electrodes that are operated in the oxidative screen mode allow for the rapid and sensitive detection of roxithromycin in small patient samples (280).


Pyrazinamide, an analog of nicotinamide, exhibits antimycobacterial activity when it is administered with other drugs. The drug is well absorbed after oral administration and hydrolyzed and hydroxylated to the following metabolites: 5-hydroxypyrazinamide, pyrazinoic acid, and 5-hydroxypyrazinoic acid. All of these may be important in drug level monitoring to avoid the dose-related adverse effects of hyperuricemia and hepatotoxicity. Earlier assays, including an HPLC method that measures pyrazinamide and its three major metabolites, are time-consuming, tedious, and involve difficult extractions. The following method is rapid and allows simultaneous determination of pyrazinamide and its metabolites.


The chromatographic system consists of an LC-6A chromatograph (Shimadzu, Kyoto, Japan) with an RF 530 HPLC fluorescence monitor (Shimadzu, Kyoto, Japan) set at 410/365 nm. Analysis is performed using a 10-µm Bondapak C18 column (30 cm × 3.9 mm, internal diameter; Waters Associates, Milford, MA) at a column temperature of 25°C. The signal is recorded using a C-R3A Chromatopac recorder (Shimadzu, Kyoto, Japan).

Chromatographic Conditions

The mobile phase consists of 0.02 mol/L KH2PO4, pH 2.56, and is delivered at a flow rate of 2.0 mL/minute. Peaks elute between 2 and 8 minutes in the following order: 2,3-pyrazinedicarboxamide, pyrazinoic acid, and pyrazinamide.


A 100-µL aliquot of 2 mol/L perchloric acid is placed in a 5-mL glass tube that contains 0.5 mL of plasma, 0.3 mL of distilled water, and 100 µL of 10 mmol/L 2,3-pyrazinedicarboxamide (as the internal standard) and is thoroughly mixed for 10 seconds. After centrifugation at 1,500 × g for 10 minutes, a 200-µL aliquot of supernatant is neutralized with 48 µL of 1 mol/L sodium hydroxide solution. Sixty microliters of the supernatant is loaded onto the column.

The calibration graph is constructed by using 0.5 mL of drug-free plasma and adding 0.1 mL of a standard solution that contains pyrazinamide, its metabolites, and the internal standard plus 0.3 mL of distilled water to obtain a calibration between 5 and 80 µg/mL for pyrazinamide, between 2.5 and 50 µg/mL for pyrazinoic acid, between 1 and 10 µg/mL for 5-hydroxypyrazinamide, and between 2.5 and 0.25 µg/mL for 5-hydroxypyrazinoic acid. This allows creation of a calibration graph because a small peak of an endogenous compound in plasma overlaps the peak of 5-hydroxypyrazinamide under these chromatographic conditions.


Peak area versus concentration is linear with standards in aqueous solutions up to 100, 50, 10, 2.5, and 150 µg/mL for pyrazinamide, pyrazinoic acid, 5-hydroxypyrazinamide, 5-hydroxypyrazinoic acid, and 2,3-pyrazine-dicarboxamide, respectively. Detection limits for 5-hydroxypyrazinoic acid, 5-hydroxypyrazinamide, pyrazinoic acid, and pyrazinamide are 3, 3, 30, and 30 ng, respectively, with an apparatus detection limit of 0.45 ng/12 µL cell (344).


Linezolid, a new oxazolidinone antibiotic has broad activity against gram-positive organisms, including MRSA, vancomycin-resistant enterococci, and penicillin-resistant pneumococci. HPLC protocols for the measurement of linezolid levels in serum and other body fluids have been published (345350). The following is a representative example of an HPLC assay for the measurement of linezolid concentrations in human serum. (345)


The stationary phase consists of Hypersil 5ODS, 10 cm × 4.6 mm (Waters Corporation, Milford, MA). A Gina 50 autosampler (Dionex, Macclesfield, United Kingdom) for UV absorbance detection (λmax254), and the integrator is a Trilab 2000 (Trivector, Sandy, United Kingdom).

Chromatographic Conditions

The mobile phase consists of 1% ortho-phosphoric acid (BDH, Analar grade, Poole, United Kingdom), 30% methanol (Prolabs, Fontenay, France), and 2 g/L heptane sulfonic acid (Sigma-Aldrich, St. Louis, MO) adjusted to pH 5 by the addition of 10 M sodium hydroxide. The flow rate is 1.0 mL/min. Retention time for linezolid is 384 seconds.


Samples are prepared by mixing aliquots (50:50) of the specimen with acetonitrile (Prolabs Fontenay, France). After mixing, the samples are incubated at room temperate for 10 minutes then centrifuged at 5,000 g for 5 minutes. Twenty microliters of the supernatant is injected.


The assay has been demonstrated to be reproducible, accurate, and linear at a concentration range from 0.0 to 30 mg/L. Intraday and interday reproducibility were less than 6% and 12.5%, with good correlation between drug concentration and peak for both aqueous and serum samples (r = 0.9999 for both). Linezolid recovery from serum approached 100% for concentrations tested. Accuracy of the assay expressed as percentage error was 4.0 for a 2.5 mg/L sample, 1.3 for an 8 mg/L sample, and 0.0 for an 18 mg/L sample. The lowest LOQ was 0.1 mg/L. No interference from 23 commonly used antimicrobial agents or unknown compounds in linezolid-free patient sera was found. Linezolid was shown to be stable in serum after sample preparation with acetonitrile for 24 hours at room temperature and was stable in serum alone for at least 7 days at room temperature and at 4°C.


GLC has been used successfully in the past to assay for antibiotics that could be volatilized. With the application of HPLC to virtually all antibiotic classes, the need for GLC analyses has become significantly reduced. The general principle of GLC analysis of antibiotics and an abbreviated assay of one drug, CAM, are presented. The method often includes a step that involves the conversion material to be assayed into a compound of high volatility so that it may be passed through a column in the gaseous phase. The general steps involved in GLC are the following:

 1.  Extraction of the drug from the serum sample

 2.  Conversion of the drug to a highly volatile form

 3.  Separation by GLC

 4.  Detection by flame ionization electron capture and so on

 5.  Quantitation of the drug by peak height or peak area

The finding that compounds can be rapidly converted to trimethylsilyl derivatives caused the gas chromatograph to become a tool for clinical laboratories. However, HPLC is the chromatographic technology of choice for the analysis of most antibiotics. The measurement of the concentrations of cephalosporin and (351) spectinomycin (352) antibiotics, CAM (353,354) and its chemically modified analogs (249,355), and the aminoglycosides have been determined using GLC.



A model 571 OA gas chromatograph equipped with dual flame ionization detector and a 1 mV model 7123A recorder (Hewlett-Packard, Palo Alto, CA), or the equivalent, is used. Glass columns are 122 cm × 2 mm (i.d.) (configuration 5) for on-column injection (Hewlett-Packard, Palo Alto, CA), packed with 3% OV-I on 100/120 Gas Chrom Q (Applied Science Laboratory Alltech Associates, Inc, Deerfield, IL). Septa are type HT-9 (high temperature, low speed; Applied Science Laboratory, Deerfield, IL).

The instrument conditions are as follows: injector temperature, 250°C; detector temperature, 300°C; oven temperature, programmed from 190°C to 270°C at 16°C per minute; gas flow rate, 40 mL/minute nitrogen, 250 mL/minute air, and 40 mL/minute hydrogen; and recorder chart speed, 13 mm/minute (Fig. 8.18).


Serum (500 µL) is combined in 16 × 125-mm test tubes (Teflon-lined, screw cap) with 500 µL of a phosphate buffer (1 mol/L, pH 6.8) and 7 mL of ethyl acetate (Nanograde; Mallinckrodt Baker, Phillipsburg, NJ) that contains 21 µg of the internal standard. The tubes are mixed by shaking for 10 minutes in an Eberbach shaker at 350 oscillations/minute and centrifuged at 2,000 rpm in a desktop centrifuge for 10 minutes. The upper (organic) phase is transferred to fresh 16 × 125-mm tubes and is evaporated under a stream of nitrogen in a water bath at 40°C. Four milliliters of HCl (0.5 mol/L) is added and the tubes are held in an ultrasonic bath for 10 seconds to ensure that the residue completely dissolves. Seven milliliters of hexane is added and the tubes are mixed by shaking for 10 minutes in the Eberbach shaker (Eberbach Corporation, Ann Arbor, MI) and centrifuged for 3 minutes at 2,000 rpm. The upper (organic) phase is transferred into 7-mL screw-capped septum vials and evaporated under a stream of nitrogen in a 40°C water bath. Fifty microliters of Trisil (Pierce Chemical Co, Rockford, IL) is added to the residue and each vial is immediately capped. The vial contents are mixed on a vortex-type mixer and are allowed to stand for 8 minutes at room temperature. Two microliters of sample is injected into the gas chromatograph. One may sialylate the next sample immediately after injection to save time. The results are calculated by the peak height ratio method, using CAM and the acetyl analog of CAM as internal standards.

Standard curves are prepared by analyzing samples of normal serum containing known amounts of CAM. It is convenient to run 10, 20, 40, 60, and 80 µg/mL in normal serum as standards.


TLC has been used in the clinical laboratory for many years to separate high molecular weight, biologically active compounds. Like all forms of chromatography, the unknown may be identified based on a comparison of the mobility of the unknown compound in a defined matrix with the mobility of standards. A TLC method has been developed that first separates antibiotics from a mixture based on their mobilities in the chromatograph and then, using a test strain of indicator organism, quantitates them by their biologic activities.

Tobramycin (Nebramycin Complex)


Thin-layer plates (20 × 20 cm) are prepared from silica gel G. Glass plates of sizes 20 × 2 × 0.03 cm and 16 × 2 × 0.3 cm are used for framing the chromatograms.


All standard antibiotic solutions are dissolved in distilled water. Two developing solvents are employed: methylethyl ketone (96%)/ethanol (25%)/ammonium hydroxide (1:1:1) and chloroform/methanol (25%)/ammonium hydroxide (1:7:4). Solvent systems are made fresh before use (Table 8.18). Bioautograms are stained with 1% tetrazolium blue and 0.02% tetrazolium violet.


The test organism used is B. subtilis ATCC strain 6633.


Silica plates with a Camag or Desaga coater are made 0.25-mm thick, dried at room temperature for 2 days, and used without further pretreatment. Lines are drawn before use to ensure separate, 1.5-cm wide tracks. Tobramycin standard solutions are each diluted to a final content of 0.1 to 0.5 µg of the analyte. Patient specimens are dissolved in distilled water to also cover this range. Chromatography is allowed to proceed for a distance of 15 cm at room temperature. The plate is subsequently air-dried.

For the detection of microbiologic activity, nutrient agar is melted and the B. subtilis test organism is added to a final organism concentration of 1 × 108/mL. A 25-mL volume of this medium is poured onto the glass frame in order to cover the silica gel surface uniformly. The agar is covered with a 20 × 20-cm glass chromatographic plate and incubated for 10 to 16 hours at 37°C.


Quantitative determinations are performed using a calibration graph. The zones of inhibition of the B. subtilis test organism are measured with the standards and unknowns, and the concentration of tobramycin is calculated in the same manner as for the analogous electrophoresis/bioautography procedure.


The TLC/bioautography method is able to measure as little as 0.1 to 0.3 µg of tobramycin (nebramycin complex). It is able to separate tobramycin into nebramycin 2, 4, 5′, 5, and 6. Furthermore, the TLC/bioautography method is able to differentiate between closely related aminoglycoside antibiotics by their gel mobilities.

The tetrazolium blue solution gives a colorless spot on a deep red background in order to measure the zones of inhibition more efficiently. In addition to tobramycin, the method is able to detect 0.2 µg of gentamicin and 0.1 µg of neomycin, paromomycin, and kanamycin (356). TLC is a useful method for the detection of tetracyclines (357,358).


A procedure based on the ion-exchange chromatographic separation of aminoglycosides from human body specimens has been developed for the spectrofluorometric assay of aminoglycosides/aminocyclitol antibiotics. The aminoglycosides are separated by elution from the column with sulfuric acid, followed by fluorometric analysis after derivatization.

Ion-Exchange Assay


Reagents for ion-exchange chromatography include 0.1 mol/L sulfuric acid and 0.5 mol/L sulfuric acid. Reagents for the analysis of aminoglycoside concentrations include acetylacetone, formaldehyde (30%), and Britton-Robinson buffer, made by mixing 100 mL of solution 1 (0.2 mol/L phosphoric acid, 0.2 mol/L acetic acid, and 0.2 mol/L boric acid) with 15 mL of 1.0 mol/L sodium hydroxide (359). To 10 mL of the final buffer, which is at pH 2.6, 0.8 mL of acetylacetone and 2.0 mL of formaldehyde are added.


The aminoglycosides are analyzed spectrophotometrically with a fluorescent dihydrolutidine derivative that is developed by the condensation of the primary amino group with acetylacetone and formaldehyde under acidic conditions. A 13.9 × 290-mm glass column is packed with 3 mL of Amberlite IRC 50 resin (sodium form). Either human urine (2 mL) or human serum (5 mL) is diluted to 10 mL with distilled water and applied to the column. Twenty milliliters of distilled water are applied to the column and impurities are diluted with 20 mL of 0.1 mol/L sulfuric acid. The aminoglycoside antibiotics are eluted from the column with 20 mL of 0.5 mol/L sulfuric acid. Two milliliters of this eluent is analyzed for the aminoglycoside concentration. The elution rate is approximately 0.5 mL/minute.

Two milliliters of the eluent is added to 2 mL of the analysis reagent and heated at 100°C for 10 minutes. After heating, the aminoglycoside is quantitated by determining its absorbance at an excitation wavelength of 421 nm and an emission wavelength of 488 nm in a 1-cm light path.


The method should be able to analyze any of the aminoglycoside antibiotics. Tobramycin, neomycin, sisomicin, kanamycin, and amikacin have been quantitated. The method is sensitive to 0.5 µg/mL.


Micellar electrokinetic chromatography is a type of capillary zone electrophoresis in which a detergent forms micelles with electrically neutral substances, allowing their separation and detection by UV absorption (360). This method permits the quantitation of penicillin in plasma. Plasma proteins are solubilized by the detergent and thus do not interfere with the assay.

Micellar Method


Sodium dodecyl sulfate is dissolved in a buffer of 0.02 mol/L sodium dihydrogen phosphate with 0.02 mol/L sodium tetraborate, passed through a 0.45-µm membrane filter, and degassed by sonication. Standards are dissolved in water to a concentration of 1 mg/mL.


A fused silica capillary tube with dimensions of 650 mm × 50 µm is used as a separation tube. Detection of antibiotic is achieved by on-column UV absorption measurement at 210 nm, using a Uvidec 100-V1 detector (Jasco, Tokyo, Japan) with a time constant of 0.05 second.


The calibration graph for aspoxicillin is linear in the range of 25 to 300 µg/mL. The detection limit is 1.3 µg/mL. The average recovery is 94% to 104%.


Nuclear magnetic resonance techniques, while not widely available in the clinical setting, have been investigated for the measurement of antibiotic concentrations. When spinning nuclei in a magnetic field are irradiated by a second perpendicular field, they change their alignment to the new field. The amount of energy required for the transformation is characteristic of the molecule and depends on factors such as electronic configuration and intermolecular interactions. Nuclear magnetic resonance spectroscopy can provide information about the kinetic and structural aspects of the interactions between ligands and macromolecules, such as drugs and receptors. It offers both sensitivity and specific simultaneous identification and quantification of drugs and their metabolites in plasma, serum, and urine. Antibiotics that have been assayed include tetracyclines, penicillins, cephalosporins, and erythromycin (361).


Gonzalez Perez et al. (362) have described a method for the determination of levels of clavulanic acid in the presence of amoxicillin by differential pulse polarography. Clavulanic acid is hydrolyzed in a sulfuric medium to obtain an electroactive product with a reduction peak at 0.75 V. The procedure can detect clavulanic acid in the range of 8 to 2 mol/L.


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a12 CFR parts 493.803 and 493.1236 (2003).

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