Most tests in the toxicology laboratory are directed toward the identification or quantitation of xenobiotics. The primary techniques used include spot tests, spectrochemical tests, immunoassays, and chromatographic techniques. Mass spectrometry may also be used, usually in conjunction with gas chromatography (GS) or liquid chromatography. Table 6–2 compares the basic features of these methodologies. Other methodologies include ion-selective electrode measurements of lithium, atomic absorption spectroscopy or inductively coupled plasma mass spectroscopy for lithium and heavy metals, and anodic stripping methods for heavy metals. Many adjunctive tests, including glucose, creatinine, electrolytes, osmolality, metabolic products, and enzyme activities, may also be useful in the management of poisoned patients. The focus here is on the major methods used for directly measuring xenobiotics.
The simplest tests are spot tests. These rely on the rapid reaction of a xenobiotic with a chemical reagent to produce a colored product (eg, the formation of a colored complex between salicylate and ferric ions) that is visually assessed in a semiquantitative manner. Because the reagents may cause precipitation of serum proteins, spot tests are more commonly performed on urine specimens or gastric aspirates. Such tests were once a mainstay of toxicologic testing. Because of the poor selectivity of chemical reagents, as well as substantial variability in visual interpretation, these assays suffer from fairly frequent false-positive results and occasional false-negative results and are rarely used today.
Spectrochemical tests rely on measurement of a light-absorbing substance. Some analytes that are intrinsically light absorbing may be directly measured. Cooximetry (also known as hemoximetry) represents a sophisticated application of spectrophotometry to the measurement of various forms of hemoglobin in a hemolyzed blood sample. Measurement of light absorbance at multiple wavelengths allows several hemoglobin species to be simultaneously quantitated. For mathematical reasons, the number of wavelengths used must be greater than the number of different types of hemoglobin present. This is why classic pulse oximetry, which uses only two wavelengths, yields spurious results in the presence of significant amounts of methemoglobin or carboxyhemoglobin (Chaps. 29, 125, and 127). Cooximetry is relatively free of interferences because the concentrations of the hemoglobins are so much higher than other substances in the blood. However, the presence of intensely colored substances (eg, methylene blue) may cause spurious increases or decreases in the apparent percentages of the hemoglobins. Modern instruments are often able to recognize a significantly atypical pattern of absorbance and generate an error message in addition to or instead of a result.
Most analytes are neither as deeply colored nor as highly concentrated as hemoglobin species. Their detection requires a chemical reaction to produce an intensely light-absorbing product that is quantitatively measured at a specific wavelength in a spectrophotometer. Because spectrophotometers can also measure ultraviolet and infrared light, it is not necessary for the product to have a visible color. Early spectrochemical assays typically measured the absorbance after conversion of all of the analyte to the light-absorbing product. Modern assays usually use rate spectrophotometry, taking multiple absorbance measurements over time to determine the rate of change in light absorbance as the reaction proceeds. During the initial phase of the reaction, this rate is constant and proportional to the initial concentration of the analyte. This significantly reduces the time needed to obtain a result because it is not necessary for the reaction to go to completion, and it allows the averaging of multiple measurements, improving precision. Furthermore, it is unaffected by nonreacting substances that absorb light at the test wavelength because the absorbance of the nonreacting substances is constant and does not contribute to the rate of change in the absorbance.
Rate spectrophotometry remains subject to interference by substances that react to produce light-absorbing products, thereby falsely increasing the apparent concentration. Substances that inhibit the assay reaction or that consume reagents without producing a light-absorbing product give falsely low results. For example, ascorbic acid produces negative interference in many spectrophotometric assays that use oxidation reactions to generate colored products.
One way to improve the selectivity of a spectrochemical assay is to increase the selectivity of the reaction that generates the light-absorbing product. Enzymes, which can catalyze highly selective reactions, are often used for this purpose. For example, many assays for ethanol use alcohol dehydrogenase (ADH) to catalyze the oxidation of ethanol to acetaldehyde, with concomitant reduction of the cofactor NAD+ (oxidized form of nicotinamide adenine dinucleotide) to NADH (reduced form of nicotinamide adenine dinucleotide). The initial rate of increase in light absorption produced by the conversion of NAD+ to NADH is proportional to the concentration of ethanol. Although other alcohols, such as isopropanol and methanol, can also be oxidized by ADH, they are much poorer substrates for ADH with low rates of reaction and correspondingly low levels of interference.
Many other enzymatic assays also rely on measuring the change in light absorption at 340 nm when NAD+ is converted to NADH or vice versa. These include enzymatic assays for ethylene glycol, as well as some enzyme-linked immunoassays, such as EMIT (enzyme-multiplied immunoassay technique) assays. All such assays are potentially subject to interference by specimens with high concentrations of lactate. Lactate dehydrogenase, which is naturally present in serum, will oxidize this lactate to pyruvate if NAD+ becomes available for simultaneous reduction to NADH. When a serum specimen with high lactate is mixed with assay reagents that contain NAD+, oxidation of the lactate contributes to the total rate of NADH production. The increased rate of NADH production results in a false increase in the measured concentration of the target analyte.
Some enzymatic reactions do not produce a colored product. Enzymes such as glucose oxidase or lactate oxidase couple oxidation of the substrate to reduction of oxygen to hydrogen peroxide, which is colorless. A coupled second reaction is then necessary using the peroxide to convert a colorless dye to a colored one. Oxidase-based reactions may be subject to interference by compounds with high structural similarity to the target analyte. For example, glycolate, a toxic metabolite of ethylene glycol, is an excellent substrate for lactate oxidase and will give falsely high lactate results when it is present.
The need to measure very low concentrations of an analyte with a high degree of specificity led to the development of immunoassays. The combination of high affinity and high selectivity makes antibodies excellent assay reagents. There are two common types of immunoassays: noncompetitive and competitive. In noncompetitive immunoassays, the analyte is sandwiched between two antibodies, each of which recognizes a different epitope on the analyte. In competitive immunoassays, analyte from the patient’s specimen competes for a limited number of antibody binding sites with a labeled version of the analyte provided in the reaction mixture. Because most drugs are too small to have two distinct antibody binding sites, drug immunoassays are usually competitive.
In competitive immunoassays, increasing the concentration of xenobiotic in the specimen results in increased displacement of labeled xenobiotic from the antibodies. The amount of xenobiotic in the specimen can be determined by measuring either the amount of label remaining bound to the assay antibodies or the amount of label free in solution. In the earliest immunoassays, the label was a radioisotope, typically iodine-125, tritium, or carbon-14. Today, radioimmunoassays are relatively uncommon because of problems associated with handling and disposal of radioactivity. Nonisotopic immunoassays are currently the most widely used methodologies for the measurement of drugs. They offer high selectivity and good precision and are readily adapted to automated analyzers, thereby decreasing both the cost and the turnaround time of the assays. The xenobiotics for which immunoassays are available are limited to those for which there is a high demand, such as widely monitored therapeutic drugs and the drugs of abuse included in workplace drug screening. However, because production costs are relatively low, these tests are widely distributed at reasonable prices.
The most widely used nonisotopic drug immunoassays are in the category of homogenous immunoassays. Homogenous immunoassays measure differences in the properties of bound and free labels rather than directly measuring one or the other after their physical separation. Avoiding a separation step allows homogenous immunoassays to be readily adapted to automated analysis. Homogenous techniques that are in wide use include EMIT (Fig. 6–1), kinetic inhibition of microparticles in solution (KIMS), and cloned enzyme donor immunoassay (CEDIA).
Enzyme-multiplied immunoassay technique (EMIT) immunoassay. The drug to be measured is labeled by being attached to the enzyme glucose-6-phosphate dehydrogenase (G6PD) near the active site. (A) Binding of the enzyme-labeled drug to the assay antibody blocks the active site, inhibiting conversion of NAD+ (oxidized form of nicotinamide adenine dinucleotide) to NADH (reduced form of nicotinamide adenine dinucleotide). (B) Unlabeled drug from the specimen can displace the drug–enzyme conjugate from the antibody, thereby unblocking the active site and increasing the rate of reaction.
Many of the newest automated immunoassays are again using physical separation techniques. In these assays, the detection antibody is physically attached to a solid support, and separation occurs by a simple wash step. This wash step removes the patient’s serum along with many potentially interfering substances. Older assays of this type used antibodies bound to large plastic beads or wells of microtiter plates and required long incubation steps because of substantial times required for diffusion of the reactants to the antibodies. Newer assays typically use latex microparticles that have very high total surface areas, allowing rapid equilibration and short assay times.
Figure 6–2 shows a schematic magnetic microparticle enzyme-labeled chemiluminescent competitive immunoassay. A single enzyme label can generate many photons, allowing high signal amplification. Coupled with a background luminescence that is essentially zero, such assays can measure concentrations below the nanomolar level. Many variations of this approach are in use. Enzyme substrates may be used that result in fluorescent or colored products. Enzymes other than alkaline phosphatase may be used as labels, or nonenzymatic fluorescent, chemiluminescent, or electroluminescent labels may be used. These new techniques are readily automated and have higher sensitivities than homogenous immunoassays.
Magnetic microparticle chemiluminescent competitive immunoassay. (A) Unlabeled drug from the specimen competes with alkaline phosphatase–labeled drug for binding to antibody-coated magnetic microparticles. The microparticles are then held by a magnetic field while unbound material is washed away. (B) A dioxetane phosphate derivative is added and is dephosphorylated by microparticle-bound alkaline phosphatase to give an unstable dioxetane product that spontaneously decomposes with emission of light. The rate of light production is directly proportional to the amount of alkaline phosphatase bound to the microparticles and inversely proportional to the concentration of competing unlabeled drug from the specimen.
Microparticle capture assays are a type of qualitative competitive immunoassay that have become very popular, especially for urine drug screening tests. The use of either colored latex or colloidal gold microparticles enables the result to be read visually as the presence or absence of a colored band, with no special instrumentation required. Competitive binding occurs as the assay mixture is drawn by capillary action through a porous membrane. This design feature is responsible for alternate names for the technique: lateral flow immunoassay or immunochromatography.
The simplest microparticle capture design uses an antidrug antibody bound to colored microparticles and a capture zone consisting of immobilized drug (Fig. 6–3). If the specimen is xenobiotic free, the beads will bind to the immobilized analyte, forming a colored band. When the amount of drug in the patient specimen exceeds the detection limit, all of the antibody sites will be occupied by drug from the specimen, and no labeled antibody will be retained in the capture zone. The use of multiple antibodies and discrete capture zones with different immobilized analytes can allow several xenobiotics to be detected with a single device.
Microparticle capture immunoassay. (A) Diagram of a device before specimen addition. Colored microbeads (about the size of red blood cells) coated with antidrug antibodies (
) are in the specimen well. At the far end of a porous strip are capture zones with immobilized drug molecules (•) and a control zone with antibodies recognizing the antibodies that coat the microbeads. (B
) Adding the urine specimen suspends the microbeads, which are drawn by capillary action through the porous strip and into an absorbent reservoir (hatched area
) at the far end of the strip. In the absence of drug in the urine, the antibodies will bind the beads to the capture zone containing the immobilized drug and form a colored band. Excess beads will be bound by antibody–antibody interactions in the control zone, forming a second colored band that verifies the integrity of the antibodies in the device. (C
) If the urine contains the drug (•) in concentrations exceeding the detection limit, all of the antibodies on the microbeads will be occupied by drug from the specimen, and the microbeads will not be retained by the immobilized drug in the capture zone. No colored band will form. However, the beads will be bound and form a band in the control zone.
A disadvantage of this design is the potential for causing confusion because a positive test result is indicated by the absence of a band. More complex (and more expensive) variations have been developed in which a colored band denotes a positive test result.
Although immunoassays have a high degree of sensitivity and selectivity, they are also subject to interferences and problems with cross-reactivity. Cross-reactivity refers to the ability of the assay antibody to bind to xenobiotics other than the target analyte. Xenobiotics with similar chemical structures may be efficiently bound, which can lead to falsely elevated results. In some situations, cross-reactivity can be beneficially exploited. For example, some immunoassays effectively detect classes of drugs rather than one specific drug. Immunoassays for opioids use antibodies to morphine that cross-react to varying degrees with structurally related substances, including codeine, hydrocodone, and hydromorphone. Oxycodone typically has low cross-reactivity, and higher concentrations are required to give a positive result. The cross-reactivity of non-morphine opiates varies with manufacturer. Consult with your lab for the relative sensitivities of the immunoassay it uses. Structurally unrelated synthetic opioids, such as meperidine and methadone, have little or no cross-reactivity and are not detected by opiate immunoassays. Immunoassays for the benzodiazepine class react with a wide variety of benzodiazepines but with varying degrees of sensitivity.12,16 Because of the highly variable response of immunoassays to the various opiates and benzodiazepines, methods based on mass spectrometry should be used for definitive results.
Class specificity can be a two-edged sword. Assays for the TCA family have similar reactivity with amitriptyline, nortriptyline, imipramine, and desipramine and can be used to provide an estimate of the total concentration of any combination of these drugs. To account for nonuniform cross-reactivity, such results of these assays are usually reported as concentration ranges (eg, <100 ng/mL, 100–300 ng/mL). A large number of other drugs with tricyclic structures, including carbamazepine, many phenothiazines, and diphenhydramine, also cross-react and generate a signal, particularly at concentrations found in patients who overdose. Qualitative tests, such as microparticle capture assays, may then yield false-positive results if the signal generated by the cross-reacting drug (eg, carbamazepine) exceeds the detection limit of the immunoassay. With quantitative or semiquantitative assays, however, the apparent concentration produced by a cross-reacting drug is generally well below TCA concentrations associated with toxicity.
Even when an antibody is selected to be specific to a single drug, it is common that metabolites of the target drug show some cross-reactivity. This, too, may be beneficial. When the metabolite is an active one (eg, carbamazepine epoxide), the contribution of its cross-reactivity may yield results that correlate better with the drug effect than the true concentration of the parent drug alone.
Immunoassays are also subject to interference by substances that impair detection of the label. Elevated lactate concentrations may lead to spuriously increased drug concentrations in specimens tested by EMIT, as described earlier. Immunoassays that rely on enzyme labels are particularly sensitive to nonspecific interference because enzyme activity is highly dependent on reaction conditions. A number of substances that can inhibit the enzyme reaction in EMIT assays are used to adulterate urine submitted for drug abuse testing with the intent of producing false-negative results (see the discussion of drug-abuse screening tests under “Special Considerations for Drug Abuse Screening Tests” later). Such adulteration may be detected when the rate of reaction is lower than the rate observed with a drug-free control.
Chromatography encompasses several related techniques in which analyte specificity is achieved by physical separation. The unifying mechanism for separation is the partition of the analytes between a stationary phase and a moving phase (mobile phase). In most instances, the stationary phase consists of very fine particles arranged in a thin layer or enclosed within a column. The mobile phase flows through the spaces between the particles. Analytes are in a rapid equilibrium between solution in the mobile phase and adsorption to the surfaces of the particles. They move when in the mobile phase and stop when adsorbed to the stationary phase. The average velocity of the analyte xenobiotics depends on the relative time spent in the moving versus stationary phase. Xenobiotics that partition primarily into the mobile phase have average velocities slightly lower than the mobile phase velocity. Average velocity decreases as the proportion of time adsorbed to the stationary phase increases. Under controlled conditions, these average velocities are highly reproducible. Xenobiotics may be provisionally identified based on their characteristic velocity, as measured by the amount of time required to traverse the length of a chromatography column (retention time). Chromatography is a separation method and must be combined with a detection method to allow identification and measurement of the separated substances.
Chromatographic behavior is sufficiently reproducible that the failure to detect a signal at the retention time characteristic of a compound effectively excludes the presence of that compound in amounts greater than the detection limit. On the other hand, a number of different substances may have migration velocities that are identical or nearly so. A positive finding is therefore not completely specific. Definitive identification depends on having additional information, which may be obtained through selective detection techniques or by confirmatory testing using a second method. The sensitivity of chromatographic methods depends on both the amount of specimen available and the sensitivity of the detection method. A major advantage of chromatographic techniques is that multiple xenobiotics may be detected and measured in a single procedure. Nor is it necessary to know in advance the specific xenobiotic to be looked for. For this reason, chromatographic techniques have a major role in screening for multiple xenobiotics.
Most chromatographic procedures require extraction and concentration of the xenobiotics to be analyzed before the chromatography is done. Extraction results in removal of salts, proteins, and other materials that may exhibit unfavorable interactions with either of the chromatographic phases. Concentration allows the substances to be introduced in a narrow “band,” so that compounds with slightly different relative mobilities become completely resolved, or separated from one another, rather than overlapping. This also results in a more intense signal as a band passes through the detector and increases sensitivity.
Extraction of drugs is most commonly done with organic solvents, but “solid-phase extraction” is also very popular.11 Solid-phase extraction is itself a modified chromatographic procedure in which a urine or serum specimen is passed through a short chromatography column with a hydrophobic stationary phase. Most drugs are sufficiently hydrophobic to partition almost completely into the stationary phase and are retained on the column. Subsequently, the retained xenobiotics are eluted with an organic solvent. The organic solvents from either extraction technique are evaporated to concentrate the extract. The extraction process allows the analyte from a large volume of specimen to be concentrated. Detection sensitivity can thereby be increased provided large-volume specimens can be readily obtained, as is true with urine.
Often a preextraction treatment is used to increase the hydrophobicity of the substances to be extracted. The most common manipulation is pH adjustment, either upward or downward, to convert charged forms of drugs into uncharged, hydrophobic, and therefore extractable ones. In other instances, enzymatic or chemical hydrolysis may be used to convert water-soluble glucuronide metabolites back to their more readily extracted parent compounds, for example, conversion of morphine glucuronide to morphine.
In the technique of high-performance liquid chromatography (HPLC), a stationary phase is packed into a column and the mobile phase is pumped through under high pressure (Fig. 6–4). This allows good flow rates to be achieved even when solid phases with very small particle sizes are used. Smaller particle size increases surface area, decreases diffusion distances, and improves resolution, but the spaces between the particles are also smaller, increasing the resistance to flow. The use of high pressure and small particles allows good separations while keeping assay time short.
High-performance liquid chromatography (HPLC). HPLC is schematically shown. (A) A mixture of three compounds (
) is injected into a column filled with a spherical reversed-phase packing. (B
) The compounds move through the column at characteristic speeds. The most hydrophilic compound (
) moves most quickly, and the most hydrophobic compound (
) moves most slowly. (C
) The compound of intermediate polarity (
) has reached the detection cell, where it absorbs light directed through the cell and generates a signal proportional to its concentration. (D
) Illustration of the HPLC tracing that might result: 1
indicates the time of injection. The artifact at 2
results when the injection solvent reaches the detector and indicates the retention time of a completely unretained compound. The peaks at 3
, and 5
correspond to the separated compounds. For example, peak 4
might be amitriptyline
, peak 3
might be the more polar metabolite, nortriptyline
, and peak 5
could be the more hydrophobic internal standard N
-ethylnortriptyline. Later-emerging peaks are typically wider and shorter because of more time for diffusive forces to spread out the molecules.
HPLC drug assays typically use “reverse-phase” chromatography. Early chromatographic techniques typically used relatively polar silica gel particles as the stationary phase, with organic solvent mobile phases. Reverse-phase chromatography uses stationary phases consisting of silica gel particles that have had hydrocarbon molecules covalently linked to the outer surface. This coats the particles with a permanently bonded oil-like layer. Mobile phases are primarily aqueous with varying amounts of organic solvent. Because of these modifications, hydrophobic xenobiotics are more strongly adsorbed by the stationary phase, but hydrophilic ones tend to remain in the mobile phase. This results in an order of elution from the column that is approximately the reverse of that seen with unmodified silica gel and organic solvents. Thus, the term reverse-phase chromatography is used. HPLC can be done using either “normal-phase” or “reverse-phase” conditions, but reverse-phase conditions are much more commonly used. A variety of hydrocarbons can be used to derivatize the silica gel. By far, the most common reverse-phase columns use an octadecyl hydrocarbon as the outer coating and are often referred to as C-18 columns.
In HPLC, the xenobiotics are detected after they exit the chromatographic column. In this case, they are identified by their retention time (the characteristic time required to traverse the column). Because most xenobiotics absorb ultraviolet light, detection is commonly by ultraviolet spectroscopy using specially designed flow-through cuvettes. Measuring light absorbance at a selected wavelength allows the amount of the xenobiotic to be determined. Accuracy is often enhanced by comparing the absorbance of the target analyte with absorbance of an internal standard (ie, a compound with a different retention time that is added in a fixed amount to all specimens). The ratio of the drug absorbance to the internal standard absorbance is proportional to the drug concentration in the specimen.
Although most HPLC detectors allow a selection of the detection wavelength, only one wavelength is commonly used during a given run. Some detectors, however, allow absorbance at multiple wavelengths to be determined by breaking white light into its component wavelengths after it has passed through the detection cuvette and then making measurements at multiple wavelengths simultaneously using a light-sensitive chip similar to those used in digital cameras. This allows the absorbance spectrum of a compound to be determined as it elutes from the column. This information can supplement the retention time and allow more specific identifications to be made.
HPLC is often the method of choice for measuring serum concentrations of xenobiotics for which no immunoassay is available. However, it is limited by an inability to analyze drugs with a wide range of polarities in a single assay or to fully resolve substances with very similar polarity, both of which limit its usefulness as a broad drug-screening technique.
GC is similar in principle to HPLC except that the moving phase is a gas, usually the inert gas helium but occasionally nitrogen. The schematic illustration of HPLC in Fig. 6–4 is also applicable to GC. The low flow resistance of gas allows high flow rates that make possible substantially longer columns than are used in HPLC. This offers the dual advantages of high resolution and fast analysis. As was true in HPLC, most GC assays incorporate an internal standard to increase precision.
Because the inert carrier gas does not engage in intermolecular interactions, partition of the analytes into the moving gas phase depends primarily on their natural volatility. Elevated column temperatures are required to achieve sufficient volatility for analysis of most xenobiotics. The use of a temperature gradient (the column temperature is programmed to increase throughout the course of the analysis) can allow xenobiotics with a wide range of volatility to be analyzed in a single run. This feature, coupled with excellent resolution, makes GC suitable for screening assays that encompass a broad range of drugs.
GC is limited to xenobiotics that are reasonably volatile at temperatures below 572°F (300°C), above which the stationary phase may begin to break down. Two principal attributes of a xenobiotic limit its volatility: its size and its ability to form hydrogen bonds. Xenobiotics that form hydrogen bonds via amino, hydroxyl, and carboxylate moieties can be made more volatile by replacing hydrogens on oxygen and nitrogen atoms with a nonbonding, preferably large, substituent. (Large substituents sterically hinder access to the acceptor electron pairs on the nitrogen and oxygen atoms.) A number of derivatizing agents can be used to add appropriate substituents. The most common derivatives involve the trimethylsilyl (TMS) group. Although derivatization with TMS substantially increases the molecular weight, the resulting derivative is much more volatile as a consequence of the loss of hydrogen bonding.
In traditional packed-column GC, the packing may consist of inert support particles with a thin coating of nonvolatile, high-molecular-weight oil that comprises the stationary phase. It is increasingly common for the stationary phase to be covalently bonded to the support particles. A highly useful variant of GC is capillary chromatography. A long, thin capillary tube of fused silica is coated on the inside with a covalently bonded stationary phase. The mobile gas phase flows through the tiny channel in the middle. These capillaries are flexible, allowing very long columns (≥10 m) to be coiled into a small space. The long column length, coupled with highly uniform conditions throughout the column, results in extremely high resolution. The small diameter of the column allows rapid thermal equilibration and the use of steep temperature gradients that can speed analysis. The major drawback to capillary chromatography is a very limited column capacity. Special techniques are needed to restrict the amount of material introduced into the column and thereby to avoid overloading it. High-sensitivity detectors are required to measure the small quantities that can be chromatographed.
A number of detectors are available for GC. The most common detector, particularly for packed columns, is the flame ionization detector. This involves directing the outflow of the column into a hydrogen flame. Organic molecules emerging from the column are burned, creating charged combustion intermediates that can be measured as a current. The amount of current flow is largely determined by the mass of carbon that is being burned. Nitrogen–phosphorus detectors are also widely used in drug analysis. In this modification of a flame ionization detector, a heated bead coated with an alkali metal salt is used to selectively generate ions from xenobiotics containing nitrogen or phosphorus. These devices detect broad ranges of substances but do not identify them. The identity of the compounds detected must be inferred from the retention time.
The mass spectrometer can serve as a highly sensitive GC detector and possesses the ability to generate highly characteristic mass spectra from the compounds it is detecting. A special requirement of the mass spectrometer is that it requires a high vacuum to prevent the ionic particles that it creates from interacting with other molecules or ions. This requires removal of the inert carrier gas and is easiest when there is a low total gas flow, such as occurs with capillary GC. The mass spectrometer in turn provides good sensitivity for the small amounts of analyte that can be accommodated in capillary GC.
The detection process begins by generating ions from the analyte. This is usually done using electron impact ionization. The gas phase analyte is separated from the bulk of the carrier gas and introduced into an ionization chamber, where it is bombarded by a stream of electrons. Electron impact can dislodge an electron from the analyte, creating a positively charged ion and frequently imparting sufficient energy to the ion to break it into pieces. If fragmentation occurs, conservation of charge requires that one of the resulting fragments be a positively charged ion. The fragments into which a molecular ion can break are characteristic of the xenobiotic as are the relative probabilities that any given fragment will carry the positive charge.
The mass spectrometer then uses electromagnetic filtering to direct only ions of a specified mass-to-charge (m/z) ratio to a detector. Because most of the ions produced have a single positive charge, the observed peaks generally correspond to the mass of the ions. The detector has sufficient electronic amplification that a single ion could theoretically be detected, accounting for the high sensitivity of mass spectrometric detection. By rapidly scanning through a range of masses that are sequentially allowed to reach the detector, a mass spectrum may be generated. The mass spectrum records the masses of the pieces produced by fragmentation of the parent ion, as well as the relative frequency with which these fragments are produced and detected. The highest mass observed in the spectrum usually corresponds to the mass of intact parent ions generated from collisions that were not energetic enough to cause fragmentation.
Figure 6–5 shows the mass spectrum obtained from a gas chromatograph at a time when the TMS derivative of the cocaine metabolite benzoylecgonine was emerging from the capillary column. The mass spectrum of any compound is highly distinctive and usually unique. The primary exception involves optical enantiomers, both of which have the same mass spectrum. Toxicologically significant examples of enantiomers include d-methamphetamine, a drug of abuse, and l-methamphetamine, which is found in decongestant inhalers. It is also important to distinguish dextrorphan, the major metabolite of the cough suppressant dextromethorphan, and levorphan (levorphanol), a controlled substance.
Mass spectrum of the trimethylsilyl derivative of benzoylecgonine (TMS-BE). (A) Mass spectrum of effluent from a gas chromatography (GC) column at the retention time of TMS-BE. The unfragmented parent ion of TMS-BE is at a mass-to-charge (m/z) ratio of 361. The two fragment peaks at m/z 243 and 259 result from fracture of the bonds at X and Y, respectively, in structure of TMS-BE (inset in B). Additional peaks at m/z 243, 259, and 364 are derived from trideuterated TMS-BZE (d3-TMS-BE) added as an internal standard. The mass spectrometer can identify and quantify TMS-BE and d3-TMS-BE independently of one another by measuring the heights of the peaks unique for each compound. The peak at m/z 425 is from a coeluting contaminant. (B) Mass spectrum of pure TMS-BE.
To avoid the need to scan the full range of masses in a typical mass spectrum, selected ion monitoring is often used. Here, the mass spectrometer is typically programmed to filter and detect only three of the larger and more characteristic peaks in the mass spectrum. In the case of TMS-benzoylecgonine (TMS-BE), the peaks at m/z 240, 256, and 361 are used. The concentration of TMS-BE in the specimen is determined from the ratio of the peak height at m/z 240 to height of a peak at m/z 243 that results from a corresponding fragment of the internal standard, which is TMS-BE that is labeled with three deuterium atoms (d3-TMS-BE) to give it a mass distinct from that of TMS-BE from the specimen (Fig. 6–5). The specificity of the identification is verified by finding peaks at m/z 256 and m/z 361, with peak height ratios to the peak at m/z 240 comparable to the ratios seen with authentic TMS-BE. The detection at the correct retention time of a xenobiotic producing all three peaks in the correct ratios produces an extremely specific identification.
The high sensitivity and specificity afforded by GS/mass spectrometry is being further extended by the related hybrid technique of liquid chromatography/tandem mass spectrometry, often abbreviated as LC/MS/MS.18 This technique is being used in an increasing number of toxicology laboratories. In LC/MS/MS, a tandem mass spectrometer is used as the detector for liquid chromatography system. The initial ionization is done under conditions that do not promote fragmentation and is commonly achieved by adding or removing a proton rather than forcefully dislodging an electron. The resulting ions have a mass that differs from that of the parent molecule by one mass unit ([M+H]+ or [M–H]−). The first mass spectrometer is used to selectively filter only unfragmented ions with the expected molecular mass. As the selected ions exit the first mass spectrometer at high speed, they are allowed to collide with molecules of an inert gas. These collisions cause the ions to break apart to create the fragment mass spectrum that is detected by the second mass spectrometer. The additional selection step provided by the first mass spectrometer greatly enhances specificity and reduces background signal, enhancing sensitivity.