Gregory L. McIntire

Lurasidone is an atypical antipsychotic that was approved by the FDA in 2010 to treat bipolar depression and schizophrenia. Like other antipsychotics, adherence to lurasidone is critical for successful disease treatment. Thus, therapeutic drug monitoring (blood testing) is often employed by clinicians to monitor adherence. Urine drug testing, with its advantages over blood testing, is another method used to confirm medication adherence. However, analytes used in blood testing are often very different than those used for testing in urine, where nonactive metabolites are often most prevalent. Choosing metabolites in urine that are relatively prevalent affords optimal method sensitivity, and thus improved testing results for adherence. To ensure optimal lurasidone adherence testing, an untargeted high-resolution mass spectrometry method was employed, using known positive human urine samples to identify the lurasidone metabolites and their relative abundance in urine. This testing identified a different primary urine metabolite from what has been reported in blood. The higher prevalence of this metabolite will improve lurasidone urine adherence monitoring.

	Medication monitoring has become increasingly important for successful treatment of patients with mental health diseases because adherence to treatment is generally poor, especially in the schizophrenic population (1–8). Urine has become an alternative to blood or plasma medication monitoring due to its noninvasive nature and ease of collection. Whereas blood or plasma drug testing usually involves the identification and quantitation of the parent compound or active metabolites, or both, the success of urine drug testing (UDT) is largely dependent on analysis of any metabolites of the parent compound. Although the parent compounds may be present in urine, often they are at very low concentrations relative to metabolites, and therefore do not provide the sensitivity required for medication monitoring. Urine metabolites are often predicted from identification in blood, plasma, or specific testing methods, such as gas chromatography-mass spectrometry (GC-MS), extractions, radioactivity, and using in vitro or animal samples. However, it has been shown that these methods are not always successful in identifying the most abundant urinary metabolite (9–12). Without suitable metabolites to test, a negative UDT result could prompt a clinician to alter treatment for a patient when treatment need not be altered. Therefore, a generic, untargeted approach is useful for the successful identification of urinary metabolites suitable for highly sensitive medication monitoring. Liquid chromatography–high resolution mass spectrometry (LC–HRMS) provides a sensitive and nonspecific detection method for setting up such an experiment.

	Lurasidone (Latuda) is an atypical antipsychotic that was approved for the treatment of acute symptoms of schizophrenia (13,14) and bipolar depression (15,16) in 2010 and 2013, respectively. It is commercially available as 20 mg, 40 mg, 60 mg, 80 mg, and 120 mg tablets, and is typically prescribed or administered at 40 or 80 mg per day. It is absorbed after oral administration with a bioavailability of 9–19%. Dosing is designed to be with food, which can increase the bioavailability by 100%. The mean elimination half-life is 18 h. Steady state serum concentrations for lurasidone are typically achieved after seven days of dosing (17–20). Lurasidone is metabolized in the liver primarily by CYP3A4. Metabolism includes oxidative N-dealkylation, hydroxylation of the norborane ring, S-oxidation, and reductive cleavage of the isothiazole ring, followed by S-methylation. Nearly two dozen metabolites of lurasidone have been previously identified, and only ~9% of the dose is excreted in urine (17–20). Typically, adherence to lurasidone therapy is monitored by evaluating levels of lurasidone and M11/ID-20219 (one of its metabolites) that were each predicted to be present in urine at approximately 12 and 24%, respectively. The structures for lurasidone and many of the confirmed metabolite structures can be seen in Table I.

	Previously, we reported the identification of novel metabolites for monitoring aripiprazole, brexpiprazole, haloperidol, and quetiapine in urine that were not originally predicted (9–12). Because there are some similarities of these antipsychotics to lurasidone, we decided to determine if the urinary lurasidone compound(s) predicted from plasma studies were indeed the most abundant prior to development of a confirmation method. This work reports the identification of lurasidone and prevalent lurasidone metabolites in urine using LC–HRMS from patients prescribed lurasidone. Additionally, confirmation of the most prominent metabolites was tested in a validated, targeted, quantitative liquid chromatography–tandem mass spectrometry method (LC–MS/MS), which are at odds with current reports of urine metabolites (17–20).

 were purchased from Cerilliant (Round Rock, Texas). Hydroxylurasidone was a custom synthesis product purchased from 

C Molecular (Greensboro, North Carolina). All solvents, including methanol (optima grade), formic acid (88%), acetonitrile (optima grade), ammonium acetate (optima grade), and isopropanol (optima grade), were purchased from VWR (Radnor, Pennsylvania, USA). Drug-free human urine was acquired from UTAK Laboratories (Valencia, California). Standards for S-methyl lurasidone and S-methyl hydroxylurasidone were not commercially available, and synthesis requests were unsuccessful.

	Identification of lurasidone metabolites using LC–HRMS was completed on 13 authentic urine samples from patients who were prescribed the medication. After metabolite identification was complete, an LC–MS/MS confirmation was validated. An additional 56 patients were prescribed lurasidone at different doses, with specimens collected over three separate days for each patient used to confirm the accuracy of the method. These samples were provided voluntarily, and anonymously, to assist with the development of a lurasidone confirmation method. No identifying or demographic information was collected on these volunteers, other than the prescribed lurasidone dose. There was an alphanumeric code from the clinic that was provided to track the patients who provided samples over the course of the three separate days. None of the results were shared with the clinician to assist with treatment. Ameritox is accredited by the College of American Pathologists (CAP) and abides by CAP, Clinical Laboratory Improvement Amendments (CLIA), and Health Insurance Portability and Accountability Act (HIPAA) requirements. Due to the secondary analysis nature of this work and the absence of clinical conclusions, neither the United States Food and Drug Administration (FDA) nor other clinical trial review or approval was obtained by Ameritox. Writing this manuscript did not involve human subjects, as defined by the U.S. Code of Federal Regulations (45 

46.102); thus, an Institutional Review Board (IRB) approval of these specific research activities was unnecessary.

	Thirteen patient urine specimens (100 µL) were diluted 5X with 400 µL of a reference standard, (0.25 µg/mL of hydrocodone-d

 was used as an internal reference standard for all LC–HRMS injections, to guarantee successful injection of the sample, and provide a retention time marker. Prepared samples were injected (5 µL) and separated on a Phenomenex Kinetex Phenyl-Hexyl, 2.1 x 50-mm, 2.6-µm column (Torrance, California) at 50 °C, and analyzed on an Agilent 6530 Q-TOF (quadrupole time-of-flight mass spectrometer) with an Agilent 1290 LC system (Santa Clara, California). The LC–QTOF method conditions are detailed in a previous publication (12). A lurasidone control in drug-free urine (75 ng/mL) was run, along with the patient samples, to assist in positive identification of the parent compound, if present. No other standards were available or purchased to assist in identification, until a confirmation method was developed. Each sample was injected and analyzed twice.

	The MS-only data were processed using Agilent Mass Hunter Qualitative Analysis and PCDL (Personal Compound Database and Library) manager software. A database of lurasidone and 11 of its possible metabolites' chemical formulas (Table I) was compiled, and used to search against the samples. The software matched compounds based on retention time (if available), mass (±20 parts per million or ppm), the isotopic distribution pattern, and the isotopic spacing theoretically derived from the chemical formula. To be identified as positive and a potential lurasidone metabolite, a compound had to have consistent retention times across multiple patient samples when a known retention time was lacking; otherwise, the retention times had to be within ±0.05 minutes of a control. The mass accuracy had to be within ±20 ppm; and the composite score of the mass accuracy and isotopic features had to be ≥70 (out of a possible 100). Compounds that had the highest area counts were also ranked and noted as the most abundant. To assist with differentiation of structural isomers, such as M8/M9 and M10, fragmentation spectra were obtained and reviewed to identify which isomer was present at an identified metabolite peak, as needed.

	Hydroxylurasidone was received as a neat solid that was dissolved into methanol at a concentration of 1 mg/mL, and lurasidone was received as a 100 µg/mL methanolic standard. Hydroxylurasidone and lurasidone were combined and diluted into a methanolic stock that was then further diluted into normal, drug-free human urine, to reach the appropriate calibrator (5, 25, 100, 500, and 1000 ng/mL) and quality control levels (75 ng/mL). Lurasidone-d8, 1 mg/mL methanolic stock, was diluted to 900 ng/mL in 0.1% formic acid in water solution. A 100 µL aliquot of the sample (patient sample, calibrator, or quality control stock) and 400 µL of lurasidone-d8 internal standard in 0.1% formic acid were added to a vial. Vials were then capped and vortexed for 10 s prior to injection of 5 µL.

	Samples were analyzed by LC–MS/MS on a Waters Acquity UPLC Xevo TQ-MS system (Waters Corporation, Milford, Massachusetts), a Waters Acquity UPLC CSH Phenyl-Hexyl 2.1 x 50-mm, 1.7-µm UPLC column. The LC method and MS conditions can be found in Strickland and associates (12). Analyte transitions are listed in Table II. The acquisition method was run in dynamic multiple reaction monitoring (MRM) mode, in order to maximize the number of points across the various analyte peaks. The validation of this method followed CAP and CLIA guidelines (21–25), and an internal SOP (standard operating procedure) that has been described in detail elsewhere (26). It should be noted that, due to the lack of standards for S-methyl lurasidone and S-methyl hydroxylurasidone, they were unable to be validated, and estimates of their concentration were made by comparing the quantiative peak area ratio to the lurasidone calibration curve. These compounds were included as a proof of concept, to show their estimated prevalence and relative importance for lurasidone compliance in UDT for when standards might be available. Also, due to the lack of standards, the transition parameters were estimated from hydroxylurasidone and are not optimized.

	The metabolite identification from the 13 patients analyzed by LC-QTOF can be seen in Table III. Compounds identified with the highest confidence (>90%) are highlighted in green, while less confident (>70% but <90%) compounds are highlighted in yellow. The three most abundant compounds for each specimen and replicate are noted with a star-asterisk in the respective square. For unidentified or not confidently identified compounds (<70%) for a given specimen replicate, the field is blank. It is clear that, although lurasidone was identified in almost all of the samples, it was not consistently among the most abundantly identified compounds. Metabolite M11, the predicted major metabolite, was rarely confidently detected in these samples. Instead of lurasidone and M11, metabolites M21 (S-methyl lurasidone), M22 (S-methyl hydroxylurasidone), and isomer M8/M9 (hydroxylurasidone), or isomer M10 (lurasidone sulfoxide), were frequently detected. To determine whether hydroxylurasidone or the lurasidone sulfoxide (isomers) was present, the collected fragmentation data from the QTOF were analyzed, and are shown in Figure 1. The identification of a peak at 

182 (red circle in Figure 1) confirmed the isomer as hydroxylurasidone by indicating a fragmentation of the hydroxylated norborane ring. If the identity was the lurasidone sulfoxide, expected fragmentation peaks of 

 152 or 237 from the oxidized sulfur atom on the isothiazole ring structure would be present. Additionally, the unhydroxylated norborane ring would have an expected 

 of 166. The absence of those expected peaks (

 152, 237, and 166) in the spectra confirms the identity of the metabolite as hydroxylurasidone, and a custom synthesis of the molecule was requested to validate a confirmation method. S-methyl lurasidone and S-methyl hydroxylurasidone were also requested as custom synthesis products, but attempts to synthesize for the method were unsuccessful.

	Upon receiving the hydroxylurasidone standard, an LC–MS/MS method was developed and validated. The results of validation of lurasidone and hydroxylurasidone are shown in Table IV. Although S-methyl lurasidone and S-methyl hydroxylurasidone were included in the method, without standards, validation was unable to be completed, and is the reason for their exclusion from Table IV. To ensure the ability to successfully detect and quantify lurasidone and hydroxylurasidone in patient specimens, samples from 56 additional patients (from three separate collection days) were provided for testing with the validated method. The results of these patient analyses are summarized in Table V, and separated by the prescribed dose. It is clear that testing for hydroxylurasidone helps with positive confirmation of taking lurasidone medication. It also appears, from the estimated concentrations of the S-methyl lurasidone and S-methyl hydroxylurasidone, that confirmation would be easier with these metabolites, because they are more abundant than both lurasidone and hydroxylurasidone. However, the lack of standards for these compounds makes it impossible to currently validate a method for reporting UDT results for these compounds.


	Figure 1: Fragmentation data from sample 8 Identified M8-M9-M10 Peak.

	The advantages of HRMS analysis have been reviewed in the literature, including the extreme selectivity of such methods (10–12,27–30). Using this method, authentic urine samples of human subjects who were known to be taking chronic doses of lurasidone were tested for the presence of lurasidone and 11 possible metabolites. Due to the high mass resolving power and low mass error on the QTOF, compounds that have similar mass to charge ratios, but different chemical formulas, were differentiated with the searching algorithm (hydroxylurasidone and S-methyl lurasidone). Also, by eliminating extraction preparation methods, compound loss was mitigated. Using liquid as opposed to gas chromatography also helped ensure that compounds with low volatility can still be accurately analyzed. Surprisingly, neither of the predicted major urinary metabolites M5 nor M11 were found to be consistently excreted through human urine in large detectable amounts. This result could be due to the fact that M5 and M11 both have a carboxylic acid moiety that would be a possible glucuronidation target. This was considered, and the glucuronidated versions were searched for in patient samples and poorly identified. While this may indicate that the glucuronide metabolites of M5 and M11 are not present in urine in any appreciable amount, it could also be due to the poor ionizability of glucuronidated compounds. To confirm, an additional hydrolysis study could be completed to see if there is an increase in the prevalence of M5 and M11 in patients after hydrolysis. However, with the significant presence of hydroxylurasidone (M8/M9), S-methyl lurasidone (M21), and S-methyl hydroxylurasidone (M22), it seemed unnecessary to pursue hydrolysis as a means for analyzing for lurasidone compliance.

	The presence of these metabolites in urine was not well predicted from results in blood. Hydroxylurasidone was estimated to have a prevalence of 2.8% as the M8 isomer, and 0.4% as the M9 isomer; S-methyl lurasidone and S-methyl hydroxylurasidone were not predicted at any measurable amount (17–20). It does appear that the estimate of lurasidone at 12% might be reasonable, as all but one of the 13 patients in metabolite discovery had detectable amounts of the parent compound (17–20).

	To better understand the relative amounts of each metabolite, lurasidone, hydroxylurasidone, S-methyl lurasidone, and S-methyl hydroxylurasidone present in urine, the results from the 56 patients used during method validation were analyzed. These results in Table V show that hydroxylurasidone, S-methyl lurasidone, and S-methyl hydroxylurasidone are present at approximately 2x, 7x, and 5x times, respectively, relative to lurasidone. All of the compounds show a general increase in concentration and percent positivity rate with increasing doses. It is clear that S-methyl lurasidone and S-methyl hydroxylurasidone provide slightly better positivity correlations at lower doses, but hydroxylurasidone does appear to provide enough benefit to help compensate for lower prevalence of lurasidone. Therefore, with the lack of available standards for S-methyl lurasidone and S-methyl hydroxylurasidone, hydroxylurasidone was validated to assist in UDT for lurasidone compliance.

	We successfully identified prevalent lurasidone metabolites in urine. Based on those identifications, we successfully validated a method for the purpose of UDT monitoring of lurasidone. The hydroxylurasidone metabolite provides benefit for lurasidone UDT monitoring by being more prevalent in the urine than lurasidone by ~2x, and providing more consistent positivity correlation at lower lurasidone doses. Although other metabolites are present in the urine in large concentrations, standards for those compounds are not available at this time. However, the proof-of-concept work with S-methyl lurasidone and S-methyl hydroxylurasidone shows that they are ~5x and ~7x greater in abundance than lurasidone, respectively, and would provide even better positivity correlation at low doses.

	(1) M. Ko, and T. Smith, Urine Drug Monitoring in Patients on Prescribed Antipsychotic Medications. 29th Annual US Psychiatric and Mental Health Congress Oct 21-24 (

http://www.ingenuityhealth.com/wpcontent/uploads/2016/11/Mental_Health_Populations_Study_US_Psych_2016_Poster.pdf

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	(8) R.A. Millet, P. Woster, M. Ko, M. DeGeorge, and T. Smith, Adherence to Treatment with Antipsychotic Medications Among Patients with Schizophrenia, Major Depressive Disorder, or Biopolar Disorder. Poster Presentation. (US Psychiatric and Mental Heal Congress (USPMHC) San Diego, CA,2015).

	(9) J. McEvoy, R.A. Millet, K. Dretchen, A.A. Morris, M.J. Corwin, and P. Buckley, 

	(10) J.R. Enders, S.G. Reddy, E.C. Strickland, and G.L. McIntire, 

	(11) O.T. Cummings, E.C. Strickland, J.R. Enders, and G.L. McIntire, 

	(12) E.C. Strickland, O.T. Cummings, A,A. Morris, A. Clinkscales, and G.L. McIntire, 

	(14) S. Caccia, L. Pasina, and A. Nobili, 

	(15) R. Franklin, S. Zorowitz, A.K. Corse, A.S. Widge, and T. Deckersbach, 

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The World Journal of Biological Psychiatry

http://doi.org/10.1080/15622975.2017.1285050

	(17) Product Information: LATUDA oral tablets, lurasidone hydrochloride oral tablets. Sunovion Pharmaceuticals Inc. Marlborough, MA (2018).

Disposition of Toxic Drugs and Chemicals in Man 

(Biomedical Publications, Seal Beach, CA, 11th Ed., 2010), pp 1232–1233.

	(19) Sunovion Pharmaceuticals Inc. Center for Drug Evaluation and Research Approval Package (Lurasidone). 

https://www.accessdata.fda.gov/drugsatfda_docs/nda/2013/200603Orig1s010.pdf

	(20) T. Ishibashi, T. Horisawa, K. Tokuda, T. Ishiyama, M. Ogasa, R. Tagashira, K. Matsumoto, H. Nishikawa, Y. Ueda, S. Toma, H. Oki, N. Tanno, I. Saji, A. Ito, Y. Ohno, and M. Nakamura, 

	(21) Association of Public Health Laboratories. CLIA-Compliant Analytical Method Validation Plan and Template for LRN-C Laboratories. December 2013.

	(22) F.T. Peters, O.H. Drummer, and F. Musshoff, 

 (AACC Press, Washington, DC, 2nd edition 2003) pp 114–115.

	(24) U.S. Department of Health and Human Services, Food and Drug Administration. Guidance for Industry-Bioanalytical Method Validation (2001).

	(25) National Laboratory Certification Program (NLCP). Manual for Urine Laboratories. October 2010.

	(27) J.M. Colby, K.L. Thoren, and K.L. Lynch, 

	(28) E. Partridge, S. Trobbiani, P. Stockham, T. Scott, and C.A. Kostakis, 

	(29) J.M. Colby,K.L. Thoren, and K.L. Lynch, 

	(30) S.K. Manier, A. Keller, J. Schaper, and M.R. Meyer, 

 is with Ameritox, LLC, in Greensboro, North Carolina. 

 is with the Molecular Education, Technology and Research Innovation Center of North Carolina State University, in Raleigh, North Carolina. 

 is with Premier Biotech, in Minneapolis, Minnestota. Direct correspondence to: 

Articles

For lurasidone treatment adherence testing, an untargeted high-resolution mass spectrometry method was employed, using known positive human urine samples to identify the lurasidone metabolites and their relative abundance in urine.

Determination of the Relative Prevalence of Lurasidone Metabolites in Urine Using Untargeted HRMS

A novel "dilute-and-shoot" method is described for the analysis of "bath salts" sold as "legal" highs, including mitragynine and nine synthetic cathinones. The cycle time for this method is 2.5 min. No sample clean-up or extraction was performed. This assay monitors two tandem mass spectrometry (MS/MS) transitions for each of the following 10 analytes: alpha-PVP, butylone, ethylone, MDPV, mephedrone, methcathinone, methedrone, methylone, naphyrone, and mitragynine; and two internal standards: methylone D3 and alpha-PVP D8. This straightforward "dilute-and-shoot" liquid chromatography (LC)–MS/MS method produces acceptable limit of detection (LOD) and limit of quantitation (LOQ) limits without costly extraction and time-consuming concentration protocols.

	Cathinone is a monoamine alkaloid found in the Khat plant (

 (1). Khat is a shrub grown in eastern Africa and the southern Arabian Peninsula, which can have mild stimulant effects when the leaves are chewed (2). Cathinone is also a beta ketone analog of amphetamine and as such is reported to cause sympathomimetic effects as well as psychoactive effects common with amphetamine and amphetamine derivatives such as MDMA ("Ecstasy") and MDA ("Eve") (3). Chewing of the Khat plant has been linked to increased risk of heart-related diseases and duodenal ulcers (4). Synthetic variants of cathinone, commonly referred to as "bath salts" or as "novel psychoactive substances" (NPS) by public health personnel, are stronger and more dangerous than their natural counterpart (5). The first synthetic derivatives of cathinone were reported during the 1920s (6-8). These initial derivatives were designed for medical purposes; however, most were withdrawn as a result of abuse and obvious dependency among users. Interestingly, bupropion (Wellbutrin) is a "synthetic cathinone" and is widely used as an antidepressant and anti-anxiety medicine (2).

	More recently, synthetic cathinones (NPS) have expanded as popular drugs of abuse. Their ready availability in the local drug market and so-called head shops make them easy "legal" highs. These active agents are sold under various names such as pesticides and insect repellants. They are often labeled as "not for human consumption," to avoid regulatory control. However, the use of bath salts has significantly increased in recent years with poison control centers in the United States reporting 304 calls related to bath salts in 2010 and 1782 calls in the first four months of 2011 (9,10). It is believed that the increase in their use is a result of their availability, lack of proper methods for drug testing, and media coverage.

	Mitragynine is a naturally occurring substance found in the leaves of the kratom (

) plant, a member of the coffee family found in Southeast Asia, Indochina, and Malaysia (11). This "legal" high is readily available and is often used to mitigate pain, aid in opiate withdrawal, and for recreational purposes (12). This natural drug is regulated in a number of countries including Malaysia but not currently scheduled in the United States (13).

	Urine drug testing is a common means of monitoring for the use of these drugs. While forensic laboratories often have access to blood, urine, and other tissues to test in post-mortem subjects, pain management laboratories tend to focus on urine or oral fluids because of the ease of sampling and minimal stability requirements. As well as ease of sample collection, the sample preparation required is comparatively easier for urine than other patient specimens, for example, blood. Often such preparation consists of sample matrix removal using solid-phase extraction (SPE); however, reports of “dilute-and-shoot” testing confirm that direct analysis without sample preparation is possible for some analytes (14). In this study, a unique “dilute-and-shoot” liquid chromatography–tandem mass spectrometry (LC–MS/MS) method was developed and validated for the analysis of nine synthetic cathinones and mitragynine (Kratom) in patient urine samples. This method is ideal for a high-throughput production environment because it has a modest 2.5-min cycle time and requires low sample preparation time and cost while still preserving sensitivity and specificity of the quantitative analytical result. Above all, this method achieved complete chromatographic resolution of the isobars, ethylone and butylone, both of which are analyzed in this method, by lowering column temperature to 20 °C and cutting flow rate to 0.4 mL/min to maintain system pressure.

	All reference standard compounds, including internal standards, were purchased from Cerilliant. Solvents, including methanol, acetonitrile, and formic acid, were purchased from VWR. Normal drug-free urine was obtained from UTAK.

	Aliquots of normal drug-free urine were fortified with the reference standards at concentrations of 25 ng/mL, 250 ng/mL, and 1000 ng/mL as calibrators. All samples, calibrators, and controls were diluted 5× with the internal standard solution (0.5 µg/mL methylone D3 and alpha-PVP D8) by adding 400 µL of internal standard to 100 µL of sample.

	All analyses were performed via LC–MS/MS on Acquity TQD UPLC/MS/MS system (Waters). A 2.1 × 50 mm, 1.7-µm Acquity UPLC BEH C18 analytical column (Waters) was used for chromatographic separations. The run time for this method was 2.0 min with a total cycle time of 2.5 min. No sample extraction or clean-up was required before LC–MS/MS analysis. Two transitions were monitored for each of 10 analytes (alpha-PVP, butylone, ethylone, MDPV, mephedrone, d,l-methcathinone, methedrone, methylone, mitragynine, and naphyrone), plus two internal standards (methylone D3 and alpha-PVP D8), as shown in Table I. Elution solvents used in the gradient were 0.1% formic acid in 10% methanol (mobile phase A) and 0.1% formic acid in 50:50 methanol– acetonitrile (mobile phase B), with a flow rate of 0.4 mL/min (Table II).

	The method was validated by examining various parameters such as limits, linearity, precision and accuracy (P&A: Ten replicates of each of three concentrations each day for three days with coefficient of variance [%CV] of each set of P&A point replicates each day and across all P&A days falling within +/- 15%), interferences, and matrix effects. Drugs of abuse (benzoylecgonine, THC-A, amphetamine, methamphetamine, MDMA, MDEA, MDA, and phentermine) and therapeutic drugs (oxazepam, morphine, imipramine, buprenorphine, fentanyl, meprobamate, methadone, tramadol, gabapentin, pregabalin, and tapentadol) were tested as possible sources of interferences. Patient samples were evaluated by comparing the quantitative results of a previously used method with those of the new method as described herein. Since mitragynine and alpha-PVP were not evaluated in the previous method, a comparison was not possible. The previous method was developed and validated as described here but lacked critical analytes, for example, mitragynine and alpha-PVP. This method has not been reported in the literature. Detailed discussion of these validation parameters can be found in a paper by Enders and McIntire (14).

	The tolerance for all analytes at each concentration level was ± 25%. All daily calibration curves had correlation coefficient values (r

) greater than or equal to 0.99. The matrix blanks following the highest calibrator points did not show any signs of carryover, that is, no response greater than 50% of the limit of quantitation (LOQ).


	Figure 1: Overlaid chromatogram of 10 analytes determined in this assay. The chromatogram was obtained after injecting a sample at the limit of quantification (LOQ) of 25 ng/mL. Internal standards (IS) are not shown.

	Two mass transitions were selected for each of the analytes following infusion studies of pure standards dissolved in matrix. The first transition served as the quantifier ion transition and the second transition served as a qualifier ion transition (Table I). The ratio of the areas of the quantifier ion transitions for the analyte of interest and the internal standard (IS) were used to obtain the relative response by plotting the area ratio of the analyte-to-IS versus analyte concentration. The area of the second ion transition of the analyte was used to generate a specific relative ratio to the area of the quantifier peak. This generates a compound specific “ion ratio” for each analyte that together with retention time helps to confirm that the peak results from the compound of interest.


	Figure 2: Chromatogram showing the separation of the isomers ethylone and butylone. 

	This method generated calibration data with consistent peak areas and ion ratios for all analytes and internal standards. Daily three-point calibration curves were completed with correlation coefficient values (r

) of at least 0.99 for all analytes. Consistent accuracies of 100 ± 25% were obtained for all calibration points, negative and positive controls, as well as points generated daily for both calibration and controls. The limit of quantification (LOQ) was determined to be 25 ng/mL, which was sufficient to achieve consistent precision and accuracies as well as acceptable ion ratios, while the upper limits of linearity and of carryover (ULOL, ULOC) were 1000 ng/mL. The chromatogram of all analytes shown in Figure 1 provides clear evidence that there is good chromatographic separation and resolution of all analytes assessed in this assay even at the LOQ. Furthermore, Figure 2 shows the separation achieved for the ethylone and butylone isobars, while Table III shows a summary of the validation results.


	Table IV: Positive NPS data from a 14-month period of testing

	This work documents the successful development and validation of an improved LC–MS/MS method for routine, high-throughput analysis of alpha-PVP, butylone, ethylone, MDPV, mephedrone, d,l-methcathinone, methedrone, methylone, mitragynine, and naphyrone, using methylone D3 and alpha-PVP D8 internal standards. During the initial stages of method development, the column temperature was held at 50 °C with a flow rate of 0.8 mL/min, which required a much longer 4.0 min cycle time for complete resolution of all analytes of interest. This was especially true for the isobars butylone and ethylone, which did not exhibit robust separation even with the longer elution time. To ensure complete resolution (< 10% peak height overlap) of these two isobars while still achieving the desired 2.5 min cycle time, the column temperature was lowered to 20 °C and the flow rate was decreased to 0.4 mL/min. Traditional chromatography paradigms indicate that higher temperatures lead to better resolution (15). However, the unique combination of column, solvents, analytes, and flow rates used here responded to lowering the temperature for improved resolution. Lowering the column temperature coupled with decreasing the gradient flow rates proved vital to ensure that appropriate system pressures were maintained while facilitating the appropriate degree of separation of the isomers, ethylone and butylone.

	The synthetic cathinone compounds of interest were successfully analyzed in urine samples by this “dilute-and-shoot” LC–MS/MS method. This particular method is robust and useful because it reduces analysis time and does not incur additional costs for up-front sample preparation while still providing accurate results. As there were no interferences observed from the major drugs of abuse and therapeutic drugs, this method is an excellent tool for the clinical, medicinal, and toxicology communities, especially in high sample volume settings.

	Results achieved with this method are summarized in Table IV where positive data from over a year of testing are summarized. Mitragynine was the most prevalent component in the tested urine samples, while alpha-PVP and methcathinone were close seconds. Although tests for these analytes are positive for just a small fraction of all pain medication monitoring samples, these results are important to both physicians and patients in their ongoing treatments for chronic pain, substance abuse, and mental illness. For example, patient ages associated with at least one positive bath salt result ranged from 19 to 72 years old, with equal positivity rates across all age groups. Lastly, the overwhelming positivity of mitragynine in this group more than likely reflects the legality of this natural product in many states in the USA, while attempts to schedule this agent at the federal level have been delayed by consumer activist protests (13).

	Any method that tests for illicit synthetic substances must be updated frequently to keep up with the changes on “the street” that result in novel pharmaceutical substances being consumed with known and unknown effects. However, this method offers advantages in terms of cost and time and has been shown to be capable of detecting many patient positives when implemented for large numbers of samples.

	The authors would like to thank the National Science Foundation (NSF-GOALI Award 1611072) for financial support of this work, together with Wake Forest University and Ameritox.

	(4) A. Al-Motarreb, M. Al-Habori, and K.J. Broadley, 

	(6) J.F. Hyde, E. Browning, and R. Adams, 

	(7) C.D. Rosenbaum, S.P. Carreiro, and K.M. Babu, 

	(8) C.L. German, A.E. Fleckenstein, and G.R. Hanson, 

https://www.medicinenet.com/bath_salts_abuse_and_addiction/article.htm

	(11) W.C. Prozialeck, J.K. Jivan, and S.V. Andurkar,

	(12) M.A. Rech, E. Donahey, J.M. Cappiello Dziedzic, L. Oh, and E. Greenhalgh, 

are with the Chemistry Department at Wake Forest University in Winston-Salem, North Carolina and Ameritox, LLC, in Greensboro, North Carolina. 

Oneka T. Cummings, Erin C. Strickland, and Gregory L. McIntire 

is with the Chemistry Department at Wake Forest University. Direct correspondence to: 

A novel “dilute-and-shoot” LC–MS/MS method is described for the analysis of “bath salts” sold as “legal” highs, including mitragynine and nine synthetic cathinones, in urine.

Liquid Chromatography Tandem Mass Spectrometry Method for Novel Psychoactive Substances: Kratom and Synthetic Cathinones in Urine

Naltrexone (Depade, Re Via, and Trexan) is a potent narcotic antagonist structurally similar to oxymorphone and naloxone. It blocks the subjective effects of heroin and other opiates and is primarily used in the management of opioid dependence and alcohol dependence. While conjugated 6β-naltrexol is the major urinary metabolite in man, conjugated naltrexone and free 6β-naltrexol are also major urinary species. To assist with monitoring substance abuse patients who are prescribed naltrexone, a rapid, selective, and sensitive liquid chromatography–tandem mass spectrometry (LC–MS-MS) method was developed to analyze for both naltrexone and 6β-naltrexol post-enzymatic hydrolysis. The validation of this method and some representative patient data are discussed in this article.

	Naltrexone is a pure narcotic antagonist structurally similar to oxymorphone and naloxone. It blocks the subjective effects of heroin and other opiates and is primarily used in the management of opioid and alcohol dependence (1). The drug is supplied as 25-mg or 50-mg tablets (under the brands Depade, ReVia, and Trexan) (2) and as an extended-release injectable formulation (Vivitrol) (3), which is administered monthly as an intramuscular injection (1). Naltrexone undergoes extensive first-pass metabolism, and up to 70% of a dose can be recovered in urine within 24 h (1,4). Conjugated 6β-naltrexol (naltrexol) is the major urinary metabolite in humans, and conjugated naltrexone and free naltrexol are other known major urinary species (5). Naltrexol has significant pharmacological activity with a half-life that is greater than three times its parent, naltrexone (6).

	Many of the published analytical methods used to detect naltrexone and its metabolites were reported in plasma, which is a more complex matrix and usually requires more sample processing and cleanup when compared to urine (5,7–9). In a high-throughput laboratory environment, the emphasis is on developing methods that are not only rapid but also sensitive and selective. In view of that, this work demonstrates a method that requires dilution with internal standard, hydrolysis, and centrifugation. The subsequent analysis of naltrexone demonstrates toxicologically acceptable limits of quantitation (LOQs) and limits of detection (LODs) and robust instrument performance that minimizes instrument utilization time when analyzing naltrexone in urine.

	The resulting method reported herein has a 1.7-min run time and a 2.7-min cycle time using a liquid chromatography–tandem mass spectrometry (LC–MS-MS) system. Limits of quantitation for both naltrexone and 6β-naltrexol were determined to be 5 ng/mL. The development and validation of this method are discussed in further detail in this article.

	Reference standards for naltrexone, naltrexone-D3, 6β-naltrexol, oxymorphone glucuronide, and oxymorphone were purchased from Cerilliant. All solvents, including methanol (Optima grade), acetonitrile (Optima grade), isopropanol (Optima grade), 0.2 M phosphate buffer (pH 7.5), and formic acid (88%) were purchased from VWR. Drug free urine was obtained from Utak Laboratories. IMCSzyme, a recombinant β-glucuronidase, was purchased from IMCS.

	The analytical method was developed using an Agilent 6460 LC–MS-MS system. After testing several ultrahigh-pressure liquid chromatography (UHPLC) columns, a 50 mm x 4.6 mm, 100-Å, 2.6-µm 

 Phenomenex Phenyl Hexyl column coupled with a Phenomenex guard cartridge (Phenyl) was used to validate this method. The column temperature was maintained at 50 °C during analysis.

	Patient urine specimens (50 µL) were diluted 10x with a mastermix consisting of 1.6 µg/mL of naltrexone-D3 and 2000 units of a recombinant β-glucuronidase from IMCS (IMCSzyme) in 0.02 M phosphate buffer, pH 7.5. Samples were incubated in an oven for 60 min at 60 °C for hydrolysis. Earlier studies using oxymorphone glucuronide (naltrexone glucuronide is not commercially available) demonstrated the time course of hydrolysis for these analytes required >60 min for complete hydrolysis. It was determined that a hydrolysis time of 60 min provided stable and consistent results with a control. A correction factor of 1.14-adjusting for approximately 88% hydrolysis efficiency-was applied to results to mimic complete hydrolysis. Samples were removed from the oven and centrifuged before analysis by LC–MS-MS.

	Reference standards were diluted to appropriate calibrator level concentrations (5, 10, 50, 250, 1000, and 5000 ng/mL) in normal human urine and were then diluted an additional 10x as detailed above. The standards were meant to mimic the already 10x diluted patient samples. An oxymorphone glucuronide hydrolysis control was used to ensure the enzyme–mastermix was acceptable for every batch in addition to a positive and negative control.

	This method was validated for naltrexone and naltrexol as per an internal standard operating procedure (SOP) based upon College of American Pathologist requirements (10) and applied to authentic positive urine samples to evaluate efficacy for adherence drug testing. For a standard at any concentration level to be considered passing, during the validation of this method, quantitation must be within ±25% of the expected value and ion ratios and chromatographic requirements must pass for all analytes assessed. Quantitation values are based on a passing calibration curve (no points excluded and 

 value greater than or equal to 0.99), which must be freshly prepared and analyzed on the same day that the particular validation study was conducted. Furthermore, the coefficient of variation (CV) between sets of replicates must be ≤15%. Experiments performed during the method validation process are explained in greater detail below.

	The lower and upper concentration limits at which the method could accurately identify and quantify analytes were evaluated. To establish reliable LODs and LOQs, four out of five replicates must pass for the necessary criteria when compared to a passing calibration curve (detailed above). The LOD and LOQ both require the analyte signal to demonstrate reproducible ion ratio stability, while the LOQ requires the analyte signal to demonstrate reproducible quantitation.

is the reproducible regression or fit of the calibration curve compared to expected concentration values. Four of the five replicates of each linearity point-usually comprising the calibrators and a minimum of five additional points within the analytical range-must pass all validation criteria.

 is the highest quantitative level of analytes present that do not produce a concentration level above the lower limit of quantitation in a subsequent blank injection. Carryover is investigated to ensure false positives are alleviated in a production environment. This experiment requires four of five blank injections following each of the five replicates at the potential carryover limit concentration to quantitate lower than the established LOQ.

	The precision and accuracy test measures the capability of the method to yield reproducibly accurate results over a period of multiple days at concentrations spanning the concentration range of interest. A total of 10 replicates of three concentrations (omitting calibration points) within the analytical range of interest are chosen and run on three separate days. Out of these 10 injections, nine must pass within 25% of the target concentration.

	The ability of the method to be unaffected by the presence of other medications or compounds is the 

. Interference would be considered confirmed if a false signal met or exceeded LOQ requirements (that is, was higher than the LOQ and had a passing ion ratio) in greater than one of the five replicates that must be run for each individual test.

 are the suppression or enhancement of analyte signals of interest because of the presence of matrix-in this case urine. The effect of matrix on the method was evaluated by running the quality control (QC) concentration of 750 ng/mL both in matrix and in the LC starting conditions (98% 5 mM ammonium formate in 10% methanol: 2% methanol).

	This test is used to confirm whether a new method accurately quantitates actual patient samples when compared to the currently in-use and accepted method for these analytes (10). The previously validated method used a Waters Acquity TQD system and had been in regular daily use for more than a year. To make this comparison, all recent patient positive urine samples for the analytes in this method were prepared using the new procedure and compared to the previous method.

	Daily calibration curves (5, 10, 50, 250, 1000, and 5000 ng/mL) routinely demonstrated 100 ± 25% accuracy for each point and excellent goodness of fit (

 > 0.99). Calibration curves for oxymorphone only included 250, 1000, and 5000 ng/mL as the hydrolysis control was set to liberate 1577.8 ng/mL. Consistent peak shapes and ion ratios were achieved for naltrexone, naltrexol, and oxymorphone. A typical chromatogram is shown in Figure 1 for the 100 ng/mL calibrator demonstrating the separation achieved for this 1.7-min method; note that naltrexol, the latest eluted, is retained well before 1 min of the gradient. The longer run time (1.7 min) and cycle time (2.7 min) are necessary for column equilibration and needle washing–carryover reduction, respectively. Details about the LC gradient and MS ion fragmentation transitions used are described in Tables I and II, respectively. Compiled results for all conducted method validation experiments are shown in Table III.

					Figure 1: Example chromatogram of a 100-ng/mL sample injection (internal standard excluded).

	The LOD and LOQ for naltrexone and naltrexol were determined to be 5 ng/mL and 100 ng/mL for oxymorphone. No appreciable carryover was observed following the highest curve point of 5000 ng/mL (see Table III). The precision and accuracy was evaluated by assessing 10 replicates of standards at 75, 750, and 2500 ng/mL on three separate days-usually there is at least one skipped day between precision and accuracy experiments. The precision and accuracy results shown in Table III demonstrate the robustness of this method for all three analytes at the three concentration levels tested.

	All analytes experienced suppressed signal response when dissolved in normal human urine compared to a neat preparation in mobile phase initial conditions (98% 5 mM ammonium formate in 10% methanol: 2% methanol). Different methods can be used to calculate the effect of the matrix on the signal response. To calculate the percentages presented in Table III the relationship between the absolute responses is used since it is believed to present a more thorough picture than comparison of relative responses or calculated concentrations.

	The interference study was conducted and there were no identified interferences for any of the analytes in this method. Interference analytes included structurally similar compounds such as naloxone, 6α-naloxol, codeine, oxycodone, noroxycodone, hydromorphone, hydrocodone, and norhydrocodone as well as panels of common drugs of abuse such as opiates, opioids, and benzodiazepines.

	In addition to the validation, the patient samples tested showed 100% qualitative correlation between the original method and the newly validated method. Results from patient samples submitted who tested positive using this method and had known prescriptions for naltrexone (tablets or injectables) are listed in Table IV. Of the 7158 patient samples analyzed over a one-month period in a production setting using this method, 125 had known prescriptions and 90 tested positive while also passing all the specimen validity criteria (11,12). The summary of this sample set (positive patients with a known prescription) is presented in Table IV.

	Analysis of naltrexone dosages show a range of 15 ng/mL to 42,017 ng/mL with a mean of 4506 ng/mL for daily tablets (50 mg taken between once and thrice daily) and a narrower range of 18 ng/mL to 18,389 ng/mL for the injectable Vivitrol formulation with a mean of 1381 ng/mL. It is difficult to assess a mean for these data because of the large discrepancies between measured concentration values for patients with both dosage forms. The median is a more meaningful measure for these data and is higher for patients taking the injectable form of naltrexone when compared to those taking the daily tablets.

	This work demonstrates that the analysis of naltrexone and its major metabolite, 6β-naltrexol, can be adequately assessed in a rapid LC–MS-MS quantitative method following hydrolysis. Key features of the method include hydrolysis using a recombinant enzyme and a cycle time of 2.7 min while achieving the high level of selectivity and specificity required for naltrexone and naltrexol. The results from this method compare well with the same samples run with an LC–MS-MS method on a different system. This newly developed method uses different transitions and a comparable column and reduces the cycle time by approximately 1 min.

Disposition of Toxic Drugs and Chemicals in Man

, 10th Edition (Biomedical Publications, Seal Beach, California, 2014), pp. 1419–1420.

		Depade (package insert), Hazelwood, Missouri, Mallinckrodt Pharmaceuticals Inc. (2014).

		Vivitrol (package insert), Waltham, Massachusetts, Alkermes Inc. (2010).

		K. Verebay, M.J. Kogan, A. de Pace, and S.J. Mule, 

		W. Huang, D.E. Moody, R.L. Foltz, and S.L. Walsh, 

		E.J. Cone, C.W. Gorodetzky, and S.Y. Yeh, 

		A.F. Davidson, T.A. Emm, and H.J. Pieniaszek, 

		Substance Abuse and Mental Health Services Administration Department of Health and Human Services, “Mandatory Guidelines for Federal Workplace Drug Testing Programs,” Federal Register 

 are with Ameritox, Ltd., in Greensboro, North Carolina. Direct correspondence to: 

Naltrexone (Depade, Re Via, Trexan) is a potent narcotic antagonist structurally similar to oxymorphone and naloxone. It blocks the subjective effects of heroin and other opiates and is primarily used in the management of opioid dependence and alcohol dependence. While conjugated 6β-naltrexol is the major urinary metabolite in man, conjugated naltrexone and free 6β-naltrexol are also major urinary species. To assist with monitoring substance abuse patients who are prescribed naltrexone, a rapid, selective, and sensitive LC–MS-MS method was developed to analyze for both naltrexone and 6β-naltrexol post-enzymatic hydrolysis. The validation of this method and some representative patient data are discussed in this report.

A Rapid, Selective, and Sensitive Method for the Analysis of Naltrexone and 6β-Naltrexol in Urine

A new method was developed and validated using automated on-line solid-phase extraction (SPE) with tandem mass spectrometry (MS). Urine samples were enzyme-hydrolyzed and diluted before detection. The validated method was applied to positive authentic urine samples to evaluate concordance with high performance liquid chromatography (HPLC)–MS-MS results.

	Benzodiazepines were first synthesized in the 1950s and successfully marketed in 1960 (1,2), leading to a rapid rise in prescriptions of the compounds for the treatment of anxiety, insomnia, and seizures, among other conditions (3). They were seen as safer than, but just as effective as, barbiturates and other older drugs used as sedatives and hypnotics. However, by the 1980s it became clear that benzodiazepines were also saddled with the same liabilities for abuse and dependence as the older medications they replaced (1,4). Despite this risk, as well as the availability of other safer medications with anxiolytic properties, the number of prescriptions written for benzodiazepines has not significantly dropped (1,4). In 2012, three benzodiazepines (alprazolam, lorazepam, and clonazepam) were among the top 100 drugs prescribed in the United States, while two others (diazepam and temazepam) made it onto the top 200 list (5). Because benzodiazepines are so widely available, both by prescription and illicitly, and have the potential to be abused, there is a need for patients to be monitored for therapeutic drug compliance or diversion.

	The class of drugs termed benzodiazepines share the core structural components of a benzene ring fused with a diazepine ring (3). The unique side chains found in distinct benzodiazepine compounds give each drug its pharmacological properties, modulating its binding to the GABA

 receptor (2,4). Benzodiazepines derive their pharmacological actions from their interactions with the GABA

 receptor, increasing the efficiency of GABA, thereby reducing communication between neurons and yielding a calming effect on many brain functions (6). In addition to these analgesic effects, benzodiazepines are often misused by poly-drug abusers to enhance and prolong the "high" generated by other drugs, and ease the withdrawal effects when other drugs are scarce (2,6,7). These unintended uses have resulted in widespread abuse of benzodiazepines and have led to their regulation by the U.S. Drug Enforcement Agency (DEA) as Schedule IV drugs (2), thereby supporting the need for urine drug testing for compliance and abuse of these compounds.

	There are more than 50 benzodiazepine compounds marketed worldwide (4), but the focus of this work is on seven of the more commonly prescribed compounds or their major metabolites: 7-aminoclonazepam, α-hydroxyalprazolam, alprazolam, lorazepam, nordiazepam, oxazepam, and temazepam (structures depicted in Figure 1). In this study, quantitative results for the analysis of these seven benzodiazepines and metabolites using a solid-phase extraction tandem mass spectrometry (SPE–MS-MS) method are compared with earlier results obtained using a conventional high performance liquid chromatography (HPLC)–MS-MS method. Because these drugs undergo extensive metabolism, samples are hydrolyzed to return compounds to their nonglucuronidated state before confirmation analysis. The previous HPLC–MS-MS method used to analyze these drugs required a 6.4-min cycle time with a 96-s data window, such that quadruple multiplexing yielded a net time per sample of 96 s. The analytical columns for this method were replaced every 1000 injections of running in a production environment, while the guard columns were replaced every 1500 injections. The SPE-MS-MS method reported herein has been tested extensively, affording a comparison with the previous method for analysis time, cartridge lifetimes, data values, and overall cost per sample.


	Figure 1: Chemical structures for the seven benzodiazepines analyzed. Row 1 (top), left to right: alprazolam, α-hydroxyalprazolam, lorazepam. Row 2 (middle), left to right: nordiazepam, oxazepam, temazepam. Row 3 (bottom), left to right: 7-aminoclonazepam, generic benzodiazepine backbone.

	The compounds of interest and the internal standard (alprazolam, α-hydroxyalprazolam, 7-aminoclonazepam, lorazepam, nordiazepam, oxazepam, temazepam, and nordiazepam-D5), along with the hydrolysis control (oxazepam glucuronide), were purchased from Cerilliant. Acetone (Optima grade), isopropanol (Optima grade), glacial acetic acid (certified ACS grade), formic acid (88%, laboratory grade), ammonium acetate (HPLC grade), ammonium formate (certified grade), and ammonium hydroxide (certified ACS Plus grade) were obtained from Fisher Scientific. Acetonitrile (OmniSolv gradient grade) and methanol (OmniSolv gradient grade) were acquired from EMD Chemicals. Potassium hydroxide (50% w/v, analytical reagent grade) was purchased from Ricca Chemical Company. Normal human urine was acquired from UTAK Laboratories. β-Glucuronidase (100,000 units/mL, from abalone) was purchased from Campbell Science. Water used for all reagents, mobile phases, and solvents was obtained from a Barnstead Nanopure analytical laboratory water system (18.2 MΩ·cm). The internal standard compound was diluted to a concentration of 1 μg/mL with water, while all other reference standards were diluted to concentrations of 100 and 10 μg/mL with methanol. These stock solutions were then used to prepare standard curve solutions and controls by diluting known amounts with normal human urine.

	A stock enzyme buffer was prepared by combining 800 mL of water and 111 mL of glacial acetic acid and filling a 1-L flask to volume with the potassium hydroxide solution. This buffer was stored in the refrigerator and was stable for one month. The stock buffer was further diluted 1:10 in water for use in samples. The purchased β-glucuronidase was also diluted 1:5 in water for use in samples. To 0.5-mL aliquots of patient urine samples, 100 ng of internal standard was added, followed by 100 μL of diluted enzyme buffer (pH 4.7), and 20 μL of diluted enzyme. Samples were mixed by vortex and hydrolyzed at 60 °C for 30 min, then centrifuged for 10 min at 2500 rpm. For HPLC–MS-MS analysis, 96-well sample plates were placed directly in the autosampler. For on-line SPE-MS-MS analysis, samples were further diluted 20-fold in water before being loaded into the sample queue.


	Table I: Gradient conditions and flow rates used for benzodiazepine assay on the Thermo Quantum Access Max MS system

	Liquid chromatography was conducted on an Agilent 1200 series HPLC system equipped with a 50 mm × 3 mm, 5-μm Thermo Hypersil GOLD aQ column fitted with a 50 mm × 0.5 mm Thermo Cyclone-P TurboFlow guard column. The injection volume was 10 μL with a total run time of 6.4 min and an injection-to-injection MS detection cycle time of 96 s. The flow rates and gradient conditions are detailed in Table I. Tandem MS was conducted using a Thermo TSQ Quantum Access Max MS system with electrospray ionization performed in the positive mode. The source temperature was 270 °C and the desolvation temperature was 375 °C. The compounds were analyzed in selected reaction monitoring (SRM) mode with argon as the collision gas. One quantitative transition and one qualitative transition were monitored for each analyte and internal standard. A summary of the MS parameters for each compound of interest are detailed in Tables II and III.


	Table II: Detailed summary of transition-specific settings on the Thermo Quantum Access Max MS system. Quantification transitions are shown in bold type.

	On-line SPE was conducted on an Agilent RapidFire 300 on-line SPE system. The SPE system uses a series of valves, high flow rate solvents, and proprietary cartridges to load a sample, rinse away the matrix, and then bulk elute or inject into a mass spectrometer in approximately 20 s or less. This method performs no chromatographic separation and therefore cannot resolve isobaric species. In this experiment, the injection volume was 10 μL onto an A2 C4 packed cartridge, with a total cycle time of 13 s. The on-line SPE cleanup of each sample was achieved with a sample load in water (mobile-phase A) at a flow rate of 1.5 mL/min and a wash of 10% acetonitrile in water (mobile-phase B) at a flow rate of 1.25 mL/min. The sample was eluted from the cartridge using 25:75 methanol–isopropanol (mobile-phase C) at a flow rate of 0.75 mL/min. Tandem MS was performed using either an Agilent 6460 triple-quadrupole system or an Agilent 6550 quadrupole time-of-flight (QTOF) system. Electrospray ionization was conducted in the positive mode. The gas temperature was 200 °C and the sheath gas temperature was 400 °C. The gas flow and sheath gas flow were set to 5 L/min and 11 L/min, respectively. The analytes of interest were detected using multiple reaction monitoring (MRM) mode with nitrogen as the collision gas. Two transitions, one quantitative and one qualitative, were monitored for each compound. A summary of the MS settings for each analyte are shown in Tables II and III. Quantitation by the SPE–MS-MS method was completed by correlating the ratio of the area counts of the analyte of interest to the area counts of the nordiazepam internal standard to the concentration of standards at varying concentrations using a quadratic curve and 1/

 data weighting for each compound. The HPLC–MS-MS method was quantitated in a similar way, though there were matching deuterated internal standards for each analyte.


	Table III: Detailed summary of transition-specific settings on the Agilent 6460 MS system. Quantification transitions are shown in bold type.

	All analytes displayed unique mass transitions, allowing the method to be easily converted to the automated SPE format, where no chromatographic separation occurs. A summary of the validation results from both the HPLC and the automated SPE methods are presented in Tables IV and V. To transition to the automated SPE method, a full validation procedure recommended by the College of American Pathologists (CAP) was performed, which included determination of the limit of detection (LOD), lower limit of quantitation (LLOQ), upper limit of quantitation (ULOQ), and an examination of carryover, matrix effects, interference, accuracy, intra- and inter-run precision, and patient comparison to the production method.


	Table IV: Comparison of concentration limits from HPLC and SPE methods

	The LLOQ was identified by the analysis of calibration curves in five replicates. It was defined as the lowest non-zero calibrator with acceptable ion ratio and suitable precision (<15% CV) and accuracy (±25%) included in the calibration curve that gave an acceptable coefficient of determination (

 ≥ 0.99). The RF LLOQ was 50 ng/mL, and the LOD was 25 ng/mL. The ULOQ was set to 10,000 ng/mL based on the therapeutic range.


	Table V: Comparison of precision and accuracy for HPLC and SPE methods

	The absence of carryover was demonstrated by injecting a negative urine sample after a sample spiked with 10,000 ng/mL of each analyte. Carryover was considered insignificant if none of the analytes was detected at or above the LLOQ. This method was free from carryover at 10,000 ng/mL.

	The matrix effect on each analyte was assessed by comparing the relative response ratio of each compound prepared in neat urine as well as in water at a mid-level concentration in the calibration curve. Five replicates of each analyte in each preparation solution were run. The average relative response ratio was determined for each set of samples and the matrix effect was calculated as follows: % matrix effect = ([average matrix response – average water response]/average water response) × 100. A positive result indicated ion enhancement, and a negative result showed ion suppression. Results are summarized in Table VI. Both alprazolam and temazepam demonstrated large matrix effects, which did not appear to affect the limitations of the method.


	Table VI: Summary of matrix effect seen during validation of both HPLC and SPE methods

	Interference from endogenous matrix components, the internal standard, and other possible drug compounds was determined through a variety of studies. To rule out interference on the internal standard, five replicates of each individual analyte were prepared at the ULOQ, substituting water for the internal standard. No interference was detected at the internal standard retention time. Matrix interference was excluded by analyzing five replicates of neat urine with no internal standard. Potential interference from other drug compounds was assessed by examining the responses of a variety of related pain medications. None of the analytes of interest were detected above the LLOQ in any of these tests. Because the automated SPE method also eliminates chromatography, an additional interference study was performed to determine if any of the possible interfering drug compounds contributed to significant ion suppression of the analytes of interest. Each benzodiazepine analyte in the new method was made up individually at a mid-level concentration (1000 ng/mL) and spiked into the various interference drug mixes containing compounds at extremely high concentrations. None of the potential interfering compounds demonstrated a suppressive effect on any of the benzodiazepines.

	Precision (intra- and inter-run) and accuracy were determined by examining 10 replicates of each of the three concentrations across the calibration range over the course of three days and calculating the percent of target for accuracy and percent CV for precision. Acceptable accuracy (<14%) and intra- and inter-run precision (<11% CV) were found at each concentration for each analyte of interest.

	A comparison of patient samples was performed to assess how the results correlated between the automated SPE method and the production HPLC method. First, 60 patient samples that were positive for a variety of the benzodiazepines of interest were analyzed by LC–MS-MS, then diluted 20-fold and analyzed by SPE–MS-MS. The data between the two methods correlated well (percent deviation did not exceed 30%), but peak interferences were observed for nordiazepam and temazepam in patient samples. While the nature of these interferences is under investigation, it was found that the use of an exact mass, high-resolution QTOF MS system eliminated the interferences from these samples. Figure 2 (top) shows the response for a sample with the interferent using a triple-quadrupole detector. This sample was not positive for nordiazepam by the current HPLC–MS-MS method, yet a positive peak is clearly visible using the automated SPE-MS-MS method. Figure 2 (bottom) shows the same sample with detection via the QTOF system. Note the total absence of signal between 8 s and 12 s of the elution time exactly where the "false" peak is eluted. Thus, the exact mass resolution of the QTOF system results in the absence of signal for this transition (271.1→139.9) and shows the absence of the false positive peak seen with the HPLC–MS-MS method. Migration of this method from triple-quadrupole MS to exact mass MS maintained the validation criteria observed in the validation of this method.


	Figure 2: (a) A chromatogram depicts the nordiazepam interference (false positive) seen when using a triple-quadrupole MS system with the on-line SPE system for benzodiazepine detection in patient samples. (b) The same sample injected into a QTOF shows the lack of a peak (that is, noise) for nordiazepam, demonstrating exact mass resolution of the interferent.

	Aside from opiate testing, benzodiazepine panels account for the predominant bulk of all samples tested in the clinical pain management laboratory, especially in a pain medication monitoring facility. Because of the volume of the panel, a faster, more efficient, and ideally cheaper method was desired. The existing HPLC–MS-MS method was multiplexed to allow for a 96-s sample injection to sample injection cycle time. On-line sample cleanup makes injection of enzyme-containing samples possible without preceding enzyme removal. Although this method appears fairly rapid, consumable costs (columns, internal standards, and solvents) were mounting, and other options to increase efficiency were explored.

	An SPE-based front-end system was selected because of its speed in sample introduction. With a cycle time of 13 s from sample injection to sample injection, the method developed on the SPE system allowed for a sevenfold increase in throughput while using a smaller number of MS instruments to handle the volume. The cost savings were also significant because of SPE cartridge lifetime and lower overall operating cost, leading to at least a 10-fold savings over the traditional HPLC–MS-MS analysis. The cartridge lifetime is notable in spite of simply diluting the enzyme-containing samples before injection. It could reasonably be expected that enzymes or proteins may clog or otherwise adversely impact the efficiency of the cartridge much like an analytical column. However, the design of the SPE cartridge, with its relatively large particle size, enables this type of dilute-and-shoot methodology.

	Method development and validation were performed as described above using seven compounds of interest (7-aminoclonazepam, α-hydroxyalprazolam, alprazolam, lorazepam, nordiazepam, oxazepam, and temazepam) and one internal standard (nordiazepam-D5), with two transitions monitored for each compound. The use of a single internal standard resulted from limitations associated with the lack of chromatography and the number of total transitions the mass spectrometer can monitor at a single time while retaining enough points across each peak to ensure peak fidelity. For similar reasons, dwell times on the triple-quadrupole and QTOF-MS systems had to be carefully monitored. Data from the method validation demonstrated comparable responses and quantitation when compared to the HPLC–MS-MS method where an internal standard was used for each compound, leading to the conclusion that only one internal standard was sufficient for accurate results. The use of a single internal standard contributed to the decrease in cost observed in the transition to the new SPE–MS-MS method.

	Interestingly, all analytes demonstrated positive matrix effects, with those for alprazolam and temazepam as the most significant. It is possible that the urine matrix assists the adsorption onto the SPE cartridge and thus results in increased responses relative to those seen in water (neat preparation) alone. Salt content and pH would be expected to have an impact on partitioning onto the SPE and inasmuch as these samples are all buffered for enzyme efficiency, it would seem to point to increased salt content as the driving force in this enhanced response.

	Comparison of patient data obtained using the SPE–MS-MS method to data obtained using the conventional HPLC–MS-MS method demonstrates good correlation, despite a cycle time that is seven times shorter. However, it is worth noting that the elimination of chromatography with the SPE–MS-MS method led to the appearance of an interferent in patient samples for the nordiazepam and temazepam transitions. Patients who were observed positive for those two compounds by HPLC–MS-MS were also observed positive by SPE–MS-MS; however, patients that had not previously demonstrated a response for either compound by HPLC–MS-MS did show one when analyzed by SPE–MS-MS. Although these "false positives" were not in fact positives by quality control criteria (ion ratios), the appearance of the interferent casts doubt on the confirmation of those two compounds and implies that the SPE–MS-MS method cannot be used in a production environment with a triple-quadrupole detector.

	Interestingly, use of a QTOF detector resolves these interferences and offers a way forward for this method. Even though initial parent ion selection on a QTOF system has the same mass resolution limitation as a triple-quadrupole system (approximately 1 Da), the use of an exact mass TOF analyzer on the back end allows for more definitive detection. This effectively removes the false fragment selection, thus eliminating the interference. As illustrated in Figure 2, the exact mass detector offered clean peaks without the interferent observed with the triple-quadrupole instrument. Repeat studies with calibrators demonstrated equivalent LLOQ and LOD values, as well as dynamic range and precision.

	The newly developed SPE–MS-MS method for the analysis of select benzodiazepine compounds returns results comparable to those found by the traditional HPLC–MS-MS method, but with a much faster analysis time and at a much lower cost per sample. Despite the absence of chromatography in the SPE–MS-MS method, the limits of analysis for the benzodiazepine panel were retained or improved on when compared to the HPLC method; however, though patient samples correlated well, peak interferences were observed for two of the compounds, precluding the SPE–triple-quadrupole MS method from entering the production workflow. The use of the QTOF detector with the SPE system showed the interferences can be resolved, and this SPE–MS-MS method may prove to be a good alternative to HPLC–MS-MS for the analysis of select benzodiazepine compounds, thereby greatly increasing efficiency while reducing cost.

	All funding was provided by Ameritox, Ltd.

	The authors wish to thank Dr. Michelle Romm of Agilent Technologies, as well as Dr. Maxine Jonas, for their expertise and assistance with method development on the RapidFire system.

Jennifer C. Hitchcock, Jeffrey R. Enders, Ayodele A. Morris, and Gregory L. McIntire

Disposition of Toxic Drugs and Chemicals in Man, 7th ed.

 (Biomedical Publications, Foster City, California, 2004), pp. 45, 376, 471, 940, 1251, and 1621.

www.pharmacytimes.com/publications/issue/2013/July2013/Top-200-Drugs-of-2012

Benzodiazepine Abuse. Drugs and Dependence,

 (Harwood Academic Publishers, New York, New York, 2002), pp. 197–212.

Rapid Analysis of Selected Benzodiazepines by Automated SPE–MS-MS

Enzyme immunoassay (EIA) is a conventional drug screening technique, but it can be limited by cross-reactivity that can lead to high false positive rates. To improve the selectivity and efficiency of a urine screen for drugs of abuse, mass spectrometry (MS) was investigated as an alternative. An existing multianalyte MS method for amphetamine, benzoylecgonine, 3,4-methylenedioxyamphetamine (MDA), methylenedioxyethylamphetamine (MDEA), 3,4-methylenedioxymethamphetamine (MDMA), methamphetamine, and phencyclidine (PCP) was modified to optimize for speed and to be a competitive screen. 

	Recently, many new direct-analysis mass spectrometry (MS) techniques have been developed primarily for screening purposes, such as high-throughput systems (1–10), laser diode thermal desorption (LDTD) (11–19), direct analysis in real time (DART, JEOL) (20–32), and desorption electrospray ionization (DESI) (20,33–44). These techniques have proven to be time efficient and useful in drug sample analysis (2,6,10,13,15,16,20–23,26,28,32,34,35,37,41,42,44). However, all of these techniques require special and expensive equipment in addition to a typical MS system that most often is purchased in tandem with a liquid chromatography (LC) system. Another downside is that these techniques can require a specific sample "holder." For example, one high-throughput approach currently only uses 96- or 384-well plates, and the LDTD system has its own special plate as well, which may not be ideal for small-volume assays because the need for automated liquid handling becomes a necessary additional expense. This additional equipment cost for smaller laboratories may not always be feasible. Therefore, a less-expensive and time-efficient LC–MS-MS alternative screening technique was investigated.

	Using conventional LC system components and short columns or commercially available guard columns, a rapid on-line sample preparation and injection (ROSPI) method was developed. ROSPI was developed using the following guidelines:

		The method cycle time had to be less than 2 min and ideally ~1 min.

		Short columns were used to help reach the method cycle time and maintain system pressures.

		Complete separation of the peaks or interferences was not necessary.

		Only one transition was used per analyte or internal standard (IS) to ensure a sufficient number of points across the peak.

		The number of IS compounds was minimized.

		Two-point calibration near the cutoff was used for semiquantitation to determine the positivity of samples.

	A current LC–MS-MS confirmation method that included commonly abused drugs was used as a proof of concept for the ROSPI method. This method included seven analytes, shown in Figure 1: amphetamine, benzoylecgonine (a cocaine metabolite), 3,4-methylenedioxyamphetamine (MDA), methylenedioxyethylamphetamine (MDEA), 3,4-methylenedioxymethamphetamine (MDMA), methamphetamine, phencyclidine (PCP). The method also included five IS compounds: amphetamine-D5, benzoylecgonine-D3, MDMA-D5, methamphetamine-D5, and PCP-D5. Typically, the compounds in this method would be screened by four different enzyme immunoassay (EIA) tests: amphetamines (amphetamine and methamphetamine), MDMA (MDA, MDEA, and MDMA), benzoylecgonine, and PCP. However, with MS all of these compounds can be screened with a single injection in less than 2 min. Furthermore, implementing with existing equipment and less expensive guard columns affords rapid screening and improved cost efficiency. Using the aforementioned goals, a ROSPI method was developed for these drugs and compared to EIA for selectivity and sensitivity using authentic patient samples.


	Figure 1: Structures of drugs analyzed.

	Amphetamine, amphetamine-D5, benzoylecgonine, benzoylecgonine-D3, MDA, MDEA, MDMA, MDMA-D5, methamphetamine, methamphetamine-D5, PCP, and PCP-D5 controls were all purchased from Cerilliant. LC solvents, acetonitrile, and formic acid were purchased from Fisher Scientific and methanol was purchased from EMD Millipore. Drug-free, normal human urine was purchased from UTAK Laboratories. All water was acquired from an in-house Barnstead Nanopure purification unit from Thermo Scientific. Polypropylene 700-μL vials and snap caps with septa were purchased from VWR International.

	Samples were run on an Agilent 6460 QQQ mass spectrometer with an Agilent 1200 Series Binary Pump, a column compartment with a CTCPal autosampler, and an Agilent Jet Stream electrospray ionization (ESI) source. Columns were purchased from Phenomenex and included a 30 mm × 3.00 mm, 2.6-μm 

, 100-Å Kinetex C18 column for the ROSPI method and a 50 mm × 3.00 mm, 2.6-μm 

, 100-Å Kinetex C18 column for the confirmation method. Both columns were used in tandem with a Phenomenex SecurityGuard Ultra C18 guard column and holder. Data were analyzed on Agilent MassHunter Quantitative Analysis software.

	Aliquots of drug-free urine were fortified with amphetamine, benzoylecgonine, MDA, MDEA, MDMA, methamphetamine, and PCP at various concentrations for the calibration curves and a quality control (QC). For the confirmation method, all compounds except PCP were prepared at concentrations of 25, 50, 100, 150, 500, 1000, and 2500 ng/mL. PCP was prepared at additional concentrations of 4 and 10 ng/mL and excluded from the 2500-ng/mL point. For the screen method, all compounds except PCP were prepared at concentrations of 100, 150, and 300 ng/mL; PCP was prepared at concentrations of 25, 50, and 150 ng/mL. All samples were diluted 20× with 500-ng/mL amphetamine-D5, benzoylecgonine-D3, MDMA-D5, methamphetamine-D5, and PCP-D5 in water and centrifuged before a 20-μL injection onto the column. The gradient for the confirmation method was 1.87 min with a 3.12 min cycle time. The gradient was improved to 0.8 min with a ~1.7 min cycle time for the screening method. The flow rate was the same for both methods at 0.6 mL/min and the solvents used were 0.1% formic acid in water for solvent A and 50:50 acetonitrile–methanol + 0.1% formic acid for solvent B. Additionally, a wash 1 solvent of 50:50 methanol–water and a wash 2 solvent of 100% methanol were used on the CTCPal system. The additional cycle time for the methods is related to the washing and injection cycle of the CTCPal system.

	In the confirmation method, two multiple reaction monitoring (MRM) transitions were used for every analyte, including the IS compounds. In the screening method, the quantifier ion transition, or the most abundant transition, was used for every analyte, including the IS compound. For the simplicity of the screening method, only benzoyl-ecgonine-D3 was used as the IS compound in the processing method, even though all IS compounds were present in the sample. This was done to demonstrate the ability to decrease cost for the screen by using a single IS compound instead of five. Data were acquired in positive ESI mode and all MS conditions remained the same for both methods.

	Because the confirmation method was previously validated, validation experiments were performed only for this screening method and included limits, precision and accuracy, and patient comparison. For the limits experiment, five replicates of each point "lower" limit of quantitation (LOQ), screen cutoff, QC, and upper limit of linearity (ULOL/carryover) were averaged and the %CV was calculated for each analyte. Precision and accuracy were assessed across three days using three replicates of only the screen cutoff and the QC for each analyte. Finally, a comparison of 344 authentic patient positive samples was correlated to the EIA results and confirmation results to determine the false positive and false negative rates as applicable.

	All analytes seemed to work well with a linear fit through the origin using the single-point calibrator to provide a two-point calibration and benzoylecgonine-D3 as the sole internal standard. Data for the limits of validation can be seen in Table I. The LOQ was 25 ng/mL for every analyte except PCP, which was 4 ng/mL. The MS screen cutoff was set at 100 ng/mL for all analytes except PCP, which was set at 25 ng/mL. All analytes had a concentration of 150 ng/mL in the QC, except PCP, which was at 50 ng/mL. Carryover/ULOL was established at 10,000 ng/mL for amphetamine, benzoylecgonine, MDMA, and methamphetamine; 2500 ng/mL for MDA and MDEA; and 1000 ng/mL for PCP. Because the screen method was concerned more with accuracy in quantitation at the screen cutoff and used a lower calibration point, the carryover/ULOL was just assessed for precision (coefficient of variation [%CV]) and carryover criteria. If the accuracy failed outside of the tolerance, it was deemed acceptable as long as it would not change the positivity of the sample. However, it appeared that even though the single-point calibrator was at a significantly lower concentration than the ULOL, 2–3× the cutoff, the linear range held up to be the same as the original method, >1000 ng/mL depending on the analyte. Requirements for passing included that accuracy of the quantitation be within ±25% and %CV be within ±15%. The passing of the limits of validation indicated that using a two-point calibration with a single IS compound could be successfully used for semiquantitation at least for the analytes in this method.