Journal of Chromatographic Science, 2016, Vol. 54, No. 4, 507–515 doi: 10.1093/chromsci/bmv172 Advance Access Publication Date: 10 December 2015 Article
Article
Development and Validation of a Sensitive LC–MS-MS Method for the Determination of Adefovir in Human Serum and Urine: Application to a Clinical Pharmacokinetic Study Ye Zhang1,†, Lu Shen1,†, Ying Zhan2, Qing-Qing Xiao3, and Jin Yang1,* 1 Center of Drug Metabolism and Pharmacokinetics, China Pharmaceutical University, 24 Tong Jia Xiang, Nanjing, China, 2Department of Pharmacy, Zhongda Hospital of Southeast University, No. 87 Ding Jia Qiao, Nanjing 210009, China, and 3Department of Clinical Pharmacy, China Pharmaceutical University, No. 24 Tong Jia Xiang, Nanjing 210009, China
*Author to whom correspondence should be addressed. Email: [email protected] †
These authors contributed equally to the paper.
Received 6 February 2015; Revised 24 August 2015
Abstract A rapid and sensitive liquid chromatography–tandem mass spectrometry method was developed and validated for the quantification of adefovir (PMEA,9-(2-phosphonylmethoxyethyl) adenine) concentration in human serum and urine. The analysis was performed on a negative ionization electrospray mass spectrometer via multiple reaction monitoring. The monitored transitions were set at m/z 272.0 → 134.0 and m/z 276.0 → 149.8 for PMEA and internal standard, respectively. After protein precipitation, samples were separated by high-performance liquid chromatography on a reversedphase Dikma Diamonsil C18 (250 × 4.6 mm; 5 µm) column with a mobile phase of 0.1 mM ammonium formate buffer–methanol. The calibration curves were linear over the serum concentration range 0.5–1,000 ng/mL and urine concentration range 2.0–1,000 ng/mL. The intra- and interday precision values of PMEA in both serum and urine were lower than 18.16% for low quality control and 13.70% for medium and high quality control. The accuracy, recovery, matrix factor and stability were also within the acceptable limits. The developed method was successfully applied to the pharmacokinetic study of following oral administration of single dose of pradefovir mesylate (10, 30, 60, 90 and 120 mg) and adefovir dipivoxil (10 mg) to healthy Chinese volunteers.
Introduction Pradefovir mesylate is designed as an oral target prodrug of 9-(2-phosphonylmethoxyethyl) adenine nucleotide analog adefovir (or adefovir, PMEA), containing a cyclic phosphonate diester linkage (see Figure 1). Pradefovir employed the HepDirect™ technology and could be activated to PMEA by CYP3A4, which exists primarily in liver cells (1–3). However, another prodrug of adefovir, the broad-spectrum antiviral anti-Hepatitis B virus (HBV) agent, adefovir dipivoxil, converted to adefovir by widespread cellular esterases (see Figure 1). It has been manifested that pradefovir has a better liver targeting property than adefovir
dipivoxil in rats and cynomolgus monkeys (4). Pradefovir mesylate yielded 15 times higher concentrations of radioactivity in the liver than adefovir dipivoxil, but only one-third of the concentrations in the kidney after oral administration to rats. In addition, after oral dosing the same radiolabeled agents in cynomolgus monkeys, pradefovir mesylate yielded 60 times higher levels of total radioactivity in the liver, but only two-thirds of total radioactivity levels in the kidney than adefovir dipivoxil (4). Since the liver is the target organ for HBV infection and the kidney is the site of toxicity, pradefovir is expected to have better efficacy and lower toxicity compared with adefovir dipivoxil.
© The Author 2015. Published by Oxford University Press. All rights reserved. For Permissions, please email: [email protected]
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Figure 1. Chemical structures of pradefovir mesylate (A), adefovir dipivoxil (B) and PMEA (C).
Pradefovir and PMEA are the main circulatory substrate after oral dosing of pradefovir mesylate. So far, only one study reported the method of simultaneous determination of pradefovir and PMEA after solid-phase extraction (SPE) from human serum and urine by liquid chromatography–tandem mass spectrometry (LC–MS-MS) under positive and negative ions with switch scanning technology (5). But the authors have not published their method validation data. We tried to develop a method for the determination of pradefovir and PMEA simultaneously in positive ionization mode. However, PMEA produced a much lower intensity in positive ionization mode than pradefovir that could not reach the enough lower limit of quantification (LLOQ) for clinical pharmacokinetic trial, while in negative ionization mode, PMEA produced a higher intensity. Hence, PMEA was analyted in negative ionization mode and was detected separately with pradefovir. The determination of PMEA using high-performance liquid chromatography (HPLC) with ultraviolet detection and HPLC with fluorescence detection has been proved to be too complicated and/or not sensitive enough for the clinical pharmacokinetic study (6–9). So far, several methods have been reported for the assay of PMEA using LC–MS-MS (10–15). Liu et al. (10) reported a LC–MS-MS method that gave a satisfactory sensitivity with LLOQ of 0.10 ng/mL, whereas the serum samples were treated by a complex SPE procedure. Xie et al. (14) developed a LC–MS-MS method with the long analysis time of 14 min. Furthermore, peaks of analytes in the above-mentioned methods were not perfect enough due to their relatively broad shapes. The purpose of our research was to develop and validate a quantitative LC–MS-MS method for the analysis of PMEA with simple pretreatment process and perfect peak shape in human serum and urine for clinical pharmacokinetic studies of pradefovir mesylate.
Experimental Chemicals and reagents Pradefovir mesylate ( purity: 99.54%) and PMEA ( purity: 94.2%) were obtained from Xi’an New Drug Research Co., Ltd (Xi’an, China). Entecavir ( purity: 99.9%) as internal standard (IS) was obtained from Wuhan Belka Biological Pharmaceutical Co., Ltd (Wuhan, China). HPLC-grade methanol and acetonitrile were purchased from Merck (Darmstadt, Germany). Formic acid was purchased from ROE (Newark, New Castle, USA). Ultrapure water was obtained from a UPH Ultrapure Water System (UPH-II-5T, Chengdu, China). Ammonium formate, trichloroacetic acid and other reagents were of analytical grade.
Instruments Chromatographic analysis was performed on a Shimadzu LC-20A series chromatographic system (Shimadzu Corporation UFLC XR, Kyoto, Japan) with two LC-20AD binary pumps, a DGU-20A3 degasser, a SIL20AC autosampler and a CTO-20A column oven. The separation was performed on a Dikma Diamonsil C18 (250 × 4.6 mm; 5 µm) column, at a temperature of 30°C. The mass spectrometer was operated on an API 4000 triple-quadrupole mass spectrometer equipped with an electrospray ionization (ESI) source (AB Sciex, Foster City, CA, USA).
LC–MS-MS analytical conditions The mobile phase was composed of 0.1 mM ammonium formate buffer in water (pH 2.5 adjusted with formic acid, mobile phase A) and methanol (mobile phase B) at a flow rate of 1.0 mL/min. The gradient elution program was as follows: 0–2.0 min, 5% B, 2.0–2.5 min, 5–30% B, 2.5–6.0 min, 30% B, 6.0–6.1 min, 30–5% B, 6.1–8.0 min, 5% B. The eluate from the column was diverted into the MS from 5.0 to 7.5 min. The total analytical run time per sample, including equilibration time, was 8.0 min. The autosampler was conditioned at 4°C and the injection volume was 20 µL for analysis. The mass spectrometer was operated in negative ESI mode with multiple reaction monitoring (MRM), monitoring the transition m/z 272.0 to m/z 134.0 for PMEA and m/z 276.0 to m/z 149.8 for IS. The MS conditions were as follows: curtain gas, 20 psi; source temperature, 550°C; dissociation gas, 6 psi; ion source gas 1, 60 psi; ion source gas 2, 50 psi; ion spray voltage, −4,200 V; declustering potential, −73 V; collision energy, −26 V; entrance potential, −10 V and collision cell exit potential, −10 V. All data were acquired and processed using Analyst 1.5.2 version software (AB Sciex).
Preparation of the stock and working solutions Primary stock solutions of PMEA were prepared in purified water at a concentration of 1.0 mg/mL. A series of working standard solutions (5, 10, 20, 50, 100, 200, 500, 1,000, 2,000, 5,000 and 10,000 ng/mL) and quality control (QC) samples (10, 1,000 and 8,000 ng/mL for serum samples, and 50, 500 and 8,000 ng/mL for urine samples) were obtained by further diluting the stock solutions in methanol/ water (1 : 1, v/v). Stock solutions of IS at 1.0 mg/mL were prepared in methanol/water (1 : 1, v/v). The IS working solution (10 µg/mL) was obtained by diluting the stock solution in methanol/water (1 : 1, v/v). All the solutions were stored at 4°C and were brought to room temperature before use.
Determination of Adefovir in Human Serum and Urine
Preparation of calibration samples and QC samples The calibration serum samples were prepared by spiking 30 µL of the standard solutions into 270 µL of blank human serum to the final concentrations of 0.5, 1.0, 2.0, 5.0, 20, 50, 200, 500 and 1,000 ng/mL. QC samples (low, medium and high concentration) at 1, 100 and 800 ng/mL were prepared in the same way as the calibration samples and stored at −20°C. The calibration urine samples were prepared by spiking 10 µL of the standard solutions into 90 µL of blank urine (100 µL blank urine and 900 µL purified water were mixed and vortexed for 3 min to achieve 10-fold dilute urine) to the final concentrations of 2.0, 10, 20, 100, 200 and 1,000 ng/mL. QC samples were prepared in a similar manner at the concentrations of 5, 80 and 800 ng/mL.
Sample preparation For serum samples, 10 µL IS (10 µg/mL) working solutions were added into a 2.0-mL centrifuge tube. After evaporation to dryness, an aliquot of 300 µL human serum was mixed and vortexed for 30 s. Then, 900 µL methanol was added and the mixture was vortexed for 3 min. These samples were centrifuged at 12,000 r.p.m. for 6 min. The supernatant was transferred, evaporated to dryness under a gentle stream of nitrogen gas. The residue was reconstituted with 100 µL 5% (w/v) trichloroacetic acid by vortexing for 2 min and then transferred to another clean 1.5 mL centrifuge tube. After further centrifugation at 12,000 r.p.m. for 6 min, 20 µL supernatant was subjected to LC–MS-MS for analysis. For urine samples, 10 µL IS (10 µg/mL) working solutions were added into a 2.0-mL centrifuge tube. After evaporation to dryness, an aliquot of 100 µL dilute human urine (100 µL urine and 900 µL purified water were mixed and vortexed for 3 min to achieve 10-fold dilute urine) was mixed and vortexed for 30 s. Then, 100 µL trichloroacetic acid 10% (w/v) was added and the mixture was vortexed for 2 min. These samples were centrifuged at 12,000 r.p.m. for 6 min. An aliquot of 20 µL of the supernatant was injected into the LC–MS-MS system for analysis.
509 Precision and accuracy The precision and accuracy of the method was evaluated by analyzing serum and urine QC samples, prepared separately from calibration standards at serum concentrations of 1, 100 and 800 ng/mL and urine concentrations of 5, 80 and 800 ng/mL, respectively. The intraday precision and accuracy of the method were assessed by determining the QC samples five times on a single day, and the interday precision and accuracy were estimated by determining the QC samples over three consecutive days. CV and RE were used to express the precision and accuracy, respectively. Recovery and matrix factor Extraction recovery was determined by comparing the peak areas obtained from serum or urine samples with the analytes spiked before and after extraction. The matrix factor was evaluated by dividing the peak area ratio of serum or urine samples spiked after extraction by an equal peak area ratio corresponding to the concentration in neat solution. Stability Stability was determined by analyzing triplicate of spiked samples at each QC level under different conditions. Short-term stability study was performed by analyzing QC samples at the room temperature for 4 h. For freeze and thaw stability, QC samples were subjected to three freeze–thaw cycles (−80°C to room temperature). Long-term stability was assessed by analyzing QC samples stored at −80°C for 153 days. Postpreparative storage stability was assessed by analyzing QC samples left in autosampler vials at 4°C for 24 h. The stability of stock solutions was confirmed by comparing the peak area from stock solutions stored at 4°C for 15 days and at the room temperature (24°C) for 3 h with freshly prepared stock solution. Carry over Carry over was tested by injecting two processed blank matrix samples sequentially after injecting an upper limit of quantification (ULOQ) sample.
Method validation The method validation assays were performed according to the United States Food and Drug Administration guidelines (16). Selectivity Selectivity was investigated by comparing the chromatograms of six different batches of blank human serum or urine with the corresponding samples spiked with PMEA, as well as samples collected from subjects to exclude interference of endogenous substances and metabolites. Linearity and LLOQ The calibration curves for serum were performed with nine concentrations (0.5, 1.0, 2.0, 5.0, 20, 50, 200, 500 and 1,000 ng/mL). The calibration curves for urine were performed with six concentrations (2, 10, 20, 100, 200 and 1,000 ng/mL). The linearity of each calibration curve was determined by plotting the peak area ratio (y) of analytes to IS versus the nominal concentration (x) of analytes with weighted (1/x 2) least square linear regression. The LLOQ was determined as the lowest concentration on the calibration curve at which accuracy (percentage relative error, RE %) within ±20% and a precision (percentage coefficient of variation, CV %) below 20%, and the signal-to-noise ratio was at least 10.
Dilution integrity Dilution integrity experiment was performed for study sample concentrations crossing the ULOQ (the highest standard of the calibration curve). Urine samples at the concentration of 80 µg/mL were diluted 10-fold with human blank urine and purified water to obtain the final test concentrations of 8 µg/mL (n = 6), respectively, and then analyzed by LC–MS-MS. Accuracy and precision should be within ±15%. Application to pharmacokinetic study The validated LC−MS-MS method was applied to investigate the clinical pharmacokinetic profiles of PMEA after oral administration single dose of pradefovir mesylate (10, 30, 60, 90 and 120 mg) and adefovir dipivoxil (ADV) (10 mg) to healthy Chinese volunteers. Fifty healthy volunteers were enrolled in this study, and randomly divided into five groups. Subjects in each group balanced by gender and body mass index, of which six received pradefovir mesylate, two received 10 mg adefovir dipivoxil and two received placebo. Whole blood samples (6–12 mL) were collected by intravenous catheter at ∼0 h ( predosing) and 15 min, 30 min, 45 min, 1 h, 1.5, 2, 3, 4, 6, 8, 12, 16, 24, 36, 48, 72, 96 and 120 h postdosing. The samples were clot at the room temperature for 30 min and centrifuged for 15 min at 3,500 r.p.m. Then upper layer was transferred into a labeled tube. Serum samples were stored at −70°C until analyzed. Urine samples were collected at
510 −12 to 0 h predose, and 0–6, 6–24, 24–48, 48–72, 72–96 and 96– 120 h postdosing. The urine samples were transferred into labeled tubes and stored at −70°C until analyzed. The total volume of urine in each interval was recorded. The study protocol was approved by the First Hospital of Jilin University Ethics Committee (reference number: 140109-007), and conducted in accordance with the Declaration of Helsinki and the principles of Good Clinical Practice. Written informed consents were obtained from all subjects before the study.
Results IS selection The rationale for selecting IS is that basic structure of IS is similar to our target compound. We attempted tenofovir as IS at first, while the chromatographic peak was so bad and hardly separated from PMEA under the premise of optimizing the chromatographic condition. Then entecavir showed not only the perfect chromatographic peak but also good retention. In addition, entecavir could be separated from PMEA in samples. Based on above reasons, we selected entecavir as IS.
Method development The optimum mobile phase was found to be 0.1 mM ammonium formate buffer in water ( pH 2.5 adjusted with formic acid, mobile phase A) and methanol (mobile phase B) at a flow rate of 1.0 mL/min. The retention times of PMEA and IS in serum samples were 5.83 and 6.77 min, respectively. The retention times of PMEA and IS in urine samples were 5.77 and 6.89 min, respectively. The total analytical run time was 8.0 min. ESI provided the optimum sensitivity for PMEA and IS in negative-ion mode. The negative product ion scan spectra of [M−H]− for PMEA and IS are shown in Figure 2. PMEA and IS gave protonated parent ion [M−H]− at m/z 272.0 and m/z 276.0, and the fragment ions of the most significant intensity were observed at m/z 134.0 for PMEA and m/z 149.8 for IS, respectively. So the mass transitions chosen for quantitation were m/z 272.0 → 134.0 for PMEA and m/z 276.0 → 149.8 for IS.
Method validation Selectivity It was proved that under current conditions, no endogenous interferences were observed at the retention times of PMEA and IS. Typical
Figure 2. Product ion mass spectra of [M–H]– ion from PMEA and IS.
Zhang et al. chromatograms of blank serum, serum spiked with PMEA and IS (LLOQ) and subjects after oral administration of pardefovir mesylate are shown in Figure 3, and those of urine samples are shown in Figure 4. Linearity and LLOQ Calibration curves for serum were linear over the concentration range of 0.5–1,000 ng/mL for PMEA with a typical calibration curve equation of y = 0.0554x + 0.00413 (R 2 = 0.9992), where y represents the peak area ratios of PMEA to the IS and x represents the serum concentrations of PMEA. Calibration curves for urine were linear over the concentration range of 2.0–1,000 ng/mL with a typical calibration curve equation of y = 0.0178x − 0.000475 (R 2 = 0.9994). The LLOQ of PMEA was 0.5 ng/mL in human serum samples and 2 ng/mL in human urine samples, respectively, with acceptable accuracy and precision. Precision and accuracy The precision and accuracy data for serum and urine samples are presented in Table I. The intra- and interday precision (CV) value for serum samples was
Article
Development and Validation of a Sensitive LC–MS-MS Method for the Determination of Adefovir in Human Serum and Urine: Application to a Clinical Pharmacokinetic Study Ye Zhang1,†, Lu Shen1,†, Ying Zhan2, Qing-Qing Xiao3, and Jin Yang1,* 1 Center of Drug Metabolism and Pharmacokinetics, China Pharmaceutical University, 24 Tong Jia Xiang, Nanjing, China, 2Department of Pharmacy, Zhongda Hospital of Southeast University, No. 87 Ding Jia Qiao, Nanjing 210009, China, and 3Department of Clinical Pharmacy, China Pharmaceutical University, No. 24 Tong Jia Xiang, Nanjing 210009, China
*Author to whom correspondence should be addressed. Email: [email protected] †
These authors contributed equally to the paper.
Received 6 February 2015; Revised 24 August 2015
Abstract A rapid and sensitive liquid chromatography–tandem mass spectrometry method was developed and validated for the quantification of adefovir (PMEA,9-(2-phosphonylmethoxyethyl) adenine) concentration in human serum and urine. The analysis was performed on a negative ionization electrospray mass spectrometer via multiple reaction monitoring. The monitored transitions were set at m/z 272.0 → 134.0 and m/z 276.0 → 149.8 for PMEA and internal standard, respectively. After protein precipitation, samples were separated by high-performance liquid chromatography on a reversedphase Dikma Diamonsil C18 (250 × 4.6 mm; 5 µm) column with a mobile phase of 0.1 mM ammonium formate buffer–methanol. The calibration curves were linear over the serum concentration range 0.5–1,000 ng/mL and urine concentration range 2.0–1,000 ng/mL. The intra- and interday precision values of PMEA in both serum and urine were lower than 18.16% for low quality control and 13.70% for medium and high quality control. The accuracy, recovery, matrix factor and stability were also within the acceptable limits. The developed method was successfully applied to the pharmacokinetic study of following oral administration of single dose of pradefovir mesylate (10, 30, 60, 90 and 120 mg) and adefovir dipivoxil (10 mg) to healthy Chinese volunteers.
Introduction Pradefovir mesylate is designed as an oral target prodrug of 9-(2-phosphonylmethoxyethyl) adenine nucleotide analog adefovir (or adefovir, PMEA), containing a cyclic phosphonate diester linkage (see Figure 1). Pradefovir employed the HepDirect™ technology and could be activated to PMEA by CYP3A4, which exists primarily in liver cells (1–3). However, another prodrug of adefovir, the broad-spectrum antiviral anti-Hepatitis B virus (HBV) agent, adefovir dipivoxil, converted to adefovir by widespread cellular esterases (see Figure 1). It has been manifested that pradefovir has a better liver targeting property than adefovir
dipivoxil in rats and cynomolgus monkeys (4). Pradefovir mesylate yielded 15 times higher concentrations of radioactivity in the liver than adefovir dipivoxil, but only one-third of the concentrations in the kidney after oral administration to rats. In addition, after oral dosing the same radiolabeled agents in cynomolgus monkeys, pradefovir mesylate yielded 60 times higher levels of total radioactivity in the liver, but only two-thirds of total radioactivity levels in the kidney than adefovir dipivoxil (4). Since the liver is the target organ for HBV infection and the kidney is the site of toxicity, pradefovir is expected to have better efficacy and lower toxicity compared with adefovir dipivoxil.
© The Author 2015. Published by Oxford University Press. All rights reserved. For Permissions, please email: [email protected]
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Zhang et al.
Figure 1. Chemical structures of pradefovir mesylate (A), adefovir dipivoxil (B) and PMEA (C).
Pradefovir and PMEA are the main circulatory substrate after oral dosing of pradefovir mesylate. So far, only one study reported the method of simultaneous determination of pradefovir and PMEA after solid-phase extraction (SPE) from human serum and urine by liquid chromatography–tandem mass spectrometry (LC–MS-MS) under positive and negative ions with switch scanning technology (5). But the authors have not published their method validation data. We tried to develop a method for the determination of pradefovir and PMEA simultaneously in positive ionization mode. However, PMEA produced a much lower intensity in positive ionization mode than pradefovir that could not reach the enough lower limit of quantification (LLOQ) for clinical pharmacokinetic trial, while in negative ionization mode, PMEA produced a higher intensity. Hence, PMEA was analyted in negative ionization mode and was detected separately with pradefovir. The determination of PMEA using high-performance liquid chromatography (HPLC) with ultraviolet detection and HPLC with fluorescence detection has been proved to be too complicated and/or not sensitive enough for the clinical pharmacokinetic study (6–9). So far, several methods have been reported for the assay of PMEA using LC–MS-MS (10–15). Liu et al. (10) reported a LC–MS-MS method that gave a satisfactory sensitivity with LLOQ of 0.10 ng/mL, whereas the serum samples were treated by a complex SPE procedure. Xie et al. (14) developed a LC–MS-MS method with the long analysis time of 14 min. Furthermore, peaks of analytes in the above-mentioned methods were not perfect enough due to their relatively broad shapes. The purpose of our research was to develop and validate a quantitative LC–MS-MS method for the analysis of PMEA with simple pretreatment process and perfect peak shape in human serum and urine for clinical pharmacokinetic studies of pradefovir mesylate.
Experimental Chemicals and reagents Pradefovir mesylate ( purity: 99.54%) and PMEA ( purity: 94.2%) were obtained from Xi’an New Drug Research Co., Ltd (Xi’an, China). Entecavir ( purity: 99.9%) as internal standard (IS) was obtained from Wuhan Belka Biological Pharmaceutical Co., Ltd (Wuhan, China). HPLC-grade methanol and acetonitrile were purchased from Merck (Darmstadt, Germany). Formic acid was purchased from ROE (Newark, New Castle, USA). Ultrapure water was obtained from a UPH Ultrapure Water System (UPH-II-5T, Chengdu, China). Ammonium formate, trichloroacetic acid and other reagents were of analytical grade.
Instruments Chromatographic analysis was performed on a Shimadzu LC-20A series chromatographic system (Shimadzu Corporation UFLC XR, Kyoto, Japan) with two LC-20AD binary pumps, a DGU-20A3 degasser, a SIL20AC autosampler and a CTO-20A column oven. The separation was performed on a Dikma Diamonsil C18 (250 × 4.6 mm; 5 µm) column, at a temperature of 30°C. The mass spectrometer was operated on an API 4000 triple-quadrupole mass spectrometer equipped with an electrospray ionization (ESI) source (AB Sciex, Foster City, CA, USA).
LC–MS-MS analytical conditions The mobile phase was composed of 0.1 mM ammonium formate buffer in water (pH 2.5 adjusted with formic acid, mobile phase A) and methanol (mobile phase B) at a flow rate of 1.0 mL/min. The gradient elution program was as follows: 0–2.0 min, 5% B, 2.0–2.5 min, 5–30% B, 2.5–6.0 min, 30% B, 6.0–6.1 min, 30–5% B, 6.1–8.0 min, 5% B. The eluate from the column was diverted into the MS from 5.0 to 7.5 min. The total analytical run time per sample, including equilibration time, was 8.0 min. The autosampler was conditioned at 4°C and the injection volume was 20 µL for analysis. The mass spectrometer was operated in negative ESI mode with multiple reaction monitoring (MRM), monitoring the transition m/z 272.0 to m/z 134.0 for PMEA and m/z 276.0 to m/z 149.8 for IS. The MS conditions were as follows: curtain gas, 20 psi; source temperature, 550°C; dissociation gas, 6 psi; ion source gas 1, 60 psi; ion source gas 2, 50 psi; ion spray voltage, −4,200 V; declustering potential, −73 V; collision energy, −26 V; entrance potential, −10 V and collision cell exit potential, −10 V. All data were acquired and processed using Analyst 1.5.2 version software (AB Sciex).
Preparation of the stock and working solutions Primary stock solutions of PMEA were prepared in purified water at a concentration of 1.0 mg/mL. A series of working standard solutions (5, 10, 20, 50, 100, 200, 500, 1,000, 2,000, 5,000 and 10,000 ng/mL) and quality control (QC) samples (10, 1,000 and 8,000 ng/mL for serum samples, and 50, 500 and 8,000 ng/mL for urine samples) were obtained by further diluting the stock solutions in methanol/ water (1 : 1, v/v). Stock solutions of IS at 1.0 mg/mL were prepared in methanol/water (1 : 1, v/v). The IS working solution (10 µg/mL) was obtained by diluting the stock solution in methanol/water (1 : 1, v/v). All the solutions were stored at 4°C and were brought to room temperature before use.
Determination of Adefovir in Human Serum and Urine
Preparation of calibration samples and QC samples The calibration serum samples were prepared by spiking 30 µL of the standard solutions into 270 µL of blank human serum to the final concentrations of 0.5, 1.0, 2.0, 5.0, 20, 50, 200, 500 and 1,000 ng/mL. QC samples (low, medium and high concentration) at 1, 100 and 800 ng/mL were prepared in the same way as the calibration samples and stored at −20°C. The calibration urine samples were prepared by spiking 10 µL of the standard solutions into 90 µL of blank urine (100 µL blank urine and 900 µL purified water were mixed and vortexed for 3 min to achieve 10-fold dilute urine) to the final concentrations of 2.0, 10, 20, 100, 200 and 1,000 ng/mL. QC samples were prepared in a similar manner at the concentrations of 5, 80 and 800 ng/mL.
Sample preparation For serum samples, 10 µL IS (10 µg/mL) working solutions were added into a 2.0-mL centrifuge tube. After evaporation to dryness, an aliquot of 300 µL human serum was mixed and vortexed for 30 s. Then, 900 µL methanol was added and the mixture was vortexed for 3 min. These samples were centrifuged at 12,000 r.p.m. for 6 min. The supernatant was transferred, evaporated to dryness under a gentle stream of nitrogen gas. The residue was reconstituted with 100 µL 5% (w/v) trichloroacetic acid by vortexing for 2 min and then transferred to another clean 1.5 mL centrifuge tube. After further centrifugation at 12,000 r.p.m. for 6 min, 20 µL supernatant was subjected to LC–MS-MS for analysis. For urine samples, 10 µL IS (10 µg/mL) working solutions were added into a 2.0-mL centrifuge tube. After evaporation to dryness, an aliquot of 100 µL dilute human urine (100 µL urine and 900 µL purified water were mixed and vortexed for 3 min to achieve 10-fold dilute urine) was mixed and vortexed for 30 s. Then, 100 µL trichloroacetic acid 10% (w/v) was added and the mixture was vortexed for 2 min. These samples were centrifuged at 12,000 r.p.m. for 6 min. An aliquot of 20 µL of the supernatant was injected into the LC–MS-MS system for analysis.
509 Precision and accuracy The precision and accuracy of the method was evaluated by analyzing serum and urine QC samples, prepared separately from calibration standards at serum concentrations of 1, 100 and 800 ng/mL and urine concentrations of 5, 80 and 800 ng/mL, respectively. The intraday precision and accuracy of the method were assessed by determining the QC samples five times on a single day, and the interday precision and accuracy were estimated by determining the QC samples over three consecutive days. CV and RE were used to express the precision and accuracy, respectively. Recovery and matrix factor Extraction recovery was determined by comparing the peak areas obtained from serum or urine samples with the analytes spiked before and after extraction. The matrix factor was evaluated by dividing the peak area ratio of serum or urine samples spiked after extraction by an equal peak area ratio corresponding to the concentration in neat solution. Stability Stability was determined by analyzing triplicate of spiked samples at each QC level under different conditions. Short-term stability study was performed by analyzing QC samples at the room temperature for 4 h. For freeze and thaw stability, QC samples were subjected to three freeze–thaw cycles (−80°C to room temperature). Long-term stability was assessed by analyzing QC samples stored at −80°C for 153 days. Postpreparative storage stability was assessed by analyzing QC samples left in autosampler vials at 4°C for 24 h. The stability of stock solutions was confirmed by comparing the peak area from stock solutions stored at 4°C for 15 days and at the room temperature (24°C) for 3 h with freshly prepared stock solution. Carry over Carry over was tested by injecting two processed blank matrix samples sequentially after injecting an upper limit of quantification (ULOQ) sample.
Method validation The method validation assays were performed according to the United States Food and Drug Administration guidelines (16). Selectivity Selectivity was investigated by comparing the chromatograms of six different batches of blank human serum or urine with the corresponding samples spiked with PMEA, as well as samples collected from subjects to exclude interference of endogenous substances and metabolites. Linearity and LLOQ The calibration curves for serum were performed with nine concentrations (0.5, 1.0, 2.0, 5.0, 20, 50, 200, 500 and 1,000 ng/mL). The calibration curves for urine were performed with six concentrations (2, 10, 20, 100, 200 and 1,000 ng/mL). The linearity of each calibration curve was determined by plotting the peak area ratio (y) of analytes to IS versus the nominal concentration (x) of analytes with weighted (1/x 2) least square linear regression. The LLOQ was determined as the lowest concentration on the calibration curve at which accuracy (percentage relative error, RE %) within ±20% and a precision (percentage coefficient of variation, CV %) below 20%, and the signal-to-noise ratio was at least 10.
Dilution integrity Dilution integrity experiment was performed for study sample concentrations crossing the ULOQ (the highest standard of the calibration curve). Urine samples at the concentration of 80 µg/mL were diluted 10-fold with human blank urine and purified water to obtain the final test concentrations of 8 µg/mL (n = 6), respectively, and then analyzed by LC–MS-MS. Accuracy and precision should be within ±15%. Application to pharmacokinetic study The validated LC−MS-MS method was applied to investigate the clinical pharmacokinetic profiles of PMEA after oral administration single dose of pradefovir mesylate (10, 30, 60, 90 and 120 mg) and adefovir dipivoxil (ADV) (10 mg) to healthy Chinese volunteers. Fifty healthy volunteers were enrolled in this study, and randomly divided into five groups. Subjects in each group balanced by gender and body mass index, of which six received pradefovir mesylate, two received 10 mg adefovir dipivoxil and two received placebo. Whole blood samples (6–12 mL) were collected by intravenous catheter at ∼0 h ( predosing) and 15 min, 30 min, 45 min, 1 h, 1.5, 2, 3, 4, 6, 8, 12, 16, 24, 36, 48, 72, 96 and 120 h postdosing. The samples were clot at the room temperature for 30 min and centrifuged for 15 min at 3,500 r.p.m. Then upper layer was transferred into a labeled tube. Serum samples were stored at −70°C until analyzed. Urine samples were collected at
510 −12 to 0 h predose, and 0–6, 6–24, 24–48, 48–72, 72–96 and 96– 120 h postdosing. The urine samples were transferred into labeled tubes and stored at −70°C until analyzed. The total volume of urine in each interval was recorded. The study protocol was approved by the First Hospital of Jilin University Ethics Committee (reference number: 140109-007), and conducted in accordance with the Declaration of Helsinki and the principles of Good Clinical Practice. Written informed consents were obtained from all subjects before the study.
Results IS selection The rationale for selecting IS is that basic structure of IS is similar to our target compound. We attempted tenofovir as IS at first, while the chromatographic peak was so bad and hardly separated from PMEA under the premise of optimizing the chromatographic condition. Then entecavir showed not only the perfect chromatographic peak but also good retention. In addition, entecavir could be separated from PMEA in samples. Based on above reasons, we selected entecavir as IS.
Method development The optimum mobile phase was found to be 0.1 mM ammonium formate buffer in water ( pH 2.5 adjusted with formic acid, mobile phase A) and methanol (mobile phase B) at a flow rate of 1.0 mL/min. The retention times of PMEA and IS in serum samples were 5.83 and 6.77 min, respectively. The retention times of PMEA and IS in urine samples were 5.77 and 6.89 min, respectively. The total analytical run time was 8.0 min. ESI provided the optimum sensitivity for PMEA and IS in negative-ion mode. The negative product ion scan spectra of [M−H]− for PMEA and IS are shown in Figure 2. PMEA and IS gave protonated parent ion [M−H]− at m/z 272.0 and m/z 276.0, and the fragment ions of the most significant intensity were observed at m/z 134.0 for PMEA and m/z 149.8 for IS, respectively. So the mass transitions chosen for quantitation were m/z 272.0 → 134.0 for PMEA and m/z 276.0 → 149.8 for IS.
Method validation Selectivity It was proved that under current conditions, no endogenous interferences were observed at the retention times of PMEA and IS. Typical
Figure 2. Product ion mass spectra of [M–H]– ion from PMEA and IS.
Zhang et al. chromatograms of blank serum, serum spiked with PMEA and IS (LLOQ) and subjects after oral administration of pardefovir mesylate are shown in Figure 3, and those of urine samples are shown in Figure 4. Linearity and LLOQ Calibration curves for serum were linear over the concentration range of 0.5–1,000 ng/mL for PMEA with a typical calibration curve equation of y = 0.0554x + 0.00413 (R 2 = 0.9992), where y represents the peak area ratios of PMEA to the IS and x represents the serum concentrations of PMEA. Calibration curves for urine were linear over the concentration range of 2.0–1,000 ng/mL with a typical calibration curve equation of y = 0.0178x − 0.000475 (R 2 = 0.9994). The LLOQ of PMEA was 0.5 ng/mL in human serum samples and 2 ng/mL in human urine samples, respectively, with acceptable accuracy and precision. Precision and accuracy The precision and accuracy data for serum and urine samples are presented in Table I. The intra- and interday precision (CV) value for serum samples was
