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Validation of a rapid and sensitive liquid chromatography-tandem mass spectrometry method for free and total mycophenolic acid.

The potential utility of monitoring plasma total mycophenolic acid (tMPA) [3] concentrations for optimizing therapy with the prodrug mycophenolate mofetil (MMF) has been demonstrated in several studies (1-5). Because MPA is highly protein bound, there is also a need to investigate free MPA (fMPA). In vitro investigations with recombinant human inosine monophosphate dehydrogenase II-and phytohemagglutinin A-stimulated human peripheral blood mononuclear cells support the concept that the free fraction of MPA is the pharmacologically active form of the drug (6). A case of severe leukopenia associated with renal failure showing a substantially increased fMPA area under the concentration-time curve (AUC) despite a tMPA AUC within the putative therapeutic range was observed in a pancreas/renal transplant recipient (7). The mean fMPA fraction in renal transplant patients with chronic renal insufficiency was found to be more than double that of patients with normal renal function (8). An investigation involving pediatric renal transplant recipients receiving a triple therapy of cyclosporin A, MMF, and steroids revealed a significant correlation between the fMPA AUC and the occurrence of leukopenia and/or severe infections (5). Surprisingly, no association was observed in this same study between the fMPA AUC and the incidence of acute rejection episodes, although the tMPA AUC showed a significant negative correlation with acute rejection (5). Current HPLC methods that use ultraviolet (UV) detection for fMPA have detection limits of 5 [micro]g/L at 215 run and 15 [micro]g/L at 254 run (9). In the latter report, -9% of samples drawn for the determination of MPA pharmacokinetic profiles had a fMPA concentration below the detection limit of 5 [micro]g/L. Furthermore, the proportion of samples with fMPA values below the UV detection limit is higher in patient populations in which tMPA concentrations are generally lower, e.g., liver transplant recipients during the early posttransplantation phase or bone marrow transplant recipients, than in kidney recipients.

Because of its flexibility and high specificity, liquid chromatography-tandem mass spectrometry (LC-MS/ MS) is finding increasing application for the quantification of numerous analytes and would appear to be ideally suited to establishing a highly sensitive and specific assay for fMPA determination. A LC-MS/MS procedure has recently been described that has a lower limit of quantification (LLOQ) of 2.5 [micro]g/L but requires a chromatography time of 12 min (10). The aim of the present study was to develop a selective method for quantification of fMPA with a LLOQ of 0.5 [micro]g/L, which would allow more accurate quantification of fMPA concentrations at the lower end of the measuring range and thus enable better assessment of any association between fMPA and acute rejection. In developing such a method, consideration was made in particular to two potential sources of interference: ion suppression (11-13) and in-source fragmentation of MPA metabolites and internal standard (14). We now describe the validation of a reliable, simple, sensitive, and rapid procedure for determining MPA in plasma and ultrafiltrates by LC-MS/MS.

Materials and Methods


MPA was obtained from Sigma-Aldrich and 7-O-mycophenolic acid [beta]-glucuronide (MPAG) from Analytical Services International Limited. The acyl glucuronide of MPA (AcMPAG) and the carboxybutoxy ether of MPA (MPAC) were kind gifts of Hoffmann-La Roche AG (Basel, Switzerland).

Stock solutions of MPA and the internal standard MPAC, each at a concentration of 1 g/L, were prepared separately in acetonitrile and stored at -20[degrees]C. For routine tMPA determinations, a single-point calibration was performed by diluting an aliquot of the MPA stock solution in drug-free EDTA plasma from blood donors to yield a final MPA concentration of 3.0 mg/L. To check the linearity of the method, additional calibrators with the following tMPA concentrations were prepared by appropriate dilution of the stock solution in drug-free EDTA plasma: 0.05, 1.0, 10.0, and 50.0 mg/L.

For fMPA determinations, a single-point calibration was also routinely used; an aliquot of the stock solution was diluted to a final MPA concentration of 50.0 [micro]g/L in a protein-free solution of KHZP04 (67 mmol/L) and NaCl (9 g/L) adjusted to pH 7.4 with NaCH. To test for linearity, additional calibrators were prepared in the above phosphate buffer with the following MPA concentrations: 0.5, 1.0, 5.0, 10, 100, 500, and 1000 [micro]g/L. In preliminary experiments we had established that the protein-free phosphate calibrator could be used as a practical and economical alternative to a calibrator in drug-free ultrafiltrates from EDTA plasma obtained from blood donors.

In-house quality-control samples were prepared by appropriate dilution of a separate MPA stock solution (1 g/L) in the phosphate buffer (final MPA concentrations, 0.5, 50, and 333.0 mg/L) as well as in drug-free plasma pools (final MPA concentrations, 0.2, 2.0, and 25.0 mg/L).


HPLC-grade methanol, ammonium acetate, acetic acid, acetonitrile, perchloric acid, and sodium tungstate were obtained from Merck.

fMPA. We added 300 [micro]L of plasma to the sample reservoir of a Centrifree[R] Micropartition system (Amicon) (6), and the tube was centrifuged for 40 min at 2000g and 20[degrees]C. We then mixed 100 [micro]L of the resulting ultrafiltrate with 10 [micro]L of the internal standard solution (2.5 mg/L MPAC in acetonitrile) in an autosampler vial, which was then placed on the autosampler for direct injection (20 [micro]L) on the LC column.

fMPA. We treated 200 [micro]L of plasma, calibrator, or quality-control sample with 20 [micro]L of perchloric acid (150 g/L), 20 [micro]L of sodium tungstate (250 g/L), and 100 [micro]L of internal standard (15 mg/L MPAC in acetonitrile) as described previously (9). After centrifugation at 2000g for 5 min, the supernatant was diluted 1:10 (by volume) with deionized water and transferred to an autosampler vial, which was then placed on the autosampler; 20 [micro]L of each supernatant was injected on the LC column. We investigated extraction efficiency by adding MPA to drug-free plasma to a final concentration of 1 mg/L. Peak areas obtained for the extracted samples were then compared with peak areas obtained for a methanol solution containing MPA at a concentration of 1 mg/L.

To test for potential carryover from previous samples, we performed the following experiments. MPA was added to a drug-free ultrafiltrate to a nominal concentration of 1000 [micro]g/L. Control samples with a MPA concentration of 5 g/L were then placed in the autosampler before and after the "high" MPA ultrafiltrate. This experiment was repeated six times. A similar protocol was used for total MPA. In this case MPA was added to drug-free plasma to a nominal concentration of 50 [micro]g/L, and the sample was extracted as described above. Controls with MPA concentrations of 0.5 mg/L were then placed in the autosampler before and after this high MPA plasma extract. The experiment was repeated six times.


An online extraction method with a column-switching technique combined with analytical LC and electrospray MS/MS detection was used for quantification of fMPA and tMPA. The extraction column was an Oasis[R] HLB column (2.1 x 20 mm; Waters), and the analytical column was an Aqua Perfect [C.sub.18] column (3.0 x 150 mm; MZ-Analysentechnik) maintained at 40[degrees]C with a DuPont column oven. The system also included a Series 200 binary pump (Perkin-Elmer), a M480 pump (Dionex), and a 10-port Rheodyne valve. Samples were injected with a Series 200 autoinjector (Perkin-Elmer) fitted with a 200-[micro]/L sample loop through the Rheodyne valve onto the extraction column; the extraction column was then washed for 1 min (flow rate, 1 mL/min) with 10 g/L acetic acid. The valve position was then switched to allow the bound material to be eluted from the extraction cartridge in back-flush mode onto the analytical column with acetonitrile-methanol-10 g/L acetic acid (21:63:16 by volume) containing 0.1 mg/L ammonium acetate at a flow rate of 500 [micro]L/min. After 1.5 min, the Rheodyne valve position was again switched to allow the extraction column to be cleansed with methanol at a flow rate of 2 mL/min for 0.5 min and then reequilibrated with 10 g/L acetic acid at a flow rate of 4 mL/min for 0.5 min. Total run time was 4 min. The corresponding valve positions, elution solvents, and pump flow rates for the chromatographic step are given in Table 1. The analytes that eluted from the HPLC column were introduced into the turbo-ion spray source with a split of 1:10. High-purity argon was used as the collision gas. Ionization was achieved in the positive-ion mode with an ionization voltage of 5500 V, an orifice voltage and collision energy of 20 eV, and a heater probe temperature of 400[degrees]C. Sample analysis was performed in the multiple-reaction monitoring (MRM) mode using the following transitions: m/z 338.2[right arrow]207.1 for the ammonium adduct ion of MPA and m/z 438.2[right arrow]207.1 for the internal standard MPAC. A personal computer running Applied Biosystems Analyst (Ver. 1.2) software was used to control the LC-MS/MS and to record the output signals from the detector. Integration of peak areas, calculation of peak-area ratios, calculation of the calibration line, and calculation of the MPA concentrations were performed with the Analyst 1.2 software.


Ion suppression attributable to matrix effects is a well-known phenomenon in LC-MS/MS quantification methods (11-13). Therefore, to investigate potential ion suppression in our system, we performed the following postcolumn infusion experiment (11). A continuous infusion of MPA (1g/L) was introduced at a flow rate of 5 [micro]L/min into the effluent from the HPLC column before introduction into the electrospray tandem mass spectrometer. Supernatants from different pretreated drug-free plasma samples (n = 5) and drug-free plasma ultrafiltrates (n = 5), as well as samples of deionized water (n = 5), were injected separately into the LC-MS/MS system, and the MS/MS response of the MRM transition for the ammonium adduct of MPA (m/z 338.2[right arrow]207.1) was recorded.


We used 106 plasma samples from routine MPA monitoring of patients treated with MMF for a method comparison with a validated HPLC procedure for fMPA and tMPA (9). tMPA was measured in all samples, and fMPA was determined in 52 samples after ultrafiltration.


We collected four patient plasma pools and divided them into three portions. Two portions were immediately stored at -20[degrees]C, and one portion was subjected to ultrafiltration at baseline for measurement of fMPA. The frozen plasma portions were thawed at 2 and 6 months for ultrafiltration and measurement of fMPA.


The nonparametric regression procedure of Passing and Bablok (15) was used for comparison of the LC-MS/MS and the HPLC procedures. The regression equations are given in the Results, together with the 95% confidence intervals for the estimates of slope and intercept. The dispersion of the residuals are documented as the 95% median distance. For comparison, the standard deviation of the residuals, [S.sub.y|x], calculated using the standard principal component procedure, is also given. Agreement between the methods was also assessed by plotting the method differences against the method means (16).



Mass interference attributable to in-source fragmentation of drug metabolites or internal standard can be a problem if these substances are not chromatographically separated from the parent drug before introduction of the sample into the ion source (14,17). To confirm that adequate separation of MPA from its metabolites and MPAC had been achieved, we injected purified samples of the respective substances on the HPLC column and monitored the elution profiles from the column (Fig. 1). Typical elution times were as follows: MPAG, 2.73 min; AcMPAG, 2.95 min; MPA, 3.31 min; and internal standard (MPAC), 3.20 min. Shown in Fig. 2 are the MRM transitions for the ammonium adduct (m/z 338.1[right arrow]207.1) and the protonated adduct (m/z 321.1[right arrow]207.1) of MPA that were obtained after injection of an extracted plasma sample and an ultrafiltrate (only the m/z 338.1[right arrow]207.1 transition is shown) from a patient being treated with MMF. MRM transitions are seen not only at the elution time of MPA but also at the elution times of MPAG and AcMPAG because of insource fragmentation of these metabolites to MPA. Because MPA is effectively separated on the chromatographic column (Fig. 1), no interference from MPAG, AcMPAG, or MPAC would be expected with our procedure.



To avoid interference by ion suppression, appropriate chromatographic separation is also necessary before introduction of the analyte into the ion source. Chromatographic run times should be long enough to ensure that matrix effects do not alter the MS/MS response of either MPA or the internal standard. To determine ion suppression, we used the postcolumn infusion experiment described by King et al. (11), in which a constant infusion of MPA was introduced into the effluent from the HPLC column before introduction into the electrospray tandem mass spectrometer. Shown in Fig. 3A is a typical ion chromatogram in which the response of the MRM transition of the ammonium adduct of MPA was continuously monitored. At the time point 0 min, pure deionized water was injected on the HPLC column. Suppression of the MS/MS response was observed between 2.3 and 2.5 min (Fig. 3A). This results from the frontline water fraction, which elutes earliest from the column. Shown in Fig. 3B is a typical ion chromatogram after injection of a drug-free plasma ultrafiltrate at 0 min. In addition to the ion suppression observed as a result of the high water content in the extraction column, additional ion suppression was seen between 2.5 and 2.8 min, presumably resulting from matrix effects caused by substances in the plasma. Similar profiles with identical ion suppression windows were observed when the ultrafiltrates from different individuals (n = 5) and when the diluted supernatants (1:10 with deionized water) from pretreated drug-free plasma samples (n = 5) were investigated. Fig. 3C shows the total ion current of an extracted plasma sample from a patient receiving MMF. As can be seen, both MPA and the internal standard elute well after the ion suppression window.



To further investigate potential interference of other drugs with this analytical procedure, we examined the ion chromatograms of 30 samples from transplant recipients not treated with MMF but who were receiving other immunosuppressive drugs, such as sirolimus, everolimus, cyclosporin, tacrolimus, and azathioprine, as well as other drugs commonly prescribed to transplant recipients, such as ketoconazole and clarithromycin. No MS/MS responses were observed for the MRM transitions corresponding to the ammonium adduct ion of MPA (m/z 338.1[right arrow]207.1) and the internal standard MPAC (m/z 438.2[right arrow]207.1).



The LLOQs were 0.5 [micro]g/L (CV = 7.0%; n = 20) for fMPA and 0.05 mg/L (CV = 6.7%; n = 20) for tMPA with an injection volume of 20 [micro]L. The fMPA method was linear over the working range between 0.5 and 1000 [micro]g/L (r >0.999), and the tMPA method was linear between 0.05 and 50 mg/L (r >0.999). Performance characteristics were tested at several concentrations of MPA added to the phosphate buffer or drug-free plasma (Table 2). The measured values for the in-house control samples were within 10% of the nominal values. The within- and between-run CVs were all <10%. The extraction efficiency (SD) for tMPA was 72.4 (2.7)% (n = 5), a value similar to the range (73-77%) we reported after detection with HPLC-UV (18).

Because carryover can be an issue with column-switching methods, we investigated this phenomenon using the following protocol. In the case of fMPA measurement, we placed a low control (nominal concentration, 5 [micro]g/L) in the autosampler before and after an ultrafiltrate to which MPA was added to give a final concentration of 1000 [micro]g/L. We observed no significant difference (P = 0.30; n = 6) between the mean fMPA values measured before [5.00 (0.37) [micro]g/L] or after [4.81 (0.21) [micro]g/L] the high-concentration MPA ultrafiltrate. For tMPA measurements, we measured a low control before and after an extracted plasma to which MPA had been added to a final concentration of 50 mg/L. Again, we observed no significant difference (P = 0.42; n = 6) between the values measured before [0.46 (0.02) mg/L] or after [0.47 (0.02) mg/L] the high-concentration MPA plasma extract.


We compared the present LC-MS/MS procedure with a validated HPLC-UV method (9), using fresh plasma samples from patients being treated with MMF. tMPA concentrations were determined in 106 plasma samples by both methods. We subjected 52 plasma samples to ultrafiltration and quantified the fMPA concentrations. Method difference plots for fMPA (Fig. 4, A and B) and tMPA (Fig. 4C) revealed good agreement between both methods over the clinically relevant ranges. The fMPA concentrations measured by HPLC-UV were 5.3-271 [micro]g/L (median, 27.2 [micro]g/L), and those measured by LCMS/MS were 4.3-281.4 [micro]g/L (median, 26.6 [micro]g/L). The mean absolute difference between the two methods was 1.87 [micro]g/L (mean relative difference, 2.56%) over the range of 5.3-281 [micro]g/L (Fig. 4A) as measured with HPLC-UV. The equation for the Passing-Bablok regression line was: y = 0.95x + 0.27 [micro]g/L ([r.sup.2] = 0.99; 95% median distance of the residuals of the Passing-Bablok regression, 8.36 [micro]g/L; Sy1X = 4.28 [micro]g/L). To check that agreement remained good in the lower range, we separately analyzed (Fig. 4B) the 37 ultrafiltrates with fMPA values between 5.3 and 47.5 [micro]g/L (HPLC-UV). The mean absolute difference between the two methods over this range was 0.23 [micro]g/L (mean relative difference, 1.40%). The equation for the Passing-Bablok regression line was: y = 1.00x - 0.51 [micro]g/L ([r.sup.2] = 0.97; 95% median distance of the residuals of the Passing-Bablok regression, 3.44 [micro]g/L; Syl, = 1.63 [micro]g/L).


In the case of tMPA (Fig. 4C), concentrations ranged from 0.2 to 24 mg/L (median, 2.0 mg/L) as measured by HPLC-UV and from 0.2 to 25.3 mg/L (median, 2.1 mg/L) as measured by LC-MS/MS. The mean absolute difference between the two methods was 0.07 mg/L (relative difference, 0.5%). The equation for the Passing-Bablok regression line was: y = 0.98x + 0.03 mg/L (r z = 0.99; 95% median distance of the residuals of the Passing-Bablok regression, 0.53 mg/L; Syl, = 0.23 mg/L).


We tested the effect of storage of plasma at -20[degrees]C and one freeze-thaw cycle before ultrafiltration on fMPA concentrations using four patient pools. Stored frozen samples were thawed after 2 or 6 months, and the fMPA concentrations were determined. The results are presented in Table 3. We observed good agreement (<15% deviation) between the fMPA concentrations at baseline and after storage.


Despite the inherent high specificity of MS/MS for analytical quantification of drugs in biological specimens, results can be subject to interferences caused by in-source fragmentation, in particular of conjugated drug metabolites (14,17), and by ion suppression (11-13). When establishing an analytical procedure using this instrumentation, careful consideration must therefore be given to these phenomena.

In-source fragmentation is a routinely used procedure in MS to obtain additional structural information by inducing low-energy collision dissociation, in which the neutral loss of small molecules such as water and methanol, as well as glucuronides and sulfates, can be observed. It has been shown that MPAG, the major glucuronide metabolite of MPA, as well as the internal standard MPAC (butoxy ether of MPA) undergo such in-source fragmentation to MPA (14). The quasimolecular ion of AcMPAG can also undergo in-source fragmentation to yield the MPA ion. This in-source fragmentation produces additional peaks in the ion current of the ammonium or protonated adduct ions of MPA (Fig. 2). Coelution of MPAG or AcMPAG with MPA would therefore give falsely increased MPA concentrations. As can be seen from Figs. 1 and 2, the chromatographic step of our procedure effectively separates the MPA glucuronides from both MPA and MPAC, thereby avoiding interference from in-source fragmentation.

Matrix effects have been described that can significantly affect the ionization of the analyte, thereby causing a reduction in the MS/MS response. These effects are apparently a result of extraction of endogenous substances from plasma that coelute with the analytes (11, 12). It is therefore essential that such substances are separated from the analyte and internal standard. In the present procedure, a chromatographic run time of 4 min was found to be sufficient to effectively separate MPA and the internal standard MPAC from matrix components that cause ion suppression.

The LC-MS/MS method described here for the determination of fMPA and tMPA is rapid, reproducible, and specific. Despite the column-switching steps, we observed no significant carryover. The present procedure offers substantial advantages over available LC-MS/MS and HPLC-UV methods. It has a LLOQ of 0.5 [micro]g/L and at the same time a short analysis time of only 4 min. Willis et al. (10) described a LC-MS/MS procedure with an atmospheric pressure chemical ionization interface in negative ionization mode for the quantification of fMPA that has a total chromatographic analysis time of 12 min and a LLOQ of 2.5 [micro]g/L. A LC-MS procedure with electrospray ionization in the negative-ion mode has been used to detect trace quantities of MPA in preparations of MPAG (19, 20). The LLOQ of this method was 9.2 [micro]g/L, and the retention time was 8.4 min (19). The detection limit for fMPA using HPLC-UV is 5 [micro]g/L, and run times are generally longer (9). Because of the high sensitivity of the LC-MS/MS procedure, only 20 [micro]L of ultrafiltrate, as opposed to 100 [micro]L for HPLC-UV, is needed for fMPA analysis. A rerun of the sample is therefore possible if required. This is of particular interest, for example, for pediatric samples.

In a recently published automated method for determination of fMPA by dialysis across a membrane followed by concentration of the dialysate on a trace enrichment column and LC, a limit of quantification of 6 [micro]g/L was reported (21). This automated method requires a sample volume of 370 [micro]L and eliminates the necessity of an ultrafiltration step. The method of Mandla et al. (21) does have the advantage that free MPAG concentrations can also be obtained in the same run. The chromatographic run time was 25 min, and up to 50 samples could be processed in 1 day. With our method, at least 90 samples can be processed within a working day, making this method suitable for routine use.

Plasma samples could be stored for up to 6 months at -20[degrees]C without any substantial change (<15%) in fMPA concentrations compared with baseline. Mandla et al. (21) also reported good stability for samples that were stored at -20[degrees]C for 2 weeks and exposed to three freeze-thaw cycles. In one lipemic sample, fMPA and free MPAG were significantly increased when the sample was stored at room temperature for 20 h. They attributed this to a possible displacement of MPA and MPAG from their binding sites on albumin by free fatty acids. We have previously reported (22) that storage of plasma at room temperature for 24 h or longer can lead to a significant increase in tMPA concentrations, presumably as a result of cleavage of MPAG by plasma glucuronidases or other hydrolases. For short-term storage, plasma should be kept at 4[degrees]C, whereas for longer periods plasma should be stored at -20[degrees]C.

The LC-MS/MS method was further validated by comparison with an established HPLC-UV method, using plasma samples from patients receiving MMF. We obtained very good agreement for the determination of both fMPA and tMPA. We observed no relevant bias between the two procedures, and the slopes of the regression lines were <5% from the line of identity.

In conclusion, we describe a rapid, sensitive LC-MS/MS procedure for the quantification of both fMPA and tMPA that is not subject to interference from ion suppression or in-source fragmentation. Because plasma samples can be stored frozen at -20[degrees]C for up to 6 months, this method could be suitable for the measurement of fMPA and tMPA not only in routine clinical practice but also in multicenter outcome studies.

We gratefully acknowledge the expert technical assistance of Melanie Fischer, Sandra Gotze, and Hamza Sinanoglu.


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[1] Department of Clinical Chemistry, George-August University Gottingen, D-37075 Gottingen, Germany.

[2] Central Institute for Clinical Chemistry and Laboratory Medicine, Klinikum Stuttgart, Stuttgart, Germany.

[3] Nonstandard abbreviations: MPA, mycophenolic acid; MMF, mycophenolate mofetil; AUC, area under the concentration-time curve; UV, ultraviolet; LC-MS/MS, liquid chromatography-tandem mass spectrometry; LLOQ, lower limit of quantification; MPAG, 7-O-mycophenolic acid-/3-glucuronide; AcMPAG, acyl glucuronide of mycophenolic acid; MPAC, carboxybutoxy ether of mycophenolic acid; and MRM, multiple reaction monitoring.

* Address correspondence to this author at: Abteilung Klinische Chemie, Zentrum Innere Medizin, Georg-August-Universitit Gottingen, Robert-KochStrasse 40, 37075 Gottingen, Germany. Fax 49-551-398551; e-mail varmstro@

Received July 8, 2003; accepted October 13, 2003.

Previously published online at DOI: 10.1373/clinchem.2003.024323
Table 1. Elution protocol for the online extraction method
with a column-switching technique combined with
analytical LC.

 Binary pump (b)

Time, Valve Solvent Solvent Flow rate,
min position (a) A B [micro]L/min

0.0-1.0 B 100% 1000
1.0-2.5 A 100% 1000
2.5-3.0 B 100% 2000
3.0-3.5 A 100% 5000
3.5-4.0 B 100% 4000

 M480 pump

Time, Solvent Flow rate,
min C (c) [micro]L/min

0.0-1.0 100% 500
1.0-2.5 100% 500
2.5-3.0 100% 500
3.0-3.5 100% 500
3.5-4.0 100% 500

(a) In valve position B, the extraction cartridge (binary pump)
and the analytical column (M480 pump) are in separate elution
circuits. In position A, the extraction cartridge and the analytical
column are in line in the same elution circuit (M480 pump).

(b) Solvent A, 10 g/L acetic acid; solvent B, methanol.

(c) Solvent C, acetonitrile-methanol-10 g/L acetic acid (21:63:16
by volume) containing 0.1 mg/L ammonium acetate.

Table 2. Analytical recovery and imprecision of the
LC-MS/MS method.

 Within run (n = 20) Between run (n = 10)

 Recovery, % CV, % Recovery, % CV, %
 0.5 [micro]g/L 103.4 7.0 109.2 8.1
 50.0 [micro]g/L 99.1 3.7 99.2 3.6
 333.0 [micro]g/L 103.6 2.5 98.4 6.9
 0.2 mg/L 99.4 2.0 100 6.6
 2.0 mg/L 99.4 1.8 97.9 4.7
 25.0 mg/L 102.1 4.0 102.3 4.5

Table 3. fMPA concentrations determined after storage of
plasma pools at 20[degrees]C for 2 or 6 months.

 fMPA, [micro]g/L

Plasma pool Baseline 2 months 6 months

 1 15.9 17.5 16.3
 2 11.3 11.7 10.3
 3 27.3 31.2 28.2
 4 9.2 8.9 7.9
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Title Annotation:Drug Monitoring and Toxicology
Author:Streit, Frank; Shipkova, Maria; Armstrong, Victor William; Oellerich, Michael
Publication:Clinical Chemistry
Date:Jan 1, 2004
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