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Quantification of riboflavin, flavin mononucleotide, and flavin adenine dinucleotide in human plasma by capillary electrophoresis and laser-induced fluorescence detection.

Riboflavin is a water-soluble vitamin that serves as a precursor for flavin mononucleotide (FMN) [1] and FAD (1, 2). These coenzymes are involved in several reduction-oxidation reactions and take part in the metabolism of other vitamins, e.g., folate and vitamin [B.sub.6] (2).

Riboflavin deficiency has been claimed to be quite common, and high prevalences have been reported, particularly in developing countries (3-5). The riboflavin status in humans has been assessed by clinical signs, by determination of the urinary excretion of the vitamin (6), and by measurement of the activity ratio of glutathione reductase in erythrocytes (7, 8).

For the determination of vitamin concentrations, several techniques have been developed, including microbiological (9), fluorometric (9), and liquid chromatographic methods (10). Most HPLC methods have been designed for the detection of high concentrations of riboflavin in food, pharmaceutical preparations, and urine (10). Only a few have been used to measure riboflavin in whole blood (11-14), serum (11, 14,15), or plasma (16-18).

During the last decade, capillary electrophoresis (CE) methods have been developed for several biomedical applications (19). Compared with HPLC, CE generally has the advantage of small sample requirements, short separation times, and high resolution (20). The consumption of organic solvents and other chemicals is usually lower than in HPLC.

We describe here a sensitive and robust CE method for the quantification of low physiological concentrations of riboflavin, FMN, and FAD in human plasma. The method is based on micellar electrokinetic capillary chromatography combined with laser-induced fluorescence (LIF) detection.

Materials and Methods

CHEMICALS

Riboflavin, FMN, FAD, and lumiflavin were purchased from Sigma-Aldrich Norway. Isoriboflavin was kindly supplied as a gift by F. Hoffmann-La Roche & Co. (Basel, Switzerland). Sodium dodecyl sulfate, trichloroacetic acid (TCA), and boric acid were obtained from Merck. Methanol (HPLC grade) was purchased from Rathburn Chemicals, and N-methylformamide was purchased from Fluka. Doubly distilled water purified on a MilliQ Plus Water Purification System (Millipore) was used for preparation of all aqueous solutions.

INSTRUMENTATION

Solid-phase extraction was performed on a programmable Gilson ASPEC sample processor (Gilson Medical Electronics) equipped with a Rheodyne Model 7010 injector valve and a 3000-[micro]L sample loop.

CE was performed on a Beckman P/ACE System 2210 coupled to a P/ACE LIF Detector (Beckman Instruments). The LIF detector was connected to a 20 mW 488 nm argon laser from Uniphase. The fluorescence was monitored using a 530 nm DF30 band-pass filter from Omega Optical. Beckman System Gold software (Ver. 8.10) was used for system control and data collection and processing.

CAPILLARY ELECTROPHORESIS

Uncoated fused-silica capillaries (Composite Metal Services) with external and internal diameters of 375 [micro]m and 75 p,m, respectively, were used. The total capillary length was 67 cm, and the effective separation length was 60 cm. The separation buffer consisted of 100 mmol/L sodium borate, pH 7.9, containing 50 mmol/L sodium dodecyl sulfate, 100 mL/L methanol, and 20 mL/L N-methylformamide. All solutions were passed through 0.2 [micro]m membrane filters (Schleicher & Schnell) before use. Capillaries were preconditioned before each electrophoretic run by rinsing in the high-pressure mode (137 kPa) with 0.1 mol/L NaOH for 60 s and the separation buffer for 120 s. Samples were injected for 7 s in the low-pressure mode (3.5 kPa), corresponding to an injection volume of ~25 nL. This was followed by a second injection of separation buffer (for 5 s), to prevent loss of sample material. Electrophoresis was performed at 24 kV (358 V/cm) in the normal polarity mode (positive potential at capillary inlet). The capillary temperature was set at 29 [degrees]C. At the end of each electrophoretic run, the capillary was rinsed (137 kPa) with separation buffer for 60 s.

SAMPLE COLLECTION AND STORAGE

EDTA plasma was obtained by collecting blood into Vacutainer Tubes (Becton Dickinson), giving a final EDTA concentration of 4 mmol/L. The samples were immediately placed on ice and centrifuged (2000g for 10 min) within 60 min. The plasma was then processed further or stored at -80 [degrees]C until use. Sample handling was carried out under dim light to avoid photodegradation of the analytes.

SAMPLE PROCESSING

Plasma samples (500 [micro]L) were mixed with 1500 [micro]L of ice-cold 100 g/L TCA containing 15 nmol/L isoriboflavin, which served as an internal standard. Precipitated protein was removed by centrifugation, and 1500 [micro]L of the supernatant was neutralized by the addition of 480 [micro]L of 2 mol/L [K.sub.2]HP[O.sub.4]. Solid-phase extraction using Bond Elut C-18 columns (1 mL/50 mg; Varian) was then carried out automatically on a Gilson ASPEC sample processor. The columns were preconditioned with 500 [micro]L of methanol followed by 1000 [micro]L of 25 mmol/L sodium phosphate buffer, pH 7.0. The columns were then loaded with 1900 [micro]L of neutralized TCA-treated plasma and washed with the same phosphate buffer as used for column conditioning. The analytes were eluted with 200 [micro]L of 600 mL/L methanol in phosphate buffer, and the columns were then loaded with 220 [micro]L of phosphate buffer to reduce the methanol concentration in the pooled eluate, which was used for CE analysis. Between steps, the columns were dried by air pressurization.

CALCULATION OF PLASMA CONCENTRATIONS

Pooled EDTA plasma containing 7.2 nmol/L riboflavin, 6.1 nmol/L FMN, and 56.6 nmol/L FAD was supplemented with 20 nmol/L riboflavin, 20 nmol/L FMN, and 50 nmol/L FAD and used for calibration. The calibrator plasma was processed in the same way as the samples and was run approximately once for every five samples. Calculation of the vitamer concentrations was based on peak area. For each of the vitamers, the sample concentration was calculated using the formula:

[C.sub.s] = [C.sub.c] X [A.sub.s]/[A.sub.c] X [A.sub.IC]/[A.sub.IS] where [C.sub.S] is the vitamer concentration in the sample, [C.sub.c] is the vitamer concentration in the calibrator plasma, [A.sub.S] is the peak area of the vitamer in the sample, [A.sub.IC] is the peak area of the vitamer in the calibrator, [A.sub.IS] is the peak area of the internal standard in the calibrator, and AIs is the peak area of the internal standard in the sample.

In addition, we collected plasma in which we obtained the flavin concentration by repeated analysis against the calibrator. This plasma was stored at -80 [degrees]C and used for quality control.

STABILITY OF THE INTERNAL STANDARD AND THE RIBOFLAVIN VITAMERS

Because the internal standard was added to the TCA used for plasma protein precipitation, we investigated its stability in 100 g/L TCA. In addition, the stability of this compound was tested at all stages of sample preparation. The stability of riboflavin, FMN, and FAD was investigated in EDTA whole blood, hemolyzed blood, and EDTA plasma and at all stages of sample preparation.

In all stability experiments, samples were placed in the dark at 23 [degrees]C, and polypropylene tubes were used for sample processing and storage. To avoid interference from the interconversion of the vitamers, each of the analytes was added separately to aliquots from one donor. Isoriboflavin served as the reference calibrator for riboflavin, FMN, and FAD, and riboflavin was used as the reference calibrator for isoriboflavin. The reference calibrators were added before the elution step to correct for variations in solid-phase extraction and sample injection. Samples were analyzed in duplicate. Plasma concentrations were log-transformed and plotted against incubation time. Data were analyzed by simple linear regression, and the decomposition was given in terms of first-order rate constants.

To study the stability of the internal standard in TCA used for plasma protein precipitation, we studied aliquots of 100 g/L TCA containing 150 nmol/L isoriboflavin incubated for 1, 2, 4, 7, and 14 days. These aliquots were used for precipitating identical plasma samples. Riboflavin was added as a reference calibrator, and the samples were processed according to standard procedures.

We investigated the stability of endogenous riboflavin, FMN, and FAD in whole blood by incubating aliquots from one donor for 1, 2, and 5 h.

To study the stability of the riboflavin vitamers in plasma, we divided a sample containing endogenous vitamin concentrations into four batches, with 500 nmol/L riboflavin, FMN, or FAD added to three of the batches. Plasma aliquots were stored for 1, 4, 8, and 14 days before analysis.

We investigated the stability of the vitamers and the internal standard in TCA-treated plasma before neutralization by adding 500 nmol/L riboflavin, FMN, FAD, or isoriboflavin to the samples. The samples were then incubated for 1, 2, 4, 7, and 12 h before additional processing. The stability of the vitamers and the internal standard was also tested in neutralized TCA-treated plasma. Plasma samples to which 500 nmol/L riboflavin, FMN, FAD, or isoriboflavin had been added were analyzed after incubation for 1, 2, 4, and 7 days. In addition, the stability of the riboflavin vitamers and the internal standard was tested in the methanol-phosphate buffer obtained by solid-phase extraction using the same concentrations of riboflavin, FMN, FAD, or isoriboflavin and incubation for 1, 2, 4, and 7 days.

HEMOLYSIS

Hemolyzed EDTA blood was prepared by sonication (20 kHz for 5 s at 150 W). The hemoglobin concentration of the plasma was measured (Technicon H2E [TM] Technicon Instruments), and the percentage of hemolysis was calculated by comparing the hemoglobin concentration of the plasma to the total hemoglobin concentration of the blood. Sonicated and nonsonicated blood from the same donor was mixed in different proportions to obtain different degrees of hemolysis. Samples were incubated for 1, 2, and 5 h.

RECOVERY AND ANALYTICAL VARIATION

The recovery was determined by the addition of riboflavin (10 and 30 nmol/L), FMN (10 and 30 nmol/L), and FAD (50 and 100 nmol/L) to plasma containing endogenous analyte. At each of the three concentrations, 15 replicates were analyzed in one analytical run.

To determine within-day precision, we used the data from the recovery experiments. Between-day precision was determined by assaying the same samples on 15 different days over a period of 3 weeks.

LINEARITY AND LIMIT OF DETECTION

The linearity of the assay at vitamin concentrations greater than the endogenous concentrations was determined by the addition of increasing amounts of the vitamers, corresponding to plasma concentrations between 0.3 and 1000 nmol/L greater than the endogenous concentrations, to plasma containing endogenous concentrations of riboflavin, FMN, and FAD.

To determine the limit of detection and assay linearity at concentrations less than the endogenous concentrations, we used photodegradation to prepare plasma devoid of riboflavin. Plasma proteins were precipitated with 100 g/L TCA without internal standard and then exposed to ultraviolet irradiation at 370 nm for 6 h; neutralizing solution and internal standard were then added. Plasma containing endogenous concentrations of the analytes was not irradiated, but otherwise was prepared in the same way and mixed with irradiated plasma in different proportions ranging from 0% to 100% to obtain different concentrations of the vitamers.

[FIGURE 1 OMITTED]

Results

CAPILLARY ELECTROPHORESIS

Electropherograms of a sample containing endogenous concentrations of riboflavin vitamers; a sample with added riboflavin, FMN, lumiflavin, and FAD (2 nmol/L each); and a sample containing 25% of the endogenous concentrations are shown in Fig. 1. The analytes eluted in the order riboflavin, lumiflavin, FMN, and FAD and were baseline-resolved within 12 min and clearly separated from the internal standard. The migration profile was stable over time, and the within- and between-day relative standard deviations (RSD) of the migration times were 1.5% and 2.0%, respectively.

SAMPLE PROCESSING AND STABILITY

Stability of the internal standard and the riboflavin vitamers. The stability of the internal standard was investigated in 100 g/L TCA and at all stages of sample preparation. The stability of riboflavin, FMN, and FAD was tested in EDTA whole blood, hemolyzed blood, and EDTA plasma and at all stages of sample preparation. In all experiments, samples were incubated in the dark at 23 [degrees]C.

The internal standard was stable for 14 days when dissolved in the 100 g/L TCA solution used for protein precipitation.

The vitamers were stable for at least 5 h in whole blood and for at least 1 h in blood with 1% hemolysis. This degree of hemolysis did not affect plasma flavin concentrations.

Plasma containing endogenous vitamin concentrations or supplemented with 500 nmol/L riboflavin, FMN, or FAD could be stored for 14 days without changes in the concentrations of riboflavin or FMN. In nonsupplemented plasma, FAD was stable for 14 days, whereas the added FAD degraded at a rate of ~0.03/day.

Riboflavin, FMN, and the internal standard were stable for 12 h in TCA-treated plasma before neutralization, whereas FAD was degraded at a rate of ~0.17/h under these conditions. All vitamers and the internal standard were stable for at least 2 days in neutralized TCA-treated plasma and in the methanol-phosphate buffer obtained by solid-phase extraction.

[FIGURE 2 OMITTED]

[FIGURE 3 OMITTED]

ASSAY PERFORMANCE

Recovery and analytical variation. The recoveries of riboflavin, FMN, and FAD added to plasma at two concentrations were 90-103% (Table 1). The within-day and between-day CVs were 4-9% and 6-12%, respectively (Table 2).

Linearity and lower limit of detection. The linearity of the assay was tested at concentrations greater than and less than the endogenous vitamer concentrations (Fig. 2), which were 14.0 nmol/L for riboflavin, 7.5 nmol/L for FMN, and 85.5 nmol/L for FAD. The linearity was documented for concentrations up to 200 nmol/L greater than the endogenous concentrations for all vitamers. At concentrations lower than the endogenous concentrations, linearity was observed down to concentrations approaching the limit of detection, which was defined as a signal-to-noise ratio >5:1.

APPLICATION OF THE METHOD

Vitamer concentrations in healthy adults. Riboflavin, FMN, and FAD were measured in plasma from 63 healthy volunteers: 35 men and 28 women. Thirty-six subjects were blood donors, and 27 subjects were recruited from laboratory staff. The mean age was 42 years (range, 23-64 years).

The median concentration (5-95 percentiles) was 8.6 nmol/L (2.7-42.5 nmol/L) for riboflavin, 7.0 nmol/L (3.5-13.3 nmol/L) for FMN, and 57.9 nmol/L (44.5-78.1 nmol/L) for FAD (Fig. 3). Accordingly, the interindividual variation was substantial for riboflavin, whereas it was lower for FMN and, in particular, for FAD. The vitamin concentrations were not related to age (P >0.26, Spearman correlation) or gender (P >0.11, Mest). There was a significant positive correlation between the concentrations of riboflavin and FMN (r = 0.72; P <0.0001) and between FMN and FAD (r = 0.37; P <0.003), whereas the correlation between riboflavin and FAD (r = 0.13; P = 0.3) was not significant.

Discussion

To our knowledge, this is the first CE-LIF method for the determination of riboflavin in human plasma. Its main advantages when compared with most HPLC methods (11-18) are high sensitivity, validation of analyte stability before and during sample processing, and quantification of all vitamers. Lumiflavin, which is a known photodegradation product of riboflavin (2), is clearly separated from the other analytes.

High sensitivity is obtained by exploiting the native fluorescence of riboflavin and its derivatives, the excitation spectra of which overlap the emission wavelength of 488 nm of the argon laser (21). The lower sensitivity for FAD (Figs. 1 and 2) is related to its lower fluorescence yield compared with riboflavin and FMN (21). In its present format, the method allows the detection of vitamin concentrations far below the reference values obtained by us (Fig. 2), and therefore should be suitable for the assessment of riboflavin status under deficiency states. Sensitivity enhancement might be obtained by the use of a HeCd laser emitting at 442 nm, which is closer to the excitation maximum of the vitamers. Furthermore, only ~25 nL of the 420-[micro]L eluate from the solid-phase columns is injected into the capillary, and the mass sensitivity could be enhanced by concentrating the eluate before the CE step.

A major portion of the riboflavin vitamers is protein bound (22,23), and TCA treatment of plasma offers protein precipitation and probably extraction of the protein-bound vitamin fraction. The high ionic strength obtained by this procedure is incompatible with CE, but solid-phase extraction provides a means for sample desalting and for concentration of analytes before CE. We also evaluated organic solvents, e.g., acetonitrile, as protein precipitating agents, but this matrix prevented the retention of analytes on the reversed-phase columns.

Isoriboflavin is an isomeric form of riboflavin (8-demethyl-6-methylriboflavin) and has been used previously as an internal standard in HPLC assays (15). We did not detect endogenous isoriboflavin in human plasma. Isoriboflavin is stable during sample processing and coelutes with the analytes during solid-phase extraction. Inclusion of the internal standard corrects for variable recovery during solid-phase extraction, differences in the amount of sample injected into the capillary, and long-term changes in laser output, and it reduces the within-and between-day CVs from 9-18% (data not shown) to 4-12% (Table 2).

We investigated the stability of the vitamers and the internal standard at different stages of sample processing because analyte degradation would seriously impair assay stability. In neutralized TCA-treated plasma, analytes were stable for at least 2 days at room temperature. This may partly explain the high analytical recovery, which was close to 100% (Table 1), and acceptable within- and between-day CVs (Table 2).

We also demonstrated that the riboflavin vitamers were stable for at least 5 h in whole blood and for several days in plasma at room temperature. This is in agreement with data published by Burch et al. (24), who found that the vitamers were stable in whole blood stored at 4 [degrees]C for 48 h. They also reported that endogenous FAD was stable in serum stored at room temperature for several hours, whereas 80% of FAD added to serum was hydrolyzed within 1 h. Others have found that FAD added to whole blood (25,26) or plasma (16,26) is highly unstable and is degraded within minutes. FMN added to whole blood was found to be decomposed within minutes by Nogami et al. (25), but more slowly by other investigators (26,27); it appears to be relatively stable in plasma (16, 22, 26).

The apparent discrepancies between our stability data and those published by others (16,22,24-27) can be attributed in part to different incubation temperatures or anticoagulants. Notably, we used EDTA, which has been shown to inhibit the activity of enzymes that hydrolyze FAD and FMN (28). The binding of the vitamers to proteins (22,23) may protect these compounds from degradation, which explains the different stabilities of endogenous vs added vitamers in plasma.

Hemolysis may influence analyte concentrations in plasma, either by the release of flavins from blood cells or by the leakage of cellular enzymes that catalyze the conversion of the vitamers. We found that hemolysis up to 1% does not represent an analytical problem. This percentage of hemolysis is seldom encountered in the clinical setting, and it is higher than the routinely used percentage for interference testing of analytical methods (29).

Our data on analyte stability are the basis for prolonged, unattended sample processing and automated CE injection. In addition, the stability of these analytes allows the measurement of vitamins in clinical samples transported or stored for days as well as epidemiological studies based on frozen samples. We have analyzed plasma samples kept at -80 [degrees]C for 32 months. In these samples, the vitamer concentrations, including FAD, were similar to those observed in fresh samples (data not shown).

We found vitamin concentrations similar to those reported by others (11,15, 30, 31). We also observed a low interindividual variation of FAD compared with variable amounts of riboflavin and FMN. Riboflavin, in particular, varied considerably between subjects (15, 31). This may indicate that the concentration of FAD is tightly regulated (24,32-34), whereas plasma riboflavin may vary in response to recent vitamin intake (31). Thus, the different flavin species in plasma appear to be under separate regulatory or nutritional influence, and the measurement of all vitamers in plasma may be desirable. The measurement of all vitamers may also be important in the investigation of conditions under which the activities of the enzymes that catalyze the interconversion of the different vitamers are changed, e.g., in riboflavin deficiency (34, 35) or thyroid disease (36, 37).

The widely used erythrocyte glutathione reductase activity coefficient is an indirect measure of the FAD concentration in the erythrocytes (7, 8). It does not, however, determine the concentrations of riboflavin and FMN or the relative distribution of different flavin species. It is considered a sensitive and robust index of riboflavin deficiency, but is less suitable for the assessment of riboflavin status at high riboflavin intake (6). Furthermore, the erythrocyte glutathione reductase activity coefficient may give misleading results for certain conditions in which erythrocyte flavin metabolism is altered, e.g., glucose-6-phosphate dehydrogenase deficiency (6, 7) and [beta]-thalassemia (38).

In conclusion, we have constructed a robust CE method for the determination of riboflavin species in human plasma. The monitoring of native fluorescence with LIF detection affords sufficiently high sensitivity to measure vitamin concentrations far lower than the concentrations found in healthy subjects. The stability of the vitamers during sample processing and an almost complete recovery allows precise quantification of all three vitamers. The method thus may become a valuable tool to assess riboflavin status in humans.

This work was funded by grants from the Norwegian Research Council. S.H. is a fellow of the Norwegian Research Council. We thank Ove Netland and Gry Kvalheim for technical assistance.

Received December 8, 1998; accepted March 10, 1999.

References

(1.) Cooperman JM, Lopez R. Riboflavin. In: Machlin LJ, ed. Handbook of vitamins. New York: Marcel Dekker, 1991:283-310.

(2.) Rivlin RS. Riboflavin. In: Ziegler EE, Filer LJ Jr, eds. Present knowledge in nutrition. Washington, DC: ILSI Press, 1996:167-73.

(3.) Boisvert WA, Castaneda C, Mendoza I, Langeloh G, Solomons NW, Gershoff SN, et al. Prevalence of riboflavin deficiency among Guatemalan elderly people and its relationship to milk intake. Am J Clin Nutr 1993;58:85-90.

(4.) Benton D, Haller J, Fordy J. The vitamin status of young British adults. Int J Vitam Nutr Res 1997;67:34-40.

(5.) Madigan SM, Tracey F, McNulty H, Eaton-Evans J, Coulter J, McCartney H, et al. Riboflavin and vitamin B-6 intakes and status and biochemical response to riboflavin supplementation in free-living elderly people. Am J Clin Nutr 1998;68:389-95.

(6.) Bates CJ, Thurnham DI, Bingham SA, Margetts BM, Nelson M. Biochemical markers of nutrient intake. In: Margetts BM, Nelson M, eds. Design concepts in nutritional epidemiology. Oxford: Oxford University Press, 1991:192-265.

(7.) Beutler E. Effect of flavin compounds on glutathione reductase activity: in vivo and in vitro studies. J Clin Investig 1969;48:1957-66.

(8.) Glatzle D, Komer WF, Christeller S, Wiss 0. Method for the detection of a biochemical riboflavin deficiency. Stimulation of NADPH2-dependent glutathione reductase from human erythrocytes by FAD in vitro. Investigations on the vitamin [B.sub.2] status in healthy people and geriatric patients. Int Z Vitaminforsch 1970; 40:166-83.

(9.) Baker H, Frank O. Analysis of riboflavin and its derivatives in biological fluids and tissues. In: Rivlin RS, ed. Riboflavin. New York: Plenum Press, 1975:49-79.

(10.) Nielsen P. Flavins. In: De Leenheer AP, Lambert WE, Nelis HJ, eds. Modern chromatographic analysis of vitamins. New York: Marcel Dekker, 1992:355-98.

(11.) Ohkawa H. A simple method for micro-determination of flavins in human serum and whole blood by high-performance liquid chromatography. Biochem Int 1982;4:187-94.

(12.) Speek AJ, van Schaik F, Schrijver J, Schreurs WH. Determination of the [B.sub.2] vitamer flavin adenine dinucleotide in whole blood by high-performance liquid chromatography with fluorometric detection. J Chromatogr 1982;228:311-6.

(13.) Floridi A, Palmerini CA, Fini C, Pupita M, Fidanza F. High performance liquid chromatographic analysis of flavin adenine dinucleotide in whole blood. Int J Vitam Nutr Res 1985;55:187-91.

(14.) Botticher B, Botticher D. A new HPLC-method for the simultaneous determination of [B.sub.1]-, [B.sub.2] and [B.sub.6] vitamers in serum and whole blood. Int J Vitam Nutr Res 1987;57:273-8.

(15.) Lambert WE, Cammaert PM, De-Leenheer AP. Liquid-chromatographic measurement of riboflavin in serum and urine with isoriboflavin as internal standard. Clin Chem 1985;31:1371-3.

(16.) Pietta P, Calatroni A, Rava A. Hydrolysis of riboflavin nucleotides in plasma monitored by high-performance liquid chromatography. J Chromatogr 1982;229:445-9.

(17.) Lopez-Anaya A, Mayersohn M. Quantification of riboflavin, riboflavin 5'-phosphate and flavin adenine dinucleotide in plasma and urine by high-performance liquid chromatography. J Chromatogr 1987;423:105-13.

(18.) Zempleni J. Determination of riboflavin and flavocoenzymes in human blood plasma by high-performance liquid chromatography. Ann Nutr Metab 1995;39:224-6.

(19.) Landers JP. Clinical capillary electrophoresis [Review]. Clin Chem 1995;41:495-509.

(20.) Kemp G. Capillary electrophoresis: a versatile family of analytical techniques [Review]. Biotechnol Appl Biochem 1998;27:9-17.

(21.) Weimar WR, Neims AH. Physical and chemical properties of flavins; binding of flavins to protein and conformational effects; biosynthesis of riboflavin. In: Rivlin RS, ed. Riboflavin. New York: Plenum Press, 1975:1-47.

(22.) Jusko WJ, Levy G. Plasma protein binding of riboflavin and riboflavin-5'-phosphate in man. J Pharm Sci 1969;58:58-62.

(23.) White HB III, Merrill AH Jr. Riboflavin-binding proteins [Review]. Annu Rev Nutr 1988;8:279-99.

(24.) Burch HB, Bessey OA, Lowry OH. Fluorometric measurements of riboflavin and its natural derivatives in small quantities of blood serum and cells. J Biol Chem 1948;175:457-70.

(25.) Nogami H, Hanano M, Awazu S, Iga T. Pharmacokinetic aspects of biliary excretion. Dose dependency of riboflavin in rat. Chem Pharm Bull Tokyo 1970;18:228-34.

(26.) Okumura M, Yagi K. Hydrolysis of flavin adenine dinucleotide and flavin mononucleotide by rabbit blood. J Nutr Sci Vitaminol Tokyo 1980;26:231-6.

(27.) Jusko WJ, Levy G, Yaffe SJ, Gorodischer R. Effect of probenecid on renal clearance of riboflavin in man. J Pharm Sci 1970;59:473-7.

(28.) Akiyama T, Selhub J, Rosenberg IH. FMN phosphatase and FAD pyrophosphatase in rat intestinal brush borders: role in intestinal absorption of dietary riboflavin. J Nutr 1982;112:263-8.

(29.) Koch DD, Peters T Jr. Selection and evaluation of methods. In: Burtis CA, Ashwood ER, eds. Tietz textbook of clinical chemistry. Philadelphia: WB Saunders, 1994:508-25.

(30.) Kodentsova VM, Vrzhesinskaya OA, Spirichev VB. Fluorometric riboflavin titration in plasma by riboflavin-binding apoprotein as a method for vitamin BZ status assessment. Ann Nutr Metab 1995;39:355-60.

(31.) Zempleni J, Galloway JR, McCormick DB. Pharmacokinetics of orally and intravenously administered riboflavin in healthy humans. Am J Clin Nutr 1996;63:54-66.

(32.) Burch HB, Lowry OH, Padilla AM, Combs AM. Effects of riboflavin deficiency and realimentation on flavin enzymes in tissues. J Biol Chem 1956;223:29-45.

(33.) Bessey OA, Horwitt MK, Love RH. Dietary deprivation of riboflavin and blood riboflavin levels in man. J Nutr 1956;58:367-83.

(34.) Fass S, Rivlin RS. Regulation of riboflavin-metabolizing enzymes in riboflavin deficiency. Am J Physiol 1969;217:988-91.

(35.) Ross NS, Hansen TP. Riboflavin deficiency is associated with selective preservation of critical flavoenzyme-dependent metabolic pathways [Review]. Biofactors 1992;3:185-90.

(36.) Rivlin RS. Hormones, drugs and riboflavin [Review]. Nutr Rev 1979; 37:241-5.

(37.) Lee SS, McCormick DB. Thyroid hormone regulation of flavocoenzyme biosynthesis. Arch Biochem Biophys 1985;237:197-201.

(38.) Anderson BB, Giuberti M, Perry GM, Salsini G, Casadio I, Vullo C. Low red blood cell glutathione reductase and pyridoxine phosphate oxidase activities not related to dietary riboflavin: selection by malaria? Am J Clin Nutr 1993;57:666-72.

STEINAR HUSTAD,* PER MAGNE UELAND, and JORN SCHNEEDE

Department of Pharmacology, University of Bergen, Armauer Hansens Hus, 5021 Bergen, Norway.

* Author for correspondence. Fax 47-55-974605; e-mail steinar.hustad@farm.uib.no.

[1] Nonstandard abbreviations: FMN, flavin mononucleotide; CE, capillary electrophoresis; LIF, laser-induced fluorescence; and TCA, trichloroacetic acid.
Table 1. Assay recovery.

 Concentration, (a) nmol/L

 Added Expected

 Endogenous Low High Low High

Riboflavin 6.9 10 30.0 16.9 36.9
FMN 8.8 10 30.0 18.8 38.8
FAD 64.0 50 100.0 114.0 164.0

 Concentration, (a) nmol/L % recovery (b)

 Measured

 Low High Low High

Riboflavin 16.8 36.3 99 (8) 98 (6)
FMN 18.7 39.6 99 (4) 103 (9)
FAD 109.1 163.2 90 (5) 99 (6)

(a) n = 15 for all concentrations.

(b) Data are given as means with SD in parentheses.

Table 2. Assay precision.

 Within-day (n = 15) Between-day (n = 15)

 Mean [+ or -] SD, CV, % Mean [+ or -] SD, CV, %
 nmol/L nmol/L

Riboflavin 6.9 [+ or -] 0.6 9 6.6 [+ or -] 0.8 12
 16.8 [+ or -] 1.3 8 16.6 [+ or -] 1.9 11
 36.3 [+ or -] 2.1 6 36.7 [+ or -] 3.4 9

FMN 8.8 [+ or -] 0.4 5 8.3 [+ or -] 0.8 9
 18.7 [+ or -] 0.7 4 17.9 [+ or -] 1.7 9
 39.6 [+ or -] 3.6 9 39.3 [+ or -] 3.7 9

FAD 64.0 [+ or -] 4.4 7 60.6 [+ or -] 4.7 8
 108.7 [+ or -] 5.1 5 109.1 [+ or -] 9.3 8
 163.2 [+ or -] 10.5 6 163.4 [+ or -] 10.4 6
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Title Annotation:Automation and Analytical Techniques
Author:Hustad, Steinar; Ueland, Per Magne; Schneede, Jorn
Publication:Clinical Chemistry
Date:Jun 1, 1999
Words:4915
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