Metabolic profiling of major vitamin D metabolites using Diels–Alder derivatization and ultra-performance liquid chromatography–tandem mass spectrometry
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- Aronov, P.A., Hall, L.M., Dettmer, K. et al. Anal Bioanal Chem (2008) 391: 1917. doi:10.1007/s00216-008-2095-8
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Biologically active forms of vitamin D are important analytical targets in both research and clinical practice. The current technology is such that each of the vitamin D metabolites is usually analyzed by individual assay. However, current LC-MS technologies allow the simultaneous metabolic profiling of entire biochemical pathways. The impediment to the metabolic profiling of vitamin D metabolites is the low level of 1α,25-dihydroxyvitamin D3 in human serum (15–60 pg/mL). Here, we demonstrate that liquid–liquid or solid-phase extraction of vitamin D metabolites in combination with Diels–Alder derivatization with the commercially available reagent 4-phenyl-1,2,4-triazoline-3,5-dione (PTAD) followed by ultra-performance liquid chromatography (UPLC)–electrospray/tandem mass spectrometry analysis provides rapid and simultaneous quantification of 1α,25-dihydroxyvitamin D3, 1α,25-dihydroxyvitamin D2, 24R,25-dihydroxyvitamin D3, 25-hydroxyvitamin D3 and 25-hydroxyvitamin D2 in 0.5 mL human serum at a lower limit of quantification of 25 pg/mL. Precision ranged from 1.6–4.8 % and 5–16 % for 25-hydroxyvitamin D3 and 1α,25-dihydroxyvitamin D3, respectively, using solid-phase extraction.
Keywords1α,25-Dihydroxyvitamin D325-Hydroxyvitamin D324R,25-Dihydroxyvitamin D3UPLCLC-MSMetabolic profilingDerivatization
The development of an LC-MS profiling method for vitamin D metabolites is impeded by their low concentration in human circulation, particularly for 1α,25(OH)2D3, with concentrations ranging from 15 to 60 pg/mL. Recently, an LC-tandem MS method was introduced to measure nonderivatized 1α,25(OH)2D3, but this requires 2 mL of human serum . Vitamin D metabolites have low ionization efficiencies in electrospray (ESI) or atmospheric pressure chemical ionization (APCI) sources because they lack easily charged groups, which would enhance ionization efficiencies. However, the conjugated diene group of vitamin D metabolites makes them a specific target for Diels–Alder derivatization. In fact, several highly reactive 4-substituted 1,2,4-triazoline-3,5-diones (TADs or Cookson-type reagents) have been reported in the literature for the analysis of vitamin D metabolites, their analogs and other dienes, including derivatization reagents for ESI-MS [6, 17–22]. The derivatization reagents introduce polar groups and thus typically result in a 100–1000-fold increase in sensitivity over nonderivatized compounds. However, no method for profiling the major vitamin D metabolites 1α,25(OH)2D2, 1α,25(OH)2D3, 24R,25(OH)2D3, 25(OH)D2 and 25(OH)D3 has been reported that can detect and quantify endogenous levels of 1α,25(OH)2D3. Here, we demonstrate an ultra-performance liquid chromatography (UPLC)–tandem MS method for the quantification of an array of the most biologically important vitamin D metabolites after Diels–Alder derivatization with 4-phenyl-1,2,4-triazoline-3,5-dione (PTAD). The method was validated for solid-phase extraction (SPE) and liquid–liquid extraction (LLE) of 1α,25(OH)2D3 and 25(OH)D3 from serum and applied to studies of vitamin D metabolism in humans.
Hexane, methyl tert-butyl ether, dichloromethane, acetonitrile, ethyl acetate, methanol, formic acid, and K2HPO4 were purchased from Fisher Scientific (Pittsburgh, PA, USA). Deionized water (resistivity of 18.1 MΩ/cm) and distilled water were prepared in-house and used for mobile phase preparation and SPE extraction, respectively. PTAD was purchased from Fluka (St. Louis, MO, USA). Standards of vitamin D metabolites were purchased from Fluka, Sigma–Aldrich (St. Louis, MO, USA) and BIOMOL (Plymouth Meeting, PA, USA) as indicated below. Deuterated surrogates of vitamin D metabolites were purchased from Synthetica (Oslo, Norway) and Medical Isotopes (Pelham, NH, USA). Human serum from healthy male donors for method development purposes was ordered from Fisher Scientific (Pittsburgh, PA, USA).
Stock and calibration solutions
Neat standards of vitamin D metabolites were diluted and stored at −80 °C; 10 μg 1α,25-dihydroxyvitamin D3 (Sigma–Aldrich) was dissolved in 1000 μL acetonitrile in the original vial and then transferred into an amber glass HPLC vial; 1 mg 1α,25-dihydroxyvitamin D2 (Fluka) was dissolved in 500 μL acetonitrile; 1 mg 25-hydroxyvitamin D3 (Fluka) was dissolved in 500 μL acetonitrile; 1 mg 25-hydroxyvitamin D2 (Fluka) was dissolved in 1000 μL acetonitrile and transferred into an amber glass HPLC vial; 50 μg 24R,25-dihydroxyvitamin D3 (BIOMOL) in 50 μL ethanol was diluted with 950 μL acetonitrile; 1 mg calcipotriol (Sequoia Research Products, Pangbourne, UK) was dissolved in 2000 μL acetonitrile. Calcipotriol was further diluted with acetonitrile to make a 125 ng/mL derivatization quality control spike. The deuterated internal standards (IS) 26,26,26,27,27,27-hexadeuterium-1α,25-dihydroxyvitamin D3 (Medical Isotopes) and 26,26,26,27,27,27-hexadeuterium-25-hydroxyvitamin D3 (Synthetica) were dissolved in acetonitrile to prepare stock solutions. The stock solutions were combined and diluted to obtain a 12.5 ng/mL d6 1α,25(OH)2D3 and 500 ng/mL d6 25(OH)D3 IS solution. The purity of the derivatized IS was assessed by LC-MS (full scan and MRM) up to 500 ng/mL and no interferences were found, including no traces of natural metabolites. All analytes were individually dissolved in a solution of PTAD (0.5 mg/mL) in acetonitrile at 100 ng/mL and allowed to react at room temperature for 4 h to form the corresponding PTAD Diels–Alder conjugate. No nonderivatized analytes were found in these solutions as analyzed by LC-MS (MRM). To prevent cross-contamination of the 1α,25(OH)2D-PTAD calibration solution, it was prepared separately from the rest of the analytes. Using serial dilutions, 0.1, 0.3, 1.0, 3.0, and 10.0 ng/mL calibration stocks of 1α,25(OH)2D3-PTAD and 1α,25(OH)2D2-PTAD were prepared. Similarly, we prepared calibration solutions of 25(OH)D2-PTAD and 25(OH)D3-PTAD at the levels of 0.1, 0.3, 1.0, 3.0, 10.0, 30.0, 100, 300 and 1000 ng/mL. These calibration solutions also contained 24R,25(OH)2D3-PTAD at tenfold lower levels than 25(OH)D2-PTAD and 25(OH)D3-PTAD; however, 0.01 and 0.03 ng/mL levels of 24R,25(OH)2D3 were not used in practice.
Plasma samples for the HIV study were obtained from the Reaching for Excellence in Adolescent Care and Health (REACH) repository [23, 24]. Serum samples from the sun exposure study were acquired from 17 volunteers in fall 2006 and 17 different volunteers in winter 2007 at week 0, week 4, and week 7 (week 8 in winter). Two 10-mL tubes of blood were collected from each participant after a four-hour fast from fat. The tubes were wrapped in foil and allowed to clot at room temperature for one hour. The tubes were centrifuged and then aliquoted in ten 1-mL tubes. These tubes were placed in light-proof boxes and kept frozen at −80 °C. Commercial human serum from male donors (Fisher Scientific) was used for method development and validation except in the extraction reproducibility study, where donor plasma was used.
Sample preparation was adapted from published methods . Briefly, 500 μL aliquotes of human serum in 2 mL plastic tubes (Fisher Scientific) were spiked with 20 μL solution of internal standard (12.5 ng/mL d6 1α,25-dihydroxyvitamin D3 and 500 ng/mL d6 25-hydroxyvitamin D3) in acetonitrile and allowed to equilibrate for 15 min at room temperature. Proteins were precipitated by the addition of 500 μL acetonitrile and by spinning the sample on a Vortex mixer for 1 min at maximum speed followed by 10 min centrifugation at 10,000×g.
The supernatant from protein precipitation was transferred into 2-mL plastic tubes (Fisher Scientific) containing 400 μL 0.4 M K2HPO4 and mixed using a Vortex mixer for 30 s followed by the addition of 500 μL methyl t-butyl ether (MTBE). The tubes were vigorously mixed for 2 min on a Vortex mixer, centrifuged for 5 min at 10,000×g, and the upper organic layer was transferred into 2-mL plastic tubes (Fisher Scientific). For method development, the organic extracts were spiked with 10 μL 25 ng/mL calcipotriol used as a control for derivatization efficiency. Samples were evaporated for 1 h using an RC10.22 vacuum concentrator (Jouan, Winchester, VA, USA), and 100 μL 0.75 mg/mL PTAD in acetonitrile was added to the residue followed by 1 min of mixing. Samples were left at room temperature for 1 h and then stored overnight at +4 °C to allow the derivatization reaction to proceed to completion; then the samples were mixed for 30 s, centrifuged and transferred into 150-μL vial inserts.
Solid-phase extraction of vitamin D metabolites using Oasis HLB (Waters, Milford, MA, USA) sorbent was adopted from published methods [22, 25]. Oasis HLB cartridges (3 cc 60 mg) were preconditioned with 3 mL ethyl acetate, 3 mL methanol and 3 mL water. Individual valves of cartridges were closed after the water meniscus reached the sorbent surface. Cartridges were loaded with 900 μL supernatant from the protein precipitation protocol and 1 mL 0.4 M K2HPO4. The valves were opened and samples were extracted using gravity only. Cartridges were subsequently washed with 3 mL water and 2 mL of 70% methanol and dried for 2 min by application of negative pressure. The needles of the extraction manifold were wiped to remove residual solvent droplets. Samples were eluted with 1.5 mL of acetonitrile into 2 mL plastic tubes. For method development the organic extracts were spiked with 10 μL 25 ng/mL calcipotriol used as a control for derivatization efficiency. Samples were evaporated for 2.5 h using a vacuum concentrator (RC10.22) and 100 μL 0.75 mg/mL PTAD in acetonitrile was added to the residue followed by 1 min of vigorous mixing. Samples were left at room temperature for 1 h, stored overnight at +4 °C for complete derivatization, vigorously mixed for 30 s, centrifuged and transferred into 150-μL vial inserts.
HPLC and tandem MS conditions
Mass spectrometry conditions
Cone voltage (V)
Precursor ion (m/z)
Collision energy (V)
Product ion (m/z)
d6 1 α,25(OH)2D3
d6 D3 (cholecalciferol)
Standard addition experiment
For LLE standard addition experiments 400 μL pooled human serum was aliquoted into 2 mL plastic tubes and each group (n = 4) was spiked with 10 μL blank, 1.0, 3.0 and 10.0 ng/mL 1α,25(OH)2D3 as well as 125, 250 or 500 ng/mL 25(OH)D3. Samples were extracted and measured independently as described above. For SPE standard addition experiments 500 μL pooled human serum was aliquoted into 2 mL plastic tubes and each group (n = 5) was spiked with 10 μL blank, spike 1 (0.75 ng/mL 1α,25(OH)2D3, 10.0 ng/mL 24R,25(OH)2D3, 100 ng/mL 25(OH)D3 and 100 ng/mL 25(OH)D2), spike 2 (twofold spike 1) and spike 3 (fourfold spike 1). Samples were extracted and measured independently as described above.
Quantification and data analysis
Calibration curve equation
y = 295.1x + 4.2
y = 300.3x + 3.0 (area) y = 3.46x + 121.76 (height)b
0.9999 (area) 0.9985 (height) b
y = 195.2x – 24.70
y = 421.1x + 199.2
y = 137.1x + 52.4
25(OH)D in serum was measured in the Bioanalytical Support Laboratory of the Western Human Nutrition Research Center (WHNRC) using a standard RIA protocol (DiaSorin, Stillwater, MN, USA) according to the manufacturer’s instructions with the following modification: the centrifugation following the precipitation step was performed at 3000×g for 60 min at +10 °C instead of the recommended 1800×g for 20 min at +20–25 °C. This modified procedure facilitated the aspiration of the supernatant from above the pellet containing the labeled 25(OH)D. The WHNRC participates in the DEQAS Vitamin D External Quality Assessment Scheme (http://www.deqas.org/)  and calibration standards from DEQAS analyzed during this period were all within acceptable limits.
Results and discussion
Derivatization reaction and product stability
TADs are among the most reactive dienophiles known. However, they are unstable in protic solvents. To determine derivatization rates, 1α,25(OH)2D3 and 25(OH)D3 at concentrations of 10 ng/mL (26 nM) and 10 μg/mL (25 μM), respectively, were allowed to react with 0.75 mg/mL (4.3 mM) PTAD at room temperature. Aliquots of the reaction mixtures were taken at fixed time intervals and quenched with equal volumes of methanol. The regression analysis according to a pseudo-first-order kinetics model resulted in t1/2 < 1 min for 25(OH)D3 and t1/2 ∼8 min for 1α,25(OH)2D3. Because the deuterium label is distant from the Diels–Alder reaction site, no isotope effects on the reaction kinetics are expected for isotopically labeled 1α,25(OH)2D3 and 25(OH)D3. Thus, >99% yield of derivatization products is achieved after one hour at room temperature, as predicted by the kinetic study. An increase in the PTAD concentration to over 2 mg/mL leads to a significant decrease in yield.
LC and MS optimization
Derivatization of vitamin D metabolites with PTAD produces two epimers, 6S and 6R, corresponding to the position of the dienophile relative to the plane of the A ring. The major isomer peak was used for integration and quantification. We compared results for 25(OH)D3 and 24R,25(OH)2D3 quantification in 50 individual serum samples using integration of either the major or minor peak and found a good correlation between the values (R2 = 0.9669 and R2 = 0.8838, respectively). The lower correlation for 24R,25(OH)2D3 can be explained by the low intensity of the minor isomer peak and the lack of a corresponding isotopically labeled standard to correct the measurements. Interestingly, the C18 BEH UPLC stationary phase does not separate isomers of derivatized 1α,25(OH)2D3, probably due to the anti position of hydroxyl groups on its A ring, which make this structure more symmetric. Separation of derivatized 1α,25(OH)2D3 can be achieved using phenyl BEH column chemistry, but this separation is not advantageous for quantitative purposes. While the C18 BEH phase does not separate isomers of 1α,25(OH)2D3-PTAD, it does separate the isomers of 1α,25(OH)2D2-PTAD (see the “Electronic supplementary material”).
In addition to cycloaddition to the locked C-10-19 : C-5-6 cisoid diene, the Diels–Alder reaction can theoretically occur at the C-5-6 : C-7-8 diene if the C-6–C-7 bond rotates into a cisoid conformation. However, this reaction would be unfavorable because of the activation barrier to uncoupling the conjugated triene system and the steric hindrance to forming a planar diene. PTAD is not only a potent dienophile but also a mild oxidizing reagent. Therefore, other possible by-products of derivatization can be form because of the oxidation of secondary alcohols of vitamin D metabolites into corresponding ketones. We surveyed mass chromatograms of derivatized standards and did not find an abundant signal (>1% peak height of derivatized standard) that would correspond to keto- ([M−2+H]+) and diketo- ([M−4+H]+) by-products.
Recoveries of internal standards and reproducibilities for SPE and LLE extraction methods
Calcipotriol derivatization yield (%)
83.3 ± 9.6 (n = 50)
105 ± 11 (n = 25)
d6 1α,25(OH)2D3recovery (%)
70.5 ± 6.7 (n = 50)
85.8 ± 8.8 (n = 25)
d6 25(OH)D3recovery (%)
78.1 ± 4.7 (n = 50)
81.0 ± 8.0 (n = 25)
0.5 ± 0.1 (n = 3)
0.6 ± 0.1 (n = 3)
26.8 ± 0.9 (n = 3)
29.9 ± 2.5 (n = 3)
36 ± 3 (n = 3)
41 ± 9 (n = 3)
1.2 ± 0.2 (n = 3)
1.8 ± 0.1 (n = 3)
In addition, SPE results in more precise measurements than LLE. Also, the throughput of SPE sample preparation is practically twice as fast as LLE. Thus, we suggest that SPE is the preferred technique for samples in clinical settings where precise measurements of 1α,25(OH)2D2, 1α,25(OH)2D3, 25(OH)D2 and 25(OH)D3 are critical. However, the LLE procedure is suitable for further explorations in vitamin D metabolic profiling, such as the addition of 1α,24R,25(OH)3D3 to the method, as well as the analysis of biohazardous human samples (e.g., HIV or hepatitis positive) that would otherwise require a special SPE extraction manifold.
Reverse-phase SPE is known for its very high retention of vitamin D2 and D3, which makes its application problematic. Only ∼30% of vitamin D2 and D3 are eluted from SPE cartridges under the conditions selected for our method (1.5 mL acetonitrile), as monitored by UPLC-UV. Using a stronger solvent such as ethyl acetate would decrease the purity of the sample and the yield of the derivatization reaction. Surprisingly, we found that LLE also results in poor recoveries of vitamin D2 and D3. The addition of vitamin D2 and D3 standards to extracted serum matrix prior to derivatization revealed that the reason for the poor recoveries is a low derivatization reaction yield. However, poor recoveries were not obtained for other more polar forms of vitamin D. Taking into consideration the severe matrix effect present during the derivatization of vitamin D2 and D3 and the need for longer chromatographic runs, we omitted these analytes from the method.
Standard addition experiment using liquid–liquid extraction
25(OH)D3 standard addition
25(OH)D3 measured (ng/mL)
Native serum (n = 4)
Native serum + 1.25 ngb 25(OH)D3 (n = 4)
26.1 (102. ± 2%)
Native serum + 2.5 ng 25(OH)D3 (n = 4)
28.4 (98.9 ± 4.8%)
Native serum + 5 ng 25(OH)D3 (n = 4)
34.3 (98.0 ± 3.3%)
1α,25(OH)2D3 standard addition
1α,25(OH)2D3 measured (pg/mL)
Native serum (n = 3)
Native serum + 10 pgb 1α,25(OH)2D3 (n = 4)
86 (103 ± 16%)
Native serum + 30 pg 1α,25(OH)2D3 (n = 4)
160 (120 ± 20%)
Native serum + 100 pg 1α,25(OH)2D3 (n = 3)
343 (111 ± 5%)
Standard addition experiment using SPE
3.03 (99.9 ± 11.1%)
3.08 (95.2 ± 6.5%)
3.19 (88.0 ± 5.4%)
31 (94 ± 16%)
52 (108 ± 16%)
90 (116 ± 7%)
17.4 (95.6 ± 1.9%)
18.7 (92.4 ± 2.2%)
21.5 (88.8 ± 4.0%)
2.3 (101.4 ± 6.1%)
4.3 (100.6 ± 3.9%)
8.3 (99.8 ± 8.6%)
The lowest calibration point (100 pg/mL) for all analytes produced chromatographic peaks with RMS signal to noise > 10. Because vitamin D metabolites are endogenous compounds, we studied method accuracy and precision using the addition of known quantities of vitamin D metabolites to human serum and extracted it by LLE and SPE. Separate standard addition experiments were performed for samples extracted by LLE (Table 4) and SPE (Table 5). For LLE, separate experiments for 1α,25(OH)2D3 and 25(OH)D3 standard spikes were performed, while for SPE, serum was spiked with a mixture of 24R,25(OH)2D3, 1α,25(OH)2D3, 25(OH)D2 and 25(OH)D3. The differences in metabolite concentrations in native serum between LLE and SPE experiments are due to the different batches of pooled human serum used in the experiments. We set quality control criteria for all extractions as precision (RSD) <20% and accuracy (analyte recovery corrected to the recovery of a corresponding deuterated internal standard) in the range of 75–125%. All samples passed these criteria except 1α,25(OH)2D3 measurement in native serum after SPE, which was below the LLOQ and above the 20% precision criterion. Average accuracies in the spiked samples were 99.6% for 25(OH)D3 and 111% for 1α,25(OH)2D3 in the LLE experiments and 92.3% for 25(OH)D3, 100.6% for 25(OH)D2, 106% for 1α,25(OH)2D3 and 94.4% for 24R,25(OH)2D3 in the SPE experiments. SPE results in more precise measurements of 25(OH)D3, while the accuracy of LLE for 25(OH)D3 is higher. Both extraction methods tend to overestimate 1α,25(OH)2D3 levels while precision is slightly better with SPE, which corresponds to the data obtained by direct comparison of both techniques (Table 3). Linear regression curves built for standard addition of 1α,25(OH)2D3 are more linear if the height of the peak (R2 = 0.9996) is used instead of peak area (R2 = 0.9850). For comparison, the R2 values of the linear regression curves for 24R,25(OH)2D3, 25(OH)D2 and 25(OH)D3 were 0.9985, 1.000, and 0.9993, respectively (see the “Electronic supplementary material”). The poor slope («1) obtained in the standard addition experiment for 24R,25(OH)2D3 can be explained with its higher polarity compared to 1α,25(OH)2D3, thus resulting in extraction losses that could not be accurately corrected due to the lack of the corresponding isotopically labeled standard.
Photosynthesis of vitamin D is a route parallel to dietary supplementation in humans (Fig. 1). UV light (290–315 nm) induces cleavage of the C-9–C-10 bond of 7-dehydrocholesterol with subsequent proton migration and conformational changes producing vitamin D3. For example, one minimal erythemal dose of UV exposure from sunlight resulted in the production of 10,000–20,000 IU (0.25–0.5 mg) of vitamin D3 . Typically, vitamin D photosynthesis in skin is thought to satisfy most human dietary requirements. However, the photosynthesis of vitamin D3 in skin becomes less efficient in people with darker skin living in higher latitudes, where sun exposure is insufficient to generate enough vitamin D3, or alternatively in people with mostly indoor lifestyles and those who habitually use sunscreens. An LC-MS method specific for D3 forms of vitamin D is particularly useful for studies of the photosynthesis of vitamin D3 in skin because it avoids interference with D2 metabolites from the diet. Because the major aim of our method development was to study the effect of skin pigmentation on vitamin D, we used a LLE version of the method to study the vitamin D deficiency in an HIV-positive urban Afro-American population previously reported to be 25(OH)D-deficient. The SPE version of the method was used for high-throughput phenotyping of vitamin D metabolism in subjects with different levels of sun exposure and different levels of skin pigmentation. The occurrence of D2 group metabolites was sporadic in studied groups and was therefore omitted from the discussion.
Effect of HIV infection on vitamin D profile
Seasonal variations in vitamin D metabolites
We analyzed 102 individual serum samples acquired from 34 subjects with different levels of sun exposure and different levels of skin pigmentation. The preliminary data showed that the majority of the studied population had a very low level of 25(OH)D2 (< 3.0 ng/mL), which corresponds to the fact that the diet is supplemented predominantly with the D3 form. However, in three of the 34 subjects the level of 25(OH)D2 was high (16.6, 9.2, and 6.3 ng/mL measured as the average of three time points). Those three subjects reported taking multivitamin supplements or drinking soy milk supplemented with vitamin D2. The ability to separate 25(OH)D2 and 25(OH)D3 forms of vitamin D is a specific feature of our method compared to typically used immunoassays. There is evidence that 25(OH)D2 does not have the same properties as 25(OH)D3 [9, 10]. In addition, in contrast to vitamin D3, which can be produced by photosynthesis in vivo, vitamin D2 is adsorbed solely from the diet (Fig. 1). Thus, separate measurement of the D2 and D3 forms of vitamin D is especially valuable for studying the photosynthesis of this vitamin.
While there are arguments that measuring 24R,25(OH)2D3 status has little biological value, it has been shown that this metabolite is active in bone tissue [14, 15]. Also, the level of 24R,25(OH)2D3 can be a measure of 1α,25(OH)2D3 clearance because 1α,25(OH)2D3 is also oxidized by 25-hydroxyvitamin D 24-hydroxylase, forming the inactive 1α,24R,25(OH)3D3. In addition, measurement of 24R,25(OH)2D3 can be a quality control for 1α,25(OH)2D3 analysis. It is well known that 25-hydroxyvitamin D 24-hydroxylase expression is positively regulated by 1α,25(OH)2D3 via a VDRE in its promoter . Thus, an elevated 1α,25(OH)2D3 level (>50–60 pg/mL) corresponds to high production of 24R,25(OH)2D3 if 25(OH)D3 status is sufficient.
Metabolic profiling is a promising tool for assessing entire metabolic pathways and studying their regulation. While in the field of vitamin D analysis different forms of protein binding assays remain the dominant procedures, they do not have the flexibility to measure multiple analytes in the same run or to separate the D2 and D3 forms of vitamin D. However, measurements of both 1α,25(OH)2D3 and 25(OH)D3 have become an important diagnostic factor for the assessment of dysregulated 1α,25(OH)2D3 extrarenal production in cancerous and inflammatory states. Thus, a method to measure 1α,25(OH)2D3 and 25(OH)D3 simultaneously could become a valuable tool in clinical practice. While this method represents a considerable improvement on the assessment of 25(OH)D3 by LC-MS, the very low circulating levels of 1α,25(OH)2D3, its thermal instability and its low polarity impede the direct measurement of this biologically important hormone with LC-MS or GC-MS. However, the sensitivity of 1α,25(OH)2D3 detection can be significantly improved using Diels–Alder derivatization with PTAD. Thus, the application of Diels–Alder derivatization allows the entire vitamin D cascade to be surveyed in a single LC-MS run, including routine direct measurements of 1α,25(OH)2D3, which has not been achieved before using LC-MS or GC-MS methods. In addition, the improved sensitivity of 25(OH)D detection allows the development of a method for the assessment of dietary 25(OH)D in very small plasma or serum samples (<50 μL). While current immunoassay methods for 1α,25(OH)2D3 quantification are superior in terms of their limits of detection, the reported LC-MS method can be used to detect excessive 1α,25(OH)2D2 and 1α,25(OH)2D3 production associated with some cancerous states and inflammation. It can also simultaneously assess the vitamin D dietary status via measurements of both 25(OH)D2 and 25(OH)D3 levels and estimate the rate of 1α,25(OH)2D3 clearance by 25-hydroxyvitamin D 24-hydroxylase via measurements of the 24R,25(OH)2D3 level. The method may be improved with the development of novel derivatization reagents with stronger ionic properties for more efficient electrospray ionization. By further expanding the list of analytes covered by this metabolic profiling method, we may gain unexpected insights into the biology of the vitamin D family of molecules.
We thank John Newman who provided the UPLC for the initial method development experiments. We thank Theresa Pedersen and Katrin Georgi for discussion of the extraction procedure, Mike Eskander for help with preparation of standards and MS optimization, Alina Wettstein for help with preparation of REACH samples and Leslie Woodhouse and Manuel Tengonciang for 25(OH)D RIA analysis. P.A.A. was supported by NIEHS Advanced Training in Environmental Toxicology Grant T32 ES007059. L.M.H. was supported by NIH Grant P60 MD00222-01. C.B.S. was supported by USDA-ARS Project 5306-51530-006-00D. K.D. was supported in part by BayGene. This research was supported in part by California Dairy Research Foundation Grant 07 HAB-01-NH, Bristol-Meyers/Squibb Freedom to Discover Award, NIEHS Grant R37 ES02710, NIEHS Superfund Basic Research Program P42 ES004699, NIEHS Center grant P30 ES05707, and NIEHS Center for Children’s Environmental Health & Disease Prevention Grant P01 ES11269.