Molecular and Cellular Biochemistry

, Volume 400, Issue 1, pp 125–133

Sphingosine 1-phosphate lyase inhibition by 2-acetyl-4-(tetrahydroxybutyl)imidazole (THI) under conditions of vitamin B6 deficiency

  • Mamoru Ohtoyo
  • Masakazu Tamura
  • Nobuo Machinaga
  • Fumihito Muro
  • Ryuji Hashimoto

DOI: 10.1007/s11010-014-2268-z

Cite this article as:
Ohtoyo, M., Tamura, M., Machinaga, N. et al. Mol Cell Biochem (2015) 400: 125. doi:10.1007/s11010-014-2268-z


Caramel food colorant 2-acetyl-4-(tetrahydroxybutyl)imidazole (THI) causes lymphopenia in animals through sphingosine 1-phosphate lyase (SPL) inhibition. However, this mechanism of action is partly still controversial because THI did not inhibit SPL in vitro either in cell-free or in cell-based systems. It is thought that the in vitro experimental conditions which have been used so far were not suitable for the evaluation of SPL inhibition, especially in case of cell-based experiments. We speculated that the key factor might be the coenzyme pyridoxal 5′-phosphate (PLP), an active form of vitamin B6 (VB6), because media used in cell-based assays usually contain an excess amount of VB6 which leads to the activation of SPL. By the use of VB6-deficient culture medium, we could regulate apo- (without PLP) and holo- (with PLP) SPL enzyme in cultured cells, resulting in the successful detection of SPL inhibition by THI. Although the observed inhibitory effect was not as strong as that of 4-deoxypyridoxine (a VB6 analog SPL inhibitor), these findings may be useful for further understanding the mechanism of action of THI.


2-Acetyl-4-(tetrahydroxybutyl)imidazole Sphingosine 1-phosphate S1P lyase Vitamin B6 4-Deoxypyridoxine 


2-Acetyl-4-(tetrahydroxybutyl)imidazole (THI) is a component of ammonia caramel food coloring, which is commonly used in a variety of foods and beverages. A series of studies has shown that THI induces lymphopenia and affects the immune system in animals [1, 2, 3, 4, 5]. Although the mechanism was unknown for a long time, Schwab et al. revealed that THI exhibits its biological effects through sphingosine 1-phosphate (S1P) lyase (SPL) inhibition [6]. SPL, which is a pyridoxal 5′-phosphate (PLP)-dependent enzyme, irreversibly degrades S1P into hexadecenal and phosphoethanolamine. In addition, SPL is known to play an important role in generating an S1P gradient that regulates lymphocyte egress from lymphoid organs into peripheral blood [7]. These findings suggest SPL inhibitors may be promising candidates for novel immunosuppressants. In fact, a recent effort to identify potent SPL inhibitors resulted in the discovery of LX2931, which was obtained through the derivatization of THI and was developed for rheumatoid arthritis [8].

Although it has been unequivocally confirmed that THI decreases SPL activity in vivo, the inhibitory mechanism is partly still controversial because THI did not show any inhibitory activities on SPL in vitro. For example, it has been reported so far that THI failed to inhibit SPL in several cell-free assays [6, 9, 10]. SPL inhibition can also be estimated using cell-based assays in which the enzyme reaction is monitored by accumulation of S1P or dihydroS1P (dhS1P). Using cell-based systems, the non-specific SPL inhibitor 4-deoxypyridoxine (DOP) was found to increase the cellular S1P level in several cell lines [11, 12, 13], whereas THI induced neither significant increase of S1P in cells [14] nor detectable S1P secretion from cells [15]. These findings suggest that THI may act by an indirect mechanism, may require metabolism to an active state, or may be dependent on the formation of a higher-order complex [9, 16, 17, 18].

It is known that the in vivo effects of THI are prevented by a high level of dietary vitamin B6 (VB6) [1, 2]. VB6, which is converted into PLP by cellular enzymes, is essential for SPL to exert its enzymatic activity. From this point of view, we came up with a hypothesis that experimental conditions of conventional cell-based assays were not appropriate for evaluation of SPL inhibition. In most cases, cell-based assays are performed using cells cultured in commonly used culture media such as DMEM, RPMI1640, and so on. These media typically contain excess amount of VB6 as pyridoxine (~20 µM), which is considerably higher than plasma VB6 vitamers either in rodents (0.75–1.3 µM) [19, 20] or in human (<100 nM) [21]. Since conventional cell-based assays were carried out under such a “VB6-rich” condition, we speculated that VB6 may interfere with detection of inhibitory activity of THI. If this is the case, then it is expected that regulating VB6 concentration in cell culturing would bring appropriate conditions for estimating SPL inhibitors.

In this study, we investigated the inhibitory effects of THI on SPL in both cell-based systems (LC–MS quantification of S1P and TLC analysis of [3H]-sphingolipids) and a cell-free enzymatic assay. By the use of VB6-deficient culture medium, we could regulate apo- (without PLP) and holo- (with PLP) SPL enzyme in cultured cells, resulting in the successful detection of SPL inhibition by THI. Furthermore, we examined the mechanism of action of SPL inhibition especially with a focus on phosphorylation of compounds.

Materials and methods


Mouse thymic nurse cell line IT-79MTNC3 [22] was obtained from the Human Science Research Resources Bank. VB6-deficient [VB6(−)] DMEM was a custom order medium produced by Cell Science & Technology Institute (Sendai, Japan). [4,5-3H]d-erythro-dihydrosphingosine 1-phosphate ([3H]dhS1P, 60 Ci/mmol) and [4,5-3H]d-erythro-dihydrosphingosine ([3H]dhSph, 60 Ci/mmol) were supplied by American Radiolabeled Chemicals. Fumonisin B1 (FB1), non-labeled d-erythro-dhS1P, d-erythro-dihydrosphingosine (dhSph), and d-erythro-sphingosine (Sph) were from Enzo Life Sciences. DOP hydrochloride was obtained from Sigma. DOP 5′-phosphate (DOPP) and THI were synthesized with reference to previous reports [e.g., 23, 24]. All other chemical reagents used in this paper were of analytical grade or equivalent.

Quantification of intracellular S1P and PLP after treatment with compounds

IT-79MTNC3 cells were cultured in conventional DMEM (Invitrogen) supplemented with 10 % FBS (Hyclone), at 37 °C with 5 % CO2 in a humidified atmosphere. Subconfluent cells were harvested and suspended in conventional DMEM containing 10 % FBS and 25 µM FB1, then seeded into 24-well tissue culture plates at 5 × 10cells/well and incubated overnight at 37 °C with 5 % CO2. On the next day, medium was discarded, and the cells were further incubated for 4.5 h in the presence of various concentrations of test compounds in 450 µl of conventional or VB6(−) DMEM containing 0.1 % BSA (assay medium). Fifty microliters of each assay medium containing 10 µM Sph was then added and incubated for 0.5 h. To extract intracellular S1P and PLP, 150 µl of a mixture of acetonitrile and 1 M HCl (4:1, v/v) was added to the harvested cells and the cell suspensions were sonicated by a tip-type sonicator. Fifty microliters of the homogenate were mixed with 50 µl of 80 % MeOH containing internal standard and then filtered and centrifuged (3,000 rpm, 1 min). LC–MS/MS was performed at Mitsubishi Chemical Medience (Tokyo, Japan). Briefly, a 4000QTRAP triple quadrupole ion trap mass spectrometer (Applied Biosystems) interfaced with an Agilent 1200SL liquid chromatography system with an HTC-PAL autosampler (Agilent Technologies) was employed for ESI LC–MS/MS analysis with positive ion mode. The ion source conditions and gas settings were optimized with reference to previous reports [e.g., 25]. The multiple reaction monitoring transition monitored was m/z 380.3/264.2 for S1P and 248.2/150.1 for PLP, respectively. Chromatographic separation was achieved using an Atlantis T3 column (Waters) with gradient elution using solvent A; water/methanol/difluoroacetic acid (950/50/0.75, v/v/v) and solvent B; methanol/water/difluoroacetic acid (950/50/0.75, v/v/v). The calibration curve was constructed by adding increasing concentrations of S1P or PLP to a fixed amount of an internal standard.

SPL assay using TLC analysis of [3H]-labeled sphingolipids

Cells (1 × 105 cells/well in 12-well plate) were cultured overnight in conventional DMEM containing 10 % FBS and 25 µM FB1, then treated with increasing concentrations of test compounds for 4.5 h under VB6(−) condition as described above. In order to trace sphingolipid metabolism, cells were labeled with 0.5 µCi/well of [3H]dhSph for 0.5 h. (In case of just confirmation of FB1 effect without test compounds, cells were labeled with 1 µCi/well of [3H]dhSph for 1 h under normal condition.) Harvested cells were subjected to lipid extraction by chloroform/methanol/HCl (100:200:1, v/v/v) and vortexing, followed by addition of chloroform and 1 % KCl. After phase separation, extracted lipids were further separated by TLC on Silica Gel 60 high performance TLC plates (Merck) with 1-butanol/acetic acid/H2O (3:1:1, v/v/v) and visualized by autoradiography [26].

Sgpl1 mRNA expression analysis

After cells were treated with test compounds, total RNA were extracted, followed by cDNA synthesis. Sgpl1 (SPL-encoding gene) expression levels were estimated by RT-PCR using forward (5′-CTTGATGCACTTCGGTGAGA-3′) and reverse (5′-TTTAGGAACTGGATCGCCAC-3′) primers.

Cell-free SPL enzymatic assay

To prepare membrane fractions for use as an enzyme source, IT-79MTNC3 cells were washed twice with PBS and incubated overnight in VB6(−) DMEM containing 10 % dialyzed FBS. On the next day, harvested cells were frozen by liquid N2 and stored at −80 °C before use.

Cells were thawed and added to homogenization buffer [50 mM HEPES–NaOH (pH 7.4), 150 mM NaCl, 10 % glycerol, 1 mM EDTA, 1 mM dithiothreitol, complete protease inhibitor cocktail (Roche)], and sonicated with a tip-type sonicator. After centrifugation (1,000×g, 3 min), the supernatants were further centrifuged (70,000 rpm, 1 h) using Optima TLX ultracentrifuge system and TLA-120.2 rotor (Beckman Coulter). The precipitates were washed and suspended in homogenization buffer, then flash frozen in liquid N2 and stored at −80 °C.

In order to measure the SPL activity, membrane fractions (10 µg protein/test) and PLP were pre-incubated for 15 or 30 min at room temperature, followed by addition of 0.01 µCi/test of [3H]dhS1P and incubation at 37 °C for 1 h in 50 µl of reaction buffer [100 mM potassium phosphate (pH 7.4), 25 mM sodium fluoride, 5 mM sodium orthovanadate, 1 mM EDTA, 1 mM dithiothreitol, 0.1 % Triton X-100, 2 µM dhS1P]. Negative control sample reactions were performed on ice. The reaction mixture was mixed with 50 µl of 0.2 N NaOH and 100 µl of chloroform by vigorous vortexing, followed by centrifugation (8,400×g, 3 min). An aliquot of the aqueous phase containing [3H]dhS1P was collected, and radioactivity was measured by a liquid scintillation counter. The obtained count was assumed to reflect the amount of remaining substrates by the SPL reaction.

In inhibition assays, various concentrations of test compounds were pre-incubated with membrane fractions for 30 min prior to addition of PLP (final 5 µM).

Phosphorylation of compounds by PDXK

Human PDXK [27] was cloned, fused with FLAG-tag at the C-terminus, and was transiently expressed in HEK293 cells and purified by anti-FLAG affinity resin. The purified PDXK (4 µg/ml) was incubated with test compounds (final 200 µM) in the presence of 1 mM ATP at 37 °C for 1 h. After the reaction, HPLC analysis was performed using Agilent 1100 HPLC system and Discovery BIO Widepore C18 column (150 × 2.1 mm, 3 μm) under the following conditions: sample injection volume 10 μl, isocratic elution with 50 mM sodium phosphate, pH 3.1, 0.2 % acetonitrile, at a flow rate 0.5 ml/min, with detection of absorbance at 280 nm.

Metabolomic analysis of the cultured cell extracts

7.5 × 105 cells/sample were cultured overnight in conventional DMEM containing 10 % FBS and 25 µM FB1. After cells were treated with 1 mM of test compounds for 5 h under VB6(−) condition, the culture medium was aspirated from the dish and cells were washed twice by 5 % mannitol solution (10 ml first followed by 2 ml). The cells were then treated with 1.3 ml of methanol containing 10 μM of commercial internal standard solution 1 (Solution ID H3304-1002, Human Metabolome Technologies, Inc., Tsuruoka, Japan). Cells were scraped from the plate and cell lysates were obtained. Then, 520 µl of Milli-Q water and 1,300 µl of chloroform were added to the 1 ml of lysate, thoroughly mixed, and centrifuged for 5 min at 2,300×g and 4 °C. The upper aqueous layer (975 µl) was centrifugally filtered through a Millipore 5-kDa cut-off filter to remove proteins. The filtrate was lyophilized and suspended in 30 µl of Milli-Q water and analyzed by CE–TOFMS.

Metabolome measurements were carried out by a facility at Human Metabolome Technologies, Inc. (Tsuruoka, Japan) [28, 29, 30, 31].


THI elevates S1P level in cells under VB6-deficient condition

Based on the speculation that VB6 in assay medium may interfere with detecting the inhibitory effect of THI, we first investigated whether VB6 deficiency affects SPL inhibition in cell-based system in which intracellular S1P is quantified by LC–MS/MS. However, cell cultivation in VB6(−) medium in itself is expected to reduce cellular PLP level and consequently reduce SPL activity even in the absence of inhibitors because SPL is a PLP-dependent enzyme. If SPL activity is lost by excessive long-term cultivation in VB6(−) medium, it would be difficult to detect additional inhibitory effects by other compounds tested. In fact, we found that overnight cultivation under VB6(−) conditions caused a marked reduction of SPL activity as described below. For this reason, we cultured cells in conventional DMEM containing 10 % FBS and used VB6(−) assay medium during the exposure with test compound (for 5 h) as described in “Materials and methods” section. Although this medium exchange in itself led to a slight increase (2.6-fold) in basal level of S1P, we concluded that these conditions could still give enough dynamic range for detecting SPL inhibitory effects. Further improvement of the detection range can be achieved by adding Sph to the medium because Sph is taken up by cells and converted to the substrate S1P. Meanwhile, it is also expected that Sph would be converted to ceramide (Cer) by Cer synthases. Since inhibition of Sph-to-Cer conversion might lead to enhancing Sph-to-S1P conversion in a compensatory way, we therefore pre-treated cells with Cer synthases inhibitor FB1. TLC analysis of [3H]-labeled sphingolipids revealed that Cer/dihydroceramide and sphingomyelin bands almost disappeared in the presence of FB1, indicating that Sph-to-Cer conversion was certainly inhibited by this pre-treatment (Fig. 1). Then, we examined the effects of test compounds on intracellular S1P under the conditions of both normal and VB6(−) and found that DOP caused a detectable increase in the S1P level at quite a low concentration especially under VB6(−) condition (Fig. 2). The effective concentration, which was defined as resulting in a >2-fold increase of S1P, was 10 nM, and this is >1,000-fold lower than that under normal condition (40 µM). Surprisingly, THI caused an obvious increase in S1P at 100 µM under VB6(−), although it was not effective even at 1 mM under normal condition (Fig. 2). To our knowledge, this is the first report showing SPL inhibition by THI in in vitro experiment.
Fig. 1

Inhibition of ceramide synthesis by fumonisin B1 (FB1). Cells were incubated with or without 25 μM of FB1 overnight, followed by labeling with 1 µCi/well of [3H]dhSph for 1 h under normal condition. Then lipids were extracted, separated, and visualized as described in “Materials and methods” section. Rfs were estimated from the previous report [26] and the data from our in house experiments. GlcCer glucosylceramide, PS phosphatidylserine, PI phosphatidylinositol, PC phosphatidylcholine

Fig. 2

Effects of compounds on intracellular S1P level. Cells were treated with serial dilutions of test compounds, followed by incubation with Sph under normal (a) or VB6(−) condition (b). S1P contents of each sample were determined by LC–MS/MS measurement. X mark (×) indicates basal S1P level for no compounds. Each data represents the mean ± SD (n = 3)

TLC analysis of [3H]-labeled sphingolipids reveals SPL inhibition by THI

Although THI increased the intracellular S1P level under VB6(−) condition, this result alone does not necessarily indicate SPL inhibition because the S1P (or dhS1P) content in cells is regulated not only by SPL but also by sphingosine kinases (SPHKs) and S1P phosphohydrolases (SPPs). To confirm whether the above-observed S1P increase was due to the inhibition of SPL, we performed [3H]dhSph labeling analysis. TLC separation of extracted lipids and autoradiography revealed both an increase in dhS1P and simultaneous decrease in phosphatidylcholine (PC) with a similar effective concentrations (10 nM for DOP and 100 µM for THI, respectively) as observed in LC–MS/MS (Fig. 3). This pattern strongly suggests neither increased activity of SPHKs nor decreased activity of SPPs, but rather a definite decrease in SPL activity. In addition, we examined changes in cell viabilities (Fig. 4a) and Sgpl1 (SPL-encoding gene) expression levels (Fig. 4b) under VB6(−) conditions, but neither of them were significantly affected by test compounds. These results demonstrate that THI can indeed reduce cellular SPL activity without change in the amount of enzyme.
Fig. 3

TLC analysis of 3H-labeled sphingolipids. Cells pre-treated with FB1 were incubated with serial dilutions of test compounds, followed by labeling with [3H]dhSph under VB6(−) condition. Then lipids were extracted, separated, and visualized as described in “Materials and methods” section. Rfs was estimated from the previous report [26] and the data from our in house experiments

Fig. 4

Effects of test compounds on cell viabilities (a) and Sgpl1 expression (b) under VB6(−) conditions. a Cells were treated with test compounds under VB6(−) condition, followed by cell viability tests using Cell Count Reagent SF (Nacalai tesque). Cell viabilities (%) were compared with the control (no compound). Each data represents the mean ± SD (n = 3). b Cells were treated with test compounds under VB6(−) condition, followed by RT-PCR for Sgpl1 and β-actin (control) expression analyses

THI decreases cellular SPL activity without affecting the endogenous PLP level

SPL activity in a cell-based system can be modulated not only by direct interaction with compounds but also the supply of coenzyme PLP, which includes uptake of VB6 and conversion to PLP. Therefore, we investigated whether test compounds reduce the amount of intracellular PLP in the above experimental conditions. As expected, basal PLP level was decreased from 4 to 0.5 ng/5 × 104 cells by VB6(−) condition itself (Fig. 5). Under normal condition, DOP reduced PLP level in a concentration-dependent manner, while THI did not significantly affect it (Fig. 5a). Under VB6(−) condition, however, neither THI nor DOP changed PLP levels at the concentrations of up to 1 mM and 10 µM, respectively (Fig. 5b). Although it is unclear why DOP did not affect PLP level under VB6(−) condition, these results demonstrate that the decrease in cellular SPL activity caused by both DOP and THI under VB6(−) conditions did not result from preventing the supply of PLP.
Fig. 5

Effects of compounds on intracellular PLP level. Cells were treated with serial dilutions of test compounds, followed by incubation with Sph under normal (a) or VB6(−) condition (b). Cells were homogenized, and PLP contents of each sample were determined by LC–MS/MS measurement. X mark (×) indicates basal PLP level for no compounds. The lowest limit in the quantitative determination was 0.075 ng PLP/5 × 104 cells. Each data represents the mean ± SD (n = 3)

THI does not directly inhibit SPL in a cell-free enzymatic assay

All the results described above indicate that THI (or possibly its converted form) may directly inhibit SPL. Therefore, we investigated whether test compounds can directly inhibit SPL or not, using a cell-free assay system. It was reported that DOPP inhibits PLP-dependent glutamic-aspartic apotransaminase only when DOPP was added to the enzyme prior to PLP [32]. In accordance with this protocol, we examined the direct inhibitory effect on SPL using a membrane fraction which is derived from IT-79MTNC3 cells cultured overnight in VB6(−) DMEM containing dialyzed FBS. Long-term cultivation in VB6(−) culture medium is expected to markedly reduce the cellular PLP level, resulting in enrichment of apoSPL (the inactive form without PLP) compared to holoSPL (active form with PLP) in the membrane fraction. In fact, the membrane fraction in itself showed slight enzymatic activity, whereas an increasing amount of spiked PLP restored the enzyme activity and increased [3H]dhS1P degradation product up to 6.2-fold (Fig. 6a). In this system, restoration of enzyme activity by 5 µM of spiked PLP was inhibited by the addition of 10 µM DOPP prior to PLP, while neither DOP nor THI showed any effect even at 1 mM, suggesting that THI in itself does not directly inhibit SPL (Fig. 6b).
Fig. 6

Cell-free enzymatic assays using apoSPL-enriched membrane fraction. The amounts of degraded [3H]dhS1P were compared with the corresponding negative controls and shown as fold increase. Each data represents the mean ± SD (n = 3). a Increasing amounts of PLP were pre-incubated with membrane fractions for 15 min, followed by addition of [3H]dhS1P as a substrate. b Test compounds were added to the membrane fraction and incubated for 30 min prior to the addition of PLP (final 5 μM). After additional incubation for 30 min, [3H]dhS1P was added to initiate enzymatic reactions

In vitro analysis of enzymatic phosphorylation of compounds

It is known that DOP is phosphorylated by PDXK to form DOPP, which competes for the active site of various PLP-dependent apoenzymes [33]. From this finding, we speculated that THI also might be phosphorylated by cellular enzymes and acquire inhibitory activity against apoSPL. To confirm whether THI can be phosphorylated in the same way as DOP, we first performed cell-free phosphorylation study using recombinant PDXK and HPLC analysis. After the PDXK reaction, the peak corresponding to the pyridoxal (positive control) was shifted to a longer retention time which was consistent with that of the phosphorylated form (PLP, Fig. 7a). In the case of DOP, we observed a shifted peak which was identified as DOPP (Fig. 7b) by the consistencies with chemically synthesized DOPP in terms of retention time as well as m/z obtained from MS analysis of eluted fraction (data not shown). However, THI did not show such a peak shift, indicating that THI is not phosphorylated in the same manner as DOP (Fig. 7c).
Fig. 7

In vitro phosphorylation analysis of test compounds. Pyridoxal (PL, a), DOP (b) or THI (c) were incubated with (dotted line) or without (solid line) PDXK, followed by HPLC analysis of the reaction products

Comprehensive search of phosphorylated test compounds by CE–MS

In order to investigate another possibility that THI may be phosphorylated by enzymes other than PDXK, we performed a metabolomic analysis of cultured cell extracts derived from cells treated with test compounds under VB6(−) conditions. In this analysis, 539 peaks (259 in positive ion mode and 280 in negative ion mode) were detected in total (Online Resource 1). Among all these peaks detected, peaks with m/z matched to [M+H]+ or [M−H] of test compounds including their phosphorylated forms are listed in Table 1. In the group tested with DOP, a peak corresponding to [M−H] of DOPP (m/z 232.037 in negative ion mode) as well as a peak corresponding to [M+H]+ of DOP (m/z 154.085 in positive ion mode) were observed. In the database of Human Metabolome Technologies, Inc., the later peak (m/z 154.085) was also annotated as dopamine which has the same molecular formula as DOP (C8H11NO2). However, we concluded that this peak is derived from DOP because these two peaks were not detected in both the control group and the group tested with THI. Even though peak identification by MS/MS is necessary to obtain a definitive conclusion, these results are enough to suppose that DOPP is actually generated from DOP by a cellular enzyme. In the case of THI, we detected a peak corresponding to [M+H]+ of THI (m/z 231.096 in positive ion mode). Although this peak was also observed in both the control and DOP-treated groups, the ratio to the control was over two orders of magnitude, suggesting that it overlaps with a peak of some metabolite originally existing in cells. However, unlike the case of DOP, we could not detect any peaks corresponding to [M−H] of putative mono- (m/z 309.049 in negative ion mode), di- (m/z 389.015), tri- (m/z 468.981), and tetra- (m/z 548.948) phosphorylated THI.
Table 1

Metabolomic analysis of the cultured cell extracts

Peak IDs


MT (min)

Average relative area (n = 3)






















Among all peaks detected, peaks with m/z matched to [M+H]+ or [M−H] of test compounds (and their phosphorylated form) are listed. A complete list is provided in the Online Resource 1

ND not detected


In this study, we investigated the mode of action of THI by focusing in particular on the VB6 concentration in the assay system. While excess dietary VB6 prevents biological effects of THI in vivo, most in vitro assays were carried out under VB6-rich condition. Based on this, we speculated that VB6 may interfere with detecting the inhibitory activity of THI. Consequently, the use of VB6(−) medium markedly sensitized SPL inhibition in a cell-based assay, resulting in the first clarification of the inhibitory effect of THI in in vitro experiments (Figs. 2b, 3).

Although we confirmed that THI certainly decreases SPL activity in cell-based systems under VB6(−) condition, it was still unclear whether THI inhibits SPL directly or not, because cellular SPL activity would be generally reduced not only by direct inhibition of enzyme but also by inhibition of VB6 uptake and its metabolism. In this experiment, however, we observed that the cellular PLP levels were not affected by DOP and THI (Fig. 5b), indicating that the VB6 uptake and PLP synthesis pathway were not responsible for SPL inhibition in this VB6(−) condition. Taken together, at least under VB6(−) conditions, it was suggested that the decrease in cellular SPL activity caused by both DOP and THI resulted not from preventing the supply of PLP or decreasing SPL expression, but from the direct/indirect interaction with the enzyme itself. For further confirmation of this, we should investigate precisely whether THI inhibits VB6 uptake or enzymes involved in PLP synthesis such as PDXK and pyridoxine 5′-phosphate oxidase.

With respect to DOP, the effective concentration in cell-based assays [10 nM under VB6(−) conditions] was quite different from that of DOPP in cell-free assay (10 µM). A possible reason for this discrepancy is that DOP may be accumulated in cells. It is known that DOP inhibits VB6 uptake in a competitive manner [34], implying that DOP is also a substrate of a putative VB6 transporter which has not been identified in mammals yet. In addition, it was reported that active transport of VB6 occurs in yeast, leading to intracellular concentrations of VB6 much higher than those supplied externally [35]. Taken together, it is supposed that DOP uptake would not be inhibited under VB6(−) condition, and as a result, the intracellular concentration of DOP (or DOPP) might become higher than the extracellular concentration. Additional studies will be needed to address this matter.

Since DOP is a VB6 analog SPL inhibitor, the hypersensitization under VB6(−) conditions was not surprising. The question is whether THI, which exhibited similar behavior as DOP in this study, is a VB6 analog-type inhibitor. In cell-free assay, however, we could not observe direct interaction of THI with apoSPL (Fig. 6b). Although DOP is converted into a phosphorylated form (Fig. 7b), there are no reports and clues about phosphorylation of THI. Moreover, THI exerts in vivo lymphopenic effects at a lower plasma concentration than the effective concentration used in this study [36]. These findings suggest again that THI may be converted into unidentified active metabolites or act by another mechanism. We hope our results will help in further understanding the mechanism of action of THI.


The authors are grateful to Dr. Mika Ikeda for practical advice, project team members for sharing insights, and Human Metabolome Technologies, Inc., for metabolomic analysis. The authors also thank Drs. Ryuta Koishi, Gen Kudo and Hiroshi Yokota for useful suggestions, and Drs. Tohru Takahashi and Hidehiko Furukawa for support.

Supplementary material

11010_2014_2268_MOESM1_ESM.xlsx (80 kb)
Supplementary material 1 (XLSX 79 kb)

Copyright information

© Springer Science+Business Media New York 2014

Authors and Affiliations

  • Mamoru Ohtoyo
    • 1
  • Masakazu Tamura
    • 2
  • Nobuo Machinaga
    • 3
  • Fumihito Muro
    • 4
  • Ryuji Hashimoto
    • 1
  1. 1.New Modality Research Laboratories, R&D DivisionDaiichi Sankyo Co., Ltd.TokyoJapan
  2. 2.Biologics Pharmacology Research Laboratories, R&D DivisionDaiichi Sankyo Co., Ltd.TokyoJapan
  3. 3.Medicinal Chemistry Research Laboratories, R&D DivisionDaiichi Sankyo Co., Ltd.TokyoJapan
  4. 4.Vaccine Business Strategy Department, Vaccine Business Intelligence DivisionDaiichi Sankyo Co., Ltd.TokyoJapan

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