Membrane transport of sepiapterin and dihydrobiopterin by equilibrative nucleoside transporters: A plausible gateway for the salvage pathway of Tetrahydrobiopterin biosynthesis

Molecular Genetics and Metabolism 102 (2011) 18–28

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Molecular Genetics and Metabolism j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / y m g m e

Membrane transport of sepiapterin and dihydrobiopterin by equilibrative nucleoside transporters: A plausible gateway for the salvage pathway of Tetrahydrobiopterin biosynthesis Akiko Ohashi a, Yuko Sugawara b, Kaori Mamada a, Yoshinori Harada b, Tomomi Sumi b, Naohiko Anzai c, Shin Aizawa a, Hiroyuki Hasegawa a,⁎ a b c

Department of Functional Morphology, Nihon University School of Medicine, Itabashi, Tokyo, 173-8610, Japan Department of Biosciences, Teikyo University of Science and Technology, Uenohara, Yamanashi 401-0193, Japan Department of Pharmacology and Toxicology, Kyorin University School of Medicine, Tokyo, 181-8611, Japan

a r t i c l e

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Article history: Received 3 July 2010 Received in revised form 15 September 2010 Accepted 15 September 2010 Available online 18 September 2010 Keywords: Tetrahydrobiopterin Supplement Sepiapterin Dihydrobiopterin Equilibrative nucleoside transporter Tetrahydrobiopterin Salvage pathway BH4-responsive PKU Endothelial cell NO-synthesis

a b s t r a c t Tetrahydrobiopterin (BH4) is synthesized de novo in particular cells, but in the case of a systemic or local BH4 deficiency, BH4 supplementation therapy is applied. BH4-responsive PKU has also been effectively treated with BH4 supplementation. However, the rapid clearance of the supplemented BH4 has prevented the therapy from being widely accepted. Deposition of BH4 after supplementation involves oxidation of BH4 to dihydrobiopterin (BH2) and subsequent conversion to BH4 by the salvage pathway. This pathway is known to be almost ubiquitous in the body. However, the mechanism for the redistribution and exclusion of BH4 across the plasma membrane remains unclear. The aim of this work was to search for the key transporter of the uptake precursor of the salvage pathway. Based on the observed sensitivity of pterin transport to nitrobenzylthioinosine (NBMPR), we examined the ability of ENT1 and ENT2, representative equilibrative nucleoside transporters, to transport sepiapterin (SP), BH2 or BH4 using HeLa cell and Xenopus oocyte expression systems. hENT2 was capable of transporting the pterins with an efficiency of SP N BH2 N BH4. hENT1 could also transport the pterins but less efficiently. Non-transfected HeLa cells and rat aortic endothelial cells were able to incorporate the pterins and accumulate BH4 via uptake that is likely mediated by ENT2 (SP N BH2 N BH4 ). When exogenous BH2 was given to mice, it was efficiently converted to BH4 and its tissue deposition was similar to that of sepiapterin as reported (Sawabe et al., 2004). BH4 deposition after BH2 administration was influenced by prior treatment with NBMPR, suggesting that the distribution of the administered BH2 was largely mediated by ENT2, although urinary excretion appeared to be managed by other mechanisms. The molecular basis of the transport of SP, BH2, and BH4 across the plasma membrane has now been described for the first time: ENT2 is a transporter of these pterins and is a plausible gateway to the salvage pathway of BH4 biosynthesis, at least under conditions of exogenous pterin supplementation. The significance of the gateway was discussed in terms of BH2 uptake for BH4 accumulation and the release for modifying the intracellular BH2/BH4 ratio. © 2010 Elsevier Inc. All rights reserved.

1. Introduction (6R)-L-erythro-Tetrahydrobiopterin (6RBH4 or simply BH4) is an enzymically active form of biopterin. The primary role of BH4 is that of an electron donor for a group of monooxygenase reactions such as involving phenylalanine hydroxylase [1], tyrosine hydroxylase [2], and tryptophan hydroxylase [3]. The enzyme complex nitric oxide synthase (NOS) requires BH4 both for the enzyme catalysis [4] and for functional

⁎ Corresponding author. Department of Functional Morphology, Nihon University School of Medicine, 30-1, Oyaguchi Kamimachi, Itabashi, Tokyo, 173-8610, Japan. Fax: +81 3 3972 0027. E-mail address: [email protected] (H. Hasegawa). 1096-7192/$ – see front matter © 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.ymgme.2010.09.005

dimerization [5]. In mammals, BH4 is synthesized de novo from guanosine triphosphate (GTP), and the pathway involves at least three enzymes, GTP-cyclohydrolase I, pyruvoyl-tetrahydropterin synthase, and sepiapterin reductase (reviews [6–8]). BH4 behaves as a coenzyme. That is, quinonoid dihydrobiopterin (qBH2 ), an oxidation product of BH4 in O2-reducing reactions, is replenished by in situ reduction by dihydropteridine reductase (DHPR: EC 1.6.99.7) and converted back to BH4 accompanied by consumption of NADH or NADPH [9]. The coenzyme is converted to inactive 7,8-dihydrobiopterin (7,8BH2 or simply BH2 ) through spontaneous isomerization of qBH2. It has long been known that BH4 is produced from sepiapterin (SP; 6-lactyl-7,8-dihydropterin) by two distinct enzymes, sepiapterin reductase (EC 1.1.1.153, Km to SP, 21 μM [10]) and dihydrofolate reductase (DHFR, EC 1.5.1.3, Km to 7,8BH2, 5–15 μM [11,12]), the latter

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of which converts 7,8BH2 to active BH4 [11,13–15]. The route of BH4 retrieval from SP or BH2 was termed the “salvage pathway” as opposed to the de novo pathway of BH4 biosynthesis [16]. BH4 is localized almost ubiquitously throughout most major organs as well as within the vascular system and is also detectable in cerebrospinal fluid and in urine. BH4 and its salvage pathway precursors exist in a state of dynamic redistribution between organs via the circulatory system. In our previous search for effective forms of BH4 supplementation, SP was found to be a good therapeutic choice [17]. Using various culture cells including RBL2H3 and HeLa, it was demonstrated that the transport of SP across the plasma membrane is equilibrative and bidirectional [18]. Although the process of cell entry is equilibrative, the cellular uptake of SP proceeds as an active transport owing to the following: (a) the salvage pathway strongly favors BH4 production, and (b) BH4 confined within the cell is hardly able to leak out. Since SP and 7,8BH2 as well as BH4 are inter-convertible, they are members of the “BP group” from a metabolic point of view. Hence, either SP or 7,8BH2 may be called a “BH4 precursor” of the salvage pathway. We were interested in the similarity between the role of the putative SP transporter(s) and that of the equilibrative nucleoside transporters (ENTs) as a precursor gateway for their respective salvage pathways. ENTs mediate the trans-membrane relocation of nucleosides and/or nucleobases in a bidirectional and equilibrative manner (reviews [19,20]). In this study, we examined whether ENT1 and ENT2 participate in the transport of BH4 precursors across the cell membrane in the salvage pathway of BH4 biosynthesis in mammalian cells. We then examined hENT1 or hENT2 in terms of their ability to transport SP and BH2 in a Xenopus-oocyte-expressing system. Because these ENTs were observed to transport SP and 7,8BH2, their role as the gateway to the BH4-salvage pathway was once again examined with non-transfected HeLa cells as well as for the first time with primary culture cells of rat aortic endothelium (ET). Furthermore, we surveyed the systemic effects of nitrobenzylthioinosine (NBMPR), a potential inhibitor of ENTs, in terms of the distribution and deposition of BH4 caused by exogenous administration of BH2 to individual mice. 2. Materials and methods (6R)-L-erythro-5,6,7,8-Tetrahydrobiopterin (6RBH4) was donated by Diichi-Asubio Pharma (Tokyo, Japan), and 6-lactyl-7,8-dihydropterin (SP, sepiapterin) and 7,8-dihydrobiopterin (7,8BH2) were purchased from Schircks Laboratories (Jona, Switzerland). N-Acetylserotonin (NAS), methotrexate (MTX), and collagenase (for Xenopus oocyte defolliculation) were purchased from Wako Pure Chemical Industries (Osaka, Japan). Nitrobenzylthioinosine (NBMPR, nitrobenzylmercaptopurineriboside), probenecid (4-(dipropylsulfamoyl)benzoic acid), o-coumaric acid, p-aminohippuric acid, penicillin G, tetraethylammonium sulfate, and cimetidine were obtained from Sigma-Aldrich (St. Louis, MO). Usually, 100× working solutions (100fold concentration over the final concentration) were prepared with solutions of DMSO, 0.1 M HCl or saline, and the pH of the medium was adjusted if needed. 2.1. Cell lines HeLa cells were maintained as a monolayer culture in Dulbecco's modified Eagle's medium (DMEM, GIBCO® Invitrogen) containing 10% fetal calf serum at 37 °C in 5% CO2/95% air. 2.2. Endothelial cells Endothelium was obtained as capillary outgrowth from rat aorta under 3D-culture on a collagen gel bed (Cellmatrix®, Nitta Gelatin, Osaka) essentially according to the manufacturer's instructions. In brief, the abdominal aorta was isolated from a young adult rat

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(Sprague–Dawley) under anesthesia, cut to a 5-mm length, placed on a collagen bed supplemented with an “ET-cell medium” containing VEGF, and kept at 37 °C in 5% CO2/95% air until the tissue had extended into a capillary outgrowth with dense branches (about 7 days). The “ET-cell medium” consisted of Nutrient Mixture F-12 Ham (N4888, Sigma-Aldrich) enriched with 20% fetal calf serum, MEM non-essential amino acids (Gibco), 1% glutamine, and 5 ng/mL rat recombinant VEGF (VEGF164®, R&D Systems Minneapolis, MN). The outgrowth was removed and subjected to digestion with collagenase (Collagenase S-1®, Nitta Gelatin). The dispersed cells were collected by centrifugation, washed thoroughly, plated on collagen-coated dishes, and then cultured in the “ET-cell medium”. The cells reached confluence at about 2.5 × 106 cells on a 10-cm dish 5 days after seeding at 2 × 105 cells. They were stocked at the 3 rd passage in liquid nitrogen. Cells restored from the liquid nitrogen were allowed to proliferate until numbers were sufficient for analyses at the 5th to 9th passages. 2.3. Xenopus oocytes African clawed frogs, Xenopus laevis, were purchased from Hamamatsu Seibutsu Kyozai (Hamamatsu, Japan). Each gonad was dissected under ice anesthesia and subjected to collagenase treatment (1 mg/mL, 1 h). Mature oocytes were then subjected to manual defolliculation, essentially according to Bianchi and Driscoll [21]. 2.4. Mice C57BL/6 J mice were obtained from Japan SLC (Hamamatsu, Japan). The animals were maintained on a constant 12-h light-dark cycle at 21–24 °C and 40–60% humidity with ordinary laboratory chow and tap water supplied ad libitum. 2.5. Cloning cDNA hENT1 and hENT2 were cloned from HeLa cells. The total RNA was extracted from the cell mass of HeLa cells using Isogen (Wako Chemical Industries, Osaka, Japan) as the protein denaturant. Transcripts were obtained by a reverse transcriptase reaction (PrimeScript®, TaKaRa, Shiga, Japan) using oligo-dT (17 bases) as the primer. The coding regions of hENT1 (ACCESSION NM_004955) and hENT2 (ACCESSION NM_001532) were separately amplified by PCR in the presence of a DNA polymerase (KOD-plus®, TOYOBO, Osaka, Japan) and appropriate primers with a restriction adaptor: hENT1 1–25 sense: TCCCCGCGGTTCGAAACCATGACAACCAGTCA CCAGCCTCAGG hENT1 1346–1371 antisense: GCTCTAGATCACACAATTGCCCGG AACAGGAAGG hENT2 1–21 sense: ACGCGTCGACTTCGAAACCATGGCGCGA GGAGACGC hENT2 1342–1371 antisense: CGCCTCGAGTCTAGATCAGAG CAGCGCCTGAAGA The cDNAs obtained by the following procedure were detected on agar electrophoresis and they were identified by sequencing with the following oligonucleotides as their primer: hENT1 386–405 sense: TGGTGAAGGTGCAGCTGGAT hENT2 396–415 sense: CTCCGTATGATTCATCAACT The PCR products were both cut to provide sticky 5′- and 3′-ends using Csp45I and XbaI, respectively. They were inserted into pENTER 11 (Gateway System® Life Technologies). Their mammalian expression vectors, pcDNA3.2/v5/hENT1 and pcDNA3.2/v5/hENT2, were then constructed using pcDNA3.2/v5/DEST and the LR-Clonase reaction according to the manufacturer's instructions.

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2.6. Transfection of HeLa cells Transfection of hENT1 and hENT2 was performed with their respective vectors with the aid of jetPEI (PolyPlus-Transfection, Illkirch, France). pCMV-SPORT-β-Gal (Invitrogen) was employed as a transfection control. Cells were treated with the vectors, hENT1, hENT2, and β-Gal in the presence of jetPEI in 10-cm dishes (1.5 × 106 cells) and allowed to express the relevant proteins for 20 h. The cells were then plated on 96 well analytical culture plate at 5 × 104 cells/ well with ordinary HeLa-cell culture medium. Several hours before the uptake experiment, the forced expression was assessed by visualizing β-Gal expression (pCMV-SPORT-β-Gal); the cells were rinsed with PBS, fixed with a mixture of formaldehyde (2%) and glutaraldehyde(0.2%) for 5 min, rinsed to remove the fixatives, and then added to a reaction mixture to allow the β-Gal reaction to proceed with 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (XGal) as the substrate at 37 °C for 1 h. When 30% of cells in culture sets were actively stained by blue precipitation, the protein was taken as overexpressed. 2.7. Expression of hENT1 and hENT2 in Xenopus oocytes Complementary RNAs (cRNAs) of hENT1 and hENT2 were prepared by in vitro transcription with T7 RNA polymerase in the presence of ribonuclease inhibitor and an RNA cap analog using a mMESSAGE mMACHINE kit (Ambion, Austin, TX). Defolliculated oocytes were injected with 50 ng of the respective cRNA or the same volume of water as the control and incubated in modified Barth's solution (82.5 mM NaCl, 2 mM KCl, 1 mM MgCl2, and 5 mM HEPES, pH 7.4) at 19 °C for 2 days. 2.8. Immunohistochemistry Xenopus laevis oocytes injected with cRNAs were fixed in 4% paraformaldehyde for 3 h, embedded in OCT compound, quick frozen on dry ice, sectioned (10 μm), placed on poly-L-lysine-coated slides, and stored at −20 °C. The sections were washed with PBS and given an additional two dips in surfactant-containing PBS. The slices on the slide were incubated with anti-ENT2 goat IgG (C-15, sc-48491, Santa Cruz Biotechnology, CA; diluted 1:50) in PBS overnight at 4 °C. They were washed three times with PBS for 10 min, then incubated with Texas-Red-labeled secondary rabbit anti-goat IgG (Vector Laboratories, CA; diluted 1:50) for 30 min at 4 °C, washed three times with PBS for 5 min each time and mounted with Vectabond Reagent (Vector Laboratories). 2.8.1. Transport experiment with Xenopus oocytes Five oocytes each were transferred to wells of a 96-well plate and preincubated in 100 μL ND96 buffer (96 mM NaCl, 2 mM KCl, 1 mM MgCl2, and 5 mM HEPES, pH 7.4) at 25 °C for 60 min. The pterin uptake was initiated by replacing the medium with 100 μL of the same buffer containing the desired concentration of the required ligands together with 1 mM dithiothreitol. Uptake was terminated at the designated time by washing oocytes three times with ice-cold ND96 buffer followed by the addition of 70 μL of acidI2 or alkaline-I2 solution as described below for biopterin analysis. Subsequently, the oocyes were crushed evenly using a plastic rod with a flat tip (5-mm diameter), and allowed to oxidize for 60 min. They were then mixed with 70 μL of 4% ascorbic acid in 4 M perchloric acid and cooled on ice for 1 h. Precipitates were removed by centrifugation. The supernatant retained a slight turbidity which was sucked off into a 3-mm cotton ball using a yellow-tipped pipette (Gilson) pressed against the bottom of the plastic tube (600-μL Eppendorf-type). The endogenous biopterin, 0.020–0.025 pmol/egg, was disregarded.

2.8.2. Uptake by cells under monolayer culture in 96-well culture plates HeLa or rat endothelial cells (ET cells) were plated on a 96-well analytical culture plate (Falcon 3072) and grown to a confluence of 5 × 104 cells/well with 200 μL of the appropriate culture medium the day before the experiments. Prior to the uptake experiments, cells were adapted to a “basal medium” for 15 min. The “basal medium” was a modified Hank's balanced salt solution which consisted of 137 mM NaCl, 5.37 mM KCl, 0.34 mM Na2HPO4, 0.44 mM KH2PO4, 0.34 mM K2HPO4, 5.5 mM glucose, and 5 mM HEPES, pH 7.4. Most transport experiments were conducted with reagents in the basal medium (100 μL) containing 1 mM dithiothreitol. Removal of the culture medium, leaving cells attached to the substrate, was performed by sucking off the medium with an 18-guage needle (connected to an aspirator) inserted vertically to lightly touch the bottom of the culture plate [18]. New medium was added with either a mechanical yellow-tipped pipette (Pipetman®, Gilson) or a repeating injector (Multipipette®, Eppendorf) of which the tips were first cut to create an inner diameter of 1.5 mm in order to avoid a sharp jet. Additional reagents were introduced by a thorough change of medium in individual wells after one rinse with the new medium. Particular components were also removed by a thorough change of medium following three repeated rinses with ice cold Ca2+and Mg2+-containing phosphate buffered saline. 2.9. Estimation of SP- and BH2-uptake by culture cells SP uptake was estimated by either a direct or indirect assay (see Supplementary Data 1). (A) For the direct assay by SP determination, internalized SP was determined after a 1-min release for virtual completion of “rapid” release after cells were fed SP in the presence of N-acetylserotonin (200 μM NAS, an inhibitor of sepiapterin reductase). (B) For the indirect assay by biopterin determination, SP was administered in the absence of NAS allowing the SP taken up to be converted to BH2 and subsequently to BH4 via the salvage pathway. BH2 uptake was also determined both directly and indirectly, since the uptake process proceeds in a complex fashion (see Supplementary Data 2). (A) For the direct BH2 assay, cells were fed 7,8BH2 in the presence of methotrexate (10 μM MTX, an inhibitor of DHFR, Ki ≈ 10 pM) and determination of BH2 or BPtotal (BH2 + BH4 ; without regard to BH4%) was performed after virtual completion of BH2 release via the “transient” process within 5 min. (B) For the assay for determining the BPtotal; 7,8BH2 was administered in the absence of MTX, allowing internalized BH2 to be converted to BH4 by the DHFR reaction. 2.9.1. Animal experiment Mice were administered 7,8BH2 at a dose of 5 mg/kg body weight (i.p.). The pterin was dissolved in 0.9% NaCl. NBMPR was suspended in saline containing 5% Na-carboxymethylcellulose using a mechanical mixer. The slurry was orally administered by gavage at a dose of 100 mg/kg. Under anesthesia, urine released at the time of cervical dislocation and blood in the still-beating heart were collected. The urine was immediately mixed with 9 vol of 10 mM HCl containing 5 mM ascorbic acid. The blood was diluted in 10 vol of 0.9% NaCl containing 5 mM each of EDTA and ascorbic acid. Luminal contents of the small intestine were collected by passing 2.7 mL of saline containing 5 mM ascorbic acid through the duct [22]. They were then acidified with HCl to a final concentration of 0.1 M and homogenized. Other organs were dissected, rinsed with saline, blotted, and then weighed. All samples were frozen in liquid nitrogen and stored at −80 °C until biopterin determination. 2.9.2. Determination of SP, BH2, and BH4 Pterins were determined essentially according to Fukushima and Nixon [23] with the following modification. Culture cells or Xenopus oocytes were washed, oxidized separately by the addition of 2% I2, 3% KI in 0.1 M HCl or in 0.2 M NaOH, for oxidation with acid-I2 and

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alkaline-I2, respectively. Biopterin and pterin (Ptn, 2-amino-4hydroxypteridine) were quantified by HPLC after excess I2 and proteins were removed by adding 4% ascorbic acid in 4 M perchloric acid (a volume equal to that of the I2/KI solution). The amount of biopterin after the acid-I2 oxidation was taken as the BPtotal in samples for which the BH4 content was not of concern. Since biopterin was not reproducibly detected in de-proteinized extracts without I2-oxidation from virtually any of the culture cells, the contents of fully oxidized biopterin were taken to be negligible. Hence, biopterin measured after alkaline oxidation was all taken to be BH2. When the Ptn amount was negligible after the acidic oxidation, the amount determined after the alkaline oxidation was taken as BH4 and that of biopterin, as BH2. In the case where endogenous compounds convertible to Ptn were not negligible, using the same procedure, oxidations with both acidic-I2 and alkaline-I2 were performed and the amount of Ptn in the acidic oxidation was subtracted from that in the alkaline oxidation to calculate BH4. Determination of pterin compounds in the animal tissues, including luminal contents of the small intestine as well as urine, were all performed after differential oxidation of both acid-I2 and alkaline-I2 so that BH2 and BH4 were distinctly determined. 2.9.3. HPLC analysis of pterins Sepiapterin was measured with an HPLC equipped with a fluorescence detector set at Ex = 412 and Em = 527 nm. Biopterin and Ptn were determined with the same fluorescence detector set at Ex = 350 and Em = 450 nm. In both analyses, the solid phase consisted of Fine-SIL C18T-5 (Jasco, Tokyo) and the mobile phase was 14% methanol for SP and 7% methanol for biopterin and Ptn. 3. Results 3.1. BH4 precursor: SP and BH2 uptake by ENT-transfected HeLa cells Our previous report demonstrated that SP uptake as the gateway of the salvage pathway was mainly controlled by bidirectional and equilibrative transporters in various cell types [18]. HeLa cells and rat ET cells are capable of taking up either SP or 7,8BH2 and accumulating BH4, and they were used here to assess various reagents for their effect on biopterin accumulation through the salvage pathway. The reagents included probenecid, o-coumaric acid, p-aminohippuric acid, penicillin G, tetraethylammonium sulfate, and cimetidine (1 mM each) and none of these significantly inhibited SP uptake in either type of cell. NBMPR (100 μM) inhibited about 80% of biopterin accumulation in the two cell types. Benzbromarone (50 μM, an inhibitor of a relatively wide range of transporters) was effective at inhibiting about 62% of SP uptake (50 μM) in HeLa cells and 12% of BH2 uptake (50 μM) in ET cells, but the cells detached from the culture plate at higher concentrations of this reagent. It was interesting that MTX (10 μM) did not inhibit uptake of either SP or BH2, but that it did block conversion of BH2 to BH4 resulting in a BH2 accumulation with no significant increase in BH4 in either type of cell, which was consistent with the finding that the BH4 accumulation was catalyzed by DHFR, the key enzyme of the BH4 salvage pathway. Taking the pronounced sensitivity to NBMPR into account, we focused on ENTs as a candidate for the responsible transporter. To determine whether hENTs are expressed in HeLa cells, we performed RT-PCR using gene-specific primers of 4 reported isoforms of ENT, hENT1 through hENT4, and we detected signals of ENT1, ENT2, and ENT3 in the total RNA extracts (data not shown). Using HeLa-cell RNA as the template, hENT1 and hENT2 were cloned as described in Materials and methods. Functional expression of hENT1 and hENT2 was examined using HeLa cells. As depicted in Fig. 1a, hENT2 was more potent than hENT1 in SP transport. Cellular accumulation of BPtotal caused by SP uptake (10 μM) in hENT1-transfected HeLa cells was roughly 2-fold greater than that in control cells, and the portion resulting from this increase

Fig. 1. Functional expression of hENT1 or hENT2 in HeLa cells exhibiting ability to uptake SP or BH2 as a gateway to the salvage pathway of BH4 biosynthesis. HeLa cells were transfected with pcDNA3.2/v5/hENT1 and/or pcDNA3.2/v5/hENT2 and plated on 96-well analytical culture plates (5 × 104 cells/well) 1 day before the experiment. Control cells were transfected with pCMV-SPORT-beta-gal. (a) SP uptake by ENTtransfected HeLa cells and its sensitivity to NBMPR: HeLa cells were transfected with control vector (open bars), hENT1 (black bars), or hENT2 (gray bars). Cells were preincubated with the indicated concentrations of NBMPR (0, 1, 10 or 200 μM), then SP (10 μM) was administered for 10 min. The SP uptake was expressed as BPtotal (BH2 + BH4, converted from sepiapterin in the cell). (b) Sensitivity of BH2 uptake to NBMPR in hENT1-transfected HeLa cells: hENT1-transfected HeLa cells (closed circles) and control cells (open circles) were administered the indicated concentrations of NBMPR (0 and 0.1–1000 nM) 15 min before initiation of BH2 uptake (50 μM). BH2 uptake was allowed to continue for 10 min and was terminated by the addition of alkaline I2-solution. In (a), **P b 0.01, ***P b 0.001; in (b), error bars are behind the respective symbols; all data are means ± S.D. (n = 4).

virtually disappeared in the presence of a low concentration of NBMPR (1 μM). Furthermore, hENT2-transfected cells exhibited a pronounced enhancement of SP uptake (ca. 11-fold) and the portion resulting from this increase was relatively resistant to NBPMR at 1 μM but was significantly inhibited at 200 μM (Fig. 1a). In view of the fact that BH2 is a more natural precursor of the salvage pathway than is SP, we employed both in the following experiments. Details of the NBMPR sensitivity of BH2 uptake by hENT1-transfected cells are depicted in Fig. 1b. BH2 uptake was enhanced, similar to that of SP uptake (2-fold), by transfection with hENT1. The half-maximum inhibition of BH2 uptake with hENT1-transfected HeLa cells was observed at around 30 nM NBMPR. The enhancement of both SP(Fig. 1a) and BH2-uptake (Fig. 1b) by hENT1 transfection completely disappeared at a low concentration of NBMPR, and levels were similar but slightly lower than levels in the control cells. Meanwhile, SP uptake by non-transfected HeLa cells as well as by hENT2-transfected cells was significantly inhibited with NBMPR (P b 0.01) but at a relatively high concentration, a finding which supports the idea that the endogenously expressed transporter of SP was common to the exogenously transfected hENT2.

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3.2. Transport of SP, BH2, and BH4 by Xenopus oocytes expressing hENT1 or hENT2 Xenopus oocytes were injected with the cRNA vector of ENT2. An immunochemical examination was performed to assess the proper expression of hENT2 on the plasma membrane. The fluorescent signal of hENT2 was observed in the periphery of the cRNA-injected oocytes and not at all in the control water-injected oocytes (Fig. 2a, compared to 2b). Xenopus oocytes injected with cRNA of hENT1 as well as with that of hENT2 were examined to determine their uptake of pterins. No significant amount of SP was detected in the oocytes after SP loading, however, a considerable accumulation of BH2 and BH4 was observed

in the cRNA-injected oocytes (roughly 80% was BH2 with BH4 as the remainder, Fig. 2c). The results suggested that the oocytes were furnished with high levels of sepiapterin reductase, enough to convert virtually all SP to BH2, and moderate levels of DHFR, as well as with NADPH, the co-substrate of both enzymes. Xenopus oocytes expressing either hENT1 or hENT2 progressively incorporated BH2 at much higher rates than the control oocytes (water-injected) as shown in Fig. 2d. The uptake of the respective pterins, SP, BH2 or BH4, by the hENT-expressing oocytes was almost linear at a substrate concentration of up to 500 μM (data not shown). The respective efficiencies of hENT1 and hENT2 in transporting SP, BH2 and BH4, were compared in series by measuring the BPtotal (Fig. 2e). hENT2-expressing oocytes

(b)

(c) SP Uptake BH4, BH2(pmol/h/oocyte)

(a)

50 40 30 20 10 0

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DW

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Pterin Uptake BPtotal(pmol/h/oocyte)

15 10 5 0

0

60

120

80

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60 40 20

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180

BH4

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BH2 Uptake BPtotal(pmol/oocyte)

BH2 Uptake BPtotal(pmol/oocyte)

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Hypoxathine

Uridine

Thymidine

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None

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Thymidine

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Fig. 2. Functional expression of hENT1 and hENT2 in Xenopus oocytes and the uptake of SP, BH2, and BH4 by these cells. Xenopus oocytes were injected with cRNA of hENT1 or hENT2 (50 ng per oocyte) and allowed to express the transporters at 19 °C for 2 days. Control oocytes were injected with distilled water (DW). (a) Immunohistochemical imaging was conducted with the primary antibody, anti-ENT2 goat IgG, and Texas-Red-labeled rabbit anti-goat IgG as the secondary antibody (pink fluorescence) as described in Materials and methods. The fluorescent image in the dark field was superimposed on an ordinary bright-field image. The pink fluorescence is located adjacent to and below the black margin of the egg pigment (scale bar, 100 μm). (b) Control oocytes show no significant fluorescence. (c) Conversion of incorporated SP to BH2 and BH4 by Xenopus oocytes: The hENT2-expressing oocytes were incubated in SP (50 μM) at 25 °C for 60 min, and the SP level in the oocyte was below the detection limit (data not shown). Accumulation of BH4 (black bars) and BH2 (stacked open bars) in hENT2-expressing oocytes. (d) ENT-expressing oocytes incorporated BH2 at enhanced rates: Oocytes received either water (open circles), hENT1-cRNA (closed circles), or hENT2-cRNA (gray circles). They were allowed to take up 7,8BH2 (50 μM) for the indicated times. Most error bars are hidden behind symbols. (e) Comparison of the transporters (hENT1 and hENT2) in terms of their uptake of pterins (BH4, BH2 or SP). Oocytes were treated for ENT expression as in (d); receiving either water (open bars), hENT1 (black bars), or hENT2 (gray bars). These oocytes were administered either 6RBH4, 7,8BH2, or SP (50 μM each) at 25 °C for 60 min. Biopterin uptake (BH4, BH2, and SP) by cRNAinjected oocytes was sigificantly enhanced. The rank order of biopterin transport was SPN N BH2 N BH4 in both hENT1- and hENT2-expressing oocytes. (f) Inhibition of BH2 uptake by ENT substrates in hENT1-expressing Xenopus oocytes: Oocytes were injected with hENT1-cRNA (closed bars) or water as a control (open bars). NBMPR (100 μM) or nucleic acids (1 mM each) were given to oocytes together with 7,8BH2 (50 μM). (g) Inhibition by ENT substrates in hENT2-expressing oocytes: Oocytes were injected with hENT2 (gray bars) or water (open bars). Experiments were performed as described in (f). *P b 0.05, **P b 0.01, ***P b 0.001, n.s. not significant; data are means ± S.D. (n = 4).

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took up SP and BH2 at a higher rate than did hENT1-expressing oocytes. Both of these hENT-expressing oocytes preferred SP as the substrate over BH2. Of the three pterins, BH4 was the least efficiently taken up by oocytes expressing either hENT1 or hENT2. The authentic ENT substrates, three nucleosides and one nucleobase, were examined in terms of their attenuation of BH2 uptake (hENT1, Fig. 2f; hENT2, Fig. 2g). Adenosine, thymidine and uridine (1 mM each) suppressed BH2 uptake (50 μM) in both hENT1- and hENT2-expressing oocytes. BH2 uptake by hENT1-expressing oocytes was not significantly inhibited with hypoxanthine (a nucleobase), while that by hENT2oocytes was. These results were compatible to earlier findings on the wider specificity of hENT2 which also transports nucleobases [24]. 3.3. HeLa cells: contribution of ENTs to the endogenous uptake of SP and BH2 The ability of hENT1 and hENT2 to transport SP, BH2 and BH4 has been demonstrated above. Consequently, we examined the contribution of these ENTs to the endogenous uptake of the BH4-salvage pathway precursors using non-transfected HeLa cells. HeLa cells were allowed to take up BH2 as well as SP in the presence of various ENT ligands (Fig. 3a) and the respective responses of these pterins to ENT ligand were found to be different. The increase in the BPtotal caused by

15 ***

10

Hypoxanthine

Thymidine

Uridine

0

(b) SP Uptake (%)

3.4. Rat ET cells: contribution of ENTs in uptake of SP and BH2 Rat ET cells proliferating in the presence of VEGF were examined for their ability to uptake SP, BH2 and BH4. The rank order of uptake efficiency was SP N BH2 N BH4 (Fig. 4a), similar to that of the ENTexpressing Xenopus oocytes. Their uptake of both SP and BH2 (10 μM each) was considerably inhibited by ENT substrates (1 mM each) as shown in Fig. 4b. Among ENT substrates, adenosine strongly inhibited uptake of both SP and BH2. The ENT inhibitor NBMPR was effective at BH2 uptake at concentrations higher than 1 μM, in strong accordance with the idea that the majority of BH2 uptake was caused by ENT2 rather than by ENT1 (Fig. 4c). Considerable portions of SP and BH2 uptake were resistant to a high concentration of NBMPR, suggesting that other transporters mediated the uptake and that these were insensitive to the reagent. 3.5. Individual mice: effect of NBMPR on BH4 distribution after BH2 administration

5

120 80 40 0

SP uptake (10 μM) was inhibited significantly by NBMPR (P b 0.001). ENT substrates, 1 mM each of the nucleosides, inhibited SP uptake. Hypoxanthine (1 mM), an ENT2 substrate, was also effective at inhibiting the uptake of SP. In the presence of a high concentration of NBMPR, however, a considerable uptake, representing more than 30% of SP transport, was observed, suggesting the involvement of other mediator(s). The profile of inhibition relative to the concentration suggested that the endogenous SP uptake was a multi-component process with at least two inflection points, at around 0.01 μM and 3 μM of NBMPR (Fig. 3b). On comparing BH2 uptake with that of SP (see open bars (SP) and gray bars (BH2 ) in Fig. 3a), relocation of BH2 was scarcely inhibited (most values were statistically insignificant) by the same reagents that considerably inhibited the uptake of SP.

* n.s.

Adenosine 50 µM NBMPR 0.1 µM NBMPR None

SP or BH2 Uptake BPtotal(pmol/10min/106cells)

(a)

23

0

10-8

10-6

10-4

NBMPR (M) Fig. 3. Effects of ENT ligand on endogenous uptake of SP or BH2 by HeLa cells. HeLa cells under monolayer culture (5 × 104 cells/well, 96-well analytical culture plate) were exposed to the ENT inhibitors or ENT substrates 15 min before the uptake was initiated. (a) Inhibition of NBMPR or ENT substrates (1 mM each) to SP- and BH2-uptake by HeLa cells: The cells were given 10 μM each of SP (open bars) or 7,8BH2 (gray bars) for 10 min. NBMPR did not significantly inhibit BH2 uptake. The relatively high residual transport of BH2 was only slightly inhibited by the ENT ligands. (b) Sensitivity of SP uptake to NBMPR in non-transfected HeLa cells: NBMPR was given to cells at the indicated concentrations, then SP (10 μM) was administered to commence uptake. The mean value obtained under control conditions (no NBMPR) was taken as 100%; the control value was 18.7 ± 1.2 pmol/10 min/106 cells. The thin line represents a superimposition of two dotted lines with a total height of 80% over the base line (horizontal line), one with an IC50 of 1 × 10− 8 M (downwards arrow) reaching 20% of the total SP transport, and the other with an IC50 of 3 × 10− 6 M (upwards arrow) and 60% of the total. All data are means ± S.D. (n = 8 in (a) and n = 4 in (b)).

The characteristic distribution of biopterin in mice is depicted in Fig. 5a–d. About 95% of most tissue biopterin (BPtotal) was in the form of BH4 while the remainder was BH2. In luminal contents of the small intestine, 62% of the biopterin was BH4. The endogenous biopterin content of various organs was not significantly altered by administration of NBMPR (100 mg/kg, p.o.) in 120 min. However, we noted that urinary excretion of BPtotal had decreased to 68% (P = 0.03) without any change in the BH4 percentage (BH4% N93%), as shown in Fig. 5d. When mice were administered 7,8BH2 (5 mg/kg, i.p.), a large increase in the BPtotal was observed in most organs (roughly 10-fold) at 30 min (compare upper panels, Fig. 5a–d, and bottom panels, Fig. 5e– h), while biopterin levels in the brain scarcely increased. Despite 7,8BH2 administration, the BPtotal in all tissues was mainly BH4; liver, 98.0% ± 0.6; kidney, 96.7% ± 0.9; intestinal mucosa, 90.1% ± 0.7; blood, 86.5% ± 7.4 (with BH2 as the remainder). With the prior administration of NBMPR (100 mg/kg, p.o., 90 min before BH2 administration), the deposition of BPtotal was further increased in the kidney (P b 0.001) and mucosa of the small intestine (P b 0.005) without any apparent alteration in the percentage of BH4, as shown in Fig. 5e and f. Although BH2 administration greatly increased biopterin in the liver (about 8fold, Fig. 5e), NBMPR treatment appeared to inhibit it, although only slightly (P = 0.07). NBMPR treatment after BH2 administration caused a significant increase in the blood BPtotal (P b 0.01), mainly due to a rise in BH2, while the increase in BH4 was moderate (barely significant, P = 0.08, Fig. 5g). At the same time, the luminal contents of the small intestine also showed a BH2 increase but the level of BH4 was unchanged (Fig. 5f). Both BH2 and BH4 have been reported to be mainly excreted from the liver as biliary juice, presumably via the gastrohepatic circulation at least in part [22]. The excretion of biopterin in urine (BH4, 62.3% ± 1.0 with BH2 as the remainder) increased tremendously, after BH2 administration, reaching about a 100-fold higher level. In contrast to the excretion of endogenous biopterin

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Pterin Uptake BPtotal(pmol/10 min/106cells)

(a) 25 20 15 10 5

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Hypoxanthine

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(c) BH2 Uptake (%)

120

80

40

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NBMPR (M) Fig. 4. Uptake of SP, BH2, and BH4 by rat ET cells, and effects of ENT ligand on BH2 uptake. Rat ET cells under monolayer culture (5 × 104 cells/well, 96-well analytical culture plate) were used. (a) ET cell uptake of BH4, BH2, and SP was compared. The cells were given 6RBH4, 7,8BH2, or SP (10 μM each) for 10 min. BH4 (closed bars) and BH2 (stacked open bars). (b) Competition of ENT substrates in SP- and BH2-uptake by ET cells: NBMPR (30 μM) or ENT substrates (1 mM each) were given to cells together with SP (open bars) or 7,8BH2 (gray bars). (c) Sensitivity of BH2 uptake to NBMPR in ET cells: NBMPR was given to cells at the indicated concentrations before 7,8BH2 (10 μM) was administered to commence uptake. The mean value obtained under control conditions (no NBMPR) was taken as 100%; the control value was 1.43 ± 0.03 pmol/10 min/106 cells. All data are means ± S.D. (n = 8 in (a) and (b), n = 4 in (c)).

(Fig. 5d), the increase in biopterin excretion after BH2 administration was not likely affected by the administration of NBMPR (Fig. 5h). 4. Discussion ENT1 has been classified as “es” based on its extreme sensitivity to NBMPR while other ENTs are referred to as “ei” owing to their relative indifference to the reagent. A species difference in the sensitivity to ligands was demonstrated between humans and rats. By nature, ENTs, especially ENT2, are characterized by a wide substrate specificity and by a relatively low affinity to their substrate (high Km, around 1 mM for nucleosides with ENT2) (reviews [19,20]). ENT1 and ENT2 were reported to be widely distributed in mammalian tissues (review [25]).

The respective expression of both ENTs has also been described in mice and rats [26]. In mammals, they play a fundamental role in the provision of nucleosides for salvage pathways of nucleotide synthesis in cells deficient in de novo biosynthetic pathways. Nucleoside transporters have also aroused great interest as effective but safe analogues of nucleosides for the development of anti-cancer and antiviral drugs (for reviews [27,28]). BH4 has been reported to be distributed almost ubiquitously, that is, not only in organs that synthesize it de novo but also in most cells and tissues not engaged in BH4-dependent monooxygenase reactions [29]. Along with exploration of NOS inclusion of BH4 as the essential cofactor, we have learned that BH4 is widely distributed in various locations in the body including the vascular tissues. Major BH4 deficiency causes hyperphenylalaninemia accompanied by neuronal dysfunction due to disruption of phenylalanine catabolism and aromatic monoamine biosynthesis. Local insufficiencies in BH4, particularly in the vascular system, have been reported to cause vascular dysfunction (for reviews [30–32]). There is a potential demand for BH4 supplementation as a means of treating certain lifestyle-related diseases. However, the relatively rapid turnover of BH4, and hence its short retention within the systemic circulation, has made the development of BH4 supplementation strategies very challenging. In addition, not enough is known about the kinetics and mechanism of the uptake, distribution, and excretion of biopterin after supplementation. On BH4 supplementation, many organs show a great accumulation of this pterin. Previous findings have demonstrated that when 6RBH4 was administered, systemic oxidation of BH4 to BH2 occurred before the cellular uptake of 7,8BH2 as the precursor of the salvage pathway (review [33]). In one study, the deposition of BH4 after administration of either SP or 7,8BH2, as well as 6RBH4, was efficiently suppressed by prior treatment with the DHFR inhibitor MTX [17]. In another study, when mice were administered 6SBH4, an unnatural diastereomer of BH4, the 6RBH4 that accumulated in their organs reached a level comparable to that after 6RBH4 treatment [34]. In this context, the ubiquitous distribution of BH4 might reflect the ubiquitous localization of the salvage pathway as well as its gating transporter for the uptake of precursors. In fact, in many cells that we have investigated, we detected an MTX-sensitive BH4 accumulation on administering either SP, 7,8BH2, or 6RBH4. These lines (not all data reported) include RBL2H3 (rat mast cell line), PC12 (rat adrenal medulla cell line), MBMEC (mouse brain microvascular endothelial cell line), HIT-T15 (rat pancreatic B-cell line), Cos-7 (monkey kidney cell line), and hepatocytes (rat, primary culture) [35]. Nutrient-absorbing epithelial cells, Caco-2 (porcine intestinal epithelial cell line) [35] and LLC-PK1 (porcine renal tubular epithelial cell line), and erythrocytes were exceptional in their ability to take up BH4 in a manner insensitive to MTX while they incorporated SP and BH2 and converted these to BH4 as well by the salvage pathway. In humans, SP might be derived from 6-lactoil-tetrahydropterin by oxidation, but SP was hardly detected in the healthy human body with a few exceptional cases [36,37]. Exogenous SP administration, however, has proven to be more efficient at raising BH4 levels in most organs than administration of 6RBH4 either by a p.o. or i.p. route [17]. Taking into account the presumable involvement of BH2 uptake after 6RBH4 administration, together with the remarkable efficiency brought about by SP administration, we searched for the transporter (s) responsible for the gating of the BH4 salvage pathway using SP and 7,8BH2 as the precursors in this study. 4.1. Ability of recombinant human ENT to transport SP and BH2 expressed in HeLa cells or Xenopus oocytes Cloned hENT1 and hENT2 were capable of transporting SP and BH2 when expressed in HeLa cells (Fig. 1) and Xenopus oocytes (Fig. 2). The African clawed frog, Xenopus laevis, was found to synthesize and

0 NBMPR - + - + - + - + Liver Kidney Spleen Brain

(e)

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150

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200

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6 4

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(h) 600

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500 400 300 200 100 0

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15

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(g)

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20 **

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(f) 250

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5

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(a)

25

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A. Ohashi et al. / Molecular Genetics and Metabolism 102 (2011) 18–28

- + Blood

-

+

Urine

Fig. 5. Systemic effect of NBMPR on biopterin distribution in mice with or without 7,8BH2 administration. Mice (C57BL/6 J) were administered NBMPR (100 mg/kg, p.o.) or the vehicle (−). Upper panels (a–d): Biopterin levels were determined 120 min after NBMPR administration in (a) organ tissues, (b) mucosa and luminal contents of the small intestine, (c) blood, and (d) urine. Lower panels (e–h): Mice received NBMPR as above, and were then administered 7,8BH2 (5 mg/kg, i.p.) 90 min after the NBMPR was given, followed by a biopterin determination 30 min after BH2 was given (120 min after NBMPR administration). Biopterin levels: (e) organ tissues including the brain (insert), (f) mucosa and luminal contents of the small intestine, (g) blood, and (h) urine. BPtotal (black bars) and BH4 (open bars). *P b 0.05, **P b 0.01, n.s. not significant; data are means ± S.D. (n = 4).

accumulate a tremendous amount of D-erythro-neopterin in skin chromatophores of both larvae and adults [38]. Neopterin synthesis was shown to begin a few hours after fertilization, at the time of pigment cell differentiation in the tail-bud stage of late embryogenesis [39]. Active synthesis of biopterin starts earlier than that of neopterin (unpublished observation). This made researchers curious as to whether these pterins were endogenous in the oocytes. However, no interfering amounts of SP, BH2 or BH4 were detected. When hENT1 or hENT2 was expressed, pterin uptake by these oocytes was greatly enhanced but was also inhibited by various ENT substrates, coherently reflecting the substrate specificities of hENT1 and hENT2 as reported. The ranking order of pterin transport of the BH4 group by hENT1- or hENT2-expressing Xenopus oocytes was in both cases SP N BH2 N BH4 (Fig. 2e), consistent with the observed order of these pterins in accumulating BH4 in rat ET cells (Fig. 4a) despite the difference in species in the two experiments. 4.2. Contribution of ENT1 and ENT2 to the uptake of SP and BH2 by HeLa cells and rat ET cells In the HeLa cell experiments using non-transfected cells, both hENT1 and hENT2 appeared to be involved in SP transport (Fig. 3a). The graphic profile of NBMPR inhibition of SP uptake (Fig. 3b) suggested that the participation of ENT1 and ENT2 was estimated to be around 20% and 60%, respectively, as follows. The line fitted to NBMPR

inhibition was approximated to be a superimposed line of two reverse sigmoid curves both with a Hill coefficient of (n = 1). One represented inhibition with a low concentration of NBMPR (IC50 ≈ 10−8 M) reaching less than 20% of the total SP transport, the other represented a higher NBMPR concentration (IC50 ≈ 3 × 10−6 M) to a maximum of about 60%. The source of about 20% of the residual SP uptake in the presence of 1 × 10−4 M NBMPR was unknown. The ratio of BH2 transport to SP transport (BH2/SP) in HeLa cells was ≈ 0.23 (Fig. 3a), higher than that of ENT2 suggested for hENT2expressing Xenopus oocytes with a BH2/SP of 0.12 (Fig. 2e). Curiously however, BH2 uptake by HeLa cells was virtually insensitive to ENT ligands including NBMPR and nucleic acids (Fig. 3a). Inhibition of BH2 transport by the ENT inhibitor NBMPR was not significant, and only a slight inhibition (less than 25%, P b 0.01) was observed with uridine. BH2 uptake in HeLa cells seemed, unlike SP uptake, to be largely mediated by some other transporter(s). The putative transporter of BH2 in HeLa cells remains to be explored at a later time. In any case, HeLa cells took up both SP and BH2 as the gateway of the BH4 salvage pathway despite the fact that these cells do not synthesize BH4 de novo. Endothelial cell cultures have frequently been studied in relation to NO synthesis. Biosynthesis of nitric oxide, a major component of the endothelium-derived relaxing factor, EDRF, requires BH4 as a rate limiting factor in these cells [40]. ET cells were reported to express ENT4 in addition to ENT1 and ENT2 [25]. hENT4 and mENT4 have a

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high affinity to adenosine, an anti-inflammatory mediator, and a unique preference for an acidic extracellular pH (lower than pH 7) in order to function [41]. The existence of another ENT which is moderately sensitive to NBMPR but distinct from ENT2, was also reported in human cardiac microvascular endothelial cells [42]. Our observations do not rule out the participation of such ENT homologues as these in the gateway of the BH4 salvage pathway of ET cells, but they are left unexplored at present. Meanwhile, ET cells of rat origin were demonstrated to be furnished with a potential transporter for uptake of the BH4 precursor, and in this case, it most probably is ENT2. These ET cells are able to uptake BH4 and accumulate it in that form, but with much less efficiency than with precursor supplementation. Assuming that human ET cells behave similarly, the high efficiency observed in the order of SP N BH2 N BH4 (Fig. 4a) points the way to a wider choice of therapeutics for vascular disorders caused by BH4 insufficiency. Although ET cells are capable of synthesizing BH4 de novo, the endogenous BH4 in the rat ET cells used in this study was 1–2 pmol/106 cells or less, less than 1 μM using an assumed ET cell volume of 2 pL for the calculation. The ET cell salvage pathway from sepiapterin to BH4 was first documented in relation to NOS function by Gross et al [40]. The endogenous content of BH4 in ET cells was first estimated with pieces of dissected canine cerebral arteries [43]. Both authors utilized sepiapterin successfully to increase cell/tissue biopterin, suggesting that ET cells were able to uptake SP and that their salvage pathway was functional. The in vivo involvement of the salvage pathway, i.e. via DHFR, in maintaining ET cell BH4, and BH2 was explicitly stated through the finding that reduced DHFR activity caused eNOS uncoupling in vascular tissues in lieu of NO production [44,45]. The actual concentration of the substrate of DHPR, qBH2, could be a fraction of the measurable biopterin (less than 1 μM). The Km of DHPR to the substrate qBH2, 1–5 μM [46,47] is rather large when compared with the estimated qBH2 concentration in the ET cell. Hence DHPR could not exert its full potential. The recycling by DHPR might therefore be inefficient due to the escape of qBH2 by spontaneous conversion to 7,8BH2, the substrate of DHFR. Therefore, one interpretation is that BH4 recycling in ET cells is more dependent on DHFR rather than on DHPR. Furthermore, a BH2 rise relative to BH4 has been reported to be the major cause of NOS uncoupling in vascular tissues, producing superoxide and consequent peroxynitrite formation [48–50]. ENT2 favors BH2 to BH4. In this context, by virtue of its bidirectional nature, ENT2 might play an important function as a drainer of 7,8BH2, excluding it from the cells and modifying the BH2/BH4 ratio. 4.3. Possible role of ENT in systemic distribution of BH4 after BH2 administration In individual mice, the distribution of biopterins without administration of BH2 was not altered significantly by NBMPR administration. Blood is the major carrier of BH2 as well as BH4 in the systemic circulation. The blood concentration of BPtotal determined in this work using mice was 0.83 ± 0.09 nmol/mL, similar to that of rats as reported by others [51,52]. The plasma biopterin was 15–20% of that in blood, hence, the concentration might be less than 0.2 μM. When biopterin is present at a submicromolar concentration in extracellular fluid, ENT-mediated uptake in most cells might be minute. Although attenuation of tissue biopterin levels was not detected 120 min after the drug treatment, this does not rule out the participation of ENT in tissue redistribution of biopterin. In particular, cells capable of synthesizing BH4 de novo might maintain physiological BH4 levels at around the Km values of the relevant pterindependent enzymes. Consequently, 7,8BH2 could be continuously generated via the spontaneous conversion of qBH2. A possible outward flow of 7,8BH2 via the bidirectional transporter might be the source of BH2 in the circulation. We speculate that the continuous outward flow of 7,8BH2 thus driven could be attenuated by the prior

administration of NBMPR but it was too slight to detect in the tissue. In support of this idea, we detected a significant net decrease in the urinary excretion of biopterin (Fig. 5d). BH4-responsive PKU patients [53] are known to be the most frequently and effectively treated with cofactor supplementation. These patients are believed to be normal in terms of BH4 homeostasis including regulated biosynthesis. In fact, they actually have higher than average BH4 levels owing to positive feedback from the GTPCH regulatory protein via their hyper levels of phenylalanine. It is still unknown how additional supplementation of BH4 restores the liver's ability to metabolize phenylalanine via a mutant hydroxylase protein that is unable to catalyze the reaction under normal BH4 levels. Mice with normal BH4 levels were administered BH2 (5 mg/kg) as a model for simulating how BH4 is distributed after the supplementation of BH4 in normal mice. Administration of BH2 to mice caused a pronounced deposition of BH4 without a big change in the BH4 percentage in most organs as well as blood within 30 min. This suggested that virtually none of the elevation in the BPtotal was due to flow-through of the injected BH2 but was instead caused by the cellular uptake of BH2 and its subsequent conversion to BH4 by the salvage pathway inside the cell. The transient elevation in plasma BH2 levels at the time of BH2 administration (i.p.) was considered to be high enough for ENTs, even those with low affinity to this biopterin, to translocate BH2 across the cell membrane, resulting in BH4 accumulation by the salvage pathway. Among the organs examined, the brain was the least responsive either to administration of BH2 alone or to co-administration of BH2 and NBMPR, despite the pronounced expression of ENT2 in the mouse brain [26]. We should recall that SP directly injected into the cerebral ventricle resulted in a pronounced increase in brain BH4 [54], a result indicating that SP was taken up by the brain cells and was converted to BH4 by the salvage pathway as long as the precursor reached the brain by any route. In interpreting our finding of only a faint responsiveness in the brain, we should note that it is not clear as yet exactly how BH2, BH4 and/or NBMPR are allowed to pass the blood brain barrier (BBB). Peripherally administered BH4 is well known to have difficulty in reaching the brain [55], and we considered that the BBB might have blocked entry of the pterin. Although the outflow of BH2 was not explicitly demonstrated in this report, BH2 might have been moved out of the cells owing to the bidirectional action of ENT. 7,8BH2 spontaneously converted from qBH2 might be exuded when BH4 oxidation exceeds the capacity of the DHPR recycling reaction in consumption of NADH or NADPH. The excreted BH2 after entering the circulation can thus serve as a precursor anywhere in the body. This redistribution of BH2 could be mediated by ENT2 in vivo. It was noted that the elevation in BPtotal in the urine (about 100-fold, more than 50% was BH4) after 7,8BH2 administration (5 mg/kg) far exceeded the increase in the blood (7- to 8-fold), but it was uncertain whether it was affected by NBMPR treatment (Fig. 5h). Under conditions in which the blood biopterins are elevated to unphysiologically high levels, the urinary excretion of biopterin might be driven by unknown but powerful mechanisms involving transporter(s) other than ENT. The observed increase in urinary biopterin might be a major hurdle barring wider BH4 supplementation such as in response to the needs of BH4-responsive PKU patients. 4.4. The role of ENT in BH4 metabolism Nucleoside transporters have been demonstrated to participate as the major gateway players of the salvage pathway of BH4 biosynthesis. ETN1 and ENT2 have been characterized as the biopterin transporters. The remarkable characteristics of ENT2 are (1) its equilibrative and bidirectional role as a transporter, (2) its low affinity but strong capacity to choose a substrate that can make it permeable

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across the plasma membrane, and (3) its wide distribution in various tissues in a polar or non-polar state depending on the polarity of the cell. The dual role of ENT under physiological conditions may be as follows. (A) As a gateway of the BH4 salvage pathway: cells whose de novo BH4 biosynthesis is so weak that they retain low levels of BH4, as observed with the ET cells under culture in this work, accumulate BH4 by almost constitutively expressing ENT2 and SPR as well as DHFR. When BH4 supplemented by means of exogenous administration of either SP, BH2, or BH4, all these cells might accumulate BH4. When BH4 is required by the cell, the accumulated BH4 might serve as the cofactor. Simultaneously these cells and other cells as well, which do not require BH4, function as a reservoir of the administered BH4. BH4 could remain in the cell until it is oxidized to 7,8BH2, which is made more permeable through the function of ENT2. Otherwise, exogenously administered pterins, SP, BH2, or BH4, would remain in the plasma and be directly excreted out into urine, presumably at a rate no slower than that of glomerular filtration in the kidney. (B) As a drain to remove inactive biopterin 7,8BH2 out of the cell: some particular cells maintain relatively large amounts of BH4 mainly by de novo biosynthesis. This group includes liver and kidney cells, both phenylalanine-hydroxylating, serotonin-, dopamine-, noradrenaline-neurons, enterochromaffin cells, adrenal medulla cells, all monoamine-synthesizing, and other unknown cells of BH4-rich organs [29]. Culture cells of these origins share a high capacity for BH4 synthesis, and include phenylalanine hydroxylating hepatocytes [35] and dopamine/noradrenaline-synthesizing sympathetic neurons [56], both of which were primary cultures, and cell lines of serotoninsynthesizing RBL2H3 and noradrenaline/adrenaline-synthesizing PC12 (in review [33,57]). BH4 levels in these cells might be maintained at near the Km of the relevant hydroxylase mainly by de novo synthesis and DHPR-catalyzed recycling [9]. In addition, their cellular concentration of qBH2, an oxidation intermediate of BH4, might be relatively high due to active hydroxylation and a high concentration of BH4, (roughly 20 to 40 μM). Hence DHPR activity may nearly reach its full potential, thus maintaining a good supply of biopterin for conversion to BH4. However, due to the short half-life of qBH2 (T1/2 ≈ 1.8 min [58]), the spontaneous conversion to 7,8BH2 might occur at a limited but stable rate. Thus, inefficient recycling by DHPR might occur when BH4 levels are too high, such as after BH4 supplementation. In cases where DHFR activity is too low for salvage to occur in situ, the large amounts of BH2 thus produced may be moved out of the cell by ENT2. The BH2 thus excluded might be the source of the salvage pathway precursor and may especially benefit BH4-poor cells such as ET cells of the vascular system. Although vascular endothelial cells are able to synthesize BH4 de novo, the amount of endogenous BH4 in these cells is largely dependent on the salvage pathway as mentioned with regard to eNOS function. In this case, it is uncertain whether the recovery of BH4 from 7,8BH2 by DHFR activity takes place in situ in individual cells or in other cells receiving exogenously supplied 7,8BH2 via the ENT transporter. The roles of ENT2 are thus made evident in both (1) the uptake of BH2 and (2) the modifying of the BH2/BH4 ratio under local and pathological conditions. Studies on the molecular basis of biopterin transporters have just begun with the work presented here and many aspects of biopterin transport remain elusive. 5. Conclusion ENT2, a representative nucleoside transporter, is ubiquitously expressed. This transporter can move SP and BH2 into the cell as precursors of the salvage pathway of BH4 biosynthesis. Owing to its equilibrative and bidirectional nature, in addition to its preference for BH2 over BH4, it should play a role in keeping down the intracellular BH2/BH4 ratio. Despite the low intrinsic affinity of ENT2 to its substrates, a slow migration of extracellular BH2 at a low concentra-

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tion might therefore occur across the cell membrane under physiological conditions. When the blood biopterin is raised to physiologically untenable levels, for example in the case of supplementation therapy with either SP, BH2 or BH4, ENT2 functions as a potent transporter in delivering the pterins to the intracellular salvage pathway. Moreover, the gateway also functions as a drain to remove inactive 7,8BH2 from cells when a high BH4 cannot be maintained by recycling via either DHPR or DHFR. Acknowledgments We thank Prof. Hitoshi Endou (Department of Pharmacology and Toxicology, Kyorin University School of Medicine), Kazuhiko Matsumoto (Torii Pharmaceutical Co., Ltd.), Hiroyuki Kusuhara (Graduate School of Pharmaceutical Sciences, University of Tokyo), and Michihiro Kasahara (Laboratory of Biophysics, School of Medicine, Teikyo University) for helpful discussions and critical suggestions. We appreciate our students, Atsumi Takabe, Keisuke Araki, Hiromi Hosokawa, for all their efforts in collaboration with the Student Research Program, Department of Biosciences, Teikyo University of Science and Technology. Appendix A. Supplementary data Supplementary data to this article can be found online at doi:10.1016/ j.ymgme.2010.09.005. References [1] S. Kaufman, The structure of the phenylalanine-hydroxylation cofactor, Proc. Natl. Acad. Sci. U. S. A. 50 (1963) 1085–1093. [2] T. Nagatsu, M. Levitt, S. Udenfriend, Tyrosine hydroxylase. the initial step in norepinephrine biosynthesis, J. Biol. Chem. 239 (1964) 2910–2917. [3] W. Lovenberg, E. Jequier, A. Sjoerdsma, Tryptophan hydroxylation: measurement in pineal gland, brainstem, and carcinoid tumor, Science 155 (1967) 217–219. [4] N.S. Kwon, C.F. Nathan, D.J. Stuehr, Reduced biopterin as a cofactor in the generation of nitrogen oxides by murine macrophages, J. Biol. Chem. 264 (1989) 20496–20501. [5] K.J. Baek, B.A. Thiel, S. Lucas, D.J. Stuehr, Macrophage nitric oxide synthase subunits. Purification, characterization, and role of prosthetic groups and substrate in regulating their association into a dimeric enzyme, J. Biol. Chem. 268 (1993) 21120–21129. [6] C.A. Nichol, G.K. Smith, D.S. Duch, Biosynthesis and metabolism of tetrahydrobiopterin and molybdopterin, Annu. Rev. Biochem. 54 (1985) 729–764. [7] B. Thony, G. Auerbach, N. Blau, Tetrahydrobiopterin biosynthesis, regeneration and functions, Biochem. J. 347 (Pt 1) (2000) 1–16. [8] G. Werner-Felmayer, G. Golderer, E.R. Werner, Tetrahydrobiopterin biosynthesis, utilization and pharmacological effects, Curr. Drug Metab. 3 (2002) 159–173. [9] S. Kaufman, Studies on the structure of the primary oxidation product formed from tetrahydropteridines during phenylalamine hydroxylation, J. Biol. Chem. 239 (1964) 332–338. [10] S. Kato, Sepiapterin reductase from horse liver: purification and properties of the enzyme, Arch. Biochem. Biophys. 146 (1971) 202–214. [11] S. Kaufman, Metabolism of the phenylalanine hydroxylation cofactor, J. Biol. Chem. 242 (1967) 3934–3943. [12] H.T. Abelson, R. Spector, C. Gorka, M. Fosburg, Kinetics of tetrahydrobiopterin synthesis by rabbit brain dihydrofolate reductase, Biochem. J. 171 (1978) 267–268. [13] M. Matsubara, M. Akino, On the presence of sepiapterin reductase different from folate and dihydrofolate reductase in chicken liver, Experientia 20 (1964) 574–575. [14] M. Matsubara, S. Katoh, M. Akino, S. Kaufman, Sepiapterin reductase, Biochim. Biophys. Acta 122 (1966) 202–212. [15] T. Fukushima, Biosynthesis of pteridines in the tadpole of the bullfrog, Rana catesbeiana, Arch. Biochem. Biophys. 139 (1970) 361–369. [16] C.A. Nichol, C.L. Lee, M.P. Edelstein, J.Y. Chao, D.S. Duch, Biosynthesis of tetrahydrobiopterin by de novo and salvage pathways in adrenal medulla extracts, mammalian cell cultures, and rat brain in vivo, Proc. Natl. Acad. Sci. U. S. A. 80 (1983) 1546–1550. [17] K. Sawabe, K.O. Wakasugi, H. Hasegawa, Tetrahydrobiopterin uptake in supplemental administration: elevation of tissue tetrahydrobiopterin in mice following uptake of the exogenously oxidized product 7, 8-dihydrobiopterin and subsequent reduction by an anti-folate-sensitive process, J. Pharmacol. Sci. 96 (2004) 124–133. [18] K. Sawabe, K. Yamamoto, Y. Harada, A. Ohashi, Y. Sugawara, H. Matsuoka, H. Hasegawa, Cellular uptake of sepiapterin and push–pull accumulation of tetrahydrobiopterin, Mol. Genet. Metab. 94 (2008) 410–416.

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