GNL knockout mice

GNL knockout mice

Archives of Biochemistry and Biophysics 496 (2010) 38–44 Contents lists available at ScienceDirect Archives of Biochemistry and Biophysics journal h...

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Archives of Biochemistry and Biophysics 496 (2010) 38–44

Contents lists available at ScienceDirect

Archives of Biochemistry and Biophysics journal homepage: www.elsevier.com/locate/yabbi

Ascorbic acid depletion enhances expression of the sodium-dependent vitamin C transporters, SVCT1 and SVCT2, and uptake of ascorbic acid in livers of SMP30/GNL knockout mice Akiko Amano a,b, Toshiro Aigaki a, Naoki Maruyama b, Akihito Ishigami c,* a b c

Cellular Genetics, Graduate School of Science and Engineering, Tokyo Metropolitan University, Tokyo 192-0397, Japan Aging Regulation, Tokyo Metropolitan Institute of Gerontology, Tokyo 173-0015, Japan Department of Biochemistry, Faculty of Pharmaceutical Sciences, Toho University, Chiba 274-8510, Japan

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Article history: Received 25 November 2009 and in revised form 23 January 2010 Available online 1 February 2010 Keywords: Ascorbic acid Dehydroascorbic acid Glucose transporter Senescence marker protein-30 Sodium-dependent vitamin C transporter

a b s t r a c t In this study, we examined whether ascorbic acid (AA) and dehydroascorbic acid (DHA), the oxidized form of AA, levels in tissues regulate the AA transporters, sodium-dependent vitamin C transporters (SVCT) 1 and SVCT2 and DHA transporters, glucose transporter (GLUT) 1, GLUT3, GLUT4 mRNA by using senescence marker protein-30 (SMP30)/gluconolactonase (GNL) knockout (KO) mice. These mice are incapable of synthesizing AA in vivo. AA depletion enhanced SVCT1 and SVCT2 mRNA expression in the liver and SVCT1 and GLUT4 mRNA expression in the small intestine, but not in the cerebrum or kidney. Next, we examined the actual impact of AA uptake by using primary cultured hepatocytes from SMP30/GNL KO mice. In the AA-depleted hepatocytes from SMP30/GNL KO mice, AA uptake was significantly greater than in matched cultures from wild-type mice. These results strongly affirm that intracellular AA is an important regulator of SVCT1 and SVCT2 expression in the liver. Ó 2010 Elsevier Inc. All rights reserved.

Introduction In vitro systems, ascorbic acid (AA)1 acts as an anti-oxidant by scavenging a variety of harmful radicals [1–3]. AA ionizes at the hydroxyl C-2 or C-3 positions of chemical structure and exists as a monovalent anion in vivo [1,4]. AA is also an essential substrate for various proline hydroxylases that hydroxylate proline residues in both collagen proteins and some enzymes with a transcriptional activity like hypoxia inducible factor [5]. The ascorbyl radical is initially generated from these reactions and this radical dismutates to AA and dehydroascorbic acid (DHA), the oxidized form of AA [4,6– 8]. Moreover, a protective effect of AA supplements toward oxidative DNA damage in white blood cells is also reported [7,9–11]. AA and small amounts of DHA are virtually present in all tissues under physiological conditions [12,13]. Although many animals can synthesize

* Corresponding author. Address: Department of Biochemistry, Faculty of Pharmaceutical Sciences, Toho University, Miyama 2-2-1, Funabashi, Chiba 274-8510, Japan. Fax: +81 47 472 1536. E-mail address: [email protected] (A. Ishigami). 1 Abbreviations used: AA, ascorbic acid; DHA, dehydroascorbic acid; DMEM, defined minimum essential medium; ECD, electrochemical detection; EDTA, ethylenediaminetetraacetic acid; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; Glut, glucose transporter; GNL, gluconolactonase; HPLC, high-performance liquid chromatography; KO, knockout; RT-PCR, real-time polymerase chain reaction; SMP30, senescence marker protein-30; SVCT, sodium-dependent vitamin C transporter. 0003-9861/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2010.01.012

AA in vivo, others such as humans, nonhuman primates and guinea pigs have lost the ability to make AA because of many mutations in the L-gulono-c-lactone oxidase gene, which is essential for AA synthesis in vivo, and resulting lack of that enzyme’s activity [14]. Therefore, the latter group must obtain AA from dietary sources. The sodium-dependent vitamin C transporters 1 (SVCT1) and SVCT2, both of which are 12-transmembrane proteins [15], have a functional role in sodium-dependent, secondary active transport of AA from outside to inside cells [16]. Recent study indicated that SVCT1 is involved in the whole-body homeostasis of AA and exhibits a higher maximum velocity (Vmax) value than SVCT2 [17]. The Michaelis constant (Km) value of SVCT1 ranges from 65 to 237 lM. Human SVCT1 is distributed in numerous tissues including lung, liver, kidney, intestine and skin [17]. In contrast, human SVCT2 occupies mainly the brain, eye, liver, kidney, intestine, adrenal gland, bone and skeletal muscle [17]. SVCT2 has a lower Km (8– 115 lM) value than SVCT1 and, as a high affinity AA transporter, takes up lower concentrations of AA than does SVCT1 [17]. Despite these reports of SVCT1 and SVCT2 mRNA expression in multiple mouse tissues [18], their expression levels have not been compared nor has the relationship between SVCT expression and tissues’ content of AA. Moreover, we do know that DHA (AA’s oxidized form) is transported by the glucose transporter 1 (GLUT1), GLUT3 and GLUT4 [13,19]. Glucose is also known to be a competitive inhibitor of DHA uptake in GLUT1 and GLUT3 [19], and GLUT4 is a participant

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in insulin-stimulated translocation from a subcompartment of the trans-Golgi reticulum to the plasma membrane [20]. Regarding the regulation of SVCT expression in vitro, epidermal growth factor proved to be an inducer of SVCT2 mRNA expression in the human trophoblast cell line HTR-8/SVneo [21]. Further, calcium and phosphate ions induced SVCT2 mRNA expression and increased AA uptake in MC3T3-E1 osteoblastic cells [22]. However, efforts to identify the regulatory mechanisms of SVCT and GLUT mRNA expression in vivo are still incomplete, because the experimental animal of choice is the mouse, and their ability to synthesize AA presents considerable difficulty in controlling their AA and DHA levels. We previously established a strain of senescence marker protein30 (SMP30)/gluconolactonase (GNL) knockout (KO) mice [23,24], which cannot synthesize AA, because SMP30/GNL is required for that process [25]. However, we can control the bodily status of AA in SMP30/GNL KO mice by providing AA in their food and drinking water. Thus, SMP30/GNL KO mice are a powerful human-like model for investigating the internal movements and effects of AA as well as the regulation of SVCTs and GLUTs mRNA expression in vivo. Here, we used the quantitative real-time polymerase chain reaction (RT-PCR) to determine whether AA and DHA levels in tissues regulate SVCT and GLUT mRNA expression levels. By using primary cultured hepatocytes from SMP30/GNL KO mice, we found that SVCT1 and SVCT2 mRNA expression levels and AA uptake ability were significantly enhanced in the hepatocytes of AA-depleted SMP30/GNL KO mice, indicating that AA manifests a marked impact as a regulatory element of SVCT1 and SVCT2 expression in the liver. Materials and methods Animal SMP30/GNL KO mice were generated by the gene targeting technique described previously [23]. Female KO mice (SMP30/ GNL/) were mated with male KO mice (SMP30/GNLY/) to produce KO mice. In this study, only male KO mice were used. Male wildtype (WT) (SMP30/GNLY/+) mice were purchased from Japan SLC (Shizuoka, Japan). At 30 days of age, SMP30/GNL KO and WT mice were divided into the following four groups: AA(+) SMP30/GNL KO, AA() SMP30/GNL KO, AA(+) WT, and AA() WT mice. The AA(+) group had free access to water containing 1.5 g/L AA and 10 lM ethylenediamine tetraacetic acid (EDTA), whereas the AA() group had free access to water without AA until the end of experiment when the animals were 63-days-old. Water bottles were changed every three or four days until the experiment ended. All mice were fed an AA-deficient diet (CL-2, CLEA Japan, Tokyo, Japan). Throughout the experiments, animals were maintained on a 12-h light/dark cycle in a controlled environment. All experimental procedures using laboratory animals were approved by the Animal Care and Use Committee of the Toho University and Tokyo Metropolitan Institute of Gerontology. Preparation of tissues Mice were sacrificed and systemically perfused with ice-cold phosphate buffered saline through the left ventricle to wash out remaining blood cells. Tissues were collected and stored at 80 °C until use. Measurement of AA and DHA AA was measured by using a high-performance liquid chromatography (HPLC)–electrochemical detection (ECD) method as described previously [26]. Tissues were homogenized in 14 and

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50 (only adrenal gland) volumes of 5.4% metaphosphoric acid by using Potter–Elvehjem Teflon homogenizer and centrifuged at 21,000g for 15 min at 4 °C. To measure the AA plus DHA, centrifugal supernatants obtained were suitably diluted, and samples (90 ll) were treated with 10 ll of 350 mM tris (2-carboxyethyl) phosphine hydrochloride then incubated for 2 h at 4 °C to reduce DHA to AA. After 2 h, 965 ll of 5% metaphosphoric acid was added to samples and centrifuged at 21,000g for 10 min at 4 °C. Then, the resultant supernatants were analyzed by HPLC using an Atlantis dC18 5 lM column (4.6  150 mm, Nihon Waters, Tokyo, Japan). The mobile phase consisted of 50 mM phosphate buffer (pH 2.8), 540 lM EDTA and 2% methanol at a flow rate of 1.3 mL/min, and electrical signals were recorded by using an electrochemical detector with a glassy carbon electrode at +0.6 V [27,28]. For measurement of AA alone, 90 ll of the centrifugal supernatant of tissues was diluted with 975 ll of 5% metaphosphoric acid and analyzed by HPLC–ECD. Actual DHA content in tissue samples was calculated by subtracting amounts of the AA from AA plus DHA. Extraction of total RNA and cDNA synthesis Total RNA was extracted by using ISOGENÒ (Wako Pure Chemical, Osaka, Japan) [29]. Tissue samples were homogenized with a Teflon-pestle homogenizer in ISOGEN, and total RNA was extracted according to the manufacturer’s protocol. The final RNA pellet was dissolved in diethyl pyrocarbonate-treated H2O. RNA concentrations were determined and confirmed as free from protein contamination by measuring absorbance at 260 and 280 nm. Then, cDNA was synthesized using SuperScript II Reverse transcriptase (Invitrogen, Carlsbad, CA, USA) following the manufacturer’s protocol. The cDNA was stored at 80 °C until use. Quantitative RT-PCR RT-PCR reactions were performed in duplicate by using the qPCR super mix-UDG-with ROX (Invitrogen) following the manufacturer’s protocol. The forward and reverse primers for SVCT1 (Assays Assay ID: Mm00495520_m1), SVCT2 (Assay ID: Mm00497751_m1), GLUT1 (Assay ID: Mm00441473_m1), GLUT3 (Assay ID: Mm00441483_m1) and GLUT4 (Assay ID: Mm00436615_m1) came from an inventory of TaqManÒ Gene Expression Assays (Applied Biosystems, Foster City, CA, USA). As an endogenous control gene, TaqManÒ Rodent GAPDH Control Reagents (Applied Biosystems) was used. The reactions were performed by using the RT-PCR equipment (Applied Biosystems 7300 Real Time PCR System). For quantitative analysis of SVCTs and GLUTs, a standard curve method was designed; that is, an aliquot from each experimental sample was used to generate standard curves. The expression levels in individual samples were then compared with levels from all tissues in common with any gene measurement. The correlation coefficient of the standard curve was more than 0.999. The mRNA levels of SVCTs and GLUTs were evaluated relative to mRNA levels of GAPDH, and the SVCT and GLUT expression level in the liver and small intestine of AA() WT mice was considered as 100%, respectively. Isolation and culture of mouse hepatocytes Mouse hepatocytes from AA() SMP30/GNL KO and WT mice at 63 days of age were isolated by the collagenase perfusion method as described previously [30]. Briefly, each liver was perfused in situ through the vena cava inferior with ethylene glycol-bis(2-aminoethylether)-N,N,N0 ,N0 -tetraacetic acid (EGTA) solution containing 0.5 mM EGTA, 5 mM glucose, 4.1 mM NaHCO3, 136 mM NaCl, 5.3 mM KCl, 0.3 mM Na2HPO4, 0.4 mM KH2PO4, and 10 mM Hepes, pH 7.2. Then, the solution was replaced with collagenase solution

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containing 0.03% collagenase, 4.8 mM CaCl2, 136 mM NaCl, 5.3 mM KCl, 0.3 mM Na2HPO4, 0.4 mM KH2PO4, 0.006% Trypsin inhibitor and 10 mM Hepes, pH 7.2. After the collagenase perfusion, the livers were removed and filtered through nylon mesh (100 lM) and then washed with Hanks’ Balanced Salt Solution to remove nonparenchymal cells. Final cell preparations were suspended at 2.5  105 cells/ml of defined minimum essential medium (DMEM) containing 0.4 lg/L dexamethasone, 0.1 lg/ml bovine pancreatic trypsin inhibitor, 2 mM L-glutamine supplemented with 5% fetal calf serum and then placed into culture plates coated with bovine type I collagen (Sigma–Aldrich, St. Louis, MO, USA). DMEM and fetal calf serum did not detect AA. Cells were cultured at 37 °C under 5% CO2 in air for 3 h to allow attachment to culture plates, after which the medium was replaced with serum-free DMEM. Cell viability was determined by trypan blue dye exclusion before plating. AA uptake into hepatocytes For determination of AA uptake into cells, we used primary mouse hepatocytes cultured for one day after isolation. Subsequently, cells were incubated with 100 lM AA in DMEM for 3, 6, 9, 12, 15, 20, 30, 45 and 60 min. At the appropriate time, cells were washed with phosphate buffered saline and collected with 5% metaphosphoric acid to measure the AA and DHA content. AA was measured by a HPLC–ECD method. The protein concentration was measured by Lowry’s method [31] using bovine serum albumin as a standard. Statistical analysis Results are expressed as means ± SEM. The probability of statistical differences between experimental groups was determined by

Student’s t-test or ANOVA as appropriate. One way ANOVAs were performed using Kareida Graph software (Synergy Software, Reading, PA, USA). Statistical differences were considered significant at p < 0.05. Results Comparative body weights After weaning at 30 days of age, SMP30/GNL KO and WT mice were divided into four groups: AA(+) SMP30/GNL KO, AA() SMP30/GNL KO, AA(+) WT, and AA() WT mice. To investigate, the effect of AA depletion on growth, we compared the body weights among these four groups. Until the experiment ended, when the mice were 63-days-old, there were no significantly different increases in body weights among the four groups (data not shown). At that time, the mean body weights of the AA(+) and AA() SMP30/GNL KO and WT mice were 24.9 ± 0.8, 25.3 ± 0.3, 25.0 ± 0.8 and 23.1 ± 0.6 g, respectively. AA and DHA levels in AA depleted mice To ensure AA depletion from AA() SMP30/GNL KO mice, we measured AA plus DHA contents in specific tissues compared to that in their AA(+) SMP30/GNL KO counterparts and both groups of WT mice at the age of 63 days. AA plus DHA contents in the cerebrum, liver, kidney and small intestine of AA() SMP30/GNL KO mice were 0.132 ± 0.018 lmol/g tissue, 0.147 ± 0.005 lmol/g tissue, 0.076 ± 0.002 lmol/g tissue and 0.112 ± 0.010 lmol/g tissue, respectively, constituting significant reductions of 96%, 88%, 91% and 97% when compared to AA() WT mice (Fig. 1). In the cerebrum and liver of AA(+) WT mice, values of total

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Fig. 1. AA and DHA contents in the cerebrum, liver, kidney and small intestine of AA(+) and AA() SMP30/GNL KO and WT mice at 63 days of age. After weaning at 30 days of age, the mice were divided into four groups: AA(+) SMP30/GNL KO, AA() SMP30/GNL KO, AA(+) WT and AA() WT. The AA(+) groups had free access to water containing 1.5 g/L AA and 10 lM EDTA, whereas the AA() groups had free access to water without AA until the experiment ended when the mice were 63-days-old. AA (black columns) and DHA (gray columns) contents of tissues were analyzed by HPLC–ECD. Values are expressed as means ± SEM (AA plus DHA) of five animals.  p < 0.01: AA() SMP30/GNL KO mice compared to AA(+) SMP30/GNL KO, AA(+) WT and AA() WT mice.

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150

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The effect of AA depletion on SVCTs and GLUTs mRNA expression To clarify the influence of vitamin C and glucose transporters SVCT and GLUT during AA depletion, we measured SVCT1, SVCT2, GLUT1, GLUT3 and GLUT4 mRNA expression levels in tissues from AA(+) and AA() SMP30/GNL KO and WT mice by using quantitative RT-PCR. SVCT1 mRNA expression was detected in the liver, kidney and small intestine but not in the cerebrum (Fig. 2A). The liver and small intestine of AA() SMP30/GNL KO mice expressed a significant 21% and 55% higher levels, respectively, than those of AA(+) SMP30/

SVCT1 expression level (%)

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SVCT1 expression level (%)

AA were almost the same as those of AA() WT mice; however, the kidneys and small intestines of AA(+) WT mice had a significant 27% and 22% larger amount of total AA plus DHA than AA() WT mice, respectively. Moreover, AA plus DHA levels in the cerebrum, liver, kidney and small intestine of AA(+) SMP30/GNL KO mice were similar to those of AA() WT mice. On the other hand, the DHA per AA plus DHA in the cerebrum, liver, kidney and small intestine of AA(+), AA() WT and AA(+) SMP30/GNL KO mice were 5.9–6.1%, 4.2–6.3%, 18.1–20.5% and 5.8–6.9%, respectively (Fig. 1).

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Fig. 2. SVCT1 and SVCT2 mRNA expression level in tissues of AA(+) and AA() SMP30/GNL KO and WT mice at 63 days of age. SVCT1 (A) and SVCT2 (B) mRNA expression level in tissues were measured by using quantitative RT-PCR as described in Materials and methods. The mRNA levels of SVCTs were evaluated relative to mRNA levels of GAPDH, and the SVCT expression level in the livers of AA() WT mice was considered as 100%. Data from quantitative RT-PCR represent a mean ± SEM of five animals.

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700 600

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among all four groups. GLUT3 mRNA was present with near equality in the cerebrum and small intestine of all four groups but was not found in the liver or kidney (Fig. 3B). Moreover, GLUT4 mRNA was expressed in the kidney and small intestine but not the cerebrum or liver (Fig. 3C). GLUT4 expression level in the small intestine of AA() SMP30/GNL KO mice was higher than that in AA(+) SMP30/GNL KO mice but not to a significant extent.

GLUT1 expression level (%)

A

GLUT1 expression level (%)

GNL KO mice. In contrast, SVCT2 mRNA expression was detected in the liver and kidney as well as the small intestine and cerebrum (Fig. 2B). The amount of SVCT2 in liver of AA() SMP30/GNL KO mice was 70–86% higher than in the other three groups. Moreover, SVCT2 expression level in small intestine of AA(+) WT mice was a significant 43% lower than that in AA() WT mice. GLUT1 mRNA expression was detected in the cerebrum, liver, kidney and small intestine (Fig. 3A) and at similar volumes

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Fig. 3. GLUT1, GLUT3 and GLUT4 mRNA expression in tissues of AA(+) and AA() SMP30/GNL KO and WT mice at 63 days of age. GLUT1 (A), GLUT3 (B) and GLUT4 (C) mRNA expression levels in tissues were measured by using a quantitative RT-PCR method. The mRNA levels of GLUTs were evaluated relative to mRNA levels of GAPDH, and the GLUT expression level in small intestines of AA() WT mice was considered as 100%. Data from quantitative RT-PCR represent a mean ± SEM of five animals.

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SMP30/GNL KO mice. On the other hand, although the latter’s DHA levels increased slightly by 60 min of culture, as did those of the WT mice, DHA levels in the hepatocytes of AA() SMP30/ GNL KO mice were a significant 147% higher than those of AA() WT mice at 60 min (Fig. 4).

AA ( nmol/ mg protein)

7 AA DHA AA DHA

6 5

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SVCT mRNA expression by mouse hepatocytes

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As is typical for the liver in vivo, hepatocytes from AA() SMP30/GNL KO and WT mice effectively expressed SVCT1 and SVCT2 mRNA. Notably, the levels of SVCT1 and SVCT2 expressed were significantly greater when the source was hepatocytes from AA() SMP30/GNL KO mice, i.e., 53% and 121% respectively, than those from AA() WT mice (Fig. 5).

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Time (min) Discussion Fig. 4. Time course of AA uptake by primary cultured mouse hepatocytes from AA() SMP30/GNL KO and WT mice. Cells were incubated with 100 lM AA in DMEM for 3, 6, 9, 12, 15, 20, 30, 45 and 60 min. At those time intervals, cells were collected with 5% metaphosphoric acid and measured for AA and DHA contents by HPLC–ECD. Values are expressed as means ± SEM of five animals.  p < 0.01 as compared to the AA content of WT mouse hepatocytes measured at the same time point. *p < 0.01 as compared to the DHA content of WT mouse hepatocytes measured at the same time point.

AA uptake by mouse hepatocytes To investigate more directly the effect of AA uptake into cells in an environment of AA depletion, we used primary cultured hepatocytes taken from AA() SMP30/GNL KO and WT mice when they were 63-days-old. In culture medium containing 100 lM AA, the cells’ uptake of AA was measured during a 60 min period. Hepatocytes from AA() SMP30/GNL KO mice took up AA at an almost linearly increasing rate of 0.14 nmol/mg protein/min for15 min, after which the rate decreased slightly to 0.07 nmol/mg protein/min for the next 60 min (Fig. 4). Initially, AA() WT mouse hepatocytes showed a similar linear increase of AA uptake at 0.13 nmol/mg protein/min that lasted for 15 min, but the rate then decreased decisively to 0.02 nmol/mg protein/min until the 60 min termination. At 45 and 60 min after AA supplementation, AA contents in the hepatocytes of AA() SMP30/GNL KO mice were a significant 51% and 80% higher than those of AA() WT mice, respectively. Since WT mice can synthesize AA in vivo, their hepatocytes contained 0.70 nmol/mg protein of AA at 0 min of culture. In contrast, no AA was detectable at 0 min in the hepatocytes from AA()

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Fig. 5. SVCT1 and SVCT2 mRNA expression in primary cultured hepatocytes from AA() SMP30/GNL KO and WT mice. SVCT1 (A) and SVCT2 (B) mRNA were measured by using quantitative RT-PCR. The mRNA levels of SVCTs were evaluated relative to mRNA levels of GAPDH, and the level SVCT expressed by hepatocytes from WT mice was considered as 100%. Data from quantitative RT-PCR represent a mean ± SEM of five animals.

The present study demonstrates that AA depletion enhanced SVCT1 and SVCT2 mRNA expression in the liver and SVCT1 and GLUT4 mRNA expression in the small intestine of SMP30/GNL KO mice, which were used for this purpose because they cannot synthesize AA in vivo [25]. Moreover, the ability of hepatocytes from AA-depleted SMP30/GNL KO mice to take up AA was far superior to that of hepatocytes from matched WT mice due to the former’s enhanced expression levels of SVCT1 and SVCT2. These results strongly indicate that intracellular AA content is an important regulatory element of SVCT1 and SVCT2 expression in the liver. Feeding SMP30/GNL KO mice with a diet lacking AA for 33 days after weaning caused them to develop a significant loss of AA and DHA, i.e., they contained less than 12% in the cerebrum, liver, kidney and small intestine of the content in WT mice. This low AA and DHA content did not affect their tissue-specific expression profiles of the AA transporters, SVCT1 or SVCT2, or the DHA transporters GLUT1, GLUT3 and GLUT4. In the cerebrum, SVCT2, GLUT1 and GLUT3 mRNAs were detected, but not significant difference in the amounts of these mRNAs expressed by AA depletion. Agus et al. [32] has reported that AA cannot pass the blood–brain barrier, although DHA can pass that barrier via transport by the GLUTs. Since AA was supplied to the cerebrum in the form of DHA, the expression level of SVCT2 might have been unaffected in cerebral tissues from AA-depleted SMP30/GNL KO mice. In the small intestine, SVCT and GLUT mRNA were detected, and SVCT1 and GLUT4 expression levels, but not those of SVCT2, were enhanced by AA depletion. Moreover, SVCT2 expression in AA(+) WT mice was significantly lower than in AA() WT mice. Possibly a state of AA excess in cells of the small intestine would affect their SVCT2 content, because the AA plus DHA level in small intestines from AA(+) WT mice was significantly higher than that from AA() WT mice (Fig. 1). In the kidney, SVCT1, SVCT2, GLUT1 and GLUT4 mRNAs were expressed at relatively similar levels despite AA depletion, although the AA plus DHA level in kidneys from AA(+) WT mice was significantly higher than that from AA() WT mice (Fig. 1). In the liver, SVCT1, SVCT2 and GLUT1 mRNAs were detected, with SVCT1 and SVCT2 present in AA-depleted SMP30/GNL KO mice in a significant 21% and 70% higher quantity, respectively, than in AAsufficient SMP30/GNL KO mice. AA absorbed from the small intestine was first transported to the liver via the hepatic portal vein and then distributed to multiple organs. Therefore, the liver must be far more sensitive to a state of AA deficiency than other organs. Reidling et al. [33] has also reported that SVCT2 expression is more susceptible to regulation at the RNA level in the human liver. Thus, the SVCT1 and especially SVCT2 mRNA expression level were enhanced in the liver and also in primary cultured hepatocytes from AA-depleted SMP30/GNL KO mice. Consequently, the AA uptake ability of hepatocytes from AA-depleted SMP30/GNL KO mice

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exceeded that of hepatocytes from WT mice. Although these results should be ideally confirmed at the SVCT1 and SVCT2 protein level, we were unable to demonstrate these results because of not cross-reactivity with these proteins by using commercially available antibodies. These same phenomenon were also reported by Michels et al. [34]. In conclusion, to the best of our knowledge here we found for the first time that AA depletion enhances SVCT1 and SVCT2 mRNA expression levels in the liver and strengthens AA uptake ability from outside to inside hepatic cells. Considering these overall results, intracellular AA content must be one of the major regulatory elements of SVCT1 and SVCT2 expression in the liver. Acknowledgments This study is supported by a Grant-in-Aid for Scientific Research from the Ministry of Education, Science, and Culture, Japan (to A.I.), and a Grant-in-Aid from the Smoking Research Foundation, Japan (to A.I.). We thank Ms. P. Minick for the excellent English editorial assistance. Vitamin C powder was kindly provided by DSM Nutrition Japan. References [1] R.C. Rose, A.M. Bode, FASEB J. 7 (1993) 1135–1142. [2] Y. Kondo, T. Sasaki, Y. Sato, A. Amano, S. Aizawa, M. Iwama, S. Handa, N. Shimada, M. Fukuda, M. Akita, J. Lee, K.S. Jeong, N. Maruyama, A. Ishigami, Biochem. Biophys. Res. Commun. 377 (2008) 291–296. [3] Y. Sato, S. Kajiyama, A. Amano, Y. Kondo, T. Sasaki, S. Handa, R. Takahashi, M. Fukui, G. Hasegawa, N. Nakamura, H. Fujinawa, T. Mori, M. Ohta, H. Obayashi, N. Maruyama, A. Ishigami, Biochem. Biophys. Res. Commun. 375 (2008) 346– 350. [4] D.J. Lane, A. Lawen, Free Radic. Biol. Med. 47 (2009) 485–495. [5] K. Hirota, G.L. Semenza, Biochem. Biophys. Res. Commun. 338 (2005) 610–616. [6] Y. Li, H.E. Schellhorn, J. Nutr. 137 (2007) 2171–2184. [7] A. Carr, B. Frei, FASEB J. 13 (1999) 1007–1024. [8] D. Su, J.M. May, M.J. Koury, H. Asard, J. Biol. Chem. 281 (2006) 39852–39859. [9] J. Riviere, J.L. Ravanat, J.R. Wagner, Free Radic. Biol. Med. 40 (2006) 2071–2079.

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