Advertisement
Browse Subject Areas
?

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Ascorbic Acid Biosynthesis and Brackish Water Acclimation in the Euryhaline Freshwater White-Rimmed Stingray, Himantura signifer

  • Samuel Z. H. Wong,

    Affiliation Department of Biological Sciences, National University of Singapore, Kent Ridge, Singapore, Republic of Singapore

  • Biyun Ching,

    Affiliation Department of Biological Sciences, National University of Singapore, Kent Ridge, Singapore, Republic of Singapore

  • You R. Chng,

    Affiliation Department of Biological Sciences, National University of Singapore, Kent Ridge, Singapore, Republic of Singapore

  • Wai P. Wong,

    Affiliation Department of Biological Sciences, National University of Singapore, Kent Ridge, Singapore, Republic of Singapore

  • Shit F. Chew,

    Affiliation Natural Sciences and Science Education, National Institute of Education, Nanyang Technological University, Singapore, Republic of Singapore

  • Yuen K. Ip

    dbsipyk@nus.edu.sg

    Affiliation Department of Biological Sciences, National University of Singapore, Kent Ridge, Singapore, Republic of Singapore

Ascorbic Acid Biosynthesis and Brackish Water Acclimation in the Euryhaline Freshwater White-Rimmed Stingray, Himantura signifer

  • Samuel Z. H. Wong, 
  • Biyun Ching, 
  • You R. Chng, 
  • Wai P. Wong, 
  • Shit F. Chew, 
  • Yuen K. Ip
PLOS
x

Abstract

L-gulono-γ-lactone oxidase (Gulo) catalyzes the last step of ascorbic acid biosynthesis, which occurs in the kidney of elasmobranchs. This study aimed to clone and sequence gulonolactone oxidase (gulo) from the kidney of the euryhaline freshwater stingray, Himantura signifer, and to determine the effects of acclimation from freshwater to brackish water (salinity 20) on its renal gulo mRNA expression and Gulo activity. We also examined the effects of brackish water acclimation on concentrations of ascorbate, dehydroascorbate and ascorbate + dehydroascorbate in the kidney, brain and gill. The complete cDNA coding sequence of gulo from the kidney of H. signifer contained 1323 bp coding for 440 amino acids. The expression of gulo was kidney-specific, and renal gulo expression decreased significantly by 67% and 50% in fish acclimated to brackish water for 1 day and 6 days, respectively. There was also a significant decrease in renal Gulo activity after 6 days of acclimation to brackish water. Hence, brackish water acclimation led to a decrease in the ascorbic acid synthetic capacity in the kidney of H. signifer. However, there were significant increases in concentrations of ascorbate and ascorbate + dehydroascorbate in the gills (after 1 or 6 days), and a significant increase in the concentration of ascorbate and a significant decrease in the concentration of dehydroascorbate in the brain (after 1 day) of fish acclimated to brackish water. Taken together, our results indicate that H. signifer might experience greater salinity-induced oxidative stress in freshwater than in brackish water, possibly related to its short history of freshwater invasion. These results also suggest for the first time a possible relationship between the successful invasion of the freshwater environment by some euryhaline marine elasmobranchs and the ability of these elasmobranchs to increase the capacity of ascorbic acid synthesis in response to hyposalinity stress.

Introduction

Ascorbic acid is water-soluble and has a molecular mass of 176.1. At physiological pH, ascorbic acid is present mainly as the ascorbate anion [1]. Ascorbate functions mainly as a water-soluble antioxidant and a cofactor of enzymes involved in biosynthetic reactions [2]. Physiologically, ascorbate acts as a component of the intracellular antioxidant system which includes low molecular mass substances such as tocopherol and glutathione, and antioxidant enzymes such as superoxide dismutase and glutathione peroxidase [3]. Ascorbate is oxidized (one-electron transfer) to the ascorbyl free radical which can be further oxidized with the loss of a second electron to dehydroascorbate [4]. As an antioxidant, ascorbate protects tissues from oxidative damages by reacting with and inactivating radicals and oxidants such as superoxide anion, peroxyl- and hydroxyl-radicals, singlet oxygen, and hypochlorous acid [5], [6], [7], [8]. Moreover, ascorbate can regenerate α-tocopherol from α-tocopheryl radical in membranes [9].

Biosynthesis of ascorbic acid from glucose occurs in kidney of the majority of cold-blooded vertebrates and some birds, liver of some birds and the majority of mammals, and both kidney and liver of some birds and some marsupials. L-gulono-γ-lactone oxidase (GULO/Gulo) is the enzyme required for the last step of ascorbic acid biosynthesis, and it is expressed in kidney of elasmobranchs, some non-teleost fishes, amphibians, reptiles, non-passerine birds and monotremes, kidney and liver of some passerine birds and marsupials, and liver of some birds, some marsupials and most placental mammals [10]. Fishes like elasmobranchs, lampreys, sturgeons and lungfishes express Gulo and synthesize ascorbic acid in the kidney [10]. In general, Gulo/GULO is a microsomal enzyme that catalyzes the aerobic conversion of gulonolactone to ascorbate, with the production of hydrogen peroxide [11], [12]. Invertebrates, teleost fishes, some passerine birds, guinea pigs, bats and anthropoid primates, including humans, lack functional Gulo/GULO [13], [14], and therefore need a dietary intake of ascorbic acid as vitamin C. Although the evolution of ascorbic acid biosynthesis and the related kidney/liver transition have attracted great attention in the past, there is a paucity of molecular information on gulo from elasmobranchs, stingrays in particular [15]. Furthermore, it is unclear why elasmobranchs synthesize ascorbic acid, and there is a dearth of knowledge on effects of changes in environmental conditions (e.g. salinity changes) on gulo expression in elasmobranchs.

One of the main physiological functions of ascorbate is to maintain cellular homeostasis during stress [2]. Any environmental disturbance can be considered a potential source of stress, because the animal has to deal with the physiological changes triggered by changes in the environment. As has been demonstrated in turtle [16], [17], the greater the stress an animal undergoes, the more the ascorbic acid it has to produce or obtain from the diet. Salinity changes can cause stress; therefore, aquatic organisms have to deal with salinity stress, and ascorbate can be an important anti-stress agent. The branchial ascorbate concentration of the euryhaline mud crab, Scylla serrata, decreases with an increase in salinity indicating a possible role of ascorbate in combating stress in the gills [18]. Similarly, for the freshwater Philippine catfish, Clarias batrachus, which lacks the ability to synthesize ascorbic acid, ascorbate concentrations in the liver, kidney, brain and muscle decrease with increasing salinity [19]. More importantly, due to an increase in energy demand for osmoregulation, acclimation to different salinities can result in increased metabolic rate, which can in turn lead to increased production of reactive oxygen species. Indeed, it has been established that salinity changes result in oxidative stress in sea bass [20] and sturgeon [21]. However, at present, there is a dearth of knowledge on the effects of salinity changes on the concentrations of ascorbate, and its oxidative product, dehydroascorbate, in various organs/tissues of elasmobranchs which can synthesize ascorbic acid in kidney.

The majority of elasmobranchs are marine, and there are few stenohaline freshwater elasmobranchs. For a predominantly marine group with rare lineages that have successfully invaded and fully adapted to freshwater, euryhalinity is a relatively uncommon feature of elasmobranchs (as compared with teleosts), and there are only several notable examples such as Carcharhinus leucas and some populations of Dasyatis sabina [22]. The freshwater Asian white-rimmed stingray, Himantura signifer (Compagno and Roberts 1982), belongs to Family Dasyatidae, and can be found in the Batang Hari Basin in Jambi of Sumatra in Indonesia. Although H. signifer can be found in freshwater, it may re-enter brackish/estuarine and marine environments for reproduction [23]. Unlike the non-ureogenic stenohaline freshwater Amazonian stingray, Potamotrygon motoro [24], H. signifer is ureogenic and undergoes ureosmotic osmoregulation in brackish water [24]. Thus, H. signifer has to suppress urea synthesis and retention in freshwater, but it cannot stop urea synthesis completely. As a result, the plasma and tissues of freshwater H. signifer contain higher concentrations of urea than other freshwater fishes [24]. Because of this, H. signifer could be exposed to greater salinity stress in freshwater than in brackish water. Since salinity stress is known to induce oxidative stress in fish [18], [19], H. signifer could be exposed to greater salinity-induced oxidative stress in freshwater than in brackish water. Furthermore, since H. signifer is nitrogen-limited, and conserves dietary nitrogen as urea for osmoregulatory purposes [25], it is possible that it would be confronted with postprandial osmotic stress resulting from increased urea synthesis upon feeding in freshwater.

Therefore, this study was undertaken to obtain the full cDNA sequence of gulo from the kidney of H. signifer, and to determine its mRNA expression in the kidney of fish kept in freshwater (control) or exposed to brackish water (salinity 20) for 1 or 6 days using quantitative RT-PCR. Since H. signifer could be exposed to decreased salinity stress in brackish water, we aimed to test the hypothesis that exposure to brackish water would result in a down-regulation of its renal gulo expression. In order to confirm that salinity changes indeed affected ascorbic acid synthesis in H. signifer, efforts were made to determine the renal Gulo activity and the concentrations of ascorbate and dehydroascorbate in the kidney, brain and gills of H. signifer exposed to freshwater or brackish water. The hypothesis tested was that, despite a decrease in renal Gulo activity, acclimation from freshwater to brackish water would lead to increases and decreases in the concentrations of ascorbate and dehydroascorbate, respectively, in some organs of H. signifer, reflecting the redox status due to a decrease in salinity-induced oxidative stress in brackish water.

Materials and Methods

Ethics statement

This study was approved by the Institutional Animal Care and Use Committee of the National University of Singapore (IACUC 021/10). In Singapore, no specific permission from an authority is required for collection of seawater. The current study is not an ecological/field study, and no animals were collected from the field.

Animals

Specimens of H. signifer were purchased from a local fish farm. Fish were kept in dechlorinated tap water (freshwater; pH 6.8–7.0) at 25°C in plastic tanks of appropriate sizes with aeration under a 12 h light∶12 h dark regime for at least 1 week before experiments. They were fed live shrimps daily.

Experimental conditions and collection of samples

Control fish (N = 5) were immersed in 25 volumes (v/w) of freshwater in plastic tanks, and they served as controls for the two experimental conditions: exposure to brackish water (salinity 20; pH 7.8) for 1 or 6 days. For exposure to salinity changes, fish (N = 5) were transferred from freshwater (day 0) to waters of salinity 5 on day 1, salinity 10 on day 2, salinity 15 on day 3, salinity 20 on day 4 and kept in salinity 20 for 1 or 6 days. Water was changed daily. Natural seawater was collected from the sea at least 1 km away from the coast of the Singapore main island. Waters of different salinities were prepared by mixing seawater with an appropriate quantity of freshwater. Salinity was monitored using a YSI Model 30/10 FT salinometer (Yellow Springs Instrument Co. Inc, Ohio, USA). During salinity acclimation, stingrays were fed live shrimps on alternate days. Both control and experimental fish were killed by an overdose of neutralized 0.05% MS222, and their tissues quickly excised, frozen in liquid nitrogen and stored at −80°C until analysis.

Total RNA extraction and cDNA synthesis

Total RNA was isolated from kidney samples of H. signifer using TRI REAGENT™ protocol and purified using the Qiagen RNeasy Mini Kit (Qiagen GmbH, Hilden, Germany). The RNA was quantified spectrophotometrically using Hellma TrayCell (Hellma GmbH & Co. KG, Müllheim, Germany) and its integrity checked electrophoretically to verify RNA by comparing the 18S and the 28S bands, which were visualized by a G:Box gel documentation system (Syngene, Cambridge, UK). Total RNA (1 µg) isolated was reverse transcribed into cDNA using RevertAid™ First Strand cDNA synthesis kit (Fermentas International, Inc, Burlington, ON, Canada).

Polymerase Chain Reaction (PCR)

The complete gulo sequence was obtained using primers (Forward: 5′- TYC TCM AGG TGG AYM WGG AGA -3′; Reverse: 5′- TCR STG TGW GGR AAC CAG A -3′) designed from the conserved regions of Triakis scyllium (ABO15547.1), Mustelus manazo (ABO15548.1), Scyliorhinus torazame (Q90YK3.1), Acipenser transmontanus (ABO15549.1), Rattus norvegicus (NM_022220), Mus musculus (AY453064) and Sus scrofa (AF440259), as reported in Cho et al. [15]. PCR was carried out in Biorad Peltier thermal cycler (Biorad, Hercules, CA, USA) using DreamTaq™ DNA polymerase (Fermentas International Inc.). The cycling conditions were 94°C (3 min), followed by 35 cycles of 94°C (30 s), 55°C (30 s), 72°C (2 min) and 1 cycle of final extension at 72°C (10 min). PCR products were electrophoresed in 1% agarose gel. Bands of the estimated size were extracted from the gels using QIAquick® Gel Extraction Kit (Qiagen GmbH). Purified PCR products were subjected to cycle sequencing using BigDye® Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA) and purified by ethanol/sodium acetate precipitation. Purified products were automatically sequenced using the 3130XL Genetic Analyzer (Applied Biosystems). The fragments were verified to be gulo from Genbank database.

Rapid amplification of cDNA ends (RACE)-PCR

Total RNA (1 µg) was reverse transcribed into 5′-RACE-Ready cDNA and 3′RACE-Ready cDNA using SMARTer™ RACE cDNA Amplification kit (Clontech Laboratories, Mountain View, CA, USA). RACE-PCR was performed using the Advantage® 2 PCR kit (Clontech Laboratories) to generate the 5′ and 3′ cDNA fragments, with 5′- GAA TGC TCG GAG TCA GTA AAC CGG G -3′ and 5′- GGT TCT AAC CGT CAC CAT CCA GT -3′, respectively. RACE-PCR cycling conditions were 25 cycles of 94°C for 30 s, 65°C for 30 s and 72°C for 4 min. RACE-PCR products were separated using gel electrophoresis, purified and sequenced. Multiple sequencing was performed in both directions to obtain the full-length cDNA. Sequence assembly and analysis were performed using BioEdit [26]. The cDNA sequence of gulo from H. signifer has been deposited to the Genbank with accession number (KC465465).

The Gulo amino acid sequence was translated from the nucleotide sequence of gulo using ExPASy Proteomic server [27]. The potential phosphorylation, O-GlcNAcylation and N-GlcNAcylation sites were predicted using NetPhos 2.0 [28], YinOYang 1.2 [29], [30] and NetNglyc 1.0 [31], respectively. The transmembrane regions (TMs) were predicted using MEMSATS & MEMSAT-SVA provided by PSIPRED protein structure prediction server (http://bioinf.cs.ucl.ac.uk/psipred/) [32].

Phylogenetic analysis

Amino acid sequences of Gulo/GULO from other animals were obtained from Genbank with the following accession numbers: Triakis scyllium (ABO15547.1), Mustelus manazo (ABO15548.1), Scyliorhinus torazame (Q90YK3.1), Acipenser transmontanus (ABO15549.1), Raja kenojei (EF397523.1), Dasyatis akajei (EF397524.1), Xenopus laevis (NM_001095065), Gallus gallus (XM_001234313), Meleagris gallopavo (XM_003204567.1), Pelodiscus sinensis (AET14634.1), Bos taurus (Q3ZC33.3), Sus scrofa (NP_001123420.1), Hipposideros armiger (ADP88814.1), Rousettus leschenaultii (ADP88813.1), Rattus norvegicus (P10867.3), Mus musculus (P58710.3), Ornithorhynchus anatinus (XM_001521551), and Lysinibacillus sphaericus (YP_001699013.1). These sequences were aligned using ClustalX2 and phylogenetic analysis was performed using neighbor-joining method and 100 bootstrap replicates with Phylip [33].

Tissue expression

The mRNA expression of gulo was performed on twelve different organs/tissues, including brain, heart, eye, spleen, stomach, intestine, kidney, gills, skin, liver and muscle. PCR was carried out in Biorad Peltier thermal cycler (Biorad, Hercules) using DreamTaq™ DNA polymerase (Fermentas International Inc.) and using forward primer 5′- TYC TCM AGG TGG AYM WGG AGA -3′ and reverse primer 5′- TCR STG TGW GGR AAC CAG A -3′. The cycling conditions consist of 95°C (3 min), followed by 30 cycles of 94°C (30 s), 60°C (30 s), 72°C (30 s) and 1 cycle of final extension at 72°C (10 min). PCR products were then electrophoresed in 2% agarose gel.

qPCR

RNA from kidney samples were treated with Deoxyribonuclease I, (Sigma–Aldrich Co., St Louis, MO, USA) to remove any contaminating genomic DNA. First strand cDNA was then synthesized from 1 µg of total RNA using random hexamer primer and the RevertAid™ first stand cDNA synthesis kit (Fermentas International Inc.). qPCR was performed in triplicates using a StepOnePlus™ Real-Time PCR System (Applied Biosystems). The standard cDNA (template) was serially diluted in sterile water (from 107 to 102 specific copies/2 µl). The PCR reactions contained 5 µl of 2× Fast SYBR® Green Master Mix (Applied Biosystems), 0.1 µM of forward (5′- CCT CAC ACG GAC AAG ACA G -3′) or reverse (5′- CTG GAA CTG CTG TTG TGG A -3′) qPCR primers, and cDNA (equivalent to 1 ng of RNA) or standard (2 µl) in a total volume of 10 µl. Cycling conditions were 95°C for 20 s (1 cycle), followed by 45 cycles of 95°C for 3 s and 60°C of 30 s. Data (CT values) were collected at each elongation step. Runs were followed by melt curve analysis by increasing from 60°C to 95°C in 0.3°C increments to confirm the presence of only a single product. The PCR products were separated in a 2% agarose gel to verify the presence of a single band.

In order to determine the absolute quantity of gulo transcripts in a qPCR reaction, efforts were made to produce a pure amplicon of a defined region of gulo cDNA from the kidney of H. signifer following the methods of Gerwick et al. [34]. PCR was performed with qPCR primers and cDNA as a template in a final volume of 25 µl with the following cycling conditions: initial denaturation 95°C for 3 min, followed by 35 cycles of 95°C for 30 s, 60°C for 30 s and 72°C for 30 s and 1 cycle of final extension of 72°C for 10 min. PCR product was separated in a 2% agarose gel. The product was excised and purified using QIAquick gel extraction kit (Qiagen GmbH). The gulo nucleotide fragment in the purified product was cloned using pGEM®-T Easy vector (Promega Corporation, Madison, WI, USA). The presence of the insert in the recombinant clones was confirmed by sequencing. The cloned circular plasmid was quantified using a spectrophotometer. Copy numbers were calculated from the CT values of the standards [33]. A standard curve was obtained from plotting threshold cycle (CT) on the Y-axis and the natural log of concentration (copies µl−1) on the X-axis. The CT, slope, PCR efficiency, Y-intercept and correlation coefficient (R2) were calculated using the default setting of StepOne™ Software v2.1 (Applied Biosystems). Diluted standards were stored at −20°C. The PCR efficiency for gulo was 91.6%. The quantity of transcript in an unknown sample was determined from the linear regression line derived from the standard curve and the number of copies per ng cDNA.

Determination of specific activity of Gulo

Specific activity of Gulo from the kidney of H. signifer was measured using a modification of the direct spectrophotometric assay described by Dabrowski [35]. Kidney samples from H. signifer were homogenized in 5 volumes (w/v) of homogenization buffer containing 50 mM NaHPO4 buffer (pH 7.4), 0.2% Na-deoxycholate and 1 mM EDTA three times using an Ultra-Turrax T25 homogenizer (Ika®-Labortechnik, Staufen, Germany) at 24,000 r.p.m. for 20 s each with 10 s intervals. Homogenates were centrifuged at 10,000×g for 30 min at 4°C to obtain the supernatants, which were used for determination of specific activity of Gulo at 25°C. The assay medium contained 100 mM L-gulonolactone, 50 mM reduced glutathione in 50 mM NaHPO4 buffer (pH 7.4). The reactions were stopped after 0 or 2 h by addition of ice-cold 10% metaphosphoric acid to the reaction mixtures. The reaction mixtures were then centrifuged at 10,000×g for 15 min at 4°C to obtain the supernatants, which were used directly for determination of concentrations of (ascorbate + dehydroascorbate) using a commercial Vitamin C assay kit (Cosmo Bio Co., Ltd., Tokyo, Japan). Specific activities of Gulo were expressed as µg ascorbic acid formed h−1 g−1 wet mass or µg ascorbic acid formed h−1 mg−1 protein.

Determination of concentrations of ascorbate and dehydroascorbate

Samples of the brain, kidney and gills of H. signifer were homogenized in 13 volumes (w/v) of ice-cold 5.4% metaphosphoric acid three times using an Ultra-Turrax T25 homogenizer (Ika®-Labortechnik) at 24,000 r.p.m. for 20 s each with 10 s intervals. The homogenates were centrifuged at 10,000×g for 30 min at 4°C to obtain the supernatants, which were used directly for the determination of concentrations of (ascorbate + dehydroascorbate), and dehydroascorbate alone, using a commercial Vitamin C Assay Kit (Cosmo Bio Co., Ltd., Tokyo, Japan). Total (ascorbate + dehydroascorbate) measurement involved the addition of an oxidizing reagent to the reaction, according to the manufacturer's protocol, while measurement of dehydroascorbate excludes this step. L-ascorbic acid solution was freshly prepared, using 5% metaphosphoric acid, as a standard for comparison. The standard curve was linear in the range of 0.5 to 4 µg of ascorbic acid. Ascorbate concentrations were calculated from the differences between concentrations of ascorbate + dehydroascorbate and dehydroascorbate. Concentrations were calculated and expressed in µg g−1 wet tissue mass. Preliminary results indicated that the muscle ascorbate + dehydroascorbate concentration was close to the detection limit of the assay method and therefore not investigated in this study.

Determination of thiobarbituric acid reactive substances

Thiobarbituric acid reactive substances were quantified as an index of lipid peroxidation [36] to elucidate the effect of brackish water acclimation on the redox-status of the gills and liver of H. signifer. These two organs were chosen because gills are in direct contact with the external medium and affected instantly by salinity changes, and the liver is the site of increased urea synthesis through the ornithine-urea cycle in response to brackish water acclimation. The frozen sample was homogenized three times, in 20 volumes (w/v) of ice-cold 1.1% phosphoric acid, for 20 s each with 10 s intervals using an Ultra-Turrax T25 homogenizer (Ika®-Labortechnik) at 24,000 r.p.m. Then, 0.4 ml of the homogenate was added to an equal volume of a mixture of 1% thiobarbituric acid, 0.1 mmol l−1 butylated hydroxytoluene solution, 50 mmol l−1 sodium hydroxide, and 0.2 ml of 7% phosphoric acid. After heating for 15 min at 98°C, the sample was vigorously mixed with 1.5 ml of butanol, and centrifuged at 2000×g for 5 min using a Beckman J2-21/E centrifuge (Beckman Coulter Inc., Fullerton, CA, USA). The organic layer was transferred to glass cuvettes, and the optical densities at 532 nm and 600 nm were determined using a Shimadzu UV-160A spectrophotometer (Shimadzu, Kyoto, Japan). Blanks were prepared by replacing the thiobarbituric acid with 3 mmol l−1 hydrochloric acid. The results were calculated as Sample (A532–A600) – Blank (A532–A600) as recommended by Ramos and Hermes-Lima [37]. TBARS values were obtained by using the extinction coefficient of 156 l mmol−1 cm−1.

Statistical analysis

Results are presented as means ± standard errors of the mean (S. E. M.). Differences between means are evaluated using one-way analysis of variance (ANOVA), followed by the Tukey post-hoc test for liver and kidney data, and the Dunnett T3 post-hoc test for gill. Differences were regarded as statistically significant at P<0.05.

Results

Nucleotide sequences, translated amino acid sequences and phylogenetic analysis

The complete coding cDNA sequence of gulo from the kidney of H. signifer consisted of 1323 bp (Genbank accession number KC465465), which coded for 440 amino acids (Figure 1) with a calculated molecular mass of 50.8 kDa. The deduced amino acid sequence consisted of two conserved domains, the flavin adenine dinucleotide (FAD)-binding domain at positions 21–156 and the D-arabinono-1,4-lactone oxidase (ALO) domain at positions 180–438 (Figure 2). The percentage similarities between the amino acid sequence of Gulo from H. signifer and those from other animals ranged between 66.3 and 75.9 for fish species, and 46.2 and 67.5 for non-fish species (Table 1). An analysis of the number of phosphorylation and glycosylation sites of H. signifer compared with various animals revealed 9 phosphorylation and 5 O-GlcNAcylation sites (Table 2). In addition, a phylogenetic analysis confirmed that Gulo of H. signifer was closely related to those of other elasmobranchs (Figure 3).

thumbnail
Figure 1. The cDNA sequence (GenBank accession number KC465465) of gulonolactone oxidase (gulo) and the deduced amino acid sequence of gulonolactone oxidase (Gulo) from the kidney of Himantura signifer.

https://doi.org/10.1371/journal.pone.0066691.g001

thumbnail
Figure 2. Multiple amino acid alignment of the deduced amino acid sequence of gulonolactone oxidase (Gulo) from the kidney of Himantura signifer, with eight other known sequences of Gulo from GenBank; Scyliorhinus torazame (cloudy catshark Gulo; Q90YK3.1), Triakis scyllium (banded houndshark Gulo; ABO15547.1), Mustelus manazo (starspotted smooth-hound Gulo; ABO15548.1), Acipenser transmontanus (sturgeon Gulo; ABO15549.1), Xenopus laevis (frog GULO; NM_001095065), Pelodiscus sinensis (turtle GULO; AET14634.1), Gallus gallus (chicken GULO; XM_001234313), and Mus musculus (mouse GULO; P58710.3).

Identical amino acids are indicated by shaded residues. The FAD-binding and ALO domains are underlined in bold and dotted lines respectively. P denotes potential phosphorylation sites. O denotes potential O-GlcNAcylation sites. The predicted transmembrane domains are underlined. The transmembrane domains (TM) of Gulo of H. signifer were predicted using MEMSATS & MEMSAT-SVA provided by PSIPRED protein structure prediction server.

https://doi.org/10.1371/journal.pone.0066691.g002

thumbnail
Figure 3. A phylogenetic tree that illustrates the relationship between the deduced amino acid sequence of gulonolactone oxidase (Gulo) of Himantura signifer and those of other animals with that of Lysinibacillus sphaericus as the outgroup.

https://doi.org/10.1371/journal.pone.0066691.g003

thumbnail
Table 1. The percentage similarity between the deduced amino acid sequence of gulonolactone oxidase (Gulo) from the kidney of Himantura signifer and those of other animal species obtained from Genbank.

https://doi.org/10.1371/journal.pone.0066691.t001

thumbnail
Table 2. Number of phosphorylation, O-GlcNAcylation and N-GlcNAcylation sites in the deduced amino acid sequence of gulonolactone oxidaseof various animals compared with Himantura signifer.

https://doi.org/10.1371/journal.pone.0066691.t002

A close examination of the alignment of Gulo from H. signifer with those from three species of shark, sturgeon, frog, turtle, chicken and mouse revealed that a considerable number of amino acid residues were unique to fish (Figure 2). For example, all the fish species including H. signifer have Gln3, Gln35, Glu38, Ala381 and Glu385, but all non-fish species have correspondingly Leu/His3, Glu35, Asp/Ala38, Val/Pro/Leu381 and Asn/Ser/Gly/Thr385 instead. Furthermore, all the fish Gulo sequences have Phe at C-terminus while non-fish sequences have Tyr. In addition, certain characters, which include Cys52, Tyr65, Asp225, Lys226, Thr241 and Ser307, were unique to all the four elasmobranch species compared to non-elasmobranchs (correspondingly Gly52, Phe65, Glu225, Asn226, Ser241 and Gln307). Apart from these shared features, H. signifer showed considerable unique amino acid residues in the conserved regions not found in the Gulo/GULO sequences of other vertebrates. Significant amino acid replacements might include Ser28, Lys42, Gln76, Ile103 in the FAD-binding domain and Ala185, Leu319, Glu325, Lys382 in the ALO domain.

Tissue expression of gulo

The mRNA expression of gulo in H. signifer was kidney-specific; it was not detectable in the brain, heart, eye, spleen, stomach, intestine, gills, skin, liver and muscle of fish kept in freshwater (Figure 4).

thumbnail
Figure 4. mRNA expression of gulonolactone oxidase (gulo) from the brain, heart, eye, spleen, stomach, intestine, kidney, gills, skin, liver and muscle of Himantura signifer kept in freshwater.

https://doi.org/10.1371/journal.pone.0066691.g004

mRNA expression of gulo from the kidney of fish exposed to brackish water

There were significant decreases in the mRNA expression of gulo in the kidney of H. signifer after 1 day (by 67%) or 6 days (by 50%) of exposure to brackish water as compared to the freshwater control (Figure 5).

thumbnail
Figure 5. Absolute quantification (copies of transcript per ng cDNA) of mRNA expression of gulonolactone oxidase (gulo) from the kidney of Himantura signifer kept in freshwater (FW; control) or exposed to brackish water (salinity 20) for 1 or 6 days after a 4-day progressive increase in salinity.

Results represent mean ± S. E. M. (N = 5). Means not sharing the same letter are significantly different (P<0.05).

https://doi.org/10.1371/journal.pone.0066691.g005

Specific activity of Gulo from the kidney of fish exposed to brackish water

The specific activity (µg ascorbic acid formed h−1 g−1) of Gulo from the kidney of H. signifer kept in freshwater was 125±12.9 (Figure 6). Although the renal Gulo activity was unaffected by 1 day of exposure to brackish water, it decreased significantly in fish exposed to brackish water for 6 days.

thumbnail
Figure 6. Specific activity of gulonolactone oxidase (Gulo; µg ascorbic acid formed h−1 g−1) from the kidney of Himantura signifer kept in freshwater (FW) on day 0 (control), or exposed to brackish water (salinity 20) for 1 or 6 days after a 4-day progressive increase in salinity.

Results represent mean ± S. E. M. (N = 4). Means not sharing the same letter are significantly different (P<0.05).

https://doi.org/10.1371/journal.pone.0066691.g006

Ascorbate and dehydroascorbate concentrations in the kidney, brain and gills of fish exposed to brackish water

The concentrations (µg g−1 wet mass) of ascorbate, dehydroascorbate and ascorbate + dehydroascorbate in the kidney of H. signifer kept in freshwater were 25.8±4.32, 17.3±5.45 and 43.1±6.31, respectively (Figure 7), and they remained unchanged in fish exposed to brackish water for 1 or 6 days.

thumbnail
Figure 7. Concentrations (µg g−1 wet mass) of ascorbate (AA; ▪), dehydroascorbate (DA; ▪) or AA+DA (TAA; □) in the kidney of Himantura signifier kept in freshwater (FW) on day 0 (control), or exposed to brackish water (salinity 20) for 1 or 6 days after a 4-day progressive increase in salinity.

There was no statistical difference between different groups. Results represent mean ± S. E. M. (N = 4).

https://doi.org/10.1371/journal.pone.0066691.g007

The concentrations (µg g−1 wet mass) of ascorbate, dehydroascorbate and ascorbate + dehydroascorbate in the brain of H. signifer kept in freshwater were the highest among the three organs studied (203±4.07, 19.2±1.94 and 222±3.86, respectively; Figure 8). For fish exposed to brackish water for 1 day, there were a significant increase in the ascorbate concentration (by 1.1-fold) and a significant decrease in the dehydroascorbate concentration (by 62%) in the brain.

thumbnail
Figure 8. Concentrations (µg g−1 wet mass) of ascorbate (AA; ▪), dehydroascorbate (DA; ▪) or AA+DA (TAA; □) in the brain of Himantura signifer kept in freshwater (FW) on day 0 (control), or exposed to brackish water (salinity 20) for 1 or 6 days after a 4-day progressive increase in salinity.

Results represent mean ± S. E. M. (N = 4). Means not sharing the same letter are significantly different (P<0.05).

https://doi.org/10.1371/journal.pone.0066691.g008

The concentrations (µg g−1 wet mass) of ascorbate, dehydroascorbate and ascorbate + dehydroascorbate in the gills of H. signifer kept in freshwater were the lowest among the three organs studied (7.40±1.42, 7.61±3.10 and 15.0±2.36, respectively; Figure 9). After 1 day of exposure to brackish water, there was a significant increase in ascorbate concentration (by 3.1-fold), and in ascorbate + dehydroascorbate concentration (by 2.5-fold) in the gills. For fish exposed to brackish water for 6 days, concentrations of ascorbate and ascorbate + dehydroascorbate increased significantly by 4.6-fold and 3.3-fold, respectively.

thumbnail
Figure 9. Concentrations (µg g−1 wet mass) of ascorbate (AA; ▪), dehydroascorbate (DA; ▪) or AA+DA (TAA; □) in the gills of Himantura signifer kept in freshwater (FW) on day 0 (control), or exposed to brackish water (salinity 20) for 1 or 6 days after a 4-day progressive increase in salinity.

Results represent mean ± S. E. M. (N = 4). Means not sharing the same letter are significantly different (P<0.05).

https://doi.org/10.1371/journal.pone.0066691.g009

Products of lipid peroxidation in the gills and liver of fish exposed to brackish water

The concentrations (nmol g−1 wet mass ±S. E. M.; N = 4) of thiobarbituric acid reactive substances in the liver of H. signifer exposed to brackish water for 1 day (33.9±7.72) or 6 days (45.5±4.79) were not significantly different from that of the freshwater control (42.7±2.23). Similarly, the concentrations of thiobarbituric acid reactive substances in the gills of fish exposed to brackish water for 1 day (18.9±1.0) or 6 days (14.0±0.5) were not significantly different from that of the freshwater control (14.0±1.8).

Discussion

Molecular characterization of gulo/Gulo from the kidney of H. signifer

Similar to other elasmobranchs, e.g. Raja kenojei and Dasyatis akajei [15], gulo was expressed only in the kidney of H. signifer. The cDNA sequence of gulo from the kidney of H. signifer coded for a protein of 440 amino acids, and the molecular mass of Gulo (50.8 kDa) from H. signifer was comparable to that (50.96 kDa) from Scyliorhinus torazame [38]. In general, Gulo/GULO amino acid sequences contain several strongly hydrophobic regions which may be associated with the endoplasmic reticulum membrane [39]. Indeed, Gulo/GULO are concentrated in the microsomal fraction during cellular fractionation, indicating that they are membrane-bound [40], [41]. Similarly, the Gulo from H. signifer comprised two transmembrane regions, indicating that it is an integral membrane protein.

The deduced amino acid sequence of Gulo from H. signifer contained two conserved domains, the FAD-binding domain at positions 21–156 and the ALO domain at positions 180–428. Since Gulo/GULO is an oxygen-dependent oxidoreductase using FAD as a co-factor, the FAD binding domain includes a covalent FAD-binding site, with a conserved His54 residue to which FAD binds via an 8-alpha-(N3-histidyl)-riboflavin linkage [41]. During the catalytic reaction, flavin first undergoes reduction by accepting two electrons from the substrate L-gulono-γ-lactone, followed with oxidation by O2 which presumably involves the ALO domain. The process produces H2O2 and 2-oxo-L-gulono-γ-lactone, which forms L-ascorbic acid after non-enzymatic isomerisation [42].

Cho et al. [15] analysed Gulo/GULO from fish and mammals and reported that a significant number of amino acid residues were exclusive to either mammals or fishes. An example would be the Pro98 residue in mammalian GULO versus Glu98 in fish Gulo. However, the Gulo from H. signifer did not conform to amino acid residues exclusive to fish, and it contained several amino acid residues in the conserved regions not found in Gulo/GULO of other animals. These amino acids included Ser28, Lys42, Gln76 and Ile103 in the FAD-binding domain, and Ala185, Leu319, Glu325 and Lys382 in the ALO domain. In spite of these replacements, the Gulo of H. signifer is phylogenetically closer to elasmobranchs than to bony fishes and amphibians. It shares the highest percentage similarity, in a descending order, with those of T. scyllium, S. torazame and M. manazo. Among the residues found only in H. signifer, Ser28 could be related to the regulation of Gulo in this stingray because it is a potential phosphorylation site located in the FAD-binding domain. Furthermore, analysis of the phosphorylation and glycosylation sites across the Gulo/GULO of different species (Table 2) revealed that those from the euryhaline freshwater H. signifer and the stenohaline freshwater Potamotrygon motoro possess the least number of phosphorylation sites. These findings indicate that successful invasion of the freshwater environment could have induced changes in the post-translational regulation of Gulo activity in the kidney of elasmobranchs.

Salinity changes and oxidative stress

In aquatic organisms, salinity change causes a variety of physiological responses such as increases in plasma concentrations of stress-related hormones, stimulation of energy metabolism, and perturbation of steady state concentrations of electrolytes. It has been established that salinity-induced stress is associated with increased production of reactive oxygen species, causing oxidative damage. Liu et al. [43] reported that acute salinity stress caused changes in activities of superoxide dismutase, catalase, glutathione peroxidase in shrimp (Litopenaeus vannamei). In addition, they [43] reported that the resistance of L. vannamei to acute salinity changes could be enhanced by moderate doses of dietary supplement of vitamin E. For fish, Roche and Boge [20] reported changes in antioxidant enzymes in the red blood cells of sea bass (Dicentrarchus labrax) subjected to hypoosmotic shock. Choi et al. [44] demonstrated that salinity changes could lead to changes in the mRNA expression of glutathione peroxidase and glutathione-S-transferase in the olive flounder (Paralichthys olivaceus). They [44] concluded that these two enzymes played an important role in the detoxification of reactive oxygen species and might serve as indicators of salinity-induced oxidative stress in P. olivaceus. Furthermore, Martínez-Alvarez et al. [21] reported that acclimation from freshwater to seawater resulted in salinity-induced oxidative stress in the sturgeon (Acipenser naccarii) as reflected by increases in activities of catalase, glutathione peroxidase and superoxide dismutase, and in lipid peroxide concentrations, in blood, liver and heart.

In comparison, the concentrations of thiobarbituric acid reactive substances remained statistically unchanged in the gills and liver of H. signifer after 1 day or 6 days of transfer form freshwater to brackish water. Hence it can be concluded that acclimation from freshwater to brackish water did not lead to increased oxidative stress in H. signifer. However based on thiobarbituric acid reactive substances alone, it would be difficult, if not impossible, to judge whether H. signifer would experience less salinity-induced oxidative stress in brackish water. Since thiobarbituric acid reactive substances represent terminal products of lipid peroxidation [45], an increase in oxidative stress would lead to their accumulation, but a reduction in oxidative stress does not necessarily result in an increase in their removal. Indeed, after 20 days of exposure to seawater, plasma osmolality, erythrocyte constants and muscle water content in A. naccarii return to levels comparable to those of the freshwater control [21]. This would mean that successful acclimation from freshwater to seawater has been achieved, leading to a reduction in salinity stress caused by seawater exposure. However, the concentration lipid peroxidation products did not return to control level [21].

Why would acclimation from freshwater to brackish water lead to decreases in renal gulo mRNA expression and Gulo activity in H. signifer?

Molecular phylogeny suggests that freshwater stingrays from Southeast Asia, including H. signifer, have only marine species as their nearest relative, and they underwent multiple colonizations rather than speciations within freshwater [46]. Since the invasion of the H. signifer lineage into freshwater habitats appears to be relatively recent [47], it is not unexpected that H. signifer would experience higher salinity stress in freshwater than in brackish water. Indeed, freshwater H. signifer has to enter a brackish or marine environment at some point in its life history, possibly for reproduction [23]. Unlike the stenohaline freshwater P. motoro, H. signifer is ureogenic and ureotelic in freshwater [24]. Although H. signifer down-regulates the ornithine-urea cycle capacity and reduces the retention of urea in freshwater, its plasma urea concentration (43.8 mmol l−1) is much higher than that of P. motoro (0.65 mmol l−1). Consequently, in freshwater, the blood plasma osmolality of H. signifer (416 mosmol kg−1) is considerably higher than that of P. motoro (349 mosmol kg−1) [24]. In addition, urea concentrations in muscle, liver and brain of freshwater H. signifer (70.9, 49.4 and 59.3 µmol g−1 wet mass, respectively) are much higher than those of P. motoro (0.38, 0.04 and 0.38 µmol g−1 wet mass, respectively) [24]. Consequently, H. signifer would experience greater osmotic stress in freshwater than in brackish water due to the steep ionic and urea gradients between the blood and the environment. After transfer from freshwater to brackish water, H. signifer would logically experience a decrease in osmotic stress and hence a decrease in salinity-induced oxidative stress, leading to a significant decrease in renal gulo mRNA expression and Gulo activity.

In rodents, exposure to xenobiotics leads to increased oxidative stress, and there are increases in ascorbate concentrations in tissues and urine due to up-regulation of ascorbic acid biosynthesis [48], [49]. In mouse [50], but not rat [48], exposure to xenobiotics also leads to increases in GULO expression and GULO activity in the liver. In addition, a negative feedback of ascorbic acid intake on the synthesis of GULO has been reported in mouse [51]. Although Moreau et al. [52] reported that Chondrostei fish showed unchanged Gulo activity with increasing dietary ascorbic acid, the slow rate of ascorbic acid synthesis in poikilotherms may account for this observation. Subsequently, Moreau and Dabrowski [53] reported that an increase in dietary levels of α-tocopherol and/or ascorbic acid significantly raised the concentrations of these two antioxidants in the liver, and concomitantly lowered the Gulo activity in the kidney, of the white sturgeon Acipenser transmontanus. Being potent free radical scavengers, α-tocopherol and ascorbic acid could affect the overall redox state of the cell, which in turn modulated the level of Gulo expression and activity in the white sturgeon [53]. Therefore, like mammals, changes in redox status can lead to changes in gulo/Gulo expression in fish. Since decreases in renal gulo mRNA expression and Gulo activity in H. signifer exposed to brackish water indicate a decrease in the production of ascorbic acid, and since salinity-induced oxidative stress has been demonstrated in fish (sea bass, [20]; sturgeon, [21]; olive flounder [44]), it can be deduced that H. signifer was in an improved redox status, probably due to a reduction in salinity-induced oxidative stress after transfer from freshwater to brackish water.

Changes in tissue ascorbate and/or dehydroascorbate concentrations indicate that H. signifer encounters less salinity-induced oxidative stress in brackish water

The concentration of ascorbate + dehydroascorbate in the kidney of H. signifer (∼43 µg g−1 wet mass) falls in the range of 37–123 µg g−1 wet mass as reported for a variety of fishes [54], [55]. The steady state concentration of ascorbate in the kidney is maintained by a balance between its synthesis through Gulo and its oxidation to dehydroascorbate plus its transport out to the blood. Results on gulo mRNA expression and Gulo activity indicate a decrease in ascorbate production in the kidney of H. signifer exposed to brackish water, but yet the concentrations of total ascorbate + dehydroascorbate, ascorbate and dehydroascorbate in the kidney remained unchanged. Hence, it can be deduced that there was a decrease in the overall demand of ascorbate as an antioxidant in non-renal organs/tissues of H. signifer during brackish water acclimation, corroborating the proposition that H. signifer experienced less salinity stress in brackish water than in freshwater.

Ascorbate is a vital antioxidant in the mammalian brain [4]. It is transported into the brain and neurons via sodium-ascorbate co-transporter 2, and accumulated within these cells against a concentration gradient [56]. The highest concentration of ascorbate in the human body is found in the brain (∼2–10 mmol l−1) and in neuroendocrine tissues such as adrenal, and it is difficult to deplete ascorbate from the brain [57]. Since human cannot synthesize ascorbic acid, a high brain ascorbate concentration is attributed to (a) an adequate dietary supplementation of vitamin C, (b) effective ascorbate/dehydroascorbate transport mechanisms [58], and (c) efficient reducing systems to maintain the vitamin in the form of ascorbate [59], [60], [61], [62]. Ascorbate probably also serves as an important antioxidant in fish brains because high brain ascorbate concentrations have been reported for Oplegnathus fasciatus (254 µg g−1) [63] and Anguilla japonica (251 µg g−1) [64]. In addition, the brain shows the slowest rate of decline in ascorbate concentration when teleost fishes, which cannot synthesize ascorbic acid, were subjected to dietary vitamin C deficiency [65], [66]. Similarly, a high concentration of ascorbate + dehydroascorbate was detected in the brain of H. signifer (approximately 222 µg g−1 wet mass), with much higher concentration of ascorbate than dehydroascorbate. Therefore, it is logical to deduce that the brain of H. signifer possesses transport mechanisms for ascorbate and/or dehydroascorbate, and biochemical mechanisms/systems to maintain ascorbate in the reduced state. Since there were a significant increase and a significant decrease in the concentrations of ascorbate and dehydroascorbate, respectively, with the total ascorbate + dehydroascorbate concentration remained unchanged in the brain of H. signifer after 1 day of exposure to brackish water, it can be concluded that brackish water acclimation indeed led to a decrease in oxidative stress in the brain.

The gills of freshwater elasmobranchs play an important role in osmoregulation [22], and are presumably one of the major epithelia that are directly exposed to oxidative stress due to environmental disturbances. In comparison, the concentration (15 µg g−1 wet mass) of ascorbate + dehydroascorbate in the gills of H. signifer was much lower than those in the brain and kidney. It is possible that besides ascorbate, other antioxidant mechanisms such as superoxide dismutase, catalase and glutathione could also play an important role in branchial oxidative defence [67]. There were significant increases in concentrations of ascorbate and total ascorbate + dehydroascorbate, with the dehydroascorbate concentration remained unchanged, in the gills of H. signifer after exposure to brackish water for 1 or 6 days. Since gulo was not expressed in the gills, it would imply that there was an increase in the uptake of ascorbate from the blood, and that the excess ascorbate was not converted to dehydroascorbate due to a decrease in salinity-induced oxidative stress when H. signifer was acclimated from freshwater to brackish water. If indeed H. signifer would experience increased oxidative stress in freshwater, the accumulation of ascorbate in the gills during brackish water acclimation can be viewed as a possible preparation for it to readjust back to freshwater when opportunity arises.

Conclusions

H. signifer has a short history of freshwater invasion [47] and is apparently confronted with greater salinity stress in freshwater than in brackish water [24]. Our results demonstrate for the first time that brackish water acclimation could lead to a down-regulation of renal gulo expression and a decrease in renal Gulo activity, indicating a decrease in the capacity of ascorbic acid synthesis, in H. signifer. Yet, concentrations of ascorbate increased in some organs, which in some cases were accompanied with a decrease in dehydroascorbate concentration, indicating that H. signifer indeed experienced lower salinity-induced oxidative stress in brackish water. Taken altogether, these results suggest that the ability to upregulate ascorbic acid production to combat hyposalinity-induced oxidative stress could have contributed in part to the successful invasion of the freshwater habitat by some euryhaline marine elasmobranchs.

Author Contributions

Conceived and designed the experiments: SFC YKI. Performed the experiments: SZHW BC. Analyzed the data: SZHW BC. Contributed reagents/materials/analysis tools: YRC WPW. Wrote the paper: SZHW YKI.

References

  1. 1. Davies MB, Austin J, Partridge DA (1991) Vitamin C: Its Chemistry and Biochemistry. Cambridge: The Royal Society of Chemistry. 154.
  2. 2. Mandl J, Szarka A, Banhegyi G (2009) Vitamin C: update on physiology and pharmacology. Br J Pharmacol 157: 1097–1110.
  3. 3. Cohen G (1994) Enzymatic/nonenzymatic sources of oxyradicals and regulation of antioxidant defences. Ann N Y Acad Sci 738: 8–14.
  4. 4. Rice ME (2000) Ascorbate regulation and its neuroprotective role in the brain. Trends Neurosci 23: 209–216.
  5. 5. Nishikimi M (1975) Oxidation of ascorbic acid with superoxide anion generated by the xanthine-xanthine oxidase system. Biochem Biophys Res Commun 63: 463–468.
  6. 6. Bodannes RS, Chan PC (1979) Ascorbic acid as a scavenger of singlet oxygen. FEBS Lett 105: 195–196.
  7. 7. Machlin LJ, Bendich A (1987) Free radical tissue damage: protective role of antioxidant nutrients. FASEB J 1: 441–445.
  8. 8. Jenner AM, Ruiz JE, Dunster C, Halliwell B, Mann GE, et al. (2002) Vitamin C protects against hypochlorous acid-induced glutathione depletion and DNA base and protein damage in human vascular smooth muscle cells. Arterioscler Thromb Vasc Biol 22: 574–580.
  9. 9. Bisby RH, Parker AW (1995) Reaction of ascorbate with the alpha-tocopheroxyl radical in micellar and bilayer membrane systems. Arch Biochem Biophys 317: 170–178.
  10. 10. Drouin G, Godin JR, Pagé B (2011) The genetics of vitamin C loss in vertebrates. Curr Genomics 12: 371–378.
  11. 11. Chatterjee IB, Chatterjee GC, Ghosh NC, Ghosh JJ, Guha BC (1960a) Biological synthesis of L-ascorbic acid in animal tissues: conversion of L-gulonolactone into L-ascorbic acid. Biochem J 74: 193–203.
  12. 12. Chatterjee IB, Chatterjee GC, Ghosh NC, Ghosh JJ, Guha BC (1960b) Biological synthesis of L-ascorbic acid in animal tissues: conversion of D-glucuronolactone and L-gulonolactone into L-ascorbic acid. Biochem J 76: 279–292.
  13. 13. Nishikimi M, Fukuyama R, Minoshima S, Shimizu N, Yagi K (1994) Cloning and chromosomal mapping of the human nonfunctional gene for L-gulono-γ-lactone oxidase, the enzyme for L-ascorbic acid biosynthesis missing in man. J Biol Chem 269: 13685–13688.
  14. 14. Nandi A, Mukhopadhyay CK, Ghosh MK, Chattopadhyay DJ, Chatterjee IB (1997) Evolutionary significance of vitamin C biosynthesis in terrestrial vertebrates. Free Radic Biol Med 22: 1047–1054.
  15. 15. Cho YS, Douglas SE, Gallant JW, Kim KY, Kim DS, et al. (2007) Isolation and characterization of cDNA sequences of L-gulono-gamma-lactone oxidase, a key enzyme for biosynthesis of ascorbic acid, from extant primitive fish groups. Comp Biochem Physiol B Biochem Mol Biol 147: 178–190.
  16. 16. Zhou X, Xie M, Niu C, Sun R (2003) The effects of dietary vitamin C on growth, liver vitamin C and serum cortisol in stressed and unstressed juvenile soft-shelled turtles (Pelodiscus sinensis). Comp Biochem Physiol A Mol Integr Physiol 135: 263–270.
  17. 17. Zhou X, Niu C, Sun R (2005) The effect of vitamin C on stress withstanding capability in the juvenile soft-shelled turtle (Pelodiscus sinensis). Aquacult Nutr 11: 169–174.
  18. 18. Paital B, Chainy GB (2010) Antioxidant defenses and oxidative stress parameters in tissues of mud crab (Scylla serrata) with reference to changing salinity. Comp Biochem Physiol C Toxicol Pharmacol 151: 142–151.
  19. 19. Sarma K, Prabakaran K, Krishnan P, Grinson G, Kumar AA (2013) Response of a freshwater air-breathing fish, Clarias batrachus to salinity stress: an experimental case for their farming in brackish water areas in Andaman, India. Aquacult Int 21: 183–196.
  20. 20. Roche H, Bogé G (1996) Fish blood parameters as a potential toll for identification of stress caused by environmental factors and chemical intoxication. Mar Environ Res 41: 27–43.
  21. 21. Martínez-Alvarez RM, Hidalgo MC, Domezain A, Morales AE, García-Gallego M, et al. (2002) Physiological changes of sturgeon Acipenser naccarii caused by increasing environmental salinity. J Exp Biol 205: 3699–3706.
  22. 22. Ballantyne JS, Robinson JW (2010) Freshwater elasmobranches: a review of their physiology and biochemistry. J Comp Physiol B 180: 475–493.
  23. 23. Otake T, Ishii T, Tanaka S (2005) Otolith Sr:Ca ratios in a freshwater stingray, Himantura signifer (Compagno and Roberts, 1982), from the Chao Phraya River, Thailand. Coast Mar Sci 29: 147–153.
  24. 24. Tam WL, Wong WP, Loong AM, Hiong KC, Chew SF, et al. (2003) The osmotic response of the Asian freshwater stingray (Himantura signifer) to increased salinity: a comparison with marine (Taeniura lymma) and Amazonian freshwater (Potamotrygon motoro) stingrays. J Exp Biol 180: 475–493.
  25. 25. Chew SF, Poothodiyil NK, Wong WP, Ip YK (2006) Exposure to brackish water, upon feeding, leads to enhanced conservation of nitrogen and increased urea synthesis and retention in the Asian freshwater stingray Himantura signifer. J Exp Biol 209: 484–492.
  26. 26. Hall TA (1999) BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic Acids Symp Ser 41: 95–98.
  27. 27. Gasteiger E, Gattiker A, Hoogland C, Ivanyi I, Appel RD, et al. (2003) ExPASy: the proteomics server for in-depth protein knowledge and analysis. Nucleic Acids Res 31: 3784–3788.
  28. 28. Blom N, Gammeltoft S, Brunak S (1999) Sequence- and structure-based prediction of eukaryotic protein phosphorylation sites. J Mol Biol 294: 1351–1362.
  29. 29. Gupta R (2001) Prediction of glycosylation sites in proteomes: from post-translational modifications to protein function. PhD thesis. Technical University of Denmark, Center for Biological Sequence Analysis.
  30. 30. Gupta R, Brunak S (2003) Prediction of glycosylation across the human proteome and the correlation to protein function. Pac Symp Biocomput 7: 310–322.
  31. 31. Gupta R, Jung E, Brunak S (2004) Prediction of N-glycosylation sites in human proteins. In preparation..
  32. 32. Bryson K, McGuffin LJ, Marsden RL, Ward JJ, Sodhi JS, et al. (2005) Protein structure prediction servers at University College London. Nucleic Acids Res 33: W36–38.
  33. 33. Felsenstein J (1989) PHYLIP–Phylogeny Inference Package (Version 3.2). Cladistics 5: 164–166.
  34. 34. Gerwick L, Corley-Smith G, Bayne CJ (2007) Gene transcript changes in individual rainbow trout livers following an inflammatory stimulus. Fish Shellfish Immunol 22: 157–171.
  35. 35. Dabrowski K (1990) Gulonolactone oxidase is missing in teleost fish – the direct spectrophotometric assay. Biol Chem Hoppe Seyler 371: 207–214.
  36. 36. Hermes-Lima M, Willmore WG, Storey KB (1995) Quantification of lipid peroxidation in tissue extracts based on Fe(III)xylenol orange complex formation. Free Radic Biol Med 19: 271–280.
  37. 37. Ramos-Vasconcelos GR, Hermes-Lima M (2003) Hypometabolism, antioxidant defenses and free radical metabolism in the pulmonate land snail Helix aspersa. J Exp Biol 206: 675–685.
  38. 38. Nam YK, Cho YS, Douglas SE, Gallant JW, Reith ME, et al. (2002) Isolation and transient expression of a cDNA encoding L-gulono-gamma-lactone oxidase, a key enzyme for L-ascorbic acid biosynthesis, from the tiger shark Scyliorhinus torazame. Aquaculture 209: 271–284.
  39. 39. Koshizaka T, Nishikimi M, Ozawa T, Yagi K (1988) Isolation and sequence analysis of a complementary DNA encoding rat liver L-gulono-gamma-lactone oxidase, a key enzyme for L-ascorbic acid biosynthesis. J Biol Chem 263: 1619–1621.
  40. 40. Sato P, Nishikimi M, Udenfriend S (1976) Is L-gulonolactone-oxidase the only enzyme missing in animals subject to scurvy? Biochem Biophys Res Commun 71: 293–299.
  41. 41. Kiuchi K, Nishikimi M, Yagi K (1982) Purification and characterization of L-gulonolactone oxidase from chicken kidney microsomes. Biochemistry 21: 5076–5082.
  42. 42. Nishikimi M, Yagi K (1991) Molecular basis for the deficiency in humans of gulonolactone oxidase, a key enzyme for ascorbic acid biosynthesis. Am J Clin Nutr 54: 1203S–1208S.
  43. 43. Liu Y, Wang WN, Wang AL, Wang JM, Sun RY (2007) Effects of dietary vitamin E supplementation on antioxidant enzyme activities in Litopenaeus vannamei (Boone, 1931) exposed to acute salinity changes. Aquaculture 265: 351–358.
  44. 44. Choi CY, An KW, An MI (2008) Molecular characterization and mRNA expression of glutathione peroxidase and glutathione S-transferase during osmotic stress in olive flounder (Paralichthys olivaceus). Comp Biochem Physiol A 149: 330–337.
  45. 45. Lushchak VI, Bagnyukova TV, Husak VV, Luzhna LI, Lushchak OV, et al. (2005) Hyperoxia results in transient oxidative stress and an adaptive response by antioxidant enzymes in goldfish tissues. Int J Biochem Cell Biol 37: 1670–1680.
  46. 46. Sezaki K, Begum RA, Wongrat P, Srivastava MP, SriKantha S, et al. (1999) Molecular phylogeny of Asian freshwater and marine stingrays based on the DNA nucleotide and deduced amino acid sequences of the cytochrome b gene. Fish Sci 65: 563–570.
  47. 47. Otake T (1999) Adaptation of freshwater stingrays collected from Thailand, Laos and India. – Serum composition, rectal gland and nephron structures, and otolith Sr:Ca ratios. In: Tanaka S, Report of studies of adaptability and conservation of freshwater elasmobranchs. pp. 79–91.
  48. 48. Horio F, Shibata T, Makino S, Machino S, Hayashi Y, et al. (1993) UDP glucuronosyltransferase gene-expression is involved in the stimulation of ascorbic acid biosynthesis by xenobiotics in rats. J Nutr 123: 2075–2084.
  49. 49. Horio F, Horie T (1997) The role of microsomal beta-glucuronidase in ascorbic acid biosynthesis stimulated by xenobiotics in rats. Biosci Biotechnol Biochem 61: 109–112.
  50. 50. Braun L, Kardon T, El Koulali K, Csala M, Mandl J, et al. (1999) Different induction of gulonolactone oxidase in aromatic hydrocarbon-responsive or -unresponsive mouse strains. FEBS Lett 463: 345–349.
  51. 51. Tsao CS, Young M (1989) Effect of exogenous ascorbic acid intake on biosynthesis of ascorbic acid in mice. Life Sci 45: 1553–1557.
  52. 52. Moreau R, Dabrowski K, Sato PH (1999) Renal L-gulono-1,4-lactone oxidase activity as affected by dietary ascorbic acid in lake sturgeon (Acipenser fulvescens). Aquaculture 180: 359–372.
  53. 53. Moreau R, Dabrowski K (2003) Alpha-tocopherol downregulates gulonolactone oxidase activity in sturgeon. Free Radic Biol Med 34: 1326–1332.
  54. 54. Fracalossi DM, Allen ME, Yuyama LK, Oftedal OT (2001) Ascorbic acid biosynthesis in Amazonian fishes. Aquaculture 192: 321–332.
  55. 55. Papp ZG, Saroglia M, Jeney Z, Jeney G, Terova G (1999) Effects of dietary vitamin C on tissue ascorbate and collagen status in sturgeon hybrids (Acipenser ruthenus X Acipenser baeri). J Appl Ichthyol 15: 258–260.
  56. 56. Agus DB, Gambhir SS, Pardridge WM, Spielholz C, Baselga J, et al. (1997) Vitamin C crosses the blood-brain barrier in the oxidized form through the glucose transporters. J Clin Invest 100: 2842–2848.
  57. 57. Harrison FE, May JM (2009) Vitamin C function in the brain: vital role of the ascorbate transporter SVCT2. Free Radic Biol Med 46: 719–730.
  58. 58. Tsukaguchi H, Tokui T, Mackenzie B, Berger UV, Chen X, et al. (1999) A family of mammalian Na+-dependent L-ascorbic acid transporters. Nature 399: 70–75.
  59. 59. Meister A (1994) Glutathione-ascorbic acid antioxidant system in animals. J Biol Chem 269: 9397–9400.
  60. 60. Winkler BS, Orselli SM, Rex TS (1994) The redox couple between glutathione and ascorbic acid: a chemical and physiological perspective. Free Radic Biol Med 17: 333–349.
  61. 61. Rose RC (1993) Cerebral metabolism of oxidized ascorbate. Brain Res 628: 49–55.
  62. 62. Fornai F, Saviozzi M, Piaggi S, Gesi M, Corsini GU, et al. (1999) Localization of a glutathione-dependent dehydroascorbate reductase within the central nervous system of the rat. Neuroscience 94: 937–948.
  63. 63. Wang X, Kim KW, Bai SC, Huh MD, Cho BY (2003) Effects of the different levels of dietary vitamin C on growth and tissue ascorbic acid changes in parrot fish (Oplegnathus fasciatus). Aquaculture 215: 203–211.
  64. 64. Ren T, Koshio S, Teshima SI, Md MI, Alam S, et al. (2005) Optimum dietary level of L-ascorbic acid for Japanese Eel, Anguilla japonica. J World Aquacult Soc 36: 437–443.
  65. 65. Dabrowski K (1990) Absorption of ascorbic acid and ascorbic sulfate and ascorbate metabolism in stomachless fish, common carp. J Comp Physiol B 160: 549–561.
  66. 66. Dabrowska H, Dabrowski K (1990) The influence of the dietary magnesium on the minerals, ascorbic acid and glutathione concentrations in tissues of common carp. Magnes Trace Elem 9: 101–109.
  67. 67. Ching B, Chew SF, Wong WP, Ip YK (2009) Environmental ammonia exposure induces oxidative stress in gills and brain of Boleophthalmus boddarti (mudskipper). Aquat Toxicol 95: 203–212.
  68. 68. Klein SL, Strausberg RL, Wagner L, Pontius J, Clifton SW, et al. (2002) Genetic and genomic tools for Xenopus research: The NIH Xenopus initiative. Dev Dyn 225: 384–391.
  69. 69. Gong X, Niu C, Zhang Z (2011) cDNA cloning and tissue expression for L-gulonolactone oxidase gene in soft-shelled turtle Pelodiscus sinensis a species with the ability to synthesize ascorbic acid. Fish Sci 77: 547–555.
  70. 70. Hillier LW, Miller W, Birney E, Warren W, Hardison RC, et al. (2004) Sequence and comparative analysis of the chicken genome provide unique perspectives on vertebrate evolution. Nature 432: 695–716.
  71. 71. Ha MN, Graham FL, D'Souza CK, Muller WJ, Igdoura SA, et al. (2004) Functional rescue of vitamin C synthesis deficiency in human cells using adenoviral-based expression of murine L-gulono-gamma-lactone oxidase. Genomics 83: 482–492.