Departments of 1 Cellular Biology and Anatomy, 2 Obstetrics and Gynecology, 3 Biochemistry and Molecular Biology, and 4 Ophthalmology, Medical College of Georgia, Augusta, Georgia 30912
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Taurine is actively
transported at the retinal pigment epithelial (RPE) apical membrane in
an Na+- and Cl-dependent manner. Diabetes may
alter the function of the taurine transporter. Because nitric oxide
(NO) is a molecule implicated in the pathogenesis of diabetes, we asked
whether NO would alter the activity of the taurine transporter in
cultured ARPE-19 cells. The activity of the transporter was stimulated
in the presence of the NO donor 3-morpholinosydnonimine. The
stimulatory effects of 3-morpholinosydnonimine were not observed during
the initial 16-h treatment; however, stimulation of taurine uptake was
elevated dramatically above control values with 20- and 24-h
treatments. Kinetic analysis revealed that the stimulation was
associated with an increase in the maximal velocity of the transporter
with no significant change in the substrate affinity. The NO-induced increase in taurine uptake was inhibited by actinomycin D and cycloheximide. RT-PCR analysis and nuclear run-on assays provided evidence for upregulation of the transporter gene. This study provides
the first evidence of an increase in taurine transporter gene
expression in human RPE cells cultured under conditions of elevated
levels of NO.
cell culture
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
TAURINE, a
-aminosulfonic acid, is the most abundant free amino acid in the
retina (38). A high concentration of taurine is essential
for maintenance of the structural and functional integrity of the
retina (20, 34). The precise physiological role of taurine
in the retina has yet to be established, although it has been suggested
to function in calcium modulation (23, 29, 34) and
osmoregulation (7, 46, 50). Taurine is an unusual
biological molecule. It is an amino acid that is not required for
protein synthesis, it accumulates in excitable tissues, and its
turnover rate is low (7-10 days). The only known metabolic role of
taurine is in the synthesis of taurine-conjugated bile acids. Taurine
is a zwitterion at physiological pH and, hence, carries no net charge.
Because the intracellular concentration of taurine in tissues such as
retina, heart, and placenta is 20-40 mM, taurine has been proposed
to function as an osmolyte (39). This proposed role is
intriguing, because retinal cell edema is associated with pathologies
such as ischemia and reperfusion during diabetic retinopathy,
macular edema, and neurodegeneration (15, 24). Taurine has
been proposed also to function as an antioxidant. Taurine, in concert
with zinc, protects rod outer segment membranes from ion and/or water
entry occurring as a consequence of membrane lipid peroxidation
(37). Recently, Keyes and Zimmerman (28) showed that taurine, in combination with retinol, protects
lipids from oxidative damage and may play a key role in protecting
retinal pigment epithelial (RPE) lipids during exposure to cyclic light.
In the retina, the concentration of taurine is highest in photoreceptor
and RPE cells (27, 35, 55). These observations have
prompted numerous investigations of transport mechanisms for taurine.
Functional studies suggest that a transporter for taurine is present on
the apical membrane of RPE (32, 33) and in cultured human
RPE cells (30). Recently cDNAs for the human taurine
transporter have been cloned, and sequence homology places it within
the gene family of Na+- and Cl-dependent
neurotransmitter transporters (25, 41). The human taurine
transporter cDNA encodes a protein of 619 amino acids with 12 putative
transmembrane domains. The taurine transporter in various cell types,
including RPE cells, is regulated by signal transduction pathways
(3, 13, 26, 40), hypertonicity (50, 51), and
extracellular taurine levels (19, 26).
Recently, the effect of diabetes on taurine transporter activity was investigated. In vitro studies by Stevens et al. (47) reported that high glucose levels rapidly and specifically decreased the activity and mRNA levels of the transporter in cultured RPE cells. In contrast, in vivo studies of RPE from diabetic rats showed that uptake of taurine was elevated, rather than decreased, suggesting that diabetes stimulates the activity of the taurine transporter (53). A molecule implicated in the pathogenic complications of diabetes, including diabetic retinopathy, is nitric oxide (NO) (14, 42, 48). Circulating levels of NO are elevated in diabetes mellitus (2, 29). Yilmaz and co-workers (58) reported a fivefold elevation of NO in the vitreous of patients with proliferative diabetic retinopathy compared with nondiabetic control patients. NO is produced by different isoforms of NO synthases (NOS) (57). In the retina, constitutive NOS and inducible NOS (iNOS) are present, the former in amacrine and ganglion cells and the latter in RPE and Müller cells (16, 45, 57). The cytokine-inducible form of NOS has been demonstrated also in RPE cells. There is evidence that NOS activity is increased in retinas of diabetic rats (8). Given that NO levels are increased in diabetic retinopathy (42, 58) and that the diabetic condition may affect taurine transport (47, 53), we were interested in determining the effects of NO on taurine transport in vitro. In the present study, we used the well-differentiated RPE cell line ARPE-19 (9, 10) to assess the effects of various NO donors on the activity and expression of the human taurine transporter. We demonstrate that exposure of cells to the NO donor 3-morpholinosydnonimine (SIN-1) leads to a dramatic increase in taurine transport activity. We further demonstrate that exposure of these cells to SIN-1 upregulates taurine transporter gene expression.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials.
[1,2-3H]taurine, [2,3-3H]alanine,
[2,3,4,5-3H]arginine, and
[-32P]uridine triphosphate were purchased from
Amersham Pharmacia (Piscataway, NJ); [3,4-3H]glutamine
and [2,3-3H]glutamate from NEN Research Products (Boston,
MA); and [3H(N)]carnitine and
[4,5-3H]leucine from Moravek Biochemicals (Brea, CA).
TRIzol reagent and cell culture supplies were purchased from Life
Technologies (Gaithersburg, MD); Ultrahyb hybridization solution from
Ambion (Austin, TX); restriction enzymes and pGEM-T vector from Promega (Madison, WI); SIN-1, sodium nitroprusside (SNP), and
methylene blue from Research Biochemicals International (Natick, MA);
actinomycin D, cycloheximide, ascorbic acid, glutathione,
-alanine,
trypan blue solution, lipopolysaccharide (LPS, Escherichia
coli), and 8-bromoguanosine 3',5'-cyclic monophosphate (8-BrcGMP)
from Sigma Chemical (St. Louis, MO); and recombinant human
interferon-
(IFN-
) from Biosource International (Camarillo,
CA). Human RPE (ARPE-19) cells were purchased from American Type
Culture Collection (Manassas, VA). Tissue-Tek OCT embedding compound
was obtained from Miles Laboratories (Elkhart, IN); the polyclonal
antibody against rat taurine transporter as well as the immunogenic
control peptide from Alpha Diagnostic International (San Antonio, TX);
and the monoclonal anti-nitrotyrosine antibody from Upstate Biologicals (Lake Placid, NY).
Animals. Three-week-old albino (ICR) mice were obtained from Harlan Sprague Dawley (Indianapolis, IN). Animals were maintained on a 12:12-h light-dark cycle and were fed the standard Purina mouse chow diet. Care and use of the animals adhered to the principles set forth in The Guiding Principles in the Care and Use of Animals (DHEW Publication No. 80-23).
Cell culture. ARPE-19 cells were cultured in 75-cm2 flasks with Dulbecco's modified Eagle's medium-nutrient mixture F-12 (DMEM-F-12) supplemented with 10% fetal bovine serum, 100 U/ml penicillin, and 100 µg/ml streptomycin. Cultures were passaged by dissociation in 0.05% (wt/vol) trypsin and seeded in 24-well culture plates. In one set of experiments assessing the effects of NO donors in conjunction with elevated glucose levels, cells were exposed for 24 h to DMEM-F-12 containing 17 mM glucose or to DMEM-F-12 containing 45 mM glucose.
Uptake measurements in ARPE-19 cells.
For uptake experiments, the culture medium was removed from ARPE-19
cells and the cells were subsequently washed twice with uptake buffer.
The composition of the uptake buffer was 25 mM HEPES-Tris, 140 mM NaCl,
5.4 mM KCl, 1.8 mM CaCl2, 0.8 mM MgSO4, and 5 mM glucose, pH 7.5. For experiments dealing with the influence of
Na+ and Cl on the transport process, the
uptake buffers consisted of 20 mM HEPES-Tris (pH 7.5) containing 140 mM
NaCl, sodium gluconate, or N-methyl-D-glucamine
chloride. Uptake was initiated by addition of 250 µl of uptake buffer
containing radiolabeled substrates. Uptake measurements were done with
a 15-min incubation at 37°C. At the end of the incubation, uptake was
terminated by removal of the medium by aspiration followed by three
washes with ice-cold uptake buffer without the radiolabeled substrates.
The cells were then solubilized in 0.5 ml of 1% sodium dodecyl sulfate
(SDS) in 0.2 N NaOH and transferred to scintillation vials for
quantitation of radioactivity.
NO donors.
To determine the effects of NO on the activity of taurine transporter
in ARPE-19 cells, confluent cultures were incubated in the absence or
presence of NO donors SIN-1 or SNP, and the uptake of
[3H]taurine was measured. The majority of experiments
were done with SIN-1, the more potent of the two NO donors in
stimulating the taurine transporter. In one set of experiments, cells
were incubated simultaneously with the cytokines LPS (20 ng/ml) and IFN- (1 µg/ml), and the uptake of [3H]taurine was
measured. The recovery of the stimulation of taurine uptake was
assessed by incubating ARPE-19 cells with 1 mM SIN-1 for 24 h,
removing the uptake buffer, and replacing the buffer with culture
medium. Cells were then maintained in the medium for 0, 2, 4, 6, and 24 h before assessment of the recovery of stimulated uptake
of [3H]taurine. To test the specificity of NO donors in
stimulating the activity of the taurine transporter, ARPE-19 cells were
incubated with 1 mM SIN-1 for 24 h at 37°C, and then uptake of
[3H]taurine and the radiolabeled compounds glutamate,
glutamine, alanine, arginine, carnitine, and leucine was measured for
15 min. The capacity of antioxidants and NO scavengers to block the SIN-1-induced stimulation of taurine uptake by the taurine transporter was examined by incubating ARPE-19 cells in uptake buffer containing SIN-1 only vs. SIN-1 in the presence of ascorbate (10 mM), glutathione (10 mM), or methylene blue (1 mM). Cells were treated also with ascorbate (10 mM), glutathione (10 mM), or methylene blue (1 mM) independently to assess whether these agents had any effect of their
own on taurine uptake by cultured ARPE-19 cells. Cells were treated
also for 16 h with 8-BrcGMP (100 µM), a cell-permeable cGMP
analog, and the uptake of [3H]taurine was measured. To
determine whether cells incubated with SIN-1 were positive for
nitrotyrosine, ARPE-19 cells were cultured to confluency on chamber
slides. They were incubated for 2 h in the presence or absence of
SIN-1. Cells were fixed in 4% paraformaldehyde for 10 min and
permeabilized with acetone for 5 min. Slides were incubated overnight
with the polyclonal antibody against nitrotyrosine (1:100) at 4°C.
The sections were subsequently incubated overnight at 4°C with
FITC-conjugated anti-mouse IgG (1:100). Sections were examined by
epifluorescence using a Zeiss Axioplan 2 microscope equipped with a
Spot camera and Spot software (version 2.2).
Data analysis. Each uptake experiment was performed in duplicate or triplicate and was repeated two to four times. Data analysis (analysis of variance) was performed using the NCSS statistical software package (P < 0.05 was considered significant). Kinetic analysis was done using the computer program Fig.P (version 6.0, Biosoft, Cambridge, UK). Data are presented as means ± SE.
Semiquantitative RT-PCR analysis of the steady-state levels of mRNAs for taurine transporter and glyceraldehyde-3-phosphate dehydrogenase. Confluent cultures of ARPE-19 cells were treated with or without 1 mM SIN-1 for 24 h at 37°C, and poly(A)+ mRNA was then prepared from these cells using the TRIzol reagent. RT-PCR was carried out using primer pairs specific for human taurine transporter and human glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The primers specific for human taurine transporter were 5'-GCTAGCTGCATAGTAGTC-3' (sense) and 5'-TGGAACACACCTCACTGC-3' (antisense) and corresponded to nucleotide positions 872-889 and 1751-1769, respectively, of the human taurine transporter cDNA (41). The primers specific for human GAPDH were 5'-AAGGCTGAGAACGGGAAGCTTGTCATCAAT-3' (sense) and 5'-TTCCCGTTCAGCTCAGGGATGACCTTGCCC-3' (antisense), corresponding to nucleotide positions 241-270 and 711-740 in human GAPDH cDNA (49). Each of these RT-PCR products was subcloned in pGEM-T vector and sequenced to establish its identity. For semiquantitative RT-PCR, PCR following reverse transcription was carried out with various numbers of cycles (range 9-30). The products were size fractionated on an agarose gel and subjected to Southern hybridization with probes specific for the taurine transporter or GAPDH. These probes were generated by labeling the respective subcloned RT-PCR products with [32P]dCTP. The intensity of the hybridization signal was quantified using the STORM PhosphorImaging System (Molecular Dynamics, Sunnyvale, CA). The relationship between the intensity of the signal and the PCR cycle number was then analyzed to determine the linear range for the PCR product formation. The intensities of the signals within the linear range were used for data analysis.
Nuclear run-on transcription assay.
To monitor SIN-1-induced changes in the transcription rate of the
taurine transporter gene, the nuclear run-on assay was performed as
described elsewhere (18) with modifications. ARPE-19 cells were incubated for 24 h with or without 1 mM SIN-1. Nuclei were isolated by homogenization with 10 strokes in a Dounce homogenizer in 2 ml of lysis buffer [10 mM Tris · HCl, pH 7.4, 3 mM
CaCl2, 2 mM MgCl2, and 1% (vol/vol) NP-40]
followed by centrifugation at 500 g for 5 min at 4°C.
Pelleted nuclei were frozen immediately in liquid nitrogen in 200 µl
of freezing buffer [50 mM Tris · HCl, pH 8.3, 40% (vol/vol)
glycerol, 5 mM MgCl2, and 0.1 mM EDTA] until the assay was
performed. The in vitro labeling of nascent RNA was performed in 200 µl of 2× reaction buffer {10 mM Tris · HCl, pH 8.0, 5 mM
MgCl2, 0.3 M KCl, 10 mM dithiothreitol, 1 mM each ATP, GTP,
and CTP, and 10 µl of [-32P]UTP (10 mCi/ml)}. The
mixture was incubated for 30 min at 30°C. RNA was isolated using the
TRIzol reagent according to the manufacturer's instructions. RNA was
suspended in 50 µl of 10 mM
N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid, pH
7.4, 10 mM EDTA, and 0.2% (wt/vol) SDS, heated for 5 min at 90°C,
and added directly to the hybridization solution. To prepare membranes
for hybridization with radiolabeled RNA, 20 µg of plasmid DNA,
containing the human taurine transporter cDNA in pGEM-T vector, were
linearized with the restriction enzyme SalI. DNA was
denatured by addition of 0.4 M NaOH for 30 min at room temperature and
then neutralized by addition of 6× saline-sodium citrate (SSC) and
placement on ice. The slot-blot apparatus was prepared, and 5 µg of
plasmid DNA were applied to each slot under low vacuum. Each slot was
rinsed with 500 µl of 6× SSC. Membrane strips were prehybridized for
2 h at 68°C in Ultrahyb hybridization buffer. Membranes were
hybridized overnight at 68°C using 1 ml of the same buffer containing
radiolabeled RNA. After hybridization, membranes were washed twice for
5 min each in 2× SSC and 0.1% SDS at 68°C and twice for 15 min each
in 0.1% SSC and 0.1% SDS at 68°C and exposed to X-ray film.
Laser scanning confocal microscopic analysis of taurine transporter in cultured ARPE-19 cells and in intact RPE. Immunohistochemical methods were used to localize taurine transporter in cultured human ARPE-19 cells and in RPE of intact mouse eyes. Eyes from 3-wk-old albino mice were enucleated and frozen immediately in Tissue-Tek OCT, and 10-µm-thick cryosections were prepared. The cells were fixed with ice-cold methanol, and the cryosections were fixed with ice-cold acetone. Cells and cryosections were blocked with 10% normal goat serum. Samples were incubated with an affinity-purified polyclonal antibody against rat taurine transporter. The rabbit anti-taurine transporter antibody (Alpha Diagnostics) was prepared using a 20-mer peptide (designated Tau-11) near the carboxy-terminal cytoplasmic region of rat TAUT-1 (43). The immunogenic peptide used in production of the antibody shows 100% homology to rat and mouse and 90% to human and canine taurine transporter. The antibody has been shown to cross-react with mouse, human, rat, and cat tissues. Cells were incubated with the primary antibody for 3 h at room temperature at a dilution of 1:100; tissue cryosections were incubated for 3 h at room temperature at a dilution of 1:50 and incubated overnight at 4°C. To demonstrate the specificity of the antibody, the primary antibody was neutralized with an excess of the antigenic peptide before use for incubation with tissue sections. Additional negative controls included using buffer only and 0.1% normal rabbit serum in place of the primary antibody. After they were rinsed, all samples were incubated overnight at 4°C with an FITC-conjugated AffiniPure goat anti-rabbit IgG at a dilution of 1:100. Cells and cryosections were optically sectioned (z-series) using a Nikon Diaphot 200 Laser Scanning Confocal Imaging System (Molecular Dynamics). Images were analyzed using the Image Display 3.2 software package (Silicon Graphics, Mountain View, CA).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present study focused on the role of NO in the regulation of
the human taurine transporter in cultured human ARPE-19 cells. Before
testing the activity of the transporter in ARPE-19 cells, we sought to
establish its presence using immunohistochemical methods. ARPE-19 cells
were cultured to confluency on plastic chamber supports. The cells were
incubated with a commercially available, affinity-purified antibody
against the taurine transporter. As shown in Fig.
1, laser scanning confocal optical
sections of cells demonstrated intense labeling with the antibody. In
Fig. 1A, cells were sectioned optically in the vertical
plane, allowing the distribution of the protein to be viewed from the
sides of the cells. There appeared to be protein distributed throughout the cell cytoplasm. In Fig. 1B, cells were scanned in a
horizontal plane, allowing the cells to be viewed from above. There was
an abundance of bright fluorescence across the apical surface of the
cells. These data supported the use of ARPE-19 cells for study of
taurine uptake activity. Incubation of cells with the antibody that had
been incubated with an excess of immunogenic control peptide resulted
in no positive staining (Fig. 1C). Although it is necessary
to use cultured cells to study the activity of the taurine transporter,
we considered it essential also to confirm the presence of the taurine
transporter in RPE cells in an intact mammalian retina, inasmuch as
there have been no immunohistochemical studies documenting the presence
of the transporter in RPE. To this end, we used cryosections of mouse
retina (Fig. 2). A
hematoxylin-and-eosin-stained cryosection of the outer retina (Fig.
2A) is provided for comparison to the photomicrograph of the
immunolabeled section (Fig. 2B). The outer portion of the
retina, including the outer nuclear layer of photoreceptor cell nuclei,
and the inner segments of these cells are labeled. The RPE is the
single layer of epithelial cells between choroid and outer segments.
Figure 2B shows a cryosection of mouse retina incubated with
the antibody against the taurine transporter. The taurine transporter
was detected abundantly in RPE and was present throughout much of the
RPE cell cytoplasm. The taurine transporter was expressed also in the
photoreceptor cell inner segments, in which the endoplasmic reticulum,
Golgi apparatus, and other organelles are located (Fig. 2B).
Labeling was present in other regions of the retina, including the
retinal ganglion cell layer and outer plexiform layer (data not shown).
|
|
Stimulation of human taurine transporter by NO
donors.
Figure 3A shows the effect of
the NO donors SIN-1 and SNP on the uptake of 80 nM
[3H]taurine by the taurine transporter in human ARPE-19
cells. Both NO donors, at 1 mM and when incubated with cells for
24 h, stimulated the uptake process. SIN-1 was a more potent
stimulator of taurine uptake than SNP. Exposure of ARPE-19 cells to
SIN-1 led to a 3.5-fold stimulation of taurine uptake. Exposure of
cells to 1 mM SNP under similar conditions resulted in an ~2.5-fold
stimulation of taurine uptake. Figure 3B shows the
dose-response relationship for SIN-1. SIN-1 was ineffective up to 100 µM. The stimulation was seen only at 1 mM. The effect was observed
with freshly prepared SIN-1. Incubation of cells with "spent" SIN-1
(i.e., SIN-1 that had been prepared 48 h before incubation) showed
a markedly reduced stimulatory effect (data not shown).
|
|
Time course of stimulation of taurine uptake by
SIN-1.
After determining that SIN-1 stimulated the uptake of taurine in
ARPE-19 cells after 24 h of incubation, we performed a time-course study. Figure 4 shows the data for
exposure of cells for various lengths of time to 1 mM SIN-1. During the
initial 16 h of exposure to SIN-1, the uptake of taurine was
similar to that of cells not exposed to the NO donor; however, by
18 h the uptake of taurine in SIN-1-treated cells increased and
was dramatically elevated above control values by 20 and 24 h.
|
Inhibition of NO-induced stimulation of taurine
transporter by antioxidants and NO scavengers.
To determine whether the stimulation of taurine uptake by SIN-1 could
be inhibited by antioxidants and NO scavengers, ARPE-19 cells were
incubated for 24 h with or without 1 mM SIN-1 in the presence or
absence of antioxidants, ascorbate (10 mM) or glutathione (10 mM), or
with the NO scavenger methylene blue (1 mM). The uptake of
[3H]taurine (80 nM) was then measured for 15 min in these
cells. As shown in Fig. 5, uptake of
taurine was threefold higher in cells treated with SIN-1 alone than in
cells treated similarly but in the absence of SIN-1. When cells were
incubated simultaneously with SIN-1 and ascorbate or SIN-1 and
glutathione, the SIN-1 stimulation of taurine uptake was markedly
attenuated. Similarly, methylene blue also attenuated the stimulatory
effect of SIN-1 on taurine uptake. These antioxidants and NO scavengers
by themselves had no noticeable effect on taurine transporter activity
(data not shown). Incubation of ARPE-19 cells with SIN-1 for 2 h
followed by immunohistochemical detection of nitrotyrosine revealed the presence of higher levels of nitrotyrosine in cells treated with SIN-1
(Fig. 6A) than in cells not
exposed to SIN-1 (Fig. 6B). A positive nitrotyrosine
reaction is considered an indicator of NO production. These data
further support the belief that the SIN-1-induced stimulation of
taurine transporter in ARPE-19 cells is mediated through NO. In
experiments in which ARPE-19 cells were exposed to 8-BrcGMP (100 µM),
the uptake of taurine (80 nM) was stimulated 23% (2.67 ± 0.10 pmol/mg protein) compared with control cells (2.17 ± 0.14 pmol/mg
protein), suggesting that the influence of NO on taurine transporter is
mediated, at least in part, by cGMP.
|
|
Specificity of NO-induced stimulation of taurine
transporter.
The NO-induced stimulation of [3H]taurine uptake in
ARPE-19 cells was not a nonspecific effect, because the uptake of other nutrients, glutamine, glutamate, alanine, arginine, carnitine, and
leucine, was not affected under identical experimental conditions (Table 2). Because the uptake of taurine
was actually stimulated in the presence of SIN-1, while uptake of other
compounds was not affected, it is not likely that SIN-1 damaged the RPE
cells. Total protein levels measured in cells (n = 8)
exposed to SIN-1 (0.21 ± 0.02 mg) did not differ significantly
from protein levels in cells not exposed to SIN-1 (0.23 ± 0.02, P > 0.5). Moreover, when cells were cultured in the
presence or absence of SIN-1 for 24 h and subjected to the trypan
blue exclusion assay (0.2% wt/vol) to assess cell viability, there was
no significant difference in the number of cells excluding the dye
(data not shown).
|
Kinetic analysis of NO-induced stimulation of
taurine transporter activity.
We then analyzed the kinetics of taurine transporter in control cells
and in cells treated for 24 h with 1 mM SIN-1 (Fig. 7). The analysis showed that the increase
in the transport activity of the taurine transporter observed in
SIN-1-treated ARPE-19 cells compared with control cells was primarily
associated with an increase in the maximal velocity of the transporter
with no significant change in the substrate affinity. The maximal
velocity of taurine uptake was 3.5-fold greater in SIN-1-treated cells
than in control cells (697.8 ± 61.9 vs. 194.3 ± 25.6 pmol · mg protein1 · 15 min
1). The Michaelis-Menten constant for taurine remained
almost the same in SIN-1-treated and control cells (9.4 ± 1.9 vs.
8.9 ± 2.7 µM).
|
|
Semiquantitative RT-PCR analysis of
NO-induced stimulation of the taurine transporter.
We then investigated the influence of SIN-1 treatment on the
steady-state levels of mRNA transcripts specific for the taurine transporter. mRNA samples isolated from control and SIN-1-treated ARPE-19 cells were used for semiquantitative RT-PCR for the
determination of the levels of mRNA transcripts. As an internal
control, we determined the steady-state levels of the GAPDH mRNA in the
samples in parallel. RT-PCR was done with a wide range of PCR cycles
(i.e., 9-30). The resultant products were run on a gel and then
subjected to Southern hybridization with 32P-labeled cDNA
probes specific for the taurine transporter and GAPDH. The
hybridization signals were quantified using the STORM PhosphorImaging
System, and the intensities that were in the linear range with the PCR
cycle number were used for analysis. The results of these experiments
(Fig. 9) showed that in the cells treated with SIN-1, the steady-state levels of taurine transporter mRNA increased markedly compared with control cells. These results demonstrate that the SIN-1-induced increase in taurine transporter activity is likely due to increased expression of the gene coding for
the transporter.
|
Nuclear run-on assay.
To confirm the results of the semiquantitative RT-PCR data, we used the
nuclear run-on transcription assay, which is a sensitive method for
direct measurement and comparison of specific gene transcription in
cells (17). We isolated nuclei from ARPE-19 cells that had
been incubated for 24 h in the presence or absence of SIN-1 and
prepared 32P-labeled nascent RNA as described by Greenberg
and Bender (17). The RNA was subsequently used in
slot-blot analysis on membranes to which the human taurine transporter
cDNA had been applied to quantitate specifically the taurine
transporter mRNA in the nascent RNA samples. As shown in Fig.
10, the intensity of the signal for the
taurine transporter mRNA from ARPE-19 cells incubated with SIN-1 was
much greater than that in cells not incubated in SIN-1. Densitometric
scans of the bands revealed a 12-fold greater intensity in the
SIN-1-treated than in the control cells. This experiment was carried
out twice, and the results were the same in both cases. The data
provide strong support that transcription of the taurine transporter
gene is stimulated in the presence of the NO donor SIN-1.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The present study was designed to assess the effects of NO on the transport of taurine by cultured human RPE cells. NO is a mediator of many physiological processes, such as vasodilation and neurotransmission; it is recognized also for its toxic effects (14). NO has been implicated in the pathogenesis of diabetic retinopathy (2, 14, 29, 42, 48, 57, 58). We have shown recently that, in cultured RPE cells, NO stimulates the transport of cystine, the disulfide form of cysteine, which is required for synthesis of the antioxidant glutathione (4). In the present study, we asked whether the transport of taurine would be affected similarly. Taurine is thought to have antioxidant properties (28, 37). Moreover, there is convincing evidence that it functions in osmoregulation (5, 39, 50).
To determine whether NO altered the activity of the taurine transporter, we used the well-differentiated ARPE-19 cell line. These cells are a rapidly growing RPE cell line established in the laboratory of Dr. Larry Hjelmeland (University of California, Davis). They form a uniform population of polarized epithelial monolayers on porous filter supports. They retain features characteristic of RPE cells, including defined cell borders, a cobblestone appearance, noticeable pigmentation (9, 10), and the capacity to phagocytose outer segment disks (11). The usefulness of these cells in studying the transport of folate was established recently (6). In the present study, we used immunohistochemical methods to determine whether the taurine transporter is present in ARPE-19 cells. With the use of an affinity-purified antibody against the transporter, our confocal microscopic analysis clearly showed that the transporter is present in ARPE-19 cells. The horizontal scans suggested that the protein was available on the apical membrane and was present also throughout the cells. A number of functional studies from our laboratory and others had suggested the presence of the transporter on the apical surface of cultured RPE cells (30, 32, 33); however, this report represents the first immunohistochemical evidence to that effect. To confirm the localization of the taurine transporter to RPE in vivo, we used cryosections of normal mouse eye. The antibody detected a strong positive signal in several layers of the retina, including the retinal ganglion cells, outer plexiform layer, photoreceptor cell inner segments, and the RPE. Although recent in situ hybridization studies have localized the mRNA encoding mouse taurine transporter to these same retinal layers (54), our data represent the first immunohistochemical evidence of the location of the transporter in any mammalian retina. The detection of the taurine transporter in intact RPE is important confirmation of the presence of the protein in RPE in vivo and validates the use of RPE cells to study its function in vitro.
Having established the presence of the transporter in ARPE-19 cells, we
sought to address the effects of NO on taurine transport. We used the
NO donors SIN-1 and SNP. Incubation of ARPE-19 cells with SIN-1 or SNP
caused a marked stimulation of uptake of taurine by the transporter.
Inasmuch as SIN-1 was more potent than SNP in triggering this effect,
the remaining experiments were conducted with SIN-1. The effects on
taurine transporter activity by SIN-1 were not immediate. Rather, the
increased activity was observed after 18 h of incubation with the
NO donor. Incubation of cells for 24 h with SIN-1 led to a
3.5-fold stimulation of taurine transporter activity. Incubation with
SIN-1 did not affect the Na+ or Cl dependency
of the taurine transporter, the Na+- and
Cl
-activation kinetics, or the capacity of
-alanine to
compete with taurine for the uptake process. Taurine uptake in ARPE-19 cells was not only stimulated by NO donors directly, but also by
exposure of the cells to agents such as LPS that are known to induce
NOS (45). Additional experiments suggested that the SIN-1
effects on taurine transporter activity were mediated via NO. Treatment
of ARPE-19 cells with NO scavengers inhibited SIN-1 stimulation of
taurine transporter activity. Moreover, immunohistochemical analysis
assessing nitrotyrosine production in SIN-1-treated cells showed
positive reactivity, indicating that NO was produced.
Kinetic analysis showed that the increased activity of the taurine transporter in the presence of SIN-1 was not due to an increase in substrate affinity but, rather, to an increase in the maximal velocity of taurine uptake. Such data suggest that synthesis of the taurine transporter may be increased in the presence of NO donors. To test this further, we cultured ARPE-19 cells in the presence of SIN-1 plus agents that inhibit transcription and translation, actinomycin D and cycloheximide, respectively. In both cases, the SIN-1-induced increase in taurine transporter activity was completely inhibited. Semiquantitative RT-PCR provided further evidence that NO increases the steady-state levels of taurine transporter mRNA ~3.5-fold. Although NO stimulated the uptake of taurine by ARPE-19 cells, it had little effect on the transport of other amino acids such as alanine or glutamate. The observed increase in steady-state levels of taurine transporter mRNA levels may be due to an increase in the transcription rate of the taurine transporter gene or an increase in the stability of the taurine transporter mRNA. The finding that the increased mRNA levels are completely abolished by the transcription inhibitor actinomycin D argues in favor of the gene expression as the site of regulation of the NO-mediated increase in mRNA levels. Moreover, the data obtained from the nuclear run-on transcription assay provide strong evidence that the increase in taurine transporter mRNA levels is due to increased transcriptional activity.
These data provide the first evidence that taurine transporter activity is regulated by NO. They are important, because NO levels are increased in diabetic retinopathy, the leading cause of blindness among working-aged Americans (56). Other investigators have examined taurine transporter activity under hyperglycemic conditions. Stevens et al. (47) showed that taurine transporter activity decreased within 4 h of exposure of human RPE cells to 30 mM glucose. On the other hand, Vilchis and Salceda (53) studied uptake of taurine in RPE isolated from rats that had been diabetic for 3 and 6 wk. They found a stimulation of taurine uptake in diabetic RPE compared with normal RPE. The differences in the experimental outcomes from these two laboratories may reflect differences in the length of time RPE cells were exposed to hyperglycemic conditions. In the case of cultured cells (47), the earliest effect of high glucose appears to be a downregulation of the transporter. Thus the acute effect of hyperglycemia on the taurine transporter in RPE cells seems to be a downregulation of the transporter expression. This effect was accompanied by a decrease in intracellular levels of taurine. The long-term effects of hyperglycemia on the taurine transporter in the RPE cells used in that study are not known. Interestingly, in the present study, exposure of ARPE-19 cells to high glucose for 24 h did not affect taurine transporter function. In the case of the studies of diabetic rats (53), taurine transporter activity increased. In this in vivo model, the retinas would have been subjected constantly to a hyperglycemic state for several weeks. It is noteworthy that the increased taurine transporter activity was greater after 6 wk of diabetes than after 3 wk. Because animal studies have shown that chronic hyperglycemia increases taurine transport activity in RPE cells (53), we speculate that long-term diabetes may be associated with elevated NO production, which in turn stimulates the taurine transporter expression. Our experimental findings showing NO-induced stimulation of taurine uptake in cultured RPE cells support this speculation. That is, they may suggest that, in diabetic retinopathy, NO is one of the molecules responsible for the observed upregulation of taurine transporter expression and activity. Our experiments address an important aspect of the altered diabetic state on RPE cells, that of increased levels of NO. Thus our data showing an upregulation of taurine transport in cultured cells may reflect conditions of longer-standing diabetes, rather than immediate effects of the initial exposure of the cells to hyperglycemia.
Because NO is known to function as a vasodilator (14) and because diabetic retinopathy has an ischemic component in which NO levels may be decreased, it may seem paradoxical to implicate NO in the pathogenesis of diabetic retinopathy. This paradox is resolved when it is recognized that the increased level of NO observed in the vitreous of diabetic patients is not an early event (58). During the early stages of diabetic retinopathy, typically called the nonproliferative stage, platelets clump together to form small stable aggregates that can lead to capillary closure (21). During this period of ischemia, there may be a reduction of vascular NO. The nonproliferative stage is followed, however, by a proliferative stage in which new blood vessels are formed. It is this proliferative stage that is associated with increased levels of NO (57). The proliferative stage is associated also with an increase in vascular endothelial growth factor, which has been shown to upregulate NO (22, 52). In addition, proliferative diabetic retinopathy is associated with elevated intravitreal levels of cytokine tumor necrosis factor, interferon, and interleukin-1 (1, 12). These compounds have been shown also to induce production of NO by the expression of iNOS, which is found in Müller and RPE cells (16, 45, 57).
The present study did not address the specific mechanism by which NO regulates the expression of the taurine transporter gene. It is known that NO activates guanylate cyclase. Our data showing that 8-BrcGMP, a cell-permeable cGMP analog, stimulated taurine uptake suggest that the effects of NO donors may be at least partially mediated by cGMP. It is well known also that, in addition to this signaling role, NO interacts with cellular redox systems, especially thiol groups. NO exposure or synthesis may impose a nitrosative stress similar to that imposed by oxygen-derived species. The attenuation of the stimulatory effect of NO on taurine transporter expression by the antioxidants ascorbic acid and glutathione suggests that nitrosative stress may contribute also to the NO-induced action observed in the present study. In summary, these data show that NO regulates the expression and activity of the taurine transporter in RPE cells in vivo. It is becoming apparent that NO regulates other transporters in RPE as well. For example, NO stimulates the transport by RPE of cystine, the disulfide form of cysteine, which is required for synthesis of the antioxidant glutathione (4). On the other hand, NO inhibits the activity of the reduced-folate transporter in cultured RPE cells (44). Studies of the regulation of transporter activity by NO may provide important insights for altered cellular function that occurs in conditions such as diabetes, when NO is elevated.
![]() |
ACKNOWLEDGEMENTS |
---|
The authors thank Susan Johnson for assistance in preparation of the manuscript and Penny Roon for preparation of the mouse tissue sections.
![]() |
FOOTNOTES |
---|
This research was supported by an unrestricted award from Research to Prevent Blindness to the Department of Ophthalmology, Medical College of Georgia, by an award from the Medical College of Georgia Research Institute, and National Eye Institute Grants EY-13089 and EY-12830.
Address for reprint requests and other correspondence: S. B. Smith, Dept. of Cellular Biology and Anatomy, Medical College of Georgia, Augusta, GA 30912 (E-mail: sbsmith{at}mail.mcg.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 5 March 2001; accepted in final form 8 August 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Abu el Asrar, AM,
Maimone D,
Morse PH,
Gregory S,
and
Reder AT.
Cytokines in the vitreous of patients with proliferative diabetic retinopathy.
Am J Ophthalmol
114:
731-736,
1992[ISI][Medline].
2.
Bank, N,
and
Aynedjian HS.
Role of EDRF (nitric oxide) in diabetic renal hyperfiltration.
Kidney Int
43:
1306-1312,
1993[ISI][Medline].
3.
Brandsch, M,
Miyamoto Y,
Ganapathy V,
and
Leibach FH.
Regulation of taurine transport in human colon carcinoma cell lines (HT-29 and Caco-2) by protein kinase C.
Am J Physiol Gastrointest Liver Physiol
264:
G939-G946,
1993
4.
Bridges, CC,
Kekuda R,
Wang H,
Prasad P,
Mehta P,
Huang W,
Smith SB,
and
Ganapathy V.
Structure, function and regulation of human cystine/glutamate transporter in retinal pigment epithelial cells.
Invest Ophthalmol Vis Sci
42:
47-54,
2001
5.
Burg, MB,
and
Kador PF.
Sorbitol, osmoregulation, and the complications of diabetes.
J Clin Invest
81:
635-640,
1988[ISI][Medline].
6.
Chancy, CD,
Kekuda R,
Huang W,
Prasad PD,
Kuhnel J-M,
Sirotnak FM,
Roon P,
Ganapathy V,
and
Smith SB.
Expression and differential polarization of the reduced-folate transporter-1 and the folate receptor in mammalian retinal pigment epithelium.
J Biol Chem
275:
20676-20684,
2000
7.
Davidson, AN,
and
Kaczmarek LN.
Taurine a possible neurotransmitter?
Nature
234:
107-108,
1971[ISI][Medline].
8.
Do Carmo, A,
Lopes C,
Santos M,
Proenca R,
Cunha-Vaz J,
and
Carvalho AP.
Nitric oxide synthase activity and L-arginine metabolism in the retinas from streptozotocin-induced diabetic rats.
Gen Pharmacol
3:
319-324,
1998.
9.
Dunn, KC,
Aotaki-Keen AE,
Putkey FR,
and
Hjelmeland LM.
ARPE-19, a human retinal pigment epithelial cell line with differentiated properties.
Exp Eye Res
62:
155-169,
1996[ISI][Medline].
10.
Dunn, KC,
Marmorstein AD,
Bonilha VL,
Rodriguez-Boulan E,
Giordano F,
and
Hjelmeland LM.
Use of the ARPE-19 cell line as a model of RPE polarity: basolateral secretion of FGF5.
Invest Ophthalmol Vis Sci
39:
2744-2749,
1998[Abstract].
11.
Finnemann, SC,
Bonilha VL,
Marmorstein AD,
and
Rodriguez-Boulan E.
Phagocytosis of rod outer segments by retinal pigment epithelial cells requires v
5 integrin for binding but not for internalization.
Proc Natl Acad Sci USA
94:
12932-12937,
1997
12.
Franks, WA,
Limb GA,
Stanford MR,
Ogilvie J,
Wolstencroft RA,
Chignell AH,
and
Dumonde DC.
Cytokines in human intraocular inflammation.
Curr Eye Res
11 Suppl:
187-191,
1992[ISI][Medline].
13.
Ganapathy, V,
Ramamoorthy JD,
Del Monte MA,
Leibach FH,
and
Ramamoorthy S.
Cyclic AMP-dependent up-regulation of the taurine transporter in a human retinal pigment epithelial cell line.
Curr Eye Res
14:
843-850,
1995[ISI][Medline].
14.
Goldstein, IM,
Ostwald P,
and
Roth S.
Nitric oxide: a review of its role in retinal function and disease.
Vision Res
36:
2979-2994,
1996[ISI][Medline].
15.
Goto, H,
Wu GS,
Chen F,
Kristeva M,
Sevanian A,
and
Rao NA.
Lipid peroxidation in experimental uveitis: sequential studies.
Curr Eye Res
11:
489-499,
1992[ISI][Medline].
16.
Goureau, O,
Hicks D,
Courtois Y,
and
de Kozak Y.
Induction and regulation of nitric oxide synthase in retinal Müller glial cells.
J Neurochem
63:
310-317,
1994[ISI][Medline].
17.
Greenberg, ME,
and
Bender TP.
Identification of newly transcribed RNA.
In: Current Protocols in Molecular Biology. New York: Wiley, 1997, sect. 4.10.1-4.10.11.
18.
Greenberg, ME,
and
Ziff EB.
Stimulation of 3T3 cells induces transcription of the c-fos protooncogene.
Nature
311:
433-438,
1984[ISI][Medline].
19.
Han, X,
Budreau AM,
and
Chesney RW.
Adaptive regulation of MDCK cell taurine transporter (pNCT)mRNA: transcription of pNCT gene is regulated by external taurine concentration.
Biochim Biophys Acta
1351:
296-304,
1997[ISI][Medline].
20.
Hayes, KC,
Carey RE,
and
Schmidt SY.
Retinal degeneration associated with taurine deficiency in the cat.
Science
188:
949-951,
1975[ISI][Medline].
21.
Heath, H,
Brigden WD,
Canever JV,
Pollock J,
Hunter PR,
Kelsey J,
and
Bloom A.
Platelet adhesiveness and aggregation in relation to diabetic retinopathy.
Diabetologia
7:
308-315,
1971[ISI][Medline].
22.
Hood, JD,
Meininger CJ,
Ziche M,
and
Granger HJ.
VEGF upregulates ecNOS message, protein, and NO production in human endothelial cells.
Am J Physiol Heart Circ Physiol
274:
H1054-H1058,
1998
23.
Huxtable, RJ.
From heart to hypothesis: a mechanism for the calcium modulatory actions of taurine.
In: The Biology of Taurine: Methods and Mechanisms, , edited by Huxtable RJ,
Franconi F,
and Giotti A.. New York: Plenum, 1987, p. 371-388.
24.
Ito, T,
Nakano M,
Yamamoto Y,
Hiramitsu T,
and
Mizuno Y.
Hemoglobin-induced lipid peroxidation in the retina: a possible mechanism for macular degeneration.
Arch Biochem Biophys
316:
864-872,
1995[ISI][Medline].
25.
Jhaing, SM,
Fithian L,
Smanik P,
McGill J,
Tong Q,
and
Mazzaferri EL.
Cloning of the human taurine transporter and characterization of taurine uptake in thyroid cells.
FEBS Lett
318:
139-144,
1993[ISI][Medline].
26.
Jones, DP,
Miller LA,
Dowling C,
and
Chesney RW.
Regulation of taurine transporter activity in LLC-PK1 cells: role of protein synthesis and protein kinase C activation.
J Am Soc Nephrol
2:
1021-1029,
1991[Abstract].
27.
Kennedy, AJ,
and
Voaden MJ.
Free amino acids in the photoreceptor cells of the frog retina.
J Neurochem
23:
1093-1095,
1974[ISI][Medline].
28.
Keys, SA,
and
Zimmerman WF.
Antioxidant activity of retinol, glutathione, and taurine in bovine photoreceptor cell membranes.
Exp Eye Res
68:
693-702,
1999[ISI][Medline].
29.
Komers, R,
Allen TJ,
and
Cooper ME.
Role of endothelium-derived nitric oxide in the pathogenesis of renal hemodynamic changes of experimental diabetes.
Diabetes
43:
1190-1197,
1994[Abstract].
30.
Leibach, JW,
Cool DR,
Del Monte MA,
Ganapathy V,
Leibach FH,
and
Miyamoto Y.
Properties of taurine transport in a human retinal pigment epithelial cell line.
Curr Eye Res
12:
29-36,
1993[ISI][Medline].
31.
Li, Y-P,
and
Lombardini JB.
Inhibition by taurine of the phosphorylation of specific synaptosomal proteins in the rat cortex: effects of taurine on the stimulation of calcium uptake in the mitochondria and inhibition of phosphoinositide turnover.
Brain Res
553:
89-96,
1991[ISI][Medline].
32.
Miller, SS,
and
Steinberg RH.
Potassium modulation of taurine transport across the frog retinal pigment epithelium.
J Gen Physiol
74:
237-259,
1979[Abstract].
33.
Miyamoto, Y,
Kulanthaivel P,
Leibach FH,
and
Ganapathy V.
Taurine uptake in apical membrane vesicles from the bovine retinal pigment epithelium.
Invest Ophthalmol Vis Sci
32:
2542-2551,
1991[Abstract].
34.
Neuringer, M,
and
Sturman J.
Visual acuity loss in rhesus monkey infants fed a taurine-free human infant formula.
J Neurosci Res
18:
597-601,
1987[ISI][Medline].
35.
Orr, HT,
Cohen AI,
and
Lowry OH.
The distribution of taurine in the vertebrate retina.
J Neurochem
26:
609-611,
1976[ISI][Medline].
36.
Pasantes-Morales, H.
Current concepts on the role of taurine in the retina.
Prog Retinal Res
5:
207-229,
1986.
37.
Pasantes-Morales, H,
and
Cruz C.
Protective effect of taurine and zinc on peroxidation-induced damage in photoreceptor outer segments.
J Neurosci Res
11:
303-311,
1984[ISI][Medline].
38.
Pasantes-Morales, H,
Klethi J,
Ledig M,
and
Mandel P.
Free amino acids of chicken and rat retina.
Brain Res
41:
494-497,
1972[ISI][Medline].
39.
Pasantes-Morales, H,
Ochoa de la Paz LD,
Sepulveda J,
and
Quesada O.
Amino acids as osmolytes in the retina.
Neurochem Res
24:
1339-1346,
1999[ISI][Medline].
40.
Ramamoorthy, S,
Del Monte MA,
Leibach FH,
and
Ganapathy V.
Molecular identity and calmodulin-mediated regulation of the taurine transporter in a human retinal pigment epithelial cell line.
Curr Eye Res
13:
523-529,
1994[ISI][Medline].
41.
Ramamoorthy, S,
Leibach FH,
Mahesh VB,
Han H,
Yang-Feng T,
Blakely RD,
and
Ganapathy V.
Functional characterization and chromosomal localization of a cloned taurine transporter from human placenta.
Biochem J
300:
893-900,
1994[ISI][Medline].
42.
Schmetterer, L,
Findl O,
Fasching P,
Ferber W,
Strenn K,
Breiteneder H,
Adam H,
Eichler HG,
and
Wolzt M.
Nitric oxide and ocular blood flow in patients with IDDM.
Diabetes
46:
653-658,
1997[Abstract].
43.
Smith, KE,
Borden LA,
Wang CH,
Hartig PR,
Branchek TA,
and
Weinshank RL.
Cloning and expression of a high-affinity taurine transporter from rat brain.
Mol Pharmacol
42:
563-569,
1992[Abstract].
44.
Smith, SB,
Huang W,
Chancy C,
and
Ganapathy V.
Regulation of the reduced folate transporter by nitric oxide in cultured human retinal pigment epithelial cells.
Biochem Biophys Res Commun
257:
279-283,
1999[ISI][Medline].
45.
Sparrow, JR,
Nathan CF,
and
Vodovotz Y.
Cytokine regulation of nitric oxide synthase in mouse retinal pigment epithelial cells in culture.
Exp Eye Res
59:
129-139,
1994[ISI][Medline].
46.
Stevens, MJ,
Henry DN,
Thomas TP,
Killen PD,
and
Greene DA.
Aldose reductase gene expression and osmotic dysregulation in cultured human retinal pigment epithelial cells.
Am J Physiol Endocrinol Metab
265:
E428-E438,
1993
47.
Stevens, MJ,
Hosaka Y,
Masterson JA,
Jones SM,
Thomas TP,
and
Larkin DD.
Downregulation of the human taurine transporter by glucose in cultured retinal pigment epithelial cells.
Am J Physiol Endocrinol Metab
277:
E760-E771,
1999
48.
Tilton, RG,
Chang K,
Hasan KS,
Smith SR,
Petrash JM,
Misko TP,
Moore WM,
Currie MG,
Corbett JA,
and
McDaniel ML.
Prevention of diabetic vascular dysfunction by guanidines. Inhibition of nitric oxide synthase versus advanced glycation end-product formation.
Diabetes
42:
221-232,
1993[Abstract].
49.
Tokunaga, K,
Nakamura Y,
Sakata K,
Fujimori K,
Ohkubo M,
Sawada K,
and
Sakiyama S.
Enhanced expression of a glyceraldehyde-3-phosphate dehydrogenase gene in human lung cancers.
Cancer Res
47:
5616-5619,
1987[Abstract].
50.
Uchida, S,
Moo Kwon HM,
Preston AS,
and
Handler JS.
Taurine behaves as an osmolyte in MDCK cells: protection by polarized, regulated transport of taurine.
J Clin Invest
88:
656-662,
1991[ISI][Medline].
51.
Uchida, S,
Moo Kwon H,
Yamauchi A,
Preston AS,
Marumo F,
and
Handler JS.
Molecular cloning of the cDNA for an MDCK cell Na+- and Cl-dependent taurine transporter that is regulated by hypertonicity.
Proc Natl Acad Sci USA
89:
8230-8234,
1992[Abstract].
52.
Van der Zee, R,
Murohara T,
Luo Z,
Zollmann F,
Passeri J,
Lekutat C,
and
Isner JM.
Vascular endothelial growth factor/vascular permeability factor augments nitric oxide release from quiescent rabbit and human vascular endothelium.
Circulation
95:
1030-1037,
1997
53.
Vilchis, C,
and
Salceda R.
Effect of diabetes on levels and uptake of putative amino acid neurotransmitters in rat retina and retinal pigment epithelium.
Neurochem Res
21:
1167-1171,
1996[ISI][Medline].
54.
Vinnakota, S,
Qian X,
Egal H,
Sarthy V,
and
Sarkar HK.
Molecular characterization and in situ localization of a mouse retinal taurine transporter.
J Neurochem
69:
2238-2250,
1997[ISI][Medline].
55.
Voaden, MJ,
Lake N,
Marshall J,
and
Morjaria B.
Studies on the distribution of taurine and other neuroactive amino acids in the retina.
Exp Eye Res
24:
249-257,
1977[ISI][Medline].
56.
Wu, G.
Diabetic retinopathy.
In: Retina: The Fundamentals. Philadelphia, PA: Saunders, 1995, p. 31.
57.
Yamamoto, R,
Bredt DS,
Snyder SH,
and
Stone RA.
The localization of nitric oxide synthase in the rat eye and related cranial ganglia.
Neuroscience
54:
189-200,
1993[ISI][Medline].
58.
Yilmaz, G,
Esser P,
Kociek N,
Aydin P,
and
Heimann K.
Elevated vitreous nitric oxide levels in patients with proliferative diabetic retinopathy.
Am J Ophthalmol
130:
87-90,
2000[ISI][Medline].