Vol. 274, Issue 6, F1102-F1108, June 1998
Decreased endothelial nitric oxide synthase in gastric mucosa
of rats with chronic renal failure
M.
Tomikawa,
M.
Ohta,
N. D.
Vaziri,
J. D.
Kaunitz,
R.
Itani,
Z.
Ni, and
A. S.
Tarnawski
Department of Medicine, Veterans Affairs Medical Center, Long Beach,
California 90822; University of California, Irvine, California;
Department of Surgery II, Kyushu University, Fukuoka 812-0054, Japan; and Veterans Affairs Medical Center, Los Angeles, California and
the University of California, Los Angeles, California 90073
 |
ABSTRACT |
According to recent reports, chronic renal failure (CRF)
increases the susceptibility of gastric mucosa to injury. Since nitric oxide plays a major role in gastric mucosal defense and injury, we
investigated, in rats with CRF produced by five-sixths nephrectomy and
in control rats, the expression of nitric oxide synthase (NOS) in the
stomach and measured mucosal and submucosal gastric blood flow. In CRF
rats, gastric mucosal blood flow was significantly reduced compared
with control rats, whereas submucosal and serosal blood flow was
significantly increased. CRF significantly decreased endothelial NOS
(eNOS) mRNA abundance by 53% (P < 0.01) and reduced expression of eNOS protein by 42%
(P < 0.01) compared with
the controls. Enzyme activity of eNOS was significantly reduced in gastric mucosa of CRF rats (P < 0.05). These data are consistent with reduced gastric mucosal blood
flow in CRF rats and can explain altered susceptibility of gastric
mucosa to injury in CRF rats.
gastric mucosal blood flow; mucosal injury
| |
INTRODUCTION |
THE MORTALITY FROM UPPER gastrointestinal hemorrhage is
significantly increased in patients with chronic renal failure (CRF) compared with normal subjects (22). The propensity for upper gastrointestinal bleeding in patients with CRF has been primarily attributed to mucosal vascular malformations and peptic ulcer disease,
which are both more prevalent in this population (28). Since CRF does
not cause gastric acid hypersecretion (14), the impaired defense
mechanisms are frequently implicated in the pathogenesis of nonvascular
mucosal lesions in CRF. The CRF rat has been frequently used as a model
system for evaluating gastric defense mechanism in CRF. Recent studies
demonstrated that CRF rats have increased susceptibility to gastric
mucosal acid-mediated damage in an aspirin injury model (17). The
pathogenesis of these mucosal abnormalities has not been fully defined.
Nitric oxide (NO), a potent vasodilator, is generated from the terminal
guanidino-nitrogen atoms of
L-arginine by a family of
enzymes known as NO synthase (NOS) (20). The role of NO in gastric
mucosal defense and mucosal susceptibility to injury is complex and
both increased and decreased NO generation have been linked to
increased gastric mucosal susceptibility to injury. NO plays a major
role in gastric mucosal defense by modulating the mucosal
microcirculation (21). It has been demonstrated that endogenously
generated or exogenously added NO protects the gastric mucosa against
injury from ethanol or endothelin-1, whereas inhibition of NOS
increases susceptibility to gastric mucosal injury (8, 10). On the
other hand, an excessive production of NO can be toxic to the gastric
mucosa and can cause mucosal injury. In fact, we have
demonstrated that the reversal of overexpressed NOS with
N
-nitro-L-arginine methyl
ester (L-NAME) can
mitigate gastric injury in rats with portal hypertension (13).
Effect of CRF on NO metabolism is controversial, with both increased
and decreased NO synthesis reported in rats and humans (9, 11, 19, 25).
Vallance et al. (25) showed that accumulation of endogenous
NG, NG-dimethylarginine,
an NOS inhibitor, may impair NO synthesis in CRF patients. Recently,
Vaziri et al. (26) have demonstrated a marked downregulation of NOS
expression in the remnant kidney and vascular tissues of rats with CRF.
To our knowledge, the effect of CRF on gene expression, enzyme
activity, or tissue localization of NOS in the stomach has not been
investigated. Since, as stated above, marked changes of NO production
can enhance the susceptibility of the gastric mucosa to injury (8, 10,
13), we postulated that increased gastric mucosal susceptibility to
injury in CRF may be, in part, due to altered NOS expression and enzyme
activity in this tissue. Therefore, we determined mRNA expression,
protein expression, and NOS enzyme activity of both endothelial NOS
(eNOS) and inducible NOS (iNOS), as well as eNOS protein localization in gastric mucosa. We also measured gastric blood flow.
| |
MATERIALS AND METHODS |
Animal models. Sixty male
Sprague-Dawley rats (Harlan Sprague Dawley, Indianapolis, IN) with an
average body weight of 300 g were used. The animals were fed a standard
laboratory diet (Purina Rat Chow; Purina Mills, Brentwood, MO) and
water ad libitum. Animals were randomly assigned to the CRF and
sham-operated control groups. Animals assigned to the CRF group
underwent five-sixths nephrectomy. Under general anesthesia (with
pentobarbital sodium, 50 mg/kg ip), the animals were subjected to a
two-thirds left nephrectomy followed by a right nephrectomy 4 days
later to produce CRF. The nephrectomies were carried out
extraperitoneally through a dorsal incision. Strict hemostasis and
aseptic measures were observed during the surgical procedures. Systolic
arterial blood pressure was monitored using a tail sphygmomanometer
(Harvard Apparatus, South Natick, MA). At the end of the 6-wk
observation period, the animals were placed in metabolic cages for a
24-h urine collection. Urine was collected for the measurement of
combined nitrate and nitrite
(NO
2/NO3)
and creatinine. Plasma and urinary creatinine were measured using
standard laboratory techniques. After measuring gastric blood flow, the
animals were killed by decapitation, and stomachs were immediately
harvested and frozen in liquid nitrogen or fixed in 10% Formalin or in
4% paraformaldehyde.
Measurements of urine
NO3/NO3.
The concentration of total
NO2/NO3
in the test samples was determined by a modification of the procedure
described by Braman and Hendrix (4) using the purge system of an NO
analyzer (NOA, model 270B; Sievers Instruments, Boulder, CO). Briefly,
the urine samples were diluted 10 times in distilled water prior to
analysis. A saturated solution of VCl3 in 1 M HCl was prepared and
filtered prior to use. A quantity of 5 ml of this reagent was purged
with nitrogen gas for 5-10 min prior to use. To minimize foaming
of the residual proteins in the samples, 100 µl of the 1/30 dilution
of antifoam C (Sigma Chemical, St. Louis, MO) were added to the
VCl3 reagent. The purge vessel was
equipped with a cold water condenser and a water jacket to permit
heating of the reagent to 90°C, using a circulating water bath. The
hydrochloric acid vapors were removed by a gas bubbler containing ~15
ml of 1 M NaOH. The gas flow rate into the chemiluminescence detector
was controlled using a needle valve adjusted to yield a cell pressure
of ~7 Torr. The flow rate of nitrogen into the purge vessel was
adjusted to prevent vacuum distillation of the reagent.
Samples were injected into the purge vessel to react with the
VCl3/HCl reagent, which converted
NO2,
NO3, and
S-nitroso compounds to NO. The NO
produced was stripped from the reaction chamber (by purging with
nitrogen and vacuum) and detected by ozone-induced chemiluminescence in
the chemiluminescence detector. The signal generated (NO peak and peak
area) was recorded and processed by a Hewlett-Packard model 3390 integrator. In a typical assay, a quantity of 5 µl of the test sample
was injected to the purge vessel, and all samples were run in
triplicate.
Standard curves were constructed using various concentrations of
NO3 (5-100 µM) relating the
luminescence produced to the given
NO3 concentrations of the standard
solutions. The amount of
NO2/NO3 in the test sample was determined by interpolation of the result into
the standard curve. The intra- and interassay variabilities for this
assay were 4.59% and 5.91%, respectively.
Measurements of gastric blood flow.
Gastric blood flow was measured under laparotomy in rats anesthetized
with pentobarbital sodium (50 mg/kg ip). The measurements were
performed using the laser-Doppler flow meter (model BLF 21; Transonic
Systemic, Ithaca, NY) as previously described (13). The selected probe
of the flowmeter with modification can measure blood flow in the tissue to the depth of 0.5 mm. To measure submucosal blood flow including muscular and serosal blood flow, the fiber-optic probe (model HL-P1002)
was applied gently to the gastric serosa. For the measurements of
mucosal blood flow, the same probe was inserted through a small incision in the forestomach and applied gently to the gastric mucosa of
the same gastric area, where serosal blood flow was measured. Blood
flow measurements were performed with a time constant of 1.0 s and were
recorded using a computer program (Windaq/200; Dataq Instruments,
Akron, OH). A measurement was considered satisfactory when
1) it was stable for at least 10 s,
2) it was free of motion artifacts,
and 3) the reading was reproducible.
Ten measurements were performed in each animal. Gastric mucosal
blood flow was expressed as tissue perfusion units (TPU) (3).
RNA isolation and RT-PCR for NOS.
Gastric specimens for analysis of RNA were immediately frozen in liquid
nitrogen, then stored at 
80°C. Frozen specimens were
homogenized with a Polytron homogenizer (Kinematica, Littau,
Switzerland) in 4 mol/l guanidinium isothiocyanate, and total RNA was
isolated using the guanidinium isothiocyanate-phenol-chloroform method
(5). The total RNA concentration in each sample was determined from
absorbance at 260 nm, and the quality of each RNA preparation was
determined by 1% agarose-formaldehyde gel electrophoresis and ethidium
bromide staining.
RT-PCR were carried out using a GeneAmp RNA PCR kit and a DNA thermal
cycler (Perkin-Elmer, Norwalk, CT) according to standard techniques
(1). Briefly, 0.3 µg of total RNA was used as the template to
synthesize complementary DNA with 2.5 U of Moloney murine leukemia
virus reverse transcriptase, in 10 µl of buffer containing 10 mmol/l
Tris · HCl, pH 8.3, 50 mmol/l KCl, 5 mmol/l MgCl2, 1 mmol/l of each
deoxyribonucleoside triphosphate, 2.5 mmol/l of random hexamer, and 1.4 U/µl of ribonuclease inhibitor. RT was performed at room temperature
for 20 min, then at 42°C for 15 min, at 94°C for 5 min, and at
5°C for 5 min. The resulting complementary DNA was used as a
template for subsequent PCR.
The PCR specific primer set used for rat eNOS (24) was 5'
TACGGAGCAGCAAATCCAC 3' (forward) and 5'
CAGGCTGCAGTCCTTTGATC 3' (reverse). The PCR for 
-actin was used
as a positive control and an internal standard. The specific primer set
for rat -actin (rat -actin control Amplimer set; Clontech
Laboratories, Palo Alto, CA) (12) was 5' TTGTAACCAACTGGGACGATATGG
3' (forward) and 5' GATCTTGATCTTCATGGTGCAGG 3'
(reverse). The PCR was performed in 50 µl of buffer containing 10 mmol/l Tris · HCl, pH 8.3, 2 mmol/l
MgCl2, 50 mmol/l KCl, 0.2 mmol/l
of each deoxyribonucleoside triphosphate, 0.25 µg of each primer, 2.5 µCi of
[
-32P]dCTP, and 1.2 U of Taq DNA polymerase. To define the
optimal amount of cDNA, increasing quantities of the RT cDNA products (2-12 µl) for both eNOS and -actin were added to the PCR
reaction (total 50 µl), and the resulting increase in PCR products
was determined. This study demonstrated that the RT products were within the linear range of the PCR detection system.
Therefore, the amplification of 10 µl of the RT cDNA product was used
in this study. To define the optimal amplification cycle, another study
was performed with 20-30 cycles of PCR amplification (27). Since
the amplified products for both eNOS and -actin during 24-30
cycles were also within the linear range of the PCR detection system,
the amplification of 28 cycles was used in this study. The temperature
profile of amplification consisted of 94°C for 1 min, 63°C for
1 min, and 72°C for 2 min.
Ten-microliter aliquots of PCR-amplified mixture were electrophoresed
using a 1.25% agarose gel. The gel was dried using a gel dryer (model
543; Bio-Rad Laboratories, Hercules, CA) and subjected to
autoradiography at room temperature for 3 h. The amplified cDNA
products were identified by restriction enzyme analysis as
reported in our previous study (13).
The intensity of bands on the X-ray film was measured by densitometric
scanning (UltroScan XL laser densitometer; Pharmacia LKB Biotechnology,
Uppsala, Sweden). The eNOS signal was standardized against -actin
signal for each sample, and results are expressed as the eNOS/-actin
ratio.
Western blot analysis. Frozen gastric
specimens were homogenized with a Polytron homogenizer (Kinematica) in
a lysis buffer containing 62.5 mmol/l EDTA, 50 mmol/l Tris, pH 8.0, 0.4% deoxycholic acid, 1% Nonidet P-40, 0.5 µg/ml leupeptin, 0.5 µg/ml pepstatin, 0.5 µg/ml aprotinin, 0.2 mmol/l
phenylmethylsulfonyl fluoride, 0.05 mmol/l aminoethylbenzenesulfonyl
fluoride. The homogenates were then centrifuged (14,000 rpm for 10 min
at 4°C) to remove tissue debris without precipitating membrane
fragments. The protein content of the homogenate was determined by the
bicinchoninic acid protein assay (23), using a commercial kit (BCA
protein assay reagent; Pierce Chemical, Rockford, IL). Proteins were
eluted from the supernatant directly into SDS sample buffer and
separated with 7.5% SDS-PAGE. Protein loading was equal (200 µg/well) in all lanes. Transfer blotting to nitrocellulose was
performed in a buffer containing 25 mmol/l
3-[(1,1-dimethyl-2-hydroxyethyl)amino]-2-hydroxypropanesulfonic acid, pH 9.5, in 20% methanol. Filters were blocked in a buffer containing 5% powdered nonfat milk, incubated for 1 h with specific polyclonal antibodies against eNOS (Affinity BioReagents, Neshanic Station, NJ) or iNOS (Transduction Laboratories, Lexington, KY), diluted 1:1,000, and incubated for 1 h with anti-rabbit IgG peroxidase conjugate (Sigma Chemical). The signal was visualized by the
chemiluminescence method, using ECL Western blotting detection reagents
and Hyperfilm-ECL (Amersham Life Science, Arlington Heights, IL). The
lysate of endothelial cells (EA · hy
926; provided by Dr. Edgell, Department of Pathology,
University of North Carolina, Chapel Hill, NC) (6) and the
electrophoresis standard protein for iNOS (Cayman Chemical, Ann Arbor,
MI), respectively, were used as positive controls for eNOS and iNOS.
Quantification of protein signals was performed by densitometric
scanning of autoradiographs (UltroScan XL Laser Densitometer, Pharmacia
LKB Biotechnology).
Immunofluorescence staining for eNOS.
Gastric specimens were fixed in 4% paraformaldehyde for 4 h and
subsequently transferred to 0.5 mol/l sucrose in phosphate-buffered
saline for 24 h. Then they were frozen at 80°C until
cutting. Cryostat sections (10-µm thick, Jung Cryocut 1800; Leica,
Deerfield, IL) were digested with 0.1% trypsin (Sigma Chemical) at
37°C for 10 min and incubated overnight with the specific
polyclonal antibody against eNOS (Affinity BioReagents) diluted 1:100.
This antibody was also used for Western blot analysis. For control
studies, gastric sections were incubated overnight with
phosphate-buffered saline instead of the primary antibody. After
washing with phosphate-buffered saline, sections were incubated for 30 min with fluorescein-conjugated immunoglobulin (Sigma Chemical) diluted
1: 50. Immunofluorescence of coded sections was evaluated using a Nikon
Optiphot epifluorescence microscope with B filter composition (Nikon,
Garden City, NY). For the quantitative assessment of fluorescence
intensity we used a Nikon TMD Diaphot microscope connected to a video
analysis system (Image-1/FL; Universal Imaging, Westchester, PA) (13).
The Image-1 system allows an image to be entered into computer memory
in a fraction of second, therefore fading of fluorescence is not a
problem. This system distinguishes density of staining on a scale of
0-255 units. All measurements of fluorescence intensity were made
by an investigator unaware of the code. Fluorescence intensity in the
endothelia of mucosal veins (collecting venules) and submucosal veins
was measured in standardized rectangles under ×400 magnification
in 10 randomly selected fields of each section. Each measurement was
standardized by subtracting the background intensity in each slide. All
samples were processed and immunostained at the same time, and
fluorescence intensity was measured on coded sections during the same
session and under the same conditions.
NOS activity assay. NOS activity was
measured by determining the conversion of
L-[3H]arginine
to [3H]citrulline
according to the method described by Fernández et al. (7). Frozen
gastric specimens obtained from operated rats were homogenized with a
Polytron homogenizer (Kinematica) in a buffer (250 mg/ml) containing
320 mmol/l sucrose, 0.1 mmol/l EDTA, 10 mmol/l HEPES, pH 7.4, 1 mmol/l
DL-dithiothreitol, 10 µg/ml leupeptin, and 2 µg/ml aprotinin. The homogenates were then
centrifuged (14,000 rpm for 20 min at 4°C). Such centrifugation
removes tissue debris but not the membrane fraction. The protein
normalized supernatants were assayed in duplicate.
The supernatant (40 µl) was added to 100 ml of a buffer consisting of
40 mmol/l potassium phosphate, pH 7.4, 8 mmol/l
L-valine, 1 mmol/l reduced
nicotinamide adenine dinucleotide phosphate, 1 mmol/l
MgCl2, 2 mmol/l
CaCl2, 40 µmol/l
L-arginine, 10 mg/l calmodulin,
and 5.12 mCi/l
L-[3H]arginine
monohydrochloride (specific activity 64 Ci/mmol; Amersham Life
Science). Assays were also performed in the presence of 10 mmol/l
L-NAME
(L-NAME buffer) or in the
presence of 10 mmol/l EGTA (EGTA buffer). Samples were incubated for 10 min at 37°C before termination of the reaction by the addition of
860 µl of H2O at 4°C. Then
250 µl of the diluted incubate solution was added to 250 µl of the
Na+ form of Dowex 50W-X8
(Bio-Rad). The resin-incubate mix was settled for 15 min at 25°C,
and 100 µl of supernatant was pipetted into scintillation vials. Four
milliliters of counting fluid (Cytoscint; ICN
Pharmaceuticals, Costa Mesa, CA) were added to each vial. Measurements of radioactivity (in cpm), corresponding to
[3H]citrulline, were
made on a liquid scintillation counter (model LS5801; Beckman
Instruments, Palo Alto, CA). Results were expressed as picomoles per
milligram of total protein per minute. The eNOS activity was determined
from the difference between activities obtained in control and EGTA
buffers, and the iNOS activity was determined from the difference
between activities obtained in EGTA and
L-NAME buffers.
Statistical Analysis. Results are
expressed as the means ± SD. Student's
t-test was used to compare data
between CRF and control groups. One-way analysis of variance and
Bonferroni correction were used to compare differences between
localized fluorescence intensity in the immunostaining.
| |
RESULTS |
General data and urinary excretion of NO
metabolites. Weight gain and hematocrit in the CRF
group were significantly lower, compared with the control rats, at the
conclusion of the study (Table 1). A
significant rise in plasma creatinine and a significant decline in
creatinine clearance were observed in the CRF group. Moreover, arterial
blood pressure was significantly higher in CRF animals than in the
control group. Urinary excretion of the NO metabolites,
NO2/NO3,
was significantly decreased by 43% in CRF rats (Table 1). The
intra- and interassay variabilities for this assay were 4.59% and
5.91%, respectively.
Gastric blood flow. In CRF rats,
gastric mucosal blood flow was significantly decreased (from 21.2 ± 2.8 in controls to 15.8 ± 4.6 TPU in CRF rats;
P < 0.05), whereas gastric
submucosal blood flow was significantly increased in CRF rats compared
with the control rats (from 11.5 ± 5.0 in controls to 24.8 ± 6.7 TPU in CRF rats; P < 0.05) (Fig.
1).
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Fig. 1.
Gastric submucosal and mucosal blood flow. Values are means ± SD (n = 12 animals per each
group); TPU, tissue perfusion units.
* P < 0.05 compared with
controls. Control, control rats; CRF, chronic renal failure rats.
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|
Expression of eNOS mRNA in stomach.
RT-PCR demonstrated expression of gastric eNOS mRNA in CRF and control
groups (Fig.
2A). The
abundance of eNOS mRNA in the CRF stomachs was reduced by 53%,
compared with the controls (P < 0.01; Fig. 2B).
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Fig. 2.
A: RT-PCR analysis for endothelial NO
synthase (eNOS) mRNA and -actin mRNA in gastric tissues.
B: quantitative analysis of eNOS mRNA
in the gastric tissues using densitometric scanning of amplified PCR
products. Each eNOS signal was standardized against the corresponding
-actin signal, and results are expressed as eNOS/-actin ratio.
Values are means ± SD (n = 18 animals per each group). * P < 0.01 compared with controls.
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Expression and localization of eNOS and iNOS proteins
in stomach. Western blots demonstrated the presence of
eNOS (140-kDa bands) in stomachs of CRF and control groups (Fig.
3A). No
iNOS protein (130-kDa bands) was detected by Western blots in the
gastric mucosa of either normal or CRF rats (Fig.
3B). Gastric eNOS protein expression
was reduced by 42% in the CRF group, compared with the controls
(P < 0.01, Fig.
3C).
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Fig. 3.
Western blot analysis for eNOS (A)
and inducible NOS (iNOS; B) in
gastric tissues. eNOS and iNOS proteins are expressed as 140- and
130-kDa bands, respectively. C:
quantitative analysis of eNOS protein expression in the gastric tissues
using densitometric scanning of Western blots. Values are means ± SD in densitometric units (n = 12 animals per each group). * P < 0.01 compared with controls. EC, lysate of endothelial cells (positive
control); P, electrophoresis standard protein for iNOS (positive
control).
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Immunofluorescence microscopy demonstrated that the distribution of
eNOS was mostly localized to the endothelium of mucosal and submucosal
vessels (Fig. 4,
A and
B).
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Fig. 4.
Photomicrographs of immunofluorescence staining for eNOS in the gastric
wall of control rats (A) and CRF
rats (B). Staining is mainly
localized to endothelium of mucosal and submucosal vessels.
C: intensity of immunofluorescence
signal for eNOS in stomachs of control rats (open bars) and CRF rats
(solid bars); mv, endothelium of mucosal veins; m, muscularis mucosae;
smv, endothelium of submucosal veins; pm, muscularis propria. Values
are means ± SD in image intensity units
(n = 6 animals per each group).
* P < 0.01 compared with
control rats.
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The expression of eNOS fluorescence signal in the endothelium of
mucosal and submucosal vessels was significantly reduced in CRF rats
compared with the control rats (mucosal vein, 24% reduction;
submucosal vein, 24% reduction; P < 0.01) (Fig. 4C).
NOS enzyme activity. Gastric eNOS
enzyme activity in the CRF rats was reduced by 21% compared with
controls (P < 0.05), whereas iNOS
enzyme activity was similar in the two groups (Fig.
5).
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Fig. 5.
Enzyme activity of gastric eNOS
(left) and iNOS
(right). Values are means ± SD
in pmol · mg of total
protein1 · min1
(n = 6 animals per each group).
* P < 0.05 compared with
control rats.
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| |
DISCUSSION |
This study demonstrated for the first time that eNOS mRNA abundance,
protein expression, and enzyme activity are decreased in the stomachs
of rats with CRF. Expression of eNOS protein signal detected by
immunofluorescence staining was predominantly localized to the
endothelium of mucosal and submucosal vessels and was significantly decreased in CRF rats compared with control rats. Although the methods
used are semiquantitative, the results are internally consistent.
Urinary excretion of
NO2/NO3 at steady-state condition was reduced in the CRF group, suggesting decreased total body NO production in these animals. Since NO plays an
important role in the maintenance of mucosal integrity by increasing
mucosal blood flow (21), the decreased expression of eNOS may lead to
decreased gastric mucosal flow and influence the susceptibility of the
gastric mucosa to injury.
Utilizing direct measurements of mucosal blood flow, this study
demonstrated that in CRF rats gastric mucosal blood flow is significantly reduced, whereas gastric serosal + submucosal
blood flow is significantly increased compared with control rats. These new data, i.e., selectively reduced gastric mucosal blood flow, are
consistent with and can be explained by reduced eNOS expression in
gastric mucosa of rats. These data can also explain increased susceptibility of gastric mucosa of CRF rats to injury. The increased submucosal blood flow is consistent with submucosal blood shunting and/or development of arteriovenous malformations, which are
present with increased frequency in clinical CRF. Quintero et al. (17, 18) have demonstrated increased basal gastric mucosal blood flow and
increased susceptibility to gastric damage in CRF rats. Furthermore,
they found that perfusion of stomach with acid further increased
mucosal blood flow. This effect was abolished by specific NOS
inhibitors, suggesting an NO-dependent mechanism governing the
hyperemic response to luminal acid in CRF (18). They also found that
systemic vascular resistance and peripheral blood flow were unchanged,
and mean arterial pressure was increased (18). These findings
indirectly suggest that CRF specifically affects gastric mucosal NOS
activity. Since other examples of enhanced NO synthesis, such as sepsis
and portal hypertension, are associated with systemic hyperdynamic
circulation and decreased peripheral vascular resistance (16), the
circulatory changes in CRF reported by Quintero et al. (17, 18)
are much different from the above conditions.
In contrast to the indirect observations made by Quintero and
co-workers (18), suggesting activation of gastric NO system, we found
reduced eNOS mRNA abundance, protein expression, and enzyme activity in
the stomachs of the CRF rats. Our data of gastric blood flow obtained
with a selective measurements of blood flow in the mucosa and
separately in the submucosa can explain differences between our studies
and those of Quintero et al., because the indirect method used by
Quintero et al. measures the blood flow in gastric wall (mucosa plus
submucosa) rather than selectively in the mucosa. The laser-Doppler
method used for the blood flow measurements in our study is based on
the principle that light scattered by moving erythrocytes experiences a
shift in its frequency. Thus this method measures directly flow of
erythrocytes in the tissue. There are also several additional possible
explanations for the increased submucosal blood flow in CRF rats. The
negative effect of decreased NO production on gastric blood flow may
have been offset by CRF-associated volume expansion, hypertension, and
anemia, which tend to increase blood flow. It is also possible that
other vasoactive dilatory mediators (e.g., adrenomedullin or atrial
natriuretic peptide) are involved in the process of increased
submucosal blood flow.
The increased submucosal blood flow could also be explained by
"denervation hypersensitivity" to NO in CRF, in which low basal NO production may upregulate overall NO sensitivity, producing exaggerated physiological responses to stimuli (2). This phenomenon is
suggested to be present in patients with essential hypertension (15),
but its relevance to CRF remains speculative until definitively studied. It should be noted that in addition to NO, several other factors including hypervolemia, anemia, and hypertension frequently present in CRF, can affect local and systemic circulation. This can
potentially account for a possible increase in submucosal gastric blood
flow, despite depressed local NO production in the CRF animals shown
here.
Another novel finding in this study was that only gastric eNOS
expression was reduced, whereas iNOS expression was unaffected by CRF.
There are few previous studies concerning the regulation of NOS
activity in CRF. Noris and co-workers (11) demonstrated that platelets
obtained from CRF subjects generated more NO than did platelets
obtained from control subjects and that CRF plasma induced NO synthesis
by normal platelets in vitro. On the basis of these data, the authors
speculated that in CRF, iNOS activity was increased in response to
elevated blood concentrations of tumor necrosis factor-, which in
turn were the result of decreased renal cytokine clearance in CRF
(11). In contrast, Vaziri et al. (26) have shown marked downregulation
of vascular and renal tissue NOS in CRF rats. The results of the
present study demonstrating reduced NOS expression in gastric mucosa
of CRF rats are consistent with those of Vaziri et al. (26) in the
kidney and vascular tissue in this model.
Our previous study demonstrated that eNOS mRNA expression and enzyme
activity in gastric mucosa are much higher than those of iNOS (13).
This study did not demonstrate (by Western blot analysis) marked
presence of iNOS protein in gastric mucosa of either normal or CRF
rats. The iNOS activity assay, however, demonstrated low iNOS enzyme
activity in both normal and CRF rats. These results are internally
consistent, because the iNOS protein levels corresponding to such a low
activities are below the range of detection levels by Western blot
analysis.
In summary, rats with CRF have reduced expression of eNOS mRNA and
protein, decreased eNOS enzyme activity in the stomach, and reduced
gastric mucosal blood flow. These findings suggest that reduction of
eNOS activity (by inference, reduced NO generation) causes reduced
mucosal blood flow and affects susceptibility to gastric mucosal injury
in CRF rats.
| |
ACKNOWLEDGEMENTS |
We thank Dr. Cora-Jean S. Edgell, Department of Pathology,
University of North Carolina, Chapel Hill, for kindly providing the
permanent endothelial cell line (EA · hy 926).
| |
FOOTNOTES |
This study was supported by the Medical Research Service of the
Department of Veterans Affairs. M. Ohta was visiting scientist from
Department of Surgery II, Kyushu University, Fukuoka, Japan, and was
the recipient of the Uehara Memorial Foundation (Tokyo, Japan)
postdoctoral research fellowship.
Address for reprint requests: A. S. Tarnawski, Gastroenterology Section
(111G), DVA Medical Center, 5901 E. Seventh St., Long Beach, CA 90822.
Received 20 March 1997; accepted in final form 23 February 1998.
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