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Department of Pediatric Surgery, Osaka University Medical School, Suita, Osaka 565, Japan
1To whom correspondence should be addressed.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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KEY WORDS: • nitric oxide synthase inhibitor • apoptosis • goblet cell hyperplasia • zinc deficiency • intestine • rats
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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), indicating that the intestine is one of the tissues most
sensitive to zinc deficiency. Our previous study (Cui et al. 1996
), consistent with other reports (Nobili et al. 1997
, Southon et al. 1984
, Vallee and Falchuk 1993
), demonstrated that zinc deficiency induces
changes in intestinal morphology, including decreases in villous height
and crypt depth, inflammatory cell infiltration of the lamina propria
and lesions of intestinal mucosa.
Nitric oxide, a free radical gas, is an important regulatory factor in
physiologic processes (Vallance and Moncada 1994
). The
intestine possesses both the calcium-dependent constitutive nitric
oxide synthase
(NOS)2
and the
calcium-independent inducible NOS (iNOS), which has been demonstrated
under lipopolysaccharide stimulation (Tepperman et al.
1993
). iNOS, when induced, produces a large amount of nitric
oxide (Iadecola et al. 1995
), resulting in a decrease in
cellular viability and local intestinal damage (Tepperman et al.
1993
). Recently, we reported that zinc-deficient (ZD) rats had
an iNOS gene expressed in the intestine and that interleukin-1
treatment caused many fold enhancement in expression and induced
diarrhea (Cui et al. 1997
). Accordingly, we reasoned
that nitric oxide produced by iNOS may play a role in the mechanisms of
zinc deficiency–induced damage in the intestine.
Evans blue, when injected into the circulation, binds within seconds to
serum proteins (mainly albumin), forming a dissociable complex. An
increase in local capillary permeability to macromolecules, caused by
inflammation or other types of damage, will therefore be detected as an
extravasation and deposition of the protein-Evans blue complex in the
interstitial tissues (Lange et al. 1994
). Evans blue
leakage technique has been used to evaluate the role of NO in
microvascular permeability of the skin (Lippe et al. 1993
).
Apoptosis is a physiologically essential mechanism of cell death that
together with cell proliferation is responsible for the precise
regulation of cell numbers for a variety of cell populations during
normal development (Raff et al. 1993
, Steller 1995
). It also serves as a defense mechanism to remove unwanted
and potentially dangerous cells (Steller 1995
). Evidence
from both in vivo and in vitro experiments with rodents indicates that
zinc deficiency induces apoptosis (Rogers et al. 1995
,
Sunderman 1995
). It was demonstrated by an
ultrastructural study that zinc deficiency increased apoptotic bodies
in the intestinal mucosa (Elmes and Jones 1980
). On the
other hand, NO, especially when produced excessively, causes apoptosis.
It has been documented that cytokines, including interleukin-1 and
tumor necrosis factor-, upregulate Fas, a death signal, through a
nitric oxide-dependent mechanism in vascular smooth muscle cells
(Fukuo et al. 1997
). NO may also induce apoptosis by
inhibiting glyceraldehyde-3-phosphate dehydrogenase (Nakazawa et al. 1997
).
In this study, we used L-NAME, a NOS inhibitor, to determine whether inhibition of NO production could affect vascular permeability, apoptosis and morphologic changes in the intestine of rats with zinc deficiency.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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All experiments involving animals were conducted in accordance with NIH
guidelines (NRC 1985
) and were approved by the Osaka
University Medical School Animal Care and Use Committee. Male
Sprague-Dawley rats (n = 40, Charles River, Yokohama,
Japan), at 3 wk of age, were individually housed in acid-washed,
stainless steel cages at 23°C with a 12-h light:dark cycle. Rats were
allowed free access to glass-distilled deionized water and fed a
semipurified zinc-supplemented (50.8 mg zinc/kg) diet for 1 wk to allow
acclimation to our laboratory conditions before being divided into
three groups. One group was given free access to zinc-supplemented diet
(ad libitum group, AL, n = 10); the second group was
given a zinc-deficient (ZD) (n = 20) diet (2 mg
zinc/kg), based on the AIN-76A formulation as previously described
(Cui et al. 1997
). The third group was pair-fed (PF,
n = 10) the zinc-supplemented diet at a level of food
intake equal to the daily mean amount of the ZD group.
Experimental protocol.
AL, PF and ZD rats were fed their respective diets for 4 wk. After receiving the ZD diet for 1 wk, rats were further randomly assigned to ZD-NAME (+) and ZD-NAME (-) groups and had free access to drinking water with or without the addition of 0.3 g/L L-NAME (n = 10), respectively, for 3 wk. After the diets were fed for 4 wk, rats were anesthetized by diethyl ether. Heparinized blood was collected from the abdominal aorta, and the small intestine was taken. In total, 15 cm of intestinal sample from the upper jejunum, including the mucosal, muscular and plasma layers, was used for histologic study, the TdT-mediated dUTP-biotin nick end labeling (TUNEL) test, RNA analysis and determination of zinc content and iNOS activity. Plasma was separated and stored at -20°C until analysis. The intestinal samples for RNA analysis were processed immediately.
Determination of zinc content.
Plasma was digested with 1 mol/L hydrochloric acid as described
previously (Takagi et al. 1986
). Intestine was digested
in 5 volumes of concentrated nitric acid in a tightly capped tube at
room temperature for 24 h and then at 100°C overnight, as
described previously (Cui et al. 1997
). By using atomic
absorption spectrophotometry (Z-6100 simultaneous multielement atomic
absorption spectrophotometer, Hitachi Instrument, Tokyo, Japan), zinc
concentrations were calculated from a standard curve generated using
Zinc Std. Soln. (Wako Pure Chemical, Osaka, Japan).
Quantification of Evans blue extravasation in intestine.
Intestinal microvascular permeability was evaluated by the Evans blue
leakage technique (Lange et al. 1994
). After rats were
fed their diets for 4 wk, a subgroup of AL, PF, ZD-NAME (+), and
ZD-NAME (-) rats, (n = 5 for each group) was injected
with 20 mg Evans blue/kg (0.01 mg/L, dissolved in physiologic saline
containing 0.1 U heparin/L) through the femoral vein under
ethyl ether anesthesia. At 30 min after injection, rats were killed by
bleeding; the jejunum was removed, washed with physiologic saline and
blotted with filter paper. Extravasated Evans blue was extracted from
the intestine with formamide at 50°C for 24 h. The amount of dye
eluted in formamide was determined by spectrophotometry at 630 nm.
The calibration curve was made by dilution of Evans blue with formamide
at concentrations varying from 312.5 x
10-3 to 10-2 ng/L. The
correlation between concentration and the absorbance at 630 nm was
linear, giving the following equation:
 |
where y represents the concentration of Evans blue and x is the corresponding absorbance at 630 nm. The extravasation of Evans blue was expressed as µg/g wet tissue.
RNA isolation.
Total RNA was extracted from tissues by using the commercial reagent
ISOGEN (NipponGene, Tokyo, Japan), as described previously (Cui et al. 1997
). The RNA was dissolved in diethyl
pyrocarbonate-treated distilled water. The concentration of RNA was
estimated from the absorbance at 260 nm (the ratio at 260/280 was
between 1.6 and 1.9).
Competitive reverse transcription-polymerase chain reaction (RT-PCR).
The expression of MT-1 mRNA was determined by competitive RT-PCR as
described previously (Cui et al. 1998a
).
Briefly, MT-1 gene-specific primers (upper primer: 5'-CCC AAC TGC TCC
TGC TCC AC-3'; lower primer: 5'-GTC ACT TCA GGC ACA GCA CG-3') and
composite (MT-1 MIMIC) primers (upper primer: 5'CCC AAC TGC TCC TGC TCC
ACC TGC TCG CTT CGC TAC TTG CA-3'; lower primer: 5'-GTC ACT TCA GGC ACA
GCA CGC GGC ACC TGT CCT ACG AGT TG-3') were used to synthesize the
target MT-1 cDNA or MT-1 MIMIC cDNA, respectively. To obtain a standard
curve of MT target quantity to target/MIMIC intensity ratio, the PCR
products were amplified again by using target cDNA primer alone from
0.01 attomole (amol) of the MIMIC products together with the target
products diluted serially.
Samples of cDNA were then added to PCR amplification reactions
containing a constant amount of MT-1 MIMIC [0.01 amol (amol =
10-18 mol)] with the following schedule:
denaturation, annealing and extension at 94, 60 and 72°C for 1 min, 1
min and 1 min 30 s, respectively, for 25 cycles. PCR products were
electrophoresed on 2% agarose gels containing ethidium bromide. The
intensities of UV-induced fluorescence were analyzed by NIH Image 1.55
software, and MT-1 quantity was calculated according to the standard
curve of MT target quantity to target/MIMIC intensity ratio (Fig. 1
).
|
iNOS mRNA expression was analyzed by a semiquantitative RT-PCR by using
a pair of gene-specific primers (forward 20-mer, 5'-GCT ACA CTT CCA ACG
CAA CA-3'; reverse 20-mer, 5'-TGG GTG GGA GGG GTA GTG AT-3') with the
schedule of denaturation, annealing, and extension at 94, 60 and 72°C
for 40 s, 1 min and 1 min 30 s, respectively, for 35 cycles.
To ensure that equal amounts of reverse-transcribed RNA were added to
the PCR reaction, a parallel amplification of
glyceraldehyde-3-phosphate-dehydrogenase (GAPDH) mRNA was performed as
an internal reference as described previously (Cui et al. 1997
) The ratio of iNOS mRNA/GAPDH mRNA intensities was used to
evaluate the relative levels.
Assay of NOS activity.
Intestinal samples were homogenized in 5 volumes of ice-cold
homogenized buffer. NOS activity was determined as the conversion of
radiolabeled L-arginine to L-citrulline by the
methods described previously (Seo et al. 1994
). Briefly,
10 µL of a sample was incubated with
L-[Guanidino-14C] arginine (NEN,
Tokyo Japan) added with 2 mmol/L of guanidinoethyldisulphide (GED+) or
without GED (GED-), which is a selective inhibitor of iNOS
(Szabo et al. 1996
), for 10 min at 37°C. The whole
reaction mixture was then applied to a 0.4 mL Dowex 50 WX column (Na+
form, 200–400 mesh). Citrulline was eluted with 0.5 mL of buffer, and
its radioactivity was determined with a liquid scintillation counter.
The specific activity was expressed as pmol/(min · mg protein) of
citrulline formed. The arbitrary iNOS activity was calculated by the
following formula: NOS activity (GED-) - NOS activity (GED+). Then,
the activity ratio of iNOS vs. total NOS, i.e., NOS activity (GED-),
was calculated as follows: [NOS activity (GED-) - NOS activity
(GED+)]/NOS activity (GED-) x 100.
Identification of apoptosis.
The TUNEL method was performed with the Apop Tag in Situ Apoptosis Detection Kit- Peroxidase (Oncor, Gaithersburg, MD). Sections were washed twice with xylene for 5 min, with 95 and 70% ethanol for 3 min each wash, and then with double distilled water (DDW). They were then treated with 20 mg proteinase K/L (Sigma Chemical, St. Louis, MO) for 15 min at room temperature and washed four times with DDW for 2 min. Endogenous peroxidase was inactivated by covering the sections with 2% H2O2 for 5 min at room temperature. The sections were rinsed with PBS, and immersed in TdT buffer for 15 min at room temperature. TdT enzyme were then added to incubate at 37°C for 60 min. The reaction was terminated with stop buffer for 30 min at 37°C. After being washed with PBS 3 times for 5 min, the sections were treated with antidigoxigenin-peroxidase for 30 min at room temperature, washed again with PBS and visualized by DAB chromagen. After methyl green staining, the sections were washed again, dried and mounted. For a slide serving as a negative control, procedures were identical to those used for the experiment except that TdT enzyme was omitted (distilled water substituted for enzyme). The samples from at least three rats from each group were evaluated for count of the apoptotic-positive cells in the villus. Counts were made in 10 viewing fields with 10 villi from each sample, but the field with the highest number of apoptotic-positive cells was selected, photographed and then analyzed.
Morphologic study.
Rat intestines were fixed in 10% buffered formalin. Samples were embedded in paraffin, and 3-µm thick sections were made and stained with hematoxylin and eosin. Samples from at least three rats from each group were evaluated by counting the goblet cells of the villi. Counts were made in 10 viewing fields with 10 villi in each specimen, but the specimen with the highest number of goblet cells was selected, photographed and then analyzed.
Statistical analysis.
Data were expressed as mean ±SD, n = 5 unless specified otherwise. Differences between groups were determined by using one-way ANOVA with post-hoc testing by Fisher's protected least significant difference (PLSD). Statview-J 4.1 software (Abacus Concepts, Berkeley, CA) was used on an Apple Macintosh computer. Differences were considered significant at a level of P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Plasma zinc concentrations were 21.89 ± 2.72 and 15.88 ± 1.16 µmol/L in AL and PF rats, respectively (n = 5, P < 0.01). In ZD rats, the concentration was 3.00 ± 2.67 µmol/L (n = 5, P < 0.01 vs. both AL and PF rats). Administration of L-NAME to ZD rats did not affect zinc concentration (3.15 ± 0.87 µmol/g, P > 0.1. The concentration of zinc in the intestine did not differ among groups (data not shown).
As shown in Figure 2
, the relative concentration of MT-1 mRNA was 3.02 ± 0.44
amol/µg RNA or 4.89 ± 0.96 amol/µg RNA in the intestine of AL
and PF rats, respectively.The level was lower in ZD rats by 47 and 38%
compared with AL and PF rats, respectively. There were no significant
differences in the relative concentration of MT-1 mRNA between ZD-NAME
(+) and (-) rats.
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Messenger RNA of iNOS was clearly detected by RT-PCR in the intestine
of ZD-NAME (+) and (-) rats, but not in AL and PF rats (Fig. 3
).
|
As shown in Figure 4
, apoptotic-positive cells could be seen only in the tip of the
intestinal villi of AL rats. The cells were sparse in the villi of PF
rats. However, apoptotic-positive cells were identified from the crypt
to the tip in ZD-NAME (-) rats. The number of apoptotic-positive cells
in a viewing field with 10 villi (n = 3) was
significantly higher in ZD-NAME (-) rats (353 ± 36) than in AL
and PF rats (24 ± 22 and 1 ± 2; P <
0.001for both). Morphologically, most of the apoptotic-positive cells
were absorptive enterocytes. Treatment with L-NAME in ZD
rats reduced the number to 21 ± 10 (P < 0.001
vs. ZD-NAME (-) rats).
|
. The villi were more pointed toward the tip, and the brush border
of the villi was flawed in ZD rats (Fig. 5
C, D)
compared with those in AL and PF rats (Fig. 5
A,
B). In particular, hyperplasia of the goblet cells was
observed in ZD-NAME (-) rats. The number of goblet cells in a viewing
field with 10 villi (n = 3) was significantly higher in
ZD-NAME (-) rats (294 ± 5) than in AL and PF rats (148 ±
21 and 178 ± 25; P < 0.05, respectively).
Treatment with L-NAME in ZD rats reversed the number to
134 ± 3 (P < 0.05 vs. ZD-NAME (-) rats).
|
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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and 1998a
) and those of others (Huber and Cousins 1988
). Metallothionein mRNA expression is closely regulated by
the dietary supply of zinc, and the levels in some tissues are direct
reflections of zinc intake (Blalock et al. 1988
,
Huber and Cousins 1988
). MT-1 mRNA level was similar in
ZD rats with or without L-NAME treatment, suggesting that
this compound did not affect zinc status.
Three NOS isoforms (Fig. 6
) have been identified, i.e, the neuronal NOS, iNOS and endothelial
NOS; all three cell types, including intestinal nerves, macrophages
(recruited and resident) and intestinal vessels, are present in the
intestine. As shown in this and previous studies (Cui et al.
1997
), iNOS mRNA could be detected in the intestine of ZD rats
but not in normal (AL) or food-restricted (PF) rats. It is not
surprising that administration of L-NAME did not affect
iNOS mRNA abundance because it inhibits NOS pharmacologically only by a
competitive mechanism. The activity ratio of iNOS to total NOS was
significantly elevated in ZD rats, whereas L-NAME reduced
it to a level comparable to that in AL rats. However, there was
significantly greater iNOS activity in the ZD rats relative to AL rats,
but not relative to PF rats. There was no significant difference in
total enzyme activity between ZD-NAME(-) and ZD-NAME(+) groups.
Examination of the activity with the pooled samples of whole layers of
the intestine might account in part for the discrepancies. Reportedly,
the total NOS activity was sevenfold higher in the mucosa than in the
neuromuscular layers (Hogaboam et al. 1996
). Local
effects of L-NAME might also play a role. Although oral
administration of the agent is considered to be a systemic route, it is
likely that the concentration of L-NAME was much higher in
the lumen than in the tissue of intestine, which affected NOS in the
mucosa directly. By using an immunohistochemical staining technique to
detect iNOS protein, the positive staining was found to be present
mainly in the basal layer and scattered in the villous cells
(Cui et al. 1997
). Evaluating NOS activity in the mucous
layer and in other layers of the intestine separately may reveal
whether iNOS increases only in the mucous layer and whether
constitutive NOS recovers consequently due to administration of
L-NAME. Interestingly, this study demonstrated that total
activity of NOS was lower in ZD rats than in controls.
|
). Changes in
physiologic function in the intestine and feeding behavior due to
starvation as well as timing for sampling might be responsible for the
increase in PF rats.
When stained for programmed cell death by use of the TUNEL method, the
apoptotic-positive cells were restricted to the villous tips of the
normal intestine (Gavrieli et al. 1992
), which
demonstrates that DNA fragmentation is the last stage in the terminal
differentiation pathway of the epithelial cells. Our results concerning
apoptotic-positive cells in the normal intestinal epithelium (AL rats)
were consistent with the findings reported by Gavrieli et al. (1992)
and others (Leonard and Aret 1996
). As also
shown in this study, the apoptotic-positive cells were distributed
throughout the whole epithelial layer of the intestine in ZD rats and
the numbers were several fold higher than that in normal intestine,
consistent with the previous observation that zinc deficiency increased
apoptosis in the intestine (Elmes and Jones 1980
).
Treatment of rats with L-NAME abrogated this phenomenon,
limiting the apoptotic-positive cells to the villi tips and
dramatically reducing the number to that of the normal intestine. This
observation suggests that NO, especially produced by iNOS, could be
involved in zinc deficiency-induced apoptosis in the intestine of rats
because the level of iNOS was inhibited by L-NAME and also
because the location of iNOS-positive staining (Cui et al. 1997
) was near the position in which apoptotic-positive cells
were present. When NO is produced, it diffuses across cell membranes
freely and equally in all directions with an average half-life of
4 s (Lancaster 1994
). The cells in the immediate
vicinity can be affected because no efficient scavenger mechanism
exists to remove NO before it can become toxic. On the other hand, it
is possible that zinc deficiency resulted in a vulnerability of the
intestinal cells to NO. Interestingly, the apoptotic-positive cells of
the villi were few in food-restricted (PF) rats. This observation
indicates that malnutrition might be interfering with the proliferating
and differentiating processes of intestinal epithelium rather than with
initiating apoptosis.
The present morphologic study revealed goblet cell hyperplasia of the
intestine in ZD rats, which was not observed in our previous study. One
of the explanations for this discrepancy is that the duration of
treatment of rats with a zinc-deficient diet in this study (4 wk) was
much shorter than that in the previous study (7 wk, Cui et al. 1996
).
This suggests that goblet cell hyperplasia associated with zinc
deficiency is a time-dependent phenomenon. The goblet cell hyperplasia
might play a protective role or serve as a marker for intestinal damage
during zinc deficiency. On the other hand, a relative increase of
goblet cells over other villous cells cannot be ruled out. It is
possible that absorptive enterocytes die of apoptosis during zinc
deficiency, whereas the goblet cells tenaciously resist this abnormal
environment. Treatment of ZD rats with L-NAME abrogated the
goblet cell hyperplasia, suggesting that NO plays a role in the
induction of this phenomenon. Further investigations are required to
clarify the mechanisms.
Depletion of dietary zinc damages the intestine. This study was
conducted to determine whether iNOS might play a role in the pathogenic
mechanisms. A question has arisen concerning how to design controls for
multiple physiologic changes, including body weight and appetite loss
caused by zinc-deficiency, and for pharmacodynamics. Conventionally, an
AL group of the same age was used as a control, but dietary amount,
intake of water and body weight differed from the other two groups. The
PF group was generally designed as a control for dietary intake but not
for intake of water or feeding behavior. It has been pointed out that
the use of PF animals as a control may be misleading for investigation
of intestinal metabolism (Park et al. 1985
) because a
meal-eating pattern resulting from food restriction causes both
morphologic and enzymatic changes in the intestine. Furthermore, there
is an intrinsic difference of NOS components among the three groups of
rats as reported in our previous study, i.e., ZD rats already expressed
iNOS, whereas expression was not detectable in the intestine of AL and
PF rats. Reportedly, chronic administration of L-NAME in
normal rats or guinea pigs increased NO release as a result of
compensatory expression of iNOS in the intestine (Miller et al. 1996
). Accordingly, treatment of ZD rats with
L-NAME may provide a remedy for inhibiting NOS, especially
iNOS that was expressed in the intestine, whereas it may initiate a
pathogenic process of inducing iNOS in normal rats. Finally, there was
a substantial problem to be solved in choosing a route by which the
agent could be administered chronically and equally among AL, PF and ZD
groups. Intake of water, even calculated as mL/kg body weight, was
markedly different among the AL, PF and ZD groups. Thus, the present
route could have led to unequal amounts of L-NAME
administered. Administration of the agent through insertion of a
gastric tube or by daily injection is invasive. Therefore, this study
did not use AL and PF rats treated with LNAME as further
control groups.
In conclusion, treatment of ZD rats with L-NAME attenuates damage of the intestine associated with reduction of the activity ratio of iNOS relative to total NOS. The beneficial effects include reversal of increased intestinal vascular permeability, inhibition of goblet cell hyperplasia and apoptosis in the villous layer.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Manuscript received June 1, 1998. Initial review completed July 7, 1998. Revision accepted December 14, 1998.
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74 DNA. Lower panel: the ratio of
target/MIMIC intensities is linearly when plotted against the amount of
target cDNA.
