Departments of 1Human Biology and 2Pharmacology, Maastricht University and 3epartment of Gastroenterology, University Hospital Maastricht, All Nutrition and Toxicology Research Institute Maastricht, 6229 ER Maastricht, The Netherlands
Submitted 26 September 2002 ; accepted in final form 24 April 2003
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ABSTRACT |
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duodenum; epithelium; lipid peroxidation; small intestine; oxidative stress
Oral iron therapy, commonly indicated in people with an elevated risk for developing iron deficiency, such as children, adolescents, the elderly, (pregnant) women, vegetarians, and athletes, may induce considerable gastrointestinal (GI) side effects, such as nausea, vomiting, heartburn, abdominal pain, and constipation, in contrast to intravenous iron treatment (2). Iron-induced oxidative damage in the intestine after oral ingestion of iron supplements may, in part, be responsible for these GI side effects. Additionally, oral iron therapy in patients suffering from GI disorders associated with an impaired mucosal or systemic antioxidant capacity, such as inflammatory bowel disease or celiac disease, may attribute to the development of major oxidative damage (8, 13, 16, 19, 27).
Oral iron supplements were shown to aggravate lactic acid-induced damage to the barrier function of the small intestine epithelium (9) and to promote colon tumorigenesis in mice (22) and inflammatory processes in rats with colitis (1, 20). Moreover, electron paramagnetic resonance spectroscopy in rats in vivo showed that oral iron therapy with ferrous sulfate resulted in iron-mediated oxidative stress through hydroxyl radicals in the small intestine. This stress resulted in a decrease in cell turnover, shortening of microvillus height, and partial or complete erosion of the microvilli in the duodenum (24). Epidemiological studies showed a role of dietary iron in the development of colon cancer (26). Additionally, ROS may induce development of several forms of cancer (4, 18).
Because of the complexity of the direct measurement of oxidative damage in the lumen of the human small intestine in vivo, no data are available regarding the effects of clinically prescribed amounts of iron on oxidative damage in the human intestine in vivo. Also, little is known about the antioxidant defense mechanisms against ROS production in the intestine. For this purpose, we developed a perfusion technique to directly quantify oxidative damage and antioxidant capacity in vivo in the small intestine in healthy adults. This study aimed to determine iron-induced oxidative damage provoked by a single clinical dosage of ferrous sulfate in the small intestine in humans and to obtain more information about the antioxidant defense mechanisms in the human small intestine in vivo.
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MATERIALS AND METHODS |
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Six healthy nonsmoking volunteers (25 ± 5 yr, 3 males, 3 females) without a history of a GI disorder participated in the present study. Volunteers were in good physical health at the time of participation as checked by medical interview. All subjects gave their written informed consent before participation. The study protocol was approved by the Ethics Committee of the University Hospital Maastricht, Maastricht, The Netherlands.
Protocol
After an overnight fast, a perfusion catheter was ingested orogastrically by each volunteer. The catheter consisted of three silicon tubes connected to a rubber balloon on the distal end of the catheter. One of the tubes called the sample port had three small openings at 5 cm proximal of the distal end of the catheter to enable fluid sampling from the small intestine at the level of the distal end of the catheter. The other two tubes were connected in an open loop with the balloon to enable fluid and air transport in and out of the balloon. A fourth lumen had a single opening at 45 cm from the distal end. This port, referred to as the perfusion port, served to administer a test solution directly into the duodenum.
After installation of the catheter tip into the stomach, the rubber balloon
was filled with 1.5 ml of mercury. Subjects were instructed to remain in the
supine position on the right hand side with the upper body lifted 45° and
the feet raised 20 cm until the balloon entered the duodenum. Positioning of
the catheter was checked with continuous pH-registration at the tube tip. When
the pH shifted from low (12.5) to
46, the mercury was
aspirated and the balloon was subsequently inflated with 15 ml of air.
Subjects were instructed to stay in the supine position on the back with the
upper body lifted 45°. The catheter propagated subsequently by the
peristaltic movements of the GI tract. Catheter positioning was successful
when the perfusion port was placed
510 cm distally from the
pylorus. This was checked by a second pH-registration channel, which was
located at 5 cm proximal from the perfusion port. After positioning, the
balloon was deflated to stop it from progressing in the GI tract. Throughout
the experiment, the pH of the second pH channel was continuously registered to
ensure proper positioning of the perfusion port. After the perfusion
experiment, subjects removed the catheter by pulling it gently.
Via the perfusion port, a saline solution was injected at 10 ml/min during 195 min. This period was necessary to reach steady-state conditions as was observed in pilot experiments (data not shown). Subsequently, a 300-ml saline solution containing 80 mg elemental iron as ferrous sulfate (Merck, Darmstadt, Germany) was perfused at 10 ml/min during 30 min. Finally, a saline solution was perfused again for 60 min to rinse out the iron solution and intestinal secretions.
Intestinal fluid samples were obtained by using the lever properties of the catheter. After an initial adaptation period of 195 min, intestinal fluid content was levered through the sample lumen into a beaker placed in melting ice. Intestinal fluid excretions were pooled over 15-min intervals and stored at -80°C until analysis.
Biochemical Analysis of Intestinal Fluid Samples
Thiobarbituric acid reactive substances assay. The determination of thiobarbituric acid reactive substances (TBARS) was based on the formation of a colored adduct of malondialdehyde (MDA) with 2-thiobarbituric acid (TBA). A 100 µl sample was added to 900 µl of a reagent [containing 0.12 M TBA, 0.32 M o-phosphoric acid, 0.68 mM butylated hydroxytoluene (BHT), and 0.01% (mass/vol) EDTA]. The mixture was incubated for 1 h at 100°C in a water bath. After cooling, the MDA-products were extracted with 500 µl butanol. Then 30 µl of the butanol layer were injected on an HPLC system (Agilent, Palo Alto, CA) equipped with a fluorescence detector, set on an excitation wavelength of 532 nm and an emission wavelength of 553 nm, and a Nucleosil C18 column (150 x 3.2 mm) particle size 5 µm (Supelco, PA). Samples were eluted with 65% (vol/vol) 25 mM PBS (pH 4.8) and 35% (vol/vol) methanol. A calibration curve was constructed by using malonaldehyde bis(diethylacetal) as standard.
Antioxidant capacity assay. The antioxidant capacity assay was carried out as described previously (24a) with some modifications. The 2,2'-azinobis-(3-ethylbenzthiazoline-bsulfonate (ABTS·+) was produced by incubating a solution of 0.23 mM ABTS and 2.3 mM 2,2'-azobis-(2-amidinopropane) hydrochloric acid (ABAP) in 100 mM sodium phosphate buffer pH 7.4 at 70°C until the absorption of the solution at 734 nm was between 0.680 and 0.720. Deproteination was done by mixing the sample with an equal volume of a solution of 10% (wt/vol) TCA. In the reaction of antioxidants with the blue/green ABTS·+, the blue/green color disappears. This decolorization after 5 min is determined spectrophotometrically at 734 nm. The reduction in absorbance is related to that of Trolox, a synthetic, hydrophilic vitamin E analog, which gives the Trolox equivalent antioxidant capacity (TEAC) value. The TEAC value is calculated as molar Trolox equivalents of the sample.
Glutathione assay. Total glutathione was determined by using the
recycling method (25). To 50
µl of a 100 µl sample of a 0.4 mM NADPH/0.3 mM
5,5'-dithiobis(2-nitrobenzoic acid) solution in 143 mM PBS (pH 7.4)
containing 6.3 mM EDTA, is added in a 96-well plate. The reaction is then
started by adding 50 µl of a solution containing 4 U/ml glutathione
reductase. The increase of the absorption (A/min) is followed for 2 min
at 405 nm. The
A/min of the samples is compared with that of the
calibrators and the concentration of total glutathione is calculated.
Uric acid assay. Uric acid was determined in ultra filtrates by using an HPLC method described previously (15) with minor modifications. The sample was filtered over a 3-kDa filter unit (Centrex UF 0.5; Schleicher & Schuell, Dassel, Germany) by centrifugation for 45 min at 14,000 g and 4°C. Then 25 µl of the filtrates was analyzed on the HPLC. The system (Agilent) consisted of a Hypersil BDS C-18 end-capped column (125 x 4 mm), particle size 5 µm (Agilent), with a mobile phase of 5 mM sodium phosphate buffer (pH 3.3). Detection was performed by using a variable wavelength detector set on a wavelength of 292 nm. Concentrations of uric acid were calculated by using a calibration curve of uric acid.
Statistics
Differences were assessed by univariate ANOVA using SPSS version 10.0 software. Differences were regarded as significant when P < 0.05. Multiple comparisons, used to localize time differences, were adjusted with the least significant difference.
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RESULTS |
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TBARS levels were significantly higher at 15, 30, 45, and 60 min after the start of the ferrous sulfate perfusion compared with the baseline level measured at t = 0 (P < 0.05) and returned to baseline levels after 90 min (Fig. 1).
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The TEAC of the intestinal fluid was significantly higher at 30 and 45 min after the start of the ferrous sulfate perfusion compared with the baseline level measured at t = 0 (P < 0.05) (Fig. 2).
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Median alkaline phosphatase concentrations of the perfusate samples did not differ significantly between t = 0 and t = 30 min (502 ± 377 and 510 ± 363 U/l, respectively).
Uric acid concentrations of the perfusate samples were not significantly different from baseline during the first 75 min after the start of the ferrous sulfate perfusion (Table 1). At t = 90, uric acid was statistically lower than baseline values (P < 0.05).
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Glutathione concentrations in the perfusate samples were under the detection limit of the equipment (<0.10 µM).
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DISCUSSION |
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TBARS concentrations in the intestinal fluid samples after the iron
challenge are roughly 35 times higher than those observed in plasma in
healthy adults. This is a highly interesting finding in the view of the
dilution of the intestinal fluid samples. In pilot experiments, we measured
the dilution of the intestinal fluid with the saline or saline/iron solution
in the intestinal segment by the addition of an inert marker, lactulose, to
the ingoing solution and subsequent measurement of the lactulose concentration
in the perfusate samples. A dilution of 100-fold was found in all
samples. This dilution was constant throughout the experiment (data not
shown). In view of the fact that sample dilution did not change during the
experiment, independently of the presence or absence of iron, intestinal fluid
dilution did not interfere with the results of the study and hence, was not
measured in the present study.
On average, 15 mg of iron is present in the daily diet. Approximately
2.5 mg iron entering the intestine will be absorbed. Approximately 1% of the
dietary iron remains in the lumen in a form likely to participate in free
radical generation. During oral iron therapy in adults using dosages of
60195 mg iron/day, this proportion remains the same, resulting in a
much larger amount of iron available for Fenton reactions. During Fenton
chemistry, ferrous iron catalyzes the formation of hydroxyl radicals in the
presence of superoxide anion radicals and hydrogen peroxide, both produced as
a result of normal metabolism. Hydroxyl radicals are highly reactive and can
inflict damage to any biologic molecule in its direct environment. The free
electron can transfer to another molecule and hence, initiate a cascade of
damaging reactions. Previously, it was shown that the free radical generating
capacity from feces in healthy volunteers was markedly increased after iron
supplementation (19 mg/day iron as ferrous sulfate for 14 day)
(14). However, this was
measured in fecal samples ex vivo in an in vitro assay. This does not
necessarily reflect the conditions from the present study in which the true
effects of iron ingestion in the small intestine in vivo were investigated.
The results of the present study confirm the finding that iron ingestion leads
to an increase in free radical production in the gut. The model used in the
present study mimics ingestion of an oral iron supplement, which dissolves in
the gastric juice and is subsequently released gradually into the small
intestine.
From the data (Fig. 1) it is clear that oxidative damage occurs immediately after iron administration into the small intestine. This damage continues during saline perfusion after the ferrous sulfate perfusion. This is likely due to the low perfusion flow in the experimental setup. After discontinuation of the iron perfusion, it takes some time to rinse out all of the iron and the intestinal juices secreted during the iron perfusion from the small intestine. After 60 min of saline perfusion, no iron-induced rise in TBARS is observed in the intestinal fluid samples anymore. These data are in line with those of a rat study in which it was shown that high dosages of iron supplements induce lipid peroxidation in the colon (21). Any effects of iron supplements in the small intestine were not investigated in that study.
The lipid peroxidation observed in the present study preceded a marked rise in TEAC (Fig. 2). From this, it may be concluded that iron-induced oxidative stress induces the release of an antioxidative component in the small intestine. As for the TBARS concentrations, the TEAC values are also remarkably high, especially in view of the previously mentioned dilution of the intestinal fluid samples with the ingoing solution. The data of the present study indicate that a powerful physiological mechanism in the small intestine protects against iron-induced oxidative damage. In view of the fast release of the antioxidant(s), it is unlikely that the iron-induced oxidative damage directly induced production of an antioxidative agent, which is subsequently released into the lumen. It is more likely that an antioxidant already present in a storage compartment in the small intestine was released on oxidative damage. In the present study, all samples were completely deproteinized during the TEAC assay. Sample deproteination is routine practice during the TEAC assay to reduce the analysis of variance. In pilot experiments, we observed that sample deproteination did not affect the average TEAC values compared with duplicate nondeproteinized samples. Hence, the high antioxidant capacity was exclusively caused by a nonprotein species. This excludes a role for enzymatic antioxidants or metallothionines in the observed rise in antioxidant capacity.
The rise in TEAC might be explained by lysis of epithelial or mucosal cells. If the membranes of intact cells were disrupted, its cellular content was excreted into the lumen. As a result, the antioxidant capacity would predominantly originate from the cytosolic antioxidant capacity of the epithelial cells. However, this would have been accompanied by a rise in total alkaline phosphatase, which we did not observe. Also, removal of intact epithelial cells present in the intestinal fluid samples by centrifugation did not significantly affect the antioxidant capacity of the samples (data not shown). This suggests that the strong rise in TEAC after iron-induced oxidative stress was not due to cell lysis but was brought about by a specific antioxidant mechanism.
Data regarding the nonprotein antioxidant capacity of the small intestine are scarce. It was shown in a rat study (17) that the small intestine has a powerful reducing capacity, which is protective against triglyceride hydroperoxides. The underlying mechanism or the site of action (luminal, mucosal, or epithelial) was not identified. In the present study, glutathione and uric acid were determined in the intestinal fluid samples in an attempt to identify the origin of the rise in antioxidant capacity after iron perfusion. Due to the intestinal fluid dilution by the perfusing solution, glutathione concentrations were below the detection threshold of the equipment used. Uric acid, a powerful antioxidant present in lung- and intestine epithelial lining fluid (12), was present in low amounts not affected by the iron intervention. Vitamin C, present in low amounts in most samples, did not show a correlation with antioxidant capacity (data not shown). Hence, neither of these antioxidants was responsible for the rise in antioxidant capacity after the iron challenge in the present study.
The perfusion model to investigate oxidative stress and antioxidant capacity in the human small intestine enabled us to determine the effects of a prooxidative agent over a standardized segment of the small intestine. A major advantage of the technique is the rather noninvasive protocol of the catheter positioning, because this occurred using normal peristaltic movements. The technique is applicable to healthy volunteers as well as patients. The perfusion technique can only be used to study a limited segment of the proximal small intestine. Hence, results may not be directly extrapolated to the entire small intestine. However, due to a decreasing nutrient concentration along the small intestine, nutrient-associated oxidative damage probably takes place predominantly in the proximal small intestine. The rate of perfusion needs to be as low as possible to avoid effects of the perfusion experiment on normal functioning of the small intestine. Because of the low perfusion flow, it is not possible to monitor acute, short-term responses in the small intestine. Additionally, sampling occurs at 15-min intervals to ensure sufficient sample collection during each collection interval. The perfusion technique provides the unique feature to study oxidative damage and antioxidant capacity in secretions of the small intestine in vivo in humans under strictly standardized conditions. This model is applicable to a variety of intestinal disorders associated with a disturbance in the equilibrium between oxidative stress and the antioxidant defense network. Increased oxidative stress occurs in acute and chronic inflammation, certain drugs, and nutrients. Decreased antioxidant capacity may be present in disorders of mucosal integrity and chronic malabsorption disorders associated with antioxidant deficiencies (8). A disturbance in the equilibrium between oxidative stress and the antioxidant defense network in the small intestine may be an important factor in the pathophysiology of these disorders. Additionally, the perfusion technique can be used to evaluate the efficiency of nutritional antioxidant interventions in restoring a sufficient antioxidant network against oxidative attack in the small intestine.
Iron ingestion induced Fenton-driven oxidative stress in the human small intestine, followed by a massive rise in total antioxidant capacity in the lumen. This rise may reflect the release of a powerful, yet unidentified, nonprotein antioxidant. Future work should focus on identification of this antioxidant and on the possibilities to increase the antioxidant capacity against oxidative stress with nutritional interventions in disorders related to a high intestinal oxidative stress or a low intestinal antioxidant defense network.
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FOOTNOTES |
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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.
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REFERENCES |
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