Disturbances of the GIT are encountered in wild and captive dolphins, and damage to the gastric mucosa, including ulcers, has been reported.1,2 Possible causative agents for gastric ulcers include Helicobacter spp, nematodes, and other parasites. It is difficult to diagnose gastric ulcers in captive dolphins, and diagnosis is currently dependent on endoscopic evaluation. However, this poses many challenges because endoscopy requires expensive equipment, dolphins typically must be removed from the water for the evaluation, and the complex multiple-chambered stomach of dolphins makes it difficult to examine the entire stomach and proximal portion of the small intestine.
Permeability tests are used to evaluate integrity of the mucosal barrier along the entire length of the GIT of infant and adult humans and various other animals. Commonly used indicators include xylose, mannitol, and rhamnose.3,4 Regardless of the marker or markers used and area of the GIT of concern, the procedure involves oral administration of a dose of a permeability indicator or indicators and measurement of concentrations in serum or urine. Serum or urine concentrations are low when the mucosa is intact (impermeable), whereas there is a direct relationship between marker concentrations and severity of mucosal inflammation and ulceration. The use of specific permeability markers has allowed clinicians to detect the prevalence and severity of mucosal damage in regions of the GIT that are not accessible by endoscopy.
The sucrose permeability test has been validated as an indicator of gastric ulcers and other gastric pathologic conditions in humans and companion animals.5–7 It also has been used in horses,8 which have a high incidence of gastric ulcers, particularly when in training.9 Because healthy gastric epithelium is not permeable to sucrose, and because sucrase rapidly degrades sucrose that enters the small intestine, sucrose serves as an excellent marker for evaluating integrity of the gastric mucosa but not for evaluating permeability of the small intestine and more distal regions of the GIT.10 The test is sufficiently sensitive so that even slight mucosal damage not evident endoscopically can be detected.11 The sucrose permeability test has been used to study gastric damage caused by administration of non-steroidal antiinflammatory drugs to humans12,13 and dogs.6
The objective of the study reported here was to determine the feasibility of measuring concentrations of sucrose in the serum of dolphins after oral administration of a sucrose solution. Our long-term goal was to develop a noninvasive method that can be used routinely to screen captive dolphins for prevalence and severity of gastric ulcers. This would allow attending veterinarians to better monitor group health, identify specific dolphins for endoscopic evaluation, and determine the efficacy of treatment regimens.
Sucrose concentrations in serum or urine are typically measured by use of electrochemical HPLC analysis.14 To achieve concentrations of sucrose in serum and urine that are detectable by HPLC, it is necessary to administer large doses of sucrose (eg, 50 g for dogs that weigh 20 to 40 kg6 and 454 g for horses that weigh 454 to 500 kg15). However, it is not known whether disaccharidase activity of dolphins is sufficient to hydrolyze sucrose entering the small intestines, and a high dose of sucrose could pose health risks to dolphins by causing osmotic diarrhea. Therefore, the analytical approach for the study reported here combined HPLC with mass spectroscopy (ie, LC-MS-MS) to increase the sensitivity and accuracy of detection so a lower dose of sucrose could be used.
Materials and Methods
Animals—Eight captive adult bottlenose dolphins (Tursiops truncatus) housed at Marine Life Oceanarium, Gulfport, Miss, were used in the study. None of the dolphins had overt signs of gastric ulcers or other GIT disorders, and all were considered to be in good health by the attending veterinarian. However, endoscopy was not used to evaluate the gastric mucosa of the dolphins. The exception was 1 dolphin that was anorectic and had stomach contents that were suggestive of a gastric disturbance (burgundy red and containing proteinaceous clumps and WBCs). Subsequent endoscopic evaluation of this dolphin revealed gastric ulcers, which were treated by use of a combination of omeprazole magnesiuma and sucralfate.b Sucrose permeability testing was conducted in this dolphin shortly before the endoscopic procedure by oral administration of 25 g of sucrose.
Experimental design—Monthly health assessments were conducted on the dolphins. Assessments included a CBC, serum biochemical analysis, and evaluation of gastric fluid samples. Measurement of serum sucrose concentrations was performed during each monthly health assessment.
Dolphins received the first meal of the day at 8 AM and the last meal of the day at 4 PM. Each month for 9 months, up to 4 of the dolphins were assigned to receive a sterile solution containing 25 or 50 g of sucrosec in 500 mL of water; the sucrose solution was administered via gastric intubation. The remaining dolphins were used as control animals for that month and were not administered sucrose. During the study, all 8 dolphins were orally administered sucrose solutions 4 times (2 doses of 50 g and 2 doses of 25 g). The procedures used required that dolphins were trained for gastric intubation and blood collection. A volume of 500 mL was selected for the sucrose administration because it is comparable on a per-kilogram basis with the 4-L volume administered to horses and should have been sufficient to expose the lining of the entire stomach of the dolphins to the dissolved sucrose. Food coloring was added to the sucrose solution to enable investigators to monitor regurgitation.
Collection of blood samples—Blood samples used for the health assessment and sucrose measurement were collected from the ventral fluke vein before the dolphins were fed the first meal of the day and approximately 45 to 50 minutes after administration of the sucrose solution because this interval between administration and sample collection yields maximum sucrose concentrations in horses.8 Serum was separated and frozen until analyzed for sucrose; samples were analyzed within 2 weeks after collection.
Measurement of serum sucrose concentration— Analysis required a total serum volume of 200 μL, which allows for duplicate assays (100 μL each). Serum was diluted 1:9 in 2-mL microcentrifuge tubes by use of a solution consisting of 90% acetonitriled to 10% water that contained 10 ng of an internal standard (trichlormethiazidee)/μL. Diluted samples were mixed thoroughly and then centrifugedf (18,000 × g for 5 minutes). Supernatant was transferred to 2mL target vials.
Analysis was performed by use of a liquid chromatographg coupled to a mass spectrometer.h For liquid chromatography separation, 100 μL of sample was injected into a 5-μm, 100 × 2.1-mm columni by use of a mobile phase that was programmed for 2 minutes of water and acetonitrile (90:10), followed by a gradient during the next 14 minutes to achieve a final ratio of 5:95. This resulted in elution of sucrose at 1.8 minutes and trichlormethiazide at 8.4 minutes.
After separation, sucrose and internal standard were detected by use of the mass spectrometer operating in the tandem mode. An M/Z value of 365 was used for the precursor ion of sucrose (mass + 1). A product ion with an M/Z value of 182 was generated in the positive tandem mode and used to quantify sucrose. The negative tandem mode was used for detecting trichlormethiazide because of the electronegative chlorine in the molecule. An M/Z value of 379 (mass – 1) was used for the precursor ion, and an M/Z value of 305 for the product ion was used for quantification of trichlormethiazide.
A series of standard solutions was included with each analysis. These were prepared by dilution (1 part of the standard to 9 parts of the diluent), which resulted in sucrose concentrations ranging from 5 to 5,000 pg/μL; each sucrose concentration contained 10 ng of trichlormethiazide/μL. Values were not reliable when concentrations of sucrose were < 50 pg/μL. Resulting data were subjected to linear regression analysis, with a regression coefficient of 0.99 accepted as the lower limit of acceptance. The regression coefficients were routinely ≥ 0.995. Continuous calibrations by use of known sucrose concentrations were included every 10 to 20 samples to assess system performance. When a continuous calibration deviated by ≥ 20% from the actual concentration, the calibration was assessed, and samples were reanalyzed to maintain accuracy. During validation of the analysis protocol, matrix spikes prepared by adding known amounts of sucrose to control serum revealed recoveries of > 90%.
Collection of gastric contents—During the routine monthly health assessment and prior to feeding the first meal of the day and administration of the sucrose solution, each dolphin was intubated, a sample of gastric contents was removed, and pH of the gastric contents was measured. Samples of gastric contents were also examined to detect possible damage to the gastric mucosa on the basis of clarity, color (white, pink, or red), prevalence of proteinaceous clumps, and whether cytologic evaluation revealed evidence of WBCs and bacteria or fungi.
Gastric contents were also collected from 3 dolphins after the last meal of the day had been consumed. Additional samples of gastric contents were collected from those same 3 dolphins at 8 PM, midnight, and immediately before feeding of the first meal on the subsequent day. The pH was immediately recorded for each sample of gastric contents.
Analysis of data—Monthly values for serum sucrose concentration of each dolphin and the pooled value for all dolphins were evaluated by use of a 1-way ANOVA, with differences among dolphins identified by use of the Duncan test.j A similar approach was used to evaluate changes in gastric pH. Values were reported as mean ± SD. For all comparisons, values of P < 0.05 were accepted as significant.
Results
Animals—Administration of 25 and 50 g of sucrose in 500 mL of water did not cause any obvious health problems, as determined on the basis of observations of behavior and appetite. The doses were tolerated, and regurgitation was not evident.
Serum sucrose concentrations in control dolphins—Mean ± SD sucrose concentration in serum collected from control dolphins (n = 35) was 102 ± 95 ng/mL. This sucrose concentration did not differ significantly from the background concentration and did not exceed the minimum concentration that could be reliably detected.
Serum sucrose concentrations after oral administration—Mean ± SD serum sucrose concentration of dolphins administered 25 g of sucrose (n = 16) was 868 ± 452 ng/mL (range, 442 to 1,958 ng/mL; Figure 1). However, this concentration did not differ significantly (P = 0.08) from the concentrations measured in control dolphins. Mean serum concentration after administration of 50 g of sucrose to 16 dolphins (3,798 ± 2,787 ng/mL) was significantly higher than the concentration in control dolphins and in dolphins administered 25 g of sucrose.
Mean ± SD sucrose concentrations in serum samples obtained from 8 captive adult bottlenose dolphins (Tursiops truncatus) 45 minutes after oral administration of 500 mL of water that contained 25 or 50 g of sucrose and in serum samples obtained when the dolphins were not administered sucrose solution. a,b Values with different letters differ significantly (P < 0.05).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Mean ± SD sucrose concentrations in serum samples obtained from 8 captive adult bottlenose dolphins (Tursiops truncatus) 45 minutes after oral administration of 500 mL of water that contained 25 or 50 g of sucrose and in serum samples obtained when the dolphins were not administered sucrose solution. a,b Values with different letters differ significantly (P < 0.05).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Mean ± SD sucrose concentrations in serum samples obtained from 8 captive adult bottlenose dolphins (Tursiops truncatus) 45 minutes after oral administration of 500 mL of water that contained 25 or 50 g of sucrose and in serum samples obtained when the dolphins were not administered sucrose solution. a,b Values with different letters differ significantly (P < 0.05).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Wide intra- and interindividual variation in serum sucrose concentrations was evident for both sucrose doses (Figure 2). Repeating the analysis of samples for each dolphin on various dates and with other sets of serum samples yielded similar concentrations, which indicated that the variation among dolphins and sample dates was not caused by analysis error or by variation attributable to samples being collected and stored before analysis in different months. Instead, the permeability of sucrose appeared to vary among dolphins and differed among months for specific dolphins. The serum sucrose value after administration of 25 g of sucrose to the dolphin that was determined endoscopically to have gastric ulcers was 998 ng/mL. This was not the highest concentration for that dose, but it was higher than the concentration of 665 ng/mL measured the following month after the dolphin was treated with sucralfate and omeprazole magnesium.
Mean ± SD sucrose concentrations in serum samples obtained from 8 captive dolphins 45 minutes after oral administration of 500 mL of water that contained 25 g of sucrose (black bars) or 50 g of sucrose (gray bars).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Mean ± SD sucrose concentrations in serum samples obtained from 8 captive dolphins 45 minutes after oral administration of 500 mL of water that contained 25 g of sucrose (black bars) or 50 g of sucrose (gray bars).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Mean ± SD sucrose concentrations in serum samples obtained from 8 captive dolphins 45 minutes after oral administration of 500 mL of water that contained 25 g of sucrose (black bars) or 50 g of sucrose (gray bars).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Gastric pH—Gastric pH was highest immediately after the last meal of the day was consumed, but was significantly lower 4 hours later (Figure 3). The low values between midnight and feeding of the first meal at 8 AM on the subsequent day indicated that the gastric mucosa can be exposed daily for at least 8 hours to a pH of ≤ 2.0.
Mean ± SD pH of gastric contents collected from 3 captive dolphins immediately after feeding of the last meal of the day (4 PM), 4 hours later (8 PM), 8 hours later (midnight), and before feeding of the first meal of the subsequent day (8 AM). a-c Values with different letters differ significantly (P < 0.05).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Mean ± SD pH of gastric contents collected from 3 captive dolphins immediately after feeding of the last meal of the day (4 PM), 4 hours later (8 PM), 8 hours later (midnight), and before feeding of the first meal of the subsequent day (8 AM). a-c Values with different letters differ significantly (P < 0.05).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Mean ± SD pH of gastric contents collected from 3 captive dolphins immediately after feeding of the last meal of the day (4 PM), 4 hours later (8 PM), 8 hours later (midnight), and before feeding of the first meal of the subsequent day (8 AM). a-c Values with different letters differ significantly (P < 0.05).
Citation: American Journal of Veterinary Research 67, 6; 10.2460/ajvr.67.6.931
Discussion
The concentration of sucrose in urine after oral administration of a dose is considered to be a valuable screening tool for evaluating gastric permeability in children because of the high specificity (98%) and positive predictive value (96%) for disruption of the gastric mucosa.16 Analysis of the findings reported here revealed that measurable, and sometimes high, concentrations of sucrose were detectable in the sera of 8 captive dolphins 45 minutes after administration of doses of 25 and 50 g of sucrose by gastric intubation following an overnight period without food. Furthermore, the negligible concentrations of sucrose detected in the 35 samples of sera collected from control dolphins were all at or less than the limits of reliable detection and were considered to be within the background concentrations. In light of this, it was decided that an initial serum sample (before administration of sucrose) was unnecessary.
Sensitivity of the LC-MS-MS (detection limits as low as 5 ng/mL) exceeds that of HPLC methods (detection limits in the range of micrograms per milliliter). This allows for the administration of a lower dose of sucrose without compromising the ability to detect and evaluate the severity of damage to the gastric mucosa. The ability to obtain and analyze serum will allow the evaluation of gastric permeability to be included as a component of routine health assessments of captive dolphins. Moreover, the collection of blood samples and harvest of serum are less intensive and can be accomplished faster than the collection of urine. Specifically, the use of urine samples requires the bladder to be catheterized for complete evacuation before administration of sucrose and again later for collection of the sample.
More than 90% of the dolphins administered 50 g of sucrose had serum sucrose concentrations that were higher than those measured in sera of horses with no visible damage to the gastric mucosa (ulcer score of 0).8 Moreover, the serum sucrose concentrations measured in some of the dolphins after the 50-g dose were comparable to values measured after administration of 500 g of sucrose to horses with endoscopic ulcer scores of 3 (scale of 0 to 3, with 3 considered to be the most severe ulcers).8 The 10-fold higher dose administered to horses, compared with the dose administered to the dolphins, exceeds the 4- to 5-fold difference in body mass and, hence, blood volume. A limitation of the study reported here was that serum sucrose concentrations measured in the dolphins were not correlated with direct observations of the gastric mucosa, which is in contrast to the information reported for horses.8 Interestingly, the serum sucrose concentration after administration of 25 g to the dolphin that was anorectic and in which endoscopic evaluation revealed gastric ulcers was not significantly different from the mean concentration for all dolphins administered 25 g of sucrose and was only approximately half of the highest value measured. It is currently not known whether the high and variable serum sucrose values measured in the captive dolphins used for this study are indicative of a high incidence of mucosal inflammation or ulcers.
An alternate explanation is that the values measured in the captive dolphins may have been representative of normal gastric mucosa. Direct correlations of serum sucrose concentrations and observations of the gastric mucosa are needed to determine which dose of sucrose (ie, 25 or 50 g) has the greater diagnostic value. Doses higher than 50 g were not evaluated in the study because there was concern that disaccharidase activity in the small intestine may not have been sufficient, and a higher dose could have led to osmotic diarrhea.
Performance horses have a high incidence of damage to the gastric mucosa, with gastric ulcers in 70% to 94% of horses in training and in 100% of actively racing horses.17 In contrast, gastric ulcers develop in a small percentage of free-ranging horses that are allowed to graze and only rarely would have stomachs that do not contain food. An influence of feeding regimen is evident by subjecting horses to alternating periods of feeding and nonfeeding for 96 hours, which exposes the gastric mucosa to low pH during the nonfeeding periods, induces gastric ulcers,18 and, in our experience, results in higher serum sucrose concentrations. Observations of wild dolphins suggest that they are opportunistic feeders, eating when food is available but rarely going extended periods between meals. Also, the stomachs of some stranded dolphins contain food, but < 20% have evidence of mucosal ulcers.2 Similar to performance horses, captive dolphins are fed several meals during the day (between 8 AM and 4 PM), but they are not fed during the night. Gastric pH measured in the captive dolphins used for the study reported here was often < 2 after the overnight period without food. Moreover, the low values at midnight and before feeding of the first meal at 8 AM on the subsequent day indicated that the gastric mucosa of captive dolphins can be exposed daily for at least 8 hours to a pH ≤ 2.0. If the high serum sucrose values that were measured in the captive dolphins administered 50 g of sucrose are indicative of a high incidence of damage to the gastric mucosa, it will be necessary to determine whether there are relationships among feeding frequency, gastric pH, mucosal integrity, and serum sucrose concentration.
If the current feeding practices for captive dolphins were found to be associated with increased prevalence of mucosal damage, a preventive approach that could be explored would be to maintain intragastric pH above a value that does not induce ulcers. This may be accomplished by simply avoiding long overnight periods without food or including buffering agents in the last meal of the day. The most commonly used therapeutic for horses with gastric ulcers19 is the proton-pump inhibitor omeprazole.k Although effective at increasing gastric pH of horses, omeprazole is costly, requires multiple doses over an extended period, and does not necessarily address the cause of the ulcer, particularly when it is related to alternating periods of feeding and nonfeeding.
Additional studies will be necessary to validate the use of serum sucrose concentrations to screen for prevalence and severity of damage to the gastric mucosa among populations of captive dolphins. Most of the gastric damage observed in other species of dolphins is located in the main stomach,1 which is lined by squamous epithelium, and consequently is subject to ulcers. This is similar to findings in other species of animals, such as equids.18 Because the main stomach of dolphins is accessible for endoscopic evaluation, it should be possible to correlate endoscopic observations with serum sucrose measurements and determine the dose of sucrose with the highest diagnostic value. The relationship between time after administration for collection of samples and serum sucrose concentrations will also need to be defined for dolphins to establish the interval after sucrose administration that maximizes sensitivity and predictive value.
Validation of a sensitive method to monitor gastric permeability will allow veterinarians and other people who care for captive dolphins to routinely screen for evidence of ulcers or damage to the gastric mucosa. Moreover, the availability of a noninvasive method will provide opportunities to better understand the incidence and severity of gastric ulcers in captive dolphins; to search for possible causes of gastric ulcers, such as bacterial and fungal infections, disease, parasites, or diet and feeding regimens; and to evaluate the efficacy of therapeutic protocols.
ABBREVIATIONS
| GIT | Gastrointestinal tract |
| HPLC | High-performance liquid chromatography |
| LC-MS-MS | Liquid chromatography–mass spectrometry–mass spectrometry |
| M/Z | Mass to charge |
Prilosec, Proctor and Gamble, Cincinnati, Ohio.
Carafate, Hoechst, Wiesbaden, Germany.
BP-220, Fisher Biotech, Fair Lawn, NJ.
A996, Fisher Biotech, Fair Lawn, NJ.
T-1016, Sigma-Aldrich, St Louis, Mo.
Eppendorf centrifuge 5415, Brinkman Instruments Inc, Westbury, NY.
HP 1100 liquid chromatograph, Agilent Technologies, Wilmington, Del.
Esquire mass spectrometer, Bruker Daltronics, Billerica, Mass.
Allure PFP propyl 5 μm, 100 × 2.1-mm column, Restesk Corp, Bellefonte, Pa.
Proc GLM, SAS, version 8.1, SAS Institute Inc, Cary, NC.
GastroGard, Merial, Duluth, Ga.
References
- 1↑
Harper CM, Dangler CA & Xu S. et al. Isolation and characterization of a Helicobacter sp. from the gastric mucosa of dolphins, Lagenorhynchus acutus and Delphinus delphis. Appl Environ Microbiol 2000;66: 4751–4757.
- 2↑
Abollo E, Lopez A & Gestal C, et al. Long-term recording of gastric ulcers in cetaceans stranded on the Galician (NW Spain) coast. Dis Aquat Organ 1998;32: 71–73.
- 3
Abazia C, Ferrara R & Corsaro MM, et al. Simultaneous gaschromatographic measurement of rhamnose, lactulose and sucrose and their application in the testing of gastrointestinal permeability. Clin Chim Acta 2003;338: 25–32.
- 4
Steiner JM, Williams DA, Moeller EM. Development and validation of a method for simultaneous separation and quantification of 5 different sugars in canine urine. Can J Vet Res 2000;64: 164–170.
- 5
Giofre MR, Meduri G & Pallio S, et al. Gastric permeability to sucrose is increased in portal hypertensive gastropathy. Eur J Gastroenterol Hepatol 2000;12: 529–533.
- 6↑
Meddings JB, Kirk D. Olson ME. Noninvasive detection of nonsteroidal anti-inflammatory drug-induced gastropathy in dogs. Am J Vet Res 1995;56: 977–981.
- 7
Meddings JB, Sutherland LR & Byles NI, et al. Sucrose: a novel permeability marker for gastroduodenal disease. Gastroenterology 1993;104: 1619–1626.
- 8↑
Hewetson M, Cohen ND & Love S, et al. Serum sucrose concentration in blood: a new method for assessment of gastric permeability in horses with gastric ulceration. J Vet Intern Med 2006;in press.
- 9↑
Rabuffo TS, Orsini JA & Sullivan E, et al. Associations between age or sex and prevalence of gastric ulceration in Standardbred racehorses in training. J Am Vet Med Assoc 2002;221: 1156–1159.
- 10↑
DeMeo M. Sucrose permeability as a marker for nonsteroidal anti-inflammatory gastroduodenal injury: how sweet is it? Nutr Rev 1995;53: 13–16.
- 11↑
Erlacher L, Wyatt J & Pflugbeil S, et al. Sucrose permeability as a marker for NSAID-induced gastroduodenal injury. Clin Exp Rheumatol 1998;16: 69–71.
- 12
Smecuol E, Bai JC & Sugai E, et al. Acute gastrointestinal permeability responses to different non-steroidal anti-inflammatory drugs. Gut 2001;49: 650–655.
- 13
Kawabata H, Meddings JB & Uchida Y, et al. Sucrose permeability as a means of detecting diseases of the upper digestive tract. J Gastroenterol Hepatol 1998;13: 1002–1006.
- 14↑
Cox MA, Iqbal TH & Cooper BT, et al. An analytical method for the quantitation of mannitol and disaccharides in serum: a potentially useful technique in measuring small intestinal permeability in vivo. Clin Chim Acta 1997;263: 197–205.
- 15↑
O'Conner MS, Steiner JM & Roussel AJ, et al. Evaluation of urine sucrose concentration for detection of gastric ulcers in horses. Am J Vet Res 2004;65: 31–39.
- 16↑
Pinotic L, Zecic-Fijacko M & Vcev A, et al. Diagnostic value of a peroral sucrose permeability test in children with recurrent upper abdominal pain. Coll Antropol 2004;28: 775–780.
- 17↑
Begg LM, O'Sullivan CB. The prevalence and distribution of gastric ulceration in 345 racehorses. Aust Vet J 2003;81: 199–201.
- 18↑
Murray MJ, Eichorn ES, Jeffrey SC. Histological characteristics of induced acute peptic injury in equine gastric squamous epithelium. Equine Vet J 2001;33: 554–560.
- 19↑
Buchanan BR, Andrews FM. Treatment and prevention of equine gastric ulcer syndrome. Vet Clin North Am Equine Pract 2003;19: 575–597.
