Veterinarians, independently of their area of specialty, are commonly confronted to clinical disease associated with gastrointestinal (GI) motility disorders, e.g., postoperative ileus in horses, delayed gastric emptying or constipation in small animals, and abomasal displacement or cecal dilatation in cattle. The pathomechanisms leading to these disorders are, in most cases, not well understood. Therefore, motility in health and disease is an area of active research in veterinary medicine, and various methods, including in-vivo, ex-vivo and in-vitro techniques, are available for assessment of GI motility in animals. The principles, advantages and disadvantages of the methods of investigation of GI motility used most widely in veterinary research will be shortly reviewed, using examples from the ruminant species for illustration.
In-Vivo Assessment of Motility
Assessment of Flow Rate of Ingesta
Flow rate of ingesta can be measured directly by diverting gastric or intestinal contents through a cannula implanted in the GI location of interest and measuring the volume accumulated over a given period of time. Because, however, removal of digestive juices can lead to severe electrolyte and/or acid-base disturbances, flow rates are mostly measured by use of re-entrant cannulas, which allow for flow rate assessment and immediate return of the GI content to the digestive tract.1,2 Whole-tract digesta kinetics, i.e., the transit time of markers from the rumen to the rectum have been described in cattle.3 Such measurements may be used for nutrition studies but do not allow for estimation of differentiated flow rates for the various segments of the digestive tract.4
The gastric emptying rate in sheep has been determined by use of electromagnetic (flowmeter) probes implanted in the proximal duodenum. In that study, a high degree of correlation was observed between flow rate and not only mechanical activity recorded with strain gauges but also with myoelectric activity recorded from electrodes implanted in the wall of the abomasum and duodenum.5 Cannulas have further been used for estimation of abomasal outflow in sheep based on the concentration vs. time curve of a marker, e.g., CrEDTA, or for measurement of small intestinal transit time between a cannula implanted in the duodenum and one in the distal ileum after injection of a marker, e.g., phenol red or polyethylene glycol.6,7
The implantation of cannulas in the GI tract, while allowing for direct assessment of flow rate of ingesta, is an invasive method. The GI segment of interest has to be apposed to the abdominal wall in a fixed position, which may impact on normal GI activity and cause interferences with physiologic motility patterns.8
In the past, Measurements of mechanical activity in the GI tract for direct assessment of muscle contraction have been achieved with strain-gauge force transducers sutured to the gastric or intestinal wall, or with pressure probes, e.g., air- or water-filled balloons connected to a manometer.9 More recently, miniature pressure transducers mounted on catheters have been used to measure intraluminal pressures, however they have mostly have been used to monitor intrauterine pressure in cattle to date.10
Assessment of Myoelectric Activity
Myoelectric activity of the GI tract consists of slow waves (oscillations of the smooth muscle cell membrane potential which remain below the depolarization threshold and are not associated with muscular contraction), and superimposed spike potentials (depolarization of the membrane beyond the excitation threshold for contractions) which can be recorded from electrodes implanted in the muscle layer of the gastric or intestinal wall.11 Correlation of spiking activity with muscle contraction and propulsion of gut contents has been demonstrated, which allows for use of myoelectric activity in the digestive tract as an indicator of mechanical movement.9,12,13 Reliable myoelectric recordings can be obtained over extended periods of time from permanent electrodes implanted in the wall of GI organs, but this invasive technique is reserved for experimental settings.14 In contrast, thin retrievable electrodes can be implanted in patient animals during surgery for correction of disease, e.g., in cattle with cecal dilatation, for recording of myoelectric activity during the postoperative period. At the end of the measurements, the electrodes are retrieved and the animals can be returned to their owners.15 While electrode implantation remains an invasive technique, it allows to gather reliable and detailed information on GI motility in-vivo, and especially on organized patterns such as the migrating myoelectric complex (MMC). The effects of diet,16 disease,17 or drugs18 on motility patterns can be investigated by use of this technique.
The most widely used absorption test in veterinary research is the acetaminophen absorption test. Acetaminophen, best known as an analgesic and antipyretic drug in humans, is absorbed in the proximal small intestine, thus the time to appearance of acetaminophen in the blood after oral administration correlates with gastric emptying time. It has been used in preruminant calves to study the effects of oral fluid composition on abomasal emptying rate or to compare the time necessary for milk to reach the duodenum in ruminal drinkers vs. healthy calves.19,20,21,22 In adult cattle, abomasal emptying has been assessed by use of the D-xylose absorption test, whereby, in contrast to monogastric animals, D-xylose could not be administered orally but was injected directly into the abomasum in order to by-pass the forestomachs. A prolonged time to maximal D-xylose blood concentration, thus delayed abomasal emptying in cows after surgical correction of abomasal displacement vs. healthy controls was demonstrated.23
Furthermore, the breath hydrogen test can be used to assess intestinal malabsorption of carbohydrates. If increased amounts of carbohydrates pass non-absorbed through the small intestine, they are fermented by colonic bacteria to short-chain fatty acids and gases, including hydrogen which is transferred in the blood to the lungs to be excreted in the breath. Increased breath hydrogen concentrations have been measured, e.g., in preruminant calves with decreased intestinal absorptive capacity.24
While contrast radiography has been used in the past for transit studies, it has been largely replaced, at least in large animals, by ultrasonography and/or scintigraphy.
Abomasal volume and emptying rate have been assessed by ultrasonography, by measuring the maximal visible length, width and height of the stomach from the ventral right flank of calves.21,25 While ultrasonography of the intestine is widely used for diagnostic purposes in ruminant GI disease,26 use of this technique in research has been limited because it is difficult to establish protocols that ensure reproducible and thus comparable measurements.
Nuclear scintigraphy has been used with 99mTc as a marker to determine gastric emptying in calves.20,27 Scintigraphy allows for quantification of gastric emptying with indices such as t½, the time required for 50% of the initial volume to be emptied from the stomach. In adult sheep, assessment of abomasal emptying after oral administration of radiolabelled meals was not possible due to superposition of the stomach compartments, but could be achieved after administration of the marker directly into the abomasum.28
Ex-Vivo Assessment of Motility
Organ Bath Studies
In ex-vivo studies, specimens of the organ of interest are harvested from animals immediately after death and maintained viable for several hours in an oxygenated polyionic solution. Smooth muscle samples are suspended in individual organ-bath chambers and connected to a force transducer for registration of contraction and/or relaxation of the muscle specimens. The effects of various conditions, such as the effects of increasing concentrations of volatile fatty acids,29 or those of agonists and/or antagonists of specific receptors in the gut wall30 can be investigated. This relatively easy technique can be used e.g., for preliminary evaluation of potential therapeutic agents, in order to confirm their effects on the tissues of interest prior to clinical trials. However, results obtained with isolated tissue samples in absence of the complex in-vivo mechanisms regulating GI motility must be interpreted with caution because the effects of a given substance may be different in organ-bath settings or in-vivo.
In-Vitro Studies of Motility
Quantitative PCR for Measurement of mRNA Expression
Real-time RT-PCR techniques have been used to quantify the expression of mRNA coding e.g., for receptors involved in the regulation of motility.31,32 With these methods, mRNA expression can be quantified out of tiny tissue specimens, e.g., biopsy samples harvested from clinical patients at surgery for comparison with mRNA levels in healthy animals.33,34 However, interpretation of such results is difficult because mRNA expression is not necessarily correlated to receptor protein expression and receptor function.
Measurement of Receptor Protein Expression
Binding studies allow to quantify receptor protein expressed in tissues in very low levels.35,36 These methods imply to work with radioactive compounds and they do not provide information on the localization of receptor proteins in the tissue under study. Based on simultaneous quantification of expression of mRNA and receptor protein, potential correlation of these two levels of receptor expression can be investigated. In case of a high degree of correlation inferences might be made regarding protein expression based on results of measurements of mRNA expression, which are easier to perform and feasible with a few grams of tissue only.
Receptor protein can also be stained by immunohistochemistry methods, provided that specific antibodies are available.37 With this technique, receptor protein can be localized to specific layers or cell types of the stomach or intestine wall, but the results are at best semi-quantitative.
1. Horney FD, et al. Am J Vet Res 1972;33:1385.
2. Sissons JW, Smith RH. J Physiol 1978;283: 307.
3. Luginbuhl JM, et al. J Anim Sci 1994;72:201.
4. de Vega, et al. Br J Nutr 1998;80:381.
5. Malbert CH, Ruckebusch Y. J Physiol 1988;401:227.
6. Smith RH. J Physiol 1964;172:305.
7. Gregory PC, Miller SJ. J Physiol 1989;413:415.
8. MacRae JC, et al. Res Vet Sci 1973;14:78.
9. Ruckebusch Y. J Physiol 1970;210:857.
10. Hirsbrunner G, et al. Theriogenology 2000;54:291.
11. Sarna SK. Gastroenterology 1985;89:894.
12. Bueno, et al. J Physiol 1975;249:69.
13. Bolton JR, et al. Am J Vet Res 1976;37:1387.
14. Meylan M, et al. Am J Vet Res 2002;63:78.
15. Stocker S, et al. Am J Vet Res 1997;58:961.
16. Meylan M, et al. Am J Vet Res 2002;63:857.
17. Meylan M, et al. J Vet Intern Med 2003;17:571.
18. Zanolari P, et al. J Vet Med A 2004;51:456.
19. Schaer S, et al. J Vet Med A 2005;52:325.
20. Marshall TS, et al. Am J Vet Res 2005;66:364.
21. Sen I, et al. Am J Vet Res 2006;67:1377.
22. Herrli-Gygi M, et al. Vet J 2007;Epub ahead of print.
23. Wittek T, et al. J Vet Intern Med 2005;19:905.
24. Holland, et al. Am J Vet Res 1986;47:2020.
25. Wittek T, et al. Am J Vet Res 2005;66:537.
26. Braun U. Vet J 2003;166:112.
27. Nappert G, Lattimer J.C. Can J Vet Res 2001;65:54.
28. Nicholson T, et al. Res Vet Sci 1997;62:26.
29. Allemann M, et al. Am J Vet Res 2000;61:678.
30. Pfeiffer JB, et al. Am J Vet Res 2007;68:313.
31. Meylan M, et al. Am J Vet Res 2004;65:1142.
32. Meylan M, et al. Am J Vet Res 2004;65:1151.
33. Engel L, et al. Am J Vet Res 2006;67:95.
34. Kobel B, et al. Am J Vet Res 2006;67:1367.
35. Ontsouka EC, et al. J Anim Sci 2006;84:3277.
36. Ontsouka EC, et al. J Recept Signal Transduct Res. 2007;27:147.
37. Stoffel MH. Am J Vet Res 2006;67:1992.