Intestinal epithelia separate luminal contents from the interstitium and are responsible for absorption of nutrients from the lumen and for separation of potentially toxic intestinal luminal contents from systemic circulation. Their function is primarily determined by the tight junctions in intestinal epithelial cells that seal the paracellular space. Intestinal tight junctions are selectively permeable, and their permeability can increase in response to luminal nutrients or pathologically by mucosal immune cells and cytokines, the enteric nervous system, and pathogens.1 Increased intestinal permeability has been found to be associated with a variety of human diseases, including those primarily affecting gut, such as inflammatory bowel disease, celiac disease, and irritable bowel syndrome, as well as systemic diseases or diseases involving other organ systems, such as type I diabetes, graft versus host disease, human immunodeficiency virus (HIV), multiple sclerosis, rheumatoid arthritis, and autism.2–5
Intestinal permeability is a useful marker for disease diagnosis and therapeutic effect assessment. There are multiple methods available to evaluate and measure intestinal permeability in human and animal studies.6,7 The lactulose and mannitol absorption test has been most prevalently applied clinically and in research. Lactulose and mannitol are sugars with molecular weights of 342 and 182 g/mol, respectively. The larger sugar lactulose can only cross the intestinal barrier via paracellular route, and the smaller-sized mannitol is thought to freely cross the intestinal barrier through transcellular transport, independent of intestinal barrier damage or dysfunction.7 Absorption of both sugars is affected by gastric dilution, motility, bacterial degradation, and renal function in the same manner. These 2 sugars are not metabolizable in humans or animals and are excreted in urine after absorption.8 This sugar test has demonstrated a high diagnostic value to assess intestinal permeability with specificity and sensitivity to the patients of coeliac disease, liver cirrhosis, and Crohn’s disease.9,10
The sugar test procedure begins with an oral administration of a mixture of lactulose and mannitol after overnight fasting, and then all urine is collected into a container for 5 or 6 hours postdose. The lactulose and mannitol concentrations in the urine sample are measured, and the ratio of lactulose to mannitol in the urine is used to assess intestinal permeability.10–12
This procedure is more complicated in animal studies compared to human testing. In animal studies, urine collection needs to be performed in metabolism cages13–15 or using urinary bladder urethral catheterization or cystocentesis16–18 or using attached bags.19,20 As a result, the operational execution of this procedure is much more challenging in animal studies.
The objective of this study was to evaluate whether intestinal permeability can be assessed by plasma sample instead of urine sample collection and analysis following an oral administration of a lactulose and mannitol mixture in pigs. The correlation of lactulose-to-mannitol ratios in plasma and urine was evaluated in the pigs.
Materials and Methods
Materials
Lactulose, d-mannitol, 13C-lactulose, and 13C-mannitol were purchased from Sigma-Aldrich. 13C-lactulose and 13C-mannitol were used as internal standards in the liquid chromatography–mass spectrometry (LC/MS) analysis. The mixture of lactulose (300 mg/mL) and mannitol (30 mg/mL) was dissolved in distilled water for animal dosing.
Animals
Ten weaned piglets (Large White/Landrace X Duroc) were obtained from Puregenic Pork Inc (Bunker Hill). The body weights and ages of pigs varied from 9.7 to 19 kg and 4 to 7 weeks at dosing. The pigs were group housed for about 1 week before the study initiation. One day before dosing, each pig was placed into an individual metabolism cage. The pigs were fasted overnight. Each pig was dosed with a single oral gavage of lactulose and mannitol mixture at 500 mg/kg body weight lactulose and 50 mg/kg body weight mannitol using a syringe and a short plastic tube with soft edges at the back of the throat.13 No feed was supplied until completion of the study. Water was supplied 30 minutes postsugar administration. Serial blood was collected via the jugular vein of pigs at predose, 10 and 30 minutes and 2, 4, and 6 hours postdosing. Plasma samples were obtained by centrifuging blood samples in K2 EDTA tubes. Cumulated urine was collected through grid mesh floor of the cages to ice-cold containers at 6 hours postdosing. Both plasma and urine samples were stored in −20 °C freezer before analysis. The operation was conducted under the oversight of the company’s Institutional Animal Care and Use Committee and according to local, state, and national regulations.
Sample preparation for analysis
Urine samples were centrifuged at 5,000 X g for 10 min. Supernatants of 3 µL were mixed with 350 µL of 80% acetonitrile and 20% 2 mM ammonium acetate containing 2 µg/mL of 13C-mannitol and 2 µg/mL of 13C-lactulose. The lactulose and mannitol concentrations in the mixtures were quantified using LC/MS. Plasma samples of 10 µL were mixed with 350 µL of 80% acetonitrile and 20% 2 mM ammonium acetate containing 2 µg/mL of 13C-mannitol and 2 µg/mL of 13C-lactulose. The mixtures were centrifuged at 2,000 X g for 10 min. The lactulose and mannitol concentrations in the supernatants were quantified using LC/MS.
LC/MS
Lactulose and mannitol in plasma and urine were quantified using a LC/MS system consisting of a Waters Acquity UPLC and an Applied Biosystem API4000 mass spectrometer. Separation was accomplished using a Waters Acquity BEH Amide column (1.7 μm, 2.1 X 100 mm) at 40 °C. The mobile phases were 90% acetonitrile and 10% 2 mM NaOH as mobile phase B and water containing 2 mM NaOH as mobile phase A. All analytes were eluted in 7 min by a gradient from 5 to 75 percent of mobile phase B at a flow rate of 0.3 mL/min. The multiple reaction monitoring (MRM) transitions were from m/z 341 to 161 for lactulose and m/z 181 to 101 for mannitol using positive electrospray ionization mode on the mass spectrometer. The MRM transitions were from m/z 353 to 167 for 13C-lactulose and m/z 187 to 92 for 13C-mannitol. Calibration standards of mannitol and lactulose were prepared from 0.006 to 2 μg/mL in both pig plasma and urine blank controls.
Data processing
The concentration ratios of lactulose to mannitol in urine were calculated directly for each pig. The pharmacokinetic parameters of lactulose and mannitol in the plasma of each pig were assessed using a noncompartmental model in Watson LIMS software (Thermo Fisher). The lactulose-to-mannitol ratios in the plasma of pigs were calculated using the sugar ratios of the area under curves from 0 to 6 hours (AUC0–6h); extrapolated area under curves (AUCextrap); maximum plasma concentrations (Cmax.); the sugar plasma concentrations at 2, 4, or 6 hours; and mean sugar plasma concentrations collected at time points of 2, 4, and 6 hours.
Results
The plasma profiles of lactulose and mannitol following a single oral gavage administration of lactulose and mannitol mixture at 500 mg/kg lactulose and 50 mg/kg mannitol in pigs are illustrated (Figure 1). Both sugars were absorbed rapidly following the oral administration and reached maximum plasma concentrations at around 2 to 4 hours in pigs. The pharmacokinetic parameters of both sugars are listed (Table 1). The sugar concentrations in the cumulated urine samples of 6 hours are listed (Table 2).
Plasma profiles of lactulose (A) and mannitol (B) in pigs following a single oral gavage administration of mixture of lactulose and mannitol.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.01.0002
Pharmacokinetic parameters of lactulose and mannitol in piglets following a single oral administration of lactulose and mannitol mixture.
| AUC0–6h (µg·h/mL) | AUCextrap (µg·h/mL) | Cmax (µg/mL) | tmax (h) | t1/2 (h) | ||||||
|---|---|---|---|---|---|---|---|---|---|---|
| Pig no. | Lactulose | Mannitol | Lactulose | Mannitol | Lactulose | Mannitol | Lactulose | Mannitol | Lactulose | Mannitol |
| 1 | 54.8 | 90.2 | 62.9 | 105 | 12.8 | 23.2 | 2.0 | 2.0 | 1.35 | 1.41 |
| 2 | 47.7 | 92.2 | 55.4 | 101 | 12.6 | 27.8 | 2.0 | 2.0 | 1.87 | 1.26 |
| 3 | 43.2 | 53.1 | 57.7 | 62.0 | 11.2 | 15.3 | 2.0 | 2.0 | 2.61 | 1.58 |
| 4 | 29.2 | 55.6 | 35.4 | 70.4 | 8.47 | 15.9 | 2.0 | 2.0 | 2.01 | 1.98 |
| 5 | 34.7 | 43.2 | 43.4 | 55.2 | 7.82 | 10.3 | 4.0 | 2.0 | 1.72 | 1.83 |
| 6 | 57.4 | 25.1 | 88.5 | 36.5 | 14.5 | 6.22 | 4.0 | 4.0 | 2.56 | 2.31 |
| 7 | 21.7 | 55.7 | 28.9 | 81.2 | 5.46 | 12.9 | 2.0 | 4.0 | 2.32 | 2.43 |
| 8 | 78.9 | 54.7 | 95.2 | 93.7 | 20.9 | 15.4 | 2.0 | 2.0 | 1.70 | 3.71 |
| 9 | 65.3 | 50.3 | 74.9 | 61.2 | 17.8 | 14.5 | 2.0 | 2.0 | 1.51 | 2.04 |
| 10 | 72.0 | 33.5 | 117 | 59.0 | 18.0 | 8.10 | 4.0 | 4.0 | 2.81 | 3.31 |
| Mean | 50.5 | 55.4 | 65.9 | 72.5 | 13.0 | 15.0 | 2.6 | 2.6 | 2.05 | 2.19 |
| SD | 18.7 | 21.5 | 28.0 | 22.1 | 4.95 | 6.53 | 0.97 | 0.97 | 0.50 | 0.79 |
| %CV | 37.0 | 38.8 | 42.5 | 30.5 | 38.2 | 43.6 | 37.2 | 37.2 | 24.6 | 36.3 |
Total urinary volumes collected in 6 hours, lactulose and mannitol concentrations in the urine, and urinary lactulose-to-mannitol ratios (L/M).
| Pig no. | Body weight at dosing (kg) | Urine volume (mL) | Lactulose (µg/mL) | Mannitol (µg/mL) | L/M |
|---|---|---|---|---|---|
| 1 | 16.1 | 260 | 502 | 847 | 0.59 |
| 2 | 15.6 | 1,000 | 112 | 260 | 0.43 |
| 3 | 13.4 | 560 | 94 | 118 | 0.80 |
| 4 | 10 | 200 | 252 | 540 | 0.47 |
| 5 | 14.2 | 260 | 354 | 511 | 0.69 |
| 6 | 15 | 300 | 507 | 243 | 2.09 |
| 7 | 14.8 | 200 | 310 | 883 | 0.35 |
| 8 | 17.5 | 100 | 2,230 | 1,380 | 1.62 |
| 9 | 19 | 480 | 415 | 324 | 1.28 |
| 10 | 9.7 | 90 | 758 | 359 | 2.11 |
The lactulose-to-mannitol ratios of AUC0–6h, AUCextrap, and Cmax of each pig were calculated in plasma. The ratios were correlated to their urinary sugar ratios with slope values of 1.03, 0.976, and 1.02 and R2 values of 0.995, 0.965, and 0.990, respectively (Figure 2). The plasma ratios of lactulose to mannitol were a function of time in pigs (Figure 3). The ratio became relatively steady after 2 hours postdosing for each pig. The sugar ratios in plasma at 2, 4, or 6 hours and the mean values of these 3 time points were correlated to their urinary sugar ratios with the slopes and R2 values close to 1 (Figure 4).
Correlation of lactulose-to-mannitol (L/M) ratios of AUC0–6h (A), AUCextrap (B), or Cmax (C) to the ratios in urine of pigs.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.01.0002
L/M ratios in plasma collected at different times in pigs.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.01.0002
Correlation of L/M ratios in plasma at 2 hours (A), 4 hours (B), or 6 hours (C) or mean values at 2, 4, and 6 hours (D) to the ratios in urine of pigs.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.01.0002
Discussion
The lactulose and mannitol test has been considered the gold standard for functional measurement of intestinal permeability in humans, which measures urinary excretion of these 2 nonmetabolized sugars over a 6-hour period after oral uptake.7,21 Absorption of the sugars occurs predominantly within 6 hours, and only a small percentage of both sugars is absorbed relative to the amounts dosed.19,22 The urinary sugars detected reflect the absorption in the small intestine since they are predominantly degraded by colonic bacteria.23,24 The lactulose-to-mannitol ratio is the most reliable indicator to monitor intestinal permeability changes clinically.11
Experimental animals, such as mice, rats, swine, chicken, and equine, have been used in intestinal permeability assessment in a variety of disease models in human drug discovery and development.5 Intestinal permeability has also been assessed as a tool in animals for veterinary medicine research.17,25–31 When the lactulose and mannitol test has been applied in animals, metabolism cages are usually needed to collect urine samples. Metabolism cages are designed to separate and collect feces and urine for numerous qualitative and quantitative determinations. Although metabolism cages have been widely used in drug mass balance or metabolism studies, the cages are not always available in every laboratory due to their cost and operation, especially the larger cages for dogs, pigs, and other larger animals. Sugar intestinal permeability assessment has also been conducted through urinary bladder urethral catheterization or cystocentesis in pigs, dogs, and cats.16–18 For larger animals, bags that attach to animals are also used to collect urine for the sugar test.19,20 These approaches are much more technically challenging.
Although sugar test is predominantly performed using urine collection, plasma or serum assay has been explored previously. Following oral administration of rhamnose, 3-O-methyl-d-glucose, d-xylose, and lactulose to dogs with or without inflammatory bowel disease, ratios of lactulose to rhamnose and d-xylose to 3-O-methyl-d-glucose collected at 2 hours are correlated to the ratios in dog urine collected for 5 hours postdose.32 Dogs that followed a single oral administration of lactulose and mannitol demonstrated that lactulose-to-mannitol ratios in plasma are linearly correlated with the ratios in urine at 1, 2, 3, 4, and 5 hours post-administration, although the plasma ratios are twice higher than the urinary ratios.33 A single-point serum measurement of lactulose and mannitol after 60 min of an oral administration of the sugar mixture has been tested in newborn healthy and diarrheic calves. The diarrheic calves show higher serum lactulose-to-mannitol ratios than the healthy calves on days 0, 7, 14, and 21 postbirth.34 Plasma assay oral administration of lactulose and mannitol has been also performed to assess isoflurane anesthesia effect on intestinal permeability in dogs. The transitory increase of plasma lactulose-to-mannitol ratios is observed due to anesthesia.31
Blood collection is more practical than urine collection in animals. Lactulose-to-mannitol ratios of AUC0–6h, AUCextrap, and Cmax in plasma samples were linearly correlated to their urinary ratios in pigs in this study; therefore, the urine assay can be replaced with a plasma assay for lactulose and mannitol absorption or intestinal permeability. It is apparent that plasma collection for 6 hours was not enough reaching the terminal phases of elimination of lactulose and mannitol in pigs (Figure 1). The correlation of sugar urine ratios of their AUCextrap ratios is more variable relative to AUC0–6h and Cmax ratios. Extension of blood collection after 6 hours may minimize the difference. The lactulose-to-mannitol ratios in plasma were approaching a steady status at and after 2 hours (Figure 3). The individual plasma ratios at 2, 4, or 6 hours and the mean ratios of these 3 time points were comparable to the ratios in urine (Figure 4).
The usage of metabolism cages needs more space and animal care in operation, particularly for large animals such as companion animals and livestock. Individual housing in metabolism cages can also result in stress in pigs.35–37 Such stress can negatively impact intestinal integrity and function.38,39 In this study, the lactulose-to-mannitol ratios varied from 0.3 to 2.1 among the 10 pigs, compared to the ratio at around 0.01 from the pigs not caged individually in metabolism cages in our other tests (data not reported). These high lactulose-to-mannitol ratios may be the result of the stress due to individual caging. The differences in the ratios among the pigs of this study may also indicate the tolerance of individual pigs to stress. Further investigation will be needed to interpret such a phenomenon.
In summary, lactulose-to-mannitol ratios of their pharmacokinetic parameters, such as AUC0–last, AUCextrap, and Cmax, are all comparable to the sugar ratios in the urine of pigs. The plasma concentration ratios of lactulose to mannitol at single time points or the mean values of several time points of their later exposure phases are acceptable to assess intestinal permeability in pigs or other animals following oral administration of a mixture of lactulose and mannitol solution. This provides a way to conduct the intestinal permeability assessment in animals without urine collection using metabolism cages or other more technically challenging approaches.
Acknowledgments
The authors declare that there were no conflicts of interest.
The authors thank Dr. Bernard Hummel and Dr. Kevin Esch for review and suggestions.
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