A. Minuti, S. Ahmed, E. Trevisi, F. Piccioli-Cappelli, G ...

Experimental acute rumen acidosis in sheep: consequences on clinical, rumen, gastrointestinal permeability conditions, and blood chemistry A. Minuti, S. Ahmed, E. Trevisi, F. Piccioli-Cappelli, G. Bertoni, N. Jahan and P. Bani J ANIM SCI published online July 1, 2014

The online version of this article, along with updated information and services, is located on the World Wide Web at: http://www.journalofanimalscience.org/content/early/2014/07/01/jas.2014-7594

www.asas.org

Downloaded from www.journalofanimalscience.org at Cattolica Del Sacro on July 11, 2014

Left Running Head: Minuti et al. Right Running Head: Acidosis and gastrointestinal permeability

Experimental acute rumen acidosis in sheep: consequences on clinical, rumen, gastrointestinal permeability conditions, and blood chemistry 1

A. Minuti,*† S. Ahmed,*‡ E. Trevisi,*† F. Piccioli-Cappelli,* G. Bertoni,* N. Jahan,* and P. Bani*2

*Istituto di Zootecnica, Facoltà di Agraria, Università Cattolica del Sacro Cuore, via Emilia Parmense 84, 29122 Piacenza, Italy; †PRONUTRIGEN - Centro di Ricerca sulla Proteomica e Nutrigenomica, Università Cattolica del Sacro Cuore, Piacenza, Italy; and ‡Bangladesh Livestock Research Institute, Savar, Dhaka 1341, Bangladesh.

1

This manuscript is based on research supported by the “Fondazione Romeo ed Enrica

Invernizzi” Milan, Italy. 2

Corresponding author: [email protected]

Received January 14, 2014. Accepted June 14, 2014.

DownloadedOnline from www.journalofanimalscience.org Cattolica Del Sacro on July 11, 2014 Published First on July 1, 2014 atas doi:10.2527/jas.2014-7594

ABSTRACT Acute acidosis was induced in sheep, and gastrointestinal permeability was assessed by using lactulose as a permeability marker. Metabolism was evaluated by monitoring blood metabolites. Four rams (72.5 ± 4.6 kg BW) were used in a 2×2 change over design experiment. The experimental period lasted 96 hours from -24 h to 72 h. After 24 h of fasting (from -24 h to 0 h) for both controls and acidosis-induced rams (ACID), 0.5 kg of wheat flour was orally dosed at 0 h and 12 h of the experimental period to ACID, while the basal diet (grass hay, ad libitum) was restored to control. At 24 h, a lactulose solution (30 g of lactulose in 200 mL of water) was orally administered. Blood samples were collected at -24, 0, 24, 48, and 72 h of the experimental periods for the analysis of metabolic profiles and during the 10 h after lactulose dosage to monitor lactulose changes in blood. In addition, rumen and fecal samples were collected at 24 h of the experimental period. The acidotic challenge markedly reduced (P < 0.01) rumen pH and VFA but increased rumen D- and L-lactic acid (P < 0.01). Concurrently, a decrease of fecal pH and VFA occurred in ACID (P < 0.01), together with an abrupt increase (P < 0.01) of lactate and fecal alkaline phosphatase. Blood lactulose was significantly increased in ACID rams peaking 2 h after lactulose dosage. Blood glucose, β-hydroxybutyrate, Ca, K, Mg, and alkaline phosphatase showed a significant reduction (P < 0.05) at 24 h, whereas urea and NEFA declined (P < 0.05) from 48 h to 72 h. A strong inflammatory acute phase response with oxidative stress in ACID group was observed from 24 h to 72 h; higher values of haptoglobin (P < 0.01) were measured from 24 h to 72 h and of ceruloplasmin from 48 h (P < 0.05) to 72 h (P < 0.01). Among the negative acute phase reactants, plasma albumin, cholesterol, paraoxonase, and Zn concentration also decreased (P < 0.05) in ACID at different time points between 24 and 72 h after acidotic challenge start. A rise (P < 0.05) of reactive oxygen metabolites and a drop of vitamin E (P < 0.01) between 24 and 72 h were indicative of oxidative stress in ACID. The perturbation of these blood metabolites suggests that acute acidosis was effectively induced by our model. The increase of lactulose in blood in ACID rams indicates that gastrointestinal permeability for the marker increased and the large increment after 2 h from dosage suggests that most of the passage occurred through the rumen or abomasal walls.

Key words: acute ruminal acidosis, gastrointestinal permeability, lactulose test, sheep


INTRODUCTION During acute ruminal acidosis, the increase of proton concentration in the rumen challenges the integrity of the ruminal epithelial cells with the appearance of microlesions and parakeratosis (Mohamed Nour et al. 1998; Kleen et al., 2003; Ismail et al. 2010) that can affect animal metabolism. Furthermore, the acidic environment, the high osmotic pressure and the high concentration of lactate in the rumen can cause death and lysis of the gram-negative bacteria leading to an increase in cell-free lipopolysaccharide (LPS) in the forestomachs (Gozho et al., 2007; Zebeli and Ametaj, 2009). An impairment of the barrier function of the rumen epithelium resulting from feeding high grain-based diets is a possible mechanism of LPS translocation (Kleen et al., 2003). Translocation of LPS from the gastrointestinal tract (GI) to the bloodstream causes an inflammatory response with possible relevant metabolic disorders such as laminitis (Bertoni et al., 1989; Nocek, 1997), displacement of the abomasum (Zebeli et al., 2011), fatty liver, and sudden death syndrome (Ametaj et al., 2005). Liver rapidly detoxifies the circulating LPS, and attempts to quantify LPS in peripheral blood circulation have faced many difficulties (Andersen et al., 1994; Gozho et al., 2007). The LPS has only occasionally been detected in ruminants peripheral circulation after an experimental induction of subacute ruminal acidosis by high grain diets (Khafipour et al., 2009a). In our previous studies (Ahmed et al., 2013; Minuti et al., 2013), we proved that lactulose can be used as a marker to assess GI permeability in adult ruminants. We hypothesized that the changes of GI epithelium integrity during rumen acidosis condition can be evaluated by the lactulose test. The aim of this study was to evaluate, during an experimentally induced acute ruminal acidosis in rams, the GI epithelium integrity through lactulose permeability test and the related consequences on animal metabolism and inflammatory status.


MATERIALS AND METHODS All of the procedures involving animals were conducted in accordance with Italian laws on animal experimentation and ethics (DL n.116, 27/01/1992), protocol number ZN2013002.

Animals and Feeding System Four rams of the local population “Bardigiana” (72.5 ± 4.6 kg of initial BW) were used in this experiment. Animals were housed in separate cages with wooden floors and free access to water, in an environmentally controlled room (temperature 22°C, relative humidity 70%). Before and between the experimental periods the animals were fed twice daily (0700 and 1900 h) grass hay (11.4% CP and 8.11 MJ/kg ME, on a DM basis) for ad libitum intake, to ensure between 3 and 6% orts, and a vitamin-mineral supplement. During the experimental periods animals were fed according to the experimental design.

Experimental Design The experiment was carried out according to a change-over design (2x2), with two 4-d experimental periods (1 and 2) and a 28-d interval between periods. Figure 1 shows the time schedule of each experimental period. All the animals were fasted for 24 h (from -24 h to 0 h of the experimental period). In the control rams (CTR), the previous grass hay diet was then restored. In the treated rams (ACID), animals were then orally dosed with wheat flour to induce acute acidosis. Total amount of wheat flour was 1.0 kg/ram (corresponding to 12.8 ± 0.9 SD g/kg BW), administered in 2 equivalent portions of 0.5 kg at 0 h and 12 h of the experimental period. In the ACID group the grass hay diet availability was restored after 24 h. Blood samples were collected (before morning feed delivery) at -24, 0, 24, 48, and 72 h during


experimental periods. Rumen fluid through stomach tube and grab fecal samples were collected at 24 h. To assess gastrointestinal permeability, at 24 h (Lactulose Dosage Time= LDT), a solution containing 30 g of lactulose (150 g/L) was orally administered to every ram, and blood samples were collected at 0, 2, 4, 6, 8 and 10 h (0LDT, 2LDT, 4LDT, 6LDT, 8LDT and 10LDT) after LDT.

Feed Intake and Health Problems During the trial rectal temperature was measured at -24, 24, 36, 48, 60 and 72 h of the experimental period (Figure 1). At the same times animals were observed by general inspection to record development of clinical symptoms. In addition, feed intake and fecal features, were monitored.

Blood Sampling and Analysis Blood was collected from the jugular vein into evacuated tubes containing lithium-heparin (Vacutainer, Becton Dickinson, Plymouth, UK) and immediately cooled in ice water. A small amount of blood was used for packed cell volume determination (Centrifugette 4203, ALC International Srl, Cologno Monzese, Italy); the remainder was centrifuged at 3,500 × g for 15 min at 6°C, and the plasma was frozen (−20°C) until analyses. Blood samples were analyzed to assess the following profiles: metabolic (glucose, total cholesterol, urea, aspartate aminotransferase, γ-glutamyl transpeptidase, alkaline phosphatase [ALP], NEFA, β-hydroxybutyrate [BHBA], D- and L-lactate), mineral (Ca, P, Mg, Na, K, Cl, and Zn) and oxidative-inflammatory (total protein, albumin, globulin, total bilirubin, haptoglobin, ceruloplasmin, vitamins A, vitamin E, reactive oxygen metabolites [ROMs], thiol


groups, total nitric oxide metabolites , and paraoxonase. Blood metabolites were analyzed by an automated biochemistry analyzer (ILAB 600; Instrumentation Laboratory, Lexington, MA). Total protein, albumin, total cholesterol, total bilirubin, triglycerides, urea, Ca, P, Mg, aspartate aminotransferase, γ-glutamyl transpeptidase, and ALP were determined using kits purchased from

Instrumentation Laboratory

(Instrumentation Laboratory SpA-Milano, Italy). Globulin was calculated as difference between total protein and albumin. Ions (Na, K, and Cl) were measured by the potentiometric method (ion-selective electrode connected to ILAB 600). Zinc and NEFA were measured by commercial kits (Wako Chemicals GmbH, Neuss, Germany). The BHBA was determined by commercial kits Ranbut (Randox Laboratories Ltd., Crumlin, Co. Antrium, UK). Haptoglobin and ceruloplasmin were analyzed using the methods described by Skinner et al. (1991) and Sunderman and Nomoto (1970) respectively, adapted to the ILAB 600 conditions. The ROMs and SHp were measured by commercial kits (Diacron International s.r.l., Grosseto, Italy). The NOX were measured using the Griess test according to Gilliam et al. (1993) and Bouchard et al. (1999). Paraoxonasewas measured by the method of Ferré et al. (2002) adapted to the ILAB 600 as previously described by Bionaz et al. (2007). Plasma vitamins A and E were extracted with hexane and analyzed by reverse-phase HPLC using an HPLC (Varian, ProStar, Walnut Creek, CA), equipped with an Allsphere ODS-2 3µm, 150 × 4.6 mm column (Alltech, Deerfield, IL) and an UV detector set at 325 nm for vitamin A, 290 nm for vitamin E; a solution of 80:20 methanol:tetrahydrofurane was used as mobile phase. D- and L-lactate were analyzed using Megazyme L and D Lactate assay Kit (Megazyme International Ltd., Wicklow, Ireland). Lactulose Analysis The lactulose concentration in plasma samples collected at 0LDT, 2LDT, 4LDT, 6LDT, 8LDT and 10LDT


was analyzed through GLC adapting the methodology used for urinary sugar detection by Farhadi et al., (2003). Plasma samples were prepared for the analysis as follows: 250 µL of sample was added with 250 µL of solution containing internal standard (0.2 g/L of myo-inositol in 7% wt/vol trichloroacetic acid). Samples were mixed by vortexing and centrifuged (15,000 x g for 10 min at 20ºC). After that, 400 µL of the supernatant were transferred to a new tube and evaporated to dryness at 70°C under a stream of nitrogen. The dried residues were then taken up in 200 µL of anhydrous pyridine containing 25 mg/mL of hydroxylamine chlorhydrate, mixed by vortexing, heated at 70°C for 1 h, and centrifuged (15,000 x g for 10 min at 20ºC). An aliquot (100 µL) of the supernatant was transferred to a new tube, and sugar oximes were silylated with 100 µL of N-trimethylsilylimidazole for 30 min at 70°C. An aliquot of the silylated derivatives was sealed in an vial for analysis. The lactulose was quantified using a gas chromatograph (model 7820A GC, Agilent Technologies, Santa Clara, CA) equipped with a DB-1 capillary column (15 m × 530 μm × 1.5 μm; Agilent J&W GC column), and flame-ionization detection. The oven temperature was 220°C held for 5 min and then increased by 10°C/min for 2 min, 5°C/min for 4 min and 3.5°C/min for 4 min to a final temperature of 274°C, held for 9 min. The analysis time was approximately 24 min. The injector temperature was 250°C, the detector temperature was 280°C. Glass wool packed liners were used to prevent contamination of GC with dirty particles from the sample. The samples were injected by auto sampler, and 1 μL at 25:1 split ratio was injected. Hydrogen and air were used for flame ionization detection. Helium at a flow rate of 10 mL/min was used as carrier gas.

Rumen and Feces Sampling and Analysis


Rumen fluid was collected through a stomach tube (rubber tube of ~ 150 cm of length for 1.5 cm of diameter) at 24 h, before morning feeding. The pH of the fluid was measured immediately after it was withdrawn from the rumen by a pH meter (GLP 21, Crison Instruments SA, Alella, Barcelona, Spain). An aliquot of the remaining fluid was immediately cooled in ice water and centrifuged at 3,000 × g for 10 min at 10°C; 2 mL aliquots of the supernatant were transferred into tubes with 1 mL of 0.12 M oxalic acid and frozen at –20°C for later analysis of VFA and lactate. Fecal samples were collected manually at 24 h directly from the rectum immediately after rumen sampling, diluted with distilled water at 1:3 ratio, homogenized in a Stomacher 400 (Tekmar, Cincinnati, OH) for 3 min and centrifuged as for the rumen fluid. The pH measure and storing of the extract were performed as for the rumen samples. The L- and D-lactate were analyzed in rumen liquor and fecal extract by the same method used for lactate in blood. Alkaline phosphatase was analyzed in the fecal extract using a kit from Instrumentation Laboratory (Instrumentation Laboratory, Lexington, MA). The VFA concentration in rumen fluid and fecal extract was analyzed by a gas chromatograph (model 7820A, Agilent Technologies, Santa Clara, CA) equipped with a DB-FFAP capillary column (30 m × 250 μm × 0.25 μm; Agilent J&W GC column), and a flame-ionization detector. The oven temperature was 60°C held for 5 min, then increased by 5°C/min to 140°C. The injector temperature was 250°C, the detector temperature was 300°C. The injector was equipped with a glass liner of glass wool to separate particles of dirt from the sample. The samples were dosed by auto sampler at an injection size of 1 μL using the split method and a 25:1 splitting ratio. Hydrogen and air were used for flame ionization detection. The carrier gas was nitrogen, with constant flow of 1.78 mL/min. Pivalic acid was used as an internal standard.


Statistical Analysis Data of each parameter were previously tested for normal distribution using the Shapiro test (SAS Inst. Inc.) and normalized by natural log transformation, when necessary. Data were analyzed using mixed model procedures of SAS (SAS Inst. Inc., Cary, NC; release 8.0) for repeated measures. The statistical model used treatment time (from trial start or from LDT), interaction of treatment x time as fixed factors; and the individual ram were random effects. Each parameter was subjected to 4 covariance structures - autoregressive order, compound symmetry, spatial power, toeplitz - and the best covariance structure was retained (Littell et al., 2006). The effect of the period (1 and 2) was initially included in the model and then removed being not significant for any parameter. All values are presented as the means and SEM. The effect of the period (1 and 2) was initially included in the model and then removed being not significant for any parameter. Statistical significance was established by using a conventional P-value of 0.05 or 0.01.

RESULTS

Health Status of the Animals At 24 h after induction of acidosis, all ACID rams showed nervous depression and exhibited periods of trembling. All could stand up; however, mostly with head hung low and dull eyes. Parallel to this, ACID rams showed acute symptoms including watery, yellowish and acidic smelling diarrhea. Cessation of feed intake occurred in all ACID rams, and no appetite was evident until 72 h when a little interest in food and water appeared. All ACID rams showed more or less severe symptoms of foot pain and difficulty walking. Meanwhile, no clinical signs could


be observed in CTR rams. At the start of the experimental period, rectal temperature was 38.9 ± 0.23 °C in both groups, but 24 h after the induction of acidosis it increased (P < 0.01) in ACID, peaked (40.7°C) at 36 h and then declined, but remained higher than CTR rams until 72 h after the acidotic challenge (39.0 in CTR vs. 39.7°C in ACID; P < 0.01).

Ruminal and Fecal Parameters The mean pH and VFA concentrations for rumen samples collected at 24 h are presented in Table 1. Rumen pH was lower (P < 0.01) in ACID compared to CTR rams, and the pH for ACID indicated an acute acidosis condition (threshold value ≤ 5.0; Nagaraja and Town, 1990). Total VFA concentration and acid molar proportions differed between groups. In ACID compared with CTR rams, a marked decrease of total VFA concentration (P < 0.01) and a higher concentrations of L- and D-lactate (P < 0.01) were observed. Fatty acid molar proportions also differed between groups. Compared to CTR, ACID rams showed higher acetate (P < 0.05) and lower propionate (P < 0.05) proportions. Characteristics of fecal samples are shown in Table 2. Similar to the rumen, the fecal pH was lower in ACID compared to CTR rams (P < 0.01). In ACID a lower total fecal VFA concentration (P < 0.01) was also found, whereas L- and D-lactate concentrations were higher in ACID than in CTR (P < 0.01). The feces of ACID rams had a higher molar proportion of acetate (P < 0.01) and a lower proportion of propionate (P < 0.01) than those of CTR rams. In feces from ACID rams, isobutyrate, valerate, and isovalerate could not be detected, and alkaline phosphatase was higher compared to CTR rams (1,543 vs. 30 U/100 g fresh matter; P < 0.01).

Blood Parameters


At -24 h and 0 h (pre-challenge period), no differences for any blood parameters between ACID and CTR rams were detected.

Metabolic Blood Parameters The results of metabolic blood parameters are presented in Figure 2 and Table 3. Before the acidotic challenge the concentrations of D- and L-lactate in blood of ACID rams were quite similar, but their behavior after acidotic challenge was different. In ACID there was a rapid rise of D-lactate (Fig. 2A), with a maximum reached at 24 h (2.68 vs. 0.07 mmol/L in ACID and CTR; P < 0.01) and by 72 h D-lactate was restored to pre-challenge values. The rise of L-lactate was more gradual (Fig. 2B); it was higher in ACID rams from 24 h to 72 h (P < 0.01), but the peak was reached at 48 h (2.96 vs. 0.72 mmol/L in ACID vs. CTR). Glucose (Fig. 2C) and BHBA (Fig. 2D) showed a decrease, starting from 24 h, as a consequence of induction of acidosis. Plasma concentrations of urea, NEFA and bilirubin all increased in ACID rams (Table 3) and differed from CTR starting from 24 h (P < 0.05) to 48 h (bilirubin) or 72 h (urea and NEFA). On the contrary, the concentrations of Ca, K, Mg, and alkaline phosphatase (Table 3) were lower in the ACID compared to CTR rams at the different time points (P < 0.05). No changes were observed for aspartate aminotransferase and γ-glutamyl transpeptidase.

Positive and Negative Acute Phase Proteins and Oxidative Stress Reactants Haptoglobin (Fig. 3A) in the ACID rams started to increase after 24 h from the induction of acidosis (P < 0.01) and continued until 72 h (P < 0.01). Similarly, ceruloplasmin (Fig. 3B) started to rise in ACID from 24 h but differences were found at 48 h (P < 0.05) and 72 h (P < 0.01). Among negative acute phase proteins, plasma albumin (Table 3) declined in ACID


compared to CTR rams from 48 h with significant differences at 72 h (P < 0.01). Major changes were found in the plasma concentration of Zn and paraoxonase (Fig. 3C and 3D), with values significantly lower in the ACID compared to the CTR rams from 24 h to 72 h for Zn (P < 0.01) and from 48 h (P < 0.05) to 72 h (P < 0.01) for paraoxonase. Total cholesterol and vitamin A had a common trend (Table 3), decreasing in ACID after acidotic challenge, but differences with respect to CTR rams became significant from 24 h (P < 0.05) for cholesterol and from 48 h (P < 0.05) for vitamin A. Both parameters continued to decline in ACID rams until 72 h (P < 0.01). Among the parameters related to oxidative stress, ROMs started to rise in ACID rams after 24 h with differences from 48 h (P < 0.05) to 72 h (P < 0.01) (Fig. 3E). Vitamin E showed lower concentration in ACID rams (Fig. 3F) from 24 h to 72 h (P < 0.01). No changes were observed for thiol groups and total nitric oxide metabolites.

Lactulose Concentration in Blood The lactulose concentrations in blood plasma at different time points after oral dosing are presented in Figure 4. No detectable concentration of lactulose was found at 0LDT for either CTR or ACID rams. After the oral dosage, the plasma concentration of lactulose in CTR slightly increased at 2LDT and remained steady until 10LDT, with an average concentration of 3 µg/mL. In contrast, plasma concentration of lactulose rose rapidly in the ACID group after 2 h from lactulose dosage (P < 0.01) and remained higher compared to the CTR until 6LDT (P < 0.05). The highest value (39 µg/mL) was recorded 2 h after LDT whereas pre-dosage values were restored at about 8LDT.

DISCUSSION


The high energy demand for increasing productive performance often leads to feeding ruminants with highly-fermentable diets, largely based on concentrates and in particular on cereal grains. This can lead to the onset of ruminal acidosis situations with varying degrees of intensity. As a consequence of the acute rumen acidosis, an impairment of the gastrointestinal epithelium can occur, and several authors have hypothesized the translocation into the bloodstream of bacterial LPS and biogenic amines, which are more abundant in ruminal digesta during ruminal acidosis (Plaizier et al., 2008; Khafipour et al., 2009a; b; Saleem et al., 2012). When LPS translocation happens, the consequence on the animal can be an overt systemic inflammatory response with associated metabolic disorders like compromised insulin sensitivity and stimulation of lipolysis. In the present experiment the digestive epithelium integrity of the rams was challenged by inducing an acute experimental acidosis. After the induction of acidosis rams developed some typical clinical signs like anorexia, abdominal pain, periods of trembling with problems in movement, and watery diarrhea with yellowish color and acidic odor. Similar clinical symptoms were also reported by Patra et al. (1996), Mohamed Nour et al. (1998) and Ismail et al. (2010) after acute acidosis induction in sheep and goats. Our model for the induction of acidosis was based on 24-h fasting followed by administration of 1 kg of wheat flour in 2 equal doses spaced by 12 h. In such conditions the mean value of rumen pH measured 24 h after the challenge in treated animals was 4.94. In some other comparable experiments, higher doses of flour or starchy concentrates were used. Patra et al. (1996) induced acute acidosis in adult sheep after 24 h fasting by feeding soaked wheat at 90 g per kg BW and after 24 h rumen pH was reduced to 4.70. Lettat et al. (2010) and Ismail et al. (2010) induced rumen acute acidosis in rumen-cannulated sheep and goats by intraruminal supply of wheat at 12 g/kg BW in sheep and wheat flour at 50 g/kg BW in goats, and in these conditions ruminal pH


dropped to 4.85 and 4.23 after 24 h, respectively. The marked reduction in ruminal pH that occurred in the present study indicates that acute acidosis was effectively induced. This strong acidification was associated with the reduction of total VFA and the marked increase of lactic acid in the rumen (50-fold increase in ACID compared with CTR). Similar results were observed in previous studies of induction of acute acidosis (Muir et al., 1980; Ismail et al., 2010; Lettat et al., 2010). The decline of VFA seems caused by a marked impairment of the bacterial flora, whereas the marked accumulation of the lactic acid can be a consequence of its increased production but reduced utilization within the rumen (Slyter, 1976; Nagaraja and Titgemeyer, 2007). Rumen concentration of both isomers of lactate in ACID was very similar, but in feces a 3:1 ratio was found between L- and D-lactic acid, respectively. Acid-tolerant lactobacilli, mainly producing D-lactate isomer (Loew and Chaplin, 1976) may proliferate in both acute and subacute acidosis (Slyter, 1976; Nagaraja and Miller, 1989; Goad et al., 1998) leading to an increase in production of total and in particular D-lactate. Microbial changes in the large intestine associated with experimentally induced acidosis have also been reported but they didn’t parallel those taking place in the forestomachs as in that site D-lactate potentially yielding lactobacilli did not proliferate (Metzler-Zebeli at al., 2013). Zust et al. (2000) measured a cecal concentration of Llactate 8 times higher than D-lactate after having induced a mild intestinal acidosis, but a 3-fold higher L-lactate when an acute acidosis was induced. Ewaschuk et al. (2004) measured a 3-fold higher level of fecal D-lactate compared with L-lactate on milk-fed diarrheic calves but lower and much similar concentrations of the two isoforms in healthy calves. The concentrations of D- and L-lactic acid in blood (Figure 2) were markedly increased during the experiment, suggesting that our protocol likely led the animals to a condition of metabolic


acidosis. This circumstance is typically observed during acute acidosis experiments in sheep (Muir et al., 1980; Patra et al., 1996; Ismail et al., 2010). Lactic acid isomers reached similar maximum concentrations in blood but had different patterns: D-lactate peaked at 24 h after the start of the induction of the acidosis and then declined rapidly, whereas L-lactate increased more slowly reaching the peak at 48 h. Although D-lactic acid can be metabolized in humans (Connor et al., 1983) and other mammals (Giesecke and von Wallenberg, 1985), L-lactic acid is reported to be more rapidly metabolized than the D-isomer (Prior, 1983; Harmon et al., 1984). Lactic acid concentration in the rumen and feces was measured after 24 h from the onset of the acidosis induction, whereby it is not possible to assess the relative contribution to blood pool of lactic acid from the different sites of absorption, i.e. rumen or lower gastrointestinal tract. Slyter (1976) and Patra et al. (1996) observed that diarrhea is often associated with acute and with subacute acidosis. Diarrhea is mainly due to an increased osmolality in the intestine consequent to high lactic acid concentration in the digesta (Huber, 1976), which in turn drives the pH decrease in feces. High concentrations of lactic acid and diarrhea occurred in the present experiment. Similar to our results, a significant reduction of fecal pH was also reported in sheep by Al Jassim et al. (2003) at 24 h after acute acidosis induction by barley engorgement (900 g DM/animal). Interestingly, fecal ALP markedly increased after the acidotic challenge. In fact, fecal ALP concentrations were 50-fold higher in ACID rams compared to CTR. The causes of this abrupt boost of ALP in feces of ACID rams are not clear. Data regarding ALP in feces of ruminants are lacking in the literature. However, Lehmann et al. (1981) proved that in rat the administration of toxic agents for the small intestinal mucosa (bleomycin or triparanol) were able to increase the release of the intestinal ALP into the feces after 1 d of treatment. For this reason several other


studies (Davis, 2003; Lapré et al., 1993; Taché et al., 2006) used the intestinal ALP in feces as marker of in vivo epithelial cell loss during intestinal damage investigations. Thomas and Henton (1985) did not find changes in the excretion of intestinal ALP after drug treatments with bleomycin in rats, which is a molecule able to cause the intestinal damage. Altogether this research supports ALP in feces as a marker of damage of the small intestinal epithelium in ruminants, but additional studies are advisable to confirm this hypothesis. In our rams, the clinical symptoms of anorexia were confirmed from changes in metabolic blood parameters. The fall of glucose and the rise of NEFA in plasma indicated an energy deficit state due to the fall of VFA consequent to the interruption of the feed intake and also to the impairment of the microbial activity (as discussed above). Generally, in fasted animals NEFA and BHBA have a similar pattern of change and are interpreted as markers of adipose tissue mobilization during energy deficiency. The same pattern of changes and the same interpretation was reported by Odongo et al. (2009) during metabolic acidosis in sheep. In the present experiment, blood BHBA had an unusual decline in ACID rams, likely related to the lower concentration of butyrate from rumen fermentation that is an important precursor of BHBA in healthy and nourished ruminants. The rise of plasma urea in acidotic sheep, observed also by Patra et al. (1996), can be interpreted as an index of the increased catabolism of AA of muscle tissues although an increase of ruminal ammonia (not monitored in this experiment) could have also happened. Moreover, in similar conditions, the urea increase in blood could be also due to a renal failure, as previously hypothesized by Dunlop (1972). The increase of blood bilirubin concentration in ACID rams is in agreement with the results obtained by Nikolov (1998) in buffalo and by Nikolov (2003) in sheep, and it may signal the


impairment of hepatic function (i.e., reduced bilirubin clearance). A decrease of plasma concentration of Ca, K, and Mg comparable to our results has been reported after acute acidosis induction in cows (Zebeli et al., 2010) and sheep (Irwin et al., 1979). The reduction of plasma Mg mainly reflects the reduced absorption of this mineral by the rumen (Schweigel and Martens, 2000), whereas the low plasma K could be justified by the reduction of feed intake, as well as the increased intestinal losses as consequence of diarrheic condition (Gennari, 1998), while the renal conservation of K takes several days to compensate the deficient potassium intake (Carlson, 1997). The low plasma Ca and Zn in ACID sheep can be interpreted as a consequence of the inflammatory condition (Bertoni and Trevisi, 2013) and it has been previously observed for lactating cows challenged with starch (Zebeli et al., 2010) and during intravenously infusion of LPS in cows (Waldron et al., 2003) and sheep (Bertoni et al., 1989). Beside this, a possible role of hyperaldosteronism during oxidative stress and consequent alteration of Ca and Mg metabolism as well as disturbance of electrolytes level could be supposed (Sun et al., 2006). A significant rise of blood lactulose concentration in ACID rams after oral ingestion indicates its paracellular absorption into systemic circulation. This event only happens when disruption of tight junctions in epithelial barrier occurs (Bjarnason et al., 1995; Meddings et al., 1999; Melichar et al., 2005). In a previous study (Minuti et al., 2013) an indomethacin-induced enteropathy model was used in sheep and proved that the lactulose test was a suitable, noninvasive, and sensitive method to evaluate gastrointestinal permeability in adult ruminants. Minuti et al. (2013) recorded the highest values of lactulose in blood at 6 h after oral dosage. In the present experiment, the blood lactulose peaked in ACID rams just after 2 h from LDT. Ruminal acidosis is associated to reticulorumen hypomotility (Kezar and Church, 1979) and reduction of rumen outflow rate (Harmon et al., 1985). The early appearance of lactulose in


blood might consequently suggests that its absorption mostly occurred before the small intestine. Thus, our results indicate that acute ruminal acidosis impairs epithelial barrier integrity in the ruminoreticulum, the abomasum, or both, allowing lactulose passage. In human and rats studies, the increase in intestinal permeability, measured using the lactulose test, has been found to be positively correlated with the presence of LPS in the systemic circulation (Schietroma et al., 2013; Liu et al., 2008) and also related with bacterial translocation (Yue et al., 2012). If this were confirmed, we can hypnotize that LPS as well as other bacterial immunogens, some other large molecules and even microbes could enter from the forestomachs into systemic circulation as well as interact with immune cells resident in the gastrointestinal mucosa (macrophage and dendritic cell) leading to local and systemic inflammation. It was previously found that sub-acute rumen acidosis (SARA) and acute rumen acidosis increase LPS concentration in rumen and stimulate systemic inflammatory response (Gozho et al., 2007; Nagaraja and Titgemeyer, 2007; Emmanuel et al., 2008; Khafipour et al., 2009a). Ismail et al. (2010) conducted a postmortem examination of rumen epithelium samples of goats after the induction of acute acidosis and observed easily detached ruminal papillae, patchy areas of sloughed ruminal papillae, hemorrhage in different areas of ruminal and reticular wall sometimes associated with ruminal wall ulceration, and similar results were observed during acute acidosis also by Mohamed Nour et al. (1998). Rumen wall is lined by keratinized stratified squamous epithelium, but is not protected by mucus, as it occurs with abomasal epithelium, and can be vulnerable to the chemical damage by acids. Low ruminal pH has been demonstrated to lead to rumenitis and eventually to parakeratosis, erosion, and ulceration of the ruminal epithelium (Garry, 2002). Moreover, acute and repeated episodes of ruminal acidosis are associated with prominent histological alterations in rumen epithelium which strongly suggest an impaired barrier function (Steele et al., 2009) and provide the


explanation for the translocation of toxins and bacteria during this disorder (Plaizier et al., 2008). In a recent paper (Plaizier et al., 2012), results from 2 models of induction of SARA (alfalfa pellet vs. grain; Khafipour et al., 2009a; b) were reviewed and led to the suggestion that during grain based SARA the translocation of LPS mainly occurs from the large intestine. In our study, the maximum concentrations of plasma lactulose were reached just 2 h after LDT, which suggests an increased permeability of the forestomachs barrier, becoming a site for LPS translocation during acute acidosis. In this study the consequences of acute ruminal acidosis on systemic inflammatory response were evident. Rectal temperature increased significantly in the ACID rams and this increment may be due to translocation of LPS into systemic circulation. Naylor and Kronfeld (1986) and Bertoni et al. (1989) found similar temperature increments after intravenous infusion of LPS in sheep. Haptoglobin and ceruloplasmin, both positive acute phase proteins, strongly increased in blood after 24 and 48 h, respectively, from the induction of acidosis. Negative markers of acute phase reaction, like Zn, paraoxonase, retinol, and albumin, decreased in blood 24, 48, 48, and 72 h, respectively, after the start of the acidosis challenge. The inflammatory responses during ruminal acidosis have been reported in the literature. Serum amyloid A and haptoglobin increased during SARA conditions (Khafipour et al., 2009a; Gozho et al., 2006, 2007). In agreement with our observations, Danscher et al. (2011), inducing acute ruminal acidosis in cattle by oligofructose overload, demonstrated that haptoglobin concentration in blood increases after 18 to 36 h from the challenge. In our study the intense oxidative stress, highlighted by blood markers changes (i.e. strong fall of antioxidant vitamin E and a rise of the concentration of ROMs) after 48 h from the onset of the rumen acidosis induction could support the damage of tissues by acidotic insult. Altogether these


hematic changes appear to be consistent with an inflammatory response mediated by immune cell resident in the gastrointestinal tract as well as translocation of LPS into bloodstream as consequence of the increased permeability of the gastrointestinal epithelia. In fact, LPS is a potent inducer of the acute phase response (Bertoni et al., 1989; Jacobsen et al., 2004) and its concentrations increases dramatically into the ruminal fluid during acidosis events (Nagaraja et al., 1978; Zebeli and Ametaj, 2009). In conclusion, the adopted model successfully induced acute rumen acidosis in sheep as indicated by severe clinical signs, the significant changes in rumen fluid parameters, the metabolic variations and the inflammatory condition. The large increase of lactulose concentration in blood after acidosis induction indicated a disruption of the gastrointestinal epithelium integrity and the consequent increase of its permeability to lactulose and likely also to large molecules. The earlier appearance of the lactulose in blood compared to a previous work using drugs known to damage intestinal epithelium suggests also that the permeability of rumen epithelium was likely altered by acidotic conditions. The results of this experiment suggest that the rumen represents a vulnerable area for the translocation of LPS, toxic biogenic amines, and perhaps different microbes during acute acidosis conditions. Results also established that lactulose can be a valuable probe to study rumen permeability in ruminants and that best times for collection of blood are from 2 to 6 h after oral dosage of the sugar. Finally, the present study also indicated the measurement of alkaline phosphatase in feces as potential marker of gastrointestinal damage in ruminants.


Literature Cited Ahmed, S., A. Minuti, and P. Bani. 2013. In vitro rumen fermentation characteristics of some naturally occurring and synthetic sugars. Ital. J. Anim. Sci. 12:359-365. Al Jassim, R.A.M., G.L.R. Gordon, and J.B. Rowe. 2003. The effect of basal diet on lactateproducing bacteria and the susceptibility of sheep to lactic acidosis. Anim. Sci. 77:459–469. Ametaj, B.N., B.J. Bradford, G. Bobe, R.A. Nafikov, Y. Lu, J.W. Young, and D.C. Beitz. 2005. Strong relationships between mediators of the acute phase response and fatty liver in dairy cows. Can. J. Anim. Sci. 85:165–175. Andersen, P.H., M. Hesselholt, and N. Jarløv. 1994. Endotoxin and arachidonic acid metabolites in portal, hepatic and arterial blood of cattle with acute ruminal acidosis. Acta Vet. Scand. 35:223–234. Bertoni, G., and E. Trevisi. 2013. Use of the liver activity index and other metabolic variables in the assessment of metabolic health in dairy herds. Vet. Clin. N. Am-Food A. 29:413–431. Bertoni, G., F. Piccioli-Cappelli, L. Calamari, and E. Trevisi. 1989. Digestive upsets of ruminants: possible role of endotoxins and/or histamine. In Proc. 7th Int. Conf. on Production diseases in farm animals,. Ithaca, NY USA. 370–373. Bionaz, M., E. Trevisi, L. Calamari, F. Librandi, A. Ferrari, and G. Bertoni. 2007. Plasma paraoxonase, health, inflammatory conditions, and liver function in transition dairy cows. J. Dairy Sci. 90:1740–1750. Bjarnason, I., A. MacPherson, and D. Hollander. 1995. Intestinal permeability: an overview. Gastroenterology. 108:1566–1581. Bouchard, L., S. Blais, C. Desrosiers, X. Zhao, and P. Lacasse. 1999. Nitric oxide production during endotoxin-induced mastitis in the cow. J. Dairy Sci. 82:2574–2581. Carlson, G.P. 1997. Fluid, Electrolyte, and Acid-Base Balance. In: Kaneko J.J., J.W. Harvey and Bruss editors ML In Clinical Biochemistry of Domestic Animals. Academic Press, San Diego, California, U.S.A. Connor, H., H.F. Woods, and J.G. Ledingham. 1983. Comparison of the kinetics and utilisation of D(-)-and L(+)-sodium lactate in normal man. Ann. Nutr. Metab. 27:481–487. Danscher, a. M., M.B. Thoefner, P.M.H. Heegaard, C.T. Ekstrøm, and S. Jacobsen. 2011. Acute phase protein response during acute ruminal acidosis in cattle. Livest. Sci. 135:62–69. Davis, C.D. 2003. Low dietary copper increases fecal free radical production, fecal water alkaline phosphatase activity and cytotoxicity in healthy men. J. Nutr. 133:522–527.


Dunlop, R.H. 1972. Pathogenesis of ruminant lactic acidosis. Adv. Vet. Sci. Comp. Med. 16:259–302. Emmanuel, D.G.V, S.M. Dunn, and B.N. Ametaj. 2008. Feeding high proportions of barley grain stimulates an inflammatory response in dairy cows. J. Dairy Sci. 91:606–614. Ewaschuk, J.B., J.M. Naylor, R. Palmer, S.J. Whiting, and G.A. Zello. 2004. D-lactate production and excretion in diarrheic calves. J. Vet. Intern. Med. 18:744–747. Farhadi, A., A. Keshavarzian, E.W. Holmes, J. Fields, L. Zhang, and A. Banan. 2003. Gas chromatographic method for detection of urinary sucralose: application to the assessment of intestinal permeability. J. Chromatogr. B. Analyt. Technol. Biomed. Life Sci. 784:145–154. Ferré, N., J. Camps, E. Prats, E. Vilella, A. Paul, L. Figuera, and J. Joven. 2002. Serum paraoxonase activity: a new additional test for the improved evaluation of chronic liver damage. Clin. Chem. 48:261–268. Garry, F.B. 2002. Indigestion in ruminants. In: Smith, B.P. (Ed.), Large Animal Internal Medicine. S.L. and B. Mosby, St. Louis and Baltimore,722–747. Gennari, F.J. 1998. Hypokalemia. N. Engl. J. Med. 339:451–458. Giesecke, D., and P. von Wallenberg. 1985. Metabolism of D(-)lactic acid in rats given high intragastral doses. Comp. Biochem. Physiol. B. 82:255–258. Gilliam, M.B., M.P. Sherman, J.M. Griscavage, and L.J. Ignarro. 1993. A spectrophotometric assay for nitrate using NADPH oxidation by Aspergillus nitrate reductase. Anal. Biochem. 212:359–365. Goad, D.W., C.L. Goad, C.L., and T.G. Nagaraja. 1998. Ruminal microbial and fermentative changes associated with experimentally induced subacute acidosis in steers. J. Anim. Sci. 76: 234–241. Gozho, G.N., D.O. Krause, and J.C. Plaizier. 2006. Rumen lipopolysaccharide and inflammation during grain adaptation and subacute ruminal acidosis in steers. J. Dairy Sci. 89:4404–4413. Gozho, G.N., D.O. Krause, and J.C. Plaizier. 2007. Ruminal lipopolysaccharide concentration and inflammatory response during grain-induced subacute ruminal acidosis in dairy cows. J. Dairy Sci. 90:856–866. Harmon, D.L., R.A. Britton, and R.L. Prior. 1984. In vitro rates of oxidation and gluconeogenesis from L(+)- and D(-)lactate in bovine tissues. Comp. Biochem. Physiol. B. 77:365–368.


Harmon, D.L., R.A. Britton, R.L. Prior, and R.A. Stock. 1985. Net portal absorption of lactate and volatile fatty acids in steers experiencing glucose-induced acidosis or fed a 70% concentrate diet ad libitum. J. Anim. Sci. 60:560–569. Huber, T.L. 1976. Physiological effects of acidosis on feedlot cattle. J. Anim. Sci. 43:902–909. Irwin, L.N., G.E. Mitchell, R.E. Tucker, and G.T. Schelling. 1979. Histamine, tyramine, tryptamine and electrolytes during glucose induced lactic acidosis. J. Anim. Sci. 48:367–374. Ismail, M., A.. Mahmound, M.. Nasr, and Y.. Badr. 2010. Clinical and laboratory studies on experimentally induced acute ruminal lactic acidosis in male goats. Alexandria J. Vet. Sci. 31:53–62. Jacobsen, S., P.H. Andersen, T. Toelboell, and P.M.H. Heegaard. 2004. Dose dependency and individual variability of the lipopolysaccharide-induced bovine acute phase protein response. J. Dairy Sci. 87:3330–3339. Kezar, W.W., and D.C. Church. 1979. Ruminal changes during the onset and recovery of induced lactic acidosis in sheep. J. Anim. Sci. 49:1161–1167. Khafipour, E., D.O. Krause, and J.C. Plaizier. 2009a. A grain-based subacute ruminal acidosis challenge causes translocation of lipopolysaccharide and triggers inflammation. J. Dairy Sci. 92:1060–1070. Khafipour, E., D.O. Krause, and J.C. Plaizier. 2009b. Alfalfa pellet-induced subacute ruminal acidosis in dairy cows increases bacterial endotoxin in the rumen without causing inflammation. J. Dairy Sci. 92:1712–1724. Kleen, J.L., G.A. Hooijer, J. Rehage, and J.P.T.M. Noordhuizen. 2003. Subacute ruminal acidosis (SARA): a review. J. Vet. Med. A. Physiol. Pathol. Clin. Med. 50:406–414. Lapré, J.A., H.T. De Vries, and R. Van der Meer. 1993. Cytotoxicity of fecal water is dependent on the type of dietary fat and is reduced by supplemental calcium phosphate in rats. J. Nutr. 123:578–585. Lehmann, F.G., H. Hufnagel, and H. Lorenz-Meyer. 1981. Fecal intestinal alkaline phosphatase: a parameter for toxic damage of the small intestinal mucosa. Digestion. 21:156–162. Lettat, A., P. Nozière, M. Silberberg, D.P. Morgavi, C. Berger, and C. Martin. 2010. Experimental feed induction of ruminal lactic, propionic, or butyric acidosis in sheep. J. Anim. Sci. 88:3041–3046. Littell, R., G. Milliken, W. Stroup, R. Wolfinger, and O. Schabenberger. 2006. SAS for Mixed Models, Second Edition. SAS Institute.


Liu, H., W. Li, X. Wang, J. Li, and W. Yu. 2008. Early gut mucosal dysfunction in patients with acute pancreatitis. Pancreas. 36:192–196. doi:10.1097/MPA.0b013e31815a399f. Loew F.M., and R.C. Chaplin. 1976. Abnormal acid production in the rumen: new findings from around the world. Bov. Proc., S. Franc, 141-164 Meddings, J.B., J. Jarand, S.J. Urbanski, J. Hardin, and D.G. Gall. 1999. Increased gastrointestinal permeability is an early lesion in the spontaneously diabetic BB rat. Am. J. Physiol. 276:G951–957. Melichar, B., J. Dvorák, R. Hyspler, and Z. Zadák. 2005. Intestinal permeability in the assessment of intestinal toxicity of cytotoxic agents. Chemotherapy. 51:336–338. Metzler-Zebeli, B U., S. Schmitz-Esser, F. Klevenhusen, L. Podstatzky-Lichtenstein, M. Wagner, and Q. Zebeli. 2013. Grain-rich diets differently alter ruminal and colonic abundance of microbial populations and lipopolysaccharide in goats. Anaerobe, 20:65–73. doi:10.1016/j.anaerobe.2013.02.005 Minuti, A., S. Ahmed, E. Trevisi, F. Piccioli-Cappelli, G. Bertoni, and P. Bani. 2013. Assessment of gastrointestinal permeability by lactulose test in sheep after repeated indomethacin treatment. J. Anim. Sci. 91:5646-5653 Mohamed Nour, M., M. Abusamra, and B.E. Hago. 1998. Experimentally induced lactic acidosis in Nubian goats: Clinical, biochemical and pathological investigations. Small Rumin. Res. 31:7– 17. Muir, L.A., E.L. Rickes, P.F. Duquette, and G.E. Smith. 1980. Control of wheat-induced lactic acidosis in sheep by thiopeptin and related antibiotics. J. Anim. Sci. 50:547–553. Nagaraja, T.G., and E.C. Titgemeyer. 2007. Ruminal acidosis in beef cattle: the current microbiological and nutritional outlook. J. Dairy Sci. 90 Suppl 1:E17–38. Nagaraja, T.G., and G. Town. 1990. Ciliated protozoa in relation to ruminal acidosis and lactic acid metabolism. In Rumen Ecosystem: Microbial Metabolism and Regulation. R. Oneora, H. Minato, and H. Itabashi, editors. New York, NY. 194. Nagaraja, T.G., and G.W. Miller. 1989. Rumen microbial changes in ionophore antibiotic-treated steers with experimentally induced acidosis. Australas. J. Anim. Sci. 2:465–468. Nagaraja, T.G., E.E. Bartley, L.R. Fina, and H.D. Anthony. 1978. Relationship of rumen gramnegative bacteria and free endotoxin to lactic acidosis in cattle. J. Anim. Sci. 47:1329–1337. Naylor, J.M., and D.S. Kronfeld. 1986. Relationships Between Metabolic Changes and Clinical Signs in Pregnant Sheep Given Endotoxin. Can J Vet Res. 50:402–409.


Nikolov, Y. 1998. Clinical experimental studies on acute rumen acidosis in buffaloes (Bubalus bubalis L.). V. Influence on several blood and rumen biochemical parameters. Vet. Arh. 68:205– 212. Nikolov, Y. 2003. Clinical experimental studies on acute rumen acidosis in sheep with a preexisting chronic lead and organophosphorus compound intoxication. I. Clinical and haematological changes. Trakia J. Sci. 1:61–66. Nocek, J.E. 1997. Bovine acidosis: implications on laminitis. J. Dairy Sci. 80:1005–1028. Odongo, N.E., S.L. Greenwood, M.M. Or-Rashid, D. Radford, O. Alzahal, A.K. Shoveller, M.I. Lindinger, J.C. Matthews, and B.W. McBride. 2009. Effects of nutritionally induced metabolic acidosis with or without glutamine infusion on acid-base balance, plasma amino acids, and plasma nonesterified fatty acids in sheep. J. Anim. Sci. 87:1077–1084. Owens, F.N., D.S. Secrist, W.J. Hill, and D.R. Gill. 1998. Acidosis in cattle: a review. J. Anim. Sci. 76:275–286. Patra, R.C., S.B. Lal, and D. Swarup. 1996. Biochemical profile of rumen liquor , blood and urine in experimental acidosis in sheep. Small Rumin. Res. 19:177–180. Plaizier, J.C., D.O. Krause, G.N. Gozho, and B.W. McBride. 2008. Subacute ruminal acidosis in dairy cows: the physiological causes, incidence and consequences. Vet. J. 176:21–31. Plaizier, J.C., E. Khafipour, S. Li, G.N. Gozho, and D.O. Krause. 2012. Subacute ruminal acidosis (SARA), endotoxins and health consequences. Anim. Feed Sci. Technol. 172:9–21. Prior, R.L. 1983. Plasma clearance of L- and D-lactate in steers fed alfalfa hay or high concentrate diets. J. Anim. Sci. 57:1029–1036. Saleem, F., B.N. Ametaj, S. Bouatra, R.Mandal, Q. Zebeli, S. M.Dunn, and D.S. Wishart. 2012. A metabolomics approach to uncover the effects of grain diets on rumen health in dairy cows. J. Dairy Sci. 95:6606–6623. doi:10.3168/jds.2012-5403 Schietroma, M., B. Pessia, F. Carlei, E.M. Cecilia, and G. Amicucci. 2013. Intestinal permeability, systemic endotoxemia, and bacterial translocation after open or laparoscopic resection for colon cancer: a prospective randomized study. Int. J. Colorectal Dis. 28:1651–1660. doi:10.1007/s00384-013-1751-4. Schweigel, M., and H. Martens. 2000. Magnesium transport in the gastrointestinal tract. Front. Biosci. 5:D666–677. Skinner, J.G., R.A. Brown, and L. Roberts. 1991. Bovine haptoglobin response in clinically defined field conditions. Vet. Rec. 128:147–149. Slyter, L.L. 1976. Influence of acidosis on rumen function. J. Anim. Sci. 43:910–929.


Steele, M.A, O. AlZahal, S.E. Hook, J. Croom, and B.W. McBride. 2009. Ruminal acidosis and the rapid onset of ruminal parakeratosis in a mature dairy cow: a case report. Acta Vet. Scand. 51:39. doi:10.1186/1751-0147-51-39. Sun, Y., R.A. Ahokas, S.K. Bhattacharya, I.C. Gerling, L.D. Carbone, and K.T. Weber. 2006. Oxidative stress in aldosteronism. Cardiovasc Res. 71:300–309. doi:10.1016/j.cardiores.2006.03.007 Sunderman, F.W., and S. Nomoto. 1970. Measurement of human serum ceruloplasmin by its pphenylenediamine oxidase activity. Clin. Chem. 16:903–910. Taché, S., G. Parnaud, E. Van Beek, and D.E. Corpet. 2006. Polyethylene glycol, unique among laxatives, suppresses aberrant crypt foci, by elimination of cells. Scand. J. Gastroenterol. 41:730– 736. Thomas, D.W., and D.H. Henton. 1985. The use of fecal alkaline phosphatase as an indicator of intestinal damage. Digestion. 31:82–88. Waldron, M.R., B.J. Nonnecke, T. Nishida, R.L. Horst, and T.R. Overton. 2003. Effect of lipopolysaccharide infusion on serum macromineral and vitamin D concentrations in dairy cows. J. Dairy Sci. 86:3440–3446. doi:10.3168/jds.S0022-0302(03)73948-4. Yue, C., B. Ma, Y. Zhao, Q. Li, and J. Li. 2012. Lipopolysaccharide-induced bacterial translocation is intestine site-specific and associates with intestinal mucosal inflammation. Inflammation. 35:1880–1888. doi:10.1007/s10753-012-9510-1. Zebeli, Q., and B.N. Ametaj. 2009. Relationships between rumen lipopolysaccharide and mediators of inflammatory response with milk fat production and efficiency in dairy cows. J. Dairy Sci. 92:3800–3809. Zebeli, Q., S. Sivaraman, S.M. Dunn, and B.N. Ametaj. 2011. Intermittent parenteral administration of endotoxin triggers metabolic and immunological alterations typically associated with displaced abomasum and retained placenta in periparturient dairy cows. J. Dairy Sci. 94:4968–4983. Zebeli, Q., S.M. Dunn, and B.N. Ametaj. 2010. Strong associations among rumen endotoxin and acute phase proteins with plasma minerals in lactating cows fed graded amounts of concentrate. J. Anim. Sci. 88:1545–1553. doi:10.2527/jas.2009-2203. Zust, J., U. Pestevsek, and A. Vengust. 2000. Impact of lactic acid fermentation in the large intestine on acute lactic acidosis in cattle. Dtsch. Tierarztl. Wochenschr. 107:359–363.


Figure 1. Experimental design. ▲= 0.5 kg wheat flour administration; ● = blood samples; ↓ = lactulose dosage time (LDT);  rumen and fecal samples collection; ■ = rectal temperature; grey box= gastrointestinal permeability assessment.


Figure 2. Effects of induction of acute ruminal acidosis on plasma concentration of (A) D-lactate, (B) L-lactate, (C) glucose, (D) β-hydroxybutyrate (BHBA). Values are means ± SEM. The treatments are control (CTR) or induction of acute ruminal acidosis (ACID). The triangle (▲) indicates the wheat flour administration during acute acidosis induction. Significance of differences between groups at each time point indicated by * for P < 0.05 and ** for P < 0.01.


Figure 3. Effects of induction of acute ruminal acidosis on plasma concentration of (A) haptoglobin, (B) ceruloplasmin, (C) zinc, (D) paraoxonase , (E) Reactive Oxygen Metabolites or ROMs, (F) vitamin E. Values are means ± SEM. The treatments are control (CTR) or induction of acute ruminal acidosis (ACID). The triangle (▲) indicates the wheat flour administration during acute acidosis induction treatment. Significance of differences between groups at each time point indicated by * for P < 0.05 and ** for P < 0.01.


Figure 4. Effect of induction of acute ruminal acidosis on plasma concentration of lactulose after its oral administration. The treatments are control (CTR) or induction of acute ruminal acidosis (ACID). Significance of differences between groups at each time point indicated by * for P < 0.05 and ** for P < 0.01.


Table 1. Effects of induction of acute ruminal acidosis in rams on ruminal pH, L lactate, D lactate, total volatile fatty acids and their molar proportions after 24 h from induction of acidosis (mean ± SEM). The treatments are control (CTR) or induction of acute ruminal acidosis (ACID). Treatment Item

CTR

ACID

SEM

P-value

pH

6.89

4.94

0.13