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¹ Department of Veterinary Physiology, Faculty of Veterinary Medicine, Freie Universität Berlin, Oertzenweg 19b, 14163 Berlin, Germany

	Abstract

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The intention of this study was to determine the effects of mucosal osmotic pressure on transport and barrier functions of the rumen epithelium of sheep, which were fed various diets: hay ad libitum, or 600, 1200 or 1800 g day^–1 of a supplemented diet plus hay ad libitum. The experiments were conducted by using the conventional Ussing chamber technique. Mucosal osmolarity was adjusted to 300 (control), 375 or 450 mosmol l^–1. Feeding of a supplemented diet led to a significant increase of mucosal to serosal Na⁺ transport and net Na⁺ transport, probably because of an increase of apical Na⁺–H⁺ exchange activity. An increase in mucosal osmotic pressure: (a) reduced net Na⁺ transport in all feeding groups, the remaining net Na⁺ transport being higher in tissues of sheep fed a supplemented diet; (b) increased transepithelial tissue conductance, this rise being smallest with a high intake of the supplemented diet; and (c) enhanced the serosal to mucosal Na⁺ transport in tissues of hay-fed sheep and sheep fed with 600 g day^–1 of the supplemented diet, while higher intakes of the supplemented diet (1200 and 1800 g) did not produce any effect. All these changes indicate a diet-dependent adaptation to luminal hypertonicity.

(Received 31 August 2005; accepted after revision 30 January 2006; first published online 1 February 2006)
Corresponding author U. Lodemann: Department of Veterinary Physiology, Faculty of Veterinary Medicine, Freie Universität Berlin, Oertzenweg 19b, 14163 Berlin, Germany. Email: lodemann{at}zedat.fu-berlin.de

	Introduction

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The rumen is an important site for the absorption of short-chain fatty acids (SCFA) and other electrolytes, such as Na⁺, K⁺, Mg²⁺ and Cl^– (Dobson, 1959; Warner & Stacy, 1972; Ferreira et al. 1972; Chien & Stevens, 1972; Martens et al. 1978; Bugaut, 1987). Hence, the rumen epithelium exhibits a variety of transport mechanisms, and its barrier function maintains relevant electrochemical gradients, which are prerequisites for an active (net) transport against a gradient. According to the classification of Powell (1981), the rumen epithelium belongs to the family of ‘moderately tight’ epithelia, such as that found in the colon. The lamina epithelialis mucosae of the ruminal tunica mucosa consists of a stratum basale, stratum spinosum, stratum granulosum and stratum corneum.

The composition of the ruminal fluid varies considerably with the feeding regime, and the ruminal epithelium is exposed to daily cyclic changes of pH, SCFA concentrations (Allen, 1997) and osmotic pressure. Before feeding, the rumen content is hypotonic with respect to blood (< 280 mosmol kg^–1; Warner & Stacy, 1965; Engelhardt, 1969). The osmotic pressure of the rumen fluid rises after a meal to maximal values of 350 mosmol l^–1 or 400 mosmol kg^–1, depending upon the type of diet (Warner & Stacy, 1965; Engelhardt, 1969; Bennink et al. 1978). This is mainly attributable to a rise in electrolytes and an increase in ammonium and SCFA concentration by fermentation (Warner & Stacy, 1965; Engelhardt, 1969). Bennink et al. (1978) have found a linear correlation between the concentration of SCFA and the osmotic pressure of the ruminal fluid.

Luminal osmotic pressure influences the functions and integrity of epithelia, and the effect of osmotic pressure depends upon the type of epithelium (‘tight’, ‘moderately tight’ or ‘leaky’; for details see Ussing, 1965; Erlij & Martinez-Palomo, 1972; DiBona, 1972; Gemmel & Stacy, 1973; Bindslev et al. 1974; Reuss & Finn, 1977; Oshio & Tahata, 1984; Soybel et al. 1987; Gäbel et al. 1987a; Tabaru et al. 1990).

Studies with the isolated epithelia of hay-fed (HF) sheep in our laboratory have shown that an increase of the ruminal osmotic pressure (ROP) leads to an elevation of the transepithelial tissue conductance (G_t) caused by a reduction of the paracellular resistance (Freyer & Martens, 1998; Schweigel et al. 2005). The luminal osmotic pressure (350 and 450 mosmol l^–1) also decreases the net Na⁺ transport by inhibiting apical Na⁺–H⁺ exchange activity and by increasing the passive backflow of Na⁺ and fluxes of Cr-EDTA (Schweigel et al. 2005). This change of passive paracellular permeability is in agreement with in vivo observations made in cows by Dobson et al. (1976). Cr-EDTA, a marker of paracellular permeability, is significantly absorbed from the temporarily isolated rumen of a cow at hypertonic ruminal osmotic pressure (hyperROP). The effect of hyperROP on rumen epithelium has a morphological equivalent. Studies by Gemmel & Stacy (1973) have demonstrated that hyperROP induces a widening of the intercellular spaces of the stratum basale and a disruption of the cell junctions in the stratum granulosum (zonulae occludentes, which may represent the permeability barrier of the rumen epithelium; Graham & Simmons, 2005). Dilatation of the lateral intercellular spaces has also been observed in Necturus antrum upon hypertonic challenge (Soybel et al. 1987).

The diet-dependent adaptation of the rumen epithelium is well known and includes: (a) an increase in the size and number of the rumen papillae (Dirksen et al. 1984); (b) an increase in the transport rates of SCFA (Dirksen et al. 1984), Na⁺ and Mg²⁺ (Gäbel et al. 1987a; Shen et al. 2004) and of Ca²⁺ (Uppal et al. 2003a); and (c) a change in the activity of propionyl coenzyme A synthetase (Nocek et al. 1980). The underlying mechanisms of this adaptation are not known, but these alterations of morphology, functions and biochemical activities have been observed in tissues of animals fed with concentrate in addition to hay or silage. Concentrate feeding is accompanied by a pronounced increase of ruminal osmotic pressure (Bennink et al. 1978). We have therefore hypothesized that an increase in the intake of concentrate (or, in our case, a supplemented diet, SD) and the corresponding osmotic challenge of the rumen epithelium could induce reduced sensitivity or enhanced defence mechanisms against hyperosmolarity; this would reduce the negative effects of hyperosmolarity on Na⁺ transport and tissue conductance observed in the tissues of HF animals.

The aim of the present experiments has therefore been to elucidate the consequences of luminal hyperosmolarity on Na⁺ transport and tissue conductance of the isolated rumen epithelium from HF sheep and from sheep fed increasing amounts of a supplemented diet.

The results obtained confirm the stimulating effect of feeding of a supplemented diet on Na⁺ transport under isosmotic conditions (300 mosmol l^–1). Luminal hyperosmolarity (375 and 450 mosmol l^–1) causes a decrease of net Na⁺ transport (J_netNa) and an increase of tissue conductance in tissues of HF animals (paracellular permeability) and animals fed with a supplemented diet (SF animals). However, these effects are much more pronounced in the epithelia of HF sheep. There is no change of paracellular permeability with a high intake of the supplemented diet. The results indicate, for the first time, that adaptation includes mechanisms that partly shield the transport and barrier function of the rumen epithelium from osmotic challenge.

	Methods

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Animal feeding and origin of tissue specimens

Animal use was in accordance with European and German law. Twenty sheep weighing 55–70 kg were assigned at random to four groups and were kept under the following feeding regimes for at least 3 weeks: (1) hay ad libitum (HF group); (2) 600 g supplemented diet per day (SF600 group); (3) 1200 g supplemented diet per day (SF1200 group); and (4) 1800 g supplemented diet per day (SF1800 group). In addition to the supplemented diet, sheep were offered hay ad libitum. The animals also had free access to tap water and a mineral supplement block. The intake of hay (ad libitum) was 2–2.4 kg day^–1, which decreased with increasing amounts of supplemented diet and was approximately 0.5 kg day^–1 in the SF1800 group. The supplemented diet (SD) contained 16% crude protein, 3% crude fat, 13% crude fibre, 9.5% crude ash, 1.2% calcium, 0.5% phosphorus and 0.5% sodium, with an energy level of 5.9 MJ metabolizable energy (ME) kg^–1. The intake of ME was 14 MJ in HF sheep and increased up to 20 MJ in SF1200 sheep. This intake of energy covered 140% of maintenance requirement in HF and 200% in SF1200 sheep.

Tissue preparation

The preparation and incubation of rumen epithelium has been described in detail by Martens et al. (1987). Briefly, the sheep were stunned with a captive bolt and killed by exsanguination in a local slaughterhouse. After death, the abdominal cavity was opened and the ruminal sac removed. A 250 cm² piece of the rumen wall was taken from the ventral sac. After being rinsed with transport buffer solution, the epithelium was stripped from the muscle layer and taken to the laboratory in the transport buffer, gassed with Carbogen (95% CO₂, 5% O₂) and kept at 38°C. It was then cut into squares (3 x 3 cm) and mounted between the halves of a conventional Ussing chamber with an exposed area of 3.14 cm². Edge damage was avoided by placing rings of silicone rubber on both sides of the tissue. Approximately 30–60 min elapsed from the killing of the animals to the tissue being mounted in Ussing chambers.

Incubation procedure

During the equilibration period (30 min), the tissues were bathed on both sides with 18 ml control buffer solution. The temperature was held constant at 38°C by a thermostat (Haake D1, VWR international GmbH, Darmstadt, Germany) by circulating heated water between the two water jackets. The buffer solution was mixed and gassed with Carbogen by a gas lift system.

At the end of the equilibration period, the mucosal buffer solution was exchanged and replaced with buffer solution with an osmotic pressure of 375 or 450 mosmol l^–1.

Electrical measurements

Electrical measurements were obtained by a microcomputer-controlled Voltage/Current Clamp (Version 2.02 by K. Mußler, Aachen, Germany). Two KCl Agar bridges were positioned near each surface of the tissue (< 3 mm) and connected to Ag–AgCl electrodes for measurement of the transepithelial potential difference (PD_t), while two more distant (> 20 mm) KCl Agar bridges were connected to Ag–AgCl electrodes for passing direct current through the tissue. The junction potential and the fluid resistance of the buffer between the tips of the potential difference-sensing bridges were determined before the tissue was mounted and were subsequently corrected by the computer-controlled voltage clamp.

The tissue was alternatively pulsed with a positive or negative pulse of 50 µA and 200 ms duration. The displacement in PD_t caused by the pulse was measured and the G_t was calculated from the change in PD_t and pulse amplitude. A continuous recording was made of PD_t (in mV), current (in µequiv · cm^–2· h^–1) and G_t (in mS · cm^–2). After an equilibration period of 15 min under open-circuit conditions, the epithelium was short circuited.

Flux measurements (²²Na⁺ fluxes)

After reaching a steady state (approximately 20 min), two epithelia that differed by less than 25% in G_t and short-circuit current (I_sc) were paired for Na⁺ fluxes. Unidirectional fluxes of Na⁺ were measured by using ²²Na⁺. The isotope (80 kBq) was added to one side of the epithelium, and the tissues were incubated for 30 min to allow equilibration of the isotope. Fluxes were calculated from the rate of appearance of tracer on the other side of the epithelium within 60 min. In one of the paired epithelia, the ion flux from mucosal to serosal side (absorption) and, in the other, the ion flux from serosal to mucosal side (secretion) was measured. ²²Na⁺ was assayed by using a well-type crystal counter (LKB Wallace-PerkinElmer, Überlingen, Germany).

In order to differentiate the flow of Na⁺ across an epithelium into a potential-dependent (J) and a potential-independent component (J_ß; see model below of Frizzell & Schultz, 1972) the epithelia were incubated under voltage clamp conditions. The sequence of the clamp potential was randomized: –25 mV; 0 mV; +25 mV; or +25 mV; 0 mV; and –25 mV. Each clamp period had a duration of 90 min.

Solutions and reagents

The rinsing and transport buffer solution consisted of (mM): 115 NaCl, 25 NaHCO₃, 0.40 NaH₂PO₄, 2.40 Na₂HPO₄, 5 KCl, 5 glucose, 1.20 CaCl₂ and 1.20 MgCl₂, kept at a pH of 7.4. The standard buffer solution for incubation contained (mM): 60 NaCl, 25 NaHCO₃, 1 KH₂PO₄, 2 K₂HPO₄, 10 glucose, 8 Mops, 25 sodium acetate, 10 sodium propionate, 5 sodium butyrate, 35 mannitol, 1 CaCl₂ and 1 MgCl₂, and was adjusted to a pH of 7.6–7.7 with Trizma (tris-hydroxymethyl-aminomethan) base (1 M) at room temperature without gassing. All reagents were purchased from Merck (VWR International GmbH, Darmstadt, Germany) and were of analytical grade.

Mucosal osmolarity was adjusted by mannitol to 300 mosmol l^–1 in the control group and to 375 and 450 mosmol l^–1 in the experimental groups. The final pH of the buffer solution in the Ussing chamber was 7.3–7.5.

Model of Frizzell & Schultz (1972)

The potential-dependent and the potential-independent parts of a (unidirectional) ion flux were calculated according to the following equation (Frizzell & Schultz, 1972; modified according to Jackson & Norris, 1985; see equations 1 and 2 and Fig. 1).

(1)

where J is the unidirectional flux (in µequiv · cm^–2· h^–1), J· is the potential-dependent part of the flux and J_ß is the potential-independent part of the flux. The electrical driving force, , is defined as:

(2)

where PD_t is the Potential difference across the tissue in direction of the flux, , z is the charge of the ion, F is the Faraday constant (96478 C · mol ^–1), R is the gas constant (8.3143 J · K^–1· mol^–1) and T is the absolute temperature (38°C = 311 K). at 0 PD_t is defined as 1.

View larger version (8K): [in this window] [in a new window]   Figure 1.  Scheme of a plot according to the equation ofFrizzell & Schultz (1972) The intercept of the y axis (Jß) represents the potential difference-independent part of ion flux, and the slope represents the potential difference-dependent part of ion flux.

Statistical analysis

Statistical evaluations were carried out by means of SPSS program for Windows, versions 9 and 10 (Jandel, Chicago, IL, USA). The graphs were plotted with Sigma Plot 5.0.

Unless otherwise stated, results are presented as means ±S.E.M. Significant effects of the treatment were reported as P < 0.05. N refers to the number of experimental animals and n to the number of epithelial tissues per treatment group.

Analysis of variance was carried out as indicated below.

G_t curves.  A one-factorial variance analysis (independent variable, ‘osmolarity’; dependent variable, G_t) was calculated for each feeding group with Dunnett's post hoc two-sided tests, if the factor ‘osmolarity’ proved to be significant (point of reference: 300 mosmol l^–1).

G_t and J_netNa under short-circuit conditions.  A variance analysis with a two-factorial design was calculated (4 feeding groups, 3 osmolarity levels and their interaction; ‘feeding groups’ and ‘osmolarity levels’ as fixed factors). Dunnett's post hoc test T3 was used.

In a second step, one-factorial variance analyses were conducted for the comparison of the four feeding groups within the three osmolarity levels (3VAs) and the three osmolarity levels within the feeding groups (4VAs). In order to avoid the value increasing uncontrollably above the nominal value (0.05) the variance analyses were conducted on a ‘family-wise’ adjusted '-level ('=/3 or /4, i.e. '= 0.017 or 0.0125). Dunnett's test one-sided post hoc tests or Dunnett's two-sided T3 tests were conducted.

G_t (G_t at 30 min minus G_t at 0 min), Na transport from the mucosal to the serosal side (J_msNa), Na transport from the serosal to the mucosal side (J_smNa) and J_netNa (+ 25 mV).  For comparison of the four feeding groups per osmolarity level, one-factorial variance analyses (‘feeding group’ as ‘fixed factor’) with Dunnett's post hoc tests were used.

The comparision of the three osmolarity levels per feeding group was likewise performed by a variance analysis with ‘osmolarity’ as ‘fixed factor’ and animal as ‘random factor’, with Dunnett's two-sided post hoc test.

J and J_ß of J_ms and J_sm.  A regression curve was calculated for each animal to obtain slopes (J) and intercepts (J_ß) for J_ms or J_sm. Variance analyses were calculated as indicated in the paragraph above.

	Results

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Tissue conductance

After the equilibration period, the mucosal buffer solution was replaced with hypertonic solutions (375 or 450 mosmol l^–1) in the experimental groups. This osmotic challenge caused a rapid and dose-dependent increase of G_t. However, diet influenced the extent of the change in G_t. Examples are given in Figs 2 and 3. In tissues of HF sheep, an osmolarity of 375 or 450 mosmol l^–1 caused a significant increase of G_t (Fig. 2).

View larger version (18K): [in this window] [in a new window]   Figure 2.  Tissue conductance (Gt) as a function of time under control conditions (300 mosmol l–1) and mucosal hyperosmolarity (375 or 450 mosmol l–1) in tissues of hay-fed sheep The sequence of the clamp potential was randomized: –25 mV; 0 mV; +25 or +25 mV; 0 mV; and –25 mV. Each clamp period had a duration of 90 min. Asterisks indicate values that differ significantly at the respective time from the control values (P < 0.05). Values are means ±S.E.M.

View larger version (18K): [in this window] [in a new window]   Figure 3.  Tissue conductance (Gt) as a function of time under control conditions (300 mosmol l–1) and mucosal hyperosmolarity (375 or 450 mosmol l–1) in tissues of SF1200 sheep The sequence of the clamp potential was randomized: –25 mV; 0 mV; +25 or +25 mV; 0 mV; and –25 mV. Each clamp period had a duration of 90 min. Asterisks indicate values that differ significantly at the respective time from the control values (P < 0,05). Values are means ±S.E.M.

Table 1 summarizes the alterations of G_t (G_t), which were determined by subtracting the G_t value before buffer exchange from the G_t value obtained 30 min after mucosal buffer exchange (osmotic challenge). The data in Table 1 clearly show that the effects of osmotic pressure depend on the feeding regime, in that: (a) G_t in tissues of HF animals exhibited a linear and positive correlation with the osmotic pressure (r= 0.85, r²= 0.73, P < 0.0001); (b) G_t in tissues of SF sheep moderately increased at 375 mosmol l^–1 (0.4 to 0.85 mS · cm^–2) and was much more pronounced at 450 mosmol l^–1 (1.62 to 2.35mS · cm^–2); (c) G_t decreased at 375 and 450 mosmol l^–1 with increasing intakes of SD (600 or 1200 g day^–1); and (d) G_t did not follow this correlation in tissues of SF1800 sheep.

View this table: [in this window] [in a new window]   Table 1. Change of tissue conductance (Gt=Gt 30 min after buffer exchange minus Gt before buffer exchange) as a function of feeding regime and osmotic pressure

Na⁺ transport

Since Na⁺ transport has been described in detail in rumen epithelium (Chien & Stevens, 1972; Martens et al. 1991; Gäbel et al. 1991,), this transport mechanism was chosen as a model for osmotic-dependent effects on active ion transport. Previous studies in our laboratory with tissues from HF animals have shown that an increase of the luminal osmotic pressure causes no significant effect on mucosal to serosal Na⁺ transport (J_msNa) but an increase of serosal to mucosal Na⁺ transport (J_smNa), both of which lead to a significantly reduced J_netNa (Schweigel et al. 2005). Tables 2 and 3 summarize the flux rates J_msNa and J_smNa; the results of the present study (tissues of HF and SF600 sheep) closely agree with previous observations.

View larger version (31K): [in this window] [in a new window]   Figure 4.  Net Na+ transport under short-circuit conditions Values are means.

When significant osmotic-dependent changes of Na⁺ flux rates occurred, they were characterized by a decrease of J_msNa and an increase of J_smNa (Tables 2 and 3). We hypothesized that the decrease of J_msNa was caused by an inhibition of electroneutral Na⁺ transport and the enhanced J_smNa was the result of a decrease in the resistance of the paracellular pathway. Frizzell & Schultz (1972) developed a model such that the flow of an ion across an epithelium could be differentiated into a potential-dependent (J) and a potential-independent component (J_ß; see Methods). We were interested in these two parameters according to the model of Frizzell & Schultz (1972): the potential difference-independent flux of J_msNa and the potential difference-dependent flux of J_smNa.

The potential difference-independent component of J_msNa (J_ß) was assumed to be represented by electroneutral Na⁺–H⁺ exchange, and the suggested osmotic-dependent inhibition of this exchanger should have led to a reduction of J_ß (intercept of the y axis; see Fig. 1). This was indeed the case. The hypothesis of possible effects of hyperosmolarity on J_ms was confirmed by the obtained alterations of J_ß. Table 4 summarizes the effect of hyperosmotic mucosal solution on J_ß of J_msNa. Although not all changes of J_ß were significantly different, the data support the assumption of the inhibition of electroneutral Na⁺ transport via Na⁺–H⁺ exchange, because pretreatment of rumen epithelia with amiloride abolished an effect of hyperosmolarity (Schweigel et al. 2005). Furthermore, diet-dependent effects are apparent. An osmolarity of 375 mosmol l^–1 did not change J_ß in tissues of SF600 and SF1200 sheep and depressed it in epithelia of SF1800 sheep. The high osmolarity of 450 mosmol l^–1 caused a reduction of J_ß in all epithelia.

View this table: [in this window] [in a new window]   Table 5. The slopes (J; potential difference-dependent Na+ flux) of JsmNa according to the model of Frizzell & Schultz (1972) as a function of feeding regime and osmotic pressure

The applied potential difference of +25 mV (serosal side positive) caused, in all cases, a reduction of J_msNa and an increase of J_smNa and so a reduced J_netNa, which is transformed into a net secretion of Na⁺ to the lumen side in tissue of HF sheep by hyperROP (Fig. 5). The potential difference of +25 mV is at the lower range of the normal transepithelial potential difference of the rumen epithelium (Martens & Blume, 1987), which clearly shows that the impairment of the shunt resistance by hyperosmolarity facilitates the passive and potential difference-driven J_smNa and reduces J_netNa. However, this effect was attenuated by the feeding of SD. There was still a net transport of Na⁺ in tissues of SF1200 and SF1800 sheep.

View larger version (13K): [in this window] [in a new window]   Figure 5.  Net Na+ transport under voltage clamp conditions (+25 mV) for the 4 feeding groups Values are means. Exemplar ± 1 s.

	Discussion

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Diet has long been known to influence physiological and biochemical functions (Nocek et al. 1980; Dirksen et al. 1984; Shen et al. 2004) and the morphology of the rumen epithelium (Liebich et al. 1987). Ruminal transport mechanisms are significantly enhanced by concentrate feeding (Dirksen et al. 1984; Gäbel et al. 1987a; Doreau et al. 1997; Uppal et al. 2003b). We have hypothesized that the diet-dependent adaptation of the rumen epithelium also includes the protection of transport mechanisms and barrier function caused by hyperROP in the ruminal epithelium of HF sheep.

The data obtained in the present study are in agreement with results from previous investigations; Na⁺ transport is significantly enhanced in tissues of SF sheep (Uppal et al. 2003b) and hyperROP significantly reduces J_netNa and generally causes changes of G_t (Schweigel et al. 2005). However, important diet-dependent differences have been observed.

Influence of increased osmotic pressure on G_t

The transepithelial tissue conductance increases immediately after the elevation of mucosal osmotic pressure (375 and 450 mosmol l^–1). The correlation between the increase of osmotic pressure and the change of G_t is linear in the tissues of HF animals and is in agreement with results from previous studies in our laboratory (Freyer & Martens, 1998; Schweigel et al. 2005) and other investigations of rumen epithelia (Gemmel & Stacy, 1973; Dobson et al. 1976). The effect of hyperROP on the increase of G_t is attenuated in tissues of SF sheep (375 and 450 mosmol l^–1) and exhibits a non-linear increase with the elevation of osmotic pressure. The osmotic-dependent alterations of G_t are reversible after the reconstitution of isosmotic conditions; this has also been observed previously (Dobson et al. 1976; Soybel et al. 1987; Schweigel et al. 2005).

The change of G_t can be explained by alterations of the cellular, G_c, or paracellular conductance, G_p, or both. The osmotic changes of G_t, G_c or G_p seen in the present study obviously depend on the feeding regime. Hypertonic mucosal solutions induce an increase of the conductance of the paracellular pathway, G_p, in tissues of HF sheep, as indicated by the increase of J_smNa and J_ß; this is in agreement with previous results (Schweigel et al. 2005). The change in paracellular conductance is the result of two effects. Hypertonic luminal solutions induce a decrease in cell volume and, hence, an enlargement of the paracellular pathway (Ussing, 1965; Gorodeski et al. 1995). Furthermore, the osmolyte (mannitol) diffuses into the space between the epithelial cells, and the subsequent osmotic-dependent flow of water into this compartment increases the paracellular conductance (DiBona & Civan, 1973; Civan & DiBona, 1974; Gorodeski et al. 1995) and, consequently, passive permeability. The osmotic-dependent increase of J_smNa is significant in tissues of HF sheep and is of the same magnitude in SF600 epithelia, supporting the conclusion that the epithelia become more ‘leaky’ at hyperROP. However, the tissues of SF1200 and SF1800 sheep do not show a change of J_smNa upon osmotic challenge, despite an increase of G_t. This supports the assumption that G_t predominantly represents alterations of cellular conductance, G_c, i.e. the conductance of the apical or basolateral membrane in these tissues.

The increase of osmotic pressure in the ruminal fluid most probably induces cell volume regulatory mechanisms (Lang et al. 1998). Although, to our knowledge, nothing is known about these mechanisms in the rumen epithelium, activation of conductance occurs in epithelia of SF1200 and SF1800 sheep. The types of changed conductance (anion or cation), their location (apical or basolateral) and the kind of ion transport for osmotic-dependent cell volume regulation are not known. Indeed, it has been shown previously that the effect of hyperROP on I_sc is small and transient (Schweigel et al. 2005).

Na⁺ transport

Effects of intake of the supplemented diet (isosmotic conditions).  Increasing the amount of SD intake enhances J_msNa and J_netNa (see Results, Table 2 and Fig. 4), in agreement with previous studies in goats (Shen et al. 2004) and sheep (Uppal et al. 2003b) and in agreement with the general observation that all the transport mechanisms of the rumen epithelium so far studied are stimulated by concentrate feeding: SCFA (Dirksen et al. 1984; Liebich et al. 1987; Doreau et al. 1997;), Na⁺, Cl^– and Mg²⁺ (Gäbel et al. 1987, 1991a; Gäbel, 1988), and Ca²⁺ (Uppal et al. 2003a). Furthermore, the conclusion that the enhanced J_msNa results from increased Na⁺–H⁺ exchange activity (Gäbel et al. 1987a; Martens & Gäbel, 1988; Shen et al. 2004) is supported by the application of the model of Frizzell & Schultz (1972). The potential-independent component of J_msNa rises with increasing SD intake and very probably represents electroneutral Na⁺ transport via Na⁺–H⁺ exchange (see Results and Table 4). The potential-dependent component remains the same. Serosal to mucosal Na⁺ transport does not change significantly with increasing SD intake under control conditions (at 300 mosmol l^–1), but is slightly higher in tissues of SF1200 and SF1800 sheep. Because this flux direction is predominantly paracellular and passive, the permeability of the shunt pathway is not significantly influenced by the feeding regime under control conditions.

Effects of an acutely elevated mucosal osmotic pressure.  We have shown, in a study with rumen epithelia of HF sheep, that hyperROP causes an inhibition of electroneutral Na⁺ transport via Na⁺–H⁺ exchange and that inhibition of Na⁺–H⁺ exchange by amiloride abolishes the effects of hyperROP on electroneutral Na⁺ transport (Schweigel et al. 2005). Furthermore, an acid load of isolated rumen epithelial cells with SCFA caused a prompt acidification of the cytosol followed by a rapid recovery of intracellular pH (Schweigel et al. 2005). This recovery was abolished by amiloride and a combination of S3226 Na⁺/H⁺ exchanger 3 (NHE3) inhibitor and HOE 694, Na⁺/H⁺ exchanger 1 (NHE1) inhibitor or by hyperosmolarity. These observations and the data of the present study are consistent with the assumption that electroneutral Na⁺ transport (J_msNa) is most probably mediated via the Na⁺–H⁺ exchanger, which is modulated by hyperosmolarity and represented by NHE3, because its mRNA has been detected in rumen epithelia (Schweigel et al. 2005). The suggested location of NHE3 in the luminal membrane is currently under investigation by immunostaining.

Osmotic-dependent inhibition of electroneutral Na⁺ transport has been demonstrated in a variety of epithelia or cell lines, and the inhibited Na⁺ transport mechanisms have been identified in these tissues as NHE3 (Weinman et al. 1987; Watts & Good, 1994; Soleimani et al. 1994; Kapus et al. 1994; Nath et al. 1996). The assumption of the inhibition of electroneutral Na⁺ transport is further supported by our data calculated according to the model of Frizzell & Schultz (1972). A reduction of the potential-independent component of J_msNa was observed in the epithelia of all feeding groups, when the mucosal osmotic pressure was increased. Nevertheless, the effect of hyperROP on J_msNa was much more pronounced in tissues of SF sheep, because this transport mechanism is upregulated in the tissues of these animals (Shen et al. 2004). Net Na⁺ transport was, however, significantly higher in tissues of SF sheep, despite the marked reduction of J_msNa (see Fig. 4).

Two general types of regulation of NHE3 have been described: an increase of exchanger number by insertion of NHE3 molecules into the luminal membrane by trafficking from subluminal vesicles or an increase in the activity of the exchanger. There is no evidence for vesicle trafficking under hypersomotic challenge (Donowitz & Tse, 2001). The regulation of NHE3 activity is mediated via the cytoplasmic C-terminus by a variety of factors (Zachos et al. 2005). However, the C-terminal domain of NHE, which is involved in the regulation of NHE by growth factors and protein kinase, is not necessary for the effect of hyperosmolarity on NHE (Nath et al. 1996). Uncertainty still exists regarding the inhibition of NHE3 by hyperosmolarity, and our limited knowledge concerning NHE3 in rumen epithelium does not provide an explanation, which is beyond the scope of this study.

The evident effect of hyperROP on J_msNa via the inhibition of NHE (J_ß) is complicated by the finding that the increase of G_t is partly attributable to an increase in the conductance of the paracellular pathway and, hence, of the passive part of J_msNa. Consequently, the effect of hyperROP on J_msNa is a combined effect of the inhibition of NHE3 and the increase of the passive part of J_msNa in tissues of HF and SF600 sheep.

The results of the present study are in agreement with earlier observations that hyperROP disturbs transport mechanisms. HyperROP (> 340 mosmol l^–1) leads to a flow of water into the rumen (Dobson et al. 1976), which appears to reduce Na⁺ absorption at a high influx of water of 290 ml h^–1 (Warner & Stacy, 1972; Tabaru et al. 1990). In addition, the absorption of SCFA is reduced at high luminal osmotic pressure (Bennink et al. 1978; Tabaru et al. 1990; Lopez et al. 1994).

The inhibition of Na⁺ transport by hyperROP seen here is not in agreement with results from a previous study. Stacy & Warner (1966) have concluded that hyperROP enhances Na⁺ absorption when ROP is increased by intraruminal infusion of mannitol plus urea, by KCl or K⁺ salts of SCFA. However, the positive effect on Na⁺ transport under these experimental conditions can now be explained by our improved knowledge of ruminal Na⁺ transport and has been discussed in detail by Schweigel et al. (2005). Briefly, Na⁺ is predominantly absorbed via Na⁺–H⁺ exchange in the luminal membrane (Chien & Stevens, 1972; Martens et al. 1991; Sehested et al. 1996). Na⁺–H⁺ exchange is stimulated by the luminal presence of SCFA (Gäbel et al. 1991,; Sehested et al. 1996) and NH₄⁺, which is released from urea by bacterial urease (Abdoun et al. 2003, 2005). Furthermore, an increase of ruminal K⁺ activates a potential-dependent cation channel in the luminal membrane and permits increased Na⁺ absorption (Lang & Martens, 1999). The observations of Stacy & Warner (1966) can most probably be explained by other mechanisms and are not related to the increased osmotic pressure.

Mucosal hyperosmolarity causes an increase of J_smNa in tissues of HF sheep in a dose-dependent manner. Because we have previously shown that the change of J_smNa is positively and linearly correlated with G_t, the paracellular pathway has been suggested to become more ‘leaky’ upon hyperosmotic challenge. This suggestion has been verified by the determination of the conductance of the paracellular pathway and of flux rates of the paracellular marker ⁵¹Cr-EDTA (Schweigel et al. 2005). Both parameters are increased upon osmotic challenge, in agreement with results from previous studies of a variety of tight or moderately tight tissues (Ussing, 1965; Civan & DiBona, 1974; Soybel et al. 1987). The rumen epithelium is a keratinized multilayered squamous epithelium, so the histological counterpart of the paracellular pathway is more complicated than the corresponding structure (tight junctions) in epithelia of the gut (Graham & Simmons, 2004). Studies by Gemmel & Stacy (1973) have shown that a stepwise increase of luminal osmotic pressure finally causes a drop of PD_t. Histological examination of the epithelium has revealed that the osmotically induced breakdown of the stratum granulosum accompanies the change of PD_t, indicating an increase of the permeability barrier in this layer of the epithelium. These histological findings and the drop of PD_t are in an good agreement with the changes of G_t and J_smNa of the present study.

Adaptation

An increase of energy intake causes well-established changes of form and functions of the rumen epithelium: increase of size and number of papillae (Dirksen et al. 1984) by reduction of apoptosis (Mentschel et al. 2001), increase of transport rates of SCFA (Dirksen et al. 1984; Gäbel et al. 1991,), of HCO₃^– (Gäbel et al. 1991,), of Na⁺ and Ca²⁺ (Uppal et al. 2003a,b) and increase of activities of enzymes (Nocek et al. 1980). Luminal factors such as propionic and butyric acid induce these changes (Kauffold et al. 1977; Mentschel et al. 2001), and insulin-like growth factor (IGF-1) may possibly also be involved (Shen et al. 2004).

The results of the present study demonstrate a decrease of sensitivity in response to a mucosal challenge of hypertonic osmotic pressure. Several reasons can be proposed to explain the modified strength against hyperROP, as follows: (a) the thickness of the cornified layer of the epithelium is increased at higher energy intake (Liebich et al. 1987), and this layer may not be in equilibrium with the bulk solution, i.e. the epithelium is not directly exposed to hyperROP; (b) the transport of electrolytes is enhanced and could dilute the buffer solution in the cornified layer just above the luminal membrane, which again prevents direct exposure of the epithelium to hyperROP (indeed, the absorbed fluid from the rumen is hypertonic; H. Martens, unpublished observation); and (c) evidence exists that the epithelia of sheep fed on energy-rich diets appear to be more resistant to mechanical stress (unproven). However, the proposed explanations have not been established experimentally and are beyond the scope of the present study.

Secretion of Na⁺ at a potential difference of +25 mV

Net Na⁺ transport rates under voltage clamp conditions decrease in the ranking order –25 > 0 > +25 mV (‘+’ or ‘–’ refers to the voltage at the serosal side). In tissues of HF sheep, a PD_t of +25 mV causes a net Na⁺ transport in the serosal to mucosal direction (secretion) at a mucosal osmotic pressure of 375 and 450 mosmol l^–1, which is in agreement with results from Sehested et al. (1996). However, tissues of SF sheep always exhibit a net absorption at +25 mV; this is positively correlated with increasing intake of SD.

In vivo studies with isolated rumen have shown that there is always a net absorption of Na⁺ in HF and SF sheep, even with an elevated osmotic pressure (Gäbel et al. 1987a,b). This discrepancy between in vivo studies (washed rumen) and in vitro studies can be explained by: (a) increased paracellular permeability attributable to the manipulation (e.g. ‘stripping’) of the epithelium (in vitro); and (b) the hyperosmolarity of the artificial rumen fluid being rapidly diminished because inflow of water is induced and osmolytes, such as SCFA, are absorbed (H. Martens, unpublished observation). This has to be taken into account when extrapolating our data to the in vivo situation.

Conclusions

Diet-induced alterations of transport mechanisms, enzyme activities and absorptive surface area are accompanied by mechanisms that protect the tissue against luminal hyperosmolarity. The underlying mechanisms of this adaptation are unknown. This adaptation maintains an undisturbed barrier function at hyperROP and has not been previously described. Furthermore, an increase of SD intake (1200 and 1800 g day^–1) diminishes the effect of hyperROP on J_netNa and abolishes changes of J_smNa as a parameter for passive and paracellular permeability. Because the barrier function is not influenced by hyperROP at these SD intakes, transepithelial and passive passage of endotoxins and bacteria, which may result in inflammation of the rumen wall or even abscesses in the epithelium and liver, are prevented. The increase of ROP in vivo is mainly caused by SCFA. Indeed, high concentrations of SCFA and low pH impair the function of the rumen epithelium under conditions of ruminal acidosis (Dirksen, 1985). Adaptation of the rumen epithelium is speculated to help to reduce the risk of lesion of the rumen epithelium by hyperROP and low pH.

	References

Top Abstract Introduction Methods Results Discussion References

 
Abdoun K, Wolf K, Arndt G & Martens H (2003). Effect of ammonia on Na⁺ transport across isolated rumen epithelium of sheep is diet dependent. Br J Nutr 90, 751–758.[CrossRef][Medline]

Abdoun K, Wolf K, Stumpff F & Martens H (2005). Modulation of electroneutral Na transport in sheep rumen epithelium by luminal ammonia. Am J Physiol Gastrointest Liver Physiol 289, G508–G520.[Abstract/Free Full Text]

Allen MS (1997). Relationship between fermentation acid production in the rumen and the requirement for physically effective fiber. J Dairy Sci 80, 1447–1462.[Abstract]

Bennink MR, Tyler TR, Ward GM & Johnson DE (1978). Ionic milieu of bovine and ovine rumen as affected by diet. J Dairy Sci 61, 315–323.

Bindslev N, Tormey JM & Wright EM (1974). The effects of electrical and osmotic gradients on lateral intercellular spaces and membrane conductance in a low resistance epithelium. J Membr Biol 19, 357–380.[CrossRef][Medline]

Bugaut M (1987). Occurrence, absorption and metabolism of short chain fatty acids in the digestive tract of mammals. Comp Biochem Physiol B 86, 439–472.[CrossRef][Medline]

Burgos MS, Langhans W & Senn M (2000). Role of rumen fluid hypertonicity in the dehydration-induced hyophagia in cows. Physiol Behav 71, 423–430.[CrossRef][Medline]

Carter RR & Grovum WL (1990). Factors affecting the voluntary intake of food by sheep. 5. The inhibitory effect of hypertonicity in the rumen. Br J Nutr 64, 285–299.[CrossRef][Medline]

Chien WJ & Stevens CE (1972). Coupled active transport of Na and Cl across forestomach epithelium. Am J Physiol 223, 997–1003.[Free Full Text]

Civan MM & DiBona D (1974). Pathways for movement of ions and water across toad urinary bladder II. Site and mode of action of vasopressin. J Membr Biol 19, 195–220.[CrossRef][Medline]

DiBona DR (1972). Passive intercellular pathway in amphibian epithelia. Nature 238, 179–181.

DiBona DR & Civan MM (1973). Pathways for movement of ions and water across toad urinary bladder. I. Anatomic site of transepithelial shunt pathways. J Membr Biol 12, 101–128.[Medline]

Dirksen G (1985). The rumen acidosis complex – recent knowledge and experiences (1). [A review in German] Tierärztl Prax 13, 501–512.[Medline]

Dirksen G, Liebich HG, Brosi G, Hagemeister H & Mayer E (1984). Morphologie der Pansenschleimhaut und Fettsäureresorption beim Rind – bedeutende Faktoren für Gesundheit und Leistung. Zbl Vet Med A 31, 414–430.

Dobson A (1959). Active transport through the epithelium of the reticulo-rumen sac. J Physiol 146, 235–251.[Free Full Text]

Dobson A, Sellers AF & Gatewood VH (1976). Dependence of Cr-EDTA absorption from the rumen on luminal osmotic pressure. Am J Physiol 231, 1595–1600.[Abstract/Free Full Text]

Donowitz M & Tse CM (2001). Molecular physiology of mammalian Na⁺/H⁺ exchangers NHE2 and NHE3. Curr Topics Membr 50, 437–498.

Doreau M, Ferchal E & Beckers Y (1997). Effects of level of intake and of available volatile fatty acids on the absorptive capacity of sheep rumen. Small Ruminant Res 25, 99–105.

Engelhardt WV (1969). Der osmotische Druck im Panseninhalt. Zbl Vet Med A 16, 665–690.

Erlij D & Martinez-Palomo A (1972). Opening of tight junctions in frog skin by hypertonic urea solutions. J Membr Biol 9, 229–240.[CrossRef][Medline]

Ferreira HG, Harrison FA, Keynes RD & Zurich L (1972). Ion transport across an isolated preparation of sheep rumen epithelium. J Physiol 222, 77–93.[Abstract/Free Full Text]

Freyer M & Martens H (1998). In vitro Untersuchungen zur Widerstandsanalyse des Pansenepithels von Schafen in Abhängigkeit vom luminalen osmotischen Druck. Proc Soc Nutr Physiol 8, 80.

Frizzell RA & Schultz SG (1972). Ionic conductances of extracellular shunt pathway in rabbit ileum. Influence of shunt on transmural sodium transport and electrical potential differences. J Genl Physiol 59, 318–346.

Gäbel G (1988). Natrium- und Chloridtransport im Pansen von Schafen: Mechanismen und ihre Beeinflussung durch intraruminale Fermentationsprodukte. Habilitationsschrift TiHo, Hannover.

Gäbel G, Bestmann M & Martens H (1991). Influences of diet, short-chain fatty acids, lactate and chloride on bicarbonate movement across the reticulo-rumen wall of sheep. J Vet Med A 38, 523–529.

Gäbel G, Martens H, Suendermann M & Galfi P (1987a). The effect of diet, intraruminal pH and osmolarity on sodium, chloride and magnesium absorption from the temporarily isolated and washed reticulo-rumen of sheep. Q J Exp Physiol 72, 501–511.[Abstract/Free Full Text]

Gäbel G, Suendermann M & Martens H (1987b). The influence of osmotic pressure, lactic acid and pH on ion and fluid absorption from the washed and temporarily isolated reticulo-rumen of sheep. J Vet Med A 34, 220–226.

Gäbel G, Vogler S & Martens H (1991). Short-chain fatty acids and CO₂ as regulators of Na⁺ and Cl^– absorption in isolated sheep rumen mucosa. J Comp Physiol B 161, 419–426.[Medline]

Gemmel RT & Stacy BD (1973). Effects of ruminal hyperosmolality on the ultrastructure of ruminal epithelium and their relevance to sodium transport. Q J Exp Physiol 58, 315–323.[Abstract/Free Full Text]

Gorodeski GI, De Santis BJ, Goldfarb J, Utian WH & Hopfer U (1995). Osmolar changes regulate the paracellular permeabiltiy of cultured human cervical epithelium. Am J Physiol 269, C870–C877.[Medline]

Graham C & Simmons NL (2005). Functional organization of the bovine rumen epithelium. Am J Physiol 288, R173–R181.

Jackson J & Norris SH (1985). Transport of sodium and chloride across rat gastric mucosa in vitro. J Physiol 360, 293–310.

Kapus A, Grinstein S, Wasan S, Kandasamy R & Orlowski J (1994). Functional characterization of three isoforms of the Na⁺/H⁺ exchanger stably expressed in Chinese hamster ovary cells. ATP dependence, osmotic sensitivity and role in cell proliferation. J Biol Chem 269, 23544–23552.[Abstract/Free Full Text]

Kauffold P, Voigt J & Herrendörfer G (1977). Untersuchungen über den Einfluss von Ernährungsfaktoren auf die Pansenschleimhaut. Arch Anim Nutr 27, 2001–2211.

Lang F, Busch GL, Ritter M, Völkl H, Waldegger S, Gulbins E & Häussinger D (1998). Functional significance of cell volume regulatory mechanisms. Physiol Rev 78, 247–306.[Abstract/Free Full Text]

Lang I & Martens H (1999). Na transport in sheep rumen is modulated by voltage-dependent cation conductance in apical membrane. Am J Physiol 277, G609–G618.[Medline]

Liebich HG, Dirksen G, Arbel A, Dori S & Mayer E (1987). Fütterungsabhängige Veränderungen der Pansenschleimhaut von Hochleistungskühen im Zeitraum von der Trockenstellung bis acht Wochen post partum. J Vet Med A 34, 661–672.

Lopez S, Hovell FDD & MacLeod NA (1994). Osmotic pressure, water kinetics and volatile fatty acid absorption in the rumen of sheep sustained by intragastric infusions. Br J Nutr 71, 153–168.[CrossRef][Medline]

Martens H & Blume I (1987). Studies on the absorption of sodium and chloride from the rumen of sheep. Comp Biochem Physiol A 86, 653–656.

Martens H & Gäbel G (1988). Transport of Na and Cl across the epihelium of ruminant forestomachs: rumen and omasum. A review. Comp Biochem Physiol A 90, 569–575.

Martens H, Gabel G & Strozyk H (1987). The effect of potassium and the transmural potential difference on magnesium transport across an isolated preparation of sheep rumen epithelium. Q J Exp Physiol 72, 181–188.[Abstract/Free Full Text]

Martens H, Gäbel G & Strozyk B (1991). Mechanism of electrically silent Na and Cl transport across the rumen epithelium of sheep. Exp Physiol 76, 103–114.[Abstract]

Martens H, Harmeyer J & Michael H (1978). Magnesium transport by isolated rumen epithelium of sheep. Res Vet Sci 24, 161–168.[Medline]

Mentschel J, Leiser R, Mülling C, Pfarrer C & Claus R (2001). Butyric acid stimulates rumen mucosa development in the calf mainly by a reduction of apoptosis. Arch Anim Nutr 55, 85–102.

Nath SK, Yue Hang C, Levine SA, Chris Yun CH, Montrose MH, Donowitz M & Tse CM (1996). Hyperosmolarity inhibits the Na⁺/H⁺ exchanger isoforms NHE2 and NHE3: an effect opposite to that on NHE1. Am J Physiol 270, G431–G441.[Medline]

Nocek JE, Herbein JH & Polan CE (1980). Influence of ration physical form, ruminal degradable nitrogen and age on rumen epithelial and some enzymatic activities. J Nutr 110, 2355–2364.[Abstract/Free Full Text]

Oshio S & Tahata I (1984). Absorption of dissociated volatile fatty acids through the rumen wall of sheep. Can J Anim Sci supplement 64, 167–168.

Powell DW (1981). Barrier function of epithelia. Am J Physiol 241, G275–G288.[Medline]

Reuss L & Finn AL (1977). Effects of luminal hyperosmolality on electrical pathways of Necturus gallbladder. Am J Physiol 232, C99–C108.[Medline]

Schweigel M, Freyer M, Leclercq S, Etschmann B, Lodemann U, Böttcher A & Martens H (2005). Luminal hyperosmolarity decreases Na transport and impairs barrier function of sheep rumen epithelium. J Comp Physiol B 174, 575–591.

Sehested J, Diernaes L, Moller PD & Skadhauge E (1996). Transport of sodium across the isolated bovine rumen epithelium: interaction with short-chain fatty acids, chloride and bicarbonate. Exp Physiol 81, 79–94.[Abstract]

Shen Z, Seyfert HM, Löhrke B, Schneider F, itnan R, Chudy A, Kuhla S, Hammon HM, Blum JW, Martens H, Hagemeister H & Voigt J (2004). Energy rich diet causes rumen papillae proliferation associated with increased IGF-1 levels and IGF type 1 receptors in young goats. J Nutr 134, 11–17.[Abstract/Free Full Text]

Soleimani M, Bookstein C, McAteer JA, Hattabaugh YJ, Bizal GL, Musch MW, Villereal M, Rao MC, Howard RL & Chang EB (1994). Effect of high osmolality on Na⁺/H⁺ exchange in renal proximal tubule cells. J Biol Chem 269, 15613–15618.[Abstract/Free Full Text]

Soybel DI, Ashley SW, DeSchryver-Kecskemeti K & Cheung LY (1987). Effects of luminal hyperosmolality on cellular and paracellular ion transport pathways in Necturus antrum. Gastroenterology 93, 456–465.[Medline]

Stacy BD & Warner ACI (1966). Balances of water and sodium in the rumen during feeding: osmotic stimulation of sodium absorption in the sheep. Q J Exp Physiol 51, 79–93.[Abstract/Free Full Text]

Tabaru H, Ikeda K, Kadota E, Murakami Y, Yamada H, Sasaki N & Takeuchi A (1990). Effects of osmolality on water, electrolytes and VFAs absorption from the isolated ruminoreticulum in the cow. Jpn J Vet Sci 52, 91–96.

Uppal SK, Wolf K, Khahra SS & Martens H (2003a). Modulation of Na⁺ transport across isolated rumen epithelium by short-chain fatty acids in hay- and concentrate-fed sheep. J Anim Physiol Anim Nutr 87, 380–388.[Medline]

Uppal SK, Wolf K & Martens H (2003b). The effect of short chain fatty acids on calcium flux rates across isolated rumen epithelium of hay-fed and concentrate-fed sheep. J Anim Physiol Anim Nutr 87, 12–20.[Medline]

Ussing HH (1965). Relationship between osmotic reactions and active sodium transport in the frog skin epithelium. Acta Physiol Scand 63, 141–155.[Medline]

Warner ACI & Stacy BD (1965). Solutes in the rumen of the sheep. Q J Exp Physiol 50, 169–184.[Abstract/Free Full Text]

Warner ACI & Stacy BD (1972). Water, sodium and potassium movements across the rumen wall of sheep. Q J Exp Physiol 57, 103–119.[Abstract/Free Full Text]

Warner ACI & Stacy BD (1977). Influence of ruminal and plasma osmotic pressure on salivary secretion in sheep. Q J Exp Physiol 62, 133–142.[Abstract/Free Full Text]

Watts BA III & Good DW (1994). Apical membrane Na⁺/H⁺ exchange in rat medullary thick ascending limb. pH-dependence and inhibition by hyperosmolality. J Biol Chem 269, 20250–20255.[Abstract/Free Full Text]

Weinman EJ, Shenolikar S & Kahn AM (1987). cAMP-associated inhibition of Na⁺-H⁺ exchanger in rabbit brush-border membranes. Am J Physiol 252, F19–F25.[Medline]

Welch JG (1979). Rumination and particle size, rumen osmotic pressure and feed intake in ruminants. Proceedings of the Cornell Nutrition Conference, pp. 49–51.