Non-technical summary
Nerve-mediated influences on gastrointestinal motility in response to orexin A, either centrally injected or applied to isolated gut preparations, have been reported. However, the presence of orexin receptors at the gastrointestinal smooth muscle level has been found. On these grounds, in the present study we evaluated whether orexin A also exerts direct muscular effects in the duodenal smooth muscle of the mouse in an attempt to explain the possible mechanism of action involved. The experimental results from mechanical and electrophysiological studies indicate that orexin A causes direct contractile responses in the isolated preparations and evokes changes in the ionic currents of the smooth muscle cells. Thus, orexin A, in addition to its neutrally mediated influences on gastrointestinal motility, exerts direct muscular effects on the mouse duodenum. This latter mechanism, from a physiological point of view, may act in a synergic manner to reinforce the neuronal signals.
Abstract
Abstract
Orexin A (OXA) has been reported to influence gastrointestinal motility, acting at both central and peripheral neural levels. The aim of the present study was to evaluate whether OXA also exerts direct effects on the duodenal smooth muscle. The possible mechanism of action involved was investigated by employing a combined mechanical and electrophysiological approach. Duodenal segments were mounted in organ baths for isometric recording of the mechanical activity. Ionic channel activity was recorded in current- and voltage-clamp conditions by a single microelectrode inserted in a duodenal longitudinal muscle cell. In the duodenal preparations, OXA (0.3 μm) caused a TTX-insensitive transient contraction. Nifedipine (1 μm), as well as 2-aminoethyl diphenyl borate (10 μm), reduced the amplitude and shortened the duration of the response to OXA, which was abolished by Ni2+ (50 μm) or TEA (1 mm). Electrophysiological studies in current-clamp conditions showed that OXA caused an early depolarization, which paralleled in time the contractile response, followed by a long-lasting depolarization. Such a depolarization was triggered by activation of receptor-operated Ca2+ channels and enhanced by activation of T- and L-type Ca2+ channels and store-operated Ca2+ channels and by inhibition of K+ channels. Experiments in voltage-clamp conditions demonstrated that OXA affects not only receptor-operated Ca2+ channels, but also the maximal conductance and kinetics of activation and inactivation of Na+, T- and L-type Ca2+ voltage-gated channels. The results demonstrate, for the first time, that OXA exerts direct excitatory effects on the mouse duodenal smooth muscle. Finally, this work demonstrates new findings related to the expression and kinetics of the voltage-gated channel types, as well as store-operated Ca2+ channels.
Introduction
Orexin A (OXA) and orexin B (OXB) were first described as neuropeptides expressed by a specific population of neurons in the lateral hypothalamic area (Sakurai et al. 1998), a region classically implicated in feeding behaviour. However, orexin nerve fibres have been widely identified throughout the central nervous system (Date et al. 1999), accounting for the involvement of these peptides in many different physiological functions, including regulation of the sleep–wake cycle, energy homeostasis and cardiovascular functions (see Kukkonen et al. 2002; Adamantidis & de Lecea, 2009).
Like the widespread orexigenic fibres, orexin receptors also are widely distributed in the central nervous system (Okumura & Takakusaki, 2008). Actions of OXA and OXB are mediated via binding to closely related G-protein-coupled receptors (Sakurai et al. 1998), termed the orexin-1 and orexin-2 receptors (OX1R and OX2R). Orexon A has equal affinity at OX1Rs and OX2Rs, whereas OXB has an appreciably greater affinity at OX2Rs (Sakurai et al. 1998).
Orexins have been reported to affect gastrointestinal motility, and most of the investigations in this area have been focused on the effects of OXA, which appears to be more potent for inducing feeding behaviour and gastric secretion than OXB (Edwards et al. 1999; Kunii et al. 1999; Takahashi et al. 1999). Experiments using central injection of OXA have shown that this peptide influences gastrointestinal motor responses (Kobashi et al. 2002; Krowicki et al. 2002; Baccari, 2010; Bülbül et al. 2010). However, orexins and their receptors are not only present in the central nervous system, but they are abundantly distributed in the gastrointestinal tract of different species, including humans (Näslund et al. 2002; Nakabayashi et al. 2003; Ehrstrom et al. 2005), suggesting that these peptides may also exert local effects.
In particular, the presence of orexins and their receptors has been revealed in the enteric nervous system (myenteric and submucosal plexuses), as well as in mucosa and smooth muscle layers throughout the gastrointestinal tract of mammals (De Miguel & Burrell, 2002; Näslund et al. 2002; Dall'Aglio et al. 2008), supporting the local influence of these peptides in several functions, including motility.
Experiments carried out on isolated gastrointestinal preparations have shown that orexins exert both relaxant and contractile effects (Korczynki et al. 2006b), mainly acting at the neural level to activate inhibitory and excitatory enteric neurons.
In addition to the neutrally mediated effects, direct smooth muscle contractions in response to OXA have been observed in rat jejunum segments (Korczynski et al. 2006a), but the myogenic mechanism of action involved has not been elucidated.
It is well known that orexin receptors induce Ca2+ elevations both via receptor-operated Ca2+ channels (ROCs) and via the ‘conventional’ phospholipase C, Ca2+-release InsP3 channels, store-operated Ca2+ channel (SOC) pathways. Studies performed in Chinese hamster ovary cells suggest that OXA-induced Ca2+ transients originate from these two paths, depending on the ligand concentration. At low OXA concentrations (≤10 nm), the primary Ca2+ influx seems to be due to the opening of ROCs. At higher OXA concentrations, it is suggested that the increase of [Ca2+]i may be the result of Ca2+ released from the endoplasmic reticulum, as well as of an influx through SOCs (Kukkonen & Åerman, 2001; Larsson et al. 2005).
Orexin A-induced Ca2+ transients also depend on the cell type. In Chinese hamster ovary cells and recombinant neuron-like cells, no evidence was found for the involvement of voltage-gated Ca2+ channels (VGCCs; Holmqvist et al. 2002). In contrast, in rat neurons orexin-stimulated Ca2+ influx has been suggested to be related to VGCC activation. In neurons, the OXA-induced Ca2+ elevation was suggested to be due to activation of orexin receptors that would activate protein kinase C, which in turn would phosphorylate and thereby activate VGCCs, thus resulting in the following activation sequence: orexin receptor, phospholipase C, protein kinase C and N-/L-type VGCC-mediated influx of Ca2+ (Uramura et al. 2001; Kukkonen et al. 2002).
Finally, K+ channels may also be involved, because in neurons, the sustained depolarization observed following OXA stimulation was related to inhibition of K+ channels (Hwang et al. 2001; Kukkonen et al. 2002; Grabauskas & Moises, 2003).
The present study was designed to investigate whether OXA exerts direct effects on the duodenal smooth muscle and to investigate the mechanism of action underlying these responses. For this purpose, mechanical and electrophysiological studies were performed on mouse duodenal preparations.
Methods
Ethical approval
The experimental protocol was designed in compliance with the Principles of Laboratory Animal Care (NIH publication 86-23, revised 1985) and the recommendations of the European Economic Community (86/609/CEE).
Animals
Experiments were carried out on 20 albino female mice of the Swiss strain, 8–12 weeks old (Morini, Reggio Emilia, Italy). The mice were fed standard laboratory chow and water, and were housed under a 12 h–12 h light–dark photoperiod and controlled temperature (21 ± 1°C). The mice were killed by cervical dislocation. The abdomen was immediately opened, and segments of duodenum, immediately distal to the pylorus, were removed.
Mechanical studies
The contents of the excised segments were gently flushed out with Krebs–Henseleit solution. Segments (20 mm in length) were suspended in 5 ml double-jacketed organ baths containing Krebs–Henseleit solution (gassed with 95% O2–5% CO2) of the following composition (mm): NaCl, 118; KCl, 4.7; MgSO4, 1.2; KH2PO4, 1.2; NaHCO3, 25; CaCl2, 2.5; and glucose, 10 (pH 7.4). Prewarmed water (37°C) was circulated through the outer jacket of the tissue bath via a constant-temperature circulator pump. The temperature of the Krebs–Henseleit solution in the organ bath was maintained within a range of 37 ± 0.5°C.
One end of each preparation was tied to a platinum rod, while the other was connected to a force displacement transducer (Grass, Quincy, MA, USA FT03) by a silk thread for continuous recording of isometric tension. The transducer was coupled to a polygraph (Sanborn, Walthamanm, MA, USA model 7700).
Duodenal preparations were allowed to equilibrate for 30 min under an initial load of 200 mg. During this period, repeated and prolonged washes of the preparations with Krebs–Henseleit solution were done to avoid accumulation of metabolites in the organ baths.
Drugs
The following drugs were used: OXA, TTX, nifedipine, 2-aminoethyl diphenyl borate (2-APB), TEA and Ni2+. All drugs were obtained from Sigma-Aldrich (St Louis, MO, USA). Solutions were prepared on the day of the experiment, except for TTX, for which a stock solution was stored at −20°C. Drugs concentrations are given as final bath concentrations.
Experimental protocol
All recordings were obtained in the presence of 1 μm TTX.
In a first series of experiments, control responses to 0.3 μm OXA were evoked. The dose employed was that reported to give the maximal contraction in the concentration–response curves and to be effective in the gastrointestinal preparations (Satoh et al. 2001; Matsuo et al. 2002; Krowinski et al. 2006b; Baccari & Calamai, 2008; Baccari et al. 2009). The interval between two subsequent applications of OXA was not less than 30 min, during which repeated and prolonged washes of the preparations with Krebs–Henseleit solution were done to avoid desensitization phenomena (Satoh et al. 2001). In order to ascertain the possible involvement of L-type or T-type Ca2+ channels in the response to OXA, nifedipine was used to selectively block L-type Ca2+ channels or Ni2+ to block T-type Ca2+ channels (Hollywood et al. 2003). For this purpose, in a series of experiments, OXA (0.3 μm) was added to the bath medium 10 min following nifedipine (1 μm) or Ni2+ (50 μm).
In the final series of experiments, to focus on the role of K+ channels and SOCs activated by OXA, we used pharmacological inhibitors. Therefore, the response to 0.3 μm OXA was elicited in the presence of TEA (1–5 mm) or 2-APB (10 μm) about 10 min after their addition to the bath medium.
Data analysis and statistical tests
The amplitude of contractile responses to OXA is expressed in absolute values (grams). Statistical analysis was performed by means of Student's paired t test. Values were considered significantly different at P ≤ 0.05. Results are given as means ± SEM. The number of muscle preparations is designated by n in the Results.
Electrophysiological studies
In parallel experiments, the same duodenal preparations as above were used for electrophysiological recording. They were opened along the antemesenterial axis and pinned, serosal or mucosal side up, to a transparent Sylgard (Dow Corning, Midland, MI, USA) floor of a dissecting dish filled with Krebs solution. First, with the mucosal side up and the aid of a dissecting microscope, the mucosa and submucosa were carefully dissected away, taking care not to damage the inner layer of circular muscle. The remaining tissue was repinned serosal side up, and the connective tissue was removed in order to expose the duodenal longitudinal smooth muscle (DLM). Two or three muscle strips (6 mm × 20 mm) were dissected and repinned, serosal side up, at the resting length to the floor of a Sylgard-coated recording chamber, and a glass microelectrode was inserted into a cell of the DLM layer.
During the electrophysiological experiments, the tissue was superfused at a rate of 1.8 ml min−1 with a control bath solution of the same composition as the Krebs–Henseleit solution used in mechanical experiments. In order to block outward K+ currents, we used a low-TEA solution (mm): NaCl, 122.5; CaCl2, 2; TEA-OH, 10; and Hepes, 10. To evaluate the presence of Na+ current, we used a choline-containing low-TEA solution that had the same composition as the low-TEA solution except that NaCl was replaced by choline chloride. To record only Ca2+ currents, we used a high-TEA solution (Na+ and K+ free; mm): CaCl2, 10; TEA-Br, 145; and Hepes, 10. This latter high-TEA solution was suitable to block not only any Na+ and K+ currents (Morelli et al. 2008), but also Ca2+ entry through ROCs (Johansson et al. 2007) and transient receptor potential channel (McFadzean & Gibson, 2002; Larsson et al. 2005). Conventional microelectrodes were filled with the internal solution containing (mm): KCl, 130; NaH2PO4, 10; CaCl2, 0.2; EGTA, 1; MgATP, 5; and Hepes, 10. Microelectrodes were pulled from borosilicate glass (GC 150-15; Clark, Reading, UK) using a micropipette vertical puller (Narishige PC-10; Kyoto, Japan). The pH was titrated to 7.4 with NaOH and to 7.2 with TEA-OH for bath and pipette solution, respectively.
We used high-resistance (60–70 MΩ) conventional microelectrodes. The electrode capacitive currents were compensated; series resistance was determined and minimized in current clamp, whereas 80–85% of compensation was made in voltage clamp. The tip potentials were often rather high, and 70–80% of a batch of microelectrodes had to be discarded because of the excessive tip potential. Only electrodes with tip potential of less than –8 mV were used. Tip potential was compensated, but during recording in current clamp about 30% of the investigated cells showed a voltage run-down 5–20 min after insertion into the cell. Cell recordings showing a run-down over 5% were discarded. The following inhibitors were used: TTX (1 μm) for INa; 2-APB (10 μm) for SOCs; nifedipine (1–10 μm) for L-type Ca2+ current (ICa,L); and Ni2+ (5–50 μm) for T-type Ca2+ current (ICa,T; Ward & Sanders, 1992; Hollywood et al. 2003). Thapsigargin (Tg; 1 μm) was used to deplete the Ca2+ stores by inhibiting the sarco(endo)plasmic reticulum Ca2+-ATPase (Morales et al., 2004). Heptanol (1 mm) was added to all solutions in order to block gap junctional currents and to enable recording of ionic currents elicited only from the impaled cell (Squecco et al. 2006; Formigli et al. 2009a).
The microelectrode was connected to a micromanipulator (Narishige, Kyoto, Japan) and an Axopatch 200B amplifier (Axon Instruments, Union City, CA, USA). Current- and voltage-clamp stimulating protocols and data acquisition were controlled by using an output and an input of the A/D–D/A interfaces (Digidata 1200; Axon Instruments) and pCLAMP 9 software (Axon Instruments).
Values of resting membrane potential (RMP) for all the cells tested were determined in current-clamp conditions immediately after the cell impalement and upon withdrawal of the microelectrode at the end of the experiment. Pulse protocols of stimulation were applied in current-clamp conditions to evaluate the changes in membrane potential due to addition of OXA to the bath solution and the action of chemicals on such responses; in addition, the voltage-clamp mode was used to evaluate the actions of OXA on voltage-gated channels mainly selective for Na+, K+ and Ca2+ currents. These latter sets of experiments were performed in a time interval from 6 to 20 min after the addition of OXA (3.10−7m) to the bath solution (see Results).
All the passive properties parameters were estimated in the same way as by Formigli et al. (2005). To allow comparison of test current recorded from different cells, membrane current amplitude was normalized to cell linear capacitance, Cm (in picoamperes per picofarad), because Cm is an index of cell surface area, assuming that membrane-specific capacitance is constant at 1 μF cm−2.
The occurrence of the SOC was further assessed in voltage-clamp mode by measuring the current amplitudes at a potential of –50 and 50 mV, taken from currents in response to voltage ramps ranging from –120 to 50 mV over a period of 0.5 s, which were imposed every 1 min from a holding potential of 0 mV and digitized at a rate of 5 kHz; two runs repeated every 20 s were averaged. For analysis, the average of two ramps elicited in high-TEA solution, to block ROC, was used for leak subtraction for the subsequent current records after adding Tg and Tg + OXA. Seven minutes after Tg addition to the bath solution, OXA was added to the bath solution too, and a voltage ramp was applied for a further 8 min.
To analyse the activation of Na+ and Ca2+ currents, voltage pulses from –70 to 50 mV were applied to voltage-clamped cells held at –80 mV, in 10 mV increments. For the fast Na+ current activation the pulse duration was 10 ms, whereas for the slower Ca2+ current it was 4 s; the protocol used an interval of 1 or 20 s between stimulating episodes for recovery. The steady-state inactivation was studied by a two-pulse protocol, with a 10 ms or 1 s prepulse to different voltages followed by a 10 ms or 1 s test pulse fixed to 0 mV after 10 or 200 ms. The smaller pulse and interval durations were used for Na+ current recording.
The interpulse intervals to the holding potential were chosen both to prevent substantial recovery from inactivation between activating pulses and to allow the activation kinetics of Ca2+ permeability to return to its resting state. Again, in the two-pulse protocol, we used an interval of 1 or 20 s between stimulating episodes for recovery. All the activation and inactivation protocols were repeated twice. The steady-state ionic current of activation (Ia) was evaluated by Ia(V) = Gmax(V–Vr)/{1 + exp[(Va–V)/ka]} and steady-state inactivation by Ih(V) = I/{1 + exp[–(Vh–V)/kh]}, where Gmax is the maximal conductance for the Ia, Vr is the apparent reversal potential, Va and Vh are the potentials eliciting the half-maximal activation and inactivation values, respectively, and ka and kh are the steepness factors.
The properties of K+ current were studied by applying 1 s voltage pulses ranging from −70 to +50 mV, starting from a holding potential (HP) of –40 mV.
The P/4 subpulse correction of cell leakage and capacitance was used to study Na+, Ca2+ and K+ currents. This procedure also minimized voltage-independent currents, such as those flowing through intermediate-conductance Ca2+-activated K+ channels and stretch-activated channels (SACs; Formigli et al. 2009a,b;).
Mathematical and statistical analysis of data was performed by pCLAMP 9 (Axon Instruments), SigmaPlot and SigmaStat (Jandel Scientific, San Rafael, CA, USA). Experiments were done at room temperature (22–25°C). Data are expressed as means ± SEM. One-way ANOVA with repeated measures was used for multiple comparisons and a value of P < 0.05 was considered significant.
Results
Mechanical responses
Duodenal preparations exhibited spontaneous mechanical activity consisting of rhythmic changes in isometric tension. Addition of OXA (0.3 μm) to the bath medium (n = 24) caused a transient contractile response (mean amplitude, 248.3 ± 8 mg). The contractile response to OXA began to decay after 30 ± 5 s of contact time, and the tension of the preparations returned to the basal level within 1.5–2 min from the addition of the peptide to the bath medium (Fig. 1). After 5–10 min from the addition of OXA to the bath medium, without washing, the preparation no longer responded to a subsequent application of OXA (0.3 μm). The response to OXA was fully regained after 30 min, during which repeated and prolonged washes of the preparations with Krebs–Henseleit solution were performed.
Figure 1. Effects of nifedipine and 2-aminoethyl diphenyl borate (2-APB) on the mechanical response to orexin A (OXA) in duodenal preparations.
Typical traces showing the transient contractile response elicited by the addition of OXA (0.3 μm) to the bath medium (A and B, left traces). In the presence of 1 μm nifedipine (A, right trace) or 10 μm 2-APB (B, right trace), the contractile response to OXA was reduced in amplitude and shortened in duration.
The contractile response to 0.3 μm OXA (n = 6) evoked in the presence of 1 μm nifedipine was reduced in amplitude (162.2 ± 6 mg; P < 0.05) and shortened in duration (58 ± 4 s; P < 0.05; Fig. 1). Orexin A (0.3 μm) in the presence of 50 μm Ni2+ (n = 4) no longer had any effect (data not shown). Likewise, the contractile response to OXA evoked following the addition of 1 mm TEA (n = 4) to the bath medium was abolished (data not shown). A higher Ni2+ concentration, such as that used to block SOCs (5 mm; Kukkonen & Åerman, 2001; Larsson et al. 2005) or a higher TEA concentration, such as that used to block ROCs (70 mm; Larsson et al. 2005; Johansson et al. 2007) could not be tested.
The response to OXA elicited in the presence of 10 μm 2-APB (n = 6) was greatly reduced in amplitude (80.8 ± 5.5 mg; P < 0.05) and also shortened in duration (50 ± 6 s; P < 0.05; Fig. 1).
To assess the mechanical responses of DLM cells caused by exposure to OXA further, we used electrophysiological methods to evaluate the kinds of voltage-gated ionic channels affected by OXA.
Rapid and slow membrane depolarization induced by OXA in current-clamp experiments
Initially, we carried out experiments in current-clamp conditions using the control bath solution. The mean RMP recorded was –57 ± 5 mV (44 cells; 12 mice). None of the muscle cells tested exhibited spontaneous electrical activity as observed in muscle cell preparations of isolated DLM (Hara et al. 1986).
Addition of OXA to the control solution induced a depolarizing response after 20–30 s. Its time course was similar to that of the contractile response, because it reached a maximum after about 1 min, followed by a progressive slow decrease. After about 5 min the response reached a quasi-steady-state level, during which only small (1–2 mV) and slow oscillations (every 5–8 min) were recorded (Fig. 2A, Con). The peak (Vp) and steady-state depolarization values (Vss) with respect to the RMP are reported in Fig. 2B. Nifedipine (1 μm), Ni2+ (50 μm) and 2-APB (10 μm) reduced the depolarization induced by OXA. Notably, Ni2+ was more effective in reducing the transient and late depolarization, whereas 2-APB was more effective on the late depolarization, with respect to nifedipine (Fig. 2A). By using the low-TEA solution (containing 10 mm TEA), the transient response was somewhat reduced, whereas the steady-state depolarization was strongly reduced at only 2–3 mV. This could indicate that OXA inhibited the K+ channels. Finally, in high-TEA solution (containing 145 mm TEA), the whole depolarization, early and late, was strongly reduced at about 1–3 mV (Fig. 2Ab and B), denoting that the first trigger for OXA responses involved ROC. In agreement with the mechanical responses, these results indicate that only the early transient responses were able to reach the voltage threshold for eliciting the contractile responses. The RMP in low- and high-TEA solution did not change significantly, being –55 ± 6 (7 cells; 5 mice) and –54 ± 6 mV (8 cells; 5 mice), respectively.
Figure 2. Membrane depolarization induced by OXA in current-clamp recording.
A, OXA was added at time 0 to the following solutions: (a) control solution (Con), control solution with 2-APB (Con+2-APB) and control solution with Ni2+ (Con+Ni); (b) control solution with nifedipine added (Con+nif), or in low-TEA and high-TEA solutions. The representative voltage traces depicted are chosen from cells having similar RMPs before OXA treatment. B, extent of depolarization induced by OXA compared with the RMP, ΔV, related to the early voltage peak (Vp) and to the late steady-state (Vss) values. *P < 0.05, **P < 0.01 and ***P < 0.001 with respect to control; §P < 0.05, Con+Ni with respect to Con+2-APB; #P < 0.05, Con+2-APB with respect to Con+nif; and +P < 0.05, low-TEA with respect to Con+Ni. Data (means ± SEM) in control solution were from (22 cells; 12 mice); those from the other experimental conditions were from 7 or 8 cells and 4 or 5 mice.
Voltage-gated channels expressed in DLM evaluated by current injection in current-clamp conditions
To investigate the above results in depth, in another set of experiments we stimulated the cells by injecting suitable currents able to depolarize the DLM cells to about –30 mV in order to activate any voltage-gated channels. Current injection in control solution elicited different response patterns. The most complex showed an early fast transient (spike-shaped) depolarization (maximal peak size at –5 ± 0.6 mV). This was followed by a second and less rapid transient and by a delayed slower hump that gradually decayed to a hyperpolarized state (Fig. 3Aa and b). These latter components were observed in all the investigated cells. In contrast, in a total of 44 cells from 12 mice, the early fast depolarization could be clearly identified only in 14 cells from four mice (32% of cells; 33% of mice) and the second less rapid transient in 22 cells from nine mice (50% of cells; 75% of mice). These three depolarizing phases were probably due to voltage-dependent Na+ (INa), T-type (ICa,T) and L-type Ca2+ currents (ICa,L), whereas the hyperpolarizing component was probably due to K+ currents.
Figure 3. Current-clamp experiments.
Time course of voltage responses elicited by injecting currents in DLM cells in solutions without (A) and with OXA (B). A and B, traces represented in a are the same as in b, but with a different x-axis scale (milliseconds vs. seconds) to better show the early onset of the depolarization due to Na+ (Na) and to T-type Ca2+ current (T,Ca); the late slow hump depolarization due to L-type Ca2+ current (L,Ca) is more easily observed in Ab and Bb; Nif indicates the voltage traces (green) recorded in the presence of nifedipine, and 2-APB (red) those obtained in the presence of 2-APB in the bath solution; Ac and Bc show similar experiments from different DLM cells but performed in a low-TEA bath solution (*); moreover, TTX+Ni indicates the voltage traces (blue) obtained in the presence of TTX and Ni2+ in order to record only the depolarization due to L-type Ca2+. All records were made after 6–20 min from OXA stimulation. For A and B, note the different ordinate scales in a, b and c. C, the traces shown [TEA–(TEA+Nif)] are the result of the voltage responses recorded in low-TEA solution minus the response in low-TEA solution with nifedipine. The other traces shown [TEA–(TEA+2-APB)] were obtained by subtracting records made in low-TEA solution with 2-APB in from those in low-TEA solution. D, voltage responses in low-TEA+Nif and choline (Ch)-containing low-TEA solution without (a) and with OXA (b). The numbers of investigated cells are given in the main text.
The early spike was probably due to INa, because it peaked at 0.8 ± 0.09 ms and –5 ± 0.4 mV and was lost when the external Na+ was substituted with choline (Ch-low-TEA solution; 6 cells; 3 mice; Fig. 3Da) or in the presence of the high-TEA solution (8 cells; 3 mice). In contrast, it was not blocked by the L-type Ca2+ channel blocker nifedipine (10 μm; 5 cells; 3 mice). The Na+ current identified in DLM probably belongs to the TTX-sensitive group of Na+ currents, because 1 μm TTX reduced the current size to 92 ± 8% (8 cells; 4 mice; P < 0.01).
The second late transient depolarization was probably due to ICa,T, because it was blocked by the addition of Ni2+ (5 μm; 8 cells; 4 mice; Fig. 3Ac). In Ch-low-TEA solution with added nifedipine (5 cells; 3 mice) the ICa,T peak depolarization was –30 ± 4 mV at 4.2 ± 0.4 ms.
Finally, the addition of nifedipine (10 μm; 8 cells; 3 mice) did not affect INa and ICa,T, but abolished the delayed slow hump, confirming that it was prevalently due to ICa,L (Fig. 3Ab). In the presence of nifedipine, the delayed slow hump was replaced by a slow hyperpolarizing phase (that reached a potential of –80.8 ± 7.6 mV at the end of the 1 s step pulse). In its slow decay, it resembled the Ca2+-dependent K+ current (IK(Ca); BK channel). As expected, this current was blocked on changing the control solution to low-TEA solution (7 cells; 3 mice) and was replaced by a late slow depolarization with a peak depolarization at –37 ± 4 mV (16 ± 2 mV with respect to the RMP; Fig. 3Ac). This was probably due to ICa,L, because in these cells in the presence of nifedipine, only the early depolarization related to INa and ICa,T was detected, whereas the late phase was completely absent (Fig. 3Ac). Finally, a small 2-APB-sensitive depolarization appeared to be superimposed on the ICa,L depolarization, because addition of 2-APB to the bath solution (6 cells; 3 mice) slightly reduced in size the total amount of depolarization (Fig. 3Ab and c). Accordingly, to evaluate the 2-APB-sensitive current, we subtracted the voltage-dependent depolarization recorded in low-TEA solution with 2-APB from that recorded in low-TEA solution. The peak value of the depolarization trace obtained in this way was 2.8 ± 0.3 mV (Fig. 3Ca). Finally, to further estimate the ICa,L-dependent depolarization, we subtracted the voltage traces recorded in low-TEA solution with nifedipine from those recorded without nifedipine. The peak value of the depolarization trace obtained in this way was 18.2 ± 2.5 mV (Fig. 3Ca) and may be considered to be the result of Ca2+ influx through L-type Ca2+ channels with the small depolarization due to 2-APB-sensitive current superimposed.
Effects of OXA on voltage-gated channels and 2-APB-sensitive current evaluated in current-clamp conditions
We next focused on the long-lasting depolarizing effects of OXA on the different ionic currents described in the foregoing. With this aim, each DLM cell analysed in the experimental conditions illustrated in the previous section was treated with OXA, and the responses of current injection were evaluated again during the late steady-state depolarization in the range from 5 to 20 min after application of OXA. Our records showed that OXA increased the size of the depolarization due to INa (peak size, 15 ± 2.1 mV; P < 0.01; Fig. 3Da) with respect to control records (Fig. 3Aa and b). Experiments in Ch-low-TEA solution demonstrated a potentiating effect of OXA on ICa,T (peak size was –25 ± 3 compared with–29 ± 3 mV in control conditions; P < 0.01; Fig. 3D). In addition, the potentiating effect of OXA on ICa,L was evaluated in low-TEA solution with added TTX and Ni2+. Orexin A increased the peak depolarization to –32 ± 4 mV (20 ± 2 mV with respect to the RMP; P < 0.05 with respect to control conditions; Fig. 3Bc). This was also confirmed by subtracting the trace recorded in low-TEA solution with nifedipine (TEA + Nif) from that recorded without nifedipine (TEA). The depolarization traces obtained in this way increased to 26.6 ± 2.5 mV (P < 0.05 compared with experiments without OXA; Fig. 3Cb). Moreover, OXA increased 2-APB-sensitive currents, because the addition of 2-APB resulted in a more pronounced reduction of the late depolarized phase compared with control condition (compare Fig. 3Ac with Bc). This was better evaluated by comparing the difference traces in Fig. 3Cb with Ca; the depolarization related to 2-APB-sensitive current increased to 6 ± 0.7 mV (P < 0.05). Finally, OXA induced a reduction of the late hyperpolarization from –78.3 ± 8.0 to –65.4 ± 7.2 mV (P < 0.05), confirming that inhibition of IK(Ca) contributed to maintain the late sustained depolarization (cf. the traces in Fig. 3Bb with Cb).
Kinetics of the identified voltage-gated channels evaluated in voltage-clamp experiments and action of OXA
The Cm and specific membrane conductance (Gm/Cm) values were also routinely evaluated in the above set of experiments by switching the device from current- to voltage-clamp mode. Notably, another parallel effect of OXA was the reduction of Cm from 41 ± 4 to 32 ± 3 pF (P < 0.01) at the Vp time point and 35 ± 4 pF (P < 0.05) at Vss time point. This reduction in Cm, denoting a cell surface reduction, may represent a consequence of cytoskeletal contraction (Formigli et al. 2005, 2007, 2009a,b). Moreover, OXA increased the specific membrane conductance from 8.3 pS pF−1 in control solution to 35 ± 4 pS pF−1 at the Vp time point and 29.4 ± 4 pS pF−1 at the Vss time point (Table 1).
Table 1.
Resting membrane potential (RMP), specific membrane conductance (Gm/Cm), membrane capacitance (Cm) and Boltzmann parameters of INa, ICa,T and ICa,L activation and inactivation in control and orexin A-treated duodenal longitudinal muscle cells
Parameter | Control | Orexin A treated |
---|---|---|
RMP (mV) | −57 ± 5 | −49 ± 5* |
Gm/Cm (pS pF−1) | 8.3 ± 0.7 | 29.4 ± 2.9** |
Cm (pF) | 41 ± 0.4 | 36 ± 0.4* |
INa | ||
INa/Cm (pA pF−1) | 2.2 ± 0.2 | 3.2 ± 0.3** |
Gm/Cm (nS pF−1) | 76 ± 6 | 102 ± 8** |
Va (mV) | −16.7 ± 1.5 | −21.8 ± 2** |
ka (mV) | 8.0 ± 0.5 | 8.5 ± 0.6 |
Vr (mV) | 47 ± 3 | 46 ± 4 |
Vh (mV) | −50 ± 5 | −61 ± 5* |
kh (mV) | 7.5 ± 0.6 | 6.6 ± 0.5* |
Tp (ms) | 0.4 ± 0.04 | 0.5 ± 0.05* |
ICa,T | ||
ICa,T/Cm (pA pF−1) | 1.5 ± 0.2 | 2.6 ± 0.2** |
Gm/Cm (nS pF−1) | 60 ± 5 | 122 ± 10*** |
Va (mV) | −40.1 ± 0.2 | −45.1 ± 0.3* |
ka (mV) | 7.2 ± 0.4 | 7.1 ± 0.5 |
Vr (mV) | 66 ± 3 | 59 ± 3** |
Vh (mV) | −65 ± 4 | −63 ± 3 |
kh (mV) | 4.5 ± 0.5 | 4.3 ± 0.5 |
Tp (ms) | 7.3 ± 0.5 | 6.7 ± 0.5* |
ICa,L | ||
ICa,L/Cm (pA pF−1) | 3.8 ± 0.3 | 5.8 ± 0.4** |
Gm/Cm (nS pF−1) | 141 ± 12 | 228 ± 18*** |
Va (mV) | −7.9 ± 1 | −14.2 ± 3** |
ka (mV) | 7.4 ± 0.4 | 7.3 ± 0.5 |
Vr (mV) | 67 ± 4 | 61 ± 4* |
Vh (mV) | −50 ± 3 | −55 ± 3* |
kh (mV) | 7.5 ± 0.5 | 7.4 ± 0.5 |
Tp (ms) | 24 ± 3 | 23 ± 3 |
Values listed in the first column are data in control solution; those in the second column were obtained in the same cells 5–20 min after orexin A application. The RMP, Gm/Cm and Cm data are from 44 cells (12 mice). Boltzmann parameters for activation, Va and ka, inactivation, Vh and kh, reversal potential (Vr) and time-to-peak current values at the step voltage eliciting the maximal current (Tp) related to INa, are from 8 cells and 3 mice; those of ICa,T and ICa,L are from 20–22 cells and 7–9 mice.
P < 0.05
P < 0.01
P < 0.005 with respect to control values.
Data are expressed as means ± SEM.
To analyse in detail the effects induced by OXA on the kinetics of a single type of voltage-dependent ionic current in DLM cells, we worked in voltage-clamp conditions.
To study only INa, the cells described in the preceeding subsection showing the fast depolarization due to INa were clamped at –80 mV in low-TEA solution to avoid the occurrence of outward K+ currents. Moreover, we used nifedipine (10 μm) to block L-type Ca2+ current and Ni2+ (5 μm) to block T-type Ca2+ current. As shown in a typical experiment in Fig. 4A, INa at 0 mV peaked at 0.4 ± 0.04 ms. The addition of OXA induced a 1.5-fold increase of INa (Fig. 4B). The bulk of the experimental data are reported in the I–V plot (Fig. 4C), where the mean INa peak amplitude is indicated for any voltage applied, both in control conditions and in the presence of OXA. It can be clearly observed that OXA was able to cause an increase in size and a 10 mV voltage shift of the maximal peak current amplitude towards negative potentials. The voltage shift was better quantified in the steady-state normalized activation curve, fitted by a Boltzmann function by the Va parameter, and it was of about 5 mV (Fig. 4D and Table 1). Moreover, OXA shifted the activation voltage threshold from –45 ± 4 to –54 ± 5 mV (P < 0.05). A greater voltage shift (10 mV) towards negative potentials was observed in the inactivation curve obtained by the inactivating stimulation protocol (current traces not shown; Fig. 4D and Table 1). The decay of INa was fitted to a single exponential function in the whole range of potential studied, and was slightly faster in the presence of OXA (Fig. 4E).
Figure 4. Effects of OXA on TTX-sensitive Na+ current in voltage-clamp experiments.
Families of Na+ currents (INa) recorded in low-TEA solution with Ni2+ and nifedipine added without (A) and with OXA (B); the current traces elicited by voltage pulses over that inducing the maximal current are depicted as thin lines. C, I–V plots of the mean INa peak value versus voltage in control and OXA-stimulated cells; the fits of a Boltzmann function are superimposed on the data. D, the fits of normalized activation and inactivation Boltzmann functions are superimposed on the data symbols. Boltzmann function parameters are listed in Table 1. Vertical lines indicate the resting membrane potential in control conditions (dashed line) and in the presence of OXA (continuous line). E, time constant (τ) for voltage dependence of current decay from all investigated cells; the fit of a single exponential function is superimposed on the data. Orexin A hastened the current decay (τ value at –20 mV with OXA was 3.9 ± 0.5 ms and in control conditions 3.0 ± 0.4 ms; P < 0.05), but not its voltage dependence (29 ± 3 and 30 ± 3 ms, respectively). In C–E, data are means ± ESM from 8 cells (3 mice).
In another set of experiments, the DLM cells that did not in current-clamp conditions show depolarization due to INa but to ICa,T and ICa,L were clamped at –80 mV in high-TEA (Na+- and K+-free) solution to avoid IK and Ca2+ current flowing through ROCs. Both ICa,T and ICa,L were recorded by applying a depolarizing pulse protocol (1 s long) from –70 to +50 mV in 10 mV increments. In the presence of nifedipine (10 μm; 12 cells; 4 mice) we could observe only ICa,T as a low-voltage-activated inward transient current (voltage threshold was at –60 ± 6 mV). In a typical experiment, as shown in Fig. 5A, the time to peak was about 7.3 ms at –20 mV. Orexin A (0.3 μm) induced a 1.7-fold increase in the size of ICa,T peak and shortened the time to peak (by about 6.7 ms; Fig. 5B and Table 1).
Figure 5. Effects of OXA on T- and L-type currents in voltage-clamp experiments.
Typical families of T- (A and B) and L-type Ca2+ currents (C and D) in high-TEA solution without (A and C) and with OXA (B and D). The numbers indicate the voltage inducing the maximal current value and the related time to peak (tp). The current traces elicited by voltage pulses over that inducing the maximal current are depicted as thin lines.
To evaluate the effect of OXA on ICa,L, we carried out experiments in the presence of Ni2+ (5 μm; 12 cells; 4 mice). The ICa,L appeared as a high-voltage-activated (–40 ± 4 mV) current that slowly inactivated to a quasi-steady state. Orexin A induced a roughly 1.5-fold increase of both peak size and late amplitude of ICa,L (Fig. 5D) compared with the control conditions (Fig. 5C), and caused a small reduction of the time to peak as well (to 23–24 ms; Table 1).
The I–V plots of ICa,T and ICa,L peaks both in the absence and in the presence of OXA are reported in Fig. 6A and B, respectively. The increment in amplitude caused by OXA is clearly observable. Figure 6 reports the steady-state activation and inactivation curves for normalized ICa,T (Fig. 6C) and ICa,L (Fig. 6D) without (control) and with OXA. Orexin A induced an approximately 5 mV negative shift of the ICa,T activation curve, without affecting inactivation. In contrast, for ICa,L a voltage shift was observed in both activation and inactivation voltage dependence. As a consequence, OXA resulted in a negative shift of the voltage threshold that was not statistically significant for ICa,T (from –60 ± 6 to –62 ± 6 mV) but was significant for ICa,L (from 31 ± 3 to 43 ± 4 mV; P < 0.01). Figure 6D suggests that the steady-state inactivation was U-shaped and that the reduction of the degree of inactivation at positive potentials was potentiated by OXA (at +50 mV, the normalized Ih value increased from 0.28 ± 0.03 to 0.48 ± 0.05; P < 0.01).
Figure 6. Effects of OXA on Boltzmann activation and inactivation functions of T- and L-type Ca2+ currents.
Current–voltage curves related to T- (A) and L-type Ca2+ currents (B); the Boltzmann fit for activation is superimposed on the data. Normalized activation (m) and inactivation curves (h) for T- (C) and L-type Ca2+ currents (D) are superimposed on the plots. In D, the arrow indicates the change induced by OXA on the U-shaped inactivation curve at positive potentials. All the curves are related to cells without (control; open symbols) and with OXA (OXA; filled symbols). Experiments were carried out in external high-TEA solution. Numbers of experiments and Boltzmann function parameters are listed in Table 1. Vertical lines indicate the resting membrane potential in control conditions (dashed line) and in the presence of OXA (continuous line).
The Boltzmann fit parameters of the activation curve (Table 1) indicate that the increases in size of peak INa, ICa,T and ICa,L were due both to an increase of the maximal conductance (Gm/Cm) and to the negative voltage shift of the activation curves. For INa and ICa,L, the peak was further increased by the negative shift of the inactivation curves. Moreover, the shift of the activation curves towards more negative potentials suggests a greater excitability of OXA-treated cells. Notably, OXA shifted Vr of both ICa,T and ICa,L negatively by about 7 mV (Table 1), denoting that [Ca2+]i was increased by OXA. In contrast, the Vr of INa was not modified by OXA, denoting that the increase of membrane Gm/Cm in the late depolarizing phase was prevalently a result of Ca2+ entry through L-type Ca2+ channels and 2-APB-sensitive channels.
In a different set of experiments (9 cells; 4 mice), DLM cells that in current-clamp conditions did not show INa depolarization were clamped at –80 mV in the control solution with Ni2+ and nifedipine (10 μm) added to evaluate the voltage dependence of IK(Ca). The IK(Ca) was identified by its relatively rapid activation followed by small and slow inactivation and noisy traces at positive potentials (Fig. 7A). In addition, it was blocked by a low TEA concentration (2 mm). Orexin A did not change the activation voltage threshold (–25.2 ± 2.7 and –23 ± 2.5 mV for control and OXA-treated cells, respectively), nor the activation Boltzmann parameters (Va was 10 ± 2 and 11 ± 2 mV in control and OXA-treated cells, respectively; Fig. 7B). The only parameter significantly affected by OXA was the maximal current size, which was reduced from 25 ± 2.2 to 17 ± 2 pA pF−1 (P < 0.05; Fig. 7C).
Figure 7. Effects of OXA on IK(Ca) (BK current).
Typical current traces recorded in control solution with nifedipine added before (A) and 5–20 min after addition of OXA to the bath (B). C, I–V plots. Orexin A induces a positive shift of Va. The related Va and ka in control conditions were 10.0 ± 1 and 10.1 ± 1 mV, respectively and in the presence of OXA 14.9 ± 2 (P < 0.01) and 9.8 ± 1 mV, respectively. Data were from 10 cells (4 mice).
To assess the effects of OXA on thapsigargin-induced current, we carried out another set of voltage-ramp experiments in high-TEA solution to block ROC currents induced by OXA. To this end, in eight cells from four mice, the sarcoplasmic reticulum was Ca2+ depleted by Tg, and after 7 min of Tg treatment OXA (0.3 μm) was added to the bath solution. A voltage ramp was applied every 1 min. The I–V plots of thapsigargin-induced current, obtained by subtracting the average of two control voltage ramps representing the leak current, were linear or showed a small rectification and a positive shift of the reversal potential of 5.4 ± 0.6 mV (Fig. 8A). The following I–V plots recorded after adding OXA (also obtained by subtracting the average of two control voltage ramps) showed progressively larger current that reached the highest values after about 5 min from the addition of OXA (Fig. 8B), which is the time when, in current-clamp experiments, OXA reached the steady-state depolarization (Fig. 2A). This is in accord with a greater sarcoplasmic reticulum Ca2+ depletion and SOC current activation. Notably, OXA also induced a negative shift of the reversal potential of 14.4 ± 0.8 mV (from 5.4 ± 0.6 to –9 ± 1 mV; P < 0.01), denoting a change of the driving force for Ca2+ entry due to the Ca2+ influx through SOCs.
Figure 8. Voltage-ramp experiment.
Thapsigargin (Tg)- and OXA-induced SOC currents in DLM cells bathed in high-TEA solution. A, voltage ramps ranging from –90 to 50 mV from a holding potential of 0 mV (duration 0.5 s; see Methods for details) were applied every 1 min. Leak-current correction was done by subtracting the average of two ramps recorded before application of Tg. The current traces shown were elicited 2, 4 and 6 min after addition of Tg, and 2, 4, 6 and 8 min after the addition of OXA. B, specific conductance at –50 and 50 mV, elicited by all applied voltage ramps. Data are from 13 cells (4 mice).
Discussion
Direct action of OXA on mouse DLM cells
The present results, obtained by a combined mechanical and electrophysiological approach, show, for the first time to our knowledge, that OXA exerts direct excitatory effects on the mouse duodenum and explain the mechanism of action underlying these responses.
In fact, even if direct smooth muscle contractions in response to OXA have been observed in rat jejunum segments (Kroczynski et al. 2006b), the mechanism of action has not yet been elucidated.
In the present mechanical experiments, contractile responses to OXA were obtained in the presence of TTX, thus indicating a direct action of the peptide on the smooth muscle. This observation is in strong agreement with the presence of OXA receptors at the gut smooth muscle level (De Miguel & Burrell, 2002; Näslund et al. 2002; Dall'Aglio et al. 2008). However, the present results obtained in the presence of TTX do not allow us to exclude the possibility that OXA may also act at the neural level. In this respect, in addition to the influence of OXA on inhibitory enteric neurons in the isolated gastrointestinal preparations (Satoh et al. 2001; Ehrström et al. 2003; Baccari & Calamai, 2008; Baccari et al. 2009), neurally mediated contractile effects have been reported; in vitro experiments have shown that OXA enhances motility of the guinea-pig distal colon (Kirchgessner & Liu, 1999) and also induces contractile effects in the small intestine of different animal species (Kirchgessner & Liu, 1999; Matsuo et al. 2002; Korczynski et al. 2006a, b). Activation of cholinergic neurons by orexins has been reported in isolated intestinal preparations (Satoh et al. 2001), and intracellular recordings from isolated myenteric neurons of the guinea-pig ileum have demonstrated an increase of acetylcholine release by the peptide (Katayama et al. 2003, 2005).
In the present mechanical study, the reduction in amplitude of the contraction in response to OXA caused by nifedipine and 2-APB suggests the involvement of L-type Ca2+ channels and SOCs, respectively, in the effects of the peptide. In contrast, the abolition of the response to OXA by low TEA concentrations (1 mm) and by the T-type Ca2+ channel blocking agent, Ni2+ (50 μm), indicates that more than one pathway is involved in the Ca2+ elevation induced by OXA. Surprisingly, in the present experiments, lower concentrations of Ni2+ and TEA were required to abolish the response to OXA compared with those used to block SOCs (Ni2+, 5 mm; Kukkonen & Åerman, 2001; Larsson et al. 2005) and ROCs (TEA, 70 mm; Larsson et al. 2005; Johansson et al. 2007), respectively. This could indicate that the SOC and ROC Ca2+-entry paths activated by OXA in DLM showed greater susceptibility to blockade by Ni2+ and TEA or, alternatively, that other mechanisms may operate, such as inhibition of K+ channels and an increase in T-type Ca2+ current.
Electrophysiological experiments in current-clamp conditions demonstrated that OXA initially induced a depolarizing transient response lasting about 5 min, followed by a long-lasting steady-state depolarization. The transient depolarization paralleled the mechanical responses, showing that only the early transient responses were able to reach the voltage threshold for eliciting the contractile responses. This may also explain the lack of effects of a subsequent application of OXA performed after a short interval and without having washed the preparations in mechanical experiments. Notably, the trigger for the depolarization induced by OXA was predominantly due to the activation of ROCs, because the early and long-lasting depolarization was blocked in high-TEA solution. The OXA transient response was enhanced by various mechanisms, initially involving the low-voltage-activated T-type Ca2+ channel, and then the high-voltage-activated L-type Ca2+ channel, as suggested by the more pronounced reduction of the transient response induced by Ni2+ compared with nifedipine. Finally, the duration of about 5 min of the transient response was a long enough time to cause at least a partial Ca2+ depletion of the sarcoplasmic reticulum and to activate SOCs, as demonstrated by 2-APB and voltage-ramp experiments.
The depolarization induced by OXA is maintained not only by the aforementioned mechanisms, but also by inhibition of IK(Ca). In fact, in the presence of OXA the hyperpolarization due to K+ currents was reduced, confirming that OXA inhibited these channels. In accord, a low TEA concentration reduced all the depolarizing responses induced by OXA. This latter mechanism is likely to operate in particular during the late steady-state depolarization, because this is more strongly depressed. Concerning the absence of contractile responses induced by OXA in solution with TEA (1 mm) and Ni2+ (50 μm), the reduced depolarization may be unable to activate voltage-dependent ionic channels substantially. In accord, the early transient depolarization in the presence of these substances did not reach –50 mV. Therefore, the contributions of Ca2+ influx through T- and L-type Ca2+ channels and the inhibition of K(Ca) channels are essential for inducing the contractile responses. Accordingly, nifedipine and 2-APB, which caused small decrease of the early transient response, significantly reduced the mechanical response.
Previous studies suggest that the primary pathway for OX1R-mediated Ca2+ elevation is the activation of a non-store-operated Ca2+-permeable channel (Holmqvist et al. 2005), even though pharmacological distinction of non-store-operated channels from store-operated mechanisms is difficult because of the lack of specific blockers or other specific means to distinguish the pathways. However, our results in DLM showed that the contribution of SOCs is smaller than that of voltage-gated L-type Ca2+ channels in cells at resting length. Orexins have been reported to act on G-protein-coupled receptors, OX1R and OX2R (Kukkonen et al. 2002), which may couple to different messenger systems (Ferguson & Samson, 2003), and this may hamper the investigation of physiologically relevant responses to orexins.
Voltage-gated channels expressed in DLM
The first aim of this research was to evaluate whether OXA could act directly on DLM cells. Our results showed new findings, such as the presence of voltage-gated Na+ and T-type Ca2+ currents in DLM, which have rarely been observed before in longitudinal smooth muscle of other regions of the gastrointestinal tract, such as jejunum, ileum and colon. Other new findings were the peculiar behaviours of ICa,L, such as the decay to a steady state different from zero and a U-shaped inactivation, and the properties of SOC. There follows a discussion of these new findings.
The expression of voltage-gated channels in smooth intestinal muscle is species, age and intestinal tract specific (Hara et al. 1986; Kuriyama et al. 1998). Moreover, differences have been observed between circular and longitudinal smooth muscle cells (Hara et al. 1986; Strege et al. 2007). The availability of expressed Na+ channels and of their kinetics showed a greater variability (Smirnov et al. 1992; Strege et al. 2007; Zhu et al. 2010). The molecular basis of Na+ channels in smooth muscle are Nav1.5 and Nav1.6. The expression of Nav1.5 has been identified in circular but not in longitudinal smooth muscle cells of mouse jejunum (Strege et al. 2007).
Our new finding shows that the expression of TTX-sensitive Na+ channels in the longitudinal smooth muscle cells of duodenum is more consistent than that observed in longitudinal smooth muscle of other intestinal segments, because it was identified in 39% of cells and 33% mice investigated. The voltage threshold of INa activation was at a voltage near the RMP, and about 50% of the Na+ channels were capable of being activated at the RMP. Thus, as in the smooth muscle cells of the longitudinal layer of the ileum in newborn rats (Smirnov et al. 1992) and in TTX-sensitive Nav1.6 (Zhu et al. 2010), the Na+ channels of DLM were capable of being activated at RMP. Moreover, the gating properties of Na+ channels in DLM cells may suggest a role in contributing to the setting of the membrane potential and, in particular, they will play a major role in regulating the excitability, mainly in response to rapid depolarizing stimuli.
Another new finding of this work is the expression in DLM of T-type Ca2+ channels, which are typically less expressed or not expressed at all in many intestinal smooth muscle cells (Yoshino et al. 1989; Smirnov et al. 1992; Kuriyama et al. 1998). The mean activation voltage threshold was at –60 mV and the windows voltage was from –65 (Vh) and –40 mV (Va). Thus, this current could only contribute to regulation of fast changes of the RMP in hyperpolarized cells, specifically those with an RMP more negative than about –65 mV.
The voltage threshold for L-type Ca2+ channel activation in DLM is at a relatively high depolarization state, about –38 mV. Thus, the activation of T-type Ca2+ channels and of Na+ channels may play a significantly role in the activation of ICa,L in cells at RMP, because at this voltage its inactivation was only about 40–50%. Moreover, once activated the ICa,L did not completely inactivate, becausee (in voltage-clamp conditions) its decay reached a steady state different from zero; in addition, it showed a U-shaped inactivation curve. This could indicate that inactivation was Ca2+ dependent, because it was more evident in OXA-treated cells. This was probably due to the greater intracellular [Ca2+], as a consequence of the increased Ca2+ influx trough L-type Ca2+ channels, which was confirmed by the shift of ICa,T and ICa,LVr towards more negative potentials. Alternatively, in agreement with Ca2+-independent kinetic models for the U-shaped inactivation curve, we have to take into consideration the finding that when the depolarization approached the ICa reversal potential in control and OXA-treated cells the fall of inactivation was less than the reduction in Ca2+ entry (Francini et al. 1992). The present results suggest that ICa,L inactivation in DLM is mediated by two complementary processes that are internal Ca2+ dependent and potential dependent. The consequence was that DLM could retain a long-lasting depolarized state, useful for maintaining a long-lasting contractile state, and a rapid contractile rate, functional for duodenal segmental contraction.
In low- and high-TEA solutions, the RMP did not change significantly. This indicates that the RMP is predominantly maintained by the Na+–K+-ATPase pump, with a small contribution of K+ channels. In agreement, the mean voltage threshold of IK(Ca) activation was –25 mV, a voltage value less negative than RMP (–57 ± 5 mV). Thus, K+ channels in DLM cells may act, at least in part, to counteract cells in a depolarized state, whereas inhibition K+ channels, as observed in the presence of OXA, may have a role in maintaining cell depolarization.
Distinct store-dependent cationic currents have been described, suggesting a molecular heterogeneity (Liu et al. 2004). The SOC properties in DLM are similar to those of human parotid gland cells (Liu et al. 2004) and to SACs observed in C2C12 myoblasts (Formigli et al. 2005, 2007, 2009a,b). Transient receptor potential channel 1 is a strong candidate for SOCs and SACs (Maroto et al. 2005; Clark et al. 2008; Formigli et al. 2009b). The reduced value of Cm induced by OXA may be a consequence of Ca2+-dependent cytoskeletal contraction, which in turn increases the membrane tension and activates SACs (Formigli et al. 2009a,b;). Interestingly, Nav1.5 and Nav1.6 are mechanosensitive, and have a similar hyperpolarizing shift in activation voltage dependence of INa in response to mechanical stimulation. Notably, the mechanosensitivity of SACs and Na+ channels (Strege et al. 2007; Wang et al. 2009) may be useful to depolarize the DLM cells and activate ICa,L when the cells are stretched.
Action of OXA on the kinetics of voltage-gated channels and SOCs
Our results demonstrate that OXA enhanced INa, ICa,T and ICa,L not only by increasing the related Gm/Cm, but also by producing a negative shift of about 5 mV in the Va parameter of INa, ICa,T and ICa,L activation curves. The negative shift of about 7 mV of ICa,T and ICa,LVr confirmed that the OXA-induced depolarization is principally due to the increase of intracellular [Ca2+] from Ca2+ entry through ROCs, T- and L-type Ca2+ channels and SOC activation. Orexin A also affected the kinetics of ICa,T and ICa,L inactivation, in that it reduced kh of INa and resulted in a negative shift of the Vh parameter of INa and ICa,L, as well as the ICa,L Ca2+ dependence (U-shaped inactivation). The action of OXA on ICa,L inactivation, together with IK(Ca) inactivation and SOC activation, may explain the maintained depolarization in the late long-lasting phase. Moreover, OXA reduces the value Cm, indicating a reduction in cell surface as a consequence of cytoskeletal contraction (Formigli et al. 2005, 2007, 2009a,b), which in turn activates the DLM mechanosensitivity.
In conclusion, the effects of OXA in DLM result in a direct contraction, supported by complex mechanisms involving intracellular messenger systems that affect not only ROCs but also more than one conductance, which may include SOCs and voltage-gated Na+, T- and L-type Ca2+ and K(Ca) channels. In addition to the effects of OXA on the central nervous system (Hwang et al. 2001; Kukkonen et al. 2002; Grabauskas & Moises, 2003; Baccari, 2010) and on postganglionic cholinergic neurotransmission (Kirchgessner & Liu, 1999; Satoh et al. 2001; Matsuo et al. 2002; Korczynski et al. 2006a,b;), OXA also exerts direct contractile actions on the duodenal smooth muscle. This direct effect may represent a physiological mechanism, acting in a synergic manner to reinforce the neural signal and/or aimed to compensate for the few dorsal motor nucleus of vagus neurons responsive to orexins that supply the duodenum (Grabauskas & Moises, 2003).
Acknowledgments
The authors are grateful to Adrio Vannucchi for its technical assistance in preparing the figures. This work was founded by grant ex 60% of the University of Florence, Italy to F.F. and M.C.B. The authors declare no conflict of interests.
Glossary
Abbreviations
- 2-APB
2-aminoethyl diphenyl borate
- Ch
choline
- DLM
duodenal longitudinal muscle
- ICa,L
L-type Ca2+ current
- ICa,T
T-type Ca2+ current
- IK(Ca)
Ca2+-dependent K+ current
- INa
TTX-sensitive Na+ current
- OXA
orexin A receptor
- OXB
orexin B
- OX1R
orexin-1 receptor
- OX2R
orexin-2 receptor
- RMP
resting membrane potential
- ROC
receptor-operated Ca2+ channel
- SAC
stretch-activated channel
- SOC
store-operated Ca2+ channel
- Tg
thapsigargin
- VGCC
voltage-gated Ca2+ channel
Author contributions
Conception and design, collection, analysis and interpretation of data, drafting the article: F.F., R.S. and G.L. for electrophysiological studies; M.C.B. and R.G. for mechanical ones. All authors contributed to writing the manuscript, which was critically revised by F.F. and M.C.B. All authors read and approved the final version of the manuscript.
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