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CELLULAR |
1 Department of Anaesthetics, Pain Medicine and Intensive Care, Chelsea & Westminster Hospital, Imperial College London, UK
2 Biophysics Section, Blackett Laboratory, Imperial College London, UK
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(Received 4 December 2006;
accepted after revision 8 January 2007;
first published online 11 January 2007)
Corresponding author S. Trapp: Biophysics Section, Blackett Laboratory, South Kensington Campus, Imperial College London, London SW7 2AZ, UK. Email: s.trapp{at}imperial.ac.uk
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Introduction |
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It is generally accepted that ATP-sensitive K+ (KATP) channels mediate the hyperpolarization of GE cells (Ashford et al. 1990; Spanswick et al. 1997, 2000; Dallaporta et al. 2000; Miki et al. 2001; Mobbs et al. 2001; Song et al. 2001; Dunn-Meynell et al. 2002; Balfour et al. 2006). In contrast, the mechanism underlying the depolarization of GI cells is more controversial. Previous studies have suggested that inhibition of the Na+–K+-ATPase, blockade of a Cl– conductance or inhibition of an acid-sensitive two-pore-domain K+ channel might underlie the response of GI cells in hypothalamic neurones (Oomura et al. 1974; Silver & Erecinska, 1998; Song et al. 2001; Routh, 2002; Burdakov et al. 2006).
KATP channels of GE neurones open in response to brief hypoglycaemia, but they are also found in a large proportion of neurones that are not glucosensors (Liss & Roeper, 2001; Mobbs et al. 2001; Balfour et al. 2006). Mostly, KATP channels seem to serve a protective role during short periods of ischaemia or hypoxia (Yamada et al. 2001). We have recently proposed that these differences in the role of KATP channels between cells do not result from different properties of these channels, but rather from differences in energy metabolism, specifically glucose handling, between different neurones (Balfour et al. 2006). We demonstrated that, as for pancreatic 
-cells, the expression of glucokinase is essential for glucosensing vagal neurones, and that glucose metabolism to ATP determines the activity of KATP channels (Balfour et al. 2006).
By way of analogy, we hypothesize that the increase in activity seen in GI neurones with falling sugar levels does not involve unique electrical mechanisms of glucosensing cells, but rather the same components that are used to respond to anoxia or ischaemia.
In order to test this hypothesis, we explored and compared the electrical responses of dorsal vagal neurones to hypoglycaemia and to mitochondrial inhibition. These neurones provide central nervous regulation of gastrointestinal function and endocrine pancreatic secretion and can thus directly or indirectly influence appetite and blood glucose levels (Laughton & Powley, 1987; Powley, 2000; Wu et al. 2004; Travagli et al. 2006). A portion of dorsal vagal neurones are intrinsic glucosensors, with GI and GE types present in roughly equal numbers. The remaining cells are a heterogeneous population showing various responses to mitochondrial inhibition (Cowan & Martin, 1992, 1995; Trapp & Ballanyi, 1995; Ballanyi et al. 1996; Ferreira et al. 2001; Balfour et al. 2006).
In this study, we focused on early depolarizing responses to metabolic inhibition, since these have not yet been characterized in any detail. We found that mitochondrial inhibition and hypoglycaemia elicited the same electrical responses. Our results indicate the involvement of two-pore-domain K+ channels and the Na+–K+-ATPase, but not a Cl– channel, in the depolarizing response of dorsal vagal neurones to metabolic inhibition. We observed that mitochondrial inhibition and glucose deprivation activate common electrical responses and conclude that glucose is sensed via its metabolic effects on ATP production.
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Methods |
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Electrophysiological studies
Brainstem slices (200 µm) were obtained from rats after halothane anaesthesia and transcardial perfusion with low-Na+ solution containing (mM): 200 sucrose, 2.5 KCl, 28 NaHCO3, 1.25 NaH2PO4, 3 pyruvate, 7 MgCl2, 0.5 CaCl2 and 7 glucose. The pH of all bicarbonate-buffered solutions was adjusted to 7.4 by gassing with 95% O2/5% CO2. After recovery at 34°C for 30 min in a solution containing (mM): 118 NaCl, 3 KCl, 25 NaHCO3, 1.2 NaH2PO4, 7 MgCl2, 0.5 CaCl2 and 2.5 glucose, slices were kept at room temperature (20-22°c) in artificial cerebrospinal fluid (ACSF) of the following composition (mM): 118 NaCl, 3 KCl, 25 NaHCO3, 1.2 NaH2PO4, 1 MgCl2, 1.5 CaCl2 and 10 glucose. Hypoglycaemic ACSF was prepared by equimolar replacement of glucose with sucrose. Patch pipettes were pulled from thin-walled borosilicate capillaries (3–6 M
; Harvard Apparatus Ltd., Edenbridge, Kent, UK) with a horizontal puller (Zeitz, Germany). For most recordings, electrodes were filled with (mM): 120 potassium gluconate, 5 Hepes, 5 BAPTA, 1 NaCl, 1 MgCl2, 1 CaCl2 and 2 K2ATP (pH 7.2; adjusted with KOH). Some recordings were performed with a ‘high-Cl–’ pipette solution of the following composition (mM): 140 KCl, 1 NaCl, 1 MgCl2, 1 CaCl2, 5 Hepes, 5 EGTA and 2 K2ATP (pH 7.2; adjusted with KOH).
Whole-cell recordings were carried out in ACSF at room temperature. Most drugs were added directly to the ACSF. Tolbutamide and diazoxide were prepared in 0.1 M NaOH as 0.05 and 0.02 M stock solutions, respectively. Strophanthidin was prepared as a 0.5 M stock solution in DMSO. Tetrodotoxin was prepared as a 1 mM stock solution in sodium citrate buffer. The recording chamber (volume 2 ml) was perfused with ACSF at a rate of 4–5 ml min–1.
Recordings were performed in both voltage-clamp and current-clamp mode using an EPC-9 amplifier and Pulse/Pulsefit software (Heka Elektronik, Lambrecht, Germany). Currents or membrane potentials were filtered at 1 kHz and digitized at 4 kHz. Membrane resistance was monitored with 500 ms current or voltage pulses every 5 s. Current–voltage (I–V) relationships were obtained either by analysing hyperpolarizing and depolarizing 200 ms voltage or current steps, or by holding the cell at –20 mV for a minimum of 30 s and then applying a voltage ramp from –20 to –120 mV over 700 ms. Compensation for the junction potential (+10 mV for the potassium gluconate pipette solution) was performed off-line. Recordings displayed in figures are adjusted for the junction potential.
Data are presented as means ± 1 S.E.M. Statistical significance was tested using one-way ANOVA followed by the Tukey's post hoc test, unless stated otherwise. P values of < 0.05 (*) and < 0.01 (**) were taken to indicate that the data were significantly different.
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Results |
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1000 times that of Na+), it would be expected that KATP channel opening would lead to a current with a reversal potential at EK (Ashcroft, 1988).
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The KATP current elicited by cyanide, azide or hypoglycaemia caused hyperpolarization and cessation of action potential firing under current-clamp conditions (n = 3–6 each). Close inspection of the time course of the electrical response to metabolic inhibition of these cells revealed that the hyperpolarization was preceded by a small depolarization and increase in firing rate (Fig. 1). In voltage-clamp mode, a small inward current (approx. 30 pA at –60 mV) was observed prior to KATP channel opening in response to azide (n = 8), cyanide (n = 4) and hypoglycaemia (n = 10; Fig. 2).
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The inward current elicited by cyanide was independent of KATP channel activity (Fig. 3). It was observed in the presence of diazoxide (n = 4) when KATP channels were open and in the presence of tolbutamide when KATP channels were closed (n = 4) and in DVN that have no functional KATP channels (approximately 60% of DVN, as determined by the failure of cyanide or azide to elicit a KATP current; n = 37; Fig. 4A).
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We next investigated whether these responses persisted in the presence of 0.5 µM TTX. For 26 cells tested, TTX completely inhibited voltage-gated Na+ currents in DVN (Fig. 4D, inset), but the inward currents in response to cyanide, azide or glucose removal persisted (Fig. 4D). Five cells developed the inward current within 3 min (30–80 s) of perfusion of glucose-free solution, and a further three DVN were exposed to glucose-free solution for up to 30 min (onset of inward current after 240–450 s). The remaining 18 cells were exposed to 1 mM cyanide and/or 3 mM azide. For all groups, the time to onset of the inward current was not significantly different from that observed for the equivalent group in the absence of TTX (Fig. 4E).
Biophysical analysis of the inward current elicited by metabolic inhibition
Analysis of the I–V relationships for the small inward current revealed that metabolic inhibition caused a decrease in conductance with a reversal potential close to EK, or an increase in conductance with a reversal potential which varied between –58 and +10 mV, or an inward current that did not reverse within the voltage range analysed (Fig. 5A).
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Equivalent responses were also elicited by hypoglycaemia. A decrease in conductance with a reversal potential of –94 ± 3 mV was observed for three neurones after prolonged hypoglycaemia. An increase in conductance with a reversal potential between –63 and –33 mV was seen in a further five DVN under these conditions (Fig. 5A). Glucose-inhibited neurones also responded with the same types of currents. A short period of hypoglycaemia caused a decrease in conductance with a reversal of –94 ± 2 mV in five cells, an increase in conductance with reversals between –48 and +10 mV in 10 cells (Fig. 5A) and a parallel shift in one DVN.
In the presence of 0.5 µM TTX, cyanide reduced a conductance with a reversal potential of –95 mV in four DVN and caused a parallel shift in the remaining 11 cells. Under the same conditions, all three types of response were seen for azide (n = 8). Glucose removal caused a fast K+ channel block in three cells and a parallel shift in two more. Three cells were exposed to prolonged hypoglycaemia, with two exhibiting a K+ channel block and one showing an increase in conductance with a reversal potential of –41 mV.
Reversal potentials of the observed inward current between –30 and –60 mV would be consistent with the opening of Cl– channels. To test this hypothesis, we performed recordings with a pipette solution containing 145 mm Cl–, compared with 5 mM Cl– for the normal internal solution. With the high-Cl– internal solution, ECl was 0 mV, compared with
–40 mV with normal internal solution (as determined by the response to 1 mM GABA). Should the current response to metabolic inhibition be mediated by Cl–, a depolarizing shift of the reversal potential of the current would be expected. This was not observed. In recordings with ‘high-Cl–’ electrodes, cyanide elicited an inward current of –26 ± 9 pA at –80 mV, after a delay of 100 ± 27 s (n
= 4). Two cells each showed a decrease in conductance with a reversal potential of –92 mV and the other two cells displayed an increase in conductance with reversal potentials of –50 and –35 mV.
A decrease in conductance: the closure of an openly rectifying K+ channel
The decrease in conductance elicited by metabolic inhibition in approximately 25% of recordings had a reversal potential close to EK (Fig. 5A). This current was well described by the Goldman–Hodgkin–Katz (GHK) equation, indicating openly rectifying potassium channels such as the two-pore-domain K+ channels (Fig. 5B; Hille, 2001).
We have previously identified functional two-poredomain K+ channels of the TASK subfamily in DVN (Hopwood & Trapp, 2005). These channels are activated by the volatile anaesthetic halothane. To test whether TASK-like channels might be involved in the response to metabolic inhibition, we assessed the response to 1 mM cyanide and to 1 mM halothane in eight DVN.
In three of these recordings, cyanide inhibited an openly rectifying K+ channel, but halothane reduced the holding current at –20 mV by 71 ± 15 pA (Fig. 5C). An increase in current by 58 ± 14 pA at –20 mV in response to halothane, as would be expected from functional TASK-like channels, was observed in three further DVN. However, two of these exhibited an increase in conductance with a reversal potential of –30 mV in cyanide and a KATP current developed in the third. These results suggest that the cyanide-inhibited openly rectifying K+ current was not mediated by TASK-like channels.
An increase in conductance: a role for the Na+–K+-ATPase?
The large scatter in reversal potentials observed for the increase in conductance under metabolic inhibition (Fig. 5A) suggests that more than one component contributes to the inward current induced by metabolic inhibition. Furthermore, the presence of a reversal potential within the observed range indicated that the inward current was not solely due to the inhibition of the Na+–K+-ATPase. The Na+–K+ pump current should reverse at approximately –600 mV (Thomas, 1972; Senatorov et al. 1997). However, any contribution of Na+–K+-ATPase inhibition to the inward current during metabolic inhibition would right-shift the I–V relationship of this current to more positive reversal potentials. If the contribution of the pump current to the inward current varied between cells, a wide spread of reversal potentials would be expected. To test this hypothesis, we assessed the response of DVN to the established Na+–K+-ATPase inhibitors strophanthidin and ouabain.
Bath application of 100 µM strophanthidin elicited an inward current of –46 ± 7 pA (n = 3) and 50 µM ouabain led to an inward current of –40 ± 11 pA (n = 4) at –80 mV. In individual cells, the inward current induced by Na+–K+-ATPase inhibition was slightly larger than the current induced by cyanide or azide (P < 0.05, n = 8, Student's paired t test). The increase in conductance in response to metabolic inhibition had reversal potentials between –58 and +10 mV (Fig. 5A), but Na+–K+-ATPase inhibition elicited an inward current with no obvious reversal potential (Fig. 6B). In order to test what effect inhibition of the Na+–K+-ATPase has on the cellular response to metabolic inhibition, cells were exposed to 3 mM azide or 1 mM cyanide after pre-incubation with a Na+–K+-ATPase inhibitor (100 µM strophanthidin or 50 µM ouabain). However, this resulted in a very large progressive inward current (> 1 nA) and cell death (n = 6).
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Discussion |
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Previous studies have demonstrated that metabolic challenges, such as anoxia, cyanide or prolonged aglycaemia, elicit KATP currents in a proportion of DVN (Trapp & Ballanyi, 1995; Ballanyi et al. 1996; Ballanyi & Kulik, 1998; Raupach & Ballanyi, 2004). We have recently shown that, additionally, a subpopulation of DVN are GE neurones that open KATP channels in response to brief hypoglycaemia (Balfour et al. 2006).
Unexpectedly, DVN with functional KATP channels also showed a small inward current during metabolic inhibition. The inward current seemed to be more sensitive to ATP depletion than the KATP current, because it preceded KATP channel opening (Figs 1 and 2). Interestingly, a small depolarization prior to hyperpolarization has also been reported for hippocampal CA1 neurones during hypoxia (Erdemli et al. 1998), indicating that this inward current is not specific for DVN. Our results suggest that at least three different electrical mechanisms are underlying the depolarizing response of DVNs to energy depletion.
Inhibition of two-pore-domain K+ channels by metabolic inhibition
It has been proposed that hypothalamic GI neurones depolarize during hypoglycaemia owing to the closure of a Cl– conductance (Song et al. 2001). However, our data provide an argument against this possibility for vagal neurones. The only reduction in conductance that was observed in this study was the inhibition of a K+ channel. This inward current was well fitted by the GHK equation, indicating the inhibition of an openly rectifying K+ channel (Hille, 2001). The rectification properties, together with the observation that these channels were open during resting conditions, indicate that this effect was not mediated by the closure of KATP channels. KATP channels are inward rectifiers and were shown to be closed under control conditions in our in vitro slice preparation (Balfour et al. 2006). Interestingly, a recent study on glucose-inhibited orexinergic neurones of the lateral hypothalamus demonstrated the inhibition of an outwardly rectifying K+ current by hypoglycaemia (Burdakov et al. 2006). The authors showed that this current was acid-sensitive and they suggested that it was carried through TASK-like two-pore-domain K+ channels.
In situ hybridization has revealed the presence of mRNA for at least three types of two-pore-domain K+ channels in DVN: TASK-1, TASK-3 and TREK-1 (Talley et al. 2001). Functional acid-sensitive TASK-like channels that are open under resting conditions have been found in DVN (Hopwood & Trapp, 2005). These were characterized by a similar reversal potential (–95 mV), a good fit with the GHK equation, inhibition by 5-HT and sensitivity to Ba2+, but resistance to Zn2+. Since anoxia has been shown to produce an external acidification in the vagal complex (Ballanyi et al. 1996), it might be attractive to speculate that the inhibition of TASK channels by an acidic pH is the underlying mechanism. However, since the same current appears to be inhibited by hypoglycaemia despite hypoglycaemia causing an extracellular alkalinization (Ballanyi et al. 1996), it seems unlikely that pH is involved.
Interestingly, Buckler et al. (2000) demonstrated a K+ current that was inhibited by metabolic inhibition (hypoxia) in rat carotid body type-I cells. Further experiments showed that this hypoxia-induced depolarization could be mimicked by mitochondrial inhibitors, e.g. cyanide (Wyatt & Buckler, 2004). Single-channel recordings revealed that the activity of this oxygen-sensitive K+ channel runs down quickly in inside-out patches, but can be increased by physiological concentrations of ATP (2–5 mM), suggesting that this channel is ATP dependent (Williams & Buckler, 2004). The properties of the channel were consistent with a TASK-like K+ channel.
In the present study, the observation that halothane did not activate, but rather inhibited a K+ current in those cells that had a metabolically inhibited K+ current might suggest that the underlying channel belongs to the halothane-inhibited (THIK) subfamily of two-poredomain K+ channels. In fact, a study investigating a hypoxia-inhibited current in glossopharyngeal neurones suggested that THIK-like channels might account for the O2 sensitivity of these cells (Campanucci et al. 2003).
In conclusion, it seems conceivable that the current blocked by metabolic inhibition in DVN is due to a two-pore-domain K+ channel, and our evidence to date would suggest that it is a THIK-like channel. However, further analysis will be required to demonstrate unequivocally which member of this K+ channel family is responsible.
An increase in conductance induced by metabolic inhibition
Metabolic inhibition caused an increase in whole-cell conductance with a range of reversal potentials between –58 and +10 mV in the majority of DVN and an inward current over the entire voltage range examined in the remaining cells. These two types of responses were observed in both the absence and presence of TTX.
An inward current with no obvious reversal potential was also produced by inhibition of the Na+–K+-ATPase with strophanthidin or ouabain. However, combination of pharmacological inhibition of the Na+–K+-ATPase with mitochondrial inhibition to identify a possible differential current caused a very large inward current instead. Interestingly, a large inward current or depolarization has also been reported for DVN in response to ischaemia (Martin, 1999; Ballanyi et al. 1996; Raupach & Ballanyi, 2004).
In a study of the lateral hypothalamic area with ion-sensitive electrodes, GI neurones hyperpolarized in response to hyperglycaemia and, consistent with increased Na+–K+-ATPase activity, intracellular [Na+] fell and extracellular [K+] rose (Silver & Erecinska, 1998). Experiments in the insulin-secreting cell line HIT-T15 demonstrated that Na+–K+-ATPase activity was altered by changes of ATP concentration between 0.25 and 6 mM. Levels of ATP in neurones are thought to be around 1 mM, therefore it seems feasible that Na+–K+-ATPase activity is altered by physiological changes in intracellular ATP concentration (Niki et al. 1989). A previous study investigated the effects of Na+–K+-ATPase inhibition in the rat auditory thalamus (Senatorov et al. 1997). In agreement with our results, the authors found that strophanthidin and ouabain induced a small inward current (–39.0 pA at the cell resting potential of –72 mV) and this current showed no obvious reversal potential. All this evidence might indicate that inhibition of the Na+–K+-ATPase by metabolic inhibition contributes to the observed inward currents in DVNs.
We hypothesize that the observed large range of reversal potentials for the inward current arises from the combination of the opening of an ion channel and the current caused by inhibition of the Na+–K+-ATPase. Consistent with this hypothesis, the recordings of metabolic inhibition under TTX, where voltage ramps were applied every 30 s, revealed reversal potentials of –30 to –50 mV for the increase in conductance only, followed by an inward current with no obvious reversal potential, which would be consistent with inhibition of the Na+–K+-ATPase (Fig. 6C).
At present it is unclear what conductance underlies the inward current with a reversal potential around –40 mV. Such a reversal potential is similar to that observed for Cl–-mediated GABA currents in this study, and an activation of a Cl– channel has been proposed to mediate the equivalent inward current in hippocampal neurones (Krnjevic & Xu, 1990). However, our recordings with ‘high-Cl–’ internal solution failed to shift the reversal potential of the inward current to more positive potentials. It therefore seems unlikely that a Cl– channel mediates this current in DVN.
Implications for mechanisms of glucosensing
Our results strongly suggest that mitochondrial inhibition and hypoglycaemia interfere with the same pathways, i.e. activation of KATP channels, and inhibition of the Na+–K+-ATPase and of two-pore-domain K+ channels. Whilst the Na+–K+-ATPase is ubiquitously expressed, only a subpopulation of cells has functional KATP channels. From this it follows that depolarization is the default response to metabolic inhibition, and hyperpolarization is observed in that subset of cells that express KATP channels. Whilst cyanide and azide have an immediate effect on metabolism in all neurones, the impact of glucose removal is only seen immediately in glucosensing neurones. We propose that the defining feature of glucosensing neurones is the expression of glucokinase (Dunn-Meynell et al. 2002; Balfour et al. 2006; Kang et al. 2006). Their dependence on glucokinase means that these cells utilize glucose at low concentrations less efficiently than other cell populations and therefore reach the threshold for an electrical response to glucose removal earlier than other neurones. This electrical response is then the same as in other neurones, and the two subtypes, GI and GE cells, are defined by the absence versus presence of KATP channels.
The fact that the inward currents described in this study can be elicited by various different metabolic inhibitors suggests that a common factor leads to the development of the current under these conditions. Particularly, when considering the pronounced difference in the delay to onset of the inward current in response to the removal of glucose from the bath solution between the two subpopulations of DVN, one might hypothesize that differences in cellular energy metabolism cause the different responses. It seems rather unlikely that these two populations are created by access barriers for glucose within the slices, because no distinct populations are observed for the response to cyanide or azide. The metabolism hypothesis is further supported by our previous finding that only cells that respond quickly to hypoglycaemia express glucokinase (Balfour et al. 2006). Glucokinase is obligatory for the response of pancreatic -cells to variations in blood glucose levels (Sakura et al. 1998). Finally, the fact that the same inward current was elicited by cyanide or azide suggests that the interference of these drugs with cellular energy metabolism leads to the observed inward current, rather than the binding of these substances to specific extracellular receptors.
Glucosensors in the ventromedial hypothalamus are likely to rely on glucose metabolism to generate their electrical response, too. Lactate stimulates GE neurones (Yang et al. 1999; Song & Routh, 2005) and inhibits GI neurones in the ventromedial hypothalamus (Yang et al. 1999, 2004; Song & Routh, 2005) and thus can substitute for glucose in vitro. Furthermore, local lactate perfusion into the ventromedial hypothalamus prevents systemic counter-regulatory responses to hypoglycaemia in vivo (Borg et al. 2003). The simplest explanation for these findings is that lactate can be metabolized and used as fuel instead of glucose, thereby overriding the response to hypoglycaemia.
Conclusions
Our results suggest that DVN use the same electrical pathways in their response to mitochondrial inhibition (by cyanide or azide) and to hypoglycaemia. Dorsal vagal neurones responded to metabolic inhibition with a small inward current that was primarily caused by inhibition of an outwardly rectifying K+ channel, or activation of a conductance with a reversal potential of approximately –40 mV and/or inhibition of the Na+–K+-ATPase. A subset of DVN express KATP channels, and opening of this channel by metabolic inhibition overrides the inward current, leading to a net outward current or hyperpolarization. Whilst these are the two universal responses to metabolic inhibition in DVN, our earlier study on glucosensing dorsal vagal neurones (Balfour et al. 2006) suggests that the presence or absence of glucokinase determines whether a DVN is a glucosensor (fast response to glucose removal) or not (slow response to glucose removal). We suggest that DVNs can be separated into subtypes and classified using these criteria and we hypothesize that this classification can be applied to most central neurones.
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References |
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Ashford ML, Boden PR & Treherne JM (1990). Glucose-induced excitation of hypothalamic neurones is mediated by ATP-sensitive K+ channels. Pflugers Arch 415, 479–483.[CrossRef][Medline]
Balfour RH, Hansen AM & Trapp S (2006). Neuronal responses to transient hypoglycaemia in the dorsal vagal complex of the rat brainstem. J Physiol 570, 469–484.
Ballanyi K, Doutheil J & Brockhaus J (1996). Membrane potentials and microenvironment of rat dorsal vagal cells in vitro during energy depletion. J Physiol 495, 769–784.
Ballanyi K & Kulik A (1998). Intracellular Ca2+ during metabolic activation of KATP channels in spontaneously active dorsal vagal neurons in medullary slices. Eur J Neurosci 10, 2574–2585.[CrossRef][Medline]
Borg MA, Tamborlane WV, Shulman GI & Sherwin RS (2003). Local lactate perfusion of the ventromedial hypothalamus suppresses hypoglycemic counterregulation. Diabetes 52, 663–666.
Buckler KJ, Williams BA & Honore E (2000). An oxygen-, acid- and anaesthetic-sensitive TASK-like background potassium channel in rat arterial chemoreceptor cells. J Physiol 525, 135–142.
Burdakov D, Jensen LT, Alexopoulos H, Williams RH, Fearon IM, O'Kelly I, Gerasimenko O, Fugger L & Verkhratsky A (2006). Tandem-pore K+ channels mediate inhibition of orexin neurons by glucose. Neuron 50, 711–722.[CrossRef][Medline]
Campanucci VA, Fearon IM & Nurse CA (2003). A novel O2-sensing mechanism in rat glossopharyngeal neurones mediated by a halothane-inhibitable background K+ conductance. J Physiol 548, 731–743.
Cowan AI & Martin RL (1992). Ionic basis of membrane potential changes induced by anoxia in rat dorsal vagal motoneurones. J Physiol 455, 89–109.
Cowan AI & Martin RL (1995). Simultaneous measurement of pH and membrane potential in rat dorsal vagal motoneurons during normoxia and hypoxia: a comparison in bicarbonate and HEPES buffers. J Neurophysiol 74, 2713–2721.
Dallaporta M, Perrin J & Orsini JC (2000). Involvement of adenosine triphosphate-sensitive K+ channels in glucose-sensing in the rat solitary tract nucleus. Neurosci Lett 278, 77–80.[CrossRef][Medline]
Dunn-Meynell AA, Routh VH, Kang L, Gaspers L & Levin BE (2002). Glucokinase is the likely mediator of glucosensing in both glucose-excited and glucose-inhibited central neurons. Diabetes 51, 2056–2065.
Erdemli G, Xu YZ & Krnjevic K (1998). Potassium conductance causing hyperpolarization of CA1 hippocampal neurons during hypoxia. J Neurophysiol 80, 2378–2390.
Ferreira M Jr, Browning KN, Sahibzada N, Verbalis JG, Gillis RA & Travagli RA (2001). Glucose effects on gastric motility and tone evoked from the rat dorsal vagal complex. J Physiol 536, 141–152.
Hille B (2001). Ion Channals of Excitable Membranes. Sinauer Associates, Inc., Sunderland.
Hopwood SE & Trapp S (2005). TASK-like K+ channels mediate effects of 5-HT and extracellular pH in rat dorsal vagal neurones in vitro. J Physiol 568, 145–154.
Kang L, Dunn-Meynell AA, Routh VH, Gaspers LD, Nagata Y, Nishimura T, Eiki J, Zhang BB & Levin BE (2006). Glucokinase is a critical regulator of ventromedial hypothalamic neuronal glucosensing. Diabetes 55, 412–420.
Krnjevic K & Xu Y (1990). Mechanisms underlying anoxic hyperpolarization of hippocampal neurons. Can J Physiol Pharmacol 68, 1609–1613.[Medline]
Laughton WB & Powley TL (1987). Localization of efferent function in the dorsal motor nucleus of the vagus. Am J Physiol Regul Integr Comp Physiol 252, R13–R25.
Liss B & Roeper J (2001). Molecular physiology of neuronal K-ATP channels. Mol Membr Biol 18, 117–127.[CrossRef][Medline]
Martin RL (1999). Block of rapid depolarization induced by in vitro energy depletion of rat dorsal vagal motoneurones. J Physiol 519, 131–141.
Miki T, Liss B, Minami K, Shiuchi T, Saraya A, Kashima Y, Horiuchi M, Ashcroft F, Minokoshi Y, Roeper J & Seino S (2001). ATP-sensitive K+ channels in the hypothalamus are essential for the maintenance of glucose homeostasis. Nat Neurosci 4, 507–512.[Medline]
Mizuno Y & Oomura Y (1984). Glucose responding neurons in the nucleus tractus solitarius of the rat: in vitro study. Brain Res 307, 109–116.[CrossRef][Medline]
Mobbs CV, Kow LM & Yang XJ (2001). Brain glucose-sensing mechanisms: ubiquitous silencing by aglycemia vs. hypothalamic neuroendocrine responses. Am J Physiol Endocrinol Metab 281, E649–E654.
Niki I, Ashcroft FM & Ashcroft SJ (1989). The dependence on intracellular ATP concentration of ATP-sensitive K-channels and of Na,K-ATPase in intact HIT-T15 -cells. FEBS Lett 257, 361–364.[CrossRef][Medline]
Oomura Y, Ono T, Ooyama H & Wayner MJ (1969). Glucose and osmosensitive neurones of the rat hypothalamus. Nature 222, 282–284.
Oomura Y, Ooyama H, Sugimori M, Nakamura T & Yamada Y (1974). Glucose inhibition of the glucose-sensitive neurone in the rat lateral hypothalamus. Nature 247, 284–286.
Powley TL (2000). Vagal circuitry mediating cephalic-phase responses to food. Appetite 34, 184–188.[CrossRef][Medline]
Raupach T & Ballanyi K (2004). Intracellular pH and KATP channel activity in dorsal vagal neurons of juvenile rats in situ during metabolic disturbances. Brain Res 1017, 137–145.[CrossRef][Medline]
Ritter RC, Slusser PG & Stone S (1981). Glucoreceptors controlling feeding and blood glucose: location in the hindbrain. Science 213, 451–452.
Routh VH (2002). Glucose-sensing neurons: are they physiologically relevant? Physiol Behav 76, 403–413.[CrossRef][Medline]
Sakura H, Ashcroft SJ, Terauchi Y, Kadowaki T & Ashcroft FM (1998). Glucose modulation of ATP-sensitive K-currents in wild-type, homozygous and heterozygous glucokinase knock-out mice. Diabetologia 41, 654–659.[CrossRef][Medline]
Senatorov VV, Mooney D & Hu B (1997). The electrogenic effects of Na+-K+-ATPase in rat auditory thalamus. J Physiol 502, 375–385.
Silver IA & Erecinska M (1998). Glucose-induced intracellular ion changes in sugar-sensitive hypothalamic neurons. J Neurophysiol 79, 1733–1745.
Song Z, Levin BE, McArdle JJ, Bakhos N & Routh VH (2001). Convergence of pre- and postsynaptic influences on glucosensing neurons in the ventromedial hypothalamic nucleus. Diabetes 50, 2673–2681.
Song Z & Routh VH (2005). Differential effects of glucose and lactate on glucosensing neurons in the ventromedial hypothalamic nucleus. Diabetes 54, 15–22.
Spanswick D, Smith MA, Groppi VE, Logan SD & Ashford ML (1997). Leptin inhibits hypothalamic neurons by activation of ATP-sensitive potassium channels. Nature 390, 521–525.
Spanswick D, Smith MA, Mirshamsi S, Routh VH & Ashford ML (2000). Insulin activates ATP-sensitive K+ channels in hypothalamic neurons of lean, but not obese rats. Nat Neurosci 3, 757–758.[CrossRef][Medline]
Talley EM, Solorzano G, Lei Q, Kim D & Bayliss DA (2001). CNS distribution of members of the two-pore-domain (KCNK) potassium channel family. J Neurosci 21, 7491–7505.
Thomas RC (1972). Electrogenic sodium pump in nerve and muscle cells. Physiol Rev 52, 563–594.
Trapp S & Ballanyi K (1995). KATP channel mediation of anoxia-induced outward current in rat dorsal vagal neurons in vitro. J Physiol 487, 37–50.
Travagli RA, Hermann GE, Browning KN & Rogers RC (2006). Brainstem circuits regulating gastric function. Annu Rev Physiol 68, 279–305.[CrossRef][Medline]
Williams BA & Buckler KJ (2004). Biophysical properties and metabolic regulation of a TASK-like potassium channel in rat carotid body type 1 cells. Am J Physiol Lung Cell Mol Physiol 286, L221–L230.
Wu X, Gao J, Yan J, Owyang C & Li Y (2004). Hypothalamus-brain stem circuitry responsible for vagal efferent signaling to the pancreas evoked by hypoglycemia in rat. J Neurophysiol 91, 1734–1747.
Wyatt CN & Buckler KJ (2004). The effect of mitochondrial inhibitors on membrane currents in isolated neonatal rat carotid body type I cells. J Physiol 556, 175–191.
Yamada K, Ji JJ, Yuan H, Miki T, Sato S, Horimoto N, Shimizu T, Seino S & Inagaki N (2001). Protective role of ATP-sensitive potassium channels in hypoxia-induced generalized seizure. Science 292, 1543–1546.
Yang XJ, Kow LM, Funabashi T & Mobbs CV (1999). Hypothalamic glucose sensor: similarities to and differences from pancreatic -cell mechanisms. Diabetes 48, 1763–1772.[Abstract]
Yang XJ, Kow LM, Pfaff DW & Mobbs CV (2004). Metabolic pathways that mediate inhibition of hypothalamic neurons by glucose. Diabetes 53, 67–73.
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). Other cells developed an increase in conductance with a wide range of reversal potentials (
). Reversal potentials obtained in the presence of 0.5 µM TTX are given in separate columns indicated by arrows. Cells that exhibited a parallel, depolarizing shift are not represented. Ba, example of a DVN in which 1 mM cyanide inhibited a current with a reversal potential close to EK. Bb, I–V relationships of the current inhibited by cyanide were well fitted by the GHK equation (dashed line). Ca, I–V relationships, in the presence and absence of 1 mM halothane, from a cell in which 1 mM cyanide inhibited a K+ current. Cb, the halothane-sensitive current (difference between I–V relationships shown in Ca) is well described by the GHK equation (dashed line).