Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Cell Mol Life Sci. Author manuscript; available in PMC 2010 April 13.
Published in final edited form as:
PMCID: PMC2854026

General anaesthetic actions on ligand-gated ion channels


The molecular mechanisms of general anaesthetics have remained largely obscure since their introduction into clinical practice just over 150 years ago. This review describes the actions of general anaesthetics on mammalian neurotransmitter-gated ion channels. As a result of research during the last several decades, ligand-gated ion channels have emerged as promising molecular targets for the central nervous system effects of general anaesthetics. The last 10 years have witnessed an explosion of studies of anaesthetic modulation of recombinant ligand-gated ion channels, including recent studies which utilize chimeric and mutated receptors to identify regions of ligand-gated ion channels important for the actions of general anaesthetics. Exciting future directions include structural biology and gene-targeting approaches to further the understanding of general anaesthetic molecular mechanisms.

Keywords: General anaesthesia, ligand-gated ion channels, GABA, glutamine, acetylcholine, glycine, serotonin, electrophysiology


Since their introduction into clinical practice just 150 years ago, general anaesthetics have become some of the most widely used and important therapeutic agents. However, despite over a century of research, the molecular mechanisms of action for general anaesthetics in the central nervous system (CNS) remain elusive. As a result of research during the last several decades, the ligand-gated ion channels have emerged as promising molecular targets to mediate the CNS effects of general anaesthetics. In this review, we aim to describe the actions of general anaesthetics on mammalian neurotransmitter-gated ion channels.

We will begin by summarizing the history of general anaesthesia and the chemical classes of general anaesthetics and then provide background on the physiology and pharmacology of ligand-gated ion channels. We will briefly look at experimental methodology and review the pharmacological criteria which can help define which proteins represent plausible molecular targets for general anaesthetics. We will then describe the actions of general anaesthetics on the ligand-gated ion channels. The last 10 years have witnessed an explosion of studies of anaesthetic modulation of ligand-gated ion channels, and we will focus in particular on recent studies which utilize recombinant chimeric and mutated receptors to identify regions of ligand-gated ion channels important for the modulatory actions of general anaesthetics. Lastly, we will discuss future directions in this area of research.

What is a general anaesthetic?

General anaesthetics include a startling range of structurally diverse molecules that can be roughly, and somewhat arbitrarily, divided into volatile anaesthetics, anaesthetic gases, alcohols and intravenous anaesthetics (fig. 1). A surprisingly elusive question is what defines a general anaesthetic, since anaesthesia is a behavioral state easily recognized but difficult to describe precisely. Depending on the clinical procedure, effective anaesthesia requires varying degrees of immobility, amnesia, unconsciousness/hypnosis, analgesia, muscle relaxation and depression of autonomic reflexes [1]. No general anaesthetic provides all of these effects, although immobility, unconsciousness/hypnosis and amnesia are behavioral hallmarks of most general anaesthetics [2]. Modern anaesthesia involves not only general anaesthetics but also the use of multiple supplemental agents including analgesics (e.g. opiates) and neuromuscular blockers.

Figure 1
Chemical structures of selected general anaesthetics. Nitrous oxide is a molecule that exists in three resonating linear structures (although often misdrawn as a cyclical structure). For simplicity, we have drawn only one of the resonance forms.

Specific versus nonspecific mechanisms of anaesthetic action

The observation that a spectrum of chemically dissimilar agents produce general anaesthesia greatly influenced the thinking of early investigators seeking to explain anaesthetic mechanisms of action. A landmark series of experiments reported independently by Hans Meyer and Charles Ernest Overton around the turn of the century determined that the potencies of general anaesthetic molecules correlated well with their water/oil partition coefficients [35]. The so-called Meyer-Overton correlation was later extended to embrace the concept that certain molecules produce general anaesthesia by a nonspecific mechanism. The traditional view since the time of Meyer and Overton has been that general anaesthetics exert their primary effects by dissolving in cell membranes, particularly in the CNS [68]. The presence of general anaesthetic molecules is thought to perturb the structural and dynamic properties of the lipid bilayer (a ‘nonspecific’ action), so that the function of crucial but unspecified membrane proteins is affected. ‘Specific’ actions of anaesthetics generally refer to direct effects of general anaesthetics on known protein molecules which result in reversible alterations in the function of the protein (e.g. increased probability of opening of an ion channel) [9].

Research within the last several decades has demonstrated numerous inconsistencies between experimental observations and nonspecific theories of general anaesthesia [912]. The main problems include the following [1, 13]: (i) Some chemical compounds are predicted by nonspecific theories to be anaesthetics but, in fact, do not produce anaesthesia; (ii) nonspecific theories of anaesthesia cannot account for the stereoselectivity demonstrated by some anaesthetic isomers; and (iii) anaesthetic effects on lipids (such as alterations in membrane bilayer fluidity), when measured experimentally, are often negligible at clinically relevant concentrations, and are easily reproduced by very small increases in ambient temperature. In contrast, decreases in body temperature mimic the behavioral effects of general anaesthetics [1315]. Despite the numerous inconsistencies between the experimental evidence and nonspecific theories of anaesthesia, there have been attempts in the last decade to present modified nonspecific theories. The interested reader is best referred to some of the more recent experimental investigations and review articles in this area [1621]. Some prescient investigators recognized a number of decades ago that anaesthetics may act instead on specific targets. For example, Sir John Eccles and colleagues studied spinal synaptic reflexes in animals under pentobarbitone anaesthesia [22, 23] and raised the possibility of anaesthetic actions at neurotransmitter receptors important in synaptic transmission.

Ligand-gated ion channels

This review summarizes recent progress in the understanding of general anaesthetic actions on receptor proteins important in synaptic transmission in the CNS. A number of excellent reviews over the last decade have summarized work on the actions of general anaesthetics on receptor proteins in the CNS [1, 13, 2437]. We aim here to expand and update these prior reviews, with particular reference to recent studies documenting general anaesthetic actions on recombinant ligand-gated ion channels. Ligand-gated ion channels are certainly not the only possible molecular targets for general anaesthetics; other neuronal proteins such as voltage-gated ion channels and G-protein-coupled receptors may also play a role in the overall behavioral spectrum of action of general anaesthetics. However, extensive research has arrived at an almost universal consensus; voltage-gated ion channels are, in general, relatively insensitive to clinically relevant concentrations of general anaesthetics [13]. Detailed studies of general anaesthetic actions on G-protein-coupled receptors are scarce, and it can be difficult to distinguish effects on the receptor per se versus general anaesthetic perturbations of second messengers or effector molecules such as protein kinases and phospholipases.

The ligand-gated ion channels have emerged as strong candidates as molecular mediators of the CNS effects of general anaesthetics [13, 26, 27]. The ligand-gated ion channels include the γ aminobutyric acid type A (GABAA), glycine, serotonin-3 (5-HT3) and nicotinic acetylcholine (ACh) receptors, along with the α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA)-, kainate- and NMDA-sensitive subtypes of ionotropic glutamate receptors (note: γ aminobutyric acid (GABA), glutamate, 5-HT and ACh also act on ‘slow’ neurotransmitter receptors, e.g. GABAB, muscarinic acetylcholine and metabotropic glutamate receptors, which are coupled to second messenger systems). GABAA, glycine, 5-HT3 and nicotinic ACh receptors form part of an evolutionarily related ligand-gated ion channel gene superfamily [38]. Ionotropic glutamate receptors were originally thought to be part of this superfamily but are now thought to belong to a distinct ion channel class (see below). All members of the ligand-gated ion channel superfamily appear to have the basic subunit topology diagrammed in figure 2, with a large N-terminal extracellular domain, four putative membrane-spanning regions (TM1–TM4), a heterogeneous intracellular loop between TM3 and TM4, and a short extracellular C-terminal domain. Residues within the extracellular N-terminal domain form the agonist binding domains [3941], whereas amino acid residues within TM2 line the ion channel pore [42, 43] (see figs 2 and and3).3). Native receptors are composed of pentameric arrangements of individual receptor subunits [44, 45] (see fig. 2).

Figure 2
Illustration of the general subunit topology and pentameric structure of receptors from the ligand-gated ion channel superfamily (nicotinic ACh, GABAA, GABAC ρ, glycine and 5-HT3 receptors). Basic features of ligand-gated ion channel subunit topology ...
Figure 3
Location of amino acid residues within TM2 and TM3 of (a) human GABAA α1 [280] and (b) human GABAA β2 [281] receptor subunits that are critical for general anaesthetic modulation or block by the noncompetitive antagonists picrotoxinin ...

GABAA and glycine receptors

GABAA and glycine receptors are chloride-selective ion channels. These are generally considered to be inhibitory neurotransmitter receptors, since in most cells, opening of chloride channels results in membrane hyperpolarization and/or stabilization of the membrane potential away from the threshold for firing action potentials [46]. GABA and glycine are the primary fast inhibitory neurotransmitters in the CNS, with glycine abundant in the spinal cord and brainstem [40, 47] and GABA predominant in higher brain regions [46]. It has been estimated that one-third of all synapses in the CNS are GABA-ergic [48].

Subunit heterogeneity creates extensive diversity among the inhibitory ligand-gated ion channels. Multiple subunits have been cloned for GABAA1–6, β1–4, γ1–4, δ, ε and π) [4956] and glycine (α1–4, β) [40, 47, 57, 58] receptors. GABAA receptors in vivo predominantly consist of α, β and γ subunits with a proposed stoichiometry of 2α: 2β: 1γ [59, 60] (see fig. 2). The existence of six α-subunit isoforms enables considerable anatomical and functional diversity of GABAA receptors [6163]. In particular, the α-subunit isoform may influence agonist potency [64, 65], agonist efficacy [66], regulation by benzodiazepines [67] and channel kinetics [68, 69]. The most common neuronal subunit combination is α1β2γ2 [52, 56]. GABAA receptors are blocked competitively by bicuculline and noncompetitively by picrotoxinin and Zn2+ [39] (see fig. 3).

Strychnine-sensitive glycine receptors in vivo consist of both α homomers and αβ heteromers, with a switch from homomeric to heteromeric receptors occurring during development [40, 57, 58]. The best-described role for glycine receptors is in Renshaw cell inhibition of motor neurones in the spinal cord; however, glycine receptors are also widely expressed in the brainstem and throughout higher regions of the neuraxis [57, 58].

GABAC receptors are formed from ρ subunits (ρ1–3) [7072]. GABAC receptors show greatest expression in the retina but are also found in other areas of the brain [73]. The designation of ‘GABAC’ for ρ subunits, while potentially confusing [56], follows from their extensive pharmacological differences from GABAA receptors, including insensitivity to the classical GABAA competitive antagonist bicuculline [7072].

Nicotinic acetylcholine receptors

Nicotinic ACh receptors may be divided into two main groups: the ‘muscle’ subtype expressed in skeletal muscle [74, 75] and in the electroplaque of Torpedo [76], and ‘neuronal’ nicotinic ACh receptors found throughout the CNS and at autonomic ganglia [41, 7779]. These receptors contain a nonselective cation channel. Multiple subunit isoforms have been described for muscle (α1, β, γ, δ, and ε) and neuronal (α2–9, β2–4) nicotinic ACh receptors [41, 75, 79, 80]. The muscle nicotinic ACh receptors mediate synaptic excitation at the neuromuscular junction [74, 75]. The physiological roles of neuronal nicotinic ACh receptors are currently an area of intense inquiry. These receptors seem likely to participate in nicotine addiction, and may perhaps be involved in neurological and psychiatric disorders, in addition to their more traditional role in the function of the sympathetic and parasympathetic nervous systems [41, 7779]. Neuronal nicotinic ACh receptors occur at both pre- and postsynaptic loci in the CNS. Presynaptic nicotinic ACh receptors modulate the release of neurotransmitters such as GABA, 5-HT, dopamine, norepinephrine, glutamate and ACh [81]. The existence of postsynaptic neuronal nicotinic ACh receptors at sympathetic ganglia has been known for many decades [79]; more recent research has also demonstrated functional postsynaptic neuronal nicotinic ACh receptors in the CNS [82].

5-HT3 receptors

5-HT3 receptors are ligand-gated cation channels that are expressed in a number of central and peripheral nervous system areas, including the hippocampus, brainstem, dorsal root ganglia (cell bodies of sensory neurones) and vagal axons [83]. 5-HT3 receptors are expressed efficiently as homomers in heterologous expression systems such as Xenopus oocytes, but there is evidence that 5-HT3 receptors in vivo may be heteromeric, indicating that additional subunits or splice variants have yet to be characterized [84]. The most unambiguous physiological role for 5-HT3 receptors in humans is in the medullary circuitry subserving the vomiting reflex, which is consistent with robust expression of the 5-HT3 receptor in the nucleus tractus solitarius and area postrema [83, 85]. 5-HT3 receptor antagonists such as ondansetron are used clinically to prevent nausea and vomiting [86]. Activation of 5-HT3 receptors also modulates extracellular dopamine levels in the nucleus accumbens, and this may be involved in the rewarding properties of certain drugs of abuse [87]. 5-HT3 receptors may also play a role in nociception [85]; for example, some nociceptive primary afferents express 5-HT3 receptors, and activation of these receptors facilitates the response of some dorsal horn neurones to noxious stimuli [88].

Ionotropic glutamate receptors

The ionotropic glutamate receptors consist of the AMPA-,NMDA-and kainate-sensitive subtypes. Glutamate receptors were originally assumed to have a membrane topology and pentameric structure similar to that proposed for the original ligand-gated ion channel superfamily. More recent work suggests instead a subunit topology that includes a reentrant loop forming the ion channel pore [8992] analogous to the reentrant pore loop associated with the S5–S6 ‘signature sequence’ of the voltage-gated potassium channels [93]. Tetrameric structures have been proposed for AMPA [94] and NMDA receptor [95] complexes. Once again, the proposed tetrameric structure is much more similar to the voltage-gated potassium channels [96] than to the members of the ligand-gated ion channel gene superfamily discussed above.

NMDA receptors (NR1, NR2A-2D,NR3A)[97100] are ligand-gated cation channels with several unusual features including, most notably, high Ca2+ permeability [101] and strong voltage-dependent block by Mg2+ ions [102104]. NMDA receptors have attracted a great deal of attention due to their involvement in the induction of long-term potentiation in the CA1 subfield of the hippocampus and other areas of the cerebral cortex [105107]. NMDA receptors are proposed to be involved in learning and memory [108, 109]. NMDA receptors are also implicated in certain forms of neurotoxicity and in the etiology of several neurodegenerative disorders [110113]. The pharmacology of the NMDA receptors has been extensively characterized, and a number of substances modify NMDA receptor function, including the coagonist glycine [114, 115], polyamines [116, 117], Zn2+ [118, 119], protons [120, 121], fatty acids [122, 123] and oxidizing/reducing agents [124].

AMPA receptors, represented in neurones by combinations of the gene products GluR1–4, appear to serve as the major fast excitatory neurotransmitter receptors at most synapses in the CNS [113, 125]. Synaptic AMPA receptors respond to glutamate quickly and transiently, due to diffusion and rapid removal of glutamate from the synaptic cleft, in addition to fast receptor desensitization [126]. AMPA receptors are therefore ideally suited for their role in excitatory transmission on a millisecond timescale. The physiological roles of kainate glutamate receptors (GluR5–7, KA1, KA2) are less clear at present, even though kainate receptors are widely distributed throughout the brain and spinal cord. Recent work provides strong evidence for the synaptic activation of kainate receptors at both pre- and postsynaptic sites [127, 128]. The study of AMPA and kainate receptors in the CNS was hampered for a long time by a lack of selective antagonists. Recent development of selective AMPA receptor antagonists has remedied this problem to some extent [128, 129]. An experimental obstacle to study of some of the kainate and AMPA receptors is desensitization on the millisecond to submillisecond timescale [130132]. This complicates the interpretation of many studies, for example those employing heterologous expression of kainate and AMPA receptors in Xenopus oocytes, in which agonist may be applied for seconds to minutes.

Pharmacological criteria that a candidate receptor must meet to be considered as a reasonable general anaesthetic target

Before discussing the actions of specific agents on ligand-gated ion channels, it is worthwhile to define specific criteria that an anaesthetic target (receptor protein or otherwise) must fulfill in order to qualify as a candidate in mediating the behavioral actions of the general anaesthetics [1, 13].

  1. The general anaesthetic must alter the function of the receptor at clinically relevant concentrations.
  2. The receptor must be expressed in the appropriate anatomical locations to mediate the specific behavioral effects of the anaesthetic.
  3. If an anaesthetic molecule shows stereoselective effects in vivo, these should be mirrored by the in vitro actions at the receptor.
  4. The hydrophobicity of a compound within a homologous series of anaesthetics should correlate with the in vivo anaesthetic potency and that at the target receptor.

The general anaesthetic must alter the function of the receptor at clinically relevant concentrations

What is the ‘clinically relevant concentration’ for a general anaesthetic? For an inhaled anaesthetic such as isoflurane, 1 minimum alveolar concentration (MAC) conventionally refers to the concentration of inhaled anaesthetic that produces immobility in 50% of animals studied [133, 134]. Immobility, a lack of purposeful response to a noxious stimulus, represents an easily determined endpoint across a large variety of different animal species. The use of immobility as an experimental endpoint is helpful in that, for most general anaesthetics, anaesthetic concentrations two- to four-fold above the EC50 (concentration of a compound which produces 50% of the maximal effect) for producing immobility are invariably lethal [13]. The anaesthetic concentrations that produce significant inhibition of cognitive functions and cortical activity, assessed using EEG-derived indicators, are lower than those required for producing immobility [135137]. Thus, anaesthetic concentrations several-fold greater than those that produce immobility define the upper boundary of the concentration range that is clinically relevant. For a target to have any relevance for anaesthesia, it must at least be sensitive to sublethal but immobilizing concentrations of anaesthetics. This issue of relevant concentrations alone poses a severe challenge to the plausibility of ‘lipid’ theories of anaesthetic action, since ‘nonspecific’ effects of general anaesthetics (e.g., disruption of lipid bilayer fluidity) appear to be negligible at clinically relevant concentrations [1315].

While the issue of relevant concentrations is obviously of paramount importance to molecular studies of general anaesthetics, the physicochemical and pharmacokinetic properties of the various anaesthetic drugs pose some obstacles to the determination of relevant concentrations. We will therefore outline the basic issues involved in the determination of accurate clinically relevant anaesthetic concentrations. This will provide a background to our later discussion of those ligand-gated ion channels that are modulated by clinically relevant concentrations of general anaesthetics.

Volatile anaesthetic potency is usually quantified in terms of MAC [133, 134]. MAC values (often expressed in the operating room in terms of percentage of anaesthetic gas by volume) can be converted to ‘aqueous’ MAC equivalent concentrations’ by use of the appropriate water/gas (or blood/gas) partition coefficients [28, 138]. This provides an estimate for the concentration of anaesthetic in the blood that is in equilibrium with the inspired partial pressure of anaesthetic in the gas phase. Franks and Lieb [28, 138] have thoroughly discussed the conversion of MAC values to aqueous equivalents, including the nettlesome issue of experimental temperature [138, 139]. Aqueous MAC equivalents are often used as guides for in vitro experiments which involve the study of volatile anaesthetics in aqueous solution [13, 28, 138].

The issue of clinically relevant concentrations for the intravenous anaesthetics and the alcohols in mammals is considerably more complicated because of pharmacokinetic aspects of these drugs and the difficulty of ascertaining steady-state drug concentrations in the brain [13]. In some cases (e.g. for propofol and the barbiturates), detailed pharmacokinetic studies have addressed these issues, and reasonable free anaesthetic concentrations in brain can be estimated [13]. In other cases (e.g. ketamine and the steroid anaesthetic alphaxalone), only total anaesthetic concentrations in blood are known, thus invariably underestimating anaesthetic potency in the brain of this class of anaesthetics, often by as much as one to two orders of magnitude [140, 141].

Table 1 lists concentrations of general anaesthetics that represent the EC50 value for producing immobility in a variety of animal species. There is a growing database of studies that determine anaesthetic concentrations needed to produce other anaesthetic endpoints involving higher cortical functions [136, 137, 142]. However, such data are not yet available for all anaesthetics. In some cases (see table 1 and accompanying legend), no mammalian data are available, or the mammalian data are likely to be incorrect, due to significant pharmacokinetic issues. In these cases, we have reported the values for tadpoles, in which pharmacokinetic impediments are considerably attenuated [13, 143].

Table 1
Clinically relevant concentrations of general anaesthetics.

The receptor must be expressed in the appropriate anatomical locations to mediate the specific behavioral effects of the anaesthetic

This is a more difficult issue to discuss, since there is considerable debate about precisely which synaptic circuits are responsible for the various behavioral functions perturbed by general anaesthetics. The immobility produced by general anaesthetics, perhaps not surprisingly, appears to involve depression of spinal reflex pathways, since it is independent of drug actions in the brain [144146]. Receptors such as GABAA and AMPA receptors are promising general anaesthetic targets due to their ubiquitous distribution and essential physiological roles as the major fast transmitters of the CNS. However, given the uncertainty concerning the exact anatomy of the synapses that are disrupted to produce the constellation of behavioral effects seen during general anaesthesia, receptors with more limited distribution (e.g. 5-HT3 receptors) may certainly play major roles as molecular mediators of the general anaesthetic state.

If an anaesthetic molecule shows stereoselective effects in vivo, these should be mirrored by the in vitro actions at the receptor

Stereoselectivity represents an important test for the relevance of a putative anaesthetic target [13, 147]. A number of general anaesthetic molecules possess a chiral carbon atom, and some pairs of stereoisomers exert different anaesthetic potencies in vivo. Stereoselectivity for producing immobility has been documented for the isomers of etomidate [15, 148] (see fig. 4), the barbiturates [149], isoflurane [150, 151] (although see [152]), ketamine [153, 154] and steroid anaesthetics [155]. The potency differences are greatest for stereoisomers of etomidate and ketamine (greater than 10-fold), with smaller potency differences (sometimes only 2-fold or less) seen for other anaesthetic isomers. The formulation of these anaesthetics is usually based on the racemic mixture due to the difficulty of separating enantiomers in large quantities (an exception is etomidate, which is prepared by a chiral synthesis [148]). Production of pure enantiomers perhaps would improve the clinical profile for some general anaesthetics [156], although cost considerations probably preclude such an outcome.

Figure 4
The selectivity of etomidate optical isomers for producing general anaesthesia in tadpoles mirrors the selectivity for potentiation of GABAA receptor function. The main graph illustrates the concentration-response curves for immobility produced by etomidate ...

General anaesthetic stereoselectivity poses the most severe challenge yet to the ‘traditional’ lipid theories of anaesthetic action. The optical isomers of isoflurane [157] and etomidate [15], despite significant differences in their in vivo potency (see fig. 4), behave identically with respect to their ability to disorder lipid bilayers. In contrast, stereoselectivity supports the plausibility of the GABAA receptor as a target in mediating the actions of etomidate [15], barbiturates [158, 159], isoflurane [160, 161], and the steroid anaesthetics [155, 162], since in vivo potency and activity at the GABAA receptor display identical trends. The in vivo stereoselectivity of ketamine stereoisomers is paralleled by the inhibitory action of the isomers at the NMDA receptor [163]. Interestingly, two enantiomers of pentobarbitone display opposing stereoselectivity for inhibition of the muscle nicotinic ACh receptor relative to their in vivo potency [164]. The structure-activity relationships for barbiturate inhibition of the muscle nicotinic ACh receptor also correlate poorly with in vivo potency [165] which effectively eliminates the muscle-type nicotinic ACh receptor as a plausible target for barbiturate action. This is perhaps not surprising, since barbiturates (and, indeed, most other general anaesthetics) do not inhibit neuromuscular transmission to any substantia degree, suggesting little or no functional blockof the muscle nicotinic ACh receptor at anaesthetic concentrations [2, 13]. The probable lack of relevance of muscle nicotinic ACh receptors for the actions of general anaesthetics certainly does not rule out the possibility that general anaesthetic actions on neuronal nicotinic ACh receptors may play a major role in the behavioral actions of general anaesthetics. The muscle and neuronal nicotinic ACh receptors, despite sharing a common agonist, have quite distinct structural and functional properties.

Despite the rewards of studying general anaesthetic stereoisomers, exemplified by the etomidate work outlined above [15] (see fig. 4), the stereoselectivity approach has been underutilized, mainly due to the limited supply and expense of purified stereoisomers [156]. Furthermore, only limited anaesthetic endpoints (mainly immobility) have been assessed for the anaesthetic stereoisomers. It would be quite interesting to know whether the additional neurobiological actions of anaesthetics (e.g. amnesia, analgesia) display similar patterns of stereoselectivity.

The hydrophobicity of a compound within a homologous series of anaesthetics should correlate with the in vivo anaesthetic potency and that at the target receptor

The so-called Meyer-Overton hypothesis, which led to the adoption of the traditional dogma concerning lipid mechanisms of anaesthesia, arose from the fundamental observation that the in vivo potency of general anaesthetics rises in parallel with increasing hydrophobicity of the anaesthetic molecules. This trend is most noticeable with the homologous series of n-alcohols (see table 1c) but also holds true for diverse anaesthetic molecules with oil/water partition coefficients varying over numerous orders of magnitude [35]. General anaesthetic actions at a plausible receptor target should, therefore, exhibit similar trends.

The Meyer-Overton correlation was traditionally interpreted to suggest nonspecific mechanisms of action for general anaesthetics in membrane lipids; however, an alternative explanation is that anaesthetics bind to hydrophobic domains of receptor proteins [9, 13]. For example, amino acid residues of hydrophobic character within the transmembrane domains of ligand-gated ion channels would be likely candidates to interact with general anaesthetics. As will be discussed below, a number of amino acid residues have been identified within ligand-gated ion channels that are critical for the modulatory actions of some general anaesthetics. Many of these amino acid residues are proposed to lie either within a transmembrane domain or else at the membrane interface.

A major problem for traditional theories arose with the discovery of hydrophobic compounds that disobey the Meyer-Overton ‘rule’ [166]. These nonanaesthetics or nonimmobilizers would be expected to partition extensively into the lipid bilayer yet produce no general anaesthetic action. The nonimmobilizers provide additional clues as to which receptor targets might underlie the behavioral actions of general anaesthetics (see below).

Experimental approaches to studying general anaesthetic actions at ligand-gated ion channels

General anaesthetic actions at ligand-gated ion channels have been studied using a variety of methodologies, including protein chemistry, radioligand binding, ion flux studies and electrophysiology [13, 24, 27]. We will focus mainly on electrophysiological studies since these, in general, provide superior time resolution and also offer the possibility of analyzing isolated cells or even single ion channels. The general anaesthetics have properties which limit the utility of other experimental techniques. For example, specific binding of radiolabeled general anaesthetics to ligand-gated ion channels has proven exceedingly difficult to demonstrate due to low affinity and high nonspecific binding to neuronal membranes [13, 24, 27], although allosteric effects of general anaesthetics have been monitored using radioligand binding of drugs to other sites on the ligand-gated ion channels (e.g. [167, 168]). In addition, limited progress has been made in developing anaesthetic congeners useful for photoaffinity labeling or other covalent modification of receptors (although see [169]). These limitations contrast starkly with the studies of other classes of agents at ligand-gated ion channels. For instance, the high-affinity benzodiazepine binding site on the GABAA receptor has been mapped out in some detail due to the ability to perform both specific radioligand binding and photoaffinity labeling [170, 171], which powerfully complements the extensive body of literature on electrophysiological actions of benzodiazepines at GABAA receptors [170].

Another exciting tool in the quest to establish the in vivo significance of a putative anaesthetic target is the use of targeted gene manipulations in mice [172]. A variety of manipulations are possible, including introducing a gene not normally present (transgenic mice), removing an endogenous gene (‘knockout mice’), or replacing an endogenous gene with an altered copy (‘knock-in mice’) [172]. Gene targeting in mice has already been very valuable for elucidating the mechanism of action for some drugs. Knockout of the GABAA γ2 receptor subunit gene resulted in mice which were effectively insensitive to the sedative/hypnotic actions of benzodiazepines such as diazepam [173]. The γ2 subunit gene knockout, in conjunction with the dependence of benzodiazepine modulation of the GABAA receptor on the presence of a γ subunit [174], effectively demonstrates the GABAA receptor as a major target mediating the sedative/hypnotic actions of benzodiazepines. Another gene-targeting experiment in mice involved the replacement of the α2a-adrenoreceptor with a dysfunctional receptor mutant. These knock-in mice failed to show analgesic and sedative responses to α2a-adrenoreceptor agonists such as dexmedetomidine and clonidine [175].

Knockout mice lacking subunit genes for GABAA6, β3, γ2, γ2L) [173, 176179], neuronal nicotinic ACh (α7, β2) [180, 181], AMPA (GluR2) [182], NMDA (NR1, NR2A, NR2C, NR3A) [183186] and kainate receptors (GluR6) [187] have already been created, and the study of such mice has enhanced understanding of the physiological roles of the particular receptor subunit. For example, mice homozygous for a deletion of the GABAA receptor β3 subunit gene exhibit cleft palate, absence seizures, hyperexcitability [177, 188] and some resistance to the immobilizing actions of intravenous and volatile anaesthetics [189].

Actions of general anaesthetics at ligand-gated ion channels

General anaesthetics act as positive or negative allosteric modulators of agonist actions at ligand-gated ion channels. Among the ligand-gated ion channels, there is no known case in which the anaesthetic competes for the same binding site as the endogenous neurotransmitter. The most extensively examined ligand-gated ion channel target for general anaesthetics has been the GABAA receptor [13, 24, 27]. Virtually every general anaesthetic tested enhances the function of the GABAA receptor at clinically relevant concentrations [13, 27, 190] (except for ketamine [191], xenon [192] and possibly nitrous oxide [193195]). General anaesthetic enhancement of GABAA receptor function is evident in single cell electrophysiological experiments as potentiation of a submaximal GABA response (see fig. 5) or, at the synaptic level, as prolongation of inhibitory postsynaptic potentials [196, 197] or currents (see fig. 6) [160, 198200]. Potentiation of submaximal GABA-induced currents remains the most popular assay for electrophysiological experiments since it is easily reproducible and can be used to study native GABAA receptors in dissociated neurones or recombinant receptors expressed in mammalian cell lines or Xenopus oocytes [13, 24, 27].

Figure 5
Specific mutations in TM2 or TM3 of the human GABAA α2 subunit abolish positive allosteric modulation by the volatile anaesthetic isoflurane at GABAA α2β1 receptors. (A) Submaximal GABA currents in wild-type GABAA α2β ...
Figure 6
Both the volatile anaesthetic halothane and the intravenous anaesthetic pentobarbitone prolong inhibitory postsynaptic currents (IPSCs) mediated by GABAA receptors. Data were obtained from whole-cell patch-clamp recordings of rat hippocampal neurones ...

Some anaesthetics, particularly the intravenous agents, open the GABAA receptor chloride channel in the absence of agonist [201214]. This ‘direct activation’ by general anaesthetics involves a binding site completely distinct from that for classical GABAA receptor agonists such as GABA and muscimol [215]. Although direct activation usually occurs at supraclinical concentrations, direct activation effects do sometimes occur at lower concentrations for some anaesthetics (e.g. propofol), suggesting possible clinical relevance. Direct activation by anaesthetics has been observed in other ligand-gated ion channels (e.g. for the anaesthetic isoflurane at the strychinine-sensitive glycine receptor [216]) but is most pronounced at the GABAA receptor.

The cloning of multiple subunit isoforms for the ligand-gated ion channels in the last decade has precipitated an explosion of studies of general anaesthetic actions on recombinant receptors. Table 2 summarizes the electrophysiological effects of general anaesthetics on a range of ligand-gated ion channels studied in neurones or in various expression systems.

Table 2
Modulatory effects of general anaesthetics on ligand-gated ion channels.

A difficult issue to address is how much alteration of receptor function by a general anaesthetic is necessary to produce certain behavioral actions. For example, even though the EC50 or IC50 (concentration of antagonist that reduces the response to a sub-maximal concentration of agonist by 50 %) for alteration of the function of a particular receptor by an anaesthetic may be well outside the clinically relevant range (the upper limit of this range is defined by the anaesthetic concentration that produces immobility in 100% of subjects), the anaesthetic may nevertheless produce slight alteration of receptor function within the clinically relevant concentration range [13]. Thus, in table 2 we have distinguished between complete and relative lack of sensitivity of a particular receptor to clinically relevant anaesthetic concentrations. In order to qualify for inclusion in table 2, a study had to (i) assess the effects of several different anaesthetic concentrations in order to derive an estimate for the EC50 or IC50 (concentration of antagonist that reduces the response to a sub-maximal concentration of agonist by 50%) for modulation and (ii) offer a reasonable certainty of examining a ‘pure’ receptor population. The latter concern is especially acute with the AMPA and kainate subtypes of glutamate receptors, for which there has been until recently a relative dearth of selective agonists and antagonists. Kainate itself activates both AMPA and kainate receptors, and this may confound electrophysiological studies which utilize kainate application to neurones.

The advent of cloning and recombinant expression techniques has greatly accelerated and facilitated attempts to classify ligand-gated ion channel sensitivity to general anaesthetics. Molecular biology techniques may now be used to determine which regions of ligand-gated ion channels are critical for anaesthetic modulation. Sensitivity to general anaesthetics varies considerably, sometimes even among closely related receptors (table 2), and this forms the basis for the use of ‘chimeric’ receptors to isolate regions of a receptor essential for anaesthetic modulation. Chimeric receptors are created by joining together, at the complementary DNA (cDNA) level, complementary fragments of receptor subunits, in which the parental subunits exhibit markedly different anaesthetic pharmacologies. The analysis of chimeric receptors can be used to delimit a region of a receptor essential for general anaesthetic modulation, after which site-directed mutagenesis can be used to identify key residues. Chimeric receptors constructed to date include panels of GABAA/glycine [217], GABAA/GABAC ρ [218], glycine/GABAC ρ [219, 220], neuronal nicotinic ACh/5-HT3 [221, 222] and AMPA GluR3/kainate GluR6 [223] receptors. The most extensive sets of chimeras created and functionally expressed for analysis of anaesthetic modulation are glycine/GABAC ρ [219, 220] and GluR3/GluR6 [223] chimeras.

Several problems may arise in the study of such chimeric receptors, including (i) lack of functional expression (greatly reduced or absent responses to agonist), (ii) chimeric receptor function that differs radically from the constituent parent receptors, and/or (iii) ambiguous pharmacological data. The first problem has substantially limited the utility of GABAA/GABAC [224] and neuronal nicotinic ACh/5-HT3 chimeras [225]; for instance, chimeras formed between the nicotinic ACh α7 subunit and the 5-HT3 receptor show functional expression only when the nicotinic α7 receptor subunit provides the N-terminal half but not vice versa [225]. Lack of functional chimeric receptor responses could potentially be due to protein folding or assembly problems, impaired ion permeation leading to very low single-channel conductance and/or a minuscule probability of opening following agonist binding (i.e. a defect in ion channel gating). Folding and assembly problems probably predominate and seem especially likely to occur when blending heteromeric with homomeric receptors (e.g. GABAA with GABAC receptors). Despite these potential pitfalls, the use of chimeric receptors has already helped to define putative sites of general anaesthetic action on some of the ligand-gated ion channels (see below).

Actions of general anaesthetics at ligand-gated ion channels Volatile anaesthetics and anaesthetic gases

Volatile anaesthetics (e.g. halogenated ethers such as isoflurane and alkanes such as halothane) alter the function of many ligand-gated ion channels at reasonable concentrations. In general, submaximal agonist responses at GABAA, glycine, 5-HT3 and GluR6 receptors are enhanced by volatile anaesthetics, whereas agonist responses at neuronal nicotinic ACh and GluR3 receptors are inhibited (table 2a). The low potency and physicochemical properties of the volatile anaesthetics pose some technical challenges for in vitro experiments [13, 27, 28, 138]. Nevertheless, recent years have witnessed a steady increase in the quality and quantity of careful studies of volatile anaesthetic actions on ligand-gated ion channels.

Considerable progress has been made in identifying amino acid residues within GABAA, glycine and kainate receptors that are critical for volatile anaesthetic potentiation of agonist-induced currents. The use of a panel of glycine α1/GABAC ρ1 chimeric receptors allowed the implication of a 45-amino acid region encompassing TM2 and TM3 of the glycine α1 receptor as both necessary and sufficient for potentiation of agonist-induced currents by the volatile ether enflurane [219]. Extensive site-directed mutagenesis of glycine α1 and GABAA α2 and β1 subunits determined that specific amino acid positions within TM2 and TM3 are also critical for agonist potentiation by isoflurane [206, 219] (see figs 3 and and5),5), n-alcohols (including ethanol) [219, 220, 226] and trichloroethanol [227] (see fig. 3). Agonist potentiation by propofol [206] and etomidate [228, 229] is also influenced by some or all of these amino acid positions (see fig. 3). In contrast to the situation at GABAA and glycine receptors, in TM4 of GluR6 kainate receptors, residue G819 is critical for volatile anaesthetic (e.g. isoflurane, enflurane, halothane) enhancement but not ethanol or pentobarbitone inhibition of submaximal kainate responses [223].

An obvious extension of the work described above with GABAA and glycine receptors is to determine whether homologous residues in the evolutionarily related neuronal nicotinic ACh and 5-HT3 receptors also play crucial roles in volatile anaesthetic actions. Preliminary evidence suggests that such is indeed the case at the 5-HT3 receptor. Some mutations in TM2 of the 5-HT3 receptor abolish the agonist-potentiating actions of volatile ethers such as enflurane and isoflurane (S. J. Mihic, personal communication).

Most halogenated alkanes and ethers containing six or fewer carbons have anaesthetic properties, but some notable exceptions to this rule exist. The work of Eger, Koblin and colleagues has demonstrated that certain highly lipid-soluble halogenated cyclobutanes and alkanes are unable to produce immobility at concentrations predicted by the Meyer-Overton correlation to be in the anaesthetic range [166]. These compounds, originally called nonanaesthetics, are now more properly referred to as nonimmobilizers, since although they do not produce immobility [166] or analgesia [230], they may interfere with learning and memory [231]. The nonimmobilizers, which are often heavily halogenated compounds (e.g. 1,2-dichlorohexafluorocyclobutane), elicit convulsions at higher concentrations [166]. The nonimmobilizers have no modulatory actions at GABAA [232], glycine [233], GABAC ρ [234], 5-HT3 [235], neuronal nicotinic ACh [236], AMPA or kainate receptors [237]. The nonimmobilizers have, however, been shown to alter the function of muscarinic ACh [238], muscle nicotinic ACh [239] and metabotropic glutamate receptors [240] at concentrations of the nonimmobilizers predicted to be anaesthetic. These results would seem to exclude the muscle nicotinic ACh muscarinic ACh and metabotropic glutamate receptors as viable molecular targets for producing immobility. These receptors may certainly play a role in other actions important in general anaesthesia such as amnesia, since nonimmobilizers and general anaesthetics share some behavioral actions in common.

The anaesthetic gases nitrous oxide and xenon have a pattern of action on the ligand-gated ion channels different from the volatile ethers and alkanes (see Table 2a). This is perhaps not surprising since the clinical effects of xenon and nitrous oxide vary from that of the ethers and alkanes; for instance, unlike the ethers and alkanes, nitrous oxide is a potent analgesic with only weak immobilizing activity [2]. Nitrous oxide inhibits agonist responses at NMDA receptors [194, 195] but has only weak potentiating actions at GABAA receptors [193195]. Very recently, xenon has been demonstrated to inhibit NMDA receptors at clinically relevant concentrations but does not modulate the function of GABAA or AMPA receptors [192]. The anaesthetic properties of xenon and krypton have long presented a challenge for molecular theories of anaesthesia, since these noble gases are among the simplest of molecules that produce anaesthesia [241]. Intriguingly, argon, xenon and krypton all possess anaesthetic properties, whereas the smaller noble gases helium and neon do not produce anaesthesia even at hyperbaric concentrations [242]. The NMDA receptor inhibition produced by xenon and nitrous oxide, with a lack of potent actions on GABAA receptors, resembles the actions of the ‘dissociative anaesthetic’ ketamine at ligand-gated ion channels (see below).

Intravenous agents

Etomidate and propofol both appear to be relatively selective for the GABAA receptor (Table 2b). The GABAA receptor fulfills all the criteria as a plausible target underlying the anaesthetic actions of these compounds. Propofol and etomidate do not modulate other ligand-gated ion channels at clinically relevant concentrations with the exception of propofol actions at the strychnine-sensitive glycine receptor [207, 233, 243]. Amino acid residues within the β subunit of the GABAA receptor have been identified that are essential for potentiation of GABAA receptor function by etomidate [228, 229, 244] and propofol [206] (see fig. 3), consistent with previous studies suggesting that the β subunit of the GABAA receptor was likely to contain binding sites for these compounds [211, 245, 246].

Many steroid anaesthetics such as alphaxalone are relatively selective for the GABAA receptor, although certain steroids have potent actions on other ligand-gated ion channels (see table 2b). For the steroid anaesthetics, structure-activity studies comparing in vivo and in vitro potencies support a role for GABAA receptors in the actions of these compounds [32, 33, 247249]. For example, the nonanaesthetic isomer structural betaxalone does not modulate the GABAA receptor [250, 251]. There have been extensive (although as yet not completely fruitful) attempts to synthesize steroid anaesthetics with improved therapeutic properties over the prototype alphaxalone, and many of these analogs have been tested at the GABAA receptor [252254]. Critical residues for modulation by alphaxalone or other steroid anaesthetics have not yet been identified within any ligand-gated ion channel, although studies of GABAA/glycine chimeric receptors suggest a major contribution of the N-terminal extracellular domain of the GABAA receptor to GABA potentiation by alphaxalone [255].

Unlike propofol, etomidate and the steroid anaesthetics, the barbiturates are much less selective. In addition to their actions at GABAA receptors, barbiturates also potently inhibit AMPA, kainate and neuronal nACh receptors (table 2b). The inhibition of AMPA receptors by barbiturates is voltage- and use-dependent [256, 257]. Studies of recombinant AMPA receptors have revealed that the potency of pentobarbitone block is critically dependent on a glutamine/arginine site in the pore-forming loop of GluR2 subunits [258]. The presence of a glutamine or arginine at this site is determined by specific RNA editing of the GluR2 RNA and strongly influences the ion selectivity and permeation properties of receptors containing the GluR2 subunit [259, 260]. The observation that pentobarbitone block depends on the glutamine/arginine site, together with the voltage- and use-dependence of the block, indicates penetration of barbiturates deep into the ion-conducting pore of AMPA receptors. Optical isomers of pentobarbitone display the same order of potency for modulatory actions at the GABAA receptor as for their in vivo anaesthetic actions [13, 159]. A residue within TM2 of the β1 subunit of the GABAA receptor has been suggested to be necessary for GABA potentiation by pentobarbitone [261] (see fig. 3). Agonist potentiation by barbiturates is not altered by mutations in GABAA receptors that abolish potentiation by volatile anaesthetics, n-alcohols, propofol or trichloroethanol [206, 219, 227]. Similarly, a mutation within TM4 of the kainate GluR6 receptor that ablates volatile anaesthetic enhancement of submaximal kainate responses does not alter inhibition by barbiturates [223].

Compared with other intravenous anaesthetic agents discussed above, the ‘dissociative anaesthetic’ ketamine has a very different in vivo and in vitro profile of action (table 2b). Ketamine and related arylcycloalkylamines such as phencyclidine produce an atypical state of ‘dissociative’ anaesthesia, characterized by sedation, immobility, amnesia, marked analgesia and a feeling of dissociation from the environment, without true unconsciousness [262]. These compounds can also produce intense hallucinations that limit their clinical use, especially in adults [2]. In contrast with most other general anaesthetics, ketamine does not potentiate GABAA receptor function at clinically relevant concentrations [191]. Ketamine appears instead to produce anaesthesia by inhibition of NMDA receptors [163, 250, 263265], although ketamine is also a potent inhibitor of neuronal nicotinic ACh receptors so contributions from these receptors cannot be ruled out [264]. NMDA receptors satisfy all of the pharmacological criteria expected of molecular targets for ketamine and phencyclidine, including stereoselectivity [163]. A site of ketamine action on the NMDA receptor has not yet been elucidated, although single-channel studies have explored the mechanism of ketamine inhibition at NMDA receptors in detail [263].


The alcohols display very little selectivity among the ligand-gated ion channels. In fact, all of the ligand-gated ion channels considered in this review are modulated by anaesthetic concentrations of most alcohols (table 2c). This obfuscates attempts to dissect the molecular underpinnings underlying the diverse behavioral actions of the alcohols. Neuronal nicotinic ACh receptors appear to be exquisitely sensitive to alcohols, in some cases showing modulation by ethanol concentrations as low as 1–10 mM [266, 267]. Mammalian blood alcohol concentrations in this range produce only mild intoxication [268]. As described above, residues within TM2 and TM3 of GABAA and glycine receptors are critically important for the agonist-potentiating actions of the n-alcohols, trichloroethanol and the volatile ether anaesthetics [206, 219, 220, 226, 227] (see figs 3 and and55).

The demonstration of a ‘cutoff’ phenomenon for the in vivo actions of the straight-chain alcohols has presented challenges for many molecular theories of anaesthesia. Potencies of the primary alcohols in producing immobility increase with increasing number of carbon atoms (n), but only up to a certain size (the cutoff), after which alcohols with longer carbon chains decline in potency or remain equipotent with the (n−1)-alcohols [143, 269, 270]. We have followed previous suggestions [13, 220] in defining cutoff as the point at which the potency of the n-alcohol no longer increases with increasing carbon chain length. As with stereoselectivity, the alcohol cutoff poses severe problems for nonspecific theories of anaesthetic action, since there appears to be no cutoff for the disordering actions of n-alcohols on lipid bilayers [14]. In general, the immobilizing actions of n-alcohols show a cutoff around dodecanol (C12) [143, 269, 270], although the limited aqueous solubility of long-chain alcohols can complicate matters [271]. The alcohol cutoff for the ligand-gated ion channels varies between receptors (see table 2c), and this could be useful in implicating or eliminating specific receptors in the various biological effects of the alcohols.

Alcohol cutoff has recently been applied to the study of glycine and GABAC ρ1 receptors harboring mutations in TM2 and TM3. It was first noted that mutation of a smaller to a larger amino acid residue in TM2 of the glycine α1 subunit (serine-267 to glutamine) reduced the alcohol cutoff for the glycine receptor from dodecanol (C12) to propanol (C3) [220]. In contrast, a double mutation of larger to smaller residues in TM2 and TM3 of the GABAC ρ1 receptor extended the alcohol cutoff from heptanol (C7) to beyond dodecanol (C12) [220]. These data provide strong evidence that mutation of selected residues within TM2 and TM3 of glycine and GABAC receptors may actually alter the dimensions of a binding pocket for n-alcohols.

Discussion and future directions

Substantial progress has been made in the last decade in defining the actions of general anaesthetic agents on ligand-gated ion channels, particularly in the areas of molecular biology, pharmacology and electrophysiology. The coming years will surely witness more major advances, perhaps most notably from the application of structural biology and gene-targeting approaches. The use of site-directed mutagenesis and chimeric receptors has proven very helpful in identifying regions of ligand-gated ion channels that play critical roles in modulation by general anaesthetics. However, more definite evidence of the existence of general anaesthetic binding pockets probably awaits the resolution of three-dimensional structures for the ligand-gated ion channels. Structural biology approaches have already been applied to the study of general anaesthetic interactions with model soluble proteins [20], including the recent report of the 2.2-Å resolution three-dimensional structure of firefly luciferase complexed with the general anaesthetic bromoform [272].

In common with other many integral membrane proteins, ligand-gated ion channels have proved recalcitrant to structural biology approaches. However, the crystallization and determination of a high-resolution structure for a bacterial potassium channel [273] surely foreshadows the eventual determination of the three-dimensional structure of the ligand-gated ion channels. A more immediate possibility is the determination of the structure of limited domains of ligand-gated ion channels; indeed, researchers have very recently succeeded in resolving the structure of the extracellular domain of an ionotropic glutamate receptor complexed with kainate [274]. Even in the absence of detailed structures, molecular modeling may be of use in making preliminary predictions that can be tested experimentally.

Targeted gene manipulations in mice will also provide hypothesis-driven tests of the in vivo roles of certain ligand-gated ion channels in mediating the diverse behavioral actions of general anaesthetics. As described above, researchers over the last 5 years have created ‘global knockout mice’ for various subunits of the ligand-gated ion channels. Given the abundance of ligand-gated ion channel knockout mice (and the commercial availability of some of these knockouts), it would be a logical step to test anaesthetic sensitivity in some or all of these mice. However, while knockout mice may provide initial clues as to the nature of anaesthetic targets, such mice can be very difficult to analyze for anaesthetic sensitivity if they exhibit grossly abnormal motor behavior, lethality or aberrations in neural development. These problems with knockout mice may be circumvented by ‘conditional’ gene knockouts, in which the gene of interest is disrupted only in limited brain regions and/or specified developmental time periods [172].

Another elegant example of gene targeting is a ‘knock-in’ mouse. One possibility is the introduction of a mutated receptor subunit that is insensitive to anaesthetic modulation in place of the normal endogenous receptor subunit (e.g. see [175]). This type of approach has recently been applied to the benzodiazepines. These studies utilized knock-in mice expressing a mutant GABAA receptor α1 subunit that confers insensitivity to benzodiazepine modulation, in place of the benzodiazepine-sensitive wild-type α1 subunit. These preliminary studies have not only demonstrated the importance of the GABAA α1 subunit isoform for the behavioral actions of benzodiazepines but also have suggested that distinct GABAA receptor α-subunit isoforms mediate different actions of the benzodiazepines, with the α1 subunit isoform necessary for sedative and anticonvulsant effects and other α-subunit isoforms critical for myorelaxant and anxiolytic actions (U. Rudolph, F. Crestani, H. Möhler, personal communication).

Knock-in mouse experiments potentially provide an elegant bridge between in vitro experiments and whole animal behavior. Ideally, the mutated receptor subunit would differ from the normal subunit only in terms of general anaesthetic modulation (i.e. agonist binding, channel gating, voltage dependence, kinetics etc. of the receptor would be relatively normal). Recently described mutations within TM2 and TM3 of GABAA (see fig. 3) and glycine receptors, which confer insensitivity to volatile ether anaesthetics [206, 219], n-alkanols [219, 220, 226], propofol [206], trichloroethanol [227], pentobarbitone [261] and etomidate [228, 229] essentially fit this qualification, as do point mutations within GluR6 kainate receptors that abolish volatile anaesthetic potentiation [223]. A complication to gene-targeting experiments is the presence of multiple subunit isoforms for the ligand-gated ion channels. For example, there are at least 17 gene products for GABAA receptor subunits; if multiple GABAA subunit isoforms play a role in general anaesthesia, then targeting of multiple genes may be required to obtain an unambiguous change in anaesthetic sensitivity. General anaesthetics produce a range of behavioral effects in animals and humans. It appears overly simplistic to ascribe all of these to a single receptor. Current and future research should eventually define the specific receptors that underlie each of the diverse behavioral actions of every class of general anaesthetics. The upcoming decade will undoubtedly be an exciting time for research into the molecular mechanisms of general anaesthetics.


We would like to thank Drs P. Flood and C. E Rickfor careful reading of the manuscript and the reviewers for many helpful suggestions. Funding was generously provided by NIH grants GM45129, GM56850 and GM00623 to N.L.H. and by NIMH training grant MH11504 to M.D.K.


1. Harrison NL, Flood P. Molecular mechanisms of general anesthetic action. Sci. Med. 1998;5:18–27.
2. Marshall BE, Longnecker DE. General anesthetics. In: Hardman JG, Limbird LE, Molinoff PB, Ruddon RW, Gilman AG, editors. The Pharmacological Basis of Therapeutics. McGraw-Hill; New York: 1996. pp. 307–330.
3. Meyer H. Zur Theorie der Alkolnarkose: der Einfuss wechselnder Temperatur auf Wirkungsstarke und Theilungscoefficient der Narcotica. Naunyn-Schmiedebergs Arch. Exp. Path. Pharmakol. 1901;46:338–346.
4. Meyer H. Welche eigenschaft der Anasthetica bedingt ihre Narkotische wirkung? Naunyn-Schmiedebergs Arch. Exp. Path. Pharmakol. 1899;42:109–118.
5. Overton E. Studien uber die Narkose, zugleich ein Beitrag zur allgemeiner Pharmakologie. Gustav Fischer; Jena, Switzerland: 1901.
6. Meyer KH. Contribution to the theory of narcosis. Trans. Faraday Soc. 1937;33:1062–1068.
7. Mullins LJ. Some physical mechanisms in narcosis. Chem. Rev. 1954;54:289–322.
8. Seeman P. The membrane actions of anesthetics and tranquilizers. Pharmacol. Rev. 1972;24:583–655. [PubMed]
9. Franks NP, Lieb WR. Do general anaesthetics act by competitive binding to specific receptors? Nature. 1984;310:599–601. [PubMed]
10. Franks NP, Lieb WR. The pharmacology of simple molecules. Archiv. Toxicol. Suppl. 1986;9:27–37. [PubMed]
11. Franks NP, Lieb WR. Is membrane expansion relevant to anaesthesia? Nature. 1981;292:248–251. [PubMed]
12. Franks NP, Lieb WR. Molecular mechanisms of general anaesthesia. Nature. 1982;300:487–493. [PubMed]
13. Franks NP, Lieb WR. Molecular and cellular mechanisms of general anaesthesia. Nature. 1994;367:607–614. [PubMed]
14. Franks NP, Lieb WR. Partitioning of long-chain alcohols into lipid bilayers: implications for mechanisms of general anesthesia. Proc. Natl. Acad. Sci. USA. 1986;83:5116–5120. [PubMed]
15. Tomlin SL, Jenkins A, Lieb WR, Franks NP. Stereoselective effects of etomidate optical isomers on gamma-aminobutyric acid type A receptors and animals. Anesthesiol. 1998;88:708–717. [PubMed]
16. Elliott JR, Urban BW. Integrative effects of general anaesthetics: why nerve axons should not be ignored. Eur. J. Anaesth. 1995;12:41–50. [PubMed]
17. Rehberg B, Urban BW, Duch DS. The membrane lipid cholesterol modulates anesthetic actions on a human brain ion channel. Anesthesiol. 1995;82:749–758. [PubMed]
18. Cantor RS. The lateral pressure profile in membranes: a physical mechanism of general anesthetics. Biochemistry. 1997;36:2339–2344. [PubMed]
19. Halsey MJ. Molecular interactions of anaesthetics with biological membranes. Gen. Pharmacol. 1992;23:1013–1016. [PubMed]
20. Eckenhoff RG, Johansson JS. Molecular interactions between inhaled anesthetics and proteins. Pharmacol. Rev. 1997;49:343–367. [PubMed]
21. Eckenhoff RG. Do specific or nonspecific interactions with proteins underlie inhalational anesthetic action? Mol. Pharmacol. 1998;54:610–615. [PubMed]
22. Eccles JC, Malcolm JL. Dorsal root potentials of the spinal cord. J. Neurophysiol. 1946;9:139–160. [PubMed]
23. Eccles JC, Schmidt R, Willis WD. Pharmacological studies on presynaptic inhibition. J. Physiol. 1963;168:500–530. [PubMed]
24. Tanelian DL, Kosek P, Mody I, MacIver MB. The role of the GABAA receptor/chloride channel complex in anesthesia. Anesthesiol. 1993;78:757–776. [PubMed]
25. Pearce RA. Effects of volatile anesthetics on GABAA receptors: electrophysiological studies. In: Moody EJ, Skolnick P, editors. Molecular Bases of Anesthesia. CRC Press; Boca Raton, FL: 1999. in press.
26. Franks NP, Lieb WR. An anesthetic-sensitive superfamily of neurotransmitter-gated ion channels. J. Clin. Anesth. 1996;8:3S–7S. [PubMed]
27. Harris RA, Mihic SJ, Dildy-Mayfield JE, Machu TK. Actions of anesthetics on ligand-gated ion channels: role of receptor subunit composition. FASEB J. 1995;9:1454–1462. [PubMed]
28. Franks NP, Lieb WR. Selective actions of volatile general anaesthetics at molecular and cellular levels. Br. J. Anaesth. 1993;71:65–76. [PubMed]
29. Weight FF, Lovinger DM, White G, Peoples RW. Alcohol and anesthetic actions on excitatory amino acid-activated ion channels. Ann. N. Y. Acad. Sci. 1991;625:97–107. [PubMed]
30. Weight FF, Aguayo LG, White G, Lovinger DM, Peoples RW. GABA- and glutamate-gated ion channels as molecular sites of alcohol and anesthetic action. Adv. Biochem. Psychopharmacol. 1992;47:335–347. [PubMed]
31. Peoples RW, Weight FF. Anesthetic actions on excitatory amino acid receptors. In: Yaksh TL, editor. Anesthesia: Biologic Foundations. Lippincott- Raven; Philadelphia: 1997. pp. 239–258.
32. Lambert JJ, Belelli D, Hill-Venning C, Peters JA. Neurosteroids and GABAA receptor function. Trends Pharmacol. Sci. 1995;16:295–303. [PubMed]
33. Lambert JJ, Belelli D, Hill-Venning C, Callachan H, Peters JA. Neurosteroid modulation of native and recombinant GABAA receptors. Cell. Mol. Neurobiol. 1996;16:155–174. [PubMed]
34. Lovinger DM. Alcohols and neurotransmitter gated ion channels: past, present and future. Naunyn-Schmiedebergs Arch. Pharmacol. 1997;356:267–282. [PubMed]
35. Mihic SJ, Sanna E, Whiting PJ, Harris RA. Pharmacology of recombinant GABAA receptors. Adv. Biochem. Psychopharmacol. 1995;48:17–40. [PubMed]
36. Smith GB, Olsen RW. Functional domains of GABAA receptors. Trends Pharmacol. Sci. 1995;16:162–168. [PubMed]
37. Whiting PJ, McKernan RM, Wafford KA. Structure and pharmacology of vertebrate GABAA receptor subtypes. Intl. Rev. Neurobio. 1995;38:95–138. [PubMed]
38. Ortells MO, Lunt GG. Evolutionary history of the ligand-gated ion-channel superfamily of receptors. Trends Neurosci. 1995;18:121–127. [PubMed]
39. Johnston GA. GABAA receptor pharmacology. Pharmacol. Ther. 1996;69:173–198. [PubMed]
40. Kuhse J, Betz H, Kirsch J. The inhibitory glycine receptor: architecture, synaptic localization and molecular pathology of a postsynaptic ion-channel complex. Curr. Opin. Neurobiol. 1995;5:318–323. [PubMed]
41. Lindstrom J, Anand R, Gerzanich V, Peng X, Wang F, Wells G. Structure and function of neuronal nicotinic acetylcholine receptors. Prog. Brain Res. 1996;109:125–137. [PubMed]
42. Xu M, Akabas MH. Amino acids lining the channel of the γ-aminobutyric acid type A receptor identified by cysteine substitution. J. Biol. Chem. 1993;268:21505–21508. [PubMed]
43. Akabas MH, Kaufmann C, Archdeacon P, Karlin A. Identification of acetylcholine receptor channel-lining residues in the entire M2 segment of the α subunit. Neuron. 1994;13:919–927. [PubMed]
44. Langosch D, Thomas L, Betz H. Conserved quaternary structure of ligand-gated ion channels: the postsynaptic glycine receptor is a pentamer. Proc. Natl. Acad. Sci. USA. 1988;85:7394–7398. [PubMed]
45. Cooper E, Couturier S, Ballivet M. Pentameric structure and subunit stoichiometry of a neuronal nicotinic acetylcholine receptor. Nature. 1991;350:235–238. [PubMed]
46. McCormick DA. GABA as an inhibitory neurotransmitter in human cerebral cortex. J. Neurophysiol. 1989;62:1018–1027. [PubMed]
47. Zafra F, Aragon C, Gimenez C. Molecular biology of glycinergic neurotransmission. Mol. Neurobiol. 1997;14:117–142. [PubMed]
48. Bloom FE, Iversen LL. Localizing [3H]GABA in nerve terminals of cerebral cortex by electron microscopic autoradiography. Nature. 1971;229:628–630. [PubMed]
49. Davies PA, Hanna MC, Hales TG, Kirkness EF. Insensitivity to anaesthetic agents conferred by a class of GABAA receptor subunit. Nature. 1997;385:820–823. [PubMed]
50. Whiting PJ, McAllister G, Vasilatis D, Bonnert TP, Heavens RP, Smith DW, et al. Neuronally restricted RNA splicing regulates the expression of a novel GABAA receptor subunit conferring atypical functional properties. J. Neurosci. 1997;17:5027–5037. [PubMed]
51. Hedblom E, Kirkness EF. A novel class ofGABAA receptor subunit in tissues of the reproductive system. J. Biol. Chem. 1997;272:15346–15350. [PubMed]
52. McKernan RM, Whiting PJ. Which GABAA-receptor subtypes really occur in the brain? Trends Neurosci. 1996;19:139–143. [PubMed]
53. Rabow LE, Russek SJ, Farb DH. From ion currents to genomic analysis: recent advances in GABAA receptor research. Synapse. 1995;21:189–274. [PubMed]
54. Stephenson FA. The GABAA receptors. Biochem. J. 1995;310:1–9. [PubMed]
55. Macdonald RL, Olsen RW. GABAA receptor channels. Annu. Rev. Neurosci. 1994;17:569–602. [PubMed]
56. Barnard EA, Skolnick P, Olsen RW, Mohler H, Sieghart W, Biggio G, et al. International union of pharmacology. XV. Subtypes of γ-aminobutyric acidA receptors: classification on the basis of subunit structure and receptor function. Pharmacol. Rev. 1998;50:291–313. [PubMed]
57. Betz H. Glycine receptors: heterogeneous and widespread in the mammalian brain. Trends Neurosci. 1991;14:458–461. [PubMed]
58. Betz H. Structure and function of inhibitory glycine receptors. Q. Rev. Biophys. 1992;25:381–394. [PubMed]
59. Tretter V, Ehya N, Fuchs K, Sieghart W. Stoichiometry and assembly of a recombinant GABAA receptor subtype. J. Neurosci. 1997;17:2728–2737. [PubMed]
60. Chang Y, Wang R, Barot S, Weiss DS. Stoichiometry of a recombinant GABAA receptor. J. Neurosci. 1996;16:5415–5424. [PubMed]
61. Fritschy JM, Mohler H. GABAA-receptor heterogeneity in the adult rat brain: differential regional and cellular distribution of seven major subunits. J. Comp. Neurol. 1995;359:154–194. [PubMed]
62. Nusser Z, Sieghart W, Benke D, Fritschy JM, Somogyi P. Differential synaptic localization of two major γ-aminobutyric acid type A receptor α subunits on hippocampal pyramidal cells. Proc. Natl. Acad. Sci. USA. 1996;93:11939–11944. [PubMed]
63. Sieghart W. Structure and pharmacology of γ-aminobutyric acidA receptor subtypes. Pharmacol. Rev. 1995;47:181–234. [PubMed]
64. Levitan ES, Blair LA, Dionne VE, Barnard EA. Biophysical and pharmacological properties of cloned GABAA receptor subunits expressed in Xenopus oocytes. Neuron. 1988;1:773–781. [PubMed]
65. Sigel E, Baur R, Trube G, Mohler H, Malherbe P. The effect of subunit composition of rat brain GABA receptors on channel function. Neuron. 1990;5:703–711. [PubMed]
66. Ebert B, Wafford KA, Whiting PJ, Krogsgaard-Larsen P, Kemp JA. Molecular pharmacology of _-aminobutyric acid type A receptor agonists and partial agonists in oocytes injected with different α, β and γ receptor subunit combinations. Mol. Pharmacol. 1994;46:957–963. [PubMed]
67. Wafford KA, Whiting PJ, Kemp JA. Differences in affinity and efficacy of benzodiazepine receptor ligands at recombinant γ-aminobutyric acidA receptor subtypes. Mol. Pharmacol. 1993;43:240–244. [PubMed]
68. Tia S, Wang JF, Kotchabhakdi N, Vicini S. Developmental changes of inhibitory synaptic currents in cerebellar granule neurons: role of GABAA receptor α6 subunit. J. Neurosci. 1996;16:3630–3640. [PubMed]
69. Lavoie AM, Tingey JJ, Harrison NL, Pritchett DB, Twyman RE. Activation and deactivation rates of recombinant GABAA receptor channels are dependent on α-subunit isoform. Biophys. J. 1997;73:2518–2526. [PubMed]
70. Cutting GR, Lu L, O'Hara BF, Kasch LM, Montrose-Rafizadeh C, Donovan DM, et al. Cloning of the γ-aminobutyric acid (GABA) rho 1 cDNA: a GABA receptor subunit highly expressed in the retina. Proc. Natl. Acad. Sci. USA. 1991;88:2673–2677. [PubMed]
71. Cutting GR, Curristin S, Zoghbi H, O'Hara B, Seldin MF, Uhl GR. Identification of a putative γ-aminobutyric acid (GABA) receptor subunit rho2 cDNA and colocalization of the genes encoding rho2 (GABRR2) and rho1 (GABRR1) to human chromosome 6q14-q21 and mouse chromosome 4. Genomics. 1992;12:801–806. [PubMed]
72. Johnston GA. GABAC receptors: relatively simple transmitter-gated ion channels? Trends Pharmacol. Sci. 1996;17:319–323. [PubMed]
73. Wegelius K, Pasternack M, Hitunen JO, Rivera C, Kaila K, Saarma M, et al. Distribution of GABA receptor ρ subunit transcripts in the rat brain. Eur. J. Neurosci. 1998;10:350–357. [PubMed]
74. Caratsch CG, Grassi F, Eusebi F. Functional regulation of nicotinic acetylcholine receptor channels in muscle. Ion Channels. 1992;3:177–206. [PubMed]
75. Conti-Tronconi BM, McLane KE, Raftery MA, Grando SA, Protti MP. The nicotinic acetylcholine receptor: structure and autoimmune pathology. Crit. Rev. Biochem. Mol. Biol. 1994;29:69–123. [PubMed]
76. Galzi JL, Revah F, Bessis A, Changeux JP. Functional architecture of the nicotinic acetylcholine receptor: from electric organ to brain. Annu. Rev. Pharmacol. Toxicol. 1991;31:37–72. [PubMed]
77. Gotti C, Fornasari D, Clementi F. Human neuronal nicotinic receptors. Prog. Neurobiol. 1997;53:199–237. [PubMed]
78. Lindstrom J. Neuronal nicotinic acetylcholine receptors. Ion Channels. 1996;4:377–450. [PubMed]
79. McGehee DS, Role LW. Physiological diversity of nicotinic acetylcholine receptors expressed by vertebrate neurons. Annu. Rev. Physiol. 1995;57:521–546. [PubMed]
80. Boyd RT. The molecular biology of neuronal nicotinic acetylcholine receptors. Crit. Rev. Toxicol. 1997;27:299–318. [PubMed]
81. McGehee DS, Role LW. Presynaptic ionotropic receptors. Curr. Opin. Neurobiol. 1996;6:342–349. [PubMed]
82. Frazier CJ, Buhler AV, Weiner JL, Dunwiddie TV. Synaptic potentials mediated via α-bungarotoxin sensitive nicotinic acetylcholine receptors in rat hippocampal interneurons. J. Neurosci. 1998;18:8228–8235. [PubMed]
83. Tecott LH, Maricq AV, Julius D. Nervous system distribution of the serotonin 5-HT3 receptor mRNA. Proc. Natl. Acad. Sci. USA. 1993;90:1430–1434. [PubMed]
84. Fletcher S, Barnes NM. Desperately seeking subunits: are native 5-HT3 receptors really homomeric complexes? Trends Pharmacol. Sci. 1998;19:212–215. [PubMed]
85. Jackson MB, Yakel JL. The 5-HT3 receptor channel. Annu. Rev. Physiol. 1995;57:447–468. [PubMed]
86. Gyermeck L. 5-HT3 receptors: pharmacologic and therapeutic aspects. J. Clin. Pharmacol. 1995;35:845–855. [PubMed]
87. Grant KA. The role of 5-HT3 receptors in drug dependence. Drug Alcohol Depend. 1995;38:155–171. [PubMed]
88. Ali Z, Wu G, Kozlov A, Barasi S. The role of 5-HT3 receptors in nociceptive processing in the rat spinal cord: results from behavioural and electrophysiological studies. Neurosci. Lett. 1996;208:203–207. [PubMed]
89. Bennett JA, Dingledine R. Topology profile for a glutamate receptor: three transmembrane domains and a channel-lining reentrant membrane loop. Neuron. 1995;14:373–384. [PubMed]
90. Hollmann M, Maron C, Heinemann S. N-glycosylation site tagging suggests a three transmembrane domain topology for the glutamate receptor GluR1. Neuron. 1994;13:1331–1343. [PubMed]
91. Sutcliffe MJ, Wo ZG, Oswald RE. Three-dimensional models of non-NMDA glutamate receptors. Biophys. J. 1996;70:1575–1589. [PubMed]
92. Wo ZG, Oswald RE. Transmembrane topology of two kainate receptor subunits revealed by N-glycosylation. Proc. Natl. Acad. Sci. USA. 1994;91:7154–7158. [PubMed]
93. Heginbotham L, Lu Z, Abramson T, MacKinnon R. Mutations in the K+ channel signature sequence. Biophys. J. 1994;66:1061–1067. [PubMed]
94. Rosenmund C, Stern-Bach Y, Stevens CF. The tetrameric structure of a glutamate receptor channel. Science. 1998;280:1596–1599. [PubMed]
95. Laube B, Kuhse J, Betz H. Evidence for a tetrameric structure of recombinant NMDA receptors. J. Neurosci. 1998;18:2954–2961. [PubMed]
96. MacKinnon R. Determination of the subunit stoichiometry of a voltage-activated potassium channel. Nature. 1991;350:232–235. [PubMed]
97. Nakanishi S, Nakajima Y, Masu M, Ueda Y, Nakahara K, Watanabe D, et al. Glutamate receptors: brain function and spinal transduction. – Brain Res. Rev. Brain Res. 1998;26:230–235.
98. Seeburg PH. The TiPS/TINS lecture: the molecular biology of mammalian glutamate receptor channels. Trends Pharmacol. Sci. 1993;14:297–303. [PubMed]
99. Mori H, Mishina M. Structure and function of the NMDA receptor channel. Neuropharmacol. 1995;34:1219–1237. [PubMed]
100. Sucher NJ, Awobuluyi M, Choi YB, Lipton SA. NMDA receptors: from genes to channels. Trends Pharmacol. Sci. 1996;17:348–355. [PubMed]
101. MacDermott AB, Mayer ML, Westbrook GL, Smith SJ, Barker JL. NMDA-receptor activation increases cytoplasmic calcium concentration in cultured spinal cord neurones. Nature. 1986;321:519–522. [PubMed]
102. Mayer ML, Westbrook GL. The physiology of excitatory amino acids in the vertebrate central nervous system. Prog. Neurobiol. 1987;28:197–276. [PubMed]
103. Mayer ML, Westbrook GL, Guthrie PB. Voltage-dependent block by Mg2+ of NMDA responses in spinal cord neurones. Nature. 1984;309:261–263. [PubMed]
104. Nowak L, Bregestovski P, Ascher P, Herbet A, Prochiantz A. Magnesium gates glutamate-activated channels in mouse central neurones. Nature. 1984;307:462–465. [PubMed]
105. Collingridge GL, Kehl SJ, McLennan H. Excitatory amino acids in synaptic transmission in the Schaffer collateral-commissural pathway of the rat hippocampus. J. Physiol. 1983;334:33–46. [PubMed]
106. Nicoll RA, Malenka RC. Contrasting properties of two forms of long-term potentiation in the hippocampus. Nature. 1995;377:115–118. [PubMed]
107. Kirkwood A, Bear MF. Elementary forms of synaptic plasticity in the visual cortex. Biol. Res. 1995;28:73–80. [PubMed]
108. Bliss TV, Collingridge GL. A synaptic model of memory: long-term potentiation in the hippocampus. Nature. 1993;361:31–39. [PubMed]
109. Morris RG, Anderson E, Lynch GS, Baudry M. Selective impairment of learning and blockade of long-term potentiation by an N-methyl-D-aspartate receptor antagonist, AP5. Nature. 1986;319:774–776. [PubMed]
110. Rothman SM, Olney JW. Excitotoxicity and the NMDA receptor – still lethal after eight years. Trends Neurosci. 1995;18:57–58. [PubMed]
111. Choi DW. Glutamate receptors and the induction of excitotoxic neuronal death. Prog. Brain Res. 1994;100:47–51. [PubMed]
112. Lipton SA, Rosenberg PA. Excitatory amino acids as a final commonpathway for neurologic disorders. New Engl. J. Med. 1994;330:613–622. [PubMed]
113. Michaelis EK. Molecular biology of glutamate receptors in the central nervous system and their role in excitotoxicity, oxidative stress and aging. Prog. Neurobiol. 1998;54:369–415. [PubMed]
114. Johnson JW, Ascher P. Glycine potentiates the NMDAresponse in cultured mouse brain neurons. Nature. 1987;325:529–531. [PubMed]
115. Kleckner NW, Dingledine R. Requirement for glycine in activation of NMDA-receptors expressed in Xenopus oocytes. Science. 1988;241:835–837. [PubMed]
116. Ransom RW, Stec NL. Cooperative modulation of [3H]MK-801 binding to the N-methyl-D-aspartate receptorion channel complex by L-glutamate, glycine, and polyamines. J. Neurochem. 1988;51:830–836. [PubMed]
117. Sprosen TS, Woodruff GN. Polyamines potentiate NMDA induced whole-cell currents in cultured striatal neurons. Eur. J. Pharmacol. 1990;179:477–478. [PubMed]
118. Peters S, Koh J, Choi DW. Zinc selectively blocks the action of N-methyl-D-aspartate on cortical neurons. Science. 1987;236:589–593. [PubMed]
119. Westbrook GL, Mayer ML. Micromolar concentrations of Zn2+ antagonize NMDA and GABA responses of hippocampal neurons. Nature. 1987;328:640–643. [PubMed]
120. Traynelis SF, Cull-Candy SG. Proton inhibition of N-methyl-D-aspartate receptors in cerebellar neurons. Nature. 1990;345:347–350. [PubMed]
121. Vyklicky L, Vlachova V, Krusek J. The effect of external pH changes on responses to excitatory amino acids in mouse hippocampal neurones. J. Physiol. 1990;430:497–517. [PubMed]
122. Miller B, Sarantis M, Traynelis SF, Attwell D. Potentiation of NMDA receptor currents by arachidonic acid. Nature. 1992;355:722–725. [PubMed]
123. Nishikawa M, Kimura S, Akaike N. Facilitatory effect of docosahexaenoic acid on N-methyl-D-aspartate response in pyramidal neurones of rat cerebral cortex. J. Physiol. 1994;475:83–93. [PubMed]
124. Aizenman E, Lipton SA, Loring RH. Selective modulation of NMDA responses by reduction and oxidation. Neuron. 1989;2:1257–1263. [PubMed]
125. Ozawa S, Kamiya H, Tsuzuki K. Glutamate receptors in the mammalian central nervous system. Prog. Neurobiol. 1998;54:581–618. [PubMed]
126. Clements JD, Lester RA, Tong G, Jahr CE, Westbrook GL. The time course of glutamate in the synaptic cleft. Science. 1992;258:1498–1501. [PubMed]
127. Lerma J. Kainate reveals its targets. Neuron. 1997;19:1155–1158. [PubMed]
128. Lerma J, Morales M, Vicente MA, Herreras O. Glutamate receptors of the kainate type and synaptic transmission. Trends Neurosci. 1997;20:9–12. [PubMed]
129. Paternain AV, Morales M, Lerma J. Selective antagonism of AMPAreceptors unmasks kainate receptor-mediated responses in hippocampal neurons. Neuron. 1995;14:185–189. [PubMed]
130. Mosbacher J, Schoepfer R, Monyer H, Burnashev N, Seeburg PH, Ruppersberg JP. A molecular determinant for submillisecond desensitization in glutamate receptors. Science. 1994;266:1059–1062. [PubMed]
131. Colquhoun D, Jonas P, Sakmann B. Action of brief pulses of glutamate on AMPA/kainate receptors in patches from different neurones of rat hippocampal slices. J. Physiol. 1992;458:261–287. [PubMed]
132. Swanson GT, Gereau RW, Green T, Heinemann SF. Identification of amino acid residues that control functional behavior in GluR5 and GluR6 kainate receptors. Neuron. 1997;19:913–926. [PubMed]
133. Quasha AL, Eger EI, Tinker JH. Determination and applications of MAC. Anesthesiol. 1980;53:315–334. [PubMed]
134. Eger EI, Saidman LJ, Brandstater B. Minimum alveolar anesthetic concentration: a standard of anesthetic potency. Anesthesiol. 1965;26:756–763. [PubMed]
135. Iselin-Chaves IA, Flaishon R, Sebel PS, Howell S, Gan TJ, Sigl J, et al. The effect of the interaction of propofol and alfentanil on recall, loss of consciousness, and the bispectral index. Anesth. Analg. 1998;87:949–955. [PubMed]
136. Chortkoff BS, Gonsowski CT, Bennett HL, Levinson B, Crankshaw DP, Dutton RC, et al. Subanesthetic concentrations of desflurane and propofol suppress recall of emotionally charged information. Anesth. Analg. 1995;81:728–736. [PubMed]
137. Chortkoff BS, Eger EI, Crankshaw DP, Gonsowski CT, Dutton RC, Ionescu P. Concentrations of desflurane and propofol that suppress response to command in humans. Anesth. Analg. 1995;81:737–743. [PubMed]
138. Franks NP, Lieb WR. Temperature dependence of the potency of volatile general anesthetics: implications for in vitro experiments. Anesthesiol. 1996;84:716–720. [PubMed]
139. Jenkins A, Franks NP, Lieb WR. Effects of temperature and volatile anesthetics on GABAA receptors. Anesthesiol. 1999;90:484–491. [PubMed]
140. Sear JW, Prys-Roberts C. Plasma concentrations of alphaxalone during continuous infusion of Althesin. Br. J. Anaesth. 1979;51:861–865. [PubMed]
141. Cohen ML, Chan SL, Way WL, Trevor AJ. Distribution in the brain and metabolism of ketamine in the rat after intravenous administration. Anesthesiol. 1973;39:370–376. [PubMed]
142. Sonner JM, Li J, Eger EI. Desflurane and the non-immobilizer 1,2-dichlorohexafluorocyclobutane suppress learning and memory by a mechanism independent of the level of unconditioned stimulation. Anesth. Analg. 1998;87:200–205. [PubMed]
143. Alifimoff JK, Firestone LL, Miller KW. Anaesthetic potencies of primary alkanols: implications for the molecular dimensions of the anaesthetic site. Br. J. Pharmacol. 1989;96:9–16. [PMC free article] [PubMed]
144. Antognini JF, Schwartz K. Exaggerated anesthetic requirements in the preferentially anesthetized brain. Anesthesiol. 1993;79:1244–1249. [PubMed]
145. Collins JG, Kendig JJ, Mason P. Anesthetic actions within the spinal cord: contributions to the state of general anesthesia. Trends Neurosci. 1995;18:549–553. [PubMed]
146. Rampil IJ, Mason P, Singh H. Anesthetic potency (MAC) is independent of forebrain structures in the rat. Anesthesiol. 1993;78:707–712. [PubMed]
147. Harrison NL. Optical isomers open a new window on anesthetic mechanism. Anesthesiol. 1998;88:566–568. [PubMed]
148. Heykants JJ, Meuldermans WE, Michiels LJ, Lewi PJ, Janssen PA. Distribution, metabolism and excretion of etomidate, a short-acting hypnotic drug, in the rat. Comparative study of (R)-(+) and S-(−)-Etomidate. Arch. Intl. Pharmacodyn. Ther. 1975;216:113–129. [PubMed]
149. Andrews PR, Mark LC. Structural specificity of barbiturates and related drugs. Anesthesiol. 1982;57:314–320. [PubMed]
150. Lysko GS, Robinson JL, Casto R, Ferrone RA. The stereospecific effects of isoflurane isomers in vivo. Eur. J. Pharmacol. 1994;263:25–29. [PubMed]
151. Harris B, Moody E, Skolnick P. Isoflurane anesthesia is stereoselective. Eur. J. Pharmacol. 1992;217:215–216. [PubMed]
152. Eger EI, Koblin DD, Laster MJ, Schurig V, Juza M, Ionescu P, et al. Minimum alveolar anesthetic concentration values for the enantiomers of isoflurane differ minimally. Anesth. Analg. 1997;85:188–192. [PubMed]
153. White PF, Schuttler J, Shafer A, Stanski DR, Horai Y, Trevor AJ. Comparative pharmacology of the ketamine isomers. Studies in volunteers. Br. J. Anaesth. 1985;57:197–203. [PubMed]
154. Ryder S, Way WL, Trevor AJ. Comparative pharmacology of the optical isomers of ketamine in mice. Eur. J. Pharmacol. 1978;49:15–23. [PubMed]
155. Wittmer LL, Hu Y, Kalkbrenner M, Evers AS, Zorumski CF, Covey DF. Enantioselectivity of steroid-induced γ-aminobutyric acidA receptor modulation and anesthesia. Mol. Pharmacol. 1996;50:1581–1586. [PubMed]
156. Moody EJ, Harris BD, Skolnick P. The potential for safer anaesthesia using stereoselective anaesthetics. Trends Pharmacol. Sci. 1994;15:387–391. [PubMed]
157. Dickinson R, Franks NP, Lieb WR. Can the stereoselective effects of the anesthetic isoflurane be accounted for by lipid solubility? Biophys. J. 1994;66:2019–2023. [PubMed]
158. Tomlin SL, Jenkins A, Lieb WR, Franks NP. Preparation of barbiturate optical isomers and their effects on GABAA receptors. Anesthesiol. 1999 in press. [PubMed]
159. Huang LY, Barker JL. Pentobarbital: stereospecific actions of (+) and (–) isomers revealed on cultured mammalian neurons. Science. 1980;207:195–197. [PubMed]
160. Jones MV, Harrison NL. Effects of volatile anesthetics on the kinetics of inhibitory postsynaptic currents in cultured rat hippocampal neurons. J. Neurophysiol. 1993;70:1339–1349. [PubMed]
161. Hall AC, Lieb WR, Franks NP. Stereoselective and non-stereoselective actions of isoflurane on the GABAA receptor. Br. J. Pharmacol. 1994;112:906–910. [PMC free article] [PubMed]
162. Atkinson RM, Davis B, Pratt MA, Sharpe HM, Tomich EG. Action of some steroids on the central nervous system of the mouse. J. Med. Chem. 1965;8:426–432. [PubMed]
163. Lodge D, Anis NA, Burton NR. Effects of optical isomers of ketamine on excitation of cat and rat spinal neurones by amino acids and acetylcholine. Neurosci. Lett. 1982;29:281–286. [PubMed]
164. Roth SH, Forman SA, Braswell LM, Miller KW. Actions of pentobarbital enantiomers on nicotinic cholinergic receptors. Mol. Pharmacol. 1989;36:874–880. [PubMed]
165. de Armendi AJ, Tonner PH, Bugge B, Miller KW. Barbiturate action is dependent on the conformational state of the acetylcholine receptor. Anesthesiol. 1993;79:1033–1041. [PubMed]
166. Koblin DD, Chortkoff BS, Laster MJ, Eger EI, Halsey MJ, Ionescu P. Polyhalogenated and perfluorinated compounds that disobey the Meyer-Overton hypothesis. Anesth. Analg. 1994;79:1043–1048. [PubMed]
167. Olsen RW, Snowman AM. Chloride-dependent enhancement by barbiturates of γ-aminobutyric acid receptor binding. J. Neurosci. 1982;2:1812–1823. [PubMed]
168. Harris BD, Wong G, Moody EJ, Skolnick P. Different subunit requirements for volatile and nonvolatile anesthetics at γ-aminobutyric acid type A receptors. Mol. Pharmacol. 1995;47:363–367. [PubMed]
169. Eckenhoff RG. An inhalational anesthetic binding domain in the nicotinic acetylcholine receptor. Proc. Natl. Acad. Sci. USA. 1996;93:2807–2810. [PubMed]
170. Sigel E, Buhr A. The benzodiazepine binding site of the GABAA receptor. Trends Pharmacol. Sci. 1997;18:425–429. [PubMed]
171. McKernan RM, Farrar S, Collins I, Emms F, Asuni A, Quirk K, et al. Photoaffinity labeling of the benzodiazepine binding site of α1β3γ2-aminobutyric acidA receptors with flunitrazepam identifies a subset of ligands that interact directly with His102 of the α subnit and predicts orientation of these within the benzodiazepine pharmacophore. Mol. Pharmacol. 1998;54:33–43. [PubMed]
172. Homanics GE, Quinlan JJ, Mihalek RM, Firestone LL. Alcohol and anesthetic mechanisms in genetically engineered mice. Front. Biosci. 1998;3:D548–D558. [PubMed]
173. Gunther U, Benson J, Benke D, Fritschy JM, Reyes G, Knoflach F, et al. Benzodiazepine-insensitive mice generated by targeted disruption of the γ2 subunit gene of γ-aminobutyric acid type A receptors. Proc. Natl. Acad. Sci. USA. 1995;92:7749–7753. [PubMed]
174. Pritchett DB, Sontheimer H, Shivers BD, Ymer S, Kettenmann H, Schofield PR, et al. Importance of a novel GABAA receptor subunit for benzodiazepine pharmacology. Nature. 1989;338:582–585. [PubMed]
175. Lakhlani PP, MacMillan LB, Guo TZ, McCool BA, Lovinger DM, Maze M, et al. Substitution of a mutant α2a-adrenergic receptor via ‘hit and run’ gene targeting reveals the role of this subtype in sedative, analgesic and anesthetic-sparing responses in vivo. Proc. Natl. Acad. Sci. USA. 1997;94:9950–9955. [PubMed]
176. Krasowski MD, Rick CE, Harrison NL, Firestone LL, Homanics GE. A deficit of functional GABAA receptors in neurons of β3 subunit knockout mice. Neurosci. Lett. 1998;240:81–84. [PMC free article] [PubMed]
177. Homanics GE, DeLorey TM, Firestone LL, Quinlan JJ, Handforth A, Harrison NL, et al. Mice devoid of γ-aminobutyric type A receptor β3 subunit have epilepsy, cleft palate, and hypersensitive behavior. Proc. Natl. Acad. Sci. USA. 1997;94:4143–4148. [PubMed]
178. Homanics GE, Ferguson C, Quinlan JJ, Daggett J, Snyder K, Lagenaur C, et al. Gene knockout of the α6 subunit of the γ-aminobutyric acid type A receptor: lack of effect on responses to ethanol, pentobarbital, and general anesthesia. Mol. Pharmacol. 1997;51:588–596. [PubMed]
179. Homanics GE, Harrison NL, Quinlan JJ, Krasowski MD, Rick CEM, de Blas AL, et al. Normal electrophysiological and behavioral responses to ethanol in mice lacking the long splice variant of the γ2 subunit of γ-aminobutyrate type A receptor. Neuropharmacol. 1999;38:253–265. [PMC free article] [PubMed]
180. Picciotto MR, Zoli M, Rimondini R, Lena C, Marubio LM, Pich EM, et al. Acetycholine receptors containing the β2 subunit are involved in the reinforcing properties of nicotine. Nature. 1998;391:173–177. [PubMed]
181. Orr-Urtreger A, Goldner FM, Saeki M, Lorenzo I, Goldberg L, De Biasi M, et al. Mice deficient in the α7 neuronal nicotinic acetylcholine receptor lack α-bungarotoxin binding sites and hippocampal fast nicotinic currents. J. Neurosci. 1997;17:9165–9171. [PubMed]
182. Jia Z, Agopyan N, Miu P, Xiong Z, Henderson J, Gerlai R, et al. Enhanced LTP in mice deficient in the AMPA receptor GluR2. Neuron. 1996;17:945–956. [PubMed]
183. Das S, Sasaki YF, Rothe T, Premkumar LS, Takasu M, Crandall JE, et al. Increased NMDA current and spine density in mice lacking the NMDA receptor subunit NR3A. Nature. 1998;393:377–381. [PubMed]
184. Forrest D, Yuzaki M, Soares HD, Ng L, Luk DC, Sheng M, et al. Targeted disruption of NMDAreceptor 1 gene abolishes NMDA response and results in neonatal death. Neuron. 1994;13:325–338. [PubMed]
185. Kadotani H, Hirano T, Masugi M, Nakamura K, Nakao K, Katsuki M, et al. Motor discoordination results from combined gene disruption of the NMDA receptor NR2A and NR2C subunits, but not from single disruption of the NR2A or NR2C subunit. J. Neurosci. 1996;16:7859–7867. [PubMed]
186. Ebralidze AK, Rossi DJ, Tonegawa S, Slater NT. Modification of NMDA receptor channels and synaptic transmission by targeted disruption of the NR2C gene. J. Neurosci. 1996;16:5014–5025. [PubMed]
187. Mulle C, Sailer A, Perez-Otano I, Dickinson-Anson H, Castillo PE, Bureau I, et al. Altered synaptic physiology and reduced susceptibility to kainate-induced seizures in GluR6-deficient mice. Nature. 1998;392:601–605. [PubMed]
188. DeLorey TM, Handforth A, Anagnostaras SG, Homanics GE, Minassian BA, Asatourian A, et al. Mice lacking the β3 subunit of the GABAA receptor have the epilepsy phenotype and many of the behavioral characteristics of Angelman Syndrome. J. Neurosci. 1998;18:8505–8514. [PubMed]
189. Quinlan JJ, Homanics GE, Firestone LL. Anesthesia sensitivity in mice that lack the β3 subunit of the γ-aminobutyric acid type A receptors. Anesthesiol. 1998;88:775–780. [PubMed]
190. Zimmerman SA, Jones MV, Harrison NL. Potentiation of γ-aminobutyric acidA receptor Cl current correlates with in vivo anesthetic potency. J. Pharmacol. Exp. Ther. 1994;270:987–991. [PubMed]
191. Simmonds MA, Turner JP. Potentiators of responses to activation of _-aminobutyric acid (GABAA) receptors. Neuropharmacol. 1987;26:923–930. [PubMed]
192. Franks NP, Dickinson R, de Sousa SLM, Hall AC, Lieb WR. How does xenon produce anaesthesia? Nature. 1998;396:324. [PubMed]
193. Dzoljic M, Van Dujin B. Nitrous oxide-induced enhancement of γ-aminobutyric acidA-mediated chloride currents in acutely dissociated hippocampal neurons. Anesthesiol. 1998;88:473–480. [PubMed]
194. Mennerick S, Jevtovic-Todorovic V, Todorovic SM, Shen WX, Olney JW, Zorumski CF. Effect of nitrous oxide on excitatory and inhibitory synaptic transmission in hippocampal cultures. J. Neurosci. 1998;18:9716–9726. [PubMed]
195. Jevtovic-Todorovic V, Todorovic SM, Mennerick S, Powell S, Dikranian K, Benshoff N, et al. Nitrous oxide (laughing gas) is an NMDA antagonist neuroprotectant, and neurotoxin. Nature Med. 1998;4:460–463. [PubMed]
196. Nicoll RA, Eccles JC, Oshima T, Rubia F. Prolongation of hippocampal inhibitory postsynaptic potentials by barbiturates. Nature. 1975;258:625–627. [PubMed]
197. Scholfield CN. Potentiation of inhibition by general anaesthetics in neurones of the olfactory cortex in vitro. Pflugers Archiv. 1980;383:249–255. [PubMed]
198. Banks MI, Pearce RA. Dual actions of volatile anesthetics on GABAA IPSCs: dissociation of blocking and prolonging effects. Anesthesiol. 1999;90:120–134. [PubMed]
199. Harrison NL, Vicini S, Barker JL. A steroid anesthetic prolongs inhibitory postsynaptic currents in cultured rat hippocampal neurons. J. Neurosci. 1987;7:604–609. [PubMed]
200. MacIver MB, Tanelian DL, Mody I. Two mechanisms for anesthetic-induced enhancement of GABAA-mediated neuronal inhibition. Ann. N. Y. Acad. Sci. 1991;625:91–96. [PubMed]
201. Barker JL, Ransom BR. Pentobarbitone pharmacology of mammalian central neurones grown in tissue culture. J. Physiol. 1978;280:355–372. [PubMed]
202. Robertson B. Actions of anaesthetics and avermectin on GABAA chloride channels in mammalian dorsal root ganglion neurones. Br. J. Pharmacol. 1989;98:167–176. [PMC free article] [PubMed]
203. Rho JM, Donevan SD, Rogawski MA. Direct activation of GABAA receptors by barbiturates in cultured rat hippocampal neurons. J. Physiol. 1996;497:509–522. [PubMed]
204. Hill-Venning C, Belelli D, Peters JA, Lambert JJ. Subunit-dependent interaction of the general anaesthetic etomidate with the γ-aminobutyric acid type A receptor. Br. J. Pharmacol. 1997;120:749–756. [PMC free article] [PubMed]
205. Sanna E, Murgia A, Casula A, Biggio G. Differential subunit dependence of the actions of the general anesthetics alphaxalone and etomidate at γ-aminobutyric acid type A receptors expressed in Xenopus laevis oocytes. Mol. Pharmacol. 1997;51:484–490. [PubMed]
206. Krasowski MD, Koltchine VV, Rick CE, Ye Q, Finn SE, Harrison NL. Propofol and other intravenous anesthetics have sites of action on the γ-aminobutyric acidA receptor distinct from that for isoflurane. Mol. Pharmacol. 1998;53:530–538. [PubMed]
207. Hales TG, Lambert JJ. The actions of propofol on inhibitory amino acid receptors of bovine adrenomedullary chromaffin cells and rodent central neurones. Br. J. Pharmacol. 1991;104:619–628. [PMC free article] [PubMed]
208. Hara M, Kai Y, Ikemoto Y. Propofol activates GABAA receptor-chloride ionophore complex in dissociated hippocampal pyramidal neurons of the rat. Anesthesiol. 1993;79:781–788. [PubMed]
209. Adodra S, Hales TG. Potentiation, activation and blockade of GABAA receptors of clonal murine hypothalamic GT1-7 neurones by propofol. Br. J. Pharmacol. 1995;115:953–960. [PMC free article] [PubMed]
210. Jones MV, Harrison NL, Pritchett DB, Hales TG. Modulation of the GABAA receptor by propofol is independent of the γ subunit. J. Pharmacol. Exp. Ther. 1995;274:962–968. [PubMed]
211. Krasowski MD, O'Shea SM, Rick CEM, Whiting PJ, Hadingham KL, Czajkowski C, et al. α subunit isoform influences GABAA receptor modulation by propofol. Neuropharmacol. 1997;36:941–949. [PMC free article] [PubMed]
212. Callachan H, Cottrell GA, Hather NY, Lambert JJ, Nooney JM, Peters JA. Modulation of the GABAA receptor by progesterone metabolites. Proc. R. Soc. Lond. Ser. B. Biol. Sci. 1987;231:359–369. [PubMed]
213. Belelli D, Callachan H, Hill-Venning C, Peters JA, Lambert JJ. Interaction of positive allosteric modulators with human and Drosophila recombinant GABA receptors expressed in Xenopus laevis oocytes. Br. J. Pharmacol. 1996;118:563–576. [PMC free article] [PubMed]
214. Yang J, Isenberg KE, Zorumski CF. Volatile anesthetics gate a chloride current in postnatal rat hippocampal neurons. FASEB J. 1992;6:914–918. [PubMed]
215. Amin J, Weiss DS. GABAA receptor needs two homologous domains of the _-subunit for activation by GABA but not by pentobarbital. Nature. 1993;366:565–569. [PubMed]
216. Downie DL, Hall AC, Lieb WR, Franks NP. Effects of inhalational general anaesthetics on native glycine receptors in rat medullary neurones and recombinant glycine receptors in Xenopus oocytes. Br. J. Pharmacol. 1996;118:493–502. [PMC free article] [PubMed]
217. Koltchine VV, Ye Q, Finn SE, Harrison NL. Chimeric GABAA/glycine receptors: expression and barbiturate pharmacology. Neuropharmacol. 1996;35:1445–1456. [PubMed]
218. Lu L, Huang Y. Separate domains for desensitization of GABA ρ1 and β2 subunits expressed in Xenopus oocytes. J. Membr. Biol. 1998;164:115–124. [PubMed]
219. Mihic SJ, Ye Q, Wick MJ, Koltchine VV, Krasowski MD, Finn SE, et al. Sites of alcohol and volatile anaesthetic action on GABAA and glycine receptors. Nature. 1997;389:385–389. [PubMed]
220. Wick MJ, Mihic SJ, Ueno S, Mascia MP, Trudell JR, Brozowski SJ, et al. Mutations of γ-aminobutyric acid and glycine receptors change alcohol cutoff: evidence for an alcohol receptor? Proc.Natl. Acad. Sci.USA. 1998;95:6504–6509. [PubMed]
221. Yu D, Zhang L, Eisele JL, Bertrand D, Changeux JP, Weight FF. Ethanol inhibition of nicotinic acetylcholine type α7 receptors involves the amino-terminal domain of the receptor. Mol. Pharmacol. 1996;50:1010–1016. [PubMed]
222. Zhang L, Oz M, Stewart RR, Peoples RW, Weight FF. Volatile general anaesthetic actions on recombinant nACHα7, 5-HT3 and chimeric nAChα7-5-HT3 receptors expressed in Xenopus oocytes. Br. J. Pharmacol. 1997;120:353–355. [PMC free article] [PubMed]
223. Minami K, Wick SJ, Stern-Bach Y, Dildy-Mayfield JE, Brozowski SJ, Gonzales EL, et al. Sites of volatile anesthetic action on kainate (glutamate receptor 6) receptors. J. Biol. Chem. 1998;273:8248–8255. [PubMed]
224. Hackam AS, Wang TL, Guggino WB, Cutting GR. Sequences in the amino termini of GABA ρ and GABAA subunits specify their selective interaction in vitro. J. Neurochem. 1998;70:40–46. [PubMed]
225. Eisele JL, Bertrand S, Galzi JL, Devillers-Thiery A, Changeux JP, Bertrand D. Chimaeric nicotinic serotonergic receptor combines distinct ligand binding and channel specificities. Nature. 1993;366:479–483. [PubMed]
226. Ye Q, Koltchine VV, Mihic SJ, Mascia MP, Wick M, Finn SE, et al. Enhancement of glycine receptor function by ethanol is inversely correlated with molecular volume at position α267. J. Biol. Chem. 1998;273:3314–3319. [PubMed]
227. Krasowski MD, Finn SE, Ye Q, Harrison NL. Trichloroethanol modulation of recombinant GABAA, glycine and GABA ρ1 receptors. J. Pharmacol. Exp. Ther. 1998;284:934–942. [PubMed]
228. Belelli D, Lambert JJ, Peters JA, Wafford K, Whiting PJ. The interaction of the general anesthetic etomidate with the γ-aminobutyric acid type A receptor is influenced by a single amino acid. Proc. Natl. Acad. Sci. USA. 1997;94:11031–11036. [PubMed]
229. McGurk KA, Pistis M, Belelli D, Hope AG, Lambert JJ. The effect of a transmembrane amino acid on etomidate sensitivity of an invertebrate GABA receptor. Br. J. Pharmacol. 1998;124:13–20. [PMC free article] [PubMed]
230. Sonner J, Li J, Eger EI. Desflurane and nitrous oxide, but not nonimmobilizers, affect nociceptive responses. Anesth. Analg. 1998;86:629–634. [PubMed]
231. Kandel L, Chortkoff BS, Sonner J, Laster MJ, Eger EI. Nonanesthetics can suppress learning. Anesth. Analg. 1996;82:321–326. [PubMed]
232. Mihic SJ, McQuilkin SJ, Eger EI, Ionescu P, Harris RA. Potentiation of γ-aminobutyric acid type A receptor-mediated chloride currents by novel halogenated compounds correlates with their abilities to induce general anesthesia. Mol. Pharmacol. 1994;46:851–857. [PubMed]
233. Mascia MP, Machu TK, Harris RA. Enhancement of homomeric glycine receptor function by long-chain alcohols and anaesthetics. Br. J. Pharmacol. 1996;119:1331–1336. [PMC free article] [PubMed]
234. Mihic SJ, Harris RA. Inhibition of ρ1 receptor GABAergic currents by alcohols and volatile anesthetics. J. Pharmacol. Exp. Ther. 1996;277:411–416. [PubMed]
235. Machu TK, Harris RA. Alcohols and anesthetics enhance the function of 5-hydroxytryptamine3 receptors expressed in Xenopus laevis oocytes. J. Pharmacol. Exp. Ther. 1994;271:898–905. [PubMed]
236. Cardoso RA, Brozowski SJ, Chavez-Noriega LE, Harris RA. Human neuronal nicotinic acetylchloline receptors expressed in Xenopus oocytes predict efficacy of halogenated compounds that disobey the Meyer-Overton rule. Anesthesiol. 1999 in press. [PubMed]
237. Dildy-Mayfield JE, Eger EI, Harris RA. Anesthetics produce subunit-selective actions on glutamate receptors. J. Pharmacol. Exp. Ther. 1996;276:1058–1065. [PubMed]
238. Minami K, Vanderah TW, Minami M, Harris RA. Inhibitory effects of anesthetics and ethanol on muscarinic receptors expressed in Xenopus oocytes. Eur. J. Pharmacol. 1997;339:237–244. [PubMed]
239. Forman SA, Raines DE. Nonanesthetic volatile drugs obey the Meyer-Overton correlation in two molecular protein site models. Anesthesiol. 1998;88:1535–1548. [PubMed]
240. Minami K, Gereau RW, Minami M, Heinemann SF, Harris RA. Effects of ethanol and anesthetics on type 1 and 5 metabotropic glutamate receptors expressed in Xenopus laevis oocytes. Mol. Pharmacol. 1998;53:148–156. [PubMed]
241. Cullen SC, Gross EG. The anesthetic properties of xenon in animals and human beings, with additional observations on krypton. Science. 1951;113:580–582. [PubMed]
242. Koblin DD, Fang Z, Eger EI, Laster MJ, Gong D, Ionescu P, et al. Minimum alveolar concentrations of noble gases, nitrogen and sulfur hexafluoride in rats: helium and neon as nonimmobilizers (nonanesthetics). Anesth. Analg. 1998;87:419–424. [PubMed]
243. Pistis M, Belelli D, Peters JA, Lambert JJ. The interaction of general anaesthetics with recombinant GABAA and glycine receptors expressed in Xenopus laevis oocytes: a comparative study. Br. J. Pharmacol. 1997;122:1707–1719. [PMC free article] [PubMed]
244. Moody EJ, Knauer C, Granja R, Strakhova M, Skolnick P. Distinct loci mediate the direct and indirect actions of the anesthetic etomidate at GABAA receptors. J. Neurochem. 1997;69:1310–1313. [PubMed]
245. Sanna E, Garau F, Harris RA. Novel properties of homomeric _1 _-aminobutyric acid type A receptors: actions of the anesthetics propofol and pentobarbital. Mol. Pharmacol. 1995;47:213–217. [PubMed]
246. Sanna E, Mascia MP, Klein RL, Whiting PJ, Biggio G, Harris RA. Actions of the general anesthetic propofol on recombinant human GABAA receptors: influence of receptor subunits. J. Pharmacol. Exp. Ther. 1995;274:353–360. [PubMed]
247. Hu Y, Zorumski CF, Covey DF. Neurosteroid analogues: structure-activity studies of benz[e]indene modulators of GABAA receptor function. 1. The effect of 6-methyl substitution on the electrophysiological activity of 7-substituted benz[e]indene-3-carbonitriles. J. Med. Chem. 1993;36:3956–3967. [PubMed]
248. Rupprecht R, Berning B, Hauser CA, Holsboer F, Reul JM. Steroid receptor-mediated effects of neuroactive steroids: characterization of structure-activity relationship. Eur. J. Pharmacol. 1996;303:227–234. [PubMed]
249. Harrison NL, Majewska MD, Harrington JW, Barker JL. Structure-activity relationships for steroid interaction with the γ-aminobutyric acidA receptor complex. J. Pharmacol. Exp. Ther. 1987;241:346–353. [PubMed]
250. Harrison NL, Simmonds MA. Modulation of the GABA receptor complex by a steroid anaesthetic. Brain Res. 1984;323:287–292. [PubMed]
251. Cottrell GA, Lambert JJ, Peters JA. Modulation of GABAA receptor activity by alphaxalone. Br. J. Pharmacol. 1987;90:491–500. [PMC free article] [PubMed]
252. Zorumski CF, Wittmer LL, Isenberg KE, Hu Y, Covey DF. Effects of neurosteroid and benz[e]indene enantiomers on GABAA receptors in cultured hippocampal neurons and transfected HEK-293 cells. Neuropharmacol. 1996;35:1161–1168. [PubMed]
253. Hill-Venning C, Peters JA, Callachan H, Lambert JJ, Gemmell DK, Anderson A, et al. The anaesthetic action and modulation of GABAA receptor activity by the novel water-soluble aminosteroid Org 20599. Neuropharmacol. 1996;35:1209–1222. [PubMed]
254. Hawkinson JE, Acosta-Burruel M, Yang KC, Hogenkamp DJ, Chen JS, Lan NC, et al. Substituted 3β-phenylethynyl derivatives of 3α-hydroxy-5α-pregnan-20-one: remarkably potent neuroactive steroid modulators of γ-aminobutyric acidA receptors. J. Pharmacol. Exp. Ther. 1998;287:198–207. [PubMed]
255. Rick CE, Ye Q, Finn SE, Harrison NL. Neurosteroids act on the GABAA receptor at sites on the N-terminal side of the middle of TM2. Neuroreport. 1998;9:379–383. [PubMed]
256. Sawada S, Yamamoto C. Blocking action of pentobarbital on receptors for excitatory amino acids in the guinea pig hippocampus. Exp. Brain Res. 1985;59:226–231. [PubMed]
257. Marszalec W, Narahashi T. Use-dependent pentobarbital blockof kainate and quisqualate currents. Brain Res. 1993;608:7–15. [PubMed]
258. Yamakura T, Sakimura K, Mishina M, Shimoji K. The sensitivity of AMPA-sensitive glutamate receptor channels to pentobarbital is determined by a single amino acid residue of the alpha 2 subunit. FEBS Lett. 1995;374:412–414. [PubMed]
259. Lomeli H, Mosbacher J, Melcher T, Hoger T, Geiger JR, Kuner T, et al. Control of kinetic properties of AMPA receptor channels by nuclear RNA editing. Science. 1994;266:1709–1713. [PubMed]
260. Sommer B, Kohler M, Sprengel R, Seeburg PH. RNA editing in brain controls a determinant of ion flow in glutamate-gated channels. Cell. 1991;67:11019. [PubMed]
261. Birnir B, Tierney ML, Dalziel JE, Cox GB, Gage PW. A structural determinant of desensitization and allosteric regulation by pentobarbital of the GABAA receptor. J. Membr. Biol. 1997;155:157–166. [PubMed]
262. Winters WD, Ferrar-Allado T, Guzman-Flores C, Alcaraz M. The cataleptic state induced by ketamine: a review of the neuropharmacology of anesthesia. Neuropharmacol. 1972;11:303–315. [PubMed]
263. Orser BA, Pennefather PS, MacDonald JF. Multiple mechanisms of ketamine blockade of N-methyl-D-aspartate receptors. Anesthesiol. 1997;86:903–917. [PubMed]
264. Anis NA, Berry SC, Burton NR, Lodge D. The dissociative anaesthetics, ketamine and phencyclidine, selectively reduce excitation of central mammalian neurones by N-methyl-aspartate. Br. J. Pharmacol. 1983;79:565–575. [PMC free article] [PubMed]
265. Zeilhofer HU, Swandulla D, Geisslinger G, Brune K. Differential effects of ketamine enantiomers on NMDA receptor currents in cultured neurons. Eur. J. Pharmacol. 1992;213:155–158. [PubMed]
266. Nagata K, Aistrup GL, Huang CS, Marszalec W, Song JH, Yeh JZ, et al. Potent modulation of neuronal nicotinic acetylcholine receptor-channel by ethanol. Neurosci. Lett. 1996;217:189–193. [PubMed]
267. Covernton PJ, Connolly JG. Differential modulation of rat neuronal nicotinic receptor subtypes by acute application of ethanol. Br. J. Pharmacol. 1997;122:1661–1668. [PMC free article] [PubMed]
268. Deitrich RA, Harris RA. How much alcohol should I use in my experiments? Alcoholism Clin. Exp. Res. 1996;20:1–2. [PubMed]
269. McCreery MJ, Hunt WA. Physico-chemical correlates of alcohol intoxication. Neuropharmacol. 1978;17:451–461. [PubMed]
270. Lyon RC, McComb JA, Schreurs J, Goldstein DB. A relationship between alcohol intoxication and the disordering of brain membranes by a series of short-chain alcohols. J. Pharmacol. Exp. Ther. 1981;218:669–675. [PubMed]
271. Dildy-Mayfield JE, Mihic SJ, Liu Y, Deitrich RA, Harris RA. Actions of long chain alcohols on GABAA and glutamate receptors: relation to in vivo effects. Br. J. Pharmacol. 1996;118:378–384. [PMC free article] [PubMed]
272. Franks NP, Jenkins A, Conti E, Lieb WR, Brick P. Structural basis for the inhibition of firefly luciferase by a general anesthetic. Biophys. J. 1998;75:2205–2211. [PubMed]
273. Doyle DA, Cabral JM, Pfuetzner RA, Kuo A, Gulbis JM, Cohen SL, et al. The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science. 1998;280:69–77. [PubMed]
274. Armstrong N, Sun Y, Chen GQ, Gouax E. Structure of a glutamate-receptor ligand-binding core in complex with kainate. Nature. 1998;395:913–917. [PubMed]
275. Giese JL, Stanley TH. Etomidate: a new intravenous anesthetic induction agent. Pharmacotherapy. 1983;3:251–258. [PubMed]
276. Lauven PM, Schwilden H, Stoeckel H. Threshold hypnotic concentration of methohexitone. Eur. J. Clin. Pharmacol. 1987;33:261–265. [PubMed]
277. Idvall J, Ahlgren I, Aronsen KF, Stenberg P. Ketamine infusions: pharmacokinetics and clinical effects. Br. J. Anaesth. 1979;51:1167–1172. [PubMed]
278. Fang Z, Ionescu P, Chortkoff BS, Kandel L, Sonner J, Laster MJ, et al. Anesthetic potencies of n-alkanols: results of additivity and solubility studies suggest a mechanism of action similar to that for conventional inhaled anesthetics. Anesth. Analg. 1997;84:1042–1048. [PubMed]
279. McKenzie D, Franks NP, Lieb WR. Actions of general anaesthetics on a neuronal nicotinic acetylcholine receptor in isolated identified neurones of Lymnaea stagnalis. Br. J. Pharmacol. 1995;115:275–282. [PMC free article] [PubMed]
280. Hadingham KL, Wingrove P, Le Bourdelles B, Palmer KJ, Ragan CI, Whiting PJ. Cloning of cDNA sequences encoding human α2 and α3 γ-aminobutyric acida receptor subunits and characterization of the benzodiazepine pharmacology of recombinant α1-, α2-, α3- and α5-containing human γ-aminobutyric acidA receptors. Mol. Pharmacol. 1993;43:970–975. [PubMed]
281. Hadingham KL, Wingrove PB, Wafford KA, Bain C, Kemp JA, Palmer KJ, et al. Role of the β subunit in determining the pharmacology of human γ-aminobutyric acid type A receptors. Mol. Pharmacol. 1993;44:1211–1218. [PubMed]
282. Xu M, Akabas MH. Identification of channel-lining residues in the M2 membrane-spanning segment of the GABAA receptor α1 subunit. J. Gen. Physiol. 1996;107:195–205. [PMC free article] [PubMed]
283. Gurley D, Amin J, Ross PC, Weiss DS, White G. Point mutations in theM2 region of the α, β, or γ subunit of the GABAA channel that abolish block by picrotoxin. Receptors Channels. 1995;3:13–20. [PubMed]
284. Wingrove PB, Wafford KA, Bain C, Whiting PJ. The modulatory action of loreclezole at the γ-aminobutyric acid type A receptor is determined by a single amino acid in the β2 and β3 subunit. Proc. Natl. Acad. Sci. USA. 1994;91:4569–4573. [PubMed]
285. Horenstein J, Akabas MH. Location of a high affinity Zn2+ binding site in the channel of α1β1 GABAA receptors. Mol. Pharmacol. 1998;53:870–877. [PubMed]
286. Miller KW, Smith EB. Intermolecular forces and the pharmacology of simple molecules. In: Featherstone RM, editor. A Guide to Molecular Pharmacology – Toxicology, part 2. Marcel Dekker; New York: 1973. pp. 427–475.
287. Eger EI, Lundgren C, Miller SL, Stevens WC. Anesthetic potencies of sulfur hexafluoride, carbon tetrafluoride, chloroform and Ethrane in dogs: correlation with the hydrate and lipid theories of anesthetic action. Anesthesiol. 1969;30:129–135. [PubMed]
288. Steward A, Allott PR, Cowles AL, Mapleson WW. Solubility coefficients for inhaled anaesthetics for water, oil and biological media. Br. J. Anaesth. 1973;45:282–293. [PubMed]
289. Mazze RI, Rice SA, Baden JM. Halothane, isoflurane, and enflurane MAC in pregnant and nonpregnant female and male mice and rats. Anesthesiol. 1985;62:339–341. [PubMed]
290. Koblin DD. Mechanisms of action. In: Miller RD, editor. Anesthesia. Churchill Livingstone; New York: 1994. pp. 67–69.
291. Crawford MW, Lerman J, Saldivia V, Carmichael FJ. Hemodynamic and organ blood flow responses to halothane and sevoflurane anesthesia during spontaneous ventilation. Anesth. Analg. 1992;75:1000–1006. [PubMed]
292. Kazama T, Ikeda K. Comparison of MAC and the rate of rise of alveolar concentration of sevoflurane with halothane and isoflurane in the dog. Anesthesiol. 1988;68:435–437. [PubMed]
293. Scheller MS, Saidman LJ, Partridge BL. MAC of sevoflurane in humans and the New Zealand white rabbit. Can. J. Anaesth. 1988;35:153–156. [PubMed]
294. Halsey MJ, Wardley-Smith B, Wood S. Pressure reversal of alphaxalone/alphadolone and methohexitone in tadpoles: evidence for different molecular sites for general anaesthesia. Br. J. Pharmacol. 1986;89:299–305. [PMC free article] [PubMed]
295. Tonner PH, Scholz J, Lamberz L, Schlamp N, Schulte J. Inhibition of nitric oxide synthase decreases anesthetic requirements of intravenous anesthetics in Xenopus laevis. Anesthesiol. 1997;87:1479–1485. [PubMed]
296. Pringle MJ, Brown KB, Miller KW. Can the lipid theories of anesthesia account for the cutoff in anesthetic potency in homologous series of alcohols? Mol. Pharmacol. 1981;19:49–55. [PubMed]
297. Brink F, Posternak JM. Thermodynamic analysis of the relative effectiveness of narcotics. J. Cell. Comput. Physiol. 1948;32:211–233. [PubMed]
298. Marshall EK, Owens AH. Absorption, excretion, and metabolic fate of chloral hydrate and trichloroethanol. Bull. Johns Hopkins Hosp. 1954;95:1–18. [PubMed]
299. Breimer DD. Clinical pharmacokinetics of hypnotics. Clin. Pharmacokinet. 1977;2:93–109. [PubMed]
300. Garrett ER, Lambert HJ. Pharmacokinetics of trichloroethanol and metabolites and interconversions among variously referenced pharmacokinetic parameters. J. Pharm. Sci. 1973;62:550–572. [PubMed]
301. Daniels S, Roberts RJ. Post-synaptic inhibitory mechanisms of anaesthesia; glycine receptors. Toxicol. Lett. 1998;100:71–76. [PubMed]
302. Carla V, Moroni F. General anaesthetics inhibit the responses induced by glutamate receptor agonists in the mouse cortex. Neurosci. Lett. 1992;146:21–24. [PubMed]
303. Lin LH, Chen LL, Zirrolli JA, Harris RA. General anesthetics potentiate γ-aminobutyric acid actions on γ-aminobutyric acidA receptors expressed by Xenopus oocytes: lack of involvement of intracellular calcium. J. Pharmacol. Exp. Ther. 1992;263:569–578. [PubMed]
304. Dilger JP, Liu Y, Vidal AM. Interactions of general anaesthetics with single acetylcholine receptor channels. Eur. J. Anaesth. 1995;12:31–39. [PubMed]
305. Zhou Q, Lovinger DM. Pharmacologic characteristics of potentiation of 5-HT3 receptors by alcohols and diethyl ether in NCB-20 neuroblastoma cells. J. Pharmacol. Exp. Ther. 1996;278:732–740. [PubMed]
306. Lin LH, Whiting P, Harris RA. Molecular determinants of general anesthetic action: role of GABAA receptor structure. J. Neurochem. 1993;60:1548–1553. [PubMed]
307. Wakamori M, Ikemoto Y, Akaike N. Effects of two volatile anesthetics and a volatile convulsant on the excitatory and inhibitory amino acid responses in dissociated CNS neurons of the rat. J. Neurophysiol. 1991;66:2014–2021. [PubMed]
308. Jones MV, Brooks PA, Harrison NL. Enhancement of γ-aminobutyric acid-activated Cl currents in cultured rat hippocampal neurones by three volatile anaesthetics. J. Physiol. 1992;449:279–293. [PubMed]
309. Nakahiro M, Yeh JZ, Brunner E, Narahashi T. General anesthetics modulate GABA receptor channel complex in rat dorsal root ganglion neurons. FASEB J. 1989;3:1850–1854. [PubMed]
310. Wachtel RE. Relative potencies of volatile anesthetics in altering the kinetics of ion channels in BC3H1 cells. J. Pharmacol. Exp. Ther. 1995;274:1355–1361. [PubMed]
311. Jenkins A, Franks NP, Lieb WR. Actions of general anaesthetics on 5-HT3 receptors in N1E-115 neuroblastoma cells. Br. J. Pharmacol. 1996;117:1507–1515. [PMC free article] [PubMed]
312. Lin LH, Chen LL, Harris RA. Enflurane inhibits NMDA, AMPA, and kainate-induced currents in Xenopus oocytes expressing mouse and human brain mRNA. FASEB J. 1993;7:479–485. [PubMed]
313. Ikemoto Y, Yamashita M, Yano T. Volatile anesthetics and a volatile convulsant differentially affect GABAA receptor-chloride channel complex. Toxicol. Lett. 1998;101:225–231. [PubMed]
314. Mascia MP, Wick MJ, Martinez LD, Harris RA. Enhancement of glycine receptor function by ethanol: role of phosphorylation. Br. J. Pharmacol. 1998;125:263–270. [PMC free article] [PubMed]
315. Violet JM, Downie DL, Nakisa RC, Lieb WR, Franks NP. Differential sensitivities of mammalian neuronal and muscle nicotinic acetylcholine receptors to general anesthetics. Anesthesiol. 1997;86:866–874. [PubMed]
316. Dickinson R, Lieb WR, Franks NP. The effects of temperature on the interactions between volatile general anaesthetics and a neuronal nicotinic acetylcholine receptor. Br. J. Pharmacol. 1995;116:2949–2956. [PMC free article] [PubMed]
317. Weight FF, Peoples RW, Wright JM, Lovinger DM, White G. Ethanol action on excitatory amino acid activated ion channels. Alcohol Alcoholism Suppl. 1993;2:353–358. [PubMed]
318. Harrison NL, Kugler JL, Jones MV, Greenblatt EP, Pritchett DB. Positive modulation of human γ-aminobutyric acid type A and glycine receptors by the inhalation anesthetic isoflurane. Mol. Pharmacol. 1993;44:628–632. [PubMed]
319. Lees G, Edwards MD. Modulation of recombinant human _-aminobutyric acidA receptors by isoflurane: influence of the delta subunit. Anesthesiol. 1998;88:206–217. [PubMed]
320. Scheller M, Bufler J, Schneck H, Kochs E, Franke C. Isoflurane and sevoflurane interact with the nicotinic acetylcholine receptor channels in micromolar concentrations. Anesthesiol. 1997;86:118–127. [PubMed]
321. Flood P, Ramirez-Latorre J, Role L. α4β2 Neuronal nicotinic acetylcholine receptors in the central nervous system are inhibited by isoflurane and propofol, but α7-type nicotinic acetylcholine receptors are unaffected. Anesthesiol. 1997;86:859–865. [PubMed]
322. Charlesworth P, Richards CD. Anaesthetic modulation of nicotinic ion channel kinetics in bovine chromaffin cells. Br. J. Pharmacol. 1995;114:909–917. [PMC free article] [PubMed]
323. Kira T, Harata N, Sakata T, Akaike N. Kinetics of sevoflurane action on GABA- and glycine-induced currents in acutely dissociated rat hippocampal neurons. Neuroscience. 1998;85:383–394. [PubMed]
324. Wu J, Harata N, Akaike N. Potentiation by sevoflurane of the γ-aminobutyric acid-induced chloride current in acutely dissociated CA1 pyramidal neurones from rat hippocampus. Br. J. Pharmacol. 1996;119:1013–1021. [PMC free article] [PubMed]
325. Horne AL, Hadingham KL, Macaulay AJ, Whiting P, Kemp JA. The pharmacology of recombinant GABAA receptors containing bovine α1, β1, γ2L sub-units stably transfected into mouse fibroblast L-cells. Br. J. Pharmacol. 1992;107:732–737. [PMC free article] [PubMed]
326. Peters JA, Kirkness EF, Callachan H, Lambert JJ, Turner AJ. Modulation of the GABAA receptor by depressant barbiturates and pregnane steroids. Br. J. Pharmacol. 1988;94:1257–1269. [PMC free article] [PubMed]
327. Thompson SA, Whiting PJ, Wafford KA. Barbiturate interactions at the human GABAA receptor: dependence on receptor subunit combination. Br. J. Pharmacol. 1996;117:521–527. [PMC free article] [PubMed]
328. Parker I, Gundersen CB, Miledi R. Actions of pentobarbital on rat brain receptors expressed in Xenopus oocytes. J. Neurosci. 1986;6:2290–2297. [PubMed]
329. Shimada S, Cutting G, Uhl GR. _-Aminobutyric acid A or C receptor? γ-Aminobutyric acid ρ1 receptor RNA induces bicuculline-, barbiturate-, and benzodiazepine-insensitive γ-aminobutyric acid responses in Xenopus oocytes. Mol. Pharmacol. 1992;41:683–687. [PubMed]
330. Gage PW, McKinnon D. Effects of pentobarbitone on acetylcholine-activated channels in mammalian muscle. Br. J. Pharmacol. 1985;85:229–235. [PMC free article] [PubMed]
331. Dilger JP, Boguslavsky R, Barann M, Katz T, Vidal AM. Mechanisms of barbiturate inhibition of acetylcholine receptor channels. J. Gen. Physiol. 1997;109:401–414. [PMC free article] [PubMed]
332. Barann M, Gothert M, Bonisch H, Dybek A, Urban BW. 5-HT3 receptors in outside-out patches of N1E-115 neuroblastoma cells: basic properties and effects of pentobarbital. Neuropharmacol. 1997;36:655–664. [PubMed]
333. Yang J, Uchida I. Mechanisms of etomidate potentiation of GABAA receptor-gated currents in cultured postnatal hippocampal neurons. Neuroscience. 1996;73:69–78. [PubMed]
334. Uchida I, Kamatchi G, Burt D, Yang J. Etomidate potentiation of GABAA receptor gated current depends . on the subunit composition. Neurosci. Lett. 1995;185:203–206. [PubMed]
335. Wachtel RE, Wegrzynowicz ES. Kinetics of nicotinic acetylcholine ion channels in the presence of intravenous anaesthetics and induction agents. Br. J. Pharmacol. 1992;106:623–627. [PMC free article] [PubMed]
336. Appadu BL, Lambert DG. Interaction of i.v. anaesthetic agents with 5-HT3 receptors. Br. J. Anaesth. 1996;76:271–273. [PubMed]
337. Scheller M, Bufler J, Hertle I, Schneck HJ, Franke C, Kochs E. Ketamine blocks currents through mammalian nicotinic acetylcholine receptor channels by interaction with both the open and the closed state. Anesth. Analg. 1996;83:830–836. [PubMed]
338. Malone HM, Peters JA, Lambert JJ. Physiological and pharmacological properties of 5-HT3 receptors – a patch clamp-study. Neuropeptides. 1991;19:25–30. [PubMed]
339. Uchida I, Li L, Yang J. The role of the GABAA receptor α1 subunit N-terminal extracellular domain in propofol potentiation of chloride current. Neuropharmacol. 1997;36:1611–1617. [PubMed]
340. Hara M, Kai Y, Ikemoto Y. Enhancement by propofol of the γ-aminobutyric acidA response in dissociated hippocampal pyramidal neurons of the rat. Anesthesiol. 1994;81:988–994. [PubMed]
341. Orser BA, Wang LY, Pennefather PS, MacDonald JF. Propofol modulates activation and desensitization of GABAA receptors in cultured murine hippocampal neurons. J. Neurosci. 1994;14:7747–7760. [PubMed]
342. Yamakura T, Sakimura K, Shimoji K, Mishina M. Effects of propofol on various AMPA-, kainate- and NMDA selective glutamate receptor channels expressed in Xenopus oocytes. Neurosci. Lett. 1995;188:187–190. [PubMed]
343. Orser BA, Bertlik M, Wang LY, MacDonald JF. Inhibition by propofol (2,6-di-isopropylphenol) of the N-methyl-D-aspartate subtype of glutamate receptor in cultured hippocampal neurones. Br. J. Pharmacol. 1995;116:1761–1768. [PMC free article] [PubMed]
344. Barker JL, Harrison NL, Lange GD, Owen DG. Potentiation of _-aminobutyric-acid-activated chloride conductance by a steroid anaesthetic in cultured rat spinal neurones. J. Physiol. 1987;386:485–501. [PubMed]
345. Prince RJ, Simmonds MA. Steroid modulation of the strychnine-sensitive glycine receptor. Neuropharmacol. 1992;31:201–205. [PubMed]
346. Feigenspan A, Wassle H, Bormann J. Pharmacology ofGABA receptorCl− channels in rat retinal bipolar cells. Nature. 1993;361:159–162. [PubMed]
347. Ueno S, Wick MJ, Ye Q, Harrison NL, Harris RA. Subunit mutations affect ethanol actions on GABAA receptors expressed in Xenopus oocytes. Br. J. Pharmacol. 1999 in press. [PMC free article] [PubMed]
348. Ueno S, Trudell JR, Eger EI, Harris RA. Actions of fluorinated alkanols on GABAA receptors: relevance to theories of narcosis. Anesth. Analg. 1999;88:877–883. [PubMed]
349. Aguayo LG, Pancetti FC. Ethanol modulation of the γ-aminobutyric acidA- and glycine-activated Cl current in cultured mouse neurons. J. Pharmacol. Exp. Ther. 1994;270:61–69. [PubMed]
350. Mihic SJ, Whiting PJ, Harris RA. Anaesthetic concentrations of alcohols potentiate GABAA receptor-mediated currents: lack of subunit specificity. Eur. J. Pharmacol. 1994;268:209–214. [PubMed]
351. Nakahiro M, Arakawa O, Narahashi T. Modulation of γ-aminobutyric acid receptor-channel complex by alcohols. J. Pharmacol. Exp. Ther. 1991;259:235–240. [PubMed]
352. Marszalec W, Kurata Y, Hamilton BJ, Carter DB, Narahashi T. Selective effects of alcohols on γ-aminobutyric acid A receptor subunits expressed in human embryonic kidney cells. J. Pharmacol. Exp. Ther. 1994;269:157–163. [PubMed]
353. Whitten RJ, Maitra R, Reynolds JN. Modulation of GABAA receptor function by alcohols: effects of subunit composition and differential effects of ethanol. Alcoholism Clin. Exp. Res. 1996;20:1313–1319. [PubMed]
354. Aguayo LG, Tapia JC, Pancetti FC. Potentiation of the glycine-activated Cl current by ethanol in cultured mouse spinal neurons. J. Pharmacol. Exp. Ther. 1996;279:1116–1122. [PubMed]
355. Mascia MP, Mihic SJ, Valenzuela CF, Schofield PR, Harris RA. A single amino acid determines differences in ethanol actions on strychnine-sensitive glycine receptors. Mol. Pharmacol. 1996;50:402–406. [PubMed]
356. Bradley RJ, Sterz R, Peper K. The effects of alcohols and diols at the nicotinic acetylcholine receptor of the neuromuscular junction. Brain Res. 1984;295:101–112. [PubMed]
357. Cardoso RA, Brozowski SJ, Chavez-Noriega LE, Harpold M, Valenzuela CF, Harris RA. Effects of ethanol on recombinant human neuronal nicotinic acetylcholine receptors expressed in Xenopus oocytes. J. Pharmacol. Exp. Ther. 1999;289:774–780. [PubMed]
358. Lovinger DM, White G. Ethanol potentiation of 5-hydroxytryptamine3 receptor-mediated ion current in neuroblastoma cells and isolated adult mammalian neurons. Mol. Pharmacol. 1991;40:263–270. [PubMed]
359. Lovinger DM. Ethanol potentiation of 5-HT3 receptor-mediated ion current in NCB-20 neuroblastoma cells. Neurosci. Lett. 1991;122:57–60. [PubMed]
360. Aoshima H. Effects of alcohols and food additives on glutamate receptors expressed in Xenopus oocytes: specificity in the inhibition of the receptors. Biosci. Biotechnol. Biochem. 1996;60:434–438. [PubMed]
361. Dildy-Mayfield JE, Harris RA. Ethanol inhibits kainate responses of glutamate receptors expressed in Xenopus oocytes: role of calcium and protein kinase C. J. Neurosci. 1995;15:3162–3171. [PubMed]
362. Wright JM, Peoples RW, Weight FF. Single-channel and whole-cell analysis of ethanol inhibition of NMDA-activated currents in cultured mouse cortical and hippocampal neurons. Brain Res. 1996;738:249–256. [PubMed]
363. Koltchine V, Anantharam V, Wilson A, Bayley H, Treistman SN. Homomeric assemblies of NMDAR1 splice variants are sensitive to ethanol. Neurosci. Lett. 1993;152:13–16. [PubMed]
364. Lovinger DM, White G, Weight FF. Ethanol inhibits NMDA-activated ion current in hippocampal neurons. Science. 1989;243:1721–1724. [PubMed]
365. Peoples RW, White G, Lovinger DM, Weight FF. Ethanol inhibition of N-methyl-D-aspartate-activated current in mouse hippocampal neurones: whole-cell patch-clamp analysis. Br. J. Pharmacol. 1997;122:1035–1042. [PMC free article] [PubMed]
366. Nakahiro M, Arakawa O, Nishimura T, Narahashi T. Potentiation of GABA-induced Cl current by a series of n-alcohols disappears at a cutoff point of a longer-chain n-alcohol in rat dorsal root ganglion neurons. Neurosci. Lett. 1996;205:127–130. [PubMed]
367. Murrell RD, Braun MS, Haydon DA. Actions of n-alcohols on nicotinic acetylcholine receptor channels in cultured rat myotubes. J. Physiol. 1991;437:431–448. [PubMed]
368. Peoples RW, Weight FF. Cutoff in potency implicates alcohol inhibition of N-methyl-D-aspartate receptors in alcohol intoxication. Proc. Natl. Acad. Sci. USA. 1995;92:2825–2829. [PubMed]
369. McLarnon JG, Wong JH, Sawyer D, Baimbridge KG. The actions of intermediate and long-chain n-alkanols on unitaryNMDAcurrents in hippocampal neurons. Can. J. Physiol. Pharmacol. 1991;69:1422–1427. [PubMed]
370. Garrett KM, Gan JP. Enhancement of γ-aminobutyric acidA receptor activity by α-chloralose. J. Pharmacol. Exp. Ther. 1998;285:680–686. [PubMed]
371. Lovinger DM, Zimmerman SA, Levitin M, Jones MV, Harrison NL. Trichloroethanol potentiates synaptic transmission mediated by γ-aminobutyric acidA receptors in hippocampal neurons. J. Pharmacol. Exp. Ther. 1993;264:1097–1103. [PubMed]
372. Peoples RW, Weight FF. Trichloroethanol potentiation of γ-aminobutyric acid-activated chloride current in mouse hippocampal neurones. Br. J. Pharmacol. 1994;113:555–563. [PMC free article] [PubMed]
373. Zhou Q, Verdoorn TA, Lovinger DM. Alcohols potentiate the function of 5-HT3 receptor-channels on NCB-20 neuroblastoma cells by favouring and stabilizing the open channel state. J. Physiol. 1998;507:335–352. [PubMed]
374. Lovinger DM, Zhou Q. Trichloroethanol potentiation of 5-hydroxytryptamine3 receptor-mediated ion current in nodose ganglion neurons from the adult rat. J. Pharmacol. Exp. Ther. 1993;265:771–776. [PubMed]
375. Downie DL, Hope AG, Belelli D, Lambert JJ, Peters JA, Bentley KR, et al. The interaction of trichloroethanol with murine recombinant 5-HT3 receptors. Br. J. Pharmacol. 1995;114:1641–1651. [PMC free article] [PubMed]
376. Peoples RW, Weight FF. Inhibition of excitatory amino acid-activated currents by trichloroethanol and trifluoroethanol in mouse hippocampal neurones. Br. J. Pharmacol. 1998;124:1159–1164. [PMC free article] [PubMed]