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doi: 10.1097/ALN.0b013e31827e52c6
Perioperative Medicine

Drosophila Ryanodine Receptors Mediate General Anesthesia by Halothane

Gao, Shuying Ph.D.*; Sandstrom, David J. Ph.D.; Smith, Harold E. Ph.D.; High, Brigit B.A.§; Marsh, Jon W. Ph.D.; Nash, Howard A. M.D., Ph.D.#

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Background: Although in vitro studies have identified numerous possible targets, the molecules that mediate the in vivo effects of volatile anesthetics remain largely unknown. The mammalian ryanodine receptor (Ryr) is a known halothane target, and the authors hypothesized that it has a central role in anesthesia.
Methods: Gene function of the Drosophila Ryr (dRyr) was manipulated in the whole body or in specific tissues using a collection of mutants and transgenes, and responses to halothane were measured with a reactive climbing assay. Cellular responses to halothane were studied using Ca2+ imaging and patch clamp electrophysiology.
Results: Halothane potency strongly correlates with dRyr gene copy number, and missense mutations in regions known to be functionally important in the mammalian Ryrs gene cause dominant hypersensitivity. Tissue-specific manipulation of dRyr shows that expression in neurons and glia, but not muscle, mediates halothane sensitivity. In cultured cells, halothane-induced Ca2+ efflux is strictly dRyr-dependent, suggesting a close interaction between halothane and dRyr. Ca2+ imaging and electrophysiology of Drosophila central neurons reveal halothane-induced Ca2+ flux that is altered in dRyr mutants and correlates with strong hyperpolarization.
Conclusions: In Drosophila, neurally expressed dRyr mediates a substantial proportion of the anesthetic effects of halothane in vivo, is potently activated by halothane in vitro, and activates an inhibitory conductance. The authors’ results provide support for Ryr as an important mediator of immobilization by volatile anesthetics.
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What We Already Know about This Topic

* Ryanodine receptors are involved in the pathogenesis of malignant hyperthermia, but their role in neurologic effects of anesthetics is less clear
* Model organisms provide a powerful genetics-based approach for studying anesthetic targets
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What This Article Tells Us That Is New

* Halothane immobilization of Drosophila flies was bidirectionally affected by mutations in the ryanodine receptor gene
* Immobilization required neurally expressed ryanodine receptors and halothane increased intracellular calcium, a mechanism that might also be relevant to anesthetic actions in vertebrates
VOLATILE general anesthetics are simple organic compounds that rapidly and reversibly suppress responsiveness to external stimuli in all animals while leaving basal functions intact. These properties have rendered anesthetics indispensable in the clinic and made them attractive tools for studying the control of arousal. One obstacle to progress is that, despite considerable effort over many years, the molecular pathways through which volatile anesthetics act remain largely unknown. In vitro studies have identified many proteins whose activity is affected by clinical concentrations of volatile general anesthetics,1 but validating these candidates in vivo has proven difficult.
Studies in genetic model organisms such as worms, flies, and mice have been instrumental in identifying novel targets of volatile anesthetics and have provided in vivo validation of candidates identified in vitro.2–6 These mutants illustrate several features of the molecular pathways mediating anesthesia. First, voltage-insensitive (“leak”) ion channels, which act to hyperpolarize neurons, have repeatedly appeared in genetic screens for targets of volatile anesthetics,2,3,5,7 suggesting that they act, at least in part, through changes in voltage and resistance across the plasma membrane. Second, the response to each anesthetic compound is affected differently by a given mutant, suggesting the presence of agent-specific pathways.2,8 Finally, mutations in these genes reduce but do not eliminate sensitivity to volatile anesthetics, indicating that additional anesthetic targets remain unidentified.
The ryanodine receptor (Ryr), a large-conductance channel that mediates release of Ca2+ from internal stores, is known to be activated by volatile anesthetics, especially halothane.9,10 However, the study of the interaction between halothane and Ryr has been largely limited to muscle, where halothane potently induces malignant hyperthermia, a pathologic condition defined by inappropriate activation of Ryr in susceptible humans and swine.11 Despite the clear connection between Ryr and halothane, the role of neurally expressed Ryr in the immobilizing effects of halothane is unexplored.
In this study, we use Drosophila genetics to test the hypothesis that Ryr mediates halothane anesthesia. We find that dRyr is a major determinant of halothane’s anesthetic effects in flies and that these effects are mediated by expression of dRyr in the nervous system, but not muscle.
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Materials and Methods

Fly Stocks and Genetic Analysis
All flies were reared on cornmeal-molasses medium and maintained at 25°C in a 12-h light–dark cycle. The following fly strains were used in this study: dRyrk04913, Appl-GAL4, nrv2-GAL4, cn1 l(2)44Fp1 bw1 sp1/SM6a, cn1 l(2)44Fl1 bw1 sp1/SM6a, l(2)44Fa3/CyO, l(2)44Ff1/CyO, l(2)44Fg1/CyO, l(2)44Fh1/CyO, and l(2)44Fj1/CyO (Drosophila Stock Center, Bloomington, IN); dRyrGS21220 (Drosophila Genetic Resource Center, Kyoto, Japan); and UAS-dRyrRNAi (10844R-3, National Institute of Genetics, Japan). UAS-GCaMP3 was a generous gift from the Howard Hughes Medical Institute (Janelia Farm, Ashburn, VA); elav-GAL4 was a generous gift from Ravi Allada, Ph.D. (Professor, University of Chicago, Chicago, IL); repo-GAL4 was from Chi-Hon Lee, Ph.D. (Senior Investigator, National Institute of Child Health and Human Development, Bethesda, MD); MHC-GAL4 was from Benjamin White, Ph.D. (Senior Investigator, National Institute of Mental Health, Bethesda, MD); and ShakB-GAL4 and Canton-S were from the lab stock collection. dRyrk04913, dRyrGS21220, dRyrδ25, elav-GAL4, nrv2-GAL4, and UAS-dRyrRNAi were made congenic with Canton-S by outcrossing for seven generations. As described previously,12 dRyrk04913 was homozygous lethal. However, the lethality was not rescued by the genomic duplication dRyr24D03, indicating that it resulted from a closely linked mutation outside the dRyr locus.
Behavioral assays of anesthesia are subject to a number of confounding factors, including genetic background and subtle environmental variations. To avoid such confounding factors, all comparisons were made within experiments run in parallel over the same time and only between groups matched for genetic background.
The deletion mutant dRyrδ25 was generated by excising the region between transposons P{XP}d03686 and P{XP}d03830 (Exelixis Stock Collection, Boston, MA), using flippase-mediated recombination at flippase recombination target sites present in these elements.13 The deletion was verified using polymerase chain reaction amplification and DNA sequencing.
The genomic duplication dRyr24D03 was generated by integrating the P[acman] clone CH321-24D03 (Bellen Laboratory, Baylor College of Medicine, Houston, TX) into the VK33 docking site on the third chromosome via φC31-mediated site-specific integration.14 UAS-dRyr-V5 was generated by cloning a 15.6-kb PacI-PsiI fragment from DRyR-15B (kindly provided by Daniel Cordova, M.Sc., Senior Biochemist, Dupont Crop Protection, Newark, DE) into the pUAST vector, and injecting the construct into Canton-S flies that carried the w1118 mutation. All injections were performed by Rainbow Transgenic Flies, Inc. (Camarillo, CA).
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Genomic Sequencing to Identify dRyr Mutation Sites
The genomic DNA libraries from both control and mutant strains used in this study were constructed and sequenced in accordance with the manufacturer’s protocols (Illumina, San Diego, CA). Briefly, 1 μg of genomic DNA was sheared by sonication, end-repaired, A-tailed, adapter-ligated, size-fractionated by gel electrophoresis, and amplified by polymerase chain reaction. Paired-end 101-cycle sequencing on a HiSeq 2000 instrument (Illumina) yielded an average of 20 million reads (range, 13–30 million reads) equivalent to approximately 29-fold coverage (range, 19- to 43-fold). Reads were aligned to the Drosophila melanogaster reference genome (dm3; Berkeley Drosophila Genome Project Release 5) with BFAST.15 SAMTools16 was used to call sequence variants, and nonsynonymous mutations were identified using ANNOVAR.17
To generate a list of unique candidate mutations, we filtered sequence variants in the mutant strains against those present in the control strain. This process revealed that only dRyr contained sequence variants in all of the dRyr alleles carrying the ethylmethanesulfonate mutations. Pairwise alignments were carried out with mammalian Ryr isoforms using BLASTP.
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Tissue Homogenates and Western Blots
After flies were tested in the reactive climbing assay, 200 fly heads or 50–100 whole flies from each genotype were used to prepare homogenates. The samples were homogenized in 100–200 μl of homogenization buffer (0.25 M sucrose, 10 mM Tris, and 1 mM EDTA) and protease inhibitors (Roche, Indianapolis, IN). The homogenates were centrifuged for 10 min at 1,000g at 4°C. The supernatant was centrifuged again for 40 min at 48,000g at 4°C to extract membrane protein. The pellet was resuspended (50 mM Tris [pH 7.4], 150 mM NaCl, 1 mM EDTA, and protease inhibitors), and 20 to 30 μg of protein was processed, electrophoresed, and transferred according to the supplier’s instructions (Invitrogen, Carlsbad, CA). Blots were probed with anti-dRyr (rabbit polyclonal, a generous gift from Daniel Cordova, M.Sc., Senior Biochemist, Dupont Crop Protection) at a 1:1,000 dilution, and anti–Na,K-ATPase (mouse mAbα5; Developmental Studies Hybridoma Bank, Iowa City, IA) at 1.3 ng/ml. Secondary antibodies were peroxidase-linked and used according to the supplier’s instructions. Blots were developed with the enhanced chemiluminescence detection system (GE Healthcare, Piscataway, NJ). For quantification, four independent homogenates were prepared for each genotype, and two aliquots of each were analyzed, yielding eight blots for each genotype. Intensities of dRyr bands were measured and normalized to those of Na,K-ATPase using ImageJ (National Institutes of Health, Bethesda, MD).
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Reactive Climbing Assay
The reactive climbing assay, also known as the distribution test, was performed as described previously.18 Male flies aged 3–7 days were collected and sorted under carbon dioxide anesthesia and allowed to recover for 1 day. Without further anesthesia, flies were loaded into testing vials (10 per vial), consisting of 50-ml tubes perforated to allow gas exchange. Testing vials were equilibrated in a chamber with a constant flow of a fixed concentration of volatile anesthetics—halothane (Sigma, St. Louis, MO), sevoflurane and enflurane (RxElite, Boise, ID), and isoflurane (Baxter Healthcare, Deerfield, IL)—for 35–45 min. After equilibration, the flies were tapped down quickly to the bottom of the vial several times at 30-s intervals. After the last round of tapping, the flies were allowed to climb for 1 min, after which the proportion remaining at the conical bottom of the testing vial (fraction down) was recorded. Flies were tested with three or four more concentrations of anesthetic, progressively increased by 0.05% with each successive test, and equilibrated for 35–45 min before each test. Naive flies of the same genotype were then tested at the four or five higher concentrations to produce the full concentration–response curve of 8–10 concentrations, covering the full range from 0% down to 100% down. The process was repeated twice with naive flies. In total, this yielded a sample size of nine for each combination of drug and genotype.
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Countercurrent Locomotion Assay
The countercurrent assay was performed as described previously.19 Typically, a group of 30 male flies of each genotype, aged 3–7 days, were loaded into the start tube, tapped to the bottom, and given 10 s to climb up into a transfer tube at top. The flies in each transfer tube were then shifted into the next base tube by banging the device. This was repeated four more times at 10-s intervals, and at the end, the flies in each base tube were counted. A transfer probability, Pt, is calculated by the following formula: Pt = [n × (no. of flies in the base tube n)]/5 × (no. of flies in all tubes). The experiment was performed three to five times for each genotype.
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Sf9 Cell Culture and Flow Cytometry
Sf9 cells stably transfected with dRyr20 were a generous gift from Daniel Cordova, M.Sc. Wild-type Sf9 cells were purchased from Invitrogen. Cultures were maintained at 28°C in SF900-II serum-free medium (Invitrogen), without antibiotics, and subcultured at confluence by sloughing, approximately twice weekly.
To prepare Sf9 cells for Ca2+ imaging, they were suspended, centrifuged, washed in Ca2+ imaging saline (130 mM NaCl, 5.4 mM KCl, 1.2 mM MgCl2, 1 mM CaCl2, 4.2 mM NaHCO3, 7.3 mM NaH2PO4, 10 mM glucose, and 63 mM sucrose),20 and then centrifuged and resuspended in 5 ml of Ca2+ imaging saline containing 1 µM of the acetoxymethyl ester of a Fluo Calcium indicator (Fluo-5N AM in most experiments) plus Pluronic F-127 (Invitrogen). Cells were incubated in Fluo-acetoxymethyl for 30–45 min, and then washed in Ca2+ imaging saline for a minimum of 30 min before cytometry. Aliquots containing approximately 0.5 × 106 cells were placed in styrene tubes and treated with anesthetics and/or thapsigargin (Tocris Bioscience, Minneapolis, MN), and fluorescence of the Fluo indicator was measured on a BD FACScan (Becton-Dickinson, Franklin Lakes, NJ). Debris and dead cells were gated out by size and fluorescence caused by propidium iodide uptake, respectively. Intracellular Ca2+ concentration was calibrated at the end of the experiment by sequential addition of ionomycin (Tocris) and EGTA, calculated as described by Tsien et al.,21 using Kd = 2.3 µM for Fluo-5F and 90 µM for Fluo-5N. Offline analysis of cytometry data was carried out using Flow-Jo 7.6.1 (Tree Star, Inc., Ashland, OR). Anesthetic concentrations in solution were converted to partial atmospheric pressures using Ostwald water/gas partition coefficients.
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Ca2+ Imaging in Larval Central Nervous System
Changes in internal Ca2+ were visualized in the cell body of motoneuron RP2 (MNISN1s) using the genetically encoded Ca2+ indicator GCaMP322 driven by ShakB-GAL4. The central nervous system was dissected from the larva, and brain lobes were removed and mounted dorsal side down on a polyornithine-coated 25-mm coverslip, which formed the bottom of the recording chamber (Warner Instruments, Hamden, CT), mounted in a custom-fabricated plate that fit into the stage of a Nikon C-1 inverted confocal microscope (Nikon Instruments, Tokyo, Japan). This setup allowed for superfusion from above, and the preparation was imaged using a 60× oil immersion objective from below. The preparation was scanned at 1 Hz, with the laser at minimum power and the pinhole opened to 100 µm.
Recording solution (70 mM NaCl, 5 mM KCl, 0.3 mM CaCl2, 4 mM MgCl2, 10 mM NaHCO3, 5 mM trehalose, 115 mM sucrose, 5 mM HEPES, pH 7.2)23 was perfused at 2 ml/min using a peristaltic pump, with solutions selected via remote-controlled solenoid valve. Anesthetics were vortexed into solution for 1–2 min and perfused from sealed reservoirs. Concentrations of anesthetic in contact with cells were determined via head-space analysis of 50-µl samples of solutions collected from the chamber, using an Agilent 6850 gas chromatograph.24,25 Ca2+ imaging data were analyzed using Nikon EZ-C1 software. Because of their superficial positions, RP2s in multiple segments could be visualized at once, allowing analysis of four to eight cells per preparation. The entire cell body of each cell in the focal plane of a given preparation was defined as a region of interest, and the average pixel intensity versus time was determined. Because the background signal in regions outside of RP2 somata was essentially nil, background subtraction was unnecessary. δF/F was calculated by dividing the intensity at a given time by the average of samples 1–10, subtracting 1 to set the initial value to 0, and multiplying by 100 to generate a percentage change.
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Whole-cell recordings were performed largely as described previously.24 The external solution was identical (118 mM NaCl, 2 mM NaOH, 2 mM KCl, 4 mM MgCl2, 0.5 mM CaCl2, 60 mM sucrose, 5 mM trehalose, 5 mM HEPES, pH 7.1, 300–305 mOs/kg). The internal solution differed by the addition of Mg-ATP and GTP-Tris (152 mM potassium gluconate, 2 mM NaCl, 0.1 mM CaCl2, 10 mM HEPES, 1 mM EGTA, 4 mM Mg-ATP, 0.6 mM GTP-Tris, 8.4 mM KOH, pH 7.2, 290–295 mOs/kg). A larval central nervous system, expressing UAS-mcd8-GFP in RP2 under the control of ShakB-GAL4, was mounted on a polyornithine-coated chip of coverslip. The region of the sheath around RP2 somata in segment A5 was softened with collagenase type XIV (Sigma) and removed manually with a suction pipette. Gigaseals were formed under the guidance of Nomarski optics, and membrane voltage was recorded using an AxoPatch 200B (Axon Instruments, Union City, CA) in current clamp mode. Recording parameters and solutions were controlled, and data were recorded using pClamp 8 (Axon). Drugs were dissolved as described above under Ca2+ Imaging in Larval Central Nervous System and perfused at 2 ml/min.
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Statistical Analysis
Reactive climbing data were analyzed largely as described previously.18 In brief, the data were entered into SPSS 13 (SPSS Inc., Chicago, IL) and fitted with the concentration–response relationship: fraction down = Cn/(Cn + EC50n), using logit analysis,26,27 where C is the anesthetic concentration, n reflects the steepness of the curve and is shared between genotypes for each experiment, and EC50 is the anesthetic concentration at the midpoint of the curve. Each vial of flies was used for four to five anesthetic concentrations, so that the full range of eight concentrations required a pair of vials. The EC50 was determined for each pair of vials, for which the percentage shift in EC50 was derived using the following formula:
Equation (Uncited)
Equation (Uncited)
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Flow cytometry and GCaMP3 fluorescence data were analyzed statistically in SigmaPlot 11.0 (Systat Software, Inc., San Jose, CA). For fitting of concentration–response curves for Fluo-5 fluorescence, data were normalized by defining percentage of untreated cells with elevated [Ca2+]i as 0 and those treated with saturating concentrations of anesthetic as 1, and the resulting relationships were fitted with a three-parameter Hill equation. Concentration–response curves for GCaMP3 fluorescence were plotted and analyzed as for Fluo-5, but using δF/F values. A few preparations that responded with exaggerated Ca2+ flux to halothane concentrations above 2.0 mM were excluded from the concentration–response analysis by Dixon’s test for outliers.
All data fit the assumptions for parametric statistics. Differences between two groups were assessed using the Student t test. Comparisons involving a single factor were performed using one-way ANOVA. The analysis of multiple genotypes and anesthetics described below under dRyr Mutations Have Weaker Effects on Other Volatile Anesthetics was performed with two-way ANOVA. The Bonferroni–Dunn test was used for post hoc analysis. Because of the conservative nature of the Bonferroni correction, P values were equal to 1 in a small number of cases.
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Ryanodine Receptor Insertion Mutants Are Resistant to Halothane
Fig. 1
Fig. 1
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The Drosophila genome contains a single Ryr gene at cytologic position 44F.12,28 In our initial tests of the role of dRyr in the response to the volatile anesthetic halothane, we used dRyrk04913, a hypomorphic mutant allele resulting from the insertion of a transposon 399 bases upstream of the start codon of dRyr,12 and dRyrGS21220, nine bases upstream of dRyrk04913 (fig. 1A). dRyrGS21220 is homozygous viable, whereas, as described previously,12 dRyrk04913 is lethal, presumably because of a tightly linked mutation outside the dRyr locus (see above under Materials and Methods for details). Given the close proximity of the transposons in the two alleles, we expected them to produce similar phenotypes.
We examined the response of the mutants to halothane using a reactive climbing assay that evaluates the flies’ righting/climbing reflex following mechanical agitation in the presence of anesthetic.18 Under these conditions, wild-type Canton-S male flies exhibited a halothane-dependent decrease in locomotor activity, resulting in an increase in the proportion failing to climb (“Fraction Down”; fig. 1B, black curve), with flies lying immobile at the bottom of the vial at the highest anesthetic concentrations. For heterozygous dRyrGS21220/+ and dRyrk04913/+, and the transheterozygote dRyrk04913/dRyrGS21220, the concentration–response curves shifted to the right, indicating resistance to halothane relative to the wild-type. The effective concentrations (vol/vol) at which 50% of flies were down (EC50) were 0.22%, 0.22%, and 0.27% for dRyrGS21220/+, dRyrk04913/+, and dRyrk04913/dRyrGS21220, respectively, all of which were significantly higher than that of the wild-type control (0.17% vol/vol; fig. 1B; P < 0.0001 for all, two-way ANOVA and Bonferroni–Dunn post hoc tests). In separate experiments, the EC50 for dRyrGS21220 homozygotes (0.27%) was also significantly higher than for matched controls (0.20%; P < 0.0001, Student t test).
Reduction in halothane sensitivity was paralleled by reduction in dRyr protein levels (fig. 1C). There was a significant effect of genotype on protein levels (P = 0.0009, one-way ANOVA), and the allelic combination that reduced halothane sensitivity the most, dRyrk04913/dRyrGS21220, reduced dRyr protein levels significantly. dRyr protein levels in the heterozygotes were intermediate between the wild-type and dRyrk04913/dRyrGS21220, but Bonferroni-Dunn post hoc tests failed to demonstrate significant differences from control. We conclude that the insertion alleles reduce dRyr expression, but that anesthetic sensitivity discriminates between alleles more effectively than do immunoblots.
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dRyr Mutations Have Weaker Effects on Other Volatile Anesthetics
Previous work identified target genes for anesthetic action that have a distinct preference for halothane over other anesthetic agents, implying the presence of agent-specific pathways.2,8 To determine the level of anesthetic specificity of dRyr, we assayed the responses of dRyr mutants to sevoflurane, enflurane, and isoflurane, and compared them to the response to halothane using a two-way ANOVA (generalized linear model; fig. 1D). There was a significant effect of anesthetic agent on the magnitude of shifts in EC50, with halothane having the strongest effect (P < 0.0001, two-way ANOVA). In Bonferroni–Dunn post hoc tests, dRyrk04913/dRyrGS21220 mutants were resistant to sevoflurane, enflurane, and isoflurane compared with wild-type controls. dRyrk04913/+ produced small shifts in the EC50 values for sevoflurane, enflurane, and isoflurane that were not significantly different from wild-type controls, and were significantly smaller than the effect of this allele on halothane. dRyrGS21220/+ had mixed effects. It showed significant resistance to enflurane, which was not significantly different from that for halothane, but did not significantly affect the response to sevoflurane or isoflurane. dRyr mutants were therefore more resistant to the anesthetic effects of halothane than to those of other volatile agents, implicating dRyr as a component of the pathway(s) that regulates halothane sensitivity.
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Halothane Sensitivity Correlates with dRyr Copy Number
Fig. 2
Fig. 2
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To explore the relationship between dRyr gene copy number, dRyr protein levels, and halothane sensitivity more systematically, we generated flies with the dRyr gene deleted or duplicated. The deletion dRyrδ25 was created by excising the region between two transposable elements using flippase–flippase recombination target-mediated recombination (fig. 1A). The resulting lethal allele removes all but the first exon of dRyr and all of CG8272, a gene of unknown function. This allele produces significant resistance to halothane, both in a heterozygous state and in combination with the hypomorphic allele RyrGS21220 (fig. 2A; P < 0.0001 and P < 0.0001, one-way ANOVA and Bonferroni–Dunn post hoc tests). In dRyrδ25/+ animals, which possess exactly one copy of dRyr, halothane EC50 is 36% higher than in wild-type controls. Transheterozygous dRyrδ25/dRyrGS21220 mutants shift halothane EC50 by 43% (fig. 2A), which is significantly higher than the wild-type but not different from dRyrδ25/+. As might be expected for a hypomorphic mutation, dRyrGS21220/+ had intermediate effects, significantly higher than the wild-type and lower than dRyrδ25/+ or dRyrδ25/dRyrGS21220.
To increase dRyr gene copy number, we inserted an additional genomic copy of dRyr into chromosome 3 to generate the dRyr duplication strain, dRyr24D03 (fig. 1A). dRyr24D03 rescued the halothane-resistant phenotype of dRyrGS21220/+ (fig. 2B), with the shift in EC50 of dRyrGS21220/+; dRyr24D03/+ significantly reduced from dRyrGS21220/+ (P < 0.0001, one-way ANOVA and Bonferroni–Dunn post hoc tests), and not different from the wild-type. Combining dRyr24D03 into a wild-type background, resulting in three copies of dRyr, caused significant hypersensitivity to halothane (i.e., a shift in EC50 of −19%; fig. 2B). Additional copies of dRyr24D03 produced lethality, suggesting that no more than three copies of the gene can be tolerated. Importantly, the mutants’ altered response to halothane did not result from nonspecific hyperactivity or arousal, in that all performed normally or worse than wild-type animals when tested for locomotion and responsiveness to mechanical stimulation in the absence of anesthetic (fig. 2C).
As was the case for the dRyr insertion mutants (fig. 1C), there was a significant effect of dRyr genotype on dRyr protein levels (fig. 2D; P < 0.0001, one-way ANOVA). In Bonferroni–Dunn post hoc tests, dRyr protein was strongly and significantly reduced in dRyrδ25/+ and dRyrδ25/ dRyrGS21220. The increase in dRyr protein in dRyr24D03/+ did not reach significance.
Taken together, our results demonstrate that halothane sensitivity follows dRyr gene copy number and protein levels. We conclude that dRyr is a limiting factor for halothane-induced anesthesia and a likely target of halothane.
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dRyr Point Mutations Change Halothane Sensitivity
Fig. 3
Fig. 3
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Although copy number variation in humans is well-documented, most disease-associated mutations in human Ryr (hRyr) genes are amino acid substitutions that lead to dominant alleles.11,29 dRyr is 45–47% identical in primary sequence to the three mammalian Ryrs and contains conserved protein domains important to Ryr function (fig. 3A), indicating that Ryr activity and regulation may be conserved across species.
Table 1
Table 1
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Previous work had identified a collection of lethal EMS mutants in region 44D-45F.30,31 Using NextGen sequencing, we determined that five of these mutants [l(2)44Fa3, l(2)44Fg1, l(2)44Fh1, l(2)44Fj1, and l(2)44Fp1] altered the sequence of dRyr (table 1; fig. 3, A and B). Two of these dRyr alleles contained nonsense mutations at positions corresponding to amino acid 3878 (Q3878X) in l(2)44Fa3, and at 4452 (Y4452X) in l(2)44Fj1. In both alleles, mutations introduced a stop codon before the transmembrane region containing the ion channel, producing truncated and nonfunctional dRyrs. As expected for loss-of-function alleles, heterozygotes were resistant to halothane (dRyrY4452X/+, 78% shift, dRyrQ3878X/+, 48% shift compared to genetically matched control strain l(2)44Ff1; fig. 3C; P < 0.0001 and P < 0.0001, one-way ANOVA and Bonferroni–Dunn post hoc tests) and both failed to complement the lethal phenotype of dRyrδ25. The relative magnitude of halothane resistance in the point mutants appeared to be larger than that of the deletion allele dRyrδ25 (fig. 2A), but differences in genetic backgrounds of the animals in the two experiments make comparisons problematic.
The three other alleles were missense mutations, leading to substitutions of highly conserved amino acids: R4305C in l(2)44Fg1, E4340K in l(2)44Fh1, and P2773L in l(2)44Fp1 (table 1; fig. 3, A and B). All of the missense mutations were lethal in combination with dRyrδ25, two of the mutations are located in a region associated with dominant channelopathies in hRyr2 (see below under Ryr and Neuropathology for details), and all exhibited dominant hypersensitivity to halothane (fig. 3C). Halothane sensitivity was enhanced in dRyrR4305C/+ (−28% shift in EC50, P = 0.0043, one-way ANOVA and Bonferroni–Dunn post hoc tests), and strongly increased in dRyrE4340K/+ and dRyrP2773L/+ (−72% and −80%, respectively; P < 0.0001, one-way ANOVA and Bonferroni–Dunn post hoc tests; fig. 3C). Thus, the missense mutations in dRyr produced dominant effects on halothane anesthesia in Drosophila, resembling the gain-of-function phenotype caused by an extra copy of dRyr (fig. 2B).
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dRyr Function in the Nervous System Is Required for Normal Halothane Sensitivity
Fig. 4
Fig. 4
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In the human disease malignant hyperthermia, mutations in hRyr1 are associated with a potentially fatal condition in which skeletal muscle Ryrs activate in response to volatile anesthetics, particularly halothane. Because the reactive climbing assay is based on locomotion, and could therefore be influenced by dRyr action in muscle, we determined the site of action of dRyr by driving the expression of double-stranded RNA (UAS-dRyrRNAi) using tissue-specific GAL4 drivers. Importantly, driving UAS-dRyrRNAi with the muscle-specific driver MHC-GAL4 failed to alter halothane sensitivity compared to controls carrying MHC-GAL4 alone (fig. 4A; P = 1, two-way ANOVA and Bonferroni–Dunn post hoc tests). By contrast, flies expressing UAS-dRyrRNAi under the control of the pan-neuronal drivers elav-GAL4 and Appl-GAL4 were significantly resistant to halothane (36% and 25% shift in EC50, respectively; fig. 4A; P < 0.0001 and P = 0.0094), suggesting that dRyr function is required in neurons for normal halothane sensitivity. Interestingly, driving UAS-dRyrRNAi expression in glia with repo-GAL4 also produced significant resistance to halothane (22% shift in EC50; P = 0.0373), indicating that dRyr is required in glia and neurons for halothane anesthesia. When driven by nrv2-GAL4, which expresses broadly but not ubiquitously in neurons and glia, UAS-dRyrRNAi produced a small (16%) but not statistically significant shift in EC50 (fig. 4A; P = 0.0862).
To further establish the site of action of dRyr, we sought to restore halothane sensitivity in dRyr mutants by expressing a UAS-dRyr transgene in selected cell types. Although expression of the rescue construct using the exclusively muscle-, neuron-, and glia-specific drivers proved lethal, expression in neurons and glia using nrv2-GAL4 restored sensitivity to halothane in transheterozygous dRyrk04913/dRyrGS21220 mutants (−22% shift in EC50; fig. 4B; P < 0.0001, Student t test). This manipulation also restored dRyr protein expression in the brains of rescued flies, as shown by Western blots of head extracts (fig. 4C; P < 0.0001, Student t test).
In summary, tissue-specific knockdown and rescue experiments support the conclusion that dRyr functions in the nervous system and not muscle to control anesthetic responsiveness of Drosophila. In addition, the lethality resulting from UAS-dRyr overexpression in most tissues, combined with the observation that homozygous dRyr24D03 and dRyrδ25 are lethal, indicates that temporal or spatial variation in dRyr expression levels are poorly tolerated, presumably because of its critical role in cellular calcium homeostasis.
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Halothane Induces Ca2+ Release in dRyr-transfected Sf9 Cells
Fig. 5
Fig. 5
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The results described in the previous section demonstrate that dRyr in the nervous system is required for full susceptibility to halothane, but the ability of halothane to activate the Drosophila Ryr has not been tested. We therefore assayed Ca2+ flux in Sf9 cells stably transfected with dRyr (Sf9+dRyr),20 using the Ca2+-sensitive dye Fluo-5 and flow cytometry (fig. 5). Without halothane, the intensity of Fluo-5 fluorescence in Sf9+dRyr cells (fig. 5, A4) corresponded to [Ca2+]i between 50 and 100 nM. Treatment with halothane concentrations greater than 1 mM shifted fluorescence to a much higher level in almost all transfected cells (fig. 5, A1), with calibrated [Ca]2+i exceeding 100 µM. Pretreatment of transfected cells with 300 nM thapsigargin to deplete intracellular calcium stores blocked the halothane-induced [Ca2+]i increase (fig. 5B), confirming that the response to halothane depends on internal Ca2+ stores. Halothane had no effect on untransfected Sf9 cells (fig. 5C), even at high concentrations (fig. 5D, open diamonds), indicating that the response to halothane required dRyr.
Curiously, individual Sf9+dRyr cells responded to halothane in an all-or-none manner, exhibiting either basal or maximal levels of [Ca2+]i at any given halothane concentration (fig. 5, A2 and A3). Concentration–response curves were thus analyzed as the proportion of cells producing elevated [Ca2+]i in response to drug application. Analyzed in this way, the EC50 for caffeine (1.42 ± 0.96 mM), for which the response is completely dependent on dRyr expression, was similar to the previously published value.20 For anesthetics, concentration–response relationships showed strong selectivity for halothane (fig. 5D), with EC50 values of 0.26, 1.1, and 1.3 mM for halothane, sevoflurane, and isoflurane, respectively. Converted to partial pressures, these values correspond to 0.48%, 1.9%, and 2.6% atm, indicating that halothane is manyfold more potent than either sevoflurane or isoflurane. We propose that molecular interactions between halothane and the dRyr protein explain the selective sensitivity of dRyr mutants to halothane anesthesia in Drosophila (fig. 1D).
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Halothane Increases [Ca2+]i and Hyperpolarizes Motoneuron RP2
Fig. 6
Fig. 6
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To examine the effects of halothane on Ca2+ flux in central neurons of Drosophila, we drove the expression of the genetically encoded Ca2+ indicator GCaMP322 in larval motoneuron RP2 using ShakB-GAL4. RP2 responded to halothane with robust increases in GCaMP3 fluorescence, clearly visible in unprocessed images (fig. 6A).
All cells produced an immediate response to halothane application that peaked during the short period of halothane entering the chamber (fig. 6B). The amplitude of this immediate response showed a robust concentration–response relationship (fig. 6C), with an EC50 for halothane of 0.61 mM. A pronounced undershoot, during which [Ca2+]i fell below resting levels, was routinely observed, particularly in response to high concentrations of halothane (fig. 6, B2). A minority of cells produced an additional delayed response, consisting of a plateau overlaid with spiky transients, at or near the time of halothane removal (fig. 6, B3).
The response of RP2 to halothane was altered by mutations in dRyr (fig. 6D). In preparations heterozygous for the deletion mutant, dRyrδ25, and the presumed truncation, dRyrQ3878X, the GCaMP3 responses to 0.5 mM halothane were reduced by more than 30% and more than 50%, respectively. Although consistent with a reduced response, these differences did not reach statistical significance (P = 1 and P = 0.6008, one-way ANOVA and Bonferroni–Dunn post hoc tests). However, RP2 motoneurons heterozygous for point mutant dRyrE4340K, which causes dominant hypersensitivity in the reactive climbing assay (fig. 3C), responded to halothane with significantly larger Ca2+ flux (fig. 6D; P = 0.0484), indicating that this allele enhances the response of dRyr to halothane. The hypersensitivity of dRyrE4340K for both reactive climbing and Ca2+ release suggests that it is a gain-of-function allele for halothane action.
The response of RP2 motoneurons to caffeine provided additional insight into the nature of the mutations. Wild-type RP2 neurons’ responses to caffeine were similar to those of other insect neurons,20 with an EC50 of 5.4 ± 2.4 mM. When challenged with 5 mM caffeine, the change in fluorescence in dRyrQ3878X/+ was significantly lower than in the wild-type (fig. 6E; P = 0.0375, one-way ANOVA and Bonferroni–Dunn post hoc tests). Surprisingly, the caffeine response of neurons heterozygous for dRyrE4340K was also smaller than normal (fig. 6E; P = 0.0453), to an extent similar to that of dRyrQ3878X. Therefore, the E-to-K substitution enhances the response to halothane but reduces the response to caffeine, indicating that the mutation selectively alters the response of dRyr depending on the nature of the signal. This conditional loss-of-function phenotype may also explain the lethality of dRyrE4340K/dRyrδ25, in that the substitution may disable dRyr for a vital function.
In whole-cell, current clamp recordings of RP2, halothane evoked a strong, concentration-dependent hyperpolarization (fig. 6F). Importantly, there was no evidence of depolarization or spiking that would indicate Ca2+ flux through channels in the plasma membrane. Instead, the membrane hyperpolarized 10–15 mV, depending on the halothane concentration. Onset and recovery both required approximately 2 min, although recovery was delayed at the higher concentration. Therefore, the Ca2+ flux demonstrated by imaging with GCaMP3 is paralleled by a robust hyperpolarization of the motoneuron, suggesting the activation of an inhibitory conductance.
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The molecular and cellular mechanisms of volatile anesthetic action have been the subject of intensive study for decades. This interest derives not only from the great medical importance of anesthesia but also from the conviction that understanding the molecular and neural pathways that mediate anesthetic effects will provide insight into the nature of arousal and consciousness. Additional interest has been generated recently by the finding that general anesthesia contributes to various neuropathologic conditions. Despite these convergent interests, the search for biologically relevant protein targets of volatile anesthetics has produced few confirmed candidates. The present study demonstrates that in Drosophila the ryanodine receptor (dRyr) is likely to represent such an anesthetic target. We show that neurally expressed dRyr mediates the behavioral response to the volatile anesthetic halothane, with whole-animal sensitivity to halothane anesthesia strongly dependent on gene dosage of dRyr. Point mutations in the dRyr gene that generate truncated, and therefore nonfunctional, proteins cause resistance to halothane, whereas missense mutations that alter highly conserved amino acids make flies more sensitive to the anesthetic. Our demonstration that the dRyr imparts halothane sensitivity to Sf9 cells, and that neurons in central nervous systems isolated from halothane-sensitive dRyr mutants exhibit elevated Ca2+ influx specifically in response to halothane, further support the conclusion that dRyr is a bona fide target of the anesthetic.
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dRyr in the Nervous System Mediates Halothane Sensitivity
The data presented here show that the potency of halothane is proportional to dRyr function. Reduction of function, as described for heterozygous dRyrK04913, dRyrGS21220, dRyrδ25, or their heteroallelic combinations, causes resistance to halothane. Moreover, point mutants that are predicted to produce truncated, and therefore nonfunctional dRyr channels, also produce dominant resistance. Conversely, an additional genomic copy of dRyr confers hypersensitivity to halothane.
Importantly, tissue-specific knockdown and rescue experiments demonstrated that dRyr function in neurons and glia, but not muscle, is necessary for normal susceptibility to halothane anesthesia and that dRyr expression in these cells is sufficient to impart anesthetic sensitivity. The Ryr-dependent anesthetic phenotypes are therefore clearly distinct from those found in malignant hyperthermia, a condition associated with mutations that cause halothane hypersensitivity of the skeletal muscle isoform, Ryr1, in humans and swine. This conclusion is underscored by the finding that RNAi-mediated knockdown of dRyr expression in muscle fails to alter the halothane sensitivity of reactive climbing.
The observation that both the Drosophila Ryr and the mammalian muscle isoform of this receptor, Ryr1, mediate halothane-sensitive physiologic processes suggests that halothane sensitivity is a general property of Ryrs. Indeed, in cardiac myocytes and neurons, which express predominantly Ryr2 and Ryr3, respectively, halothane induces Ryr-dependent Ca2+ flux from intracellular stores.32–34 This is consistent with our observations that dRyr mediates halothane-dependent Ca2+ flux in Drosophila motoneurons, and supports a proposal that neural Ryrs mediate the immobilizing effects of anesthetics.
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Mechanisms of Ryr Activation
Although regulation of Ryr function is complex, two principal mechanisms have been defined for its activation: Ca2+-mediated activation and interaction with accessory proteins.35 If halothane activates Ryr by elevating intracellular Ca2+ levels, it must do so either by promoting Ca2+ entry from the extracellular space or by causing release from intracellular stores. Flow cytometry results reported here show that elevation of internal Ca2+ is not observed in Sf9 cells, even at high levels of halothane, unless they have been transfected with dRyr. Moreover, it has been shown previously that halothane can activate isolated Ryr1 channels in membrane preparations.36 This argues against activation of dRyr by Ca2+-induced Ca2+ release.
Although it remains possible that halothane activates Ryr indirectly by interacting with one of its many accessory proteins, this explanation would require that Sf9 cells, which do not normally express detectable Ryr,20 produce the critical, halothane-dependent accessory subunit(s) when transfected with dRyr. We instead favor the hypothesis that halothane interacts directly with the dRyr protein. The absence of the full complement of regulatory subunits in Sf9+dRyr cells may in fact explain the curious bimodal pattern of the response of Sf9+dRyr cells to halothane. If Sf9 cells do not express a protein such as sorcin, which has been postulated to help terminate Ca2+-induced Ca2+ release,35 activation of dRyr by halothane could initiate a positive feedback loop of Ca2+ release, in which a small increase of cytoplasmic Ca2+ would initiate a flood of Ca2+ in the cytoplasm.
In contrast to the all-or-none response of Sf9+dRyr cells, RP2 motoneurons responded to halothane with transient, concentration-dependent Ca2+ flux. The transient nature of the signal suggests that either dRyrs inactivate in the continued presence of halothane, or that Ca2+ is rapidly removed from the cytoplasm. The transient reduction of GCaMP3 fluorescence below baseline levels on removal of halothane is consistent with robust activation of mechanisms for extrusion and sequestration of intracellular Ca2+.
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Downstream Effectors
Ca2+ released by halothane-dependent activation of dRyr must ultimately change neuronal excitability to contribute to the anesthetic state. Observations in mammals and worms show that neuronal hyperpolarization is a common feature of halothane anesthesia,4,5 consistent with our electrophysiologic recordings of RP2 motoneurons. The ultimate effector of Ryr activation is thus likely to be an inhibitory conductance, possibly carried by a K+ channel. Ca2+-activated K+ channels are obvious candidates, but it is interesting to speculate that the rapid and robust removal of excess Ca2+ from the cytoplasm may provide a novel route to K+ channel activation. Ca2+ extrusion by the plasma membrane Ca2+ adenosine triphosphatase, which exchanges internal Ca2+ for external H+, can result in cytoplasmic acidification,37 which could in turn activate K2P channels, such as TREK-1, which are sensitive to low internal pH.38 This mechanism could also provide an additional pathway for the actions of carbon dioxide, which is membrane permeant, causes acidification through the action of carbonic anhydrase, and acts as an anesthetic.39 The merits of these and competing hypotheses remain to be tested, but the work described here shows that the genetic and physiologic tools readily available in Drosophila should be useful in doing so.
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Ryr and Neuropathology
The intimate relationship between dRyr activity and halothane anesthesia has possible implications regarding the association between volatile anesthetics and cytotoxicity, neurodegeneration, and cognitive deficits, pathologic conditions often associated with dysregulation of intracellular Ca2+ levels.40 It is also interesting in this regard that two of the three point mutations in dRyr that we found to enhance sensitivity to halothane are in a region associated with human cellular abnormality.
dRyrE4340K is adjacent to a mutation in hRyr2 (i.e., N4178S) that is associated with multiple cases of catecholaminergic polymorphic ventricular tachycardia, a condition characterized by exercise-induced cardiac arrhythmia.29 Perhaps most interestingly, dRyrR4305C is a substitution identical to that causing sudden cardiac death (hRyr2R4144C).41 All three missense mutations increase the potency of halothane in our reactive climbing assay, and one enhances the halothane-evoked release of Ca2+ in RP2 motoneurons, suggesting shared mechanisms of Ryr-dependent Ca2+ dysregulation among different isoforms and across phyla. Of particular interest is the observation that the amino acid substitution in dRyrE4340K enhances sensitivity to halothane but blocks activation by caffeine, indicating that the mutant protein is not simply hyperactive. Similar mutations in neurally expressed Ryr in humans may predispose individuals to the cytotoxic effects of anesthetics. In support of this hypothesis, halothane produces altered electroencephalographic activity during episodes of malignant hyperthermia in susceptible swine.42
The emerging picture based on investigation of the mechanisms of anesthesia in genetic model organisms is that, despite the identical endpoint of immobility, each volatile anesthetic is likely to act through its own collection of molecular targets. The collection of targets mediating halothane anesthesia now includes Ryr. At present, it is unclear how completely the mechanisms uncovered in invertebrate models, such as flies and worms, will translate to mammalian anesthesia. In mammals, immobilization by volatile anesthetics is likely to be mediated by spinal circuitry.4 The insect ventral nerve cord and motoneurons such as RP2 are functional analogs of the mammalian spinal cord and spinal motoneurons, respectively. However, the degree of homology between the segmental neuromeres of insects and mammals is not fully understood, and therefore the level of conservation of molecular targets of anesthetics remains to be established.
Remaining open questions regard the identities of the cells required to mediate immobility and the precise cellular sites of action. Our knockdown and rescue experiments manipulated dRyr expression in large numbers of neurons and glia, and it will be necessary to drive dRyr expression in subsets of these cells to determine whether anesthesia depends on the activity of a few cells or a large population. Furthermore, although halothane elicits Ca2+ flux in neuronal cell bodies that correlates with hyperpolarization, we have established neither the causal link between Ca2+ flux and membrane potential, nor the most important subcellular location (e.g., axon, dendrite, synapse) of dRyr function. These questions provide fertile territory for subsequent investigations.
The authors thank David Ide, B.A. (Research Assistant, Research Services Branch, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland), and the Research Services Branch for construction of equipment; Daniel Cordova, M.Sc. (Senior Biochemist, Dupont Crop Protection, Newark, Delaware), for the anti-dRyr antibody, Sf9+dRyr cells, and Ryr-V5 plasmid; Ted Usdin, Ph.D., M.D. (Senior Investigator, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland), for the use of culture facilities; Robert L. Scott, M.Sc. (Research Assistant, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland), for technical assistance and comments on the article; Songling Huang, Ph.D. (Senior Data Analyst, TurningPoint Global Solutions, Rockville, Maryland) and Gang Chen, Ph.D. (Mathematical Statistician, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland), for statistical consultation; Qun Gu, A.A. (Research Assistant, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland), for plasmid construction; Benjamin White, Ph.D. (Senior Investigator, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland), for use of equipment and help in assembling the article; and Philip Morgan, M.D. (Professor, Seattle Children’s Hospital, Seattle, Washington), and Joseph Campbell, Ph.D. (Program Officer, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, Maryland), for helpful comments.
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