Studying Synaptic Connectivity and Strength with Optogenetics and Patch-Clamp Electrophysiology
Abstract
:1. General Introduction
2. Optogenetically Assessing Synaptic Connectivity
2.1. Characterization of Synaptic Connectivity via Ionotropic Receptors
- Is potassium or cesium the major cation?
- Is chloride or an alternative (e.g., gluconate or methanesulphonate) the major anion?
2.1.1. Considerations for AMPAR-Mediated Synaptic Connectivity
2.1.2. Considerations for NMDAR-Mediated Synaptic Connectivity
2.1.3. Considerations for GABAAR-Mediated Synaptic Connectivity
2.2. Characterization of Synaptic Connectivity via Metabotropic Receptors
2.3. Characterizing Monosynaptic vs. Polysynaptic Connectivity
2.4. Summary and Caveats
3. Using Optogenetics and Patch-Clamp to Assess the Function and Strength of Specific Synapses
3.1. AMPAR–NMDAR Ratios and I/V Relations of AMPARs
3.1.1. AMPAR–NMDAR Ratios
3.1.2. AMPAR I/V Relationships and the Rectification Index
3.1.3. Practical Examples of Optogenetic Studies Investigating AMPAR/NMDARs and I/V Relations
3.1.4. Summary and Caveats
3.2. Paired-Pulse Ratios (PPR)
3.2.1. How PPR Relates to Presynaptic Processes Including the Probability of Release
3.2.2. Practical Examples of Optogenetic Studies Investigating PPR Differences
3.2.3. Summary and Caveats
3.3. Variance-Mean Analyses such as the 1/CV2 Metric
3.3.1. How 1/CV2 Relates to Presynaptic Processes including the Probability of Release
3.3.2. Practical Examples of Optogenetic Studies Investigating 1/CV2 Differences
3.3.3. Summary and Caveats
3.4. Quantal Responses in an Input-Specific Manner Using Strontium in the Extracellular Medium
3.4.1. How Strontium-Mediated Asynchronous Release Reflects Synaptic Quantal Sizes
3.4.2. Practical Examples of Optogenetic Studies Using Strontium to Assess Quantal Size Changes
3.4.3. Summary and Caveats
3.5. Optogenetic-Assisted Study of Opioid G Protein-Coupled Receptor Control over Specific Synapses
3.5.1. Practical Examples of Optogenetic Studies of Opioid GPCR Control over Specific Synapses
3.5.2. Summary and Alternative Strategies
4. Dual Color Optogenetics for Synapse Interrogation
4.1. Using Low (Blue) Irradiance to Obtain Specificity with Dual Color Optogenetics
4.1.1. Using the Chrimson and Chronos Opsin Pair for Dual Color Optogenetics
4.1.2. Using the Chrimson and ChR2 Opsin Pair for Dual Color Optogenetics
4.2. Using Protracted Orange Light to Reduce Red-Shifted Opsin Sensitivity to Subsequent Blue Light
4.3. Summary and Caveats of Dual Color Optogenetics
5. Discussion
5.1. Considerations when Expressing Opsins
5.2. The Bystander Effect
5.3. Troubleshooting by Limiting Viral Loads and Light Intensities
5.4. Concluding Remarks
Author Contributions
Funding
Acknowledgments
Conflicts of Interest
References
- Boyden, E.S.; Zhang, F.; Bamberg, E.; Nagel, G.; Deisseroth, K. Millisecond-Timescale, Genetically Targeted Optical Control of Neural Activity. Nat. Neurosci. 2005, 8, 1263–1268. [Google Scholar] [CrossRef] [PubMed]
- Hegemann, P.; Nagel, G. From Channelrhodopsins to Optogenetics. EMBO Mol. Med. 2013, 5, 173–176. [Google Scholar] [CrossRef] [PubMed]
- Petreanu, L.; Huber, D.; Sobczyk, A.; Svoboda, K. Channelrhodopsin-2-Assisted Circuit Mapping of Long-Range Callosal Projections. Nat. Neurosci. 2007, 10, 663–668. [Google Scholar] [CrossRef] [PubMed]
- Deubner, J.; Coulon, P.; Diester, I. Optogenetic Approaches to Study the Mammalian Brain. Curr. Opin. Struct. Biol. 2019, 57, 157–163. [Google Scholar] [CrossRef]
- Haery, L.; Deverman, B.E.; Matho, K.S.; Cetin, A.; Woodard, K.; Cepko, C.; Guerin, K.I.; Rego, M.A.; Ersing, I.; Bachle, S.M.; et al. Adeno-Associated Virus Technologies and Methods for Targeted Neuronal Manipulation. Front. Neuroanat. 2019, 13, 93. [Google Scholar] [CrossRef] [Green Version]
- Kim, C.K.; Adhikari, A.; Deisseroth, K. Integration of Optogenetics with Complementary Methodologies in Systems Neuroscience. Nat. Rev. Neurosci. 2017, 18, 222–235. [Google Scholar] [CrossRef]
- Lin, J.Y. A User’s Guide to Channelrhodopsin Variants: Features, Limitations and Future Developments. Exp. Physiol. 2011, 96, 19–25. [Google Scholar] [CrossRef]
- Manz, K.M.; Siemann, J.K.; McMahon, D.G.; Grueter, B.A. Patch-Clamp and Multi-Electrode Array Electrophysiological Analysis in Acute Mouse Brain Slices. STAR Protoc. 2021, 2, 100442. [Google Scholar] [CrossRef]
- Williams, S.R.; Mitchell, S.J. Direct Measurement of Somatic Voltage Clamp Errors in Central Neurons. Nat. Neurosci. 2008, 11, 790–798. [Google Scholar] [CrossRef]
- Beaulieu-Laroche, L.; Harnett, M.T. Dendritic Spines Prevent Synaptic Voltage Clamp. Neuron 2018, 97, 75–82.e3. [Google Scholar] [CrossRef] [Green Version]
- Neher, E. [6] Correction for Liquid Junction Potentials in Patch Clamp Experiments; Academic Press: Cambridge, MA, USA, 1992. [Google Scholar]
- Pascoli, V.; Terrier, J.; Espallergues, J.; Valjent, E.; O’Connor, E.C.; Lüscher, C. Contrasting Forms of Cocaine-Evoked Plasticity Control Components of Relapse. Nature 2014, 509, 459–464. [Google Scholar] [CrossRef] [PubMed]
- Christoffel, D.J.; Walsh, J.J.; Hoerbelt, P.; Heifets, B.D.; Llorach, P.; Lopez, R.C.; Ramakrishnan, C.; Deisseroth, K.; Malenka, R.C. Selective Filtering of Excitatory Inputs to Nucleus Accumbens by Dopamine and Serotonin. Proc. Natl. Acad. Sci. USA 2021, 118, e2106648118. [Google Scholar] [CrossRef] [PubMed]
- Nuno-Perez, A.; Trusel, M.; Lalive, A.L.; Congiu, M.; Gastaldo, D.; Tchenio, A.; Lecca, S.; Soiza-Reilly, M.; Bagni, C.; Mameli, M. Stress Undermines Reward-Guided Cognitive Performance through Synaptic Depression in the Lateral Habenula. Neuron 2021, 109, 947–956.e5. [Google Scholar] [CrossRef] [PubMed]
- Cerniauskas, I.; Winterer, J.; de Jong, J.W.; Lukacsovich, D.; Yang, H.; Khan, F.; Peck, J.R.; Obayashi, S.K.; Lilascharoen, V.; Lim, B.K.; et al. Chronic Stress Induces Activity, Synaptic, and Transcriptional Remodeling of the Lateral Habenula Associated with Deficits in Motivated Behaviors. Neuron 2019, 104, 899–915.e8. [Google Scholar] [CrossRef] [Green Version]
- Lee, B.R.; Ma, Y.Y.; Huang, Y.H.; Wang, X.; Otaka, M.; Ishikawa, M.; Neumann, P.A.; Graziane, N.M.; Brown, T.E.; Suska, A.; et al. Maturation of Silent Synapses in Amygdala-Accumbens Projection Contributes to Incubation of Cocaine Craving. Nat. Neurosci. 2013, 16, 1644–1651. [Google Scholar] [CrossRef] [Green Version]
- Wright, W.J.; Dong, Y. Silent Synapses in Cocaine-Associated Memory and Beyond. J. Neurosci. 2021, 41, 9275–9285. [Google Scholar] [CrossRef]
- Ito, W.; Erisir, A.; Morozov, A. Observation of Distressed Conspecific as a Model of Emotional Trauma Generates Silent Synapses in the Prefrontal-Amygdala Pathway and Enhances Fear Learning, but Ketamine Abolishes Those Effects. Neuropsychopharmacology 2015, 40, 2536–2545. [Google Scholar] [CrossRef] [Green Version]
- Traynelis, S.F.; Wollmuth, L.P.; McBain, C.J.; Menniti, F.S.; Vance, K.M.; Ogden, K.K.; Hansen, K.B.; Yuan, H.; Myers, S.J.; Dingledine, R. Glutamate Receptor Ion Channels: Structure, Regulation, and Function. Pharmacol. Rev. 2010, 62, 405–496. [Google Scholar] [CrossRef] [Green Version]
- Nuno-Perez, A.; Mondoloni, S.; Tchenio, A.; Lecca, S.; Mameli, M. Biophysical and Synaptic Properties of NMDA Receptors in the Lateral Habenula. Neuropharmacology 2021, 196, 108718. [Google Scholar] [CrossRef]
- Meye, F.J.; Soiza-Reilly, M.; Smit, T.; Diana, M.A.; Schwarz, M.K.; Mameli, M. Shifted Pallidal Co-Release of GABA and Glutamate in Habenula Drives Cocaine Withdrawal and Relapse. Nat. Neurosci. 2016, 19, 1019–1024. [Google Scholar] [CrossRef]
- Biedermann, J.; Braunbeck, S.; Plested, A.J.; Sun, H. Nonselective Cation Permeation in an AMPA-Type Glutamate Receptor. Proc. Natl. Acad. Sci. USA 2021, 118, e2012843118. [Google Scholar] [CrossRef] [PubMed]
- Voipio, J.; Kaila, K. GABAergic Excitation and K+-Mediated Volume Transmission in the Hippocampus. In Progress in Brain Research; Elsevier: Amsterdam, The Netherlands, 2000; Volume 125. [Google Scholar]
- Tan, D.; Nuno-Perez, A.; Mameli, M.; Meye, F.J. Cocaine Withdrawal Reduces GABABR Transmission at Entopeduncular Nucleus—Lateral Habenula Synapses. Eur. J. Neurosci. 2019, 50, 2124–2133. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Schöne, C.; Apergis-Schoute, J.; Sakurai, T.; Adamantidis, A.; Burdakov, D. Coreleased Orexin and Glutamate Evoke Nonredundant Spike Outputs and Computations in Histamine Neurons. Cell Rep. 2014, 7, 697–704. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Qiu, J.; Nestor, C.C.; Zhang, C.; Padilla, S.L.; Palmiter, R.D.; Kelly, M.J.; Rønnekleiv, O.K. High-Frequency Stimulation-Induced Peptide Release Synchronizes Arcuate Kisspeptin Neurons and Excites GnRH neurons. eLife 2016, 5, e16246. [Google Scholar] [CrossRef]
- Edwards, N.; Tejeda, H.A.; Pignatelli, M.; Zhang, S.; McDevitt, R.A.; Wu, J.; Bass, C.E.; Bettler, B.; Morales, M.; Bonci, A. Circuit Specificity in the Inhibitory Architecture of the VTA Regulates Cocaine-Induced Behavior. Nat. Neurosci. 2017, 20, 438–448. [Google Scholar] [CrossRef]
- Damonte, V.M.; Pomrenze, M.B.; Manning, C.E.; Casper, C.; Wolfden, A.L.; Malenka, R.C.; Kauer, J.A. Somatodendritic Release of Cholecystokinin Potentiates GABAergic Synapses onto Ventral Tegmental Area Dopamine Cells. Biol. Psychiatry 2022. [Google Scholar] [CrossRef]
- Qi, J.; Zhang, S.; Wang, H.L.; Wang, H.; de Jesus Aceves Buendia, J.; Hoffman, A.F.; Lupica, C.R.; Seal, R.P.; Morales, M. A Glutamatergic Reward Input from the Dorsal Raphe to Ventral Tegmental Area Dopamine Neurons. Nat. Commun. 2014, 5, 5390. [Google Scholar] [CrossRef] [Green Version]
- Geddes, S.D.; Assadzada, S.; Lemelin, D.; Sokolovski, A.; Bergeron, R.; Haj-Dahmane, S.; Béïque, J.C. Target-Specific Modulation of the Descending Prefrontal Cortex Inputs to the Dorsal Raphe Nucleus by Cannabinoids. Proc. Natl. Acad. Sci. USA 2016, 113, 5429–5434. [Google Scholar] [CrossRef] [Green Version]
- Cho, J.H.; Deisseroth, K.; Bolshakov, V.Y. Synaptic Encoding of Fear Extinction in MPFC-Amygdala Circuits. Neuron 2013, 80, 1491–1507. [Google Scholar] [CrossRef] [Green Version]
- Petreanu, L.; Mao, T.; Sternson, S.M.; Svoboda, K. The Subcellular Organization of Neocortical Excitatory Connections. Nature 2009, 457, 1142–1145. [Google Scholar] [CrossRef] [Green Version]
- Shu, Y.; Yu, Y.; Yang, J.; Mccormick, D.A. Selective Control of Cortical Axonal Spikes by a Slowly Inactivating K Current. Proc. Natl. Acad. Sci. USA 2007, 104, 11453–11458. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Kuniishi, H.; Yamada, D.; Wada, K.; Yamada, M.; Sekiguchi, M. Stress Induces Insertion of Calcium-Permeable AMPA Receptors in the OFC–BLA Synapse and Modulates Emotional Behaviours in Mice. Transl. Psychiatry 2020, 10, 154. [Google Scholar] [CrossRef] [PubMed]
- Scheefhals, N.; MacGillavry, H.D. Functional Organization of Postsynaptic Glutamate Receptors. Mol. Cell. Neurosci. 2018, 91, 82–94. [Google Scholar] [CrossRef] [PubMed]
- Zucker, R.S.; Regehr, W.G. Short-Term Synaptic Plasticity. Annu. Rev. Physiol. 2002, 64, 355–405. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Fioravante, D.; Regehr, W.G. Short-Term Forms of Presynaptic Plasticity. Curr. Opin. Neurobiol. 2011, 21, 269–274. [Google Scholar] [CrossRef] [Green Version]
- Lanore, F.; Silver, R.A. Extracting Quantal Properties of Transmission at Central Synapses. In Neuromethods; Humana Press Inc.: Totowa, NJ, USA, 2016; Volume 113, pp. 193–211. [Google Scholar]
- Malenka, R.C.; Bear, M.F. LTP and LTD: An Embarrassment of Riches. Neuron 2004, 44, 5–21. [Google Scholar] [CrossRef] [Green Version]
- Díaz-Alonso, J.; Nicoll, R.A. AMPA Receptor Trafficking and LTP: Carboxy-Termini, Amino-Termini and TARPs. Neuropharmacology 2021, 197, 108710. [Google Scholar] [CrossRef]
- Bredt, D.S.; Nicoll, R.A. Review AMPA Receptor Trafficking at Excitatory Synapses Characterization of Silent Synapses A Resolution Came with the Discovery of “Silent Syn-Apses”. Neuron 2003, 40, 361–379. [Google Scholar] [CrossRef] [Green Version]
- Suska, A.; Lee, B.R.; Huang, Y.H.; Dong, Y.; Schlüter, O.M. Selective Presynaptic Enhancement of the Prefrontal Cortex to Nucleus Accumbens Pathway by Cocaine. Proc. Natl. Acad. Sci. USA 2013, 110, 713–718. [Google Scholar] [CrossRef] [Green Version]
- Arroyo, S.; Bennett, C.; Hestrin, S. Nicotinic Modulation of Cortical Circuits. Front. Neural Circuits 2014, 8, 30. [Google Scholar] [CrossRef] [Green Version]
- Vieira, M.; Yong, X.L.H.; Roche, K.W.; Anggono, V. Regulation of NMDA Glutamate Receptor Functions by the GluN2 Subunits. J. Neurochem. 2020, 154, 121–143. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Isaac, J.T.R.; Ashby, M.; McBain, C.J. The Role of the GluR2 Subunit in AMPA Receptor Function and Synaptic Plasticity. Neuron 2007, 54, 859–871. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Lu, W.; Shi, Y.; Jackson, A.C.; Bjorgan, K.; During, M.J.; Sprengel, R.; Seeburg, P.H.; Nicoll, R.A. Subunit Composition of Synaptic AMPA Receptors Revealed by a Single-Cell Genetic Approach. Neuron 2009, 62, 254–268. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Wolf, M.E.; Tseng, K.Y.; Zukin, R.S.; Einstein, A. Calcium-Permeable AMPA Receptors in the VTA and Nucleus Accumbens after Cocaine Exposure: When, How, and Why? Front. Mol. Neurosci. 2012, 5, 72. [Google Scholar] [CrossRef] [Green Version]
- Lalanne, T.; Oyrer, J.; Farrant, M.; Sjöström, P.J. Synapse Type-Dependent Expression of Calcium-Permeable AMPA Receptors. Front. Synaptic Neurosci. 2018, 10, 34. [Google Scholar] [CrossRef] [Green Version]
- Mameli, M.; Bellone, C.; Brown, M.T.C.; Lüscher, C. Cocaine Inverts Rules for Synaptic Plasticity of Glutamate Transmission in the Ventral Tegmental Area. Nat. Neurosci. 2011, 14, 414–416. [Google Scholar] [CrossRef]
- Polli, F.S.; Kohlmeier, K.A. Prenatal Nicotine Exposure Alters Postsynaptic AMPA Receptors and Glutamate Neurotransmission within the Laterodorsal Tegmentum (LDT) of Juvenile Mice. Neuropharmacology 2018, 137, 71–85. [Google Scholar] [CrossRef]
- Salpietro, V.; Dixon, C.L.; Guo, H.; Bello, O.D.; Vandrovcova, J.; Efthymiou, S.; Maroofian, R.; Heimer, G.; Burglen, L.; Valence, S.; et al. AMPA Receptor GluA2 Subunit Defects Are a Cause of Neurodevelopmental Disorders. Nat. Commun. 2019, 10, 3094. [Google Scholar] [CrossRef] [Green Version]
- Asztely, F.; Erdemli, G.; Kullmann, D.M. Extrasynaptic Glutamate Spillover in the Hippocampus: Dependence on Temperature and the Role of Active Glutamate Uptake. Neuron 1997, 18, 281–293. [Google Scholar] [CrossRef] [Green Version]
- Jackman, S.L.; Regehr, W.G. The Mechanisms and Functions of Synaptic Facilitation. Neuron 2017, 94, 447–464. [Google Scholar] [CrossRef] [Green Version]
- Dobrunz, L.E.; Stevens, C.F. Heterogeneity of Release Probability, Facilitation, and Depletion at Central Synapses. Neuron 1997, 18, 995–1008. [Google Scholar] [CrossRef] [Green Version]
- Hanse, E.; Gustafsson, B. Paired-Pulse Plasticity at the Single Release Site Level: An Experimental and Computational Study. J. Neurosci. 2001, 21, 8362–8369. [Google Scholar] [CrossRef] [PubMed]
- Jackman, S.L.; Beneduce, B.M.; Drew, I.R.; Regehr, W.G. Achieving High-Frequency Optical Control of Synaptic Transmission. J. Neurosci. 2014, 34, 7704–7714. [Google Scholar] [CrossRef] [Green Version]
- Südhof, T.C. The Synaptic Vesicle Cycle. Annu. Rev. Neurosci. 2004, 27, 509–547. [Google Scholar] [CrossRef] [Green Version]
- Blitz, D.M.; Foster, K.A.; Regehr, W.G. Short-Term Synaptic Plasticity: A Comparison of Two Synapses. Nat. Rev. Neurosci. 2004, 5, 630–640. [Google Scholar] [CrossRef] [PubMed]
- Liu, W.Z.; Zhang, W.H.; Zheng, Z.H.; Zou, J.X.; Liu, X.X.; Huang, S.H.; You, W.J.; He, Y.; Zhang, J.Y.; Wang, X.D.; et al. Identification of a Prefrontal Cortex-to-Amygdala Pathway for Chronic Stress-Induced Anxiety. Nat. Commun. 2020, 11, 2221. [Google Scholar] [CrossRef] [PubMed]
- Creed, M.; Ntamati, N.R.; Chandra, R.; Lobo, M.K.; Lüscher, C. Convergence of Reinforcing and Anhedonic Cocaine Effects in the Ventral Pallidum. Neuron 2016, 92, 214–226. [Google Scholar] [CrossRef] [Green Version]
- Britt, J.P.; Benaliouad, F.; McDevitt, R.A.; Stuber, G.D.; Wise, R.A.; Bonci, A. Synaptic and Behavioral Profile of Multiple Glutamatergic Inputs to the Nucleus Accumbens. Neuron 2012, 76, 790–803. [Google Scholar] [CrossRef] [Green Version]
- Brock, J.A.; Thomazeau, A.; Watanabe, A.; Li, S.S.Y.; Sjöström, P.J. A Practical Guide to Using CV Analysis for Determining the Locus of Synaptic Plasticity. Front. Synaptic Neurosci. 2020, 12, 11. [Google Scholar] [CrossRef] [Green Version]
- Van Huijstee, A.N.; Kessels, H.W. Variance Analysis as a Tool to Predict the Mechanism Underlying Synaptic Plasticity. J. Neurosci. Methods 2020, 331, 108526. [Google Scholar] [CrossRef]
- Hogrefe, N.; Blom, S.M.; Valentinova, K.; Ntamati, N.R.; Jonker, L.J.E.; Nevian, N.E.; Nevian, T. Long-Lasting, Pathway-Specific Impairment of a Novel Form of Spike-Timing-Dependent Long-Term Depression by Neuropathic Pain in the Anterior Cingulate Cortex. J. Neurosci. 2022, 42, 2166–2179. [Google Scholar] [CrossRef] [PubMed]
- Asede, D.; Joseph, A.; Bolton, M.L.M. Deletion of NRXN1α Impairs Long-Range and Local Connectivity in Amygdala Fear Circuit. Transl. Psychiatry 2020, 10, 242. [Google Scholar] [CrossRef] [PubMed]
- Anastasiades, P.G.; Marques-Smith, A.; Butt, S.J.B. Studies of Cortical Connectivity Using Optical Circuit Mapping Methods. J. Physiol. 2018, 596, 145–162. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Sciamanna, G.; Ponterio, G.; Mandolesi, G.; Bonsi, P.; Pisani, A. Optogenetic Stimulation Reveals Distinct Modulatory Properties of Thalamostriatal vs Corticostriatal Glutamatergic Inputs to Fast-Spiking Interneurons. Sci. Rep. 2015, 5, 16742. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Marom, M.; Sebag, A.; Atlas, D. Cations Residing at the Selectivity Filter of the Voltage-Gated Ca2+-Channel Modify Fusion-Pore Kinetics. Channels 2007, 1, 377–386. [Google Scholar] [CrossRef] [Green Version]
- Xu-Friedman, M.A.; Regehr, W.G. Probing Fundamental Aspects of Synaptic Transmission with Strontium. J. Neurosci. 2000, 20, 4414–4422. [Google Scholar] [CrossRef] [Green Version]
- Shin, O.-H.; Rhee, J.-S.; Tang, J.; Sugita, S.; Rosenmund, C.; Sü Dhof, T.C. Sr 2 Binding to the Ca 2 Binding Site of the Synaptotagmin 1 C 2 B Domain Triggers Fast Exocytosis without Stimulating SNARE Interactions. Neuron 2003, 37, 99–108. [Google Scholar] [CrossRef] [Green Version]
- Babai, N.; Kochubey, O.; Keller, D.; Schneggenburger, R. An Alien Divalent Ion Reveals a Major Role for Ca2+ Buffering in Controlling Slow Transmitter Release. J. Neurosci. 2014, 34, 12622–12635. [Google Scholar] [CrossRef] [Green Version]
- MacAskill, A.F.; Cassel, J.M.; Carter, A.G. Cocaine Exposure Reorganizes Cell Type- and Input-Specific Connectivity in the Nucleus Accumbens. Nat. Neurosci. 2014, 17, 1198–1207. [Google Scholar] [CrossRef] [Green Version]
- Vautrin, J.; Barker, J.L. Presynaptic Quantal Plasticity: Katz’s Original Hypothesis Revisited. Synapse 2003, 47, 184–199. [Google Scholar] [CrossRef]
- Edwards, R.H. The Neurotransmitter Cycle and Quantal Size. Neuron 2007, 55, 835–858. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Meye, F.J.; Ramakers, G.M.J.; Adan, R.A.H. The Vital Role of Constitutive GPCR Activity in the Mesolimbic Dopamine System. Transl. Psychiatry 2014, 4, e361. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Fields, H.L.; Margolis, E.B. Understanding Opioid Reward. Trends Neurosci. 2015, 38, 217–225. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Winters, B.L.; Gregoriou, G.C.; Kissiwaa, S.A.; Wells, O.A.; Medagoda, D.I.; Hermes, S.M.; Burford, N.T.; Alt, A.; Aicher, S.A.; Bagley, E.E. Endogenous Opioids Regulate Moment-to-Moment Neuronal Communication and Excitability. Nat. Commun. 2017, 8, 14611. [Google Scholar] [CrossRef] [Green Version]
- Cusulin, J.I.W.; Füzesi, T.; Inoue, W.; Bains, J.S. Glucocorticoid Feedback Uncovers Retrograde Opioid Signaling at Hypothalamic Synapses. Nat. Neurosci. 2013, 16, 596–604. [Google Scholar] [CrossRef] [Green Version]
- Meye, F.J.; van Zessen, R.; Smidt, M.P.; Adan, R.A.H.; Ramakers, G.M.J. Morphine Withdrawal Enhances Constitutive μ-Opioid Receptor Activity in the Ventral Tegmental Area. J. Neurosci. 2012, 32, 16120–16128. [Google Scholar] [CrossRef] [Green Version]
- Tejeda, H.A.; Wu, J.; Kornspun, A.R.; Pignatelli, M.; Kashtelyan, V.; Krashes, M.J.; Lowell, B.B.; Carlezon, W.A.; Bonci, A. Pathway- and Cell-Specific Kappa-Opioid Receptor Modulation of Excitation-Inhibition Balance Differentially Gates D1 and D2 Accumbens Neuron Activity. Neuron 2017, 93, 147–163. [Google Scholar] [CrossRef] [Green Version]
- Siuda, E.R.; Copits, B.A.; Schmidt, M.J.; Baird, M.A.; Al-Hasani, R.; Planer, W.J.; Funderburk, S.C.; McCall, J.G.; Gereau, R.W.; Bruchas, M.R. Spatiotemporal Control of Opioid Signaling and Behavior. Neuron 2015, 86, 923–935. [Google Scholar] [CrossRef] [Green Version]
- Birdsong, W.T.; Jongbloets, B.C.; Engeln, K.A.; Wang, D.; Scherrer, G.; Mao, T. Synapse-Specific Opioid Modulation of Thalamo-Cortico-Striatal Circuits. eLife 2019, 8, e45146. [Google Scholar] [CrossRef]
- Matsui, A.; Jarvie, B.C.; Robinson, B.G.; Hentges, S.T.; Williams, J.T. Separate GABA Afferents to Dopamine Neurons Mediate Acute Action of Opioids, Development of Tolerance, and Expression of Withdrawal. Neuron 2014, 82, 1346–1356. [Google Scholar] [CrossRef] [Green Version]
- Muñoz, B.; Fritz, B.M.; Yin, F.; Atwood, B.K. Alcohol Exposure Disrupts Mu Opioid Receptor-Mediated Long-Term Depression at Insular Cortex Inputs to Dorsolateral Striatum. Nat. Commun. 2018, 9, 1318. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Banghart, M.R.; Sabatini, B.L. Photoactivatable Neuropeptides for Spatiotemporally Precise Delivery of Opioids in Neural Tissue. Neuron 2012, 73, 249–259. [Google Scholar] [CrossRef] [PubMed] [Green Version]
- Joffe, M.E.; Maksymetz, J.; Luschinger, J.R.; Dogra, S.; Ferranti, A.S.; Luessen, D.J.; Gallinger, I.M.; Xiang, Z.; Branthwaite, H.; Melugin, P.R.; et al. Acute Restraint Stress Redirects Prefrontal Cortex Circuit Function through MGlu5 Receptor Plasticity on Somatostatin-Expressing Interneurons. Neuron 2022, 110, 1068–1083.e5. [Google Scholar] [CrossRef] [PubMed]
- Hooks, B.M.; Lin, J.Y.; Guo, C.; Svoboda, K. Dual-Channel Circuit Mapping Reveals Sensorimotor Convergence in the Primary Motor Cortex. J. Neurosci. 2015, 35, 4418–4426. [Google Scholar] [CrossRef] [Green Version]
- Prasad, A.A.; Xie, C.; Chaichim, C.; Nguyen, J.H.; McClusky, H.E.; Killcross, S.; Power, J.M.; McNally, G.P. Complementary Roles for Ventral Pallidum Cell Types and Their Projections in Relapse. J. Neurosci. 2020, 40, 880–893. [Google Scholar] [CrossRef]
- Xia, S.H.; Yu, J.; Huang, X.; Sesack, S.R.; Huang, Y.H.; Schlüter, O.M.; Cao, J.L.; Dong, Y. Cortical and Thalamic Interaction with Amygdala-Toaccumbens Synapses. J. Neurosci. 2020, 40, 7119–7132. [Google Scholar] [CrossRef]
- Klapoetke, N.C.; Murata, Y.; Kim, S.S.; Pulver, S.R.; Birdsey-Benson, A.; Cho, Y.K.; Morimoto, T.K.; Chuong, A.S.; Carpenter, E.J.; Tian, Z.; et al. Independent Optical Excitation of Distinct Neural Populations. Nat. Methods 2014, 11, 338–346. [Google Scholar] [CrossRef] [Green Version]
- Lin, J.Y.; Knutsen, P.M.; Muller, A.; Kleinfeld, D.; Tsien, R.Y. ReaChR: A Red-Shifted Variant of Channelrhodopsin Enables Deep Transcranial Optogenetic Excitation. Nat. Neurosci. 2013, 16, 1499–1508. [Google Scholar] [CrossRef] [Green Version]
- Zeng, H.; Madisen, L. Mouse Transgenic Approaches in Optogenetics. In Progress in Brain Research; Elsevier: Amsterdam, The Netherlands, 2012; Volume 196, pp. 193–213. [Google Scholar]
- Nectow, A.R.; Nestler, E.J. Viral Tools for Neuroscience. Nat. Rev. Neurosci. 2020, 21, 669–681. [Google Scholar] [CrossRef]
- Ferenczi, E.A.; Vierock, J.; Atsuta-Tsunoda, K.; Tsunoda, S.P.; Ramakrishnan, C.; Gorini, C.; Thompson, K.; Lee, S.Y.; Berndt, A.; Perry, C.; et al. Optogenetic Approaches Addressing Extracellular Modulation of Neural Excitability. Sci. Rep. 2016, 6, 23947. [Google Scholar] [CrossRef] [Green Version]
- Shemesh, O.A.; Tanese, D.; Zampini, V.; Linghu, C.; Piatkevich, K.; Ronzitti, E.; Papagiakoumou, E.; Boyden, E.S.; Emiliani, V. Temporally Precise Single-Cell-Resolution Optogenetics. Nat. Neurosci. 2017, 20, 1796–1806. [Google Scholar] [CrossRef] [PubMed] [Green Version]
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Linders, L.E.; Supiot, L.F.; Du, W.; D’Angelo, R.; Adan, R.A.H.; Riga, D.; Meye, F.J. Studying Synaptic Connectivity and Strength with Optogenetics and Patch-Clamp Electrophysiology. Int. J. Mol. Sci. 2022, 23, 11612. https://doi.org/10.3390/ijms231911612
Linders LE, Supiot LF, Du W, D’Angelo R, Adan RAH, Riga D, Meye FJ. Studying Synaptic Connectivity and Strength with Optogenetics and Patch-Clamp Electrophysiology. International Journal of Molecular Sciences. 2022; 23(19):11612. https://doi.org/10.3390/ijms231911612
Chicago/Turabian StyleLinders, Louisa E., Laura. F. Supiot, Wenjie Du, Roberto D’Angelo, Roger A. H. Adan, Danai Riga, and Frank J. Meye. 2022. "Studying Synaptic Connectivity and Strength with Optogenetics and Patch-Clamp Electrophysiology" International Journal of Molecular Sciences 23, no. 19: 11612. https://doi.org/10.3390/ijms231911612