Patch Clamp Electrophysiology
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Patch-clamp (ex vivo) electrophysiology is a foundational technique in neuroscience and Meaghan's favourite thing to do in the lab! The Patch-clamp technique is a versatile electrophysiological tool for understanding ion channel behavior, cellular excitability, synaptic plasticity and connectivity within neural circuits.
To patch a neuron, a glass micropipette (with a tip 3-10 microns in diameter) containing an electrode is first filled with electrolyte solution. It is then fitted to a headstage connected to a digitizer and amplifier. A tissue slice is placed in a bath of aCSF, bubbled with carbogen to maintain neuronal health, and slices are visualized on a microscope. The glass micropipette is manipulated next to the membrane of a cell, and then negative pressure is applied to make a high-resistance seal between the cellular membrane and the pipette; this "cell-attached" configuration can be used to record firing activity of the neuron. Next, more negative pressure is applied to "break-in" to the cell, rupturing the membrane and causing the intracellular space to be contiguous with the solution in the pipette. This "whole-cell" configuration is then used to measure changes in current or voltage changes across the cellular membrane.
In the Creed lab, we use patch-clamp electrophysiology to record from individual, identified neurons in brain slices. We integrate this approach with optogenetic manipulations, pharmacology and light-evoked uncaging of neuromodulatory compounds. This allows us to determine how excitability, activity, or synaptic communication between neurons is modulated by dopamine or endogenous opioids, or how these parameters are changed in a disease state or by exposure to addictive drugs.
More unique to the Creed Lab, we use this technique to screen deep brain stimulation protocols designed to re-wire neural circuits, in order to validate their efficacy and underlying molecular mechanisms before moving to in vivo studies.
To patch a neuron, a glass micropipette (with a tip 3-10 microns in diameter) containing an electrode is first filled with electrolyte solution. It is then fitted to a headstage connected to a digitizer and amplifier. A tissue slice is placed in a bath of aCSF, bubbled with carbogen to maintain neuronal health, and slices are visualized on a microscope. The glass micropipette is manipulated next to the membrane of a cell, and then negative pressure is applied to make a high-resistance seal between the cellular membrane and the pipette; this "cell-attached" configuration can be used to record firing activity of the neuron. Next, more negative pressure is applied to "break-in" to the cell, rupturing the membrane and causing the intracellular space to be contiguous with the solution in the pipette. This "whole-cell" configuration is then used to measure changes in current or voltage changes across the cellular membrane.
In the Creed lab, we use patch-clamp electrophysiology to record from individual, identified neurons in brain slices. We integrate this approach with optogenetic manipulations, pharmacology and light-evoked uncaging of neuromodulatory compounds. This allows us to determine how excitability, activity, or synaptic communication between neurons is modulated by dopamine or endogenous opioids, or how these parameters are changed in a disease state or by exposure to addictive drugs.
More unique to the Creed Lab, we use this technique to screen deep brain stimulation protocols designed to re-wire neural circuits, in order to validate their efficacy and underlying molecular mechanisms before moving to in vivo studies.
In Vivo Electrophysiology
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We use in vivo electrophysiology in acute or chronic applications. Acute preparations involve using silicone probes in head-fixed mice, and allow for a high yield of isolated single units (neurons) during optogenetic tagging and manipulation experiments. In chronic preparations, electrode assemblies are implanted permanently, and changes in neuronal activity or in vivo functional connectivity can be tracked over longer time periods (weeks to months). In both cases, a microelectrode array is inserted into a specific brain region, where it can record voltage changes. During an action potential, electric current flowing across the plasma membrane can be measured as a change in voltage potential. Each microelectrode will record a signal that is the summed voltage change from activity of multiple neurons and processes. The signal is first filtered to isolate spike activity from the lower frequency local field potential (LFP). Spikes are then detected, and identifying features (i.e. amplitude, waveform characteristics) are used to cluster spikes and determine which spikes come from which neuron, a process known as spike sorting. In the Creed Lab, we combine optogenetic manipulations with in vivo electrophysiology to record from genetically-identified neurons within the basal ganglia as animals perform behavioral tasks. We also use the technique to assess connectivity between brain regions. This approach lets us ask how neural activity and network dynamics change over the course of pain chronification or drug withdrawal, and whether our neuromodulation interventions affect neural circuit function during behaviors. |
Fluorescence-Based Biosensors
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Fluorescence-based biosensors are powerful tools that can be used to monitor neural activity and neuromodulator signalling in vivo.
The biosensor is often genetically-encoded, which allows it to be targeted to specific cell types. Optic fibers or GRIN lenses are implanted into the brain region of interest along with the biosensor, in order to visualize dynamic changes in fluorescence over time in freely moving mice.
One commonly used biosensor is gCamp, an ultrasensitive, green fluorescent indicator that emits fluorescence based on calcium-binding. Since calcium conentration in neurons increases during an action potential, the fluorescence activity can be used as an indirect proxy measurement of neural activity (measured at cell bodies) or synaptic output (if measured at axons). Neuromodulator sensors are another exciting area of development: these are platforms that couple signalling through specific g-protein couple receptors (i.e. the receptors for dopamine or endogenous opioids) to conformational changes in intracellular GFP, which can then be detected as an increase in fluorescence (i.e. Patriarchi et al., Science 2018). Alternatively, platforms such as iTango (Lee*, Creed* et al., Nat Meth 2017) can be used to drive opsin or biosensor expression in functionally-defined ensembles of neurons based on their GPCR signalling, which can be subsequently manipulated (in the case of opsins) or monitored (biosensors).
In the Creed Lab, we are using sensors for dopamine and endogenous opioids to detect release of these neuromodulators in conjunction with neural spiking activity during behavioral assays. This allows us to determine how modulation of neural circuits changes over the course of drug withdrawal or with the emergence of chronic pain symptoms.
The biosensor is often genetically-encoded, which allows it to be targeted to specific cell types. Optic fibers or GRIN lenses are implanted into the brain region of interest along with the biosensor, in order to visualize dynamic changes in fluorescence over time in freely moving mice.
One commonly used biosensor is gCamp, an ultrasensitive, green fluorescent indicator that emits fluorescence based on calcium-binding. Since calcium conentration in neurons increases during an action potential, the fluorescence activity can be used as an indirect proxy measurement of neural activity (measured at cell bodies) or synaptic output (if measured at axons). Neuromodulator sensors are another exciting area of development: these are platforms that couple signalling through specific g-protein couple receptors (i.e. the receptors for dopamine or endogenous opioids) to conformational changes in intracellular GFP, which can then be detected as an increase in fluorescence (i.e. Patriarchi et al., Science 2018). Alternatively, platforms such as iTango (Lee*, Creed* et al., Nat Meth 2017) can be used to drive opsin or biosensor expression in functionally-defined ensembles of neurons based on their GPCR signalling, which can be subsequently manipulated (in the case of opsins) or monitored (biosensors).
In the Creed Lab, we are using sensors for dopamine and endogenous opioids to detect release of these neuromodulators in conjunction with neural spiking activity during behavioral assays. This allows us to determine how modulation of neural circuits changes over the course of drug withdrawal or with the emergence of chronic pain symptoms.
Viral-Genetic Tracing and Imaging
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Viral tracing manipulations are powerful tools for visualizing neuronal connectivity and giving us unprecedented insight into neuroanatomy in heterogeneous brain structures. We use recombinant viruses, such as adenoassociated virus (AAV) to label and visualize individual, genetically-defined neurons and their projection targets for anterograde neural tracing. Combining AAVs with optogenetic effectors enables us to perform functional connectivity experiments as well. For retrograde tracing, we can use retroAAVs, or non-viral approaches (i.e. cholera toxin or retrobeads) to visualize inputs to a brain area of interest. Alternatively, modified rabies virus allows us to do retrograde tracing in a cell-type specific way: rabies virus will infect cells, make new viral particles that are released and taken up ostensibly through synapses and will infect a retrogradely connected cell. To use it as a trans-synaptic tracer, a) an EnvA-pseudotyped version of the virus is used, and b) the genome of the virus has been modified to lack the rabies glycoprotein. Using an EnvA pseudotyped version means that the envelope protein, TVA, is required for the virus to initially infect starter cells. We can express the TVA using a separate virus only in genetically defined cell types of interest, limiting the rabies infection to these starter cells. By deleting the rabies glycoprotein, we ensure that the virus will only be able to undergo once cycle of replication and retrograde transmission, effectively limiting spread to monosynaptic inputs onto genetically defined starter cells.