Name: a method for high fidelity optogenetic control of individual pyramidal neurons in vivo
Uploaded: 2013-09-02T16:30:00
Duration: 13 min 44 s
Description: Watch this Scientific Journal Video about A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo at JoVE.com

This work was supported by National Institute on Drug Abuse (NIDA) grant R01 DA24040 (DCC), University of Colorado Innovative Seed Grant (DCC), and NIDA training grant T32 DA017637 (MVB).

1. Virus Storage and Preparation
<ol>
	<li>The use of replication-incompetent AAV vectors for delivering opsin genes to rodent brain is approved for Biosafety Level 1 (BSL-1) and requires proper protection and handling procedures as outlined in the U.S. Government publication, Biosafety in Microbiology and Biomedical Laboratories, available at the Centers for Disease Control's website at (<a href="http://www.cdc.gov/biosafety/publications/bmbl5/index.htm" target="_blank">http://www.cdc.gov/biosafety/publications/bmbl5/index.htm</a>).</li>
	<li>In order to avoid repeated freeze-thaw cycles, pipette virus from manufacturer's vial into smaller working aliquots in a Class II Biosafety Cabinet and store in a -80 &deg;C freezer.</li>
	<li>Prior to stereotactic injection, allow an aliquot to thaw on ice and then briefly spin down with a bench top centrifuge.</li>
	<li>Slowly backfill a Hamilton glass syringe and needle with a small amount of silicone or mineral oil. Make sure there are no visible air bubbles in the syringe barrel. Insert the syringe into a microinjector pump and attach the pump directly to the vertical stereotaxic arm (i.e. stereotaxic manipulator).</li>
	<li>Lower the stereotaxic arm until the tip of the needle touches the bottom of the virus aliquot tube. Use the controller for the microinjector pump to withdraw the desired volume of virus (1.5 &mu;l for a 1 &mu;l injection).</li>
	<li>All material and surfaces that come into contact with virus should be decontaminated with a disinfectant such as chlorine bleach (10%).</li>
</ol>
2. Surgery and Virus Injection
<ol>
	<li>Anesthetize animal with a ketamine (rat; 125 mg/kg, i.p.) - xylazine (rat; 10 mg/kg, i.p.) cocktail. To gauge effectiveness of anesthesia, check for a reflex response to a toe pinch and give supplemental doses of ketamine if needed. Using standard aseptic surgical methods, place animal into a stereotaxic apparatus (David Kopf Instruments).</li>
	<li>Make a midline incision through the skin on top of the animal skull with small surgical scissors or scalpel. Gently separate connective tissue and clean the top of the skull with a small bone scraper.</li>
	<li>Verify that the head of the animal is level such that the z coordinates for bregma and lambda are equal. Determine the stereotaxic coordinates for the target brain area from a brain atlas such as The Rat Brain in Stereotaxic Coordinates (George Paxinos and Charles Watson, Academic Press, 2007) or The Mouse Brain in Stereotaxic Coordinates (Keith B.J. Franklin and George Paxinos, Academic Press, 2007). Mark the intended site of injection with a surgical pen.</li>
	<li>Carefully thin the skull over the target area using a hand-held drill and stop when the drill bit reaches the bottom of the skull. Using a pair of extra fine forceps, remove the thinned bone in order to expose the dura.</li>
	<li>Position the syringe needle over the target area and lower the needle until it touches the dura and use this point to calculate the z coordinate. Very slowly lower the injection needle into the brain until the proper z position is reached. In this protocol, the prelimbic region of the medial prefrontal cortex (2.7 mm anterior to bregma, &plusmn; 0.5 mm lateral to the midline, and -2.2 mm from dura) or the dorsal subiculum (-6.0 mm anterior to bregma, &plusmn; 3.0 mm lateral to the midline, and -2.0 mm from dura) is targeted in an adult Sprague Dawley male rat (approx. 275 g).</li>
	<li>Inject virus at a rate of 0.1 &mu;l/min. After the injection is completed wait 10 min before withdrawing the needle to avoid backflow of the virus to the brain surface.</li>
	<li>Use a sewing suture with attached needle to seal the skin and apply antibiotics to the wound.</li>
	<li>The time course of ChR2 and NpHR functional expression from AAV vectors will vary depending on the experimental design and viral titer. For this experiment, in vivo recordings were performed 18-21 days post injection.</li>
</ol>
3. Light Delivery and In vivo Recording of ChR2 or NpHR-expressing Neurons
<ol>
	<li>Attach a tungsten electrode (~1-1.5 M&Omega;, MicroProbes) to a glass capillary tube (Fisher Scientific) using super glue.</li>
	<li>Use a fiber stripping tool (Thorlabs) to expose the bare end of a multimode optical fiber (200 &mu;m diameter core, Thorlabs) and clean fiber tip with ethanol. Very lightly score fiber tip with a wedge shaped diamond knife (Thorlabs) and carefully remove the excess fiber (e.g. with fine forceps). The fiber should cleave easily at the score.</li>
	<li>Connect the FC end of the fiber to the PC connector on the output port of a 200 mW single diode laser (<a href="http://www.lumiphy.com" target="_blank">www.Lumiphy.com</a>) with the desired wavelength. Make sure that appropriate protective eyewear is worn at all times when working with lasers.</li>
	<li>Measure the laser intensity at the tip of the optic fiber using an optical power meter (<a href="http://www.lumiphy.com/" target="_blank">www.Lumiphy.com</a> ). For in vivo recordings, light intensity as little as 20-100 mW/mm2 can reliably evoke ChR2- or NpHR-mediated activation or silencing, respectively, of neuronal activity.</li>
	<li>Insert the optical fiber into the capillary tube. Position the fiber tip approximately 500 &mu;m above the tip of tungsten electrode and tie a thin suture thread twice at a few points near the tip so that the electrode and the fiber are straight. Make sure that the distance between the tip of electrode and the nearest knot is long enough for insertion to the target region.</li>
	<li>Prepare the animal as in steps 2.1 and 2.2 above. Use the initial craniotomy as a guide to make a new clean small window in the skull for positioning the optrode (<a href="http://www.lumiphy.com/" target="_blank">www.Lumiphy.com</a> ). Make a small incision on the dura using a fine needle.</li>
	<li>Carefully lower the optrode through the skull window and into the brain to the transduced region using a micromanipulator (Narishige). Then advance the optrode slowly in 10-50 &mu;m steps until the emergence of a light-responsive cell.</li>
	<li>A custom software user interface (NeuroLux Pro <a href="http://www.lumiphy.com/" target="_blank">www.Lumiphy.com</a>) written in LabVIEW (National Instruments) is used to control the single diode laser and to visualize and record ongoing neural activity. Recorded signals are captured with an ExAmp-20K (Kation Scientific) amplifier band-pass filtered (0.3-8 kHz), and a National Instruments analog to digital board (NI USB-6009). The NeuroLux Pro user interface allows the choice of analog inputs (choose two channels for the neuronal and TTL signal) and digital output. The user may define the sampling rate (20 kHz), the baseline period (pre-light), and post-light period. The user may define the TTL laser stimulation pulse width, frequency and stimulation period (if frequency is 20 Hz and stimulation period is 500 msec, 10 laser pulses with defined pulse width are delivered). The user also can choose continuous light delivery (e.g., Figure 1). For repeated recordings, the user also may define the number of pulse trains and the interval between trains. Data can be acquired with or without recording to disk.</li>
	<li>Following recording, animals are anesthetized with the ketamine/xylazine cocktail and transcardially perfused with physiological saline and paraformaldehyde (4%) in order to verify optrode placement and opsin expression.</li>
</ol>

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P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multimodal+fast+optical+interrogation+of+neural+circuitry.">Multimodal fast optical interrogation of neural circuitry.</a> Nature. 446 (7136), 633-639 (2007).</li><li>Nagel, G., Brauner, M., Liewald, J. F., Adeishvili, N., Bamberg, E., Gottschalk, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light+activation+of+channelrhodopsin-2+in+excitable+cells+of+Caenorhabditis+elegans+triggers+rapid+behavioral+responses.">Light activation of channelrhodopsin-2 in excitable cells of Caenorhabditis elegans triggers rapid behavioral responses.</a> Curr Biol. 15 (24), 2279-2284 (2005).</li><li>Adamantidis, A. R., Zhang, F., Aravanis, A. 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The authors declare that they have no competing financial interests.

A wide array of techniques are available for genetically-targeting microbial opsin genes to discrete brain regions in rodents. Viral gene delivery provides a relatively inexpensive and quick approach for mediating ChR2 and NpHR expression with cell-type specificity. AAV vector systems are a common choice for use in optogenetic experiments due to high production titers that remain stable during storage, its lack of pathogenicity, and its ability to produce long-term gene expression 10. Many of the opsin constructs with cell-type specific promoters are commercially available in a variety of AAV serotypes from vector core facilities such as the University of Pennsylvania (<a href="http://www.med.upenn.edu/gtp/vectorcore" target="_blank">http://www.med.upenn.edu/gtp/vectorcore</a>) or the University of North Carolina at Chapel Hill (<a href="http://genetherapy.unc.edu" target="_blank">http://genetherapy.unc.edu</a>). One drawback to AAV technology is the limited packaging capacity (~4.7 kb), which places a restriction on the transgene cassette size that can be used for cell-specific targeting. As an alternative, lentiviral vectors, which have a larger packaging capacity, are able to accommodate opsin targeting under the control of larger promoter sequences.
The use of an optrode allows for reliable detection of electrophysiological signals in combination with temporally-precise light delivery. As described above, however, sufficient care is needed in its construction. Since the cleaved optical fiber is fragile, tying the suture thread too tightly may cause the fiber to break. Even though the optrode can be used for multiple recordings, the electrode and optical fiber should be cleaned (or manually cleaved in the case of the optical fiber bare end) prior to insertion because the impedance of the electrode and quality of the delivered light will degrade with repeated use.
During electrophysiological recordings it is important that the optrode be advanced slowly. The optrode used here has approximately 350 &mu;m thickness (tungsten electrode: 125 &mu;m; core of optic fiber: 200 &mu;m) and lowering it fast could lead to movement of brain tissue. Light-induced artifacts also might be considered with particular recording and light delivery set-ups 11-12. Although we found no light-induced artifact in our experimental condition, recordings outside the transduced brain area can potentially be used as a control for light artifacts. Another consideration during the recording process is the required light intensity for observing changes in ChR2- or NpHR-expressing neurons, especially for long duration recordings. Strong laser intensity and long laser illumination could result in the tissue damage and/or a decreased response to subsequent delivered light 12-13. For behavioral experimentation the use of a control viral vector is required for eliminating any effect due to heat delivered from the fiber. For many of the commercially available optogenetic constructs there are complementary control constructs in which the opsin gene sequence has been removed.
It should be noted that engineered ChR2 variants with improved properties and kinetics have been developed along with other silencing opsins that may be better suited for certain experimental questions 14-15. For instance, a new ChR2 variant established via targeted mutagenesis, termed "ChETA", can drive high-fidelity neuronal spiking at frequencies (up to at least 200 Hz) above that of the original ChR2 14.
The combination of in vivo light delivery and simultaneous recording of neuronal responses is a critical step in establishing causal relationships between patterned activity in genetically-targeted cell populations and corresponding time-locked behavioral events. The procedures and hardware outlined here provide a straightforward approach for implementing optogenetics for in vivo head-fixed recordings in rodents.

The ability to activate or silence a specific cell type within a neural circuit in a temporally precise fashion is critical for understanding how neural circuits process different types of information underlying emotion and cognition. Experimental control over intact neural activity has employed loss-/gain-of-function tools (e.g., electrical stimulation, pharmacological modulation, lesion) that do not provide the required selectively for controlling specific populations of neurons, either on a temporal or spatial scale. Directly addressing these technological challenges, the development and application of genetically-encodable light-sensitive tools has enabled neuroscientists to control the electrical activity of select cell types during well-defined behavioral events. Many of these light-sensitive proteins are of microbial origin, with the light-gated cation channel, channelrhodopsin-2 (ChR2) 1, and the light-driven chloride pump, halorhodopsin (NpHR) 2,3, in widespread use.
A major advantage of optogenetics is the ability to genetically-target specific cell populations in heterogeneous brain regions and the technique has been successfully applied to a number of model organisms (invertebrates to nonhuman primates) 4-6. Many optogenetic transgenic animals have been generated and are commercially available 7,8, however establishing transgenic lines can be labor intensive and cost prohibitive. Here we describe a protocol for virally-mediated delivery of ChR2 and NpHR genes under the CaMKII&alpha; promoter in rat prelimbic cortex and dorsal subiculum using a recombinant adeno-associated virus (AAV) vector. Within the forebrain, CaMKII&alpha; expression is exclusive to glutamatergic pyramidal neurons 9. AAV is commonly used for basic research due to its relative ease of production and lack of pathogenicity, as well as the strong and persistent transgene expression that has been achieved with these vectors 10. Additionally we outline the steps and hardware for simultaneous light delivery and recording in anesthetized head-fixed rats.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Syringe </td>
 <td>Hamilton</td>
 <td>7653-01</td>
 <td>10 &mu;l</td>
 </tr>
 <tr>
 <td>Removable Needle</td>
 <td>Hamilton</td>
 <td>7803-03</td>
 <td>31 gauge, beveled tip</td>
 </tr>
 <tr>
 <td>Microinjector Pump</td>
 <td>World Precision Instruments</td>
 <td>UMP3</td>
 <td></td>
 </tr>
 <tr>
 <td>Pump Controller</td>
 <td>World Precision Instruments</td>
 <td>SYS-MICRO4</td>
 <td></td>
 </tr>
 <tr>
 <td>Silicone Oil</td>
 <td>Alfa Aesar</td>
 <td>A12728</td>
 <td></td>
 </tr>
 <tr>
 <td>Laser Protective Eyeware</td>
 <td>Kentek </td>
 <td>KMT-4501</td>
 <td></td>
 </tr>
 <tr>
 <td>Multimode Optical Fiber</td>
 <td>Thorlabs</td>
 <td>BFL37-200</td>
 <td>200 &mu;m diameter core, 0.37 NA</td>
 </tr>
 <tr>
 <td>Fiber Stripping Tool</td>
 <td>Thorlabs</td>
 <td>T12S21</td>
 <td>for 200 &mu;m diameter core multimode fiber</td>
 </tr>
 <tr>
 <td>Optical Power Meter</td>
 <td>Lumiphy LLC</td>
 <td><a href="http://www.lumiphy.com/" target="_blank">www.lumiphy.com</a></td>
 <td></td>
 </tr>
 <tr>
 <td>Photodetector</td>
 <td>Newport</td>
 <td>818-SL/DB</td>
 <td></td>
 </tr>
 <tr>
 <td>Blue Laser (445-473 nm, 100-200 mW)</td>
 <td>Lumiphy LLC</td>
 <td><a href="http://www.lumiphy.com" target="_blank">www.lumiphy.com</a></td>
 <td>coupled to a 200 &mu;m multimode fiber with FC/PC adapter </td>
 </tr>
 <tr>
 <td>Green Laser (532 nm, 100-200 mW)</td>
 <td>Lumiphy LLC</td>
 <td><a href="http://www.lumiphy.com" target="_blank">www.lumiphy.com</a></td>
 <td>coupled to a 200 &mu;m multimode fiber with FC/PC adapter </td>
 </tr>
 <tr>
 <td>Tungsten/Fiber Optrode</td>
 <td>Lumiphy LLC</td>
 <td><a href="http://www.lumiphy.com/" target="_blank">www.lumiphy.com</a></td>
 <td>Lumitrode</td>
 </tr>
 <tr>
 <td>Glass Capillary Tube</td>
 <td>Fisher Scientific</td>
 <td>22-362-566</td>
 <td></td>
 </tr>
 <tr>
 <td>Hydraulic Micromanipulator</td>
 <td>Narishige</td>
 <td>MO-22</td>
 <td></td>
 </tr>
 <tr>
 <td>Amplifier</td>
 <td>Kation Scientific</td>
 <td>ExAmp-20K</td>
 <td></td>
 </tr>
 <tr>
 <td>Data Acquistion Device</td>
 <td>National Instruments</td>
 <td>NI USB-6009</td>
 <td></td>
 </tr>
 <tr>
 <td>Rodent Head Restraint for Recording</td>
 <td>Lumiphy LLC</td>
 <td><a href="http://www.lumiphy.com/" target="_blank">www.lumiphy.com</a></td>
 <td></td>
 </tr>
 <tr>
 <td>Small Animal Stereotaxic Unit</td>
 <td>David Kopf Instruments</td>
 <td>Model 963</td>
 <td></td>
 </tr>
 </tbody>
</table>

a method for high fidelity optogenetic control of individual pyramidal neurons in vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

Figure 1 depicts a screenshot of the customized software (NeuroLux Pro) developed in LabVIEW environment (National Instruments) used for simultaneous recording of neuronal activity and controlling light pulse parameters (i.e. pulse frequency, pulse width, stimulation period). The software program sends a command signal to a digital acquisition device that generates TTL pulses for control over the single diode laser. Additionally the software program displays and stores the recorded neuronal activity and delivered TTL signals. These signals are digitized through the acquisition device at a defined sample rate (e.g., 20 kHz). Recorded signals are saved as a two-dimensional double-precision array in a binary file.
Figure 2 shows the custom optrode consisting of a tungsten electrode attached to an optical fiber. An optical fiber is fixed to the electrode using thin suture thread at three points. If needed, super glue can be used to adhere them. However, avoid permanently fixing the optical fiber to the electrode because although the electrode may be reused several times, light transmission through the fiber will degrade with repeated use and should be cleaved and cleaned or replaced with every insertion into the brain.
Left panels of Figure 3 show fluorescent images of enhanced yellow fluorescent protein, indicating actual ChR2 and NpHR expression. These photographs show representative NpHR expression in rat dorsal subiculum (Figure 3A) and ChR2 expression in prelimbic cortex (Figure 3B). Viral expression was restricted mostly to the dorsal subiculum and prelimbic cortex (e.g. some fluorescence was observed in dentate gyrus (Figure 3A, left)). The tracks of the electrode (thin track, arrow head) and optical fiber (thick track, arrow) were also observed.
Left panel of Figure 4 shows that ChR2 induced temporally precise spiking in rat prelimbic cortex. Figure 4A is an example trace of ChR2-triggered action potentials in response to 20 Hz delivery of 10 msec blue light pulses to rat prelimbic cortex. Raster plot (Figure 4B) shows ChR2-induced activation in six representative neurons. All recorded neurons showed light-evoked spiking with perfect fidelity. In contrast to ChR2, NpHR rapidly and reversibly silenced spontaneous activity in vivo in rat prelimbic cortex (right panel of Figure 4). Figure 4C is an example trace showing that continuous 532 nm illumination (10 s) of the prelimbic cortex expressing NpHR under the CaMKll&alpha; promoter eliminates spontaneous single-unit activity in vivo. Note that the complete silencing of prelimbic pyramidal cell activity is time-locked to the 10 sec continuous light delivery. Lower raster plot (Figure 4D) shows NpHR-induced silencing in six representative neurons. Both in vivo recordings are typical of those obtained from adult Sprague-Dawley rats in which AAV-mediated delivery of microbial opsin genes occurred 18-21 days prior.
Figure 5 shows the result of repeated ChR2 stimulation of a rat prelimbic pyramidal neuron. Blue light pulse (10 msec) was delivered at 20 Hz for 5 sec (100 pulses total). Voltage traces of the light-evoked spiking were repeatedly acquired every 2 min during a 2 hr recording session (61 repetitions total). Even when using this repetitive stimulation protocol, ChR2 induced stable and robust spiking in response to the light delivery.
Figures 6 and 7 show the results of NpHR-induced photoinhibition and ChR2-driven photoactivation of rat subicular neuron activity. As is the case in prelimbic cortex, NpHR and ChR2 were able to mediate the light-induced time-locked inhibition and activation of the opsin-transduced subicular neurons with high reproducibility.
<img alt="Figure 1" src="/files/ftp_upload/50291/50291fig1.jpg" /> 
Figure 1. Screenshot of the NeuroLux Pro software interface for simultaneous light delivery and electrophysiological recording. Pictured trace shows NpHR induced silencing of spontaneous activity of rat prelimbic pyramidal cell in response to 10 sec of continuous 532 nm light delivery. <a href="https://www.jove.com/files/ftp_upload/50291/50291fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 2" src="/files/ftp_upload/50291/50291fig2.jpg" /> 
Figure 2. High power image of custom optrode. An optical fiber is inserted into glass capillary tube attached to a tungsten electrode and fixed to the electrode using suture thread. The center-to-center distance between the electrode tip and the fiber tip is approximately 500 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/50291/50291fig3.jpg" /> 
Figure 3. Viral expression and optrode placement. (Left) Photographs show representative NpHR expression in dorsal subiculum (A) and ChR2 expression in prelimbic cortex (B). (Right) Schematic that represents the location where each photograph was taken. The arrowhead and arrow in each photograph indicate the location of the tungsten electrode and optical fiber, respectively. SUB: subiculum, DG: dentate gyrus, PL: prelimbic cortex. <a href="https://www.jove.com/files/ftp_upload/50291/50291fig3large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 4" src="/files/ftp_upload/50291/50291fig4.jpg" /> 
Figure 4. In vivo electrophysiological recordings from ChR2 or NpHR-transduced rat prelimbic cortex. (A) Example trace of ChR2-triggered action potentials in response to 20 Hz delivery of blue (473 nm) light pulses (10 msec, blue bar). (B) Raster plot showing ChR2-induced spiking in six representative neurons. Each unit activity is plotted as a dot. (C) Example trace of NpHR-induced suppression of spontaneous activity during continuous green (532 nm) light illumination (10 sec, green bar). (D) Raster plot showing NpHR-induced silencing in six representative neurons. Each unit activity is plotted as a dot. <a href="https://www.jove.com/files/ftp_upload/50291/50291fig4large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 5" src="/files/ftp_upload/50291/50291fig5.jpg" /> 
Figure 5. Repetitive 20 Hz ChR2-driven spiking of a rat prelimbic pyramidal neuron in vivo. (A) Voltage traces of the light-evoked spiking acquired at the time points 0, 30, 60, 90, 120 min after the beginning of the recordings. (B) Raster plot showing all 61 repetitions (2 min inter-trial interval, 120 min total) of the light-induced activation. Each unit activity is plotted as a dot. <a href="https://www.jove.com/files/ftp_upload/50291/50291fig5large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 6" src="/files/ftp_upload/50291/50291fig6.jpg" /> 
Figure 6. In vivo electrophysiological recordings from NpHR-transduced rat dorsal subiculum. (A) Example trace showing that 10 sec continuous 532 nm illumination (green bar) of the dorsal subiculum expressing NpHR eliminates spontaneous activity. (B) Average waveforms of the two recorded units from the above trace. The amplitude threshold was used for identifying two distinct neurons. (C) Raster plot showing five repetitions of NpHR-induced silencing of these units. Each unit activity is plotted as a dot.
<img alt="Figure 7" src="/files/ftp_upload/50291/50291fig7.jpg" /> 
Figure 7. Repetitive 20 Hz CR2-driven spiking of a rat dorsal subicular neuron in vivo. (A) Voltage traces of the light-evoked spiking acquired at the time points 0, 30, 60, 90, 120 min after the beginning of the recordings. Inset: typical bursting activity of this cell. (B) Raster plot showing all 61 repetitions (2 min inter-trial interval, 120 min total) of the light-induced activation. Each unit activity is plotted as a dot. <a href="https://www.jove.com/files/ftp_upload/50291/50291fig7large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo at JoVE.com

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

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Research

JoVE Journal

Neuroscience

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic hippocampal neurons can often grow successfully on glass coverslips at high density under serum-free conditions, but low density cultures typically require a supply of trophic factors by co-culturing them with a glia feeder layer, preparation of which can be time-consuming and laborious. In addition, the presence of glia may confound interpretation of results and preclude studies on neuron-specific mechanisms. Here, a simplified method is presented for ultra-low density (~2,000 neurons/cm2), long-term (&#62;3 months) primary hippocampal neuron culture that is under serum free conditions and without glia cell support. Low density neurons are grown on poly-D-lysine coated coverslips, and flipped on high density neurons grown in a 24-well plate. Instead of using paraffin dots to create a space between the two neuronal layers, the experimenters can simply etch the plastic bottom of the well, on which the high density neurons reside, to create a microspace conducive to low density neuron growth. The co-culture can be easily maintained for &#62;3 months without significant loss of low density neurons, thus facilitating the morphological and physiological study of these neurons. To illustrate this successful culture condition, data are provided to show profuse synapse formation in low density cells after prolonged culture. This co-culture system also facilitates the survival of sparse individual neurons grown in islands of poly-D-lysine substrates and thus the formation of autaptic connections.

Low density cultures of primary hippocampal neurons usually require glia feeder layer to supply neurotrophic factors and sustain longevity. We describe here a simplified method to culture ultra-low density neurons on glass coverslips in the presence of a high density neuronal feeder layer, which facilitates investigation of specific neuronal-autonomous mechanisms.

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

Method for High Speed Stretch Injury of Human Induced Pluripotent Stem Cell-derived Neurons in a 96-well Format

Traumatic brain injury (TBI) is a major clinical challenge with high morbidity and mortality. Despite decades of pre-clinical research, no proven therapies for TBI have been developed. This paper presents a novel method for pre-clinical neurotrauma research intended to complement existing pre-clinical models. It introduces human pathophysiology through the use of human induced pluripotent stem cell-derived neurons (hiPSCNs). It achieves loading pulse duration similar to the loading durations of clinical closed head impact injury. It employs a 96-well format that facilitates high throughput experiments and makes efficient use of expensive cells and culture reagents. Silicone membranes are first treated to remove neurotoxic uncured polymer and then bonded to commercial 96-well plate bodies to create stretchable 96-well plates. A custom-built device is used to indent some or all of the well bottoms from beneath, inducing equibiaxial mechanical strain that mechanically injures cells in culture in the wells. The relationship between indentation depth and mechanical strain is determined empirically using high speed videography of well bottoms during indentation. Cells, including hiPSCNs, can be cultured on these silicone membranes using modified versions of conventional cell culture protocols. Fluorescent microscopic images of cell cultures are acquired and analyzed after injury in a semi-automated fashion to quantify the level of injury in each well. The model presented is optimized for hiPSCNs but could in theory be applied to other cell types.

Here we present a method for a human in vitro model of stretch injury in a 96-well format on a timescale relevant to impact trauma. This includes methods for fabricating stretchable plates, quantifying the mechanical insult, culturing and injuring cells, imaging, and high content analysis to quantify injury.