This work was supported by National Institutes of Health Grants (R01NS085215 to K.F., T32 GM107000 and F30MH122146 to A.C.). The authors thank Ms. Naoe Watanabe for skillful technical assistance.

All animal protocols were reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Massachusetts Medical School. Slice culture preparation, plasmid preparation, and electroporation are also detailed in our previously published methods and can be referred to for additional information10.
1. Slice culture preparation
<ol>
	<li>Prepare mouse organotypic hippocampal slice cultures as previously described11, using postnatal 6- to 7-day old mice of either sex.
	<ol>
		<li>Prepare dissection media for organotypic slice culture consisting of (in mM): 238 sucrose, 2.5 KCl, 1 CaCl2, 4 MgCl2, 26 NaHCO3, 1 NaH2PO4, and 11 glucose in deionized water, then gas with 5% CO2/95% O2 to a pH of 7.4.</li>
		<li>Prepare organotypic slice culture media consisting of: 78.8% (v/v) Minimum Essential Medium Eagle, 20% (v/v) horse serum, 17.9 mM NaHCO3, 26.6 mM glucose, 2 M CaCl2, 2 M MgSO4, 30 mM HEPES, insulin (1 &#181;g/mL), and 0.06 mM ascorbic acid, pH adjusted to 7.3. Adjust the osmolarity to 310-330 osmol using an osmometer.</li>
		<li>Dissect hippocampi out from the whole brain by using two spatulas, and slice (400 &#181;m) using a tissue chopper. Separate slices by using two forceps and transfer to 30 mm cell culture inserts in a 6 well plate filled with culture media (950 &#181;L) underneath the inserts.</li>
	</ol>
	</li>
	<li>Store organotypic slice cultures in a tissue culture incubator (35&#176;C, 5% CO2) and change the slice culture media every two days.</li>
</ol>
2. Plasmid preparation
<ol>
	<li>Prepare the plasmid for the gene of interest.
	<ol>
		<li>Subclone enhanced green fluorescent protein (EGFP) gene into a pCAG vector.</li>
		<li>Purify pCAG-EGFP plasmid with an endotoxin-free purification kit and dissolve in an internal solution that consists of diethyl pyrocarbonate-treated water containing 140 mM K-methanesulfonate, 0.2 mM EGTA, 2 mM MgCl2, and 10 mM HEPES, adjusted to pH 7.3 with KOH (plasmid concentration: 0.1 &#181;g/&#181;L).</li>
	</ol>
	</li>
</ol>
3. Glass pipette preparation
<ol>
	<li>Pull borosilicate glass pipettes (4.5 &#8211; 8 M&#937;) on a micropipette puller (Figure 1A). 
	NOTE: The glass pipettes used for whole-cell patch clamp recordings are ideal for electroporation.</li>
	<li>Bake glass pipettes overnight at 200&#176;C to sterilize.</li>
	<li>Check the size of the pipette tip under a dissection microscope to approximate the electrical resistance.
	<ol>
		<li>Optional:Verify pipette resistance by attaching the pipette to the electroporation electrode and use the micromanipulator to maneuver the pipette tip into filter-sterilized artificial cerebrospinal fluid (aCSF) containing (in mM): 119 NaCl, 2.5 KCl, 0.5 CaCl2, 5 MgCl2, 26 NaHCO3, 1 NaH2PO4 and 11 glucose in deionized water, gassed with 5% CO2/95% O2 to a pH of 7.4. Confirm the actual resistance using the readout on the electroporator. 
		NOTE: The sharper the pipette tip, the larger the electrical resistance. The pipette resistance should be below 10 M&#937;. Glass pipettes with high pipette resistance (Figure 1B) often clog at the tip during repeated electroporation.</li>
	</ol>
	</li>
</ol>
4. Electroporation rig setup
<ol>
	<li>Install the electroporator to a standard whole-cell electrophysiology rig, equipped with an upright microscope mounted on a shifting table with a micromanipulator and peristaltic pump.</li>
	<li>Install the headstage of the electroporator onto a micromanipulator and connect a pair of speakers to the electroporator. Connect the electroporator to a foot pedal which can be used to send a pulse when ready. 
	NOTE: The speakers emit a tone when turned on, which is an indicator of the electrical resistance at the electrode. This makes it possible to determine relative changes in resistance without pulling attention away from the procedure.</li>
</ol>
5. Electroporation preparation
<ol>
	<li>Transfer slice culture inserts from 6 well plates to 3 cm Petri dishes loaded with 900 &#181;L of culture media and store in a tabletop CO2 incubator until ready to perform electroporation.
	<ol>
		<li>Preincubate fresh culture inserts with slice culture media (1 mL) for at least 30 min in a 3.5 cm Petri dish to culture slices after electroporation.</li>
	</ol>
	</li>
	<li>Clean and prepare the rig for electroporation.
	<ol>
		<li>Perfuse the lines with 10% bleach for 5 min to sterilize the tubing and chamber prior to beginning the experiment for the day.</li>
		<li>Perfuse the lines with deionized autoclaved water for at least 30 min to rinse completely.</li>
		<li>Perfuse the lines with filter-sterilized aCSF containing 0.001 mM tetrodotoxin (TTX). 
		NOTE: TTX minimizes cellular toxicity and death due to overexcitation of interneurons10.</li>
	</ol>
	</li>
	<li>Set the electroporator&#8217;s pulse parameters: amplitude of &#8211;5 V, square pulse, train of 500 ms, frequency of 50 Hz, and a pulse width of 500 &#181;s.</li>
	<li>Fill glass pipette with 5 &#181;L of plasmid-containing internal solution.
	<ol>
		<li>Remove any trapped air bubbles from the pipette tip by flicking and gently tapping the tip multiple times.</li>
		<li>Check the tip for damage by visualizing it under a dissection microscope or by repeating step 3.3.1 to check the pipette resistance. 
		NOTE: If the tip is damaged, the glass pipette must be discarded, and this step must be repeated with a new glass pipette previously prepared in step 3.</li>
	</ol>
	</li>
	<li>Securely attach the pipette tip to the electrode and turn the speakers on. Record the readout (the pipette&#8217;s resistance) of the electroporator when the tip has made contact with&#160;the aCSF medium.</li>
	<li>Cut the culture insert membrane using a sharp blade and isolate one slice culture. Carefully transfer the slice culture to the electroporation chamber by using sharp angled forceps and fix its position with a slice anchor.
	<ol>
		<li>Do not keep the slice culture outside of the incubator for more than 30 min at a time to prevent side effects such as changes in neuronal health or function12.</li>
	</ol>
	</li>
</ol>
6. Electroporate cells of interest
<ol>
	<li>Apply positive pressure to the pipette with mouth or by using a 1 mL syringe (0.2 - 0.5 mL pressure) attached to the tubing.</li>
	<li>Use the micromanipulator&#8217;s 3-dimensional knob controls to maneuver the pipette tip near the surface of the slice culture.</li>
	<li>Choose a target cell and approach it, keeping the positive pressure applied until a dimple forms on the cell surface, visible on the microscope.</li>
	<li>Perform pressure cycles.
	<ol>
		<li>Quickly apply mild negative pressure by mouth so that a loose seal forms between the pipette tip and the plasma membrane, indicated visually by the membrane going up into the pipette tip somewhat. Observe an increase (~2.5x the initial resistance) in pipette resistance by listening for an increase in tone coming from the speakers. Quickly re-apply positive pressure so that the dimple re-forms.</li>
		<li>Immediately complete at least two more pressure cycles without pausing, then hold negative pressure for 1 s. 
		NOTE: Pausing between cycles, applying too much pressure, or holding the negative pressure for too long can cause significant cell damage and possibly cause the cell to die during electroporation.</li>
	</ol>
	</li>
	<li>Quickly pulse the electroporator once using the foot pedal when the tone from the speakers reaches a stable apex in pitch, indicating peak electrical resistance. Do not wait at the peak resistance for more than 1 s before sending the pulse. 
	NOTE: We have observed no off-target electroporation when using this protocol10. Only the cells in contact with the glass pipette during pressure cycles were transfected. Positioning the pipette near other neurons does not result in gene transfection.</li>
	<li>Gently retract the pipette approximately 100 &#181;m from the cell without applying pressure.</li>
	<li>Re-apply positive pressure, verifying that the resistance is similar to the recorded readout in step 5.5, then approach the next cell.
	<ol>
		<li>Remove potential clogs, indicated visually or by a significantly increased (&#62;15% higher) pipette resistance after electroporation, by applying positive pressure. 
		NOTE: If there is no visible clog and the resistance is still significantly higher, discard the pipette and use a new one. On an average, a pipette can be used for up to 20 electroporation events if the user is careful10.</li>
	</ol>
	</li>
	<li>After electroporation, transfer the slice culture onto a fresh culture insert, and incubate at 35&#176;C in the incubator for up to 3 days.</li>
</ol>
7. Fixation, staining and imaging of organotypic hippocampal slice cultures
<ol>
	<li>Fix electroporated organotypic slice cultures 2 &#8211; 3 days after transfection with 4% paraformaldehyde and 4% sucrose in 0.01 M phosphate buffered saline (1x PBS) for 1.5 h at room temperature.</li>
	<li>Remove fixative and incubate slices in 30% sucrose in 0.1 M phosphate buffer (1x PB) for 2 h.</li>
	<li>Place slices on a slide glass and freeze them by putting the slide glass on top of crushed dry ice. Thaw the slices at room temperature and transfer them to a 6 well plate filled with 1x PBS.</li>
	<li>Stain the slices with mouse anti-GFP and rabbit anti-RFP antibodies in GDB buffer (0.1% gelatin, 0.3% Triton X-100, 450 mM NaCl, and 32% 1x PB, pH 7.4) for 2 h at room temperature.</li>
	<li>Wash slices with 1x PBS three times at room temperature for 10 min each wash.</li>
	<li>Incubate slices with anti-mouse Alexa 488-conjugated secondary antibody and anti-rabbit Alexa 594-conjugated secondary antibody in GDB buffer for 1 h at room temperature. Incubate slices with DAPI (4 &#181;g/mL) in 1x PBS for 10 min at room temperature.</li>
	<li>Wash slices with 1x PBS three times at room temperature for 3 min each wash.</li>
	<li>Mount slices on glass slides using mounting medium and perform fluorescence imaging.</li>
</ol>

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The authors declare that they have no conflicts of interest.

We describe here an electroporation method that transfects both excitatory and inhibitory neurons with high efficiency and precision. Our optimized electroporation protocol has three innovative breakthroughs to achieve highly efficient gene transfection. Our first modification was to increase pipette size compared with previously published protocols3,5,6. This change enabled us to electroporate many neurons without pipette clogg...

In molecular biology, one of the most important considerations to an investigator is how to deliver a gene of interest into a cell or population of cells to elucidate its function. The different methods of delivery can be categorized as either biological (e.g., a viral vector), chemical (e.g., calcium phosphate or lipid), or physical (e.g., electroporation, microinjection, or biolistics)1,2. Biological methods are highly efficient and can be cell type-specific but are limited by the development of specific genetic tools. Chemical approaches are very powerful in vitro, but transfections are generally random; further, these approaches are mostly reserved for primary cells only. Of the physical approaches, biolistics is the simplest and easiest from a technical point of view, but again produces random transfection results at a relatively low efficiency. For applications which require transfer into specific cells without the need for developing genetic tools, we look toward single-cell electroporation3,4.
Whereas electroporation used to refer only to field electroporation, over the past twenty years, multiple in vitro and in vivo single-cell electroporation protocols have been developed to improve specificity and efficiency5,6,7, demonstrating that electroporation can be used to transfer genes to individual cells and can, therefore, be extremely precise. However, the procedures are technically demanding, time-consuming, and relatively inefficient. Indeed, more recent papers have investigated the feasibility of mechanized electroporation rigs8,9, which can help to eliminate several of these barriers for investigators interested in installing such robotics. But for those looking for simpler means, the problems with electroporation, namely cell death, transfection failure, and pipette clogging, remain a concern.
We recently developed an electroporation method that uses larger-tipped glass pipettes, milder electrical pulse parameters, and a unique pressure cycling step, which generated a much higher transfection efficiency in excitatory neurons than previous methods, and enabled us for the first time to transfect genes in inhibitory interneurons, including somatostatin-expressing inhibitory interneurons in the hippocampal CA1 region of mouse organotypic slice culture10. However, the reliability of this electroporation method in different inhibitory interneuron types and neuronal developmental stages has not been addressed. Here, we demonstrated that this electroporation technique is capable of transfecting genes into both excitatory neurons and different classes of interneurons. Importantly, transfection efficiency was high regardless of days in vitro (DIV) slice culture age tested. This established and user-friendly technique is highly recommended to any investigator interested in using single-cell electroporation for different cell types in the context of in vitro mouse brain tissue.

<table><tbody><tr><td>&lt;strong&gt;Plasmid preparation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Plasmid Purification Kit</td><td>Qiagen</td><td>12362</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Organotypic slice culture preparation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>6 Well Plates</td><td>GREINER BIO-ONE</td><td>657160</td><td></td></tr><tr><td>Dumont #5/45 Forceps</td><td>FST</td><td>#5/45</td><td>Angled dissection forceps for organotypic slice culture preparation</td></tr><tr><td>Flask Filter Unit</td><td>Millipore</td><td>SCHVU02RE</td><td>Filtration and storage of culture media</td></tr><tr><td>Incubator</td><td>Binder</td><td>BD C150-UL</td><td></td></tr><tr><td>McIlwain Tissue Chopper</td><td>TED PELLA, INC.</td><td>10180</td><td>Tissue chopper for organotypic slice culture preparation</td></tr><tr><td>Millicell Cell Culture Insert, 30 mm</td><td>Millipore</td><td>PIHP03050</td><td>Organotypic slice culture inserts</td></tr><tr><td>Osmometer</td><td>Precision Systems</td><td>OSMETTE II</td><td></td></tr><tr><td>PTFE coated spatulas</td><td>Cole-Parmer</td><td>SK-06369-11</td><td></td></tr><tr><td>Scissors</td><td>FST</td><td>14958-09</td><td></td></tr><tr><td>Stereo Microscope</td><td>Olympus</td><td>SZ61</td><td></td></tr><tr><td>Sterile Vacuum Filtration System</td><td>Millipore</td><td>SCGPT01RE</td><td>Filtration and storage of aCSF</td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Electrode preparation&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Capillary Glasses</td><td>Warner Instruments</td><td>640796</td><td></td></tr><tr><td>Micropipetter Puller</td><td>Sutter Instrument</td><td>P-1000</td><td>Puller</td></tr><tr><td>Oven</td><td>Binder</td><td>BD (E2)</td><td></td></tr><tr><td>Puller Filament</td><td>Sutter Instrument</td><td>FB330B</td><td>Puller</td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Single-cell electroporation and fluorescence imaging #1&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>3.5 mm Falcon Petri Dishes</td><td>BD Falcon</td><td>353001</td><td></td></tr><tr><td>Airtable</td><td>TMC</td><td>63-7512E</td><td></td></tr><tr><td>CCD camera</td><td>Q Imaging</td><td>Retiga-2000DC</td><td>Camera</td></tr><tr><td>Electroporation System</td><td>Molecular Devices</td><td>Axoporator 800A</td><td>Electroporator</td></tr><tr><td>Fluorescence Illumination System</td><td>Prior</td><td>Lumen 200</td><td></td></tr><tr><td>Manipulator</td><td>Sutter Instrument</td><td>MPC-385</td><td>Manipulator</td></tr><tr><td>Metamorph software</td><td>Molecular Devices</td><td></td><td>Image acquisition</td></tr><tr><td>Peristaltic Pump</td><td>Rainin</td><td>Dynamax, RP-2</td><td>Perfusion pump</td></tr><tr><td>Shifting Table</td><td>Luigs &amp;amp; Neuman</td><td>240 XY</td><td></td></tr><tr><td>Speaker</td><td>Unknown</td><td></td><td>Speakers connected to the electroporator</td></tr><tr><td>Stereo Microscope</td><td>Olympus</td><td>SZ30</td><td></td></tr><tr><td>Table Top Incubator</td><td>Thermo Scientific</td><td>MIDI 40</td><td></td></tr><tr><td>Upright Microscope</td><td>Olympus</td><td>BX61WI</td><td></td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Fluorescence imaging #2&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>All-in-One Fluorescence Microscope</td><td>Keyence</td><td>BZ-X710</td><td></td></tr></tbody></table>

single-cell electroporation across different organotypic slice culture of mouse hippocampal excitatory and class-specific inhibitory neurons

Electroporation has established itself as a critical method for transferring specific genes into cells to understand their function. Here, we describe a single-cell electroporation technique that maximizes the efficiency (~80%) of in vitro gene transfection in excitatory and class-specific inhibitory neurons in mouse organotypic hippocampal slice culture. Using large glass electrodes, tetrodotoxin-containing artificial cerebrospinal fluid and mild electrical pulses, we delivered a gene of interest into cultured hippocampal CA1 pyramidal neurons and inhibitory interneurons. Moreover, electroporation could be carried out in cultured hippocampal slices up to 21 days in vitro with no reduction in transfection efficiency, allowing for the study of varying slice culture developmental stages. With interest growing in examining the molecular functions of genes across a diverse range of cell types, our method demonstrates a reliable and straightforward approach to in vitro gene transfection in mouse brain tissue that can be performed with existing electrophysiology equipment and techniques.

Our single-cell electroporation is capable of precisely delivering genes into visually identified excitatory and inhibitory neurons. We electroporated three different neuronal cell types at three different time points. Parvalbumin (Pv) or vesicular glutamate type 3 (VGT3) expressing neurons were visualized by crossing Pvcre (JAX #008069) or VGT3cre (JAX #018147) lines with TdTomato (a variant of red fluorescent protein) reporter line (Jax #007905), respectively named Pv/TdTomato and VGT3/TdTomato li...

Watch this Scientific Journal Video about Single-Cell Electroporation across Different Organotypic Slice Culture of Mouse Hippocampal Excitatory and Class-Specific Inhibitory Neurons at JoVE.com

Single-Cell Electroporation across Different Organotypic Slice Culture of Mouse Hippocampal Excitatory and Class-Specific Inhibitory Neurons

Presented here is a protocol for single-cell electroporation that can deliver genes in both excitatory and inhibitory neurons across a range of in vitro hippocampal slice culture ages. Our approach provides precise and efficient expression of genes in individual cells, which can be used to examine cell-autonomous and intercellular functions.

Electroporation has established itself as a critical method for transferring specific genes into cells to understand their function. Here, we describe a ...

single-cell-electroporation-across-different-organotypic-slice

Research

JoVE Journal

Neuroscience

3.6K Views.  University of Massachusetts Medical School. Hier wordt een protocol gepresenteerd voor eencellige elektroporatie dat genen kan leveren in zowel excitatory als remmende neuronen in een reeks in vitro hippocampale slice-cultuurleeftijden. Onze aanpak zorgt voor een nauwkeurige en efficiënte expressie van genen in individuele cellen, die kunnen worden gebruikt om cel-autonome en intercellulaire functies te onderzoeken.

Video: Single-Cell Electroporation across Different Organotypic Slice Culture of Mouse Hippocampal Excitatory and Class-Specific Inhibitory Neurons

Organotypic Slice Cultures for Studies of Postnatal Neurogenesis

Here we describe a technique for studying hippocampal postnatal neurogenesis in the rodent brain using the organotypic slice culture technique. This method maintains the characteristic topographical morphology of the hippocampus while allowing direct application of pharmacological agents to the developing hippocampal dentate gyrus. Additionally, slice cultures can be maintained for up to 4 weeks and thus, allow one to study the maturation process of newborn granule neurons. Slice cultures allow for efficient pharmacological manipulation of hippocampal slices while excluding complex variables such as uncertainties related to the deep anatomic location of the hippocampus as well as the blood brain barrier. For these reasons, we sought to optimize organotypic slice cultures specifically for postnatal neurogenesis research.

Here we describe a technique for studying hippocampal postnatal neurogenesis using the organotypic slice culture technique. This method allows for in vitro manipulation of adult neurogenesis and allows for the direct application of pharmacological agents to the cultured hippocampus.

Here we describe a technique for studying hippocampal postnatal neurogenesis in the rodent brain using the organotypic slice culture technique. This ...

Developmental Biology

Ex Utero Electroporation and Organotypic Slice Culture of Mouse Hippocampal Tissue

Mouse genetics offers a powerful tool determining the role of specific genes during development. Analyzing the resulting phenotypes by immunohistochemical and molecular methods provides information of potential target genes and signaling pathways. To further elucidate specific regulatory mechanisms requires a system allowing the manipulation of only a small number of cells of a specific tissue by either overexpression, ablation or re-introduction of specific genes and follow their fate during development. To achieve this ex utero electroporation of hippocampal structures, especially the dentate gyrus, followed by organotypic slice culture provides such a tool. Using this system to generate mosaic deletions allows determining whether the gene of interest regulates cell-autonomously developmental processes like progenitor cell proliferation or neuronal differentiation. Furthermore it facilitates the rescue of phenotypes by re-introducing the deleted gene or its target genes. In contrast to in utero electroporation the ex utero approach improves the rate of successfully targeting deeper layers of the brain like the dentate gyrus. Overall ex utero electroporation and organotypic slice culture provide a potent tool to study regulatory mechanisms in a semi-native environment mirroring endogenous conditions.

Ex Utero Electroporation and Organotypic Slice Culture of Mouse Hippocampal Tissue

Here we present a protocol providing a tool to examine regulatory mechanisms of specific genes during hippocampal development. Employing ex utero electroporation and organotypic slice culture allows the up- and down-regulation of the expression of genes of interest in single cells and follow their fate during development.

Mouse genetics offers a powerful tool determining the role of specific genes during development. Analyzing the resulting phenotypes by immunohistochemical ...

Organotypic Hippocampal Slice Cultures

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the strength of their connections after brief periods of strong activation. This phenomenon, known as long-term potentiation (LTP) can last for hours or days and has become the best candidate mechanism for learning and memory 2. In addition, the well defined anatomy and connectivity of the hippocampus 3 has made it a classical model system to study synaptic transmission and synaptic plasticity4.
As our understanding of the physiology of hippocampal synapses grew and molecular players became identified, a need to manipulate synaptic proteins became imperative. Organotypic hippocampal cultures offer the possibility for easy gene manipulation and precise pharmacological intervention but maintain synaptic organization that is critical to understanding synapse function in a more naturalistic context than routine culture dissociated neurons methods.
Here we present a method to prepare and culture hippocampal slices that can be easily adapted to other brain regions. This method allows easy access to the slices for genetic manipulation using different approaches like viral infection 5,6 or biolistics 7. In addition, slices can be easily recovered for biochemical assays 8, or transferred to microscopes for imaging 9 or electrophysiological experiments 10.

We describe a method to prepare organotypic hippocampal slices that can be easily adapted to other brain regions. Brain slices are laid on porous membranes and culture media is allowed to form an interface. This method preserves the gross architecture of the hippocampus for up to 2 weeks in culture.

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the ...

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the first born neurons remain closer to the ventricle while the last born neurons migrate past the first born neurons towards the surface of the brain1. In addition to neuronal migration2, a key process for normal cortical function is the regulation of neuronal morphogenesis3. While neuronal morphogenesis can be studied in vitro in primary cultures, there is much to be learned from how these processes are regulated in tissue environments.
We describe techniques to analyze neuronal migration and/or morphogenesis in organotypic slices of the cerebral cortex4,6. A pSilencer modified vector is used which contains both a U6 promoter that drives the double stranded hairpin RNA and a separate expression cassette that encodes GFP protein driven by a CMV promoter7-9. Our approach allows for the rapid assessment of defects in neurite outgrowth upon specific knockdown of candidate genes and has been successfully used in a screen for regulators of neurite outgrowth8. Because only a subset of cells will express the RNAi constructs, the organotypic slices allow for a mosaic analysis of the potential phenotypes. Moreover, because this analysis is done in a near approximation of the in vivo environment, it provides a low cost and rapid alternative to the generation of transgenic or knockout animals for genes of unknown cortical function. Finally, in comparison with in vivo electroporation technology, the success of ex vivo electroporation experiments is not dependant upon proficient surgery skill development and can be performed with a shorter training time and skill.

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

To conduct a rapid assessment of the function of genes in the development of cerebral cortex, we describe methods involving the ex vivo electroporation of plasmids co-expressing inhibitory RNA (RNAi) and GFP in murine embryonic cortex. This protocol is amenable to the study of various aspects of neurodevelopment such as neurogenesis, neuronal migration and neuronal morphogenesis including dendrite and axon outgrowth.

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, their distinct dendritic morphologies, their specific patterns of axonal connectivity, and their selective firing responses. The molecular and cellular mechanisms responsible for these aspects of differentiation during development are still poorly understood.
Here, we describe combined protocols for labeling and characterizing the structural connectivity and excitability of cortical neurons. Modification of the in utero electroporation (IUE) protocol allows the labeling of a sparse population of neurons. This, in turn, enables the identification and tracking of the dendrites and axons of individual neurons, the precise characterization of the laminar location of axonal projections, and morphometric analysis. IUE can also be used to investigate changes in the excitability of wild-type (WT) or genetically modified neurons by combining it with whole-cell recording from acute slices of electroporated brains. These two techniques contribute to a better understanding of the coupling of structural and functional connectivity and of the molecular mechanisms controlling neuronal diversity during development. These developmental processes have important implications on axonal wiring, the functional diversity of neurons, and the biology of cognitive disorders.

In Utero Electroporation Approaches to Study the Excitability of Neuronal Subpopulations and Single-cell Connectivity

This manuscript provides protocols that use in utero electroporation (IUE) to describe the structural connectivity of neurons at the single-cell level and the excitability of fluorescently labeled neurons. Histology is used to characterize dendritic and axonal projections. Whole-cell recording in acute slices is used to investigate excitability.

The nervous system is composed of an enormous range of distinct neuronal types. These neuronal subpopulations are characterized by, among other features, ...

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

GABAergic interneurons (INs) are critical components of neuronal networks that drive cognition and behavior. INs destined to populate the cortex migrate tangentially from their place of origin in the ventral telencephalon (including from the medial and caudal ganglionic eminences (MGE, CGE)) to the dorsal cortical plate in response to a variety of intrinsic and extrinsic cues. Different methodologies have been developed over the years to genetically manipulate specific pathways and investigate how they regulate the dynamic cytoskeletal changes required for proper IN migration. In utero electroporation has been extensively used to study the effect of gene repression or overexpression in specific IN subtypes while assessing the impact on morphology and final position. However, while this approach is readily used to modify radially migrating pyramidal cells, it is more technically challenging when targeting INs. In utero electroporation generates a low yield given the decreased survival rates of pups when electroporation is conducted before e14.5, as is customary when studying MGE-derived INs. In an alternative approach, MGE explants provide easy access to the MGE and facilitate the imaging of genetically modified INs. However, in these explants, INs migrate into an artificial matrix, devoid of endogenous guidance cues and thalamic inputs. This prompted us to optimize a method where INs can migrate in a more naturalistic environment, while circumventing the technical challenges of in utero approaches. In this paper, we describe the combination of ex utero electroporation of embryonic mouse brains followed by organotypic slice cultures to readily track, image and reconstruct genetically modified INs migrating along their natural paths in response to endogenous cues. This approach allows for both the quantification of the dynamic aspects of IN migration with time-lapse confocal imaging, as well as the detailed analysis of various morphological parameters using neuronal reconstructions on fixed immunolabeled tissue.

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

Here, we provide a low-cost and reliable method to generate electroporated brain organotypic slice cultures from mouse embryos suitable for confocal microscopy and live-imaging techniques.

GABAergic interneurons (INs) are critical components of neuronal networks that drive cognition and behavior. INs destined to populate the cortex migrate ...

Photostimulation and Whole-Cell Patch Clamp Recordings of Neurons in Mouse Hippocampal Slices

Source: Richevaux, L., et al. <a href="https://app.jove.com/v/59534">In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices.</a> J. Vis. Exp. (2019)
The video demonstrates a protocol to study synaptic interactions and neuronal responses in a transfected mouse hippocampal brain slice. It involves illuminating long-range sensory neurons to trigger action potentials and recording the resulting excitatory post-synaptic potentials in the recorded neuron using the whole-cell patch-clamp technique.

All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Solutions for acute slice ...

Single-Cell Electroporation across Different Organotypic Slice Culture of Mouse Hippocampal Excitatory and Class-Specific Inhibitory Neurons

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