This work was supported by NIH grants EY019312 and EY021126 (VJK), EY002687 to the Department of Ophthalmology and Visual Sciences at Washington University, and by Research to Prevent Blindness.

All experimental protocols were in accordance with the Guide for the Care and Use of Laboratory Animals and were approved by the institutional Animal Studies Committee at Washington University.
1. Setting Up Perfusion and Specimen Holder
<ol>
	<li>Prepare solution for retina perfusion fresh on the day of the experiment. Use distilled and deionized water. Use one of the following three solutions.
	<ol>
		<li>Prepare Bicarbonate-containing Ames&#8217; solution (1 L): 1 bottle of Ames&#8217; media and 1.9 g of NaHCO3,</li>
		<li>Prepare Locke&#8217;s solution (in mM): 112.5 NaCl, 3.6 KCl, 2.4 MgCl2, 1.2 CaCl2, 10.0 HEPES, 20.0 NaHCO3, 3.0 Na succinate, 0.5 Na glutamate, 0.02 EDTA, and 10.0 glucose, 0.1% MEM vitamins and amino acids</li>
		<li>Prepare HEPES-buffered Ringer solution (in mM): 133.3 NaCl, 3.3 KCl, 2.0 MgCl2, 1.0 CaCl2, 10.0 glucose, 0.01 EDTA, 12.0 HEPES, pH adjusted to 7.5 with ~5.8 ml of 1 M NaOH, add 0.72 g/L Leibovitz culture medium L-15. Use 20 - 50 &#956;M DL-AP4 and 50 &#8211; 100 &#956;M BaCl2 to isolate the photoreceptor response with any perfusion media.</li>
	</ol>
	</li>
	<li>Prepare solution for electrodes28 (in mM): 140.0 NaCl, 3.6 KCl, 2.4 MgCl2, 1.2 CaCl2, 3 HEPES, 0.01 EDTA and adjust pH to 7.4 &#8211; 7.5 with NaOH. Electrode solution can be stored at RT for several months.</li>
	<li>Prepare and test the specimen holder.
	<ol>
		<li>Glue black/grey filter paper on top of the domes of the specimen holder&#8217;s bottom part (see Figure 1A). Spread two-component 5 min epoxy glue carefully around the edges of the flat tops of the domes. If necessary, do it under dissection microscope.</li>
		<li>Wait until the glue is almost dry (about 4 min) and press the filter paper on the domes by using a flat item. Glue filter paper at least one day before the experiment. The filter paper can be used multiple times but should be replaced after a month of recordings. Use 70% ethanol to clean domes carefully from any glue residue before installing the replacement paper.</li>
		<li>Fill the electrode channels with electrode solution trying to avoid any air bubbles and screw pellet electrodes enclosed with a threaded adapter into the electrode channels (see Figures 1A and B). Connect the top and bottom pieces of the specimen holder with four screws and fill the perfusion lines with electrode solution.</li>
		<li>Measure the resistance and voltage between the leads of each electrode pair using a multimeter (Figure 1B). Resistance should be below 100 &#937;k and the voltage below 10 mV if the channels are bubble-free and the electrodes are in good conditions.</li>
	</ol>
	</li>
	<li>Pour 400 - 700 ml of perfusion media in the glass bottle. Separate another 300 ml to be used in the dissection of the retinas and store it in a fridge. Set the perfusion tubing in the heat exchanger block and place the preheated block on the heating plate (see Figure 1D).</li>
	<li>Enclose the bottle with a cap that has connections to CO2/O2 gas (if Ames&#8217; or Locke&#8217;s is used) and for perfusion tubing (see Figure 1D). Preheat the bottle with the media to 37 oC and place it on the heating plate or on the top of the light stimulation unit in water bath set to 37 &#8211; 39 oC17.
	<ol>
		<li>Prime the perfusion lines by filling them with perfusion media to initiate the gravity-driven flow.
		<ol>
			<li>In LKC system, place the bottle on the top of the stimulator unit17 to provide a significant gravitational flow that is then adjusted by flow-rate regulators without being affected by the lowering level of the perfusion solution in the bottle during the experiment.</li>
			<li>In Ocuscience system, place the perfusion bottle inside the Faraday cage in order to minimize noise and place long perfusion output lines from the specimen holder (see step 2.8) well below the level of the specimen holder (and perfusion bottle) to increase gravitational drive of the solution. Shield these output lines and connect the shield to the amplifier&#8217;s ground to prevent coupling of the electromagnetic noise to the ERG signal.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Adjust the perfusion to 3 &#8211; 5 ml/min by using flow-rate regulators. Connect 5% CO2 / 95% O2 gas from a cylinder with proper regulator and adjust the flow rate to ensure steady bubbling of the media in the bottle through an air stone.</li>
</ol>
2. Sample Preparation
<ol>
	<li>Assemble clean and sharp dissection instruments including straight-bladed microscissors, one or two 45o tweezers, razor blade and a rectangular piece of filter paper.</li>
	<li>Pour about 200 ml of cold perfusion solution in a large Petri dish so that the whole bottom part of the specimen holder (including the domes) can be immersed into the solution. This step becomes important when mounting the retinas on the domes (see step 2.6). Although some solutions are designed to be saturated with carbogen (5% CO2 / 95% O2), this is not essential for the dissection purposes and was not done in the experiments described here.</li>
	<li>For a typical experiment, keep animals in 12/12 hrs dark/light cycle and dark-adapt them for 6 &#8211; 12 hrs before the recordings. Euthanize the animal by CO2 inhalation followed by cervical dislocation under dim red light and do all the following procedures under dim red or IR light (use e.g., red filter in front of the microscope light source).
	<ol>
		<li>Pull the eyes out by using tweezers and put them in the media. Place one eye at a time on a small piece of paper (e.g., some regular filter paper) and make a slit approximately on the level of ora serrata while holding the eye with tweezers (this is done outside of the solution here).</li>
	</ol>
	</li>
	<li>Cut along the ora serrata (or closer to the equator of the eye) with microscissors and remove the cornea and the lens. Place the eye cup in the cold media in the large Petri dish and repeat the same procedure with the other eye.</li>
	<li>Cut a small incision from the top of the eye cup towards the optic nerve by keeping the scissors between the retina and sclera in order to keep the retina as intact as possible. Grip the sclera from both sides of the incision by using two tweezers and pull the tweezers away from each other to detach the retina.
	<ol>
		<li>Cut the optic nerve and try to isolate the retina with a minimum amount of physical contact to the distal surface. RPE will mostly detach automatically from the retina during the dissection process. It is more important to avoid mechanical disturbance of the retina than to perform the dissection quickly, generally retinas can be incubated at least 30 min in the solution without significant effects on the response properties.</li>
	</ol>
	</li>
	<li>Mount the retinas on the domes of the specimen holder (Figure 1C). Immerse the bottom part of the specimen holder in the Petri dish with the dissected retinas. Slide the retina, photoreceptor side (the convex surface of the isolated retina) upwards, above the dome and lift the specimen holder so that the retina attaches on the filter paper. Repeat the procedure for the other retina.</li>
	<li>Dry the holder plate carefully to prevent electrical crosstalk between the retinas as well as noise and signal shunting. Specimen holder has O-rings around the bottom part domes to prevent solution spill between the bottom and top parts of the holder as well as to help electrical isolation of the photoreceptor and ganglion cell sides of the individual retinas. Attach the top part of the holder with the four screws (see Figure 1B) and fill the perfusion channels with perfusion solution by using a syringe and a needle.</li>
	<li>Transfer the specimen holder next to the heat exchanger block and connect the input and output perfusion lines to the specimen holder (Figure 1D). Connect the electrodes to the ERG amplifier (top electrodes connect to the ground/minus pin in the amplifier) and attach the stimulator/control unit on the specimen holder by using an adaptor or slide the heating pad into the stimulator unit depending on which in vivo ERG system is used.</li>
</ol>
3. Recordings
<ol>
	<li>Configure the amplifier and stimulus settings by using the in vivo system&#8217;s software. Set the acquisition frequency to a value between 1 and 10 kHz and low-pass filtering to 300 Hz. Do not use high-pass filtering. Use 60 Hz (or 50 Hz in Europe) notch filter if necessary.</li>
	<li>Record baseline without light stimulation and with dim stimulation (e.g., green light of -35 dB or 0.3 mCd sm-2) to test for good electrical connection between the electrodes and retina sample. Wait 10 &#8211; 20 min before starting the data collection so that retinal temperature and function reach a stable state.
	<ol>
		<li>Choose stimulus parameters according to any specific experiment and start data collection. E.g., use green light from -40 up to 0 dB or from 0.1 up to 1,000 mCd sm-2 for a scotopic response family. Use about 2 to 3 log-units brighter flashes in photopic recordings where rods have to be suppressed by background light (3 &#8211; 30 Cdm-2) or a probe flash (about 1.3 Cd sm-2 = 1 dB, see Figure 3).</li>
		<li>However, remember that the exact light intensity values might be somewhat dependent on your system calibration and experimental conditions. As a rule of thumb, scale intensities down by 5 &#8211; 10-fold as compared to in vivo recordings when using green light17. A black cover with apertures above the retinas (included in the commercial adapter system) can be used to reduce light scattering in the specimen holder and facilitate homogenous and equal stimulation of both retinas by the Ganzfeld sphere of the commercial in vivo systems. 
		NOTE: For recordings from dark-adapted retina, the intensities of test flashes used in a typical experiment bleach only a negligible fraction of the pigment so that the lack of RPE-driven regeneration is not an issue. In addition, it has been previously shown that cone pigment regeneration can still take place in the isolated retina via the M&#252;ller cell visual cycle20.</li>
	</ol>
	</li>
	<li>As experiments last typically from 30 min up to several hours, monitor baseline drift, noise level and response stability during the experiment as changes in these might indicate technical problems. 
	&#8203;NOTE: Increased noise or decreased response amplitudes may indicate air bubbles in the perfusion or electrode channels, too low or high temperature, solution leak/spill or displacement of the retina.</li>
</ol>
4. Cleaning
<ol>
	<li>Detach the specimen holder from the perfusion lines, open it and flush the retinas from the filter paper. Remove the electrodes and rinse them with distilled water (do not use ethanol). Clean the specimen holder (including the perfusion channels) with ethanol and/or distilled water. Flush perfusion tubing carefully with &#62;70% ethanol.</li>
	<li>Rinse the perfusion bottle with distilled water (do not use detergents). Ethanol can be also used to clean the bottle.</li>
</ol>

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Neurosci. 19, 1203-1216 (1999).</li><li>Heikkinen, H., Vinberg, F., Nymark, S., Koskelainen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mesopic+background+lights+enhance+dark-adapted+cone+ERG+flash+responses+in+the+intact+mouse+retina:+a+possible+role+for+gap+junctional+decoupling.">Mesopic background lights enhance dark-adapted cone ERG flash responses in the intact mouse retina: a possible role for gap junctional decoupling.</a> J. Neurophysiol. 105, 2309-2318 (2011).</li><li>Gouras, P., MacKay, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+in+amplitude+of+the+human+cone+electroretinogram+with+light+adaptation.">Growth in amplitude of the human cone electroretinogram with light adaptation.</a> Invest. Ophthalmol. Vis. Sci. 30, 625-630 (1989).</li><li>Peachey, N. S., Goto, Y., al-Ubaidi, M. R., Naash, M. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Properties+of+the+mouse+cone-mediated+electroretinogram+during+light+adaptation.">Properties of the mouse cone-mediated electroretinogram during light adaptation.</a> Neurosci. Lett. 162, 9-11 (1993).</li></ol>

Washington University in St. Louis has a license agreement with Xenotec, Inc. and may receive a royalty from the sale of the ex vivo adapter.

We demonstrate here the critical steps for obtaining high-quality ex vivo ERG recordings simultaneously from two isolated mouse retinas by using in vivo ERG system components together with an ex vivo ERG adapter. In this study we perfused both retinas from the animal with the same solution (either Ames&#8217;, Locke&#8217;s or Ringer) but it is also possible to perfuse each retina with a different solution e.g., for drug testing purposes. The most important steps for obtaining high qua...

Electroretinogram (ERG) is a well-established technique that can be used to record the electrical activity of the retina triggered by light. The ERG signal is generated mainly by voltage changes caused by radial currents (along the axis of photoreceptors and bipolar cells) flowing in the resistive extracellular space of the retina. The first ERG signal was recorded in 1865 by Holmgren from the surface of a fish eye1. Einthoven and Jolly 19082 divided the ERG response to the onset of light into three different waves, called a-, b-, and c-waves, that are now known to reflect mainly the activity of photoreceptors, ON bipolar cells, and pigment epithelium cells, respectively3-8. ERG can be recorded from the eyes of anesthetized animals or humans (in vivo), from isolated eye preparation9, across isolated intact retina (ex vivo)3,10-15 or across specific retina layers with microelectrodes (local ERG)4,16. Of these, in vivo ERG is currently the most widely used method to assess retinal function. It is a noninvasive technique that can be used for diagnostic purposes or to follow the progression of retinal diseases in animals or patients. However, in vivo ERG recordings produce a complicated signal with several overlapping components, often contaminated by extraocular physiological noise (e.g., breathing and cardiac activity).
Local ERG can be used to record the signal across specific layers of the retina but it is the most invasive and has the lowest signal-to-noise ratio (SNR) as compared to the other ERG recording configurations. Local ERG is also technically demanding and requires expensive equipment (e.g., microscope and micromanipulators). Transretinal ERG from the intact, isolated retina (ex vivo ERG) offers a compromise between in vivo and local ERG methods allowing stable and high SNR recordings from intact retinas of animals or humans17. Recently, this method has been used successfully to study rod and cone photoreceptor function in mammalian, primate and human retinas18-20. In addition, due to absence of pigment epithelium in the ex vivo retina, the positive c-wave component of the ERG signal is removed and a prominent negative slow PIII component is revealed in the ex vivo recordings. The slow PIII component has been shown to originate from the activity of M&#252;ller glia cells in the retina21-23. Thus, ex vivo ERG method could also be used to study M&#252;ller cells in the intact retina. Several studies have also shown that ex vivo ERG recordings could be used to measure concentration of pharmacological agents around the retina24 and test the safety and efficacy of drugs25-27.
Multiple commercial in vivo systems are available and used in many laboratories that do not necessarily have extensive electrophysiology background. In contrast, ex vivo devices have not been available until recently17 and as a result only very few laboratories are currently taking advantage of this powerful technique. It would be beneficial to make ex vivo ERG recordings available to more laboratories in order to advance our knowledge about retinal physiology and pathology, and to develop new therapies for blinding diseases. We demonstrate here a simple and affordable ex vivo ERG device17 and show how it can be used in combination with several commercially available in vivo ERG systems to record rod- and cone-mediated signaling (a- and b-waves) and the function of M&#252;ller cells (slow PIII) from intact wild-type mouse retinas.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>In vivo ERG system</td>
			<td>OcuScience</td>
			<td>HMsERG</td>
			<td><a href="http://www.ocuscience.us/id77.html">www.ocuscience.us/id77.html</a></td>
		</tr>
		<tr>
			<td>In vivo ERG system</td>
			<td>LKC Technologies</td>
			<td>UTAS-E 3000</td>
			<td><a href="http://www.lkc.com/products/UTAS/bigshot.html">www.lkc.com/products/UTAS/bigshot.html</a></td>
		</tr>
		<tr>
			<td>Ex vivo adapter</td>
			<td>OcuScience</td>
			<td>Ex VIVO ERG adapter</td>
			<td><a href="http://www.ocuscience.us/id107.html">www.ocuscience.us/id107.html</a></td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td>North Central Instruments</td>
			<td>Leica M80</td>
			<td><a href="http://www.ncimicro.com/store/ListProduct.asp?bSub=T&amp;idCategory=27">May use any brand</a></td>
		</tr>
		<tr>
			<td>IR emitter</td>
			<td>Opto Diode Corp.</td>
			<td>OD-50L</td>
			<td><a href="http://www.optodiode.com/">www.optodiode.com</a></td>
		</tr>
		<tr>
			<td>Prowler Night Vision Scopes</td>
			<td>B.E. Meyers Electro Optics</td>
			<td>D4300-I</td>
			<td>Military grade product.</td>
		</tr>
		<tr>
			<td>Red filter</td>
			<td>Rosco Laboratories</td>
			<td>Roscolux #27 Medium Red</td>
			<td>May be used instead of IR system</td>
		</tr>
		<tr>
			<td>Red head light</td>
			<td>OcuScience</td>
			<td>ERGX011</td>
			<td><a href="http://www.ocuscience.us/catalog/i29.html">www.ocuscience.us/catalog/i29.html</a></td>
		</tr>
		<tr>
			<td>Microscissors</td>
			<td>WPI, Inc.</td>
			<td>500086</td>
			<td><a href="http://www.wpiinc.com/">www.wpiinc.com/</a></td>
		</tr>
		<tr>
			<td>Dumont tweezers #5</td>
			<td>WPI, Inc.</td>
			<td>14101</td>
			<td></td>
		</tr>
		<tr>
			<td>Razor blades</td>
			<td>Electron Microscopy Sciences</td>
			<td>72000</td>
			<td><a href="http://www.emsdiasum.com/">www.emsdiasum.com</a></td>
		</tr>
		<tr>
			<td>Scale</td>
			<td>Metler Toledo</td>
			<td>AB54-S/FACT</td>
			<td><a href="http://www.sigmaaldrich.com/">May use any brand</a></td>
		</tr>
		<tr>
			<td>pH meter and electrode</td>
			<td>Beckman Coulter</td>
			<td>pHI 350</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td><a href="http://www.sigmaaldrich.com/">May use any brand</a></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma-Aldrich</td>
			<td>60129</td>
			<td><a href="http://www.sigmaaldrich.com/">May use any brand</a></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma-Aldrich</td>
			<td>63020</td>
			<td>1.0 M solution</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma-Aldrich</td>
			<td>21114</td>
			<td>1.0 M solution</td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>431788</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>Sodium Bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S6297</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>Ames medium</td>
			<td>Sigma-Aldrich</td>
			<td>A1420</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>BaCl2</td>
			<td>Sigma-Aldrich</td>
			<td>B0750</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>DL-AP4</td>
			<td>Tocris Bioscience</td>
			<td>101</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>Succinic acid disodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>224731</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>L-Glutamic acid</td>
			<td>Sigma-Aldrich</td>
			<td>G2834</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>D-(+)-Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G7528</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>Leibovitz culture medium L-15</td>
			<td>Sigma-Aldrich</td>
			<td>L4386</td>
			<td>May use any brand</td>
		</tr>
		<tr>
			<td>MEM vitamins</td>
			<td>Sigma-Aldrich</td>
			<td>M6895</td>
			<td></td>
		</tr>
		<tr>
			<td>MEM amino acids</td>
			<td>Sigma-Aldrich</td>
			<td>M5550</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbogen</td>
			<td>Airgas</td>
			<td>UN3156</td>
			<td>5% CO2</td>
		</tr>
	</tbody>
</table>

simultaneous ex vivo functional testing of two retinas by in vivo electroretinogram system

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

We recorded flash responses from dark-adapted wild-type (WT) C57BL/6 mouse retinas by following the experimental protocols described above and illustrated in Figure 1 by using different standard perfusion solutions (Figure 2). The response waveforms and kinetics as well as sensitivity of rod photoreceptors appeared similar in Ames&#8217; and Locke&#8217;s media (Figure 2A and B). On the other hand, under HEPES-buffered Ringer solution (no bicarbonate or ...

Watch this Scientific Journal Video about Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System at JoVE.com

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

simultaneous-ex-vivo-functional-testing-two-retinas-vivo

Research

JoVE Journal

Neuroscience

9.0K Views.  Washington University in St. Louis. Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

Video: Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are tightly regulated, affecting the final dynamics of protein abundance. Various regulatory mechanisms exist at the translation step, rendering mRNA levels alone an unreliable indicator of gene expression. In addition, local regulation of mRNA translation has been particularly implicated in neuronal functions, shifting &#39;translatomics&#39; to the focus of attention in neurobiology. The presented method can be used to bridge transcriptomics and proteomics.
Here we describe essential modifications to the technique of polyribosome fractionation, which interrogates the translatome based on the association of actively translated mRNAs to multiple ribosomes and their differential sedimentation in sucrose gradients. Traditionally, working with in vivo samples, particularly of the central nervous system (CNS), has proven challenging due to the restricted amounts of material and the presence of fatty tissue components. In order to address this, the described protocol is specifically optimized for use with minimal amount of CNS material, as demonstrated by the use of single mouse spinal cord and brain. Briefly, CNS tissues are extracted and translating ribosomes are immobilized on mRNAs with cycloheximide. Myelin flotation is then performed to remove lipid rich components. Fractionation is performed on a sucrose gradient where mRNAs are separated according to their ribosomal loading. Isolated fractions are suitable for a range of downstream assays, including new genome wide assay technologies.

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

This protocol illustrates essential modifications of polyribosome fractionation in order to study the translatome of in vivo CNS samples. It allows global assessment of translation and transcription regulation through the isolation and comparison of total RNA to ribosome bound RNA fractions.

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

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 ...

Ex Vivo Pressurized Hippocampal Capillary-Parenchymal Arteriole Preparation for Functional Study

From subtle behavioral alterations to late-stage dementia, vascular cognitive impairment typically develops following cerebral ischemia. Stroke and cardiac arrest are remarkably sexually dimorphic diseases, and both induce cerebral ischemia. However, progress in understanding the vascular cognitive impairment, and then developing sex-specific treatments, has been partly limited by challenges in investigating the brain microcirculation from mouse models in functional studies. Here, we present an approach to examine the capillary-to-arteriole signaling in an ex vivo hippocampal capillary-parenchymal arteriole (HiCaPA) preparation from mouse brain. We describe how to isolate, cannulate, and pressurize the microcirculation to measure arteriolar diameter in response to capillary stimulation. We show which appropriate functional controls can be used to validate the HiCaPA preparation integrity and display typical results, including testing potassium as a neurovascular coupling agent and the effect of the recently characterized inhibitor of the Kir2 inward rectifying potassium channel family, ML133. Further, we compare the responses in preparations obtained from male and female mice. While these data reflect functional investigations, our approach can also be used in molecular biology, immunochemistry, and electrophysiology studies.

The present manuscript details how to isolate hippocampal arterioles and capillaries from the mouse brain and how to pressurize them for pressure myography, immunofluorescence, biochemistry, and molecular studies.

From subtle behavioral alterations to late-stage dementia, vascular cognitive impairment typically develops following cerebral ischemia. Stroke and ...

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent and cause visual impairment and blindness. Understanding how such diseases progress is critical to formulating new treatments. In vivo&#160;microscopy in animal models of disease is a powerful tool for understanding neurodegeneration and has led to important progress towards treatments of conditions ranging from Alzheimer&#39;s disease to stroke. Given that the retina is the only central nervous system structure inherently accessible by optical approaches, it naturally lends itself towards in vivo imaging. However, the native optics of the lens and cornea present some challenges for effective imaging access.
This protocol outlines methods for in vivo two-photon imaging of cellular cohorts and structures in the mouse retina at cellular resolution, applicable for both acute- and chronic-duration imaging experiments. It presents examples of retinal ganglion cell (RGC), amacrine cell, microglial, and vascular imaging using a suite of labeling techniques including adeno-associated virus (AAV) vectors, transgenic mice, and inorganic dyes. Importantly, these techniques extend to all cell types of the retina, and suggested methods for accessing other cellular populations of interest are described. Also detailed are example strategies for manual image postprocessing for display and quantification. These techniques are directly applicable to studies of retinal function in health and disease.

In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent ...

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in retinal diseases, and testing therapeutic interventions. The ex vivo electroretinogram (ERG) allows the quantification of contributions from individual cell types in the isolated retina by addition of specific pharmacological agents and evaluation of tissue-intrinsic changes independently of systemic influences. Retinal light responses can be measured using a specialized ex vivo ERG specimen holder and recording setup, modified from existing patch clamp or microelectrode array equipment. Particularly, the study of ON-bipolar cells, but also of photoreceptors, has been hampered by the slow but progressive decline of light responses in the ex vivo ERG over time. Increased perfusion speed and adjustment of the perfusate temperature improve ex vivo retinal function and maximize response amplitude and stability. The ex vivo ERG uniquely allows the study of individual retinal neuronal cell types. In addition, improvements to maximize response amplitudes and stability allow the investigation of light responses in retina samples from large animals, as well as human donor eyes, making the ex vivo ERG a valuable addition to the repertoire of techniques used to investigate retinal function.

Optimizing the Setup and Conditions for Ex Vivo Electroretinogram to Study Retina Function in Small and Large Eyes

Modification of existing multielectrode array or patch clamp equipment makes the ex vivo electroretinogram more widely accessible. Improved methods to record and maintain ex vivo light responses facilitate the study of photoreceptor and ON-bipolar cell function in the healthy retina, animal models of eye diseases, and human donor retinas.

Measurements of retinal neuronal light responses are critical to investigating the physiology of the healthy retina, determining pathological changes in ...

Ex Vivo Release of Calcitonin Gene-Related Peptide from the Trigeminovascular System in Rodents

Calcitonin gene-related peptide (CGRP) was first discovered in the 1980s as a splice variant from the calcitonin gene. Since its discovery, its role in migraine pathophysiology has been well established, first by its potent vasodilator properties and subsequently by its presence and function as a neurotransmitter in the sensory trigeminovascular system. The migraine-provoking ability of CGRP gave support to the pharma industry to develop monoclonal antibodies and antagonists inhibiting the effect of CGRP. A new treatment paradigm has proven effective in the prophylactic treatment of migraine. One of the useful tools to further understand migraine mechanisms is the ex vivo model of CGRP release from the trigeminovascular system. It is a relatively simple method that can be used with various pharmacological tools to achieve know-how to further develop new effective migraine treatments. The present protocol describes a CGRP release model and the technique to quantify the effect of pharmacological agents on the amount of CGRP released from the trigeminovascular system in rodents. A procedure describing the experimental approach from euthanasia to the measurement of protein levels is provided. The essential isolation of the trigeminal ganglion and the trigeminal nucleus caudalis from both mice and rats and the preparation of rat dura mater are described in detail. Furthermore, representative results from both species (rats and mice) are presented. The technique is a key tool to investigate the molecular mechanisms involved in migraine pathophysiology by using various pharmacological compounds and genetically modified animals.

Ex Vivo Release of Calcitonin Gene-Related Peptide from the Trigeminovascular System in Rodents

The present protocol describes the ex vivo calcitonin gene-related peptide (CGRP) release model and the strategy to quantify the effect of pharmacological agents on the amount of CGRP released from the trigeminovascular system in rodents.

Calcitonin gene-related peptide (CGRP) was first discovered in the 1980s as a splice variant from the calcitonin gene. Since its discovery, its role in ...