This work was supported by the Deutsche Forschungsgemeinschaft (WE849/16 1/2 to K.D. and R.W.). We are grateful to Thomas Euler (T&ouml;bingen, Germany) for the light stimulation software QDS.

The following description assumes that the experimenter has a basic understanding of retinal structure, patch-clamp recording, and two-photon microscopy. For useful information on establishing and running a patch-clamp setup and two-photon imaging system see Refs [7-12].
1. Animal and tissue preparation
<ol>
	<li>Keep mouse dark-adapted for at least 3 h. In the meantime, prepare 1-2 l of extracellular solution consisting of (in mM) 125 NaCl, 2.5 KCl, 1 CaCl2, 1.6 MgCl2, 25 NaHCO3, 10 D-glucose and equilibrate it to pH 7.4 by gassing with carbogen (5 % CO2 in O2) at room temperature.</li>
	<li>Euthanize the mouse in an air-tight chamber by CO2 overdose followed by cervical dislocation. This procedure and the following steps should be performed in darkness. Use dim long-wavelength illumination (we block short wavelengths from a cold light source by a 690 nm-longpass filter) to support personal vision while keeping the animal's eyes dark-adapted (mice only have poor vision in the red part of the light spectrum). For work in complete darkness use infrared illumination (&gt;800 nm) and wear night-vision goggles.</li>
	<li>Enucleate eyes with a pair of curved iris scissors and transfer them to a dish of extracellular solution placed under a dissecting microscope.</li>
	<li>Remove the cornea and ciliary body by opening the eye bulb along the ora serrata (the border between retina and ciliary body) with a pair of spring scissors (we first use a lancet to pierce a starting point for the cutting). Take out the lens and carefully separate the retina from the pigment epithelium. If the orientation of the retina is crucial to your study, indicate it like in Ref [12]. Cut the optical nerve between retina and pigment epithelium and remove the retina from the eyecup. Note the direction of the retinal surfaces: the inside of the curled-up retina is the ganglion cell side; the outside is the photoreceptor side.</li>
	<li>Remove the vitreous from the inner retinal surface by gently pulling it off with the aid of a wooden toothpick. For this purpose, use a small volume of extracellular solution that just covers the retina. The vitreous sticks to the toothpick and can be dragged centrifugally off the retina. Then, apply short incisions along the retinal perimeter to facilitate flattening of the tissue.</li>
	<li>Transfer the retina into the recording chamber photoreceptor-side down, spread it out on the glass bottom (we use a fine brush) and immobilize it with a nylon-strung frame of stainless steel. Prepare the second retina in the same way and keep it dark-adapted in carboxygenated extracellular solution for later use.</li>
</ol>
2. Recordings
<ol>
	<li>Install the recording chamber in darkness under an upright laser scanning microscope and superfuse the retinal preparation continuously (not less than 5 ml/min) with carboxygenated extracellular solution heated to 35 &deg;C. The microscope (also equipped with an infrared-sensitive CCD camera) is situated on a shock-absorbing air table inside a Faraday cage for electronic shielding. Cover the cage with a non-transparent curtain to keep the preparation in darkness. We also screen off light from computer monitors by red transparent film.</li>
	<li>Tune the infrared laser to 850-870 nm or longer wavelengths, switch to mode-locked condition, and use two-photon excitation to visualize GFP-expressing cells. Attenuate the laser output using neutral density filters controlled by the laser scanning software down to a degree that is just sufficient for clear recognition of the fluorescent cell</li>
	<li>For patch-clamp recordings in current-clamp mode use glass micropipettes (we use borosilicate glass tubing of 1.5 mm outer diameter and 0.225 mm wall thickness) filled with intracellular solution consisting of (in mM) 125 K-gluconate, 10 KCl, 0.5 EGTA, 10 HEPES, titrated to pH 7.4 with KOH (giving a pipette resistance of about 5 M&Omega;). Note that other experimental conditions like voltage-clamp recording require a different solution. If dye injections are desired, add a fluorescent probe (we use 10 mM Alexa Fluor 594) or a tracer molecule (3 % Neurobiotin). Insert the micropipette into the holder and make sure that the reference electrode (chlorinated silver wire) is in contact with the extracellular solution in the recording chamber.</li>
	<li>Apply pressure to the micropipette and target a GFP-expressing cell (we use a 40-fold water immersion objective; NA 1.25). Amacrine cell bodies are situated in the ganglion cell layer (directly beneath the surface in the current orientation of the retina) as well as in the proximal part of the inner nuclear layer (about 55-75 &mu;m deep within the preparation). Before micropipette contact with the targeted cell, the inner limiting membrane (a sheath made of glial endfeet) at the retinal surface has to be penetrated. Successful penetration is recognized by intracellular solution that is driven out from the micropipette tip and detaches the inner limiting membrane from the underlying retinal tissue as can be observed on an infrared transmission image captured by the IR-CCD camera.</li>
	<li>Approach the desired cell by comparing the soma position from the two-photon image with the picture of the IR-CCD camera showing the position of the recording micropipette. Note that this step is not easily achieved and requires some practice, because the GFP-expressing cell has to be recognized in the infrared transmission image in order to direct the micropipette towards it. Correct targeting is achieved when the micropipette tip causes dimpling of the cell surface that can be seen in the two-photon image. Alternative to this procedure, use a fluorescent dye (e.g. Alexa Fluor 594, 100 &mu;M) in the intracellular solution to reveal the micropipette position in the two-photon image. Two-photon infrared excitation allows for sufficient fluorescence of this dye to make the micropipette discernible. For this purpose, adjust the detection range in the laser scanning software to cover also red fluorescence.</li>
	<li>Release pressure from the micropipette and obtain a whole-cell patch-clamp configuration in current-clamp mode. A usable recording should give a membrane potential of -50 to -55 mV and last for at least 20 min.</li>
	<li>Present visual stimuli created by a stimulation software (we use QDS by Thomas Euler, University of T&uuml;bingen, Germany)11 on a computer monitor. Adjust the spatial position of the stimulation to be centered on the recorded cell: Mark the cell position on the monitor showing the transmission image of the IR-CCD camera and center a spot-like stimulus onto it. Tune the stimulus intensity by inserting neutral density filters into the beam path. For calibration of monitor spectrum and intensity see Ref [14]. Choose a prolonged interstimulus interval (e.g. 15 s) to avoid adaptation. Include a photodiode within the beam path to keep record of the stimulus timing.</li>
	<li>Start the stimulation protocol and record the light responses (we use a sampling rate of 10 kHz with filtering at 5 kHz for non-spiking cells; for spike recording a higher sampling rate like 20 kHz is recommended). Use the light stimulation software to trigger the recording software.</li>
	<li>For dye filling let the fluorescent agent diffuse into the recorded cell for 30-45 min before carefully retracting the micropipette from the cell. Take care not to pull out the cell body. If the non-fluorescent tracer Neurobiotin was used, it needs visualization by binding to streptavidin conjugated to a fluorophore (we use streptavidin-Cy3, 1:400 over night at 4 &deg;C after 20 min paraformaldehyde fixation of the retina; for dye-coupling studies streptavidin incubation can be prolonged to 3 d). Alternatively, injections can also be performed with sharp electrodes (&gt;100 M&Omega;) when the dye is driven iontophoretically into the impaled cell (rectangular pulses: 0.25-1 nA, 100 ms, interval 100 ms, total time 3-6 min).</li>
</ol>
3. Representative Results:
The following results originate from a study on a mouse expressing green fluorescent protein (GFP) under the promoter for tyrosine hydroxylase13,14, the enzyme catalyzing the rate limiting step in catecholamine synthesis (TH::GFP mouse). Based on the brightness of the GFP-signal two distinct cell populations are distinguishable (Figure 2A). Cells expressing the higher GFP level possess cell bodies located in the inner nuclear layer (INL, Figure 2A) or displaced in the ganglion cell layer (GCL, Figure 2B) and stratify in the middle of the inner plexiform layer (IPL, Figure 2C). They were identified as type 2 cells14-16 and could systematically be studied with respect to morphology (Figure 3) and electrical activity (Figure 4) even though its population density only amounts to 250 cells/mm2.
<img alt="Figure 1" src="/files/ftp_upload/3457/3457fig1.jpg" /> 
Figure 1. Schematic representation of the experimental setup. Two-photon excitation (red dotted light path) of fluorophore-expressing cells in the intact retina enables visual targeting by a micropipette (green emission light path). The retina is subjected to optical stimuli projected through the condenser of the microscope (yellow light path), and cellular light responses are recorded.
<img alt="Figure 2" src="/files/ftp_upload/3457/3457fig2.jpg" /> 
Figure 2. GFP-expressing cells in a retinal flatmount of a TH::GFP mouse. Two populations can be distinguished by the brightness of the GFP-signal: type 1 cells (DA cells) located in the INL showing weak fluorescence (A, see arrowheads) and intensely labeled type 2 cells with cell bodies either in the INL (A, see arrows) or displaced in the GCL (B) and a dendritic stratification in stratum S3 of the IPL (C). Scale bars, 50 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/3457/3457fig3.jpg" /> 
Figure 3.Morphology of a type 2 cell injected with the tracer Neurobiotin. The tracer was subsequently visualized by streptavidin-Cy3 binding (magenta). The micrograph, which is also depicting the GFP signal (green), is a projection of image stacks covering the GCL and IPL. Scale bar, 50 &mu;m.
<img alt="Figure 4" src="/files/ftp_upload/3457/3457fig4.jpg" /> 
Figure 4. Light responses of a type 2 cell located in the GCL. Response pattern to white light full-field illumination of increasing intensity in the scotopic range. Stimulus intensity is given in photoisomerizations per rod per second (Rh*/rod/s). A prolonged stimulus of 3 s was used for better distinction of the response components at stimulus onset (ON response) and offset (OFF response).

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+recording+of+light-evoked+calcium+signals+in+the+functionally+intact+retina.">Optical recording of light-evoked calcium signals in the functionally intact retina.</a> Proc. Natl. Acad. Sci. U.S.A. 96, 7035-7040 (1999).</li></ol>

Mice were handled and euthanized in accordance with institutional guidelines for animal welfare and the laws on animal experiments issued by the German government.

This method offers the possibility to study electrical properties of specific neurons in the intact retina under visual guidance without influencing the adaptational condition of the retina. It is particularly suited for the characterization of cells that are at present rather poorly studied due to low population density like most populations of amacrine cells. Two-photon excitation permits high-resolution and high-contrast imaging even from deeper parts of the tissue17, a prerequisite for accurate targeting and successful patch-clamping of cells particularly in the INL with its high density of cell bodies. 
The isolated mouse retina is viable for 3-4 h under the experimental conditions. If the second retina is stored in complete darkness under continuous carbogen gassing, it retains the purple color of unbleached photopigment and can be used after finishing experiments with the first retina. In the beginning, targeting the selected cell with the micropipette in the retinal wholemount is a bit challenging and requires some practice, especially when working in darkness. Including a fluorescent dye in the micropipette can simplify the procedure, because cell and micropipette are visible under two-photon excitation at the same time. However, adding components to the intracellular solution may impede gigaseal formation or cause the recording quality to suffer. Once successfully achieved, a good recording can last for about 1 h.
Whereas the infrared excitation light itself is only poorly absorbed by retinal photoreceptors, excited GFP-expressing cells emit light in the visible part of the spectrum. However, fluorophores are excited only in a small focal volume that is not likely to change the adaptational condition of the retina. All the more, excitation is only needed for targeting and can be switched off during recording of light responses.
The usefulness of this approach is already demonstrated by studies on specific populations of amacrine cells14,18 from which electrophysiological recordings otherwise would have been possible only by accident12. Finally, this powerful technique can further be extended by including a pharmacological approach14, Ca2+ imaging19 or by using injected cells for immunocytochemical studies or electron microscopy. That way, the functional position of a given cell type in the retinal circuitry can be unraveled.

<table border="1">
<tbody>
<tr>
<th>Reagent/Equip-ment</th>
<th>Company</th>
<th>Catalogue number</th>
<th>Comments</th>
</tr>
<tr>
<td>Night-vision goggles</td>
<td>Gutzeit GmbH, Warthausen, Germany</td>
<td>Xtron F1</td>
<td>includes a 800 nm light source</td>
</tr>
<tr>
<td>Optical filter</td>
<td>Schott, Mainz, Germany</td>
<td>RG9</td>
<td>longpass filter with cutoff at 690 nm</td>
</tr>
<tr>
<td>Iris scissors</td>
<td>Fine Science Tools</td>
<td>14061-09</td>
<td>curved with 22 mm blade</td>
</tr>
<tr>
<td>Spring scissors</td>
<td>Fine Science Tools</td>
<td>15000-00</td>
<td>straight with 3 mm blade</td>
</tr>
<tr>
<td>Recording chamber</td>
<td>Luigs &amp; Neumann GmbH, Ratingen, Germany</td>
<td>200-100 500 0180 type A(TC)</td>
<td>Teflon chamber with bottom-mounted glass cover slip (approx. 0.15 mm; No. 200-100 500 0182)</td>
</tr>
<tr>
<td>Flow heater</td>
<td>Multichannel Systems, Reutlingen, Germany</td>
<td>PH01, TC01</td>
<td>heatable perfusion cannula with temperature sensor and temperature controller </td>
</tr>
<tr>
<td>Laser scanning microscope</td>
<td>Leica Microsystems</td>
<td>DM LFS</td>
<td>controlled by Leica Confocal Software</td>
</tr>
<tr>
<td>Air table</td>
<td>Newport</td>
<td>VH 3036W-OPT</td>
<td></td>
</tr>
<tr>
<td>Laser</td>
<td>Spectra-Physics</td>
<td></td>
<td>Tsunami Mode-locked Ti:sapphire laser</td>
</tr>
<tr>
<td>CCD camera</td>
<td>pco AG, Kelheim, Germany</td>
<td>PixelFly QE</td>
<td>including control software </td>
</tr>
<tr>
<td>Micropipette glass capillaries</td>
<td>Hilgenberg, Malsfeld, Germany</td>
<td>1408411</td>
<td></td>
</tr>
<tr>
<td>Micropipette puller</td>
<td>Sutter Instrument</td>
<td>P-97</td>
<td></td>
</tr>
<tr>
<td>Alexa Fluor 594</td>
<td>Invitrogen</td>
<td>A-20004</td>
<td>fluorescent dye</td>
</tr>
<tr>
<td>Neurobiotin</td>
<td>Axxora</td>
<td>VC-SP-1120-M050</td>
<td>non-fluorescent tracer</td>
</tr>
<tr>
<td>Streptavidin-Cy3</td>
<td>Dianova</td>
<td>016-160-084</td>
<td></td>
</tr>
<tr>
<td>Micromanipulator</td>
<td>Luigs &amp; Neumann GmbH, Ratingen, Germany</td>
<td>210-100 000 0010</td>
<td>motorized Mini25 manipulator unit with display SM-5</td>
</tr>
<tr>
<td>Patch-clamp amplifier</td>
<td>npi electronic GmbH, Tamm, Germany</td>
<td>SEC-05LX npi</td>
<td></td>
</tr>
<tr>
<td>Digitizer</td>
<td>National Instruments</td>
<td>BNC-2090</td>
<td></td>
</tr>
<tr>
<td>Data acquisition software WinWCP</td>
<td>John Dempster, University of Scotland, Glasgow, UK</td>
<td></td>
<td>http://spider.science.strath.
ac.uk/sipbs/ software.htm
</td>
</tr>
<tr>
<td>Visual stimulus generator QDS</td>
<td>Thomas Euler, University of T&uuml;bingen, Germany</td>
<td></td>
<td>has to be operated on a separate computer controlling 2 monitors (user interface, stimulus monitor)</td>
</tr>
<tr>
<td>Neutral density filters</td>
<td>ITOS, Mainz, Germany</td>
<td></td>
<td></td>
</tr>
</tbody>
</table>

electrophysiological characterization of gfp-expressing cell populations in the intact retina

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack conspicuous morphological features or show a low population density. This applies particularly to retinal amacrine cells, an exceptionally multiform class of interneurons that comprise roughly 30 subtypes in mammals1. Though being a crucial part of the visual processing by shaping the retinal output2, most of these subtypes have not been studied up to now in a functional context because encountering these cells with a recording electrode is a rare event.
Recently, a multitude of transgenic mouse lines is available that express fluorescent markers like green fluorescent protein (GFP) under the control of promoters for membrane receptors or enzymes that are specific to only a subset of neurons in a given tissue3,4. These pre-labeled cells are therefore accessible to directed microelectrode targeting under microscopic control, permitting the systematic study of their physiological properties in situ. However, excitation of fluorescent markers is accompanied by the risk of phototoxicity for the living tissue. In the retina, this approach is additionally hampered by the problem that excitation light causes appropriate stimulation of the photoreceptors, thus inflicting photopigment bleaching and transferring the retinal circuits into a light-adapted condition. These drawbacks are overcome by using infrared excitation delivered by a mode-locked laser in short pulses of the femtosecond range. Two-photon excitation provides energy sufficient for fluorophore excitation and at the same time restricts the excitation to a small tissue volume minimizing the hazards of photodamage5. Also, it leaves the retina responsive to visual stimuli since infrared light (&gt;850 nm) is only poorly absorbed by photopigments6.
In this article we demonstrate the use of a transgenic mouse retina to attain electrophysiological in situ recordings from GFP-expressing cells that are visually targeted by two-photon excitation. The retina is prepared and maintained in darkness and can be subjected to optical stimuli which are projected through the condenser of the microscope (Figure 1). Patch-clamp recording of light responses can be combined with dye filling to reveal the morphology and to check for gap junction-mediated dye coupling to neighboring cells, so that the target cell can by studied on different experimental levels.

Watch this Scientific Journal Video about Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina at JoVE.com

Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Studying the physiological properties and synaptic connections of specific neurons in the intact tissue is a challenge for those cells that lack ...

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13.3K Views.  University of Oldenburg. This article depicts the recording of individual cells from fluorescently tagged neuronal populations in the intact mouse retina. By using two-photon infrared excitation transgenetically labeled cells were targeted for patch-clamp recording to study their light responses, receptive field properties, and morphology.

Video: Electrophysiological Characterization of GFP-Expressing Cell Populations in the Intact Retina

Organotypic Slice Culture of GFP-expressing Mouse Embryos for Real-time Imaging of Peripheral Nerve Outgrowth

For many purposes, the cultivation of mouse embryos ex vivo as organotypic slices is desirable. For example, we employ a transgenic mouse line (tauGFP) in which the enhanced version of the green fluorescent protein (EGFP) is exclusively expressed in all neurons of the developing central and peripheral nervous system1, allowing the possibility to both film the innervation of the forelimb and to manipulate this process with pharmacological and genetic techniques2. The most critical parameter in the successful cultivation of such slice cultures is the method by which the slices are prepared. After extensive testing of a variety of methods, we have found that a vibratome is the best possible device to slice the embryos such that they routinely result in a culture that demonstrates viability over a period of several days, and most importantly, develops in an age-specific manner. For mid-gestation embryos, this includes the normal outgrowth of spinal nerves from the spinal cord and the dorsal root ganglia to their targets in the periphery and the proper determination of skeletal and muscle tissue. 
In this work, we present a method for processing whole embryos of embryonic day (E) E10 to E12 into 300 - 400 micrometer slices for cultivation in a standard tissue culture incubator, which can be studied for up to two days after slice preparation. Critical for the success of this approach is the use of a vibratome to slice each agarose-embedded embryo. This is followed by the cultivation of the slices upon Millicell culture membrane inserts placed upon a small volume of medium, resulting in an interface culture technique. One litter with an average of 7 embryos routinely produces at least 14 slices (2-3 slices of the forelimb region per embryo), which varies slightly due to the age of the embryos as well as to the thickness of the slices. About 80% of the cultured slices show nerve outgrowth, which can be measured througout the culturing period2. Representative results using the tauGFP mouse line are demonstrated.

We present a method to prepare organotypic slices of mid-gestation mouse embryos for the cultivation and time-lapse imaging of peripheral nerve outgrowth.

For many purposes, the cultivation of mouse embryos ex vivo as organotypic slices is desirable. For example, we employ a transgenic mouse line (tauGFP) in ...

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 ...

Intact Histological Characterization of Brain-implanted Microdevices and Surrounding Tissue

Research into the design and utilization of brain-implanted microdevices, such as microelectrode arrays, aims to produce clinically relevant devices that interface chronically with surrounding brain tissue. Tissue surrounding these implants is thought to react to the presence of the devices over time, which includes the formation of an insulating "glial scar" around the devices. However, histological analysis of these tissue changes is typically performed after explanting the device, in a process that can disrupt the morphology of the tissue of interest.
Here we demonstrate a protocol in which cortical-implanted devices are collected intact in surrounding rodent brain tissue. We describe how, once perfused with fixative, brains are removed and sliced in such a way as to avoid explanting devices. We outline fluorescent antibody labeling and optical clearing methods useful for producing an informative, yet thick tissue section. Finally, we demonstrate the mounting and imaging of these tissue sections in order to investigate the biological interface around brain-implanted devices.

Here we present a histological method for capturing, labeling, optically clearing, and imaging the intact brain tissue interface around chronically implanted microdevices in rodent brain tissue. Results from the techniques comprising this method are useful for understanding the impact of various penetrating brain-implants on their surrounding tissue.

Research  into the design and utilization of brain-implanted microdevices, such as  microelectrode arrays, aims to produce clinically relevant devices ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

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 ...

Isolation of Distinct Cell Populations from the Developing Cerebellum by Microdissection

Microdissection is a novel technique that can isolate specific regions of a tissue and eliminate contamination from cellular sources in adjacent areas. This method was first utilized in the study of Nestin-expressing progenitors (NEPs), a newly identified population of cells in the cerebellar external germinal layer (EGL). Using microdissection in combination with fluorescent-activated cell sorting (FACS), a pure population of NEPs was collected separately from conventional granule neuron precursors in the EGL and from other contaminating Nestin-expressing cells in the cerebellum. Without microdissection, functional analyses of NEPs would not have been possible with the current methods available, such as Percoll gradient centrifugation and laser capture microdissection. This technique can also be applied for use with various tissues that contain either recognizable regions or fluorescently-labeled cells. Most importantly, a major advantage of this microdissection technique is that isolated cells are living and can be cultured for further experimentation, which is currently not possible with other described methods.

Nestin-expressing&#160;progenitors are a newly identified population of neuronal progenitors in the developing cerebellum. Using the microdissection technique presented here in combination with fluorescent-activated cell sorting, this cell population can be purified with no contamination from other cerebellar regions and can be cultured for further studies.

Microdissection is a novel technique that can isolate specific regions of a tissue and eliminate contamination from cellular sources in adjacent areas. ...

In-depth Physiological Analysis of Defined Cell Populations in Acute Tissue Slices of the Mouse Vomeronasal Organ

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory neurons (VSNs) express a specific type of G protein-coupled receptor (GPCR) from at least three different chemoreceptor gene families allowing sensitive and specific detection of chemosensory cues. These families comprise the V1r and V2r gene families as well as the formyl peptide receptor (FPR)-related sequence (Fpr-rs) family of putative chemoreceptor genes. In order to understand the physiology of vomeronasal receptor-ligand interactions and downstream signaling, it is essential to identify the biophysical properties inherent to each specific class of VSNs.
The physiological approach described here allows identification and in-depth analysis of a defined population of sensory neurons using a transgenic mouse line (Fpr-rs3-i-Venus). The use of this protocol, however, is not restricted to this specific line and thus can easily be extended to other genetically modified lines or wild type animals.

Here, we describe a physiological approach that allows identification and in-depth analysis of a defined population of sensory neurons in acute coronal tissue slices of the mouse vomeronasal organ using whole-cell patch-clamp recordings.

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory ...

Preparation of Horizontal Slices of Adult Mouse Retina for Electrophysiological Studies

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in these preparations is perpendicular to the retinal surface, making it ideal for the study of radially oriented neurons like photoreceptors and bipolar cells. However, the large dendritic arbors of horizontal cells, wide-field amacrine cells, and ganglion cells are mostly truncated, leaving markedly reduced synaptic activity in these cells. Whereas ganglion cells and displaced amacrine cells can be studied in a whole-mounted preparation of the retina, horizontal cells and amacrine cells located in the inner nuclear layer are only poorly accessible for electrodes in whole retina tissue.
To achieve maximum accessibility and synaptic integrity, we developed a horizontal slice preparation of the mouse retina, and studied signal transmission at the synapse between photoreceptors and horizontal cells. Horizontal sectioning allows&#160;(1) easy and unambiguous visual identification of horizontal cell bodies for electrode targeting, and (2) preservation of the extended horizontal cell dendritic fields, as a prerequisite for intact and functional cone synaptic input to horizontal cell dendrites.
Horizontal cells from horizontal slices exhibited tonic synaptic activity in the dark, and they responded to brief flashes of light with a reduction of inward current and diminished synaptic activity. Immunocytochemical evidence indicates that almost all cones within the dendritic field of a horizontal cell establish synapses with its peripheral dendrites. The horizontal slice preparation is therefore well suited to study the physiological properties of horizontally extended retinal neurons as well as sensory signal transmission and integration across selected synapses.

We developed a horizontal slice preparation of the adult mammalian retina with the plane of sectioning parallel to the retinal surface. Dendritic fields of nonradially oriented retinal neurons were not truncated, allowing the study of signal processing with patch-clamp and imaging techniques.

Vertical slice preparations are well established to study circuitry and signal transmission in the adult mammalian retina. The plane of sectioning in ...

In Vitro Characterization of the Electrophysiological Properties of Colonic Afferent Fibers in Rats

Dysfunction of the colonic sensory nerves has been implicated in the pathophysiology of several common conditions, including functional and inflammatory bowel diseases and diabetes. Here, we describe a protocol for the in vitro characterization of the electrophysiological properties of colonic afferents in rats. The colorectum, with the intact pelvic ganglion (PG) attached, is removed from the rat; superfused with carbogenated Krebs solution in the recording chamber; and cannulated at the oral and anal ends to allow for distension. A fine nerve bundle emanating from the PG is identified, and the multiunit afferent nerve activity is recorded using a suction electrode. Distension of the colonic segment elicits gradual increases in multiunit discharge. A principal component analysis is conducted to differentiate the low-threshold, the high-threshold, and the wide-dynamic range afferent fibers. Chemical sensitivity of colonic afferents can be studied through the bath or intraluminal administration of test compounds. This protocol can be modified for application to other species, such as mice and guinea pigs, and to study the differences in the electrophysiological properties of thoracolumbar/hypogastric and lumbosacral/pelvic afferents of the descending colon in normal and pathological conditions.

In Vitro Characterization of the Electrophysiological Properties of Colonic Afferent Fibers in Rats

Abnormal sensory function underlies visceral pain and other symptoms of functional and inflammatory bowel diseases. A protocol for the electrophysiological recording of the colonic afferent nerves in an ex vivo rat colorectum preparation is presented here.

Dysfunction of the colonic sensory nerves has been implicated in the pathophysiology of several common conditions, including functional and inflammatory ...