This is supported by 973 MOST grant (Grant No. 2011CB504402, 2012CB947602), National Natural Science Foundation of China (Grant No. 91132724, U1332136) and the 'Hundred Talents Project' of Chinese Academy of Science. The protocol was approved by the Committee on the Ethics of Animal Experiments of the USTC (Permit Number: USTCACUC1103013).

1. Construct the Surgery Apparatus
NOTE: To ensure the fish remains alive during and after the operation, drip anesthesia solution ethyl 3-aminobenzoate methanesulfonate (MS-222, or Tricaine) at a half concentration (0.015%) with a velocity of 1 drop/sec&#160;through the fish&#39;s mouth using a homebuilt drip system shown in Figure 1.
<ol>
	<li>Make a box as shown in Figure 1A (length is 28 cm, width&#160;is 10 cm, height&#160;is 5 cm), put a sponge (length is 6 cm, width&#160;is 9 cm, height&#160;is 4 cm) in the middle and open a gap on it.</li>
	<li>Make a smooth globular surface on the end of the needle with light curing bond (Figure 1B) which will then be inserted into the fish mouth. Ensure that the surface of the needle is smooth to prevent the fish&#39;s mouth from being damaged.</li>
	<li>Add the half concentration of MS-222 (left chamber, length is 6 cm, width&#160;is 10 cm, height is 8 cm) and saline (right chamber, length is 4 cm, width&#160;is 10 cm, height is 8 cm) to the reservoirs as shown in Figure 1C. Note: Fresh MS-222 is recommended.</li>
	<li>Anesthetize fish in 0.03% MS-222 for 2 min.</li>
	<li>Place the fish in the gap of the sponge and fix the fish with two pairs of pins, inserted behind the pectoral fin and beside the cloacal pore, respectively11.</li>
	<li>Connect the fish&#8217;s mouth, MS-222, and rearing water with a T type tube as shown in Figure 1C.</li>
	<li>Switch on the tube connected to the MS-222.</li>
</ol>
2. Retrograde Label RGCs from the Right Tectum
NOTE: Prior to beginning, disinfect all apparatuses with 70-75% ethanol, ensuring that no ethanol remains.
<ol>
	<li>Rossing: slightly remove the skin over the whole skull with sclerectome (dorsal view of the skull is shown in Figures 2A and&#160;2B).</li>
	<li>Clearing: wash the skull with saline and then dry it with ear washing bulb.</li>
	<li>Initial corrosion: corrode the skull with the etchant for 10 sec.</li>
	<li>Clearing: completely wipe away the etchant with a cotton swab and then wash the area with saline and dry it.</li>
	<li>Protection: in order to protect the&#160;skull in&#160;other areas except the right tectum from excess corrosion, cover these areas with light curing bond and then cure it by exposure to blue light with portable illuminant.</li>
	<li>Complete corrosion: put the etchant in the central area of the right tectum again for complete etching for 2 min. Note: Since skulls of older fish are calcified, corrosion will take longer.</li>
	<li>Clearing: completely wipe away the etchant with a cotton swab and then wash the area with saline and dry it again.</li>
	<li>Skull opening: carefully remove the malacic skull (Figure 2C) with forceps to expose the right tectum (Figure 2D).</li>
	<li>Clearing: wipe the mucus and blood effusing from the hole with pieces of gelfoam and wash it with saline until the bleeding has stopped.</li>
	<li>Dye placement: put the dye (tissue-labeling paste) on the whole right tectum and cover it with gelfoam.</li>
	<li>Segregating: cover the hole with a sterile scale from the same animal.</li>
	<li>Sealing: place light curing bond onto the scale and then cure it with blue light again for 40 sec.</li>
	<li>Revival: wash the surface of the bond and revive the animal with saline.</li>
	<li>Nursing and breeding: Keep the fish in 28.5 &#176;C embryo medium (ZFIN) for 5 days and feed 2 times/day. Do not include fish whose dye fails to stay in the tectum for 5 days in the experiment. Note: Embryo medium (EM) solution is vital for fish survival and temperature is the key factor for DiI labeling.</li>
</ol>
3. Wholemounting Retina and Imaging
<ol>
	<li>Place the fish in a dark field for 2 hr&#160;before retina wholemount.</li>
	<li>Anesthetize the fish with ice water.</li>
	<li>Puncture a hole in the margin of cornea and make an eye cup by removing the cornea with venus scissors.</li>
	<li>Remove the lens and then flush the center of retina with ice-cold 1x&#160;PBS.</li>
	<li>Flush the gap between retina and sclera with ice-cold 1x&#160;PBS until no pigment is left.</li>
	<li>Cut the optic nerve at the disc.</li>
	<li>Keep the RGC layer upward and hold the retina with a self-made glass spoon (Figures 3A and&#160;3B). A dropper can also be used to transfer the retina.</li>
	<li>Put the retina in a drop of icy 30% glycerin on a slide with the RGC layer upward (Figure 3C).</li>
	<li>Remove the glycerin and then flatten the retina on the glass slide before cutting retina into four quadrants (Figure 3D). Keeping the retina unfolded with RGC layer upward will be helpful for flattening the retina.</li>
	<li>Drip icy 50% glycerin on the retina to wash away the fragments of pigment.</li>
	<li>Remove the glycerin and then drop 20 &#181;l of icy 75% glycerin on the retina.</li>
	<li>Cover the sample with a 1.8 cm x&#160;1.8 cm square coverslip.</li>
	<li>Seal the coverslip with nail polish. Immediate imaging of the retina is recommended.</li>
	<li>Non-interlace image the whole retina under a 20X&#160;object lens (NA=0.70) with DP72 CCD (each image is 1360 x&#160;1024 pixels). Each field has three different focuses.</li>
	<li>Extend the depth of each field with the tool of &#8220;extended depth of field&#8221; in the software.</li>
	<li>Assemble a virtual retina with the function of photomerge.</li>
	<li>Select three fields of 680 x&#160;512 pixels (actual&#160;area is 350 &#181;m by 264 &#181;m, 0.092 mm2) in each orientation of the retina to analyze average density of RGCs (Figure 4A).</li>
	<li>Obtain the area with the function of measurements &#8211; create a polygon feature in the image software.</li>
	<li>Calculate the total number of RGCs by the equation &#8220;RGCs number = destiny x&#160;area&#8221;&#160;9.</li>
</ol>

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A., Noack, C., Boehringer, D., Biermann, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Staining+of+fluorogold-prelabeled+retinal+ganglion+cells+with+calcein-AM:+A+new+method+for+assessing+cell+vitality.">Staining of fluorogold-prelabeled retinal ganglion cells with calcein-AM: A new method for assessing cell vitality.</a> J Neurosci Methods. 192, 233-239 (2010).</li><li>Schwalb, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+factors+secreted+by+the+goldfish+optic+nerve+induce+retinal+ganglion+cells+to+regenerate+axons+in+culture.">Two factors secreted by the goldfish optic nerve induce retinal ganglion cells to regenerate axons in culture.</a> J Neurosci. 15, 5514-5525 (1995).</li><li>Watanabe, M., Inukai, N., Fukuda, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival+of+retinal+ganglion+cells+after+transection+of+the+optic+nerve+in+adult+cats:+a+quantitative+study+within+two+weeks.">Survival of retinal ganglion cells after transection of the optic nerve in adult cats: a quantitative study within two weeks.</a> Vis Neurosci. 18, 137-145 (2001).</li><li>Chiu, K., Lau, W. M., Yeung, S. C., Chang, R. C., So, K. 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Q., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+the+optokinetic+response+to+study+visual+function+of+zebrafish.">Using the optokinetic response to study visual function of zebrafish.</a> J Vis Exp. , (2010).</li><li>Tokuoka, H., Yoshida, T., Matsuda, N., Mishina, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+by+glycogen+synthase+kinase-3beta+of+the+arborization+field+and+maturation+of+retinotectal+projection+in+zebrafish.">Regulation by glycogen synthase kinase-3beta of the arborization field and maturation of retinotectal projection in zebrafish.</a> J Neurosci. 22, 10324-10332 (2002).</li><li>Xiao, T., Roeser, T., Staub, W., Baier, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+GFP-based+genetic+screen+reveals+mutations+that+disrupt+the+architecture+of+the+zebrafish+retinotectal+projection.">A GFP-based genetic screen reveals mutations that disrupt the architecture of the zebrafish retinotectal projection.</a> Development. 132, 2955-2967 (2005).</li><li>Li, L., Dowling, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+the+olfactoretinal+centrifugal+pathway+may+relate+to+the+visual+system+defect+in+night+blindness+b+mutant+zebrafish.">Disruption of the olfactoretinal centrifugal pathway may relate to the visual system defect in night blindness b mutant zebrafish.</a> Journal of Neuroscience. 20, 1883-1892 (2000).</li><li>Kassing, V., Engelmann, J., Kurtz, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+of+Single-Cell+Responses+in+the+Optic+Tectum+of+Adult+Zebrafish+with+Dextran-Coupled+Calcium+Dyes+Delivered+via+Local+Electroporation.">Monitoring of Single-Cell Responses in the Optic Tectum of Adult Zebrafish with Dextran-Coupled Calcium Dyes Delivered via Local Electroporation.</a> PLoS One. 8, (2013).</li></ol>

We have nothing to disclose.

Retrograde labeling of RGCs is important to research RGCs survival in mammalian animals, but it has not been used in zebrafish. The alternative methods, HE staining7 and antibody staining8, are not gold standards for counting RGCs number, and transgenic&#160;lines with all RGCs labeled has not yet been constructed12,13. In this video, we introduce a method to retrograde label RGCs in the retina of adult zebrafish. Though about 1% of axons project to olfactory bulb or...

As adult zebrafish have a strong ability to regenerate axons after optic nerve injury1, a proper method to count the whole RGCs is essential to evaluate RGCs survival and regeneration2. Based on the methods of retrograde labeling RGCs in mammalian and goldfish3-5, we constructed the method to label RGCs from the tectum in adult zebrafish. For adult zebrafish, two critical technical concerns should be noted: the skull of adult zebrafish is very small6; they live in a water environment. Here, we treat the skull with etchant which minimizes the hazards associated with drilling5. Then, we seal the hole with light curing bond which improves animal survival after surgery.
Previously, several other techniques were adopted to count RGCs number in indirect ways. HE staining in the retina sections labels all types of cells in the RGCL7. Antibody labeling in the whole retina, such as islet-1, can also label amacrine cells8. Though retrograde label from the optic nerve stump can label all RGCs in the retina, it cannot be adopted in the crush model because it causes extra injury to the optic nerve. Taking advantage of retrograde label from the tectum, we have researched RGCs survival and regeneration in optic nerve crush. Results show that almost all RGCs survived and over 90% of RGCs regenerated to the tectum at the first week in the crush model9.
In order to successfully label all RGCs, DiI paste was chosen after comparison with several other commercial dyes10. Firstly, it is especially designed for in vivo tissue labeling. Secondly, it is a lipophilic dye which cannot diffuse in water. Additionally, this fluorescence can persist for a long time which makes it an excellent candidate for RGCs survival research.

<table border="0" cellpadding="0" cellspacing="0" width="608">
	<tbody>
		<tr height="37">
			<td height="37" style="height:37px;width:152px;">MS222</td>
			<td style="width:152px;">Sigma Aldrich</td>
			<td style="width:152px;">E10521</td>
			<td style="width:152px;">USA</td>
		</tr>
		<tr height="37">
			<td height="37" style="height:37px;width:152px;">DiI</td>
			<td style="width:152px;">Invitrogen</td>
			<td style="width:152px;">N22880</td>
			<td style="width:152px;">USA</td>
		</tr>
		<tr height="37">
			<td height="37" style="height:37px;width:152px;">lightcuring bond</td>
			<td style="width:152px;">Heraeus Kulzer&#160;</td>
			<td style="width:152px;">Durafill bond</td>
			<td style="width:152px;">Germany</td>
		</tr>
		<tr height="55">
			<td height="55" style="height:55px;width:152px;">Gluma Etch</td>
			<td style="width:152px;">Heraeus Kulzer&#160;</td>
			<td style="width:152px;">Gluma Etch 35 Gel</td>
			<td style="width:152px;">Germany</td>
		</tr>
		<tr height="73">
			<td height="73" style="height:73px;width:152px;">Blue LED</td>
			<td style="width:152px;">Shenruo Medical Equipment Co.</td>
			<td style="width:152px;">Power Blue Light Curing Unit</td>
			<td style="width:152px;">China</td>
		</tr>
	</tbody>
</table>

retrograde labeling of retinal ganglion cells in adult zebrafish with fluorescent dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

As Figures 4B-D show, the number of DiI+ cells is two thirds of DAPI+ cells in RGCL. In a normal retina, a montage image of a whole retina (Figure 4E) shows that DiI+ RGCs are distributed over the entire retina but in a regenerated retina (Figure 4F), as RGCs in the central area have not regenerated to their target at the first week, they could not be labeled.
<img alt="Fig...

Watch this Scientific Journal Video about Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes at JoVE.com

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

retrograde-labeling-retinal-ganglion-cells-adult-zebrafish-with

Research

JoVE Journal

Neuroscience

9.4K ビュー.  University of Science and Technology of China. We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

ビデオ: Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

Labeling and Imaging Cells in the Zebrafish Hindbrain

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish embryos, injection of plasmid DNA results in mosaic expression, allowing for the visualization of single cells or small clusters of cells 1 . We describe how injection of plasmid DNA encoding membrane-targeted Green Fluorescent Protein (mGFP) under the control of a ubiquitous promoter can be used for imaging cells undergoing neurulation. Central to this protocol is the methodology for imaging labeled cells at high resolution in sections and also in real time. This protocol entails the injection of mGFP DNA into young zebrafish embryos. Embryos are then processed for vibratome sectioning, antibody labeling and imaging with a confocal microscope. Alternatively, live embryos expressing mGFP can be imaged using time-lapse confocal microscopy. We have previously used this straightforward approach to analyze the cellular behaviors that drive neural tube formation in the hindbrain region of zebrafish embryos 2. The fixed preparations allowed for unprecedented visualization of cell shapes and organization in the neural tube while live imaging complemented this approach enabling a better understanding of the cellular dynamics that take place during neurulation.

Key to understanding the morphogenetic processes that shape the early embryo is the ability to image cells at high resolution. We describe here a technique for labeling single cells or small clusters of cells in whole zebrafish embryos with membrane-targeted Green Fluorescent Protein.

Key to understanding the morphogenetic processes that shape the early vertebrate embryo is the ability to image cells at high resolution. In zebrafish ...

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1. This information is then segregated into what are known as the ON and OFF pathways. The photoreceptors signal light information to the bipolar cells (BCs), which depolarize in response to increases (On BCs) or decreases (Off BCs) in light intensity. This segregation of light information is maintained at the level of the retinal ganglion cells (RGCs), which have dendrites stratifying in either the Off sublamina of the inner plexiform layer (IPL), where they receive direct excitatory input from Off BCs, or stratifying in the On sublamina of the IPL, where they receive direct excitatory input from On BCs. This segregation of information regarding increases or decreases in illumination (the On and Off pathways) is conserved and signaled to the brain in parallel.
The RGCs are the output cells of the retina, and are thus an important cell to study in order to understand how light information is signaled to visual nuclei in the brain. Advances in mouse genetics over recent decades have resulted in a variety of fluorescent reporter mouse lines where specific RGC populations are labeled with a fluorescent protein to allow for identification of RGC subtypes 2 3 4 and specific targeting for electrophysiological recording. Here, we present a method for recording light responses from fluorescently labeled ganglion cells in an intact, isolated retinal preparation. This isolated retinal preparation allows for recordings from RGCs where the dendritic arbor is intact and the inputs across the entire RGC dendritic arbor are preserved. This method is applicable across a variety of ganglion cell subtypes and is amenable to a wide variety of single-cell physiological techniques.

This article provides a description of how to dissect and record from the isolated retinal preparation in mouse. In particular, we describe how to record light responses from a fluorescently labeled ganglion cell population and subsequently identify and analyze its morphology.

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1.  This information is then ...

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

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 ...

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

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

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

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

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

Retrograde Loading of Nerves, Tracts, and Spinal Roots with Fluorescent Dyes

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. Generally, the dyes are applied as solid crystals or by local pressure injection using glass pipettes. However, this can result in dilution of the dye and reduced labeling intensity, particularly when several hours are required for dye diffusion. Here we demonstrate a simple and low-cost technique for introducing fluorescent and ion-sensitive dyes into neurons using a polyethylene suction pipette filled with the dye solution. This method offers a reliable way for maintaining a high concentration of the dye in contact with axons throughout the loading procedure.

We describe a simple and low cost technique for introducing high concentration of fluorescent and calcium-sensitive dyes into neurons or any neuronal tract using a polyethylene suction pipette.

Retrograde labeling of neurons is a standard anatomical method1,2 that has also been used to load calcium and voltage-sensitive dyes into neurons3-6. ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

A Novel Light Damage Paradigm for Use in Retinal Regeneration Studies in Adult Zebrafish

Light-induced retinal degeneration (LIRD) is commonly used in both rodents and zebrafish to damage rod and cone photoreceptors. In adult zebrafish, photoreceptor degeneration triggers M&#252;ller glial cells to re-enter the cell cycle and produce transient-amplifying progenitors. These progenitors continue to proliferate as they migrate to the damaged area, where they ultimately give rise to new photoreceptors. Currently, there are two widely-used LIRD paradigms, each of which results in varying degrees of photoreceptor loss and corresponding differences in the regeneration response. As more genetic and pharmacological tools are available to test the role of individual genes of interest during regeneration, there is a need to develop a robust LIRD paradigm. Here we describe a LIRD protocol that results in widespread and consistent loss of both rod and cone photoreceptors in which we have combined the use of two previously established LIRD techniques. Furthermore, this protocol can be extended for use in pigmented animals, which eliminates the need to maintain transgenic lines of interest on the albino background for LIRD studies.

Multiple light damage protocols have been described to damage photoreceptors and consequently induce a retinal regeneration response in adult zebrafish. This protocol describes an improved method that can be used in pigmented animals and that damages the vast majority of rod and cone photoreceptors across the entire retina.

Light-induced retinal degeneration (LIRD) is commonly used in both rodents and zebrafish to damage rod and cone photoreceptors. In adult zebrafish, ...

Single-cell RNA-Seq of Defined Subsets of Retinal Ganglion Cells

The discovery of cell type-specific markers can provide insight into cellular function and the origins of cellular heterogeneity. With a recent push for the improved understanding of neuronal diversity, it is important to identify genes whose expression defines various subpopulations of cells. The retina serves as an excellent model for the study of central nervous system diversity, as it is composed of multiple major cell types. The study of each major class of cells has yielded genetic markers that facilitate the identification of these populations. However, multiple subtypes of cells exist within each of these major retinal cell classes, and few of these subtypes have known genetic markers, although many have been characterized by morphology or function. A knowledge of genetic markers for individual retinal subtypes would allow for the study and mapping of brain targets related to specific visual functions and may also lend insight into the gene networks that maintain cellular diversity. Current avenues used to identify the genetic markers of subtypes possess drawbacks, such as the classification of cell types following sequencing. This presents a challenge for data analysis and requires rigorous validation methods to ensure that clusters contain cells of the same function. We propose a technique for identifying the morphology and functionality of a cell prior to isolation and sequencing, which will allow for the easier identification of subtype-specific markers. This technique may be extended to non-neuronal cell types, as well as to rare populations of cells with minor variations. This protocol yields excellent-quality data, as many of the libraries have provided read depths greater than 20 million reads for single cells. This methodology overcomes many of the hurdles presented by Single-cell RNA-Seq and may be suitable for researchers aiming to profile cell types in a straightforward and highly efficient manner.

Here, we present a combinatorial approach for classifying neuronal cell types prior to isolation and for the subsequent characterization of single-cell transcriptomes. This protocol optimizes the preparation of samples for successful RNA Sequencing (RNA-Seq) and describes a methodology designed specifically for the enhanced understanding of cellular diversity.

The discovery of cell type-specific markers can provide insight into cellular function and the origins of cellular heterogeneity. With a recent push for ...