The pCAG-gapEGFP plasmid was a gift from Dr. S. McConnell (Stanford, CA). pCAG-tdTomato plasmid was a gift from Dr. M. Feller (Berkeley, CA). We thank Dr. Edward Ruthazer for suggesting the use of a two-plasmid strategy for single cell labeling and Anne Schohl (Montreal, QC) for validating the two-plasmid Cre/loxP strategy in pilot studies and Crair lab members for technical support. Supported by R01 MH62639 (MC), NIH R01 EY015788 (MC) and NIH P30 EY000785 (MC).

1. Equipment Set-up for Electroporation
<ol>
 <li>Electrodes: We modified Dumont #5 forceps to use as electrodes.
 <ol>
 <li>Separate and break apart the forceps.</li>
 <li>Solder a wire at the wider end of each prong. Wrap the wire attached and prongs with insulation tape leaving approximately 25-30 mm of the tip of the prongs exposed.</li>
 <li>Put the modified forceps back together with any suitable plastic spacer (e.g a button) between the two prongs to provide spring action.</li>
 </ol>
 </li>
 <li>Electrical Equipment: We use an electrical stimulator to deliver current pulses for electroporation and an oscilloscope and audio monitor to confirm the waveform and number of pulses being delivered.
 <ol>
 <li>Connect the wire from one prong of the forceps to a foot pedal which is also connected in series with the stimulator. The foot pedal acts as a switch. When depressed it completes the circuit for electroporation.</li>
 <li>Connect the wire from the other prong to the electrical stimulator.</li>
 <li>Connect the stimulator to the oscilloscope and audio monitor.</li>
 </ol>
 </li>
 <li>Micropipette and injector set-up: We use sharp pulled glass pipettes and the Nanoinject II injector system to inject very small volumes of dye or DNA solution directly into the retina.
 <ol>
 <li>Mount injector system on a 3-axis micromanipulator.</li>
 <li>Pull glass pipettes with a long taper and small tip.</li>
 <li>Back fill the pulled pipette with mineral oil and secure to injector (for details see Nanoinect II instruction manual).</li>
 <li>Using sharp microdissecting scissorsa cut the tip of the pipette, creating a small opening (~2-3 &mu;m).</li>
 <li>Fill the pipette with desired amount of injection solution through the tip of the pipette. As long as the integrity of the tip holds, it can be used for multiple injections and animals. 
 Note: i) Ensure enough injection solution is loaded in the tips such that backfilled mineral oil is never injected into the eye. ii) The Nanoinject II system and the specific electronic equipment used in this video are not critical for achieving successful labeling of RGCs. Injections made with other devices such as a picospritzer and electroporation with other common stimulators can also be used to label RGCs.</li>
 </ol>
 </li>
</ol>
2. Plasmid Solutions for RGC Labeling
<ol>
 <li>For labeling small clusters of RGCs: We use a construct encoding EGFP (~2-3 &mu;g/&mu;L) enhanced green fluorescent protein) under the control of a CAG promoter (chicken &beta;-actin promoter with a CMV immediate early enhancer). EGFP is tagged with the palmitoylation sequence of GAP-43 (growth associated protein-43) targeting it to the cell membrane (mut4EGFP)12. This construct is referred to as pCAG-gapEGFP.</li>
 <li>For single cell labeling: We use a combination of two constructs. The first vector is a CAG promoter driving Cre recombinase (pCAG-Cre, Addgene [Cambridge, MA] plasmid 13775)13 and the second vector contains a floxed STOP cassette followed by membrane targeted EGFP (pCAG-LNL-gapEGFP). pCAG-Cre (~0.15-ng/&mu;L) is used at approximately 1,000-10,000-fold lower concentration than pCAG-LNL-gapEGFP (~1-2 &mu;g&mu;L), confining strong EGFP expression to a small number of cells (by virtue of the relatively low pCAG-Cre concentration).</li>
</ol>
3. Retinal Injection and Electroporation Protocol
<ol>
 <li>Postnatal day 0 (P0) to P5 pups are anesthetized by hypothermia, while mice older than P5 are anesthetized with an intraperitoneal injection (0.7 ml/kg) of a cocktail of ketamine (4.28 mg/mL), xylazine (0.82 mg/ml), and acepromazine (0.07mg/ml)14.</li>
 <li>Sterilize all surgical instruments and electrode tips in a hot bead sterilizer. Subsequently cool both in sterile saline prior to use to avoid thermal damage to the eye.</li>
 <li>Place mouse under a dissection scope and position injector.</li>
 <li>For mice younger than P14 (before eye opening), surgically open the eyelid by cutting along the entire length of future eyelid opening with micro dissecting spring scissorsb.</li>
 <li>Make the eyeball partially protrude by gently applying pressure around the eye with the tips of forcepsc.</li>
 <li>Hold the eye in place by gently pinching the skin around the eyeball.</li>
 <li>Adjust micromanipulator and bring pipette close to the eyeball.</li>
 <li>Press foot pedal, which is connected to the Nanoinject II system, to expel a few drops of injection solution and thus verify that the pipette is not clogged.</li>
 <li>With one hand stabilizing the eyeball, move the micromanipulator such that the tip of the glass pipette pierces through the retinal pigment epithelium and enters the retina.</li>
 <li>Inject desired amount of solution by pressing the foot pedal for the Nanoinject II system. For example, to label single RGCs we make a single injection of 2.3-4.6nL.</li>
 <li>Retract the pipette and carefully place electrode tips directly over the injection site and depress foot pedal for stimulator to complete the circuit and electroporate the eye. We typically use square pulses with settings of 25V strength, 50msec duration, 1sec apart. 10 pulses (5 pulses of each polarity) are applied for mice older than P4 and 6 pulses (3 pulses of each polarity) are applied for P0 to P3 mice.</li>
 <li>Gently push eyeball back into socket and apply sterile ophthalmic ointment over the cut eyelids.</li>
 <li>Place animals on a temperature controlled heat pad and on complete recovery, including adequate rewarming and mobility, return animal to their mother.</li>
 <li>Monitor animals every 12 hours for any sign of pain, distress or discomfort, such as loss of mobility, abnormal posture or failure to groom. Repeated rounds of anesthesia, injections and electroporation should not be performed on the same animal.</li>
</ol>
4. Notes
<ol>
 <li>The method of injection described in this video is a 'blind' injection. It is possible to visualize the amount and location of injection when working with albino mice and colored solutions (DiI or plasmid solution with trace amounts of fast blue). However, with pigmented mice strains the injection location cannot be directly visualized and hence it is difficult to verbally describe whether one has reached the intended retinal location or how far one should move the pipette to target the RGC layer specifically. Therefore, we strongly advise new users to first start with DiI injections in albino mice since these injections can be immediately visualized in the retina and the labeling of RGC projections to the retina and brain can be used to assess quality of injection. Prepare 10% DiI in N,N-Dimethylformamide (100%) for focal injections. Once this procedure is mastered users should be able to consistently target RGCs with clear DNA plasmid solutions in pigmented mice.</li>
 <li>The method of injection described in this video can be used to make focal DiI injections to label small populations of cells to study retinotopy (4.6nL labels a few hundred RGCS)14 and to bulk label RGCs with fluorophores (Alexa555, 488 etc) conjugated to cholera toxin subunit B to study eye-specific segregation, excluding the electroporation step in #10 above. A maximal total injection volume of 2-3uL for mice older than P14 and 1-2uL for mice younger than P14 of any solution is recommended.</li>
 <li>Pipettes can be backfilled with solutions to be injected followed by backfilling with mineral oil before mounting the pipette on the injector and breaking the tip. This is especially useful when using high concentration DNA solutions (~6ug/uL), which are general very viscous and difficult to load from the tip of the pipette.</li>
 <li>If the glass pipette becomes clogged, carefully wipe the pipette tip with a cotton swab soaked in water for DNA solutions and ethanol (100%) for dye (DiI) solutions.</li>
 <li>After injecting in the eye, retract the pipette tip from the eyeball and press the foot pedal to inject again. This is to confirm that the tip did not get clogged during the injection process. If no solution is dispensed, it is quite likely that the injection was not successful. However, this should not be taken as an absolute sign that the injection was not successful. The experimenter may inject the same eye again if multiple injection sites and labeling of larger number of cells is desired.</li>
 <li>For labeling single RGCs, set the injection speed to slow on the Nanoinject control box. This further reduces the amount of solution injected into the retina as the system takes more time at this setting to dispense the solution while simultaneously the tip of the glass pipettes is being retracted quickly from the eyeball.</li>
 <li>Electroporation of postnatal day 0 to 3 (P0 to P3) mice requires additional care since it is very easy to damage the eye (this becomes obvious after a few days of recovery, when the eye is demonstrably smaller than normal). Reducing the number of pulses and voltage applied is advisable for early neonatal mice. After P4, the eyeballs are more resilient to damage from the electroporation.</li>
 <li>In our experience membrane targeted versions of EGFP or RFP ('gapEGFP', see12) are far superior to unmodified GFP for examining retinofugal projections to the brain. However, pCAG-tdTomato, which lacks a membrane targeting modification, also works well (Figure 1E). In addition, dextran conjugated fluorescent dyes can also be used to label RGCs using the electroporation protocol described above.</li>
 <li>Typically, 9 out of 10 injections will lead to labeled RGCs, though only about 15% of injections with the dual plasmid approach will result in just a single RGC being labeled. The number of successful cases can be increased by injecting DNA in multiple locations in one eye or by injecting plasmids encoding different fluorescent proteins into each eye.</li>
</ol>
5. Representative Results
RGC labeling was observed at all ages, ranging from P2 to P25, with EGFP labeling in RGCs by 24hrs after electroporation and maintained expression for at least three weeks after transfection.
Fluorescently labeled dendrites (Figure 1A, B) and axonal arbors (Figure 1F) of single RGCs can be clearly visualized and reconstructed.
Apart from RGCs, this technique can be used to label other retinal cell types such as horizontal cells, bipolar cells and various amacrine cell subtypes (Figure 1C)
This method does not interfere with the normal time course of visual map refinement as demonstrated by normal retinotopy in the SC (Figure 1E).
At all ages, in about 90% of cases, a small volume injection (~2.3-4.6nL) of pCAG-gapEGFP led to expression in a few RGCs (Figure 1D).
In approximately 15% of the trials using a single injection of the pCAG-Cre and pCAG-LNL-gapEGFP combination plasmids per animal, led to single retinal neuron labeling including RGCs (Figure 1A,B,D) and other cell types such as amacrine cells (Figure 1C). 
<img src="/files/ftp_upload/2678/2678fig1.jpg" alt="Figure 1" /> 
Figure 1 - A, B. Examples of EGFP labeled single retinal ganglion cells (arrow head pointing to axon) in a flat-mount retina at post-natal day 14 (P14). C. Example of single starburst amacrine cell at P8. D. A cluster of EGFP labeled retinal neurons including RGCs and amacrine cells in a flat-mount retina at P14. E. Dorsal and ventral RGCs in the right eye were electroporated and labeled with EGFP and temporal and ventral RGCs in the left eye were electroplated and labeled with tdTomato at P1. The target zones (asterisks) formed by the labeled RGCs can be seen in their topographically correct location in the SC at P9 (whole-mount, white outline). F. Example of a single EGFP labeled RGC arbor (2-D projection) in a sagittal section (250 &mu;m thick) of the SC (dotted line). For clarity, images in (D) and (F) have been converted to grayscale and inverted. Scale bars (&mu;m) : (A) - (D), (F): 100; (E): 500

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	<li>Huberman, A. D., Feller, M. B., Chapman, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+underlying+development+of+visual+maps+and+receptive+fields.">Mechanisms underlying development of visual maps and receptive fields.</a> Annu Rev Neurosci. 31, 479-509 (2008).</li><li>McLaughlin, T., Torborg, C. L., Feller, M. B., O'Leary, D. 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No conflicts of interest declared.

In this video we demonstrate an in vivo electroporation protocol that results in labeling of single or small clusters of retinal neurons in postnatal mice with DNA constructs encoding fluorescent proteins. Small clusters of fluorescently labeled RGC projections to the dLGN and SC reproduced similar projection patterns as previous studies using RGC labeling with lipophilic dyes, indicating that electroporation did not interfere with normal RGC axon arbor refinement. We have utilized this protocol to analyze the r...

<table border="1">
 <tbody>
 <tr>
 <th>Materials</th>
 <th>Company</th>
 <th>Catalog number</th>
 </tr>
 <tr>
 <td>Dumont #5 Forceps</td>
 <td>Fine Science Tools</td>
 <td>11252-20</td>
 </tr>
 <tr>
 <td>Electrical Stimulator</td>
 <td>Grass Instruments</td>
 <td>Model S4</td>
 </tr>
 <tr>
 <td>Oscilloscope</td>
 <td>Agilent</td>
 <td>Model 54621A</td>
 </tr>
 <tr>
 <td>Audio monitor</td>
 <td>Grass Instruments</td>
 <td>Model AM8B</td>
 </tr>
 <tr>
 <td>Puller</td>
 <td>Sutter Instruments</td>
 <td>Model P-97</td>
 </tr>
 <tr>
 <td>Vannas Scissors a</td>
 <td>World Precision Instruments</td>
 <td>14003</td>
 </tr>
 <tr>
 <td>Micro Scissors b</td>
 <td>Ted Pella</td>
 <td>1347</td>
 </tr>
 <tr>
 <td>Dumont AA Forceps c</td>
 <td>Fine Science Tools</td>
 <td>11210-20</td>
 </tr>
 <tr>
 <td>Nanoinject II System</td>
 <td>Drummond Scientific</td>
 <td>3-000-204</td>
 </tr>
 <tr>
 <td>Glass Pipettes</td>
 <td>Drummond Scientific</td>
 <td>3-000-203-G/X</td>
 </tr>
 <tr>
 <td>Foot pedal</td>
 <td>Drummond Scientific</td>
 <td>3-000-026</td>
 </tr>
 <tr>
 <td>Mineral Oil</td>
 <td>Sigma-Aldrich</td>
 <td>M3516</td>
 </tr>
 <tr>
 <td>DiI</td>
 <td>Invitrogen</td>
 <td>D-383</td>
 </tr>
 <tr>
 <td>N,N-Dimethylformamide</td>
 <td>Sigma</td>
 <td>D4551</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation at JoVE.com

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.

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 ...

transfection-mouse-retinal-ganglion-cells-vivo

Research

JoVE Journal

Neuroscience

15.4K Views.  Yale University. Ci mostrano un In vivo Protocollo di elettroporazione per trasfezione singoli o piccoli gruppi di cellule gangliari della retina (RGCs) e altri tipi di cellule della retina nei topi postnatale in un ampio intervallo di età. La capacità di etichetta e manipolare geneticamente RGCs postnatale In vivo È un potente strumento per gli studi di sviluppo.

Video: Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

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 ...

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 ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance molecules, local gene transfer or knock- out is required. Gene targeting knock-in or knock-out into local regions is possible to perform with combination with a specific CRE line, which is laborious, costly, and time consuming. Therefore, a simple transfection method, an in utero electroporation technique, which can be performed with short time, will be handy to test the possible function of candidate genes prior to the generation of transgenic animals 1,2. In addition to this, in utero electroporation targets areas of the brain where no specific CRE line exists, and will limit embryonic lethality 3,4. Here, we present a method of in utero electroporation combining two different types of electrodes for simple and convenient gene transfer into target areas of the developing brain. First, a unique holding method of embryos using an optic fiber optic light cable will make small embryos (from E9.5) visible for targeted DNA solution injection into ventricles and needle type electrodes insertion to the targeted brain area 5,6. The patterning of the brain such as cortical area occur at early embryonic stage, therefore, these early electroporation from E9.5 make a big contribution to understand entire area patterning event. Second, the precise shape of a capillary prevents uterine damage by making holes by insertion of the capillary. Furthermore, the precise shape of the needle electrodes are created with tungsten and platinum wire and sharpened using sand paper and insulated with nail polish 7, a method which is described in great detail in this protocol. This unique technique allows transfection of plasmid DNA into restricted areas of the brain and will enable small embryos to be electroporated. This will help to, open a new window for many scientists who are working on cell differentiation, cell migration, axon guidance in very early embryonic stage. Moreover, this technique will allow scientists to transfect plasmid DNA into deep parts of the developing brain such as thalamus and hypothalamus, where not many region-specific CRE lines exist for gain of function (GOF) or loss of function (LOF) analyses.

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

A gene transfer method into the developing mouse brain is described by using a unique surgical method and special shape of electrodes. This unique technique allows transfection of plasmid DNA temporally and spatially, which will aid many neuroscientists in studying brain development.

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance ...

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

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 ...

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.

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 ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

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 ...

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in vivo&#8212;rapidly, repetitively, and at a high resolution. This protocol describes OCT imaging in the mouse retina as a powerful tool to study optic neuropathies (OPN). The OCT system is an interferometry-based, non-invasive alternative to common post mortem histological assays. It provides a fast and accurate assessment of retinal thickness, allowing the possibility to track changes, such as retinal thinning or thickening. We present the imaging process and analysis with the example of the Opa1delTTAG mouse line. Three types of scans are proposed, with two quantification methods: standard and homemade calipers. The latter is best for use on the peripapillary retina during radial scans; being more precise, is preferable for analyzing thinner structures. All approaches described here are designed for retinal ganglion cells (RGC) but are easily adaptable to other cell populations. In conclusion, OCT is efficient in mouse model phenotyping and has the potential to be used for the reliable evaluation of therapeutic interventions.

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

This manuscript describes a protocol for the in vivo imaging of the mouse retina with high-resolution spectral domain optical coherence tomography (SD-OCT). It focuses on retinal ganglion cells (RGC) in the peripapillary region, with several scanning and quantifying approaches described.

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in ...

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...