We thank Humana Press for permission to publish the microinjection pipette fabrication figure which originally appeared on page 130 of reference 11; Larry Dlugas and Steven Wong, OHSU Department of Educational Communications, for videography; Larry Dlugas for video design and editing; Adam M. O'Quinn, Senior Designer, Trion/Envirco for designing our custom horizontal laminar flow hood and Les Goldsmith for providing the technical schematic; Victor Monterroso, MV, MS, PhD and Tom Chatkupt, DVM, OHSU Department of Comparative Medicine, for guidance with our animal care protocol, surgical techniques, and prophylactic analgesia regimen; Marcel Perret-Gentil, DVM, MS, for sharing his handout on veterinary suturing techniques; Edward Porsov, MS, for designing our Adobe Premiere Pro video microscopy computer workstation; and Leah White and Jonas Hinckley of LNS Captioning (Portland, OR). This work was supported by grants from the National Institute on Deafness and other Communication Disorders: DC R01 008595 and DC R01 008595-04S2 (to JB) and P30 DC005983 (Oregon Hearing Research Center Core Grant, Peter Gillespie, Principal Investigator).

1. Ventral Laparotomy
<ol>
 <li> Anesthetize a dam whose embryos are at embryonic day 11.5 (E11.5; noon on the day a vaginal plug is detected is day 0.5 of embryonic development) by intraperitoneal injection of sodium pentobarbital anesthetic solution (7.5 &mu;L per gram body weight). Working anesthetic solution: 180 &mu;L of 50 mg/mL pentobarbital sodium solution; 100 &mu;L of absolute ethanol; 320 &mu;L of 65 mg/mL aqueous magnesium sulfate (modulates uterine tone); and 400 &mu;L of propylene glycol (vehicle miscible with aqueous and organic components). </li>
 <li> Assess completeness of anesthesia by conducting noxious stimuli tests: paw squeeze; tail pinch; and blink response to cheek and vibrissae touch. Apply sterile ophthalmic ointment to the corneas. </li>
 <li> Shave the abdominal fur from the suprapubic area to the rib cage with shears and a fine blade (Oster #40 blade). Disinfect the abdomen with 70% ethanol, 10% povidone iodine (Betadine), and 70% ethanol, sequentially. Place mouse abdomen-side down on a sterile drape and then set on a heating pad or warm plate (37 &deg;C) for 2-5 mins. </li>
 <li> Incise the abdominal skin in the ventral midline with ball-tipped scissors. Extend the incision for 10-14 mm. Identify the linea alba, an avascular, white connective tissue band along the ventral midline of the abdominal wall. Incise the linea alba with ball tipped scissors and extend the incision 10-14 mm. Immediately irrigate the abdomen with prewarmed (37 &deg;C) lactated Ringer's solution. </li>
 <li> Externalize the two horns of the uterus with ring forceps without applying excessive pressure on the implantation sites. Irrigate the externalized uterine horns with prewarmed, lactated Ringer's solution. </li>
</ol>
2. Transuterine Microinjection
<ol>
 <li> Fabricate a thick-walled, borosilicate glass capillary microinjection pipette with the following characteristics: 12-16 &mu;m outer diameter and a 20 degree bevel. On a Sutter P-97 pipette puller with small box filament, use the following program: heat = ramp test plus 3 units; pressure = 200; pull = 0; velocity = 46; time = 100). With the Sutter BV-10 beveler, use the 104C (gold) abrasive disk for large diameter pipettes. Resuspend expression plasmid at 3-4 &mu;g/&mu;L in calcium free phosphate buffered saline (pH 7.2-7.4). Add crystalline fast green to the concentrated DNA, vortex gently for 30 seconds, and spin at 10,000 g for 10 seconds. The minimum amount of fast green required to visually track the efficacy of the injection is determined empirically. Backfill the beveled pipette with the DNA/fast green solution. Connect the loaded microinjection pipette to the pipette holder of the pressure injector (Picospritzer using &gt;99% pure nitrogen as source gas). </li>
 <li> Transilluminate the uterus with low intensity, halogen light to visualize the embryo within its implantation site. Identify the beating heart, brain vesicles, limb buds, nascent 4th ventricle of the hindbrain, and eye. Irrigate the uterus every 2 minutes with prewarmed, lactated Ringer's solution to maintain hydration. Apply gentle pressure on the uterus to reorient the embryo and identify the anatomical landmarks noted above. </li>
 <li> Orient the embryo to identify the primary head vein whose anterior and posterior branches flank the mesenchymal territory in which the otocyst resides. The otocyst proper cannot be seen by transillumination of the uterus. The otocyst is located midway between the anterior and posterior branches of the primary head vein which, along with the main trunk of the vein, form the shape of the uprights or goalposts on an American football field. The otocyst is midway between the uprights. </li>
 <li> Insert the injection pipette through the uterus in a trajectory in line with the presumptive location of the otocyst. Pulse the microinjector once after passing through the uterus to visualize the tracer dye and the approximate the location of the pipette tip. Advance the pipette under micrometer control and pulse again to assess depth. Further advance the pipette into the lateral head mesenchyme and pulse repeatedly. Successful otocyst targeting will reveal the tapered shape of the endolymphatic duct dorsally and the scallop shell-shape of the vestibule. Release pressure on the uterus, remove the pipette from the embryo/uterus in one motion, and immediately irrigate the uterus with prewarmed, lactated Ringer's solution. </li>
</ol>
3. In vivo Electroporation
<ol>
 <li> Irrigate the uterus with lactated Ringer's solution. Freshly applied lactated Ringer's solution is necessary to electrically couple the paddle-style electrodes to the uterus. Moisten the tungsten surfaces of the electrodes with lactated Ringer's. Center the injected otocyst in the path of the electrodes. Gently compress the uterus with the electroporation paddles. The cathode is in contact with the uterine wall lateral to the injected otocyst and the anode is in contact with the uterine wall adjacent to the uninjected otocyst. Trigger a square wave pulse train with the foot pedal switch on the electroporator. Electroporation parameters are: 5, 50 msec pulses at 43 volts per pulse and a 950 msec interpulse delay. Immediately irrigate the uterus after the pulse train is delivered. Record the current delivered to the tissue: 60-100 mAmps is sufficient to transfect otic epithelial progenitors. Inject and electroporate 4-6, E11.5 embryos per dam. </li>
 <li> Perform a second, independent transuterine microinjection of aqueous fluorescent dextran (Alexa Fluor 488 if the expression plasmid encodes a red fluorescent protein or Alex Fluor 594 if the expression plasmid encodes a green fluorescent protein) into the 4th ventricle of those embryos whose inner ears are accurately injected and at least 60 mAmps of current per pulse was delivered during electroporation. The fluorescent dextran will be detectable at birth in the hindbrain and enable selection of pups whose inner ears were manipulated during embryogenesis (see 3.5). </li>
 <li> Irrigate the uterine horns with lactated Ringer's. Reinsert the uterine horns into the abdominal cavity. Flush the uterine cavity with 2-4 mL of prewarmed, lactated Ringer's solution and allow the overflow to drain out of the incision site onto the sterile drape. Replace the draping with dry, sterile material. Suture the abdominal wall with a non-cutting needle and a 6-0 resorbable suture. We prefer a running stitch, locking every other stitch, for both the abdominal wall and skin. </li>
 <li> Dry the dams' fur and administer a non-steroidal anti-inflammatory such as Meloxicam by subcutaneous injection. Return the dam to the prewarmed recovery cage on a sterile drape. Monitor and record the dams' respirations, incision site patency, and vagina for bloody discharge. Bleeding is rare but if present and unabated, euthanize the dam while she is still under anesthesia. Note the time she regains consciousness and attempts to ambulate. Return the dam to the mouse colony when she shows signs of eating and drinking and has begun to nest build. Typically, this occurs within 12 hrs. </li>
 <li> At birth (postnatal day 0), flash the hindbrain region of each pup in the litter to detect the fluorescent dextran using a stereofluorescence dissecting microscope with a GFP or Texas Red filter set as appropriate. Return to the lactating dam only those pups that display hindbrain labeling. </li>
</ol>
4. Representative Results
 <img src="/files/ftp_upload/3653/3653fig1.jpg" alt="Figure 1" /> 
Figure 1. Electroporation-mediated gene transfer to the developing cochlea. The E11.5 otocyst was injected with an expression plasmid encoding enhanced green fluorescent protein (EGFP) and electroporated (pulse train parameters: five, 43 volt pulses at 50 msec/pulse and 950 msec interpulse delay). A) A representative inner ear from a postnatal day 6 (P6) pup whose otocyst was injected and electroporated at E11.5 demonstrating EGFP expression from the base through the middle turn of the cochlea. The lateral wall of the cochlea was removed from the middle turn and apex only. E11.5 progenitors that give rise to the apex were not transfected. B) Whole mount immunostaining of the cochlea in (A) with the hair cell marker, myosin 7a (Myo7a), indicates that EGFP expression follows the trajectory of the organ of Corti and is grossly localized to the hair cell-bearing sensory epithelium. C) A representative inner ear from an E18.5 embryo whose otocyst was injected and electroporated at E11.5. The laser confocal projection demonstrates EGFP expression in Myo7a-positive sensory hair cells. D) Laser confocal projection of the cochlear sensory epithelium from the E18.5 organ of Corti indicating EGFP expression in the inner hair cells (ihc), outer hair cells (ohc), inner phalangial cells (ipc), pillar cells (pc), and Deiters' cells (dc). The scale bar in (B) applies to (A).

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No conflicts of interest declared.

Gene transfer to the developing mouse inner ear: The mouse inner ear develops from the otic placode during the first week of postimplantation development12,13. By embryonic day 9.5 (E9.5), the placode has invaginated and morphed into a fluid-filled vesicle called the otocyst2. Otic precursors in the vesicle give rise to the sensory and nonsensory cells within the mature inner ear as well as the neurons that innervate mechanically sensitive hair cells in the vestibular and auditory s...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>Micro Sterilizing Case </td>
 <td>ROBOZ</td>
 <td>RS-9900a</td>
 <td>8X8.5X1.25 inches</td>
 </tr>
 <tr>
 <td>Ball-tipped scissors</td>
 <td>Fine Science Tools</td>
 <td>14109-09</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Ring forceps </td>
 <td>Fine Science Tools</td>
 <td>11106-09</td>
 <td>4.8mm ID/6mm OD</td>
 </tr>
 <tr>
 <td>Adson Tissue Forceps</td>
 <td>Fine Science Tools</td>
 <td>11027-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Needle driver</td>
 <td>Fine Science Tools</td>
 <td>12502-12</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Allergy Syringe Tray</td>
 <td>Becton Dickison</td>
 <td>305536</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Suture 6-0</td>
 <td>Syneture</td>
 <td>GL-889</td>
 <td>0.7 metric gastrointestinal suture</td>
 </tr>
 <tr>
 <td>Lactated Ringer's Injection USP</td>
 <td>Baxter</td>
 <td>2B2323</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Fast green</td>
 <td>Sigma Aldrich</td>
 <td>F7258</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Borosilicate glass capillary</td>
 <td>Harvard Apparatus</td>
 <td>30-0053</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Nembutal Sodium Solution</td>
 <td>OVATION Pharmaceuticals Inc.</td>
 <td>NDC 67386-501-52</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>MgSO4.7H2O</td>
 <td>Fisher Scientific</td>
 <td>M63-500</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Propylene glycol</td>
 <td>Fisher Scientific</td>
 <td>P355-1</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Ethanol</td>
 <td>Sigma Aldrich</td>
 <td>E7023-500</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Meloxicam</td>
 <td>Boehringer Ingeheim</td>
 <td>NADA 141-219</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Micropipette Puller</td>
 <td>Sutter Instruments</td>
 <td>P-97</td>
 <td>FB255B box filament; consult Pipette 			Cookbook from Sutter instruments</td>
 </tr>
 <tr>
 <td>Microelectrode Beveler</td>
 <td>Sutter Instruments</td>
 <td>BV-10</td>
 <td>104C beveling disk for large pipettes; 			consult owner's manual for beveling theory</td>
 </tr>
 <tr>
 <td>Micropipette holder</td>
 <td>Warner Instruments</td>
 <td>MP-S15T</td>
 <td>For 1.5mm outer diameter pipette and 			female pressure port for Picospritzer tubing.</td>
 </tr>
 <tr>
 <td>Tweezers-style electrode</td>
 <td>Protech International Inc.</td>
 <td>CUY650P5</td>
 <td>5 mm outer diameter</td>
 </tr>
 <tr>
 <td>Square Wave Electroporator</td>
 <td>Protech International Inc.</td>
 <td>CUY21EDIT</td>
 <td>Footpedal recommended</td>
 </tr>
 <tr>
 <td>PICOSPRITZER III</td>
 <td>Parker Hannifin</td>
 <td>051-0500-900</td>
 <td>Footpedal recommended</td>
 </tr>
 <tr>
 <td>Manual Control Micromanipulator</td>
 <td>Harvard Apparatus</td>
 <td>640056</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Horizontal laminar flow clean bench</td>
 <td>Envirco</td>
 <td>&nbsp;</td>
 <td>Custom modifications to LF 630-10554. See 			supplementary information for hood schematic.</td>
 </tr>
 <tr>
 <td>Leica stereofluorescence dissecting microcope with Lumencor 			SOLA light engine</td>
 <td>Bartels and Stout and Lumencor</td>
 <td>MZ10F with Lumencor SOLA light engine</td>
 <td>Footpedals to focus the MZ10F and to 			trigger the SOLA light engine are recommended</td>
 </tr>
 <tr>
 <td>Alexa Fluor 594 Dextran</td>
 <td>Invitrogen</td>
 <td>D22913</td>
 <td>10mg/ml, aqueous</td>
 </tr>
 <tr>
 <td>Alexa Fluor 488 Dextran</td>
 <td>Invitrogen</td>
 <td>D22910</td>
 <td>10mg/ml, aqueous</td>
 </tr>
 <tr>
 <td>Enviro-dri</td>
 <td>Shepherd Specialty Papers</td>
 <td>&nbsp;</td>
 <td><a href="http://www.ssponline.com" target="_blank">www.ssponline.com</a></td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation at JoVE.com

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.

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 ...

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JoVE Journal

Neuroscience

14.0K Views.  Oregon Health & Science University. 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.

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

Gene Delivery to Postnatal Rat Brain by Non-ventricular Plasmid Injection and Electroporation

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals are not viable in those cases where the modified gene is expressed or deleted in the whole organism. Moreover, a variety of compensatory mechanisms often make it difficult to interpret the results. The compensatory effects can be alleviated by either timing the gene expression or limiting the amount of transfected cells.
The method of postnatal non-ventricular microinjection and in vivo electroporation allows targeted delivery of genes, siRNA or dye molecules directly to a small region of interest in the newborn rodent brain. In contrast to conventional ventricular injection technique, this method allows transfection of non-migratory cell types. Animals transfected by means of the method described here can be used, for example, for two-photon in vivo imaging or in electrophysiological experiments on acute brain slices.

This protocol describes a non-viral method of delivery of genetic constructs to a certain area of living rodent brain. The method consists of plasmid preparation, micropipette fabrication, neonatal rat pup surgery, microinjection of the construct, and in vivo electroporation.

Creation of transgenic animals is a standard approach in studying functions of a gene of interest in vivo. However, many knockout or transgenic animals ...

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 ...

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to cis-regulatory elements (CREs), short (&tilde;300-600 bp) stretches of genomic DNA which can lie upstream, downstream, or within the introns of the genes they control. CREs (i.e., enhancers/promoters) typically consist of multiple clustered binding sites for both transcriptional activators and repressors1-3. They serve as logical integrators of transcriptional input giving a unitary output in the form of spatiotemporally precise and quantitatively exact promoter activity. Most studies of mammalian cis-regulation to date have relied on mouse transgenesis as a means of assaying the enhancer function of CREs4-5. This technique is time-consuming, costly and, on account of insertion site effects, largely non-quantitative. On the other hand, quantitative assays for mammalian CRE function have been developed in tissue culture systems (e.g., dual luciferase assays), but the in vivo relevance of these results is often uncertain. 
Electroporation offers an excellent alternative to traditional mouse transgenesis in that it permits both spatiotemporal and quantitative assessment of cis-regulatory activity in living mammalian tissue. This technique has been particularly useful in the analysis of cis-regulation in the central nervous system, especially in the cerebral cortex and the retina6-8. While mouse retinal electroporation, both in vivo and ex vivo, has been developed and extensively described by Matsuda and Cepko6-7,9, we have recently developed a simple approach to quantify the activity of photoreceptor-specific CREs in electroporated mouse retinas10. Given that the amount of DNA that is introduced into the retina by electroporation can vary from experiment to experiment, it is necessary to include a co-electroporated 'loading control' in all experiments. In this respect, the technique is very similar to the dual luciferase assay used to quantify promoter activity in cultured cells. 
When assaying photoreceptor cis-regulatory activity, electroporation is usually performed in newborn mice (postnatal day 0, P0) which is the time of peak rod production11-12. Once retinal cell types become post-mitotic, electroporation is much less efficient. Given the high rate of rod birth in newborn mice and the fact that rods constitute more than 70% of the cells in the adult mouse retina, the majority of cells that are electroporated at P0 are rods. For this reason, rod photoreceptors are the easiest retinal cell type to study via electroporation. The technique we describe here is primarily useful for quantifying the activity of photoreceptor CREs.

Quantifying the Activity of cis-Regulatory Elements in the Mouse Retina by Explant Electroporation

This protocol describes a simple and inexpensive way to quantify the activity of cis-regulatory elements (i.e., enhancer/promoters) in living mouse retinas via explant electroporation. DNA preparation, retinal dissection, electroporation, retinal explant culture, and post-fixation analysis and quantification are described.

Transcription factors within cellular gene networks control the spatiotemporal pattern and levels of expression of their target genes by binding to ...

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 ...

Isolating LacZ-expressing Cells from Mouse Inner Ear Tissues using Flow Cytometry

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues presents a unique challenge because of the paucity of cells of interest and few transgenic reporter mouse models. Here, we describe a protocol using fluorescence-conjugated probes to selectively label LacZ-positive cells from the neonatal cochleae.
The most common underlying pathology of sensorineural hearing loss is the irreversible damage and loss of cochlear sensory hair cells, which are required to transduce sound waves to neural impulses. Recent evidence suggests that the murine auditory and vestibular organs harbor stem/progenitor cells that may have regenerative potential1,2. These findings warrant further investigation, including identifying specific cell types with stem/progenitor cell characteristics. The Wnt signaling pathway has been demonstrated to play a critical role in maintaining stem/progenitor cell populations in several organ systems3-7. We have recently identified Wnt-responsive Axin2-expressing cells in the neonatal cochlea, but their function is largely unknown8.
To better understand the behavior of these Wnt-responsive cells in vitro, we have developed a method of isolating Axin2-expressing cells from cochleae of Axin2-LacZ reporter mice9. Using flow cytometry to isolate Axin2-LacZ positive cells from the neonatal cochleae, we could in turn execute a variety of experiments on live cells to interrogate their behavior as stem/progenitor cells. Here, we describe in detail the steps for the microdissection of neonatal cochlea, dissociation of these tissues, labeling of the LacZ-positive cells using a fluorogenic substrate, and cell sorting. Techniques for dissociating cochleae into single cells and isolating cochlear cells via flow cytometry have been described2,10-12. We have made modifications to these techniques to establish a novel protocol to isolate LacZ-expressing cells from the neonatal cochlea.

Flow cytometry is a powerful tool allowing for the isolation and study of specific cell populations. This protocol describes steps for isolating LacZ-expressing cells from cochlear tissues from neonatal transgenic mice. Dissociated cochlear cells were labeled using fluorescent-conjugated substrates of &#946;-galactosidase prior to separation via flow cytometry.

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Patch Clamp Recordings in Inner Ear Hair Cells Isolated from Zebrafish

Patch clamp analyses of the voltage-gated channels in sensory hair cells isolated from a variety of species have been described previously1-4 but this video represents the first application of those techniques to hair cells from zebrafish. Here we demonstrate a method to isolate healthy, intact hair cells from all of the inner ear end-organs: saccule, lagena, utricle and semicircular canals. Further, we demonstrate the diversity in hair cell size and morphology and give an example of the kinds of patch clamp recordings that can be obtained. The advantage of the use of this zebrafish model system over others stems from the availability of zebrafish mutants that affect both hearing and balance. In combination with the use of transgenic lines and other techniques that utilize genetic analysis and manipulation, the cell isolation and electrophysiological methods introduced here should facilitate greater insight into the roles hair cells play in mediating these sensory modalities.

The purpose of this video is to demonstrate procedures for obtaining healthy, intact hair cells from the inner ear organs of adult zebrafish and then using them for patch clamp studies aimed at characterizing the biophysical properties of their voltage-gated channels.

Patch  clamp analyses of the voltage-gated channels in sensory hair cells isolated  from a variety of species have been described previously1-4 but this  ...

In ovo Electroporation of miRNA-based Plasmids in the Developing Neural Tube and Assessment of Phenotypes by DiI Injection in Open-book Preparations

Commissural dI1 neurons have been extensively studied to elucidate the mechanisms underlying axon guidance during development1,2. These neurons are located in the dorsal spinal cord and send their axons along stereotyped trajectories. Commissural axons initially project ventrally towards and then across the floorplate. After crossing the midline, these axons make a sharp rostral turn and project longitudinally towards the brain. Each of these steps is regulated by the coordinated activities of attractive and repulsive guidance cues. The correct interpretation of these cues is crucial to the guidance of axons along their demarcated pathway. Thus, the physiological contribution of a particular molecule to commissural axon guidance is ideally investigated in the context of the living embryo. Accordingly, gene knockdown in vivo must be precisely controlled in order to carefully distinguish axon guidance activities of genes that may play multiple roles during development.
Here, we describe a method to knockdown gene expression in the chicken neural tube in a cell type-specific, traceable manner. We use novel plasmid vectors3 harboring cell type-specific promoters/enhancers that drive the expression of a fluorescent protein marker, followed directly by a miR30-RNAi transcript4 (located within the 3'-UTR of the cDNA encoding the fluorescent protein) (Figure 1). When electroporated into the developing neural tube, these vectors elicit efficient downregulation of gene expression and express bright fluorescent marker proteins to enable direct tracing of the cells experiencing knockdown3. Mixing different RNAi vectors prior to electroporation allows the simultaneous knockdown of two or more genes in independent regions of the spinal cord. This permits complex cellular and molecular interactions to be examined during development, in a manner that is fast, simple, precise and inexpensive. In combination with DiI tracing of commissural axon trajectories in open-book preparations5, this method is a useful tool for in vivo studies of the cellular and molecular mechanisms of commissural axon growth and guidance. In principle, any promoter/enhancer could be used, potentially making the technique more widely applicable for in vivo studies of gene function during development6.
This video first demonstrates how to handle and window eggs, the injection of DNA plasmids into the neural tube and the electroporation procedure. To investigate commissural axon guidance, the spinal cord is removed from the embryo as an open-book preparation, fixed, and injected with DiI to enable axon pathways to be traced. The spinal cord is mounted between coverslips and visualized using confocal microscopy.

In ovo Electroporation of miRNA-based Plasmids in the Developing Neural Tube and Assessment of Phenotypes by DiI Injection in Open-book Preparations

A method by which gene expression in the neural tube can be downregulated in a cell type-specific, traceable manner is described. We demonstrate how in ovo electroporation of microRNA-based plasmids that elicit spatiotemporally controlled RNA interference can be used to investigate commissural axon guidance in the developing neural tube.

Commissural dI1 neurons have been extensively studied to elucidate the mechanisms underlying axon guidance during development1,2. These neurons are ...

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Genetic modification of specific regions of the developing mammalian brain is a very powerful experimental approach. However, generating novel mouse mutants is often frustratingly slow. It has been shown that access to the mouse brain developing in utero with reasonable post-operatory survival is possible. Still, results with this procedure have been reported almost exclusively for the most superficial and easily accessible part of the developing brain, i.e. the cortex. The thalamus, a narrower and more medial region, has proven more difficult to target. Transfection into deeper nuclei, especially those of the hypothalamus, is perhaps the most challenging and therefore very few results have been reported. Here we demonstrate a procedure to target the entire hypothalamic neuroepithelium or part of it (hypothalamic regions) for transfection through electroporation. The keys to our approach are longer narcosis times, injection in the third ventricle, and appropriate kind and positioning of the electrodes. Additionally, we show results of targeting and subsequent histological analysis of the most recessed hypothalamic nucleus, the mammillary body.

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Despite the functional and medical importance of the hypothalamus, in utero genetic manipulation of its development has rarely been attempted. We show a detailed procedure for in utero electroporation into the mouse hypothalamus and show representative results of total and partial (regional) hypothalamic transfection.

Genetic  modification of specific regions of the developing mammalian brain is a very  powerful experimental approach. However, generating novel mouse ...