We would like to acknowledge David O'Brien for his support with data analysis, and all the members of the Canto-Soler lab for their critical discussions. This work was supported by NIH grants EY004859 and EY022631 (MVCS), Core Grant EY1765, and an unrestricted departmental grant from Research to Prevent Blindness, Inc.

All procedures described in this work were performed according to the guidelines recommended by the Animal Care and Use Committee at Johns Hopkins University.
1. Ahead of Time: Preparation of Instruments, Reagents and Dishes
<ol>
	<li>Preparation of Eggs: Incubate fertilized White Leghorn chicken eggs at 37.5 &#176;C and 60% relative humidity for 5 days in an egg incubator. 
	NOTE: An incubator with rocking capacity may be helpful for standardizing and synchronizing embryonic development, but rocking is not strictly necessary for the outcome of this protocol.</li>
	<li>Plasmid Preparation:
	<ol>
		<li>Design plasmid construct for gene overexpression or downregulation (for example through the use of RNAi technology1,34) as needed. Include a reporter gene (such as EGFP, RFP or LacZ) in the construct whenever possible. 
		NOTE: Some commonly used promoters such as CMV, can become silenced after time. The CAG promoter (chicken beta-actin promoter with CMV enhancer) is a very good alternative, providing efficient long term expression of the gene of interest35-37.</li>
		<li>Prepare a plasmid solution at a concentration of 1.5 &#181;g/&#181;l in sterile PBS. OPTIONAL:&#160;Prepare plasmid solution ahead of time and store it frozen until ready to use. &#160; 
		NOTE: Higher plasmid concentrations do not result in a significant increase in efficiency, whereas using lower plasmid concentrations lower the transfection efficiency1.</li>
	</ol>
	</li>
	<li>Electrodes:
	<ol>
		<li>For the anode use a thick gold tipped electrode (see Table of Specific Materials/ Equipment). Wipe clean with 70% ethanol. 
		NOTE: Insulation of the gold tipped electrode, leaving approximately 0.5 mm exposed, is advisable to minimize leakage of the electric current. Information for insulating material can be found in Table of Specific Materials.</li>
		<li>Make a cathode out of a 2.5 mm square box filament, 4.5 mm wide (normally used in micropipette pullers). With the aid of forceps, undo the square box and straighten filament into a long strip. Then, under dissection microscope and using a ruler as a guide, bend the filament into a square &#39;U&#39; shape 3 mm long at the base and 2 mm high at one side, with the other side the remaining length of the filament (Figure 1, C - D). Using forceps, dip the electrode in 96% ethanol and flame to sterilize. Then attach an electrical lead with a small alligator clip to the longer arm of the electrode.</li>
	</ol>
	</li>
	<li>Electroporation Chamber Setup:
	<ol>
		<li>Prepare the electroporation chamber by cutting the lid of a sterile 1.5 ml microcentrifuge tube and affixing it, flat side down, to a 100 mm Petri dish using tape (Figure 1E). 
		NOTE: Other container in the form of a well of similar size would work; however, a microcentrifuge tube lid is easily accessible and autoclavable. The well must be big enough to house a chick eye and cathode electrode, but not so big as to result in a waste of plasmid solution.</li>
	</ol>
	</li>
</ol>
2. Ex&#160;ovo Electroporation Procedure
<ol>
	<li>Preparation of Instruments:
	<ol>
		<li>Sterilize micro-dissection tools (2 pairs of big curved Moloney forceps for opening egg shells and scooping out embryos, 2 pairs of fine-tip curved Dumont tweezers for removing RPE, one pair of Bonn toothed forceps for removing lens and vitreous) by dipping the tips in 96% ethanol and flaming. Clean dissection scope using wipes dipped in 70% ethanol. Warm a bottle of sterile Calcium-Magnesium-free Hank&#39;s Balanced Salt Solution (CMF-HBSS) in a 37 &#176;C water bath. Fill 100 mm sterile Petri dishes to 3/4 full with warm CMF-HBSS.</li>
		<li>Place electroporation chamber under another clean dissection microscope and fill with 120 &#181;l of plasmid solution. Connect electrodes to electroporation apparatus, making sure custom-made electrode is connected to the negative pole and gold tipped electrode to the positive pole.</li>
	</ol>
	</li>
	<li>Embryo Collection:
	<ol>
		<li>Clean the eggs by wiping shells with wipes wetted in 70% ethanol. Maintain sterile procedure throughout the rest of the protocol to avoid carrying contamination into the cultures. Using big curved Moloney forceps carefully open a hole in the egg shell by gently tapping on the rounded tip of an egg (over the air chamber) and then gripping and pulling out shell pieces.</li>
		<li>With another pair of forceps remove membranes and collect embryo by scooping it out of the egg (being careful not to damage the embryo). Place it in a dish with CMF-HBSS. 
		NOTE: Collect as many embryos as necessary to obtain the desired number of retinal cups for electroporation as specified in 2.4.4</li>
		<li>CRITICAL STEP: Stage the embryos according to Hamburger and Hamilton38 . 
		NOTE: To obtain optimum efficiency embryos must be at stage 27 (Figure 2A).</li>
		<li>Euthanize the embryos by decapitation using Moloney forceps.</li>
	</ol>
	</li>
	<li>Obtaining Retinal Cups:
	<ol>
		<li>Using fine Dumont tweezers, carefully enucleate the eyes and transfer to a new CMF-HBSS dish.</li>
		<li>Starting from the posterior part of each eye, close to the optic nerve head, and using two fine Dumont tweezers, introduce one tip of each pair of tweezers between the neural retina and RPE and carefully dissect out the RPE being cautious to not damage the neural retina (Figure 2B). 
		NOTE: The lack of Ca2+ and Mg2+ in the solution will help the RPE detach from the neural retina more easily.</li>
	</ol>
	</li>
	<li>Electroporation:
	<ol>
		<li>Set up electroporator&#39;s parameters to deliver 5 square pulses of 15 Volts, 50 msec duration, and 950 msec interval. Place &#39;U&#39; shaped negative electrode inside electroporation chamber (microcentrifuge lid filled with 120 &#181;l of plasmid-containing solution).</li>
		<li>Using curved Dumont tweezers transfer one retinal cup to the electroporation chamber (by scooping, not pressing), and place it inside the &#39;U&#39; shaped electrode, with the posterior part of the retina towards the bottom of the electrode and the lens facing upwards (Figure 2D).</li>
		<li>Place the anode so that it touches the anterior part of the eye, next to the lens, and using the electroporator&#39;s foot pedal deliver the electric pulses.</li>
		<li>Transfer electroporated retinal cup to a new Petri dish with CMF-HBSS and repeat electroporation procedure with each of the remaining retinal cups. 
		NOTE: Up to 6 retinal cups can be electroporated with the same plasmid solution without a significant decrease in transfection efficiency.</li>
		<li>Remove the&#160;lens&#160;and vitreous body of each electroporated retinal cup by gripping them with Bonn toothed forceps and gently pulling through the pupil opening while holding the eye in place with the back of the curved Dumont tweezers.</li>
		<li>Proceed to the dissociation and culture of the electroporated retinal cells following standard procedures. 
		NOTE: For a detailed protocol on the dissociation and culture of chick retinal cells refer to Vergara and Canto-Soler, 2012, Additional file 421. In this protocol the cells are seeded at low density on polyornithine-coated 6-well plates or 35-mm dishes (6 x 105 cells/35 mm-diameter well or dish). The medium used is medium 199 with penicillin/glutamine, 10% fetal calf serum (FCS), and linoleic acid-BSA at 100 &#181;g/ml. The cultures are maintained in a humidified 37 &#176;C incubator with 5% CO2. Using this procedure, 5 - 6 x 106 cells can typically be obtained per each stage 27 electroporated eye. 
		NOTE: Ideally, no more than 3 hr&#160;should pass between embryo collection and cell plating, to ensure good cell survival and minimize stress. A typical timing for observation and further analysis is after 4 days in culture, since at this point the cells have achieved good morphological and molecular differentiation and survival is still high.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests or other conflicts of interest.

The most critical step for the success of this protocol is selecting the appropriate stage of the embryos. In previous publications, a range of embryonic stages is given for these cultures, typically defined by the days of incubation or embryonic days (ED); thus it is usually assumed that using ED 5 to ED 6 embryos will yield equivalent results. However we have found that on stage 27 (ED 5), transfection efficiency will be around 22% of the total cell population, as stated above; yet efficiency will decrease to 16% if using stage 28 embryos (ED 5.5), and to 12% at stage 29 (ED 6)1. Other critical steps include maintaining sterile conditions throughout the process; acquiring the necessary manual dexterity to avoid damaging the retina during dissection and electroporation; and avoiding long gaps from the time of embryo collection to the time of cell plating.
Another important consideration to ensure good transfection efficiencies is the concentration of the plasmid-containing solution: the recommended 1.5 &#181;g/&#181;l is the lowest concentration that we tested that did not result in a decrease in efficiency. This concentration translates into a high amount of plasmid material when considering that the retinal cups are bathed in that solution. Using the smallest size well that will accommodate the eye (such as a microcentrifuge tube lid) can minimize this problem. In addition, several eyes can be electroporated using the same solution; we have not seen a reduction in efficiency when electroporating up to 6 retinal cups consecutively in the same plasmid solution, and more could possibly be performed but we cannot attest to the difference in efficiency thereafter.
Previously published protocols have typically provided low efficiency of gene transfer for this type of culture, and thus they often had to rely on techniques that measure the product of enzymatic reactions on a cell lysate in order to increase sensitivity. Such an approach has the disadvantage of not allowing for cell-type discrimination and of relying on a low number of cells being responsible for the observed outcome, which may not always be reflective of the behavior of the larger cell population. Another way to circumvent the problem of low efficiency is to perform gene transfection in vivo, prior to eye enucleation and culture. This is a viable approach but it can usually be applied only to specific experimental paradigms, since both plasmid electroporation and RCAS virus infection (a common gene transduction vector in the chick) typically need to be performed at earlier developmental stages, creating a time lag between transfection and culture. This is an important consideration, because during that time lag the cells are exposed to the effects of both the extracellular microenvironment as well as intracellular factors, having significant implications in the interpretation of the experimental results. In contrast, the substantial increase in gene transfer efficiency achieved with the ex&#160;ovo electroporation approach allows for the study of gene function in a cell-intrinsic manner. Moreover, the advantage of this high transfection rate can be further increased when used in conjunction with automated high-throughput cell analysis1.
At the initial stages of the development of this technique, we maintained electroporated RPE-devoid eye cups in culture for 24 hr&#160;to evaluate the efficiency of electroporation achieved with different conditions1. Although we did not attempt to culture them for longer period of times, our experience indicates that the protocol described herein could also be applied to studies relying on retinal organotypic explants, provided appropriate long-term explant culture conditions are used.
Finally, it would be possible to expand the applicability of this ex&#160;ovo electroporation approach to other animal models. For example in our experience transfection of mouse eyes at postnatal day 1 or 4 using this protocol yielded qualitatively good results in explant cultures, but when dissociated cell cultures were attempted transfection efficiency was on the order of 6%, similar to that of other techniques. Thus, this could be a simple alternative to other protocols for this animal model, but it would need to be optimized for the particular system if high efficiencies are required. Of note, as stated above, embryonic stage at the time of electroporation was key to the high efficiency outcome obtained in the chick, so this parameter should be carefully considered when applying the technique to other models.
In conclusion, the ex&#160;ovo electroporation technique can expand the applicability of in vitro retinal systems as powerful tools to complement in vivo studies, offering new possibilities for retinal research.

Dissociated cell cultures from embryonic chick retinas have been widely used to study various aspects of photoreceptor cell biology, including their survival2-9, differentiation10-12, neurite outgrowth13, and more. The advantages of this system, developed in the 1980s by Ruben Adler and collaborators and perfected by his and other groups14-20, reside in the intrinsic characteristics of the chick as an animal model21. The large size of the chick eye, even at embryonic stages, provides large amounts of material for cultures. Moreover, when cultures are performed using embryonic day (ED) 5 - 6 retinas, 55 - 80% of their progenitor cells differentiate as photoreceptors14,15,18,22,23, and since approximately 86% of the photoreceptors in this animal are cones24, these cultures are particularly suitable for studies focusing on this cell type.
We have recently developed and characterized a simple technique that allows for high-efficiency plasmid transfection of the cells in these cultures, thus broadening the usefulness of this system by facilitating genetic missexpression studies1. The development of this technique stemmed from a void in the scientific literature of methods that would provide an acceptable level of transfection to allow for the study of gene function in a cell autonomous manner. This is in part because primary neuronal cultures are notoriously hard to transfect25,26. Some of the most commonly used techniques previously available for this purpose included chemical transfection methods such as lipofection or calcium phosphate-mediated transfection, which result in efficiencies in the order of 3-5% and can exert considerable toxicity27-32. Even though the use of plasmids with an enzymatic reporter system can circumvent the problem of poor transfection efficiency by amplifying the signal, they do not discriminate cell-specific effects, and their results are based on a small cell population that may not be representative of the whole. Another widely used method in the chick, RCAS virus infection, is only applicable to proliferating cells and thus not suitable for this primary retinal culture system33.
In the current protocol embryonic chick eyes are enucleated at stage 27 (ED 5), the retinal pigmented epithelium (RPE) is removed, and the retinal cup is placed in an electroporation chamber filled with a plasmid-containing solution and electroporated using custom-made electrodes, followed by retinal dissociation and culture using standard techniques21. After optimizing this procedure we have been able to consistently achieve transfection efficiencies on the order of 22% of the total number of cells in culture and 25% within the photoreceptor population alone, without compromising the survival and differentiation characteristics of the cultures1. Here we provide a detailed protocol outlining all the important steps of this procedure in order to ensure the success and reproducibility of this technique.

<table><tbody><tr><td>ECM 830 Electro Square Porator</td><td>BTX/ Harvard Apparatus</td><td>45-0052</td><td></td></tr><tr><td>Genetrode, L-Shaped, 5 mm Gold Tip</td><td>BTX/ Harvard Apparatus</td><td>45-0115 model 512</td><td>Gold tipped electrode used as anode</td></tr><tr><td>Polyimide Tubing</td><td>Vention Medical</td><td>custom made</td><td>Internal Diameter: 0.5 mm/wall thickness: 0.15 - 0.2 mm. Used for insulating gold tiped electrode</td></tr><tr><td>2.5 mm square box filament, 4.5 mm wide</td><td>Sutter Instrument Company</td><td>FB245B</td><td>Used to make cathode electrode</td></tr><tr><td>HBSS, no calcium, no magnesium, no phenol red</td><td>Gibco - Life Technologies</td><td>14175-095</td><td></td></tr><tr><td>Moloney forceps</td><td>Roboz</td><td>RS-8254</td><td>Serrated; Slight Curve; 4.5&quot; Length</td></tr><tr><td>Dumont tweezers 5/45</td><td>Roboz</td><td>RS-5058</td><td>Pattern #5, 45 Degree Angle; .10 X .06mm Tip Size; 109mm Length</td></tr><tr><td>Bonn micro forceps, 1x2 teeth</td><td>Roboz</td><td>RS-5172</td><td>Tying Platform; 1x2 Teeth, 0.12 mm Teeth; 3.75&quot; Length; 0.3 mm Tip Width</td></tr></tbody></table>

efficient gene transfer in chick retinas for primary cell culture studies: an ex-ovo electroporation approach

The cone photoreceptor-enriched cultures derived from embryonic chick retinas have become an indispensable tool for researchers around the world studying the biology of retinal neurons, particularly photoreceptors. The applications of this system go beyond basic research, as they can easily be adapted to high throughput technologies for drug development. However, genetic manipulation of retinal photoreceptors in these cultures has proven to be very challenging, posing an important limitation to the usefulness of the system. We have recently developed and validated an ex&#160;ovo plasmid electroporation technique that increases the rate of transfection of retinal cells in these cultures by five-fold compared to other currently available protocols1. In this method embryonic chick eyes are enucleated at stage 27, the RPE is removed, and the retinal cup is placed in a plasmid-containing solution and electroporated using easily constructed custom-made electrodes. The retinas are then dissociated and cultured using standard procedures. This technique can be applied to overexpression studies as well as to the downregulation of gene expression, for example via the use of plasmid-driven RNAi technology, commonly achieving transgene expression in 25% of the photoreceptor population. The video format of the present publication will make this technology easily accessible to researchers in the field, enabling the study of gene function in primary retinal cultures. We have also included detailed explanations of the critical steps of this procedure for a successful outcome and reproducibility.

Here we present a simple protocol for plasmid transfection into enucleated chick retinal cups for subsequent dissociated cell culture. Transfection is achieved by electroporation using easy to make custom electrodes (Figures 1 - 2). The parameters described in this protocol have been optimized to obtain transfection efficiencies that range between 20 and 27% (with an average of 22%) (Figure 3D). Notice that, for the reasons stated above, these results are quantified after 4 days in culture. In this context, transfection efficiency refers to the percentage of cells that express the transgene within the total cell population. If only the photoreceptor population is considered, transfection efficiencies increase to an average of 25% (Figure 3D). This is an important feature since these types of cultures are primarily used to study photoreceptors.
Cell cultures that have been electroporated using this technique are indistinguishable from their non-electroporated counterparts in their morphology under DIC illumination, cell survival characteristics, and expression of molecular markers such as visinin (a widely used marker of photoreceptors) and Pax61.
<img alt="Figure 1" src="/files/ftp_upload/52002/52002fig1.jpg" /> 
Figure 1. Making of the Electrodes. (A - D) The cathode is made out of a square box filament (A), which is first straightened (B) and then bent with forceps into a square &#39;U&#39; shape (C - D). The anode is a thick gold tipped electrode (D); Notice the insulation around the anode electrode, leaving only 0.5 mm of the tip exposed. (E) Electroporation setup. Inset shows higher magnification of boxed area. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52002/52002fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52002/52002fig2.jpg" /> 
Figure 2. Preparation of Retinal Cups for Electroporation.&#160;(A) Chick embryo at Hamburger and Hamilton embryonic stage 27. (B) The RPE is carefully removed from the enucleated eyes using sharp Dumont tweezers. (C) The retinal cup on the right is devoid of RPE and ready for electroporation. (D) The cathode electrode is positioned inside an electroporation chamber made out of a 1.5 ml microcentrifuge tube lid filled with plasmid solution. The retinal cup is carefully transferred into the cathode facing upwards, and the anode electrode is placed on top, next to the lens. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52002/52002fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52002/52002fig3.jpg" /> 
Figure 3.&#160;Efficiency of Transgene Expression in Cultured Cells after Electroporation.&#160;(A - C) Retinal cups were electroporated with a GFP expression plasmid, dissociated and cultured for 4 days. Fluorescence micrographs show DAPI stained nuclei (A) and GFP expressing retinal cells (B). Anti-Visinin antibody staining was performed to identify photoreceptor cells (C). (D) Graph illustrating transfection efficiency achieved by ex&#160;ovo electroporation, represented as the percentage of GFP expressing cells among the total cell population (DAPI positive), or among the photoreceptor population in particular (Visinin positive). Results represent the mean &#177; S.E.M. of 5 independent experiments. Scale bar in (C) represents 100 &#181;m and applies to (A - C). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52002/52002fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach at JoVE.com

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

Embryonic chick retinal cell cultures constitute valuable tools for the study of photoreceptor biology. We have developed an efficient gene transfer technique based on ex&#160;ovo plasmid electroporation of the retinas prior to culture. This technique considerably increases transfection efficiencies over currently available protocols, making genetic manipulation feasible for this system.

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

The cone photoreceptor-enriched cultures derived from embryonic chick retinas have become an indispensable tool for researchers around the world studying ...

efficient-gene-transfer-chick-retinas-for-primary-cell-culture

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

8.1K Views.  The Johns Hopkins University School of Medicine. Embryonic chick retinal cell cultures constitute valuable tools for the study of photoreceptor biology. We have developed an efficient gene transfer technique based on ex ovo plasmid electroporation of the retinas prior to culture. This technique considerably increases transfection efficiencies over currently available protocols, making genetic manipulation feasible for this system.

Video: Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair cells are aligned in one row of inner hair cells and three rows of outer hair cells that extend along the basal to apical axis of the cochlea. The explant culture technique described here provides an efficient method to isolate and maintain cochlear explants from the embryonic mouse inner ear. Also, the morphology and molecular characteristics of sensory hair cells and nonsensory supporting cells within the cochlear explant cultures resemble those observed in vivo and can be studied within its intrinsic cellular environment. The cochlear explants can serve as important experimental tools for the identification and characterization of molecular and genetic pathways that are involved in cellular specification and patterning. Although transgenic mouse models provide an effective approach for gene expression studies, a considerable number of mouse mutants die during embryonic development thereby hindering the analysis and interpretation of developmental phenotypes. The organ of Corti from mutant mice that die before birth can be cultured so that their in vitro development and responses to different factors can be analyzed. Additionally, we describe a technique for electroporating embryonic cochlear explants ex vivo which can be used to downregulate or overexpress specific gene(s) and analyze their potential endogenous function and test whether specific gene product is necessary or sufficient in a given context to influence mammalian cochlear development1-8.

We present a method that describes isolation and culture of cochlear explants from embryonic mouse inner ear. We also demonstrate a method for gene transfer into cochlear explants via square-wave electroporation. The in vitro explant culture coupled with gene transfer technique enables researchers to study the effects of altering gene expression during development.

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair ...

Whole Retinal Explants from Chicken Embryos for Electroporation and Chemical Reagent Treatments

The retina is a good model for the developing central nervous system. The large size of the eye and most importantly the accessibility for experimental manipulations in ovo/in vivo makes the chicken embryonic retina a versatile and very efficient experimental model. Although the chicken retina is easy to target in ovo by intraocular injections or electroporation, the effective and exact concentration of the reagents within the retina may be difficult to fully control. This may be due to variations of the exact injection site, leakage from the eye or uneven diffusion of the substances. Furthermore, the frequency of malformations and mortality after invasive manipulations such as electroporation is rather high.
This protocol describes an ex ovo technique for culturing whole retinal explants from chicken embryos and provides a method for controlled exposure of the retina to reagents. The protocol describes how to dissect, experimentally manipulate, and culture whole retinal explants from chicken embryos. The explants can be cultured for approximately 24 hr and be subjected to different manipulations such as electroporation. The major advantages are that the experiment is not dependent on the survival of the embryo and that the concentration of the introduced reagent can be varied and controlled in order to determine and optimize the effective concentration. Furthermore, the technique is rapid, cheap and together with its high experimental success rate, it ensures reproducible results. It should be emphasized that it serves as an excellent complement to experiments performed in ovo.

This protocol describes a method to dissect, experimentally manipulate and culture whole retinal explants from chicken embryos. The explant cultures are useful when high success rate, efficacy and reproducibility are needed to test the effects of plasmids for electroporation and/or reagent substances, i.e., enzymatic inhibitors.

The retina is a good model for the developing central nervous system. The large size of the eye and most importantly the accessibility for experimental ...

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 ...

Neuroscience

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 ...

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

Dopaminergic neurons located in the ventral midbrain control movement, emotional behavior, and reward mechanisms1-3. The dysfunction of ventral midbrain dopaminergic neurons is implicated in Parkinson's disease, Schizophrenia, depression, and dementia1-5. Thus, studying the regulation of midbrain dopaminergic neuron differentiation could not only provide important insight into mechanisms regulating midbrain development and neural progenitor fate specification, but also help develop new therapeutic strategies for treating a variety of human neurological disorders.
Dopaminergic neurons differentiate from neural progenitors lining the ventricular zone of embryonic ventral midbrain. The development of neural progenitors is controlled by gene expression programs6,7. Here we report techniques utilizing electroporation to express genes specifically in the midbrain of Hamburger Hamilton (HH) stage 11 (thirteen somites, 42 hours) chick embryos8,9. The external development of chick embryos allows for convenient experimental manipulations at specific embryonic stages, with the effects determined at later developmental time points10-13. Chick embryonic neural tubes earlier than HH stage 13 (nineteen somites, 48 hours) consist of multipotent neural progenitors that are capable of differentiating into distinct cell types of the nervous system. The pCAG vector, which contains both a CMV promoter and a chick &beta;-actin enhancer, allows for robust expression of Flag or other epitope-tagged constructs in embryonic chick neural tubes14. In this report, we emphasize special measures to achieve regionally restricted gene expression in embryonic midbrain dopaminergic neuron progenitors, including how to inject DNA constructs specifically into the embryonic midbrain region and how to pinpoint electroporation with small custom-made electrodes. Analyzing chick midbrain at later stages provides an excellent in vivo system for plasmid vector-mediated gain-of-function and loss-of-function studies of midbrain development. Modification of the experimental system may extend the assay to other parts of the nervous system for performing fate mapping analysis and for investigating the regulation of gene expression.

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

To assess the function and the regulation of genes during the development of midbrain dopaminergic neurons, we describe a method that involves in ovo electroporation of plasmid DNA constructs into embryonic chick ventral midbrain dopaminergic neuron progenitors. This technique can be used to achieve efficient expression of genes of interest to study different aspects of midbrain development and dopaminergic neuron differentiation.

Dopaminergic neurons located in the ventral midbrain control movement, emotional behavior, and reward mechanisms1-3. The dysfunction of ventral midbrain ...

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell differentiation, axon guidance, retinotopic organization and synapse formation[1]. One drawback is the inability to efficiently and reliably manipulate gene expression in RGCs in vivo, especially in the otherwise accessible murine visual pathways. Transgenic mice can be used to manipulate gene expression, but this approach is often expensive, time consuming, and can produce unwanted side effects. In chick, in ovo electroporation is used to manipulate gene expression in RGCs for examining retina and RGC development. Although similar electroporation techniques have been developed in neonatal mouse pups[2], adult rats[3], and embryonic murine retinae in vitro[4], none of these strategies allow full characterization of RGC development and axon projections in vivo. To this end, we have developed two applications of electroporation, one in utero and the other ex vivo, to specifically target embryonic murine RGCs[5, 6]. 
With in utero retinal electroporation, we can misexpress or downregulate specific genes in RGCs and follow their axon projections through the visual pathways in vivo, allowing examination of guidance decisions at intermediate targets, such as the optic chiasm, or at target regions, such as the lateral geniculate nucleus. Perturbing gene expression in a subset of RGCs in an otherwise wild-type background facilitates an understanding of gene function throughout the retinal pathway. Additionally, we have developed a companion technique for analyzing RGC axon growth in vitro. We electroporate embryonic heads ex vivo, collect and incubate the whole retina, then prepare explants from these retinae several days later. Retinal explants can be used in a variety of in vitro assays in order to examine the response of electroporated RGC axons to guidance cues or other factors. In sum, this set of techniques enhances our ability to misexpress or downregulate genes in RGCs and should greatly aid studies examining RGC development and axon projections.

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Here we present two techniques for manipulating gene expression in murine retinal ganglion cells (RGCs) by in utero and ex vivo electroporation. These techniques enable one to examine how alterations in gene expression affect RGC development, axon guidance, and functional properties.

The retina and its sole output neuron, the retinal ganglion cell (RGC), comprise an excellent model in which to examine biological questions such as cell ...

Biology

Visualization of Tangential Cell Migration in the Developing Chick Optic Tectum

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture can be recorded under video microscopy. For analyzing cell migration in the developing brain, slice culture is commonly used to observe cell migration parallel to the slice section, such as radial cell migration. However, limited information can be obtained from the slice culture method to analyze cell migration perpendicular to the slice section, such as tangential cell migration. Here, we present the protocols for time-lapse imaging to visualize tangential cell migration in the developing chick optic tectum. A combination of cell labeling by electroporation in ovo and a subsequent flat-mount culture on the cell culture insert enables detection of migrating cell movement in the horizontal plane. Moreover, our method facilitates detection of both individual cell behavior and the collective action of a group of cells in the long term. This method can potentially be applied to detect the sequential change of the fluorescent-labeled micro-structure, including the axonal elongation in the neural tissue or cell displacement in the non-neural tissue.

We describe the methods for fluorescent labeling of tangentially migrating cells by electroporation, and for time-lapse imaging of the labeled cell movement in a flat-mount culture in order to visualize migrating cell behavior in the developing chick optic tectum.

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture ...

Single Cell Transfection in Chick Embryos

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a thorough analysis of the fates of single cells under normal and experimental conditions. To this end, transfection methods that target single progenitors are a prerequisite. We describe here a technically straightforward method for transfecting single cells in chicken tissues in-ovo, allowing reliable lineage tracing as well as genetic manipulation. Specific tissue domains are targeted within the somite or neural tube, and DNA is injected directly into the epithelium of interest, resulting in sporadic transfection of single cells. Using reporters, clonal populations may consequently be traced for up to three days, and behavior of genetically manipulated clonal populations can be compared with that of controls. This method takes advantage of the accessibility of the chick embryo along with emerging tools for genetic manipulation. We compare and discuss its advantages over the widely-used electroporation method, and possible applications and use in additional in-vivo models are also suggested. We advocate the use of this method as a significant addition and complement for existing lineage tracing and genetic interference tools.

Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations.

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a ...

In Ovo Electroporation in Embryonic Chick Retina

Chicken embryonic retina is an excellent tool to study retinal development in higher vertebrates. Because of large size and external development, it is comparatively very easy to manipulate the chick embryonic retina using recombinant DNA/RNA technology. Electroporation of DNA/RNA constructs into the embryonic retina have a great advantage to study gene regulation in retinal stem/progenitor cells during retinal development. Different type of assays such as reporter gene assay, gene over-expression, gene knock down (shRNA) etc. can be performed using the electroporation technique. This video demonstrates targeted retinal injection and in ovo electroporation into the embryonic chick retina at the Hamburger and Hamilton stage 22-23, which is about embryonic day 4 (E4). Here we show a rapid and convenient in ovo electroporation technique whereby a plasmid DNA that expresses green fluorescent protein (GFP) as a marker is directly delivered into the chick embryonic subretinal space and followed by electric pulses to facilitate DNA uptake by retinal stem/progenitor cells. The new method of retinal injection and electroporation at E4 allows the visualization of all retinal cell types, including the late-born neurons1, which has been difficult with the conventional method of injection and electroporation at E1.52.

In Ovo Electroporation in Embryonic Chick Retina

The overall goal of this video is to show how to perform targeted retinal injection and in ovo electroporation of DNA/RNA constructs into the chick embryonic retina at the Hamburger and Hamilton stage 22-23, which is about embryonic day 4 (E4). This technique is very useful to study gene expression, gene regulation, and morphological change in developing chick retina.

Chicken embryonic retina is an excellent tool to study retinal development in higher vertebrates. Because of large size and external development, it is ...

Gene Transfer into Older Chicken Embryos by ex ovo Electroporation

The chicken embryo provides an excellent model system for studying gene function and regulation during embryonic development. In ovo electroporation is a powerful method to over-express exogenous genes or down-regulate endogenous genes in vivo in chicken embryos1. Different structures such as DNA plasmids encoding genes2-4, small interfering RNA (siRNA) plasmids5, small synthetic RNA oligos6, and morpholino antisense oligonucleotides7 can be easily transfected into chicken embryos by electroporation. However, the application of in ovo electroporation is limited to embryos at early incubation stages (younger than stage HH20 - according to Hamburg and Hamilton)8 and there are some disadvantages for its application in embryos at later stages (older than stage HH22 - approximately 3.5 days of development). For example, the vitelline membrane at later stages is usually stuck to the shall membrane and opening a window in the shell causes rupture of the vessels, resulting in death of the embryos; older embryos are covered by vitelline and allantoic vessels, where it is difficult to access and manipulate the embryos; older embryos move vigorously and is difficult to control the orientation through a relatively small window in the shell.
In this protocol we demonstrate an ex ovo electroporation method for gene transfer into chicken embryos at late stages (older than stage HH22). For ex ovo electroporation, embryos are cultured in Petri dishes9 and the vitelline and allantoic vessels are widely spread. Under these conditions, the older chicken embryos are easily accessed and manipulated. Therefore, this method overcomes the disadvantages of in ovo electroporation applied to the older chicken embryos. Using this method, plasmids can be easily transfected into different parts of the older chicken embryos10-12.

Gene Transfer into Older Chicken Embryos by ex ovo Electroporation

A method of gene transfer into chicken embryos at later incubation stages (older than Hamburger and Hamilton stage (HH) 22) is described. This method overcomes disadvantages of in ovo electroporation applied to older chicken embryos and is a useful technique to study gene function and regulation at older developmental stages.

The chicken embryo provides an  excellent model system for studying gene function and regulation during  embryonic development. In ovo electroporation is ...

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

Summary

Explore More Videos

Chapters in this video

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

Whole Retinal Explants from Chicken Embryos for Electroporation and Chemical Reagent Treatments

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

In vivo Electroporation of Developing Mouse Retina

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

In utero and ex vivo Electroporation for Gene Expression in Mouse Retinal Ganglion Cells

Visualization of Tangential Cell Migration in the Developing Chick Optic Tectum

Single Cell Transfection in Chick Embryos

In Ovo Electroporation in Embryonic Chick Retina

Gene Transfer into Older Chicken Embryos by ex ovo Electroporation