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 us...

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.

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			<td>ECM 830 Electro Square Porator</td>
			<td>BTX/ Harvard Apparatus</td>
			<td>45-0052</td>
			<td></td>
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			<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>
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			<td>Polyimide Tubing</td>
			<td>Vention Medical</td>
			<td>custom made</td>
			<td>Internal Diameter: 0.5mm / wall thickness: 0.15-0.2mm. Used for insulating gold tiped electrode</td>
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			<td>2.5mm square box filament, 4.5mm wide</td>
			<td>Sutter Instrument Company</td>
			<td>FB245B</td>
			<td>Used to make cathode electrode</td>
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			<td>HBSS, no calcium, no magnesium, no phenol red</td>
			<td>Gibco - Life Technologies</td>
			<td>14175-095</td>
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			<td>Moloney forceps</td>
			<td>Roboz</td>
			<td>RS-8254</td>
			<td>Serrated; Slight Curve; 4.5&#34; Length</td>
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			<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>
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			<td>Bonn micro forceps, 1x2 teeth</td>
			<td>Roboz</td>
			<td>RS-5172</td>
			<td>Tying Platform; 1X2 Teeth, 0.12mm Teeth; 3.75&#34; Length; .3mm Tip Width</td>
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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 cultu...

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 ...

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Research

JoVE Journal

Developmental Biology

8.1K Views.  The Johns Hopkins University School of Medicine.  Эмбриональные цыпленок культуры сетчатки клеток представляют собой ценный инструмент для изучения биологии фоторецепторов. Мы разработали эффективную методику переноса генов на основе экс ово плазмиды электропорации сетчатки, предшествующих культу?...

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

Optimized Ex-ovo Culturing of Chick Embryos to Advanced Stages of Development

Research in anatomy, embryology, and developmental biology has largely relied on the use of model organisms. In order to study development in live embryos model organisms, such as the chicken, are often used. The chicken is an excellent model organism due to its low cost and minimal maintenance, however they present observational challenges because they are enclosed in an opaque eggshell. In order to properly view the embryo as it develops, the shell must be windowed or removed. Both windowing and ex ovo techniques have been developed to assist researchers in the study of embryonic development. However, each of the methods has limitations and challenges. Here, we present a simple, optimized ex ovo culture technique for chicken embryos that enables the observation of embryonic development from stage HH 19 into late stages of development (HH 40), when many organs have developed. This technique is easy to adopt in both undergraduate classes and more advanced research laboratories where embryo manipulations are conducted.

Optimized Ex-ovo Culturing of Chick Embryos to Advanced Stages of Development

Viewing and accessing the chicken embryo during development can be challenging. We have developed an ex ovo method that is simple, cost effective, and can easily be used in a classroom or research setting. This method provides access to the embryo into late stages of embryonic development (HH 40).

Research in anatomy, embryology, and developmental biology has largely relied on the use of model organisms. In order to study development in live embryos ...

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 ...

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are reprogrammed from human fibroblasts using episomal vectors. Colonies of iPSCs can be observed 30 days after initiation of fibroblast reprogramming. Pluripotent colonies are manually picked and grown in neural induction medium to permit differentiation into neural progenitor cells (NPCs). iPSCs rapidly convert into neuroepithelial cells within 1 week and retain the capability to self-renew when maintained at a high culture density. Primary mouse NPCs are differentiated into astrocytes by exposure to a serum-containing medium for 7 days and form a monolayer upon which embryonic day 18 (E18) rat cortical neurons (transfected with channelrhodopsin-2 (ChR2)) are added. Human NPCs tagged with the fluorescent protein, tandem dimer Tomato (tdTomato), are then seeded onto the astrocyte/cortical neuron culture the following day and allowed to differentiate for 28 to 35 days. We demonstrate that this system forms synaptic connections between iPSC-derived neurons and cortical neurons, evident from an increase in the frequency of synaptic currents upon photostimulation of the cortical neurons. This co-culture system provides a novel platform for evaluating the ability of iPSC-derived neurons to create synaptic connections with other neuronal populations.

A protocol to generate a co-culture system consisting of neurons derived from induced pluripotent stem cells (iPSCs), primary cortical neurons and astrocytes is described. This co-culture system allows detection of the formation of synaptic contacts and circuits between new, iPSC-derived neurons and pre-existing cortical neurons expressing channelrhodopsin-2.

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are ...

Forward Genetic Approach to Uncover Stress Resistance Genes in Mice &#8212; A High-throughput Screen in ES Cells

Phenotype-driven genetic screens in mice is a powerful technique to uncover gene functions, but are often hampered by extremely high costs, which severely limits its potential. We describe here the use of mouse embryonic stem (ES) cells as surrogate cells to screen for a phenotype of interest and subsequently introduce these cells into a host embryo to develop into a living mouse carrying the phenotype. This method provides (1) a cost effective, high-throughput platform for genetic screen in mammalian cells; (2) a rapid way to identify the mutated genes and verify causality; and (3) a short-cut to develop mouse mutants directly from these selected ES cells for whole animal studies. We demonstrated the use of paraquat (PQ) to select resistant mutants and identify mutations that confer oxidative stress resistance. Other stressors or cytotoxic compounds may also be used to screen for resistant mutants to uncover novel genetic determinants of a variety of cellular stress resistance.

Forward Genetic Approach to Uncover Stress Resistance Genes in Mice &amp;#8212; A High-throughput Screen in ES Cells

Stress resistance is one of the hallmarks for longevity and is known to be genetically governed. Here, we developed an unbiased high-throughput method to screen for mutations that confer stress resistance in ES cells with which to develop mouse models for longevity studies.

Phenotype-driven genetic screens in mice is a powerful technique to uncover gene functions, but are often hampered by extremely high costs, which severely ...

Ex Vivo Culture of Chick Cerebellar Slices and Spatially Targeted Electroporation of Granule Cell Precursors

The cerebellar external granule layer (EGL) is the site of the largest transit amplification in the developing brain, and an excellent model for studying neuronal proliferation and differentiation. In addition, evolutionary modifications of its proliferative capability have been responsible for the dramatic expansion of cerebellar size in the amniotes, making the cerebellum an excellent model for evo-devo studies of the vertebrate brain. The constituent cells of the EGL, cerebellar granule progenitors, also represent a significant cell of origin for medulloblastoma, the most prevalent paediatric neuronal tumour. Following transit amplification, granule precursors migrate radially into the internal granular layer of the cerebellum where they represent the largest neuronal population in the mature mammalian brain. In chick, the peak of EGL proliferation occurs towards the end of the second week of gestation. In order to target genetic modification to this layer at the peak of proliferation, we have developed a method for genetic manipulation through ex vivo electroporation of cerebellum slices from embryonic Day 14 chick embryos. This method recapitulates several important aspects of in vivo granule neuron development and will be useful in generating a thorough understanding of cerebellar granule cell proliferation and differentiation, and thus of cerebellum development, evolution and disease.

Ex Vivo Culture of Chick Cerebellar Slices and Spatially Targeted Electroporation of Granule Cell Precursors

The cerebellar external granule layer is the site of the largest transit amplification in the developing brain. Here, we present a protocol to target genetic modification to this layer at the peak of proliferation using ex vivo electroporation and culture of cerebellar slices from embryonic Day 14 chick embryos.

The cerebellar external granule layer (EGL) is the site of the largest transit amplification in the developing brain, and an excellent model for studying ...

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. The inner ear auditory sensory organ is critical for our ability to hear. It houses a highly specialized sensory epithelium that consists of mechano-transducing hair cells (HCs) and surrounding glial-like supporting cells (SCs). Despite structural differences in the auditory organs, molecular mechanisms regulating the development of the auditory organ are evolutionarily conserved between mammals and aves. In ovo electroporation is largely limited to early stages at E1 - E3. Due to the relative late development of the auditory organ at E5, manipulations of the auditory organ by in ovo electroporation past E3 are difficult due to the advanced development of the chicken embryo at later stages. The method presented here is a transient gene transfer method for targeting genes of interest at stage E4 - E4.5 in the developing chicken auditory sensory organ via in ovo micro-electroporation. This method is applicable for gain- and loss-of-functions with conventional plasmid DNA-based expression vectors and can be combined with in ovo cell proliferation assay by adding EdU (5-ethynyl-2&#180;-deoxyuridine) to the whole embryo at the time of electroporation. The use of green or red fluorescent protein (GFP or RFP) expression plasmids allows the experimenter to quickly determine whether the electroporation successfully targeted the auditory portion of the developing inner ear. In this method paper, representative examples of GFP electroporated specimens are illustrated; embryos were harvested 18 - 96 hr&#160;after electroporation and targeting of GFP to the pro-sensory area of the auditory organ was confirmed by RNA in situ hybridization. The method paper also provides an optimized protocol for the use of the thymidine analog EdU to analyze cell proliferation; an example of an EdU based cell proliferation assay that combines immuno-labeling and click EdU chemistry is provided.

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

The auditory organ mediates hearing. Here we present a modified in ovo micro-electroporation method optimized for studying auditory progenitor cell proliferation and differentiation in the developing chicken auditory organ.

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. ...

A Novel Ex Ovo Banding Technique to Alter Intracardiac Hemodynamics in an Embryonic Chicken System

The new model presented here can be used to understand the influence of hemodynamics on specific cardiac developmental processes, at the cellular and molecular level. To alter intracardiac hemodynamics, fertilized chicken eggs are incubated in a humidified chamber to obtain embryos of the desired stage (HH17). Once this developmental stage is achieved, the embryo is maintained ex ovo and hemodynamics in the embryonic heart are altered by partially constricting the outflow tract (OFT) with a surgical suture at the junction of the OFT and ventricle (OVJ). Control embryos are also cultured ex ovo but are not subjected to the surgical intervention. Banded and control embryos are then incubated in a humidified incubator for the desired period of time, after which 2D ultrasound is employed to analyze the change in blood flow velocity at the OVJ as a result of OFT banding. Once embryos are maintained ex ovo, it is important to ensure adequate hydration in the incubation chamber so as to prevent drying and eventually embryo death. Using this new banded model, it is now possible to perform analyses of changes in the expression of key players involved in valve development and to understand the role of hemodynamics on cellular responses in vivo, which could not be achieved previously.

A Novel Ex Ovo Banding Technique to Alter Intracardiac Hemodynamics in an Embryonic Chicken System

For the first time we present here a reproducible banding procedure to alter hemodynamics in the developing heart ex ovo. This is achieved by partially constricting the outflow tract (OFT).

The new model presented here can be used to understand the influence of hemodynamics on specific cardiac developmental processes, at the cellular and ...

An Enzyme- and Serum-free Neural Stem Cell Culture Model for EMT Investigation Suited for Drug Discovery

Epithelial to mesenchymal transition (EMT) describes the process of epithelium transdifferentiating into mesenchyme. EMT is a fundamental process during embryonic development that also commonly occurs in glioblastoma, the most frequent malignant brain tumor. EMT has also been observed in multiple carcinomas outside the brain including breast cancer, lung cancer, colon cancer, gastric cancer. EMT is centrally linked to malignancy by promoting migration, invasion and metastasis formation. The mechanisms of EMT induction are not fully understood. Here we describe an in vitro system for standardized isolation of cortical neural stem cells (NSCs) and subsequent EMT-induction. This system provides the flexibility to use either single cells or explant culture. In this system, rat or mouse embryonic forebrain NSCs are cultured in a defined medium, devoid of serum and enzymes. The NSCs expressed Olig2 and Sox10, two transcription factors observed in oligodendrocyte precursor cells (OPCs). Using this system, interactions between FGF-, BMP- and TGF&#946;-signaling involving Zeb1, Zeb2, and Twist2 were observed where TGF&#946;-activation significantly enhanced cell migration, suggesting a synergistic BMP-/TGF&#946;-interaction. The results point to a network of FGF-, BMP- and TGF&#946;-signaling to be involved in EMT induction and maintenance. This model system is relevant to investigate EMT in vitro. It is cost-efficient and shows high reproducibility. It also allows for the comparison of different compounds with respect to their migration responses (quantitative distance measurement), and high-throughput screening of compounds to inhibit or enhance EMT (qualitative measurement). The model is therefore well suited to test drug libraries for substances affecting EMT.

Epithelial to mesenchymal transition (EMT) allows cancers to become invasive. To investigate EMT, a neural stem cell (NSC)-based in vitro model devoid of serum and enzymes is described. This standardized system allows quantitative and qualitative assessment of cell migration, gene and protein expression. The model is suited for drug discovery.

Epithelial to mesenchymal transition (EMT) describes the process of epithelium transdifferentiating into mesenchyme. EMT is a fundamental process during ...

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of morphogenetic events, such as gastrulation2, neurulation3-5, somitogenesis6, heart bending7,8, and brain formation9-13, during early embryogenesis. Key to understanding morphogenetic processes is to follow them dynamically by time-lapse imaging. The acquisition of time-lapse movies of chick embryogenesis ex ovo has been limited either to short time windows or to the need for an incubator to control temperature and humidity around the embryo14. Here, we present a new technique to culture chick embryos ex ovo for high-resolution time-lapse imaging using transmitted light microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique (EC-culture)1 and allows for the culturing of chick embryos without the need for a climate chamber. The embryo is sandwiched between two identical filter paper carriers and is kept fully submerged in a simple, temperature-controlled medium covered by a layer of light mineral oil. Starting from the primitive streak stage (Hamburger-Hamilton stage 5, HH5)15 up to at least the 28-somite stage (HH16)15, embryos can be cultured with either their ventral or dorsal side up. This allows the acquisition of time-lapse movies covering about 30 hr of embryonic development. Representative time-lapse frames and movies are shown. Embryos are compared morphologically to an embryo cultured in the standard EC-culture.
The submerged filter paper sandwich provides a stable environment to study early dorsal and ventral morphogenetic processes. It also allows for live fluorescence imaging and micromanipulations, such as microsurgery, bead implantation, microinjection, gene silencing, and electroporation, and has a strong potential to be combined with immersion objectives for laser-based imaging (including light-sheet microscopy).

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Here, we present a new technique to culture chick embryos ex ovo for time-lapse imaging using transmitted light and fluorescence microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique1 that allows for the culturing of chick embryos fully-submerged in a liquid culture medium.

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of ...

CRISPR-mediated Loss of Function Analysis in Cerebellar Granule Cells Using In Utero Electroporation-based Gene Transfer

Brain malformation is often caused by genetic mutations. Deciphering the mutations in patient-derived tissues has identified potential causative factors of the diseases. To validate the contribution of a dysfunction of the mutated genes to disease development, the generation of animal models carrying the mutations is one obvious approach. While germline genetically engineered mouse models (GEMMs) are popular biological tools and exhibit reproducible results, it is restricted by time and costs. Meanwhile, non-germline GEMMs often enable exploring gene function in a more feasible manner. Since some brain diseases (e.g., brain tumors) appear to result from somatic but not germline mutations, non-germline chimeric mouse models, in which normal and abnormal cells coexist, could be helpful for disease-relevant analysis. In this study, we report a method for the induction of CRISPR-mediated somatic mutations in the cerebellum. Specifically, we utilized conditional knock-in mice, in which Cas9 and GFP are chronically activated by the CAG (CMV enhancer/chicken &#223;-actin) promoter after Cre-mediated recombination of the genome. The self-designed single-guide RNAs (sgRNAs) and the Cre recombinase sequence, both encoded in a single plasmid construct, were delivered into cerebellar stem/progenitor cells at an embryonic stage using in utero electroporation. Consequently, transfected cells and their daughter cells were labeled with green fluorescent protein (GFP), thus facilitating further phenotypic analyses. Hence, this method is not only showing electroporation-based gene delivery into embryonic cerebellar cells but also proposing a novel quantitative approach to assess CRISPR-mediated loss-of-function phenotypes.

CRISPR-mediated Loss of Function Analysis in Cerebellar Granule Cells Using In Utero Electroporation-based Gene Transfer

Conventional loss-of-function studies of genes using knockout animals have often been costly and time-consuming. Electroporation-based CRISPR-mediated somatic mutagenesis is a powerful tool to understand gene functions in vivo. Here, we report a method to analyze knockout phenotypes in proliferating cells of the cerebellum.

Brain malformation is often caused by genetic mutations. Deciphering the mutations in patient-derived tissues has identified potential causative factors ...