Special thanks to James Copti for editing of the manuscript. We also gratefully acknowledge funding to JN from the German Research foundation (DFG) by the SFB 870 and SPP &#8220;Integrative Analysis of Olfaction&#8221; and SPP 1738 &#8220;Emerging roles of non-coding RNAs in nervous system development, plasticity &#38; disease&#8221;, SPP1757 &#8220;Glial heterogeneity&#8221;, and Excellence Strategy within the framework of the Munich Cluster for Systems Neurology (EXC 2145 SyNergy &#8211; ID 390857198).

All animals used in this protocol were kept under standard husbandry conditions, and experiments have been performed according to the handling guidelines and regulations of EU and the Government of Upper Bavaria (AZ 55.2-1-54-2532-0916).
1. Preparation of Plasmid Mixture for Electroporation
<ol>
	<li>Dilute the plasmid of interest in sterile water and add fast green stain stock solution [1 mg/mL]. Make sure that the final concentration of the plasmid is&#160; &#8764;1 &#181;g/&#181;L. Add the stain at a concentration of no more than 3%, as its only purpose is to color the solution and visualize ventricular injection.</li>
	<li>Once prepared, mix the plasmid solution by pipetting up and down several times or by finger tapping. Store at room temperature (RT) until usage. 
	NOTE: For co-electroporation of two plasmids into the same cell, ensure that the concentration of each individual plasmid used in the mixture is at least 0.8 ng/&#181;L with a molar ratio of 1:1 to obtain 80%&#8211;90% co-electroporation efficiency.</li>
</ol>
2. Preparations for Injection and Electroporation Procedure
<ol>
	<li>Prepare the glass capillaries (outer diameter 1 mm, inner diameter 0.58 mm) necessary for the injection in the needle pulling apparatus.</li>
	<li>In order to inject the correct amount of plasmid (see above), pull the capillary at a temperature of 68.5 &#176;C with two light and two heavy weights (see Table of Materials for puller specifications). 
	NOTE: In case a different puller is used, calibrate the capillary to deliver the appropriate volume of electroporation mix.</li>
	<li>Manually set the injection device to an injection pressure of 200 hPa (by turning the Pi knob) and constant pressure of 0 hPa. Manually set the injection time to manual mode and control the pressure with foot pedal.</li>
	<li>Set the electroporation device to &#8220;LV mode&#8221; with five pulses at 54&#8211;57 V (25 ms each with 1 s intervals). Connect the electrodes to the device.</li>
	<li>Prepare one fish tank with clean fish water, where the fish will be awakened from anesthesia after the electroporation procedure. Aerate the water by keeping the air stone attached to the air pump for the entire recovery period until the fish is fully awakened.</li>
	<li>Take a regular kitchen sponge and make a longitudinal cut in the sponge to hold fish into during the injection and electroporation procedure (see previous publication3). 
	NOTE: The kitchen sponge should be regularly washed or exchanged in order to remove potential toxic chemicals.</li>
	<li>Place a small amount of highly conductive multi-purpose ultrasound gel next to the injection and electroporation setup. 
	NOTE: This will ensure adequate electrical conductivity, and consequently, distribution of electroporated cells throughout the entire telencephalon.</li>
</ol>
3. Zebrafish Anesthesia
<ol>
	<li>Prior to anesthetization, prepare a stock solution of anesthesia with 0.2% MS222 in distilled water and adjust the pH to 7 with Tris-HCl buffer. Dilute this stock 1:10 (i.e., to 0.02% MS222) using fish water.</li>
	<li>Anesthetize the fish by keeping them in this working solution until the movement of the body and gills subsides (typically for a couple of minutes).</li>
</ol>
4. Plasmid Solution Injection
<ol>
	<li>Fill the prepared glass capillary with 10 &#181;L of plasmid solution using microloader tips. Avoid the formation of air bubbles inside the capillary.</li>
	<li>Press Menu/Change Capillary on the injection device. Insert and secure the needle into the needle holder.</li>
	<li>Under a stereomicroscope with a magnification of 3.2x or 4x, cut only the tip of the capillary using fine-end forceps. Switch the injection device from Change Capillary mode into Inject mode, then apply pressure with a foot pedal to ensure that the plasmid solution is running easily out of the needle and without hindrance.</li>
	<li>Transfer the fish from the husbandry tank to the container (plastic box) with anesthetic solution. Wait for a few minutes until movement of the gills subsides.</li>
	<li>Place fish into the pre-wetted sponge with the dorsal side facing up. Perform all the following injection steps under the stereomicroscope to ensure the accuracy of procedure.</li>
	<li>Using a dissecting micro-knife from stainless steel with 40 mm cutting edge and 0.5 mm thickness, create a small hole in the fish skull at the posterior side of the telencephalon (Figure 1B), just next to the border with optic tectum. 
	NOTE: This step should be performed carefully since the hole should be very small and superficial, penetrating solely the skull, to avoid brain damage.</li>
	<li>Tilt the fish as necessary and orient the tip of the glass capillary towards the skull in the correct angle to facilitate penetration of the capillary tip through the hole.</li>
	<li>Insert the tip of the capillary through the hole in the skull carefully until it reaches the telencephalic ventricle (see Figure 1B). This will require penetration through the dorsal ependymal cell layer. Be especially careful not to insert the capillary too deeply to avoid contact with the brain tissue. Keep the capillary precisely in between the hemispheres, remaining inside the ventricle just after piercing the dorsal ependymal layer. 
	NOTE: This is a very delicate step. Accuracy of this procedure is improved using pigmentation mutant lines such as brassy24, allowing better visualization of glass capillary position during injection.</li>
	<li>With the capillary tip inside of the ventricle, inject the plasmid solution by applying pressure with the foot pedal for about 10 s, which corresponds to approximately 1 &#181;L of plasmid solution. 
	NOTE: If changing the needle puller, capillaries or injector, the system should be calibrated in order to always deliver 1 &#181;L of plasmid solution. Calibration can be performed by measuring the diameter of the plasmid droplet expelled into a mineral oil (e.g., paraffin oil) and subsequently calculating the volume of the droplet. After 10 s of injection, there should be ~1 &#181;L of plasmid liquid expelled into the mineral oil.</li>
	<li>Confirm success of the previous step by observing the spread of liquid throughout the ventricle.</li>
</ol>
5. Electroporation
<ol>
	<li>Remove the fish from the injection set-up while still holding it in the sponge.</li>
	<li>Immerse the inner side of the tip of the electrodes in the ultrasound gel.</li>
	<li>Cover the fish telencephalon with a small amount of ultrasound gel.</li>
	<li>Position the fish head between the electrodes, placing the positive electrode at the ventral side of the fish&#8217;s head and the negative electrode on the dorsal side (Figure 1C), while still holding the fish&#8217;s body in the sponge. This sets the direction of the flow of the current necessary to electroporate ependymoglia positioned at subventricular zone.</li>
	<li>Press the electrodes gently and precisely against the telencephalon (Figure 1C). Administer the current with the foot pedal. Hold the electrodes in place until all five pulses are finished.</li>
</ol>
6. Fish Recovery
<ol>
	<li>Let the fish recover in the previously prepared, continuously aerated tank until it wakes up. Lidocaine gel could be applied on the skull in order to relieve any possibly developed pain.</li>
</ol>

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Authors have nothing to disclose.

This electroporation protocol is a reliable in vivo method of labelling individual ependymoglial cells. The protocol may need a further adaptation to label other cell types such as neurons or oligodendrocytes. To achieve successful labelling, plasmids containing different promoters can be used. Chicken-beta actin promoter, eF1alpha, CMV and ubiquitin promoter have been previously used to drive the expression of different transgenes in ependymoglia and their progeny23. However, different kinetics o...

Zebrafish are excellent animal models to examine brain regeneration after a stab wound injury. In comparison to mammals, on the evolutionary ladder, less evolved species such as zebrafish generally show higher rates of constitutive neurogenesis and broader areas of adult neural stem cell residence, leading to constant generation of new neurons throughout most brain areas in the adult life. This feature appears to correlate with significantly higher regenerative capacity of zebrafish in comparison to mammals1, as zebrafish have remarkable potential to efficiently generate new neurons in most brain injury models studied2,3,4,5,6,7,8. Here, the zebrafish telencephalon is studied, since it is a brain area with prominent neurogenesis in adulthood. These zones of adult neurogenesis are homologous to neurogenic zones in the adult mammalian brain9,10,11.
Abundant neurogenic areas in the zebrafish telencephalon are present due to the existence of radial glia like cells or ependymoglia cells. Ependymoglial cells act as resident adult neural stem cells and are responsible for generation of new neurons in both the intact and regenerating brain3,5. Lineage tracing experiments have shown that ventricular ependymoglia react to injury, then proliferate and generate new neuroblasts that migrate to the lesion site5. Due to the everted nature of zebrafish telencephalon, ependymoglial cells line the ventricular surface and build the ventral ventricular wall. The dorsal ventricular wall is formed by a dorsal ependymal cell layer (Figure 1A). Importantly, zebrafish ependymoglia embody the characteristics of both mammalian radial glia and ependymal cells. Long radial processes are a typical feature of radial glia cells, whereas cellular extensions and tight junctions (as well as their ventricular positions) are typical features of ependymal cells12,13,14. Therefore, these cells are referred to as ependymoglial cells.
To follow in vivo behavior of single ependymoglial cells during regeneration, they need to be reliably labeled. Various methods of in vivo cell labeling for fluorescent microscopy have been previously described, such as endogenous reporters or lipophilic dyes15. These methods, in contrast to electroporation, may require longer periods of time and often do not offer the possibility of single cell labeling or permanent long-term tracing. Electroporation, however (besides single cell labeling), offers the possibility of introducing new DNA into the host cell. Moreover, compared to other methods of DNA transfer into the cells, electroporation has been demonstrated to be one of the most efficient methods16,17,18,19.
Presented here is an electroporation protocol that has been refined for the purpose of labeling single ependymoglial cells in the adult zebrafish telencephalon. This protocol allows for the labelling of single ependymoglial cells in order to follow them over a long-term period20 or to manipulate specific pathways in a cell-autonomous manner21,22.

<table>
	<tbody>
		<tr>
			<td>Reagent/Material</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fast Green</td>
			<td>Sigma-Aldrich</td>
			<td>F7258-25G</td>
			<td>For coloring plasmid solution</td>
		</tr>
		<tr>
			<td>MS222</td>
			<td>Sigma-Aldrich</td>
			<td>A5040-25G</td>
			<td>MS222 should be stored at RT (up to two weeks) and protected from light</td>
		</tr>
		<tr>
			<td>Ultrasound gel</td>
			<td>SignaGel, Parker laboratories INC.</td>
			<td>15-60</td>
			<td>Electrode Gel</td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Air pump</td>
			<td>TetraTec APS 50, 10l-60l</td>
			<td></td>
			<td>Can be bought in the pet shops</td>
		</tr>
		<tr>
			<td>BTX Tweezertrodes Electrodes</td>
			<td>Platinum Tweezertrode, BTX Harvard Apparatus</td>
			<td>45-0486</td>
			<td>1mm diameter</td>
		</tr>
		<tr>
			<td>Electroporation device</td>
			<td>BTX ECM830 Square Wave Electroporation System, BTX Harvard Apparatus</td>
			<td>45-0662</td>
			<td></td>
		</tr>
		<tr>
			<td>Injection device</td>
			<td>FemtoJet 4i, Eppendorf</td>
			<td>5252000013</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard Wall Borosillicate Glass Capillary</td>
			<td>Warner Instruments</td>
			<td>64-0766</td>
			<td>Model No: G100-4</td>
		</tr>
		<tr>
			<td>Microloader tips</td>
			<td>Eppendorf</td>
			<td>5242956003</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-knife</td>
			<td>Fine Science Tools</td>
			<td>10056-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Joystick micromanipulator</td>
			<td>Narishige Japan</td>
			<td>MN - 151</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle holder</td>
			<td>FemtoJet 4i, Eppendorf</td>
			<td>5252000013</td>
			<td>Needle holder comes together with the injection device</td>
		</tr>
		<tr>
			<td>Needle pulling device</td>
			<td>Narishige Japan</td>
			<td>Model No: PC-10</td>
			<td>The PC-10 was discontinued by Narishige in 2017 and replaced by the PC-100</td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Greiner Bio-One International</td>
			<td>633161</td>
			<td></td>
		</tr>
	</tbody>
</table>

electroporation method for in vivo delivery of plasmid dna in the adult zebrafish telencephalon

Electroporation is a transfection method in which an electrical field is applied to cells to create temporary pores in a cell membrane and increase its permeability, thereby allowing different molecules to be introduced to the cell. In this paper, electroporation is used to introduce plasmids to ependymoglial cells, which line the ventricular zone of the adult zebrafish telencephalon. A fraction of these cells shows stem cell properties and generates new neurons in the zebrafish brain; therefore, studying their behavior is essential to determine their roles in neurogenesis and regeneration. The introduction of plasmids via electroporation enables long-term labeling and tracking of a single ependymoglial cell. Furthermore, plasmids such as Cre recombinase or Cas9 can be delivered to single ependymoglial cells, which enables gene recombination or gene editing and provides a unique opportunity to assess the cell&#8217;s autonomous gene function in a controlled, natural environment. Finally, this detailed, step-by-step electroporation protocol is used to obtain successful introduction of plasmids into a large number of single ependymoglial cells.

The described electroporation method allows delivery of plasmid DNA into ependymoglial cells, which are located superficially in the zebrafish telencephalon and just under the dorsal ependymal cell layer (Figure 1A).
If the result of electroporation is positive, labeled single ependymoglial cells (red cells in Figure 2A,B) can be observed among other ependymoglial cells (white in <s...

Watch this Scientific Journal Video about Electroporation Method for In Vivo Delivery of Plasmid DNA in the Adult Zebrafish Telencephalon at JoVE.com

Electroporation Method for In Vivo Delivery of Plasmid DNA in the Adult Zebrafish Telencephalon

Presented here is an electroporation method for plasmid DNA delivery and ependymoglial cell labeling in the adult zebrafish telencephalon. This protocol is a quick and efficient method to visualize and trace individual ependymoglial cells and opens up new possibilities to apply electroporation to a broad range of genetic manipulations.

Electroporation is a transfection method in which an electrical field is applied to cells to create temporary pores in a cell membrane and increase its ...

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JoVE Journal

Developmental Biology

9.6K Views.  Helmholtz Zentrum München. Presented here is an electroporation method for plasmid DNA delivery and ependymoglial cell labeling in the adult zebrafish telencephalon. This protocol is a quick and efficient method to visualize and trace individual ependymoglial cells and opens up new possibilities to apply electroporation to a broad range of genetic manipulations.

Video: Electroporation Method for In Vivo Delivery of Plasmid DNA in the Adult Zebrafish Telencephalon

Kidney Regeneration in Adult Zebrafish by Gentamicin Induced Injury

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is the nephron. Mammals are able to repair existing nephrons after injury, but lose the ability to form new nephrons soon after birth. In contrast to mammals, adult fish produce new nephrons (neonephrogenesis) throughout their lives in response to growth requirements or injury. Recently, lhx1a has been shown to mark nephron progenitor cells in the adult zebrafish kidney, however mechanisms controlling the formation of new nephrons after injury remain unknown. Here we show our method for robust and reproducible injury in the adult zebrafish kidney by intraperitoneal (i.p.) injection of gentamicin, which uses a noninvasive visual screening process to select for fish with strong but nonlethal injury. Using this method, we can determine optimal gentamicin dosages for injury and go on to demonstrate the effect of higher temperatures on kidney regeneration in zebrafish.

Here we present a reliable method to study adult kidney regeneration by inducing acute kidney injury by gentamicin injection. We show that injury is dependent on gentamicin dosage and environmental temperature using in situ hybridization to label lhx1a+ developing new nephrons.

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

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. ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

In Ovo Electroporation in the Chicken Auditory Brainstem

Electroporation is a method that introduces genes of interest into biologically relevant organisms like the chicken embryo. It is long established that the chicken embryo is an effective research model for studying basic biological functions of auditory system development. More recently, the chicken embryo has become particularly valuable in studying gene expression, regulation and function associated with hearing. In ovo electroporation can be used to target auditory brainstem regions responsible for highly specialized auditory functions. These regions include the chicken nucleus magnocellularis (NM) and nucleus laminaris (NL). NM and NL neurons arise from distinct precursors of rhombomeres 5 and 6 (R5/R6). Here, we present in ovo electroporation of plasmid-encoded genes to study gene-related properties in these regions. We show a method for spatial and temporal control of gene expression that promote either gain or loss of functional phenotypes. By targeting auditory neural progenitor regions associated with R5/R6, we show plasmid transfection in NM and NL. Temporal regulation of gene expression can be achieved by adopting a tet-on vector system. This is a drug inducible procedure that expresses the genes of interest in the presence of doxycycline (Dox). The in ovo electroporation technique &#8211; together with either biochemical, pharmacological, and or in vivo functional assays &#8211; provides an innovative approach to study auditory neuron development and associated pathophysiological phenomena.

In Ovo Electroporation in the Chicken Auditory Brainstem

Auditory brainstem neurons of avians and mammals are specialized for fast neural encoding, a fundamental process for normal hearing functions. These neurons arise from distinct precursors of embryonic hindbrain. We present techniques utilizing electroporation to express genes in the hindbrain of chicken embryos to study gene function during auditory development.

Electroporation is a method that introduces genes of interest into biologically relevant organisms like the chicken embryo. It is long established that ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain and protect cellular identities. The conversion of germ cells into specific neurons in the nematode Caenorhabditis elegans (C. elegans) is a powerful tool for performing genetic screens in order to dissect regulatory pathways that safeguard established cell identities. Reprogramming of germ cells to a specific type of neurons termed ASE requires transgenic animals that allow broad over-expression of the Zn-finger transcription factor (TF) CHE-1. Endogenous CHE-1 is expressed exclusively in two head neurons and is required to specify the glutamatergic ASE neurons fate, which can easily be visualized by the gcy-5prom::gfp reporter. A trans gene containing the heat-shock promoter-driven che-1 gene expression construct allows broad mis-expression of CHE-1 in the entire animal upon heat-shock treatment. The combination of RNAi against the chromatin-regulating factor LIN-53 and heat-shock-induced che-1 over-expression leads to reprogramming of germ cell into ASE neuron-like cells. We describe here the specific RNAi procedure and appropriate conditions for heat-shock treatment of transgenic animals in order to successfully induce germ cell to neuron conversion.

Application of RNAi and Heat-shock-induced Transcription Factor Expression to Reprogram Germ Cells to Neurons in C. elegans

This protocol describes how to study cellular processes during cell fate conversion in Caenorhabditis elegans in vivo. Using transgenic animals, allowing heat-shock promoter-driven overexpression of the neuron fate-inducing transcription factor CHE-1 and RNAi-mediated depletion of the chromatin-regulating factor LIN-53 germ cell to neuron reprogramming can be observed in vivo.

Studying the cell biological processes during converting the identities of specific cell types provides important insights into mechanism that maintain ...

Nanoparticle-mediated siRNA Gene-silencing in Adult Zebrafish Heart

Mammals have a very limited capacity to regenerate the heart after myocardial infarction. On the other hand, the adult zebrafish regenerates its heart after apex resection or cryoinjury, making it an important model organism for heart regeneration study. However, the lack of loss-of-function methods for adult organs has restricted insights into the mechanisms underlying heart regeneration. RNA interference via different delivery systems is a powerful tool for silencing genes in mammalian cells and model organisms. We have previously reported that siRNA-encapsulated nanoparticles successfully enter cells and result in a remarkable gene-specific knockdown in the regenerating adult zebrafish heart. Here, we present a simple, rapid, and efficient protocol for the dendrimer-mediated siRNA delivery and gene-silencing in the regenerating adult zebrafish heart. This method provides an alternative approach for determining gene functions in adult organs in zebrafish and can be extended to other model organisms as well.

It remains a major challenge to develop conditional gene-knockout or effective gene-knockdown in adult zebrafish organs. Here we report a protocol for performing nanoparticle-mediated siRNA gene-silencing in adult zebrafish heart, thus providing a new loss-of-function method for studying adult organs in zebrafish and other model organisms.

Mammals have a very limited capacity to regenerate the heart after myocardial infarction. On the other hand, the adult zebrafish regenerates its heart ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows within roughly two weeks after injury. Furthermore, zebrafish are able to heal bone rapidly after trepanation of the skull, and repair fractures that can be easily introduced into zebrafish bony fin rays. These injury assays represent feasible experimental paradigms to test the effect of administered drugs on rapidly forming bone. Here, we describe the use of these 3 injury models and their combined use with systemic glucocorticoid treatment, which exerts bone inhibitory and immunosuppressive effects. We provide a workflow on how to prepare for immunosuppressive treatment in adult zebrafish, illustrate how to perform fin amputation, trepanation of calvarial bones, and fin fractures, and describe how the use of glucocorticoids affects both bone forming osteoblasts and cells of the monocyte/macrophage lineage as part of innate immunity in bone tissue.

Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows ...