We thank Birgit Westernstroeer of the Centre of Reproductive Medicine and Andrology at the University of Muenster for teaching the testicular microinjection. Besides, we are grateful to Ursula Antkewitz and Petra Reckling for technical assistance. We thank the German Research Foundation (DFG) for supporting this work (WE2458/10-1).&#160;

All performed animal experiments have been approved by the local ethics committee (Landesamt f&#252;r Landwirtschaft, Lebensmittelsicherheit und Fischerei, Mecklenburg-Vorpommern, Germany).
1. Plasmid Preparation
<ol>
	<li>For plasmid preparation, use plasmid purification kits (see Materials table) or similar methods with endotoxin removal buffer so that immune reactions of the animal can be avoided. Follow the instructions of the manual. Employ ddH2O to dilute the plasmid solution.</li>
	<li>Eliminate debris by spinning the DNA solution at maximum speed (20,000 x g). Then collect the supernatant.</li>
	<li>Determine plasmid concentration with a spectrophotometer (see materials).</li>
	<li>Adjust plasmid concentration with ddH2O to 1-3 &#181;g(DNA)/&#181;l.&#160;Note: Lower concentrations will reduce transfection efficiency, while higher concentrations might cause a too viscous injection solution. Besides, the transfection efficiency depends on the size of the plasmid and has to be tested individually.</li>
	<li>Prior to injection, prepare a solution mix with for example&#160;40 &#181;l plasmid together with 5 &#181;l PBS (10x) and 5 &#181;l Fast Green (0.5%). Fast Green is needed for tracking the injection process. Preferably, use thin wall PCR-tubes or Parafilm. Note:&#160;Depending of the size of the testis, a volume of 20-50 &#181;l will be needed for each testis.&#160;</li>
</ol>
2. Preparation of Microinjection Pipette
<ol>
	<li>Use borosilicate capillaries (see &#160;Materials table) for preparing the microinjection pipettes.</li>
	<li>Pull glass capillaries with a vertical capillary puller (see&#160;Materials table).</li>
	<li>Break the capillary tip with forceps by softly banding. The tip is usually 50-80 &#181;m in diameter and about 1 cm long. Try to avoid tips that are too long as these are not stiff enough to penetrate the tissue.</li>
	<li>At last, sharpen the tip in a 30&#176; to 45&#176; angle by using a micropipette beveler (see&#160;Materials table).</li>
	<li>Load the microinjection pipette with the injection solution mix (see Plasmid Preparation). Apply the solution into the back of the glass capillary with a small syringe. Note: Try to avoid air bubbles while loading, otherwise these will be injected into the seminiferous tubules along with the plasmid solution and hence will reduce conductivity as well as possibly cause tissue damage.</li>
</ol>
3. Anesthesia and Surgery
<ol>
	<li>Perform the operation under aseptic conditions by using sterile material (i.e., syringe, needle, surgical instruments, etc.). This will reduce infection of the animal and ensure good survival rates.</li>
	<li>Use male mice for electroporation at an age of 6-8 weeks.</li>
	<li>To initialize postsurgical analgesic treatment, provide the mouse with analgesics added drinking water on the day prior to surgery. This assures a preemptive analgesia in case of highly probable postsurgical hypodipsia (diminished water intake). To this end, expose drinking water with for example a pediatric ibuprofen suspension (20 mg/ml). The medicated drinking water should also be supplied on day one and two of recovery in identical concentration. This means a daily dose of 7.5 mg/kg, valid for 5 ml/d drinking water and body weight of 25g for 8 weeks aged C57BL/6J. This analgesic regimen guarantees a fundamental pain relief and accelerated recovery along with high welfare26.</li>
	<li>For preparing&#160;the anesthesia working solution on the day of surgery, mix 10% Ketamin and 2% Xylazin in a 1:1 ratio (see&#160;Materials table).</li>
	<li>To anesthetize the animal, apply 0.25 ml/100 mg body weight of anesthetic solution subcutaneously (Ketamin = 0.125 g/kg; Xylazin = 0.025 g/kg). Use an injection site between pelvic and limb in order to prevent organ damage and injury of the mice. Usually, 10 min are needed until the mouse is deeply anesthetized.</li>
	<li>Cover the mouse eyes with vet ointment to prevent dryness during anesthesia.</li>
	<li>Test deep anesthetic arrest, which is noticeable by a total lack of response. To this end, just pinch the toe of the animal.</li>
	<li>To start the surgery, remove abdominal hair with an electric shaver or similar and disinfect the operation area with sterilium or similar.</li>
	<li>Make a ventral incision directly above the preputial glands in the center of the abdominal area. At that place, first pull the skin away and conduct a small transversal cutaneous cut of about 8-14 mm. Then, continue along the abdominal muscular layer. 
	Note: The lesion should be as small as possible to reduce harm to the animal.</li>
	<li>Pull up the abdominal fat pads carefully to expose the attached testis and place it on a prior prepared waterproof disposable paper drape just beside the incision site&#160;(Figure 2A).</li>
	<li>Use binoculars to find the efferent duct as the injection target. The ductus efferent is identified as the fine vessel junction between the testis and the epididymis. It is located adjacent to the prominent testicular artery. The ductus runs almost at an angle of 45&#176; to the artery and visibly enters the caput epididymidis. Note: Depending on the mouse&#8217;s age, the ductus is usually buried in fatty tissue.</li>
	<li>Use fine forceps to clear the efferent duct of this fatty tissue. After that, place the released ductus on a sterile&#160;paper strip to ensure the clear visibility of the duct without impairing surroundings.</li>
</ol>
4. Microinjection and DNA Application
<ol>
	<li>Connect the microinjection pipette, which is loaded with the plasmid solution to the micromanipulator/injector unit.</li>
	<li>Place the microinjection needle parallel to the efferent duct with the tip pointing towards the rete testis (Figure 2B).</li>
	<li>Use fine tweezers and strip the duct over the microinjection pipette. Make sure that the capillary is kept parallel to the duct while pulling it over. 
	Note: This procedure is more convenient than penetrating the vessel by moving the needle with the micromanipulator.</li>
	<li>Direct the needle carefully towards the testis and stop just as it penetrates the rete testis directly beneath the tunica albuginea (Figure 2C). 
	Note:&#160;In the case of overreaching the rete testis, the plasmid solution will leak into the interstitial space. This commonly leads to failure of transfecting the seminiferous tubules.</li>
	<li>For injection of the plasmid solution utilize the microinjector with following settings: pi:&#160;100 hPa, ti:&#160;0.2 sec, and pc:&#160;0 hPa (Figure 2D).</li>
	<li>Monitor the entire injection process by observing the testis filling status with the help of the green color. Take care that the testis is only filled up to 2/3 of its volume with the plasmid solution (Figure 2E). 
	Note:&#160;If the injection volume is exceeded, the testis tissue could be harmed.</li>
</ol>
5. Electroporation of Testis
<ol>
	<li>To enable effective electroporation, soak the tweezer electrodes in PBS (1x). This ensures adequate conductance.</li>
	<li>Smoothly squeeze the testis between the wet electrodes (Figure&#160;2F). Measure the electrical resistance with the electroporator.</li>
	<li>Apply current to the testis. Perform eight square pulses of 40 V at 4&#160;different testis sites with a constant duration of 50 msec pulse time and 950 msec interval time.</li>
</ol>
6. Wound Closure and Post-surgery
<ol>
	<li>When electroporation is finished, place the testis back in the original location.</li>
	<li>Sew the inner muscular layer with surgical suture (see materials).</li>
	<li>Close the skin by employing suture clips (see Materials table). Pull the skin up with tweezers. Make sure to exclude the muscular layer. Place the clips, each at a distance of not more than 5 mm. 
	Note: Because surgical suture can be swallowed by the mouse, suture clips are favored for closing the skin lesion.</li>
	<li>Allow the mouse to recover from anesthesia on sterile paper towels placed in a sterile empty mouse cage, which is tempered on a 37 &#176;C warm pad. The pad should ensure the warming of one half of the cage creating a heat gradient. For the animal this makes possible to search for the individual, most comfortable area due to the temperature variety in the cage.&#160;&#160;In addition, cover the mouse with a sheet of sterile paper towel so that stress can be further reduced. Note:&#160;Since the mouse&#8217;s body surface is uncommonly high in relation to body mass, when compared with larger animals, the thermal support is of particular importance for successful recovery of rodents.</li>
	<li>When the operated animal has fully awakened, transfer it for further recovery to a completely equipped new mouse cage. Do not put the animal back to its old cage or to a group of mice. Ensure that the cage is as clean and sterile as possible to prevent wound infection. Monitor the entire recovery process. To transfer the mouse back to the animal care facility, wait until it has fully recovered and appears to behave normally. 
	Note: Additional per- and post-surgical strategies are discussed by Pritchett-Corning KR et al6.</li>
</ol>

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Marten Michaelis, Alexander Sobczak, and Joachim M. Weitzel employed at the Institute of Reproductive Biology, Leibniz Institute for Farm Animal Biology (FBN), Dummerstorf, Germany, declare that they have no competing financial interests.

Research in the field of reproductive biology, particularly in the area of male fertility and spermatogenesis inevitably relies on living organisms. In order to examine testicular function, no adequate cell culture/in vitro system has been established capable of reflecting all the crucial morphological changes from a diploid spermatogonium to a haploid mature spermatozoon1,2. Thus, the generation of genetically modified animals is often a necessary and as such a valuable tool in male ...

Mammalian spermatogenesis is considered to be a sophisticated process of self-renewing stem cells successively undergoing mitosis, meiosis and differentiation in order to develop into mature haploid spermatozoa. These morphological changes are orchestrated by different cell types and despite profound attempts, it is still impossible to mimic these processes in cell culture1,2. Hence, research on spermatogenesis up to now relies on living organisms as in vivo models. In general, gene function studies are usually based on transgenic animals. However, generating and sustaining this kind of animal model is time-consuming, cost-intensive and quite elaborate. This is attributed to the required long breeding process for generating and maintaining the transgene over the generations. Additionally, the genetic manipulation of the entire organism by the transgenic or knockout approach is prone to cause physiological impairments when targeting genes with essential functions in multiple regions, e.g., outside the testis or systemically.
Further, some transient transfection methods are associated with some crucial disadvantages. For example, typical drawbacks of virus-mediated gene transfer are the possible provocation of immunoreactions and additional safety regulations, whereas lipofection3 and microparticle bombardment4 might damage the tissue and are limited to a certain cell depth for sufficient transfection efficiency.
In contrast, electroporation (EP) as another common way of transient transfection, seems to constitute a promising technique for enabling in vivo transfection and consistent in vivo gene analysis. In general, EP is referred to as a dynamic phenomenon that depends on local transmembrane voltage with consequently mediated pores within nanoseconds. These gaps can be maintained for milliseconds, sufficient to grant access to DNA, RNA or small molecules5. When the applied voltage&#160;is too high, the usually transient character of EP is counteracted due to heat production and induction of too comprehensive permeabilization with consequent irreversible damage of the cell5.
Here, we show that electroporation is an effective and economical transfection system which is capable of being utilized for genetic testis transformation in order to elucidate testicular gene characteristics in vivo. This article addresses plasmid preparation, microinjection via the efferent duct and the subsequent electroporation of mouse testis. This procedure can be the means of choice to achieve fast, specific and efficient transfection of seminiferous tubules of mouse testis in vivo in order to investigate processes of spermatogenesis.

<table><tbody><tr><td>Centrifuge</td><td>Sigma</td><td>1-15 PK</td><td></td></tr><tr><td>Spectrophotometer</td><td>Nanodrop</td><td>ND-1000</td><td></td></tr><tr><td>Micropipette puller</td><td>Narishige</td><td>PC-10</td><td>vertical capillary puller</td></tr><tr><td>Microinjection capillaries</td><td>Clark</td><td>GC100-10</td><td>borosilicate standard wall</td></tr><tr><td>Micropipette beveler</td><td>Bachofer</td><td>Typ 462</td><td>rotating disk beveler</td></tr><tr><td>Electroporator</td><td>Nepagene</td><td>CUY 21EDIT</td><td>square wave electroporator</td></tr><tr><td>Tweezer electrodes</td><td>Nepagene</td><td>CUY 650P5</td><td>5 mm &amp;Oslash; disk electrodes</td></tr><tr><td>Stereo microscope</td><td>Zeiss</td><td>Stemi 2000-C</td><td></td></tr><tr><td>Cold light source</td><td>Zeiss</td><td>KL 1500LCD</td><td></td></tr><tr><td>Microinjection pump</td><td>Eppendorf</td><td>Femtojet</td><td></td></tr><tr><td>Micromanipulator</td><td>homemade</td><td></td><td>XYZ cross table</td></tr><tr><td>Surgical instruments</td><td>FST</td><td></td><td>scissors, forceps, needle holder</td></tr><tr><td>Fine forceps</td><td>FST</td><td>Dumont #5, #7</td><td>efferent ducts preparation</td></tr><tr><td>Michel clip applying stapler</td><td>FST</td><td>Michel, 12029-12</td><td></td></tr><tr><td>Michel suture clips</td><td>FST</td><td>Michel, 12040-01</td><td>7.5 x 1.75 mm</td></tr><tr><td>Surgical suture</td><td>Catgut, Markneukirchen, Germany</td><td>Catgut 00</td><td></td></tr><tr><td>Syringe with 30 G needle</td><td>B. Braun</td><td>Omnican 40</td><td>to load micropipette and for anesthesia</td></tr><tr><td>Plasmid isolation kit</td><td>Promega</td><td>Cat. # A2495</td><td>plasmid Midiprep</td></tr><tr><td>Plasmid pEGFP-C1</td><td>Clontech</td><td>Cat. #6084-1</td><td>CMV-promoter + EGFP</td></tr><tr><td>Plasmid pDsRed2-N1</td><td>Clontech</td><td>Cat. #6084-1</td><td>CMV-promoter + DsRed2</td></tr><tr><td>Fast Green dye</td><td>Sigma</td><td>F7258-25G</td><td>for dilution in ddH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>10% Ketamine</td><td>Serum Werk, Bernburg, Germany</td><td>Urotamin</td><td>mix in a rate 1:1 with xylazine</td></tr><tr><td>2% Xylazine</td><td>Serum Werk, Bernburg, Germany</td><td>Xylazin</td><td>mix in a rate 1:1 with ketamine</td></tr><tr><td>Sterilium</td><td>Bode Chemie</td><td>Sterillium</td><td>disinfection</td></tr><tr><td>Vet ointment</td><td>S&amp;amp;K Pharma, GmbH</td><td>Kerato Biciron 5%, Augensalbe</td><td>opthalmic ointment to prevent eye dryness</td></tr><tr><td>To-Pro-3 iodide</td><td>Invitrogen</td><td>T3605</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;10x PBS, pH 7.4&lt;/strong&gt;</td></tr><tr><td>1.37 M NaCl</td><td>Carl Roth</td><td>3957.1</td><td></td></tr><tr><td>Ibuprofen, Dolormin</td><td>Johnson &amp;amp; Johnson Consumer Health Care Germany</td><td>01094902</td><td>analgesic pediatric solution (NSAID) for postsurgery pain relief</td></tr><tr><td>27 mM KCl</td><td>Carl Roth</td><td>6781.1</td><td></td></tr><tr><td>100 mM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Carl Roth</td><td>T106.2</td><td></td></tr><tr><td>18 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Carl Roth</td><td>3904.1</td><td></td></tr></tbody></table>

in vivo microinjection and electroporation of mouse testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

The experimental setting for performing microinjection and electroporation of mouse testis in vivo as it is used according to the protocol is illustrated in Figure 1. Even though it is possible to acquire industrially manufactured micropipettes, we preferred to generate our own pipettes by pulling (Figure 1A) and beveling (Figure 1B) glass capillaries so that they fitted our needs. The equipment for microinjection and electroporation is illustrated in Fi...

Watch this Scientific Journal Video about In Vivo Microinjection and Electroporation of Mouse Testis at JoVE.com

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

in-vivo-microinjection-and-electroporation-of-mouse-testis

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

27.8K Views.  Leibniz Institute for Farm Animal Biology (FBN). This article describes microinjection and electroporation of mouse testis in vivo as a transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

Video: In Vivo Microinjection and Electroporation of Mouse Testis

In Utero Intraventricular Injection and Electroporation of E15 Mouse Embryos

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors lining the walls of the lateral ventricle. This technique is now widely used to study the processes involved in corticogenesis by over-expressing or knocking down genes and observing the effects on cellular proliferation, migration, and differentiation. In comparison to traditional knockout strategies, in-utero electroporation provides a rapid means to manipulate a population of cells during a specific temporal window. In this video protocol we outline the experimental methodology for preparing mice for surgery, exposing the uterine horns through laporatomy, injecting DNA into the lateral ventricles of the developing embryo, electroporating DNA into the progenitors lining the lateral wall, and caring for animals post-surgery. Our laboratory uses this protocol for surgeries on E13-E16 mice, however, it is most commonly performed at E15, as shown in this video. 

In-utero in-vivo injection and electroporation of the embryonic mouse neocortex provides a powerful tool for the manipulation of individual progenitors ...

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 ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

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

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

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

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

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

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

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

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

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

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

Germline genetically engineered mouse models (G-GEMMs) have provided valuable insight into in vivo gene function in development, homeostasis, and disease. However, the time and cost associated with colony creation and maintenance are high. Recent advances in CRISPR-mediated genome editing have allowed the generation of somatic GEMMs (S-GEMMs) by directly targeting the cell/tissue/organ of interest.
The oviduct, or fallopian tube in humans, is considered the tissue-of-origin of the most common ovarian cancer, high-grade serous ovarian carcinomas (HGSCs). HGSCs initiate in the region of the fallopian tube distal to the uterus, located adjacent to the ovary, but not the proximal fallopian tube. However, traditional mouse models of HGSC target the entire oviduct, and thus do not recapitulate the human condition. We present a method of DNA, RNA, or ribonucleoprotein (RNP) solution microinjection into the oviduct lumen and in vivo electroporation to target mucosal epithelial cells in restricted regions along the oviduct. There are several advantages of this method for cancer modeling, such as 1) high adaptability in targeting the area/tissue/organ and region of electroporation, 2) high flexibility in targeted cell types (cellular pliancy) when used in combination with specific promoters for Cas9 expression, 3) high flexibility in the number of electroporated cells (relatively low frequency), 4) no specific mouse line is required (immunocompetent disease modeling), 5) high flexibility in gene mutation combination, and 6) possibility of tracking electroporated cells when used in combination with a Cre reporter line. Thus, this cost-effective method recapitulates human cancer initiation.

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

This protocol describes microinjection and in vivo electroporation for regionally restricted CRISPR-mediated genome editing in the mouse oviduct.

Germline genetically engineered mouse models (G-GEMMs) have provided valuable insight into in vivo gene function in development, homeostasis, and disease. ...

Cancer Research

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies showed that microinjection of germ cells from fertile donor mice into the seminiferous tubules of infertile recipient mice results in donor-derived spermatogenesis and sperm production by the recipient animal2. The use of donor males carrying the bacterial &beta;-galactosidase gene allowed identification of donor-derived spermatogenesis and transmission of the donor haplotype to the offspring by recipient animals1. Surprisingly, after transplantation into the lumen of the seminiferous tubules, transplanted germ cells were able to move from the luminal compartment to the basement membrane where spermatogonia are located3. It is generally accepted that only SSCs are able to colonize the niche and re-establish spermatogenesis in the recipient testis. Therefore, germ cell transplantation provides a functional approach to study the stem cell niche in the testis and to characterize putative spermatogonial stem cells. To date, germ cell transplantation is used to elucidate basic stem cell biology, to produce transgenic animals through genetic manipulation of germ cells prior to transplantation4,5, to study Sertoli cell-germ cell interaction6,7, SSC homing and colonization3,8, as well as SSC self-renewal and differentiation9,10.
Germ cell transplantation is also feasible in large species11. In these, the main applications are preservation of fertility, dissemination of elite genetics in animal populations, and generation of transgenic animals as the study of spermatogenesis and SSC biology with this technique is logistically more difficult and expensive than in rodents. Transplantation of germ cells from large species into the seminiferous tubules of mice results in colonization of donor cells and spermatogonial expansion, but not in their full differentiation presumably due to incompatibility of the recipient somatic cell compartment with the germ cells from phylogenetically distant species12. An alternative approach is transplantation of germ cells from large species together with their surrounding somatic compartment. We first reported in 2002, that small fragments of testis tissue from immature males transplanted under the dorsal skin of immunodeficient mice are able to survive and undergo full development with the production of fertilization competent sperm13. Since then testis tissue xenografting has been shown to be successful in many species and emerged as a valuable alternative to study testis development and spermatogenesis of large animals in mice14.

Protocols for germ cell transplantation and testis tissue xenografting are described. Germ cell transplantation results in donor-derived spermatogenesis in recipient testes and represents a functional reconstitution assay for identification of spermatogonial stem cells (SSCs). Testis tissue xenografting reproduces testis development and spermatogenesis of various donor species in recipient mice.

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies ...

Novel Method of Plasmid DNA Delivery to Mouse Bladder Urothelium by Electroporation

Genetically engineered mouse models (GEMMs) are extremely valuable in revealing novel biological insights into the initiation and progression mechanisms of human diseases such as cancer. Transgenic and conditional knockout mice have been frequently used for gene overexpression or ablation in specific tissues or cell types in vivo. However, generating germline mouse models can be time-consuming and costly. Recent advancements in gene editing technologies and the feasibility of delivering DNA plasmids by viral infection have enabled rapid generation of non-germline autochthonous mouse cancer models for several organs. The bladder is an organ that has been difficult for viral vectors to access, due to the presence of a glycosaminoglycan layer covering the urothelium. Here, we describe a novel method developed in lab for efficient delivery of DNA plasmids into the mouse bladder urothelium in vivo. Through intravesical instillation of pCAG-GFP DNA plasmid and electroporation of surgically exposed bladder, we show that the DNA plasmid can be delivered specifically into the bladder urothelial cells for transient expression. Our method provides a fast and convenient way for overexpression and knockdown of genes in the mouse bladder, and can be applied to building GEMMs of bladder cancer and other urological diseases.

We describe a novel method for the delivery of DNA plasmid into the urothelial cells of mouse bladder in vivo through urethra catheterization and electroporation. It offers a fast and convenient way for generating autochthonous mouse models of bladder diseases.

Genetically engineered mouse models (GEMMs) are extremely valuable in revealing novel biological insights into the initiation and progression mechanisms ...

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry

In eukaryotes, meiosis is essential for genome stability and genetic diversity in sexual reproduction. Experimental analyses of spermatocytes in testes are critical for the investigations of spindle assembly and chromosome segregation in male meiotic division. The mouse spermatocyte is an ideal model for mechanistic studies of meiosis, however, the effective methods for the analyses of spermatocytes are lacking. In this article, a practical and efficient method for the in vivo inhibition of kinesin-7 CENP-E in mouse spermatocytes is reported. A detailed procedure for testicular injection of a specific inhibitor GSK923295 through abdominal surgery in 3-week-old mice is presented. Furthermore, described here is a series of protocols for tissue collection and fixation, hematoxylin-eosin staining, immunofluorescence, flow cytometry and transmission electron microscopy. Here we present an in vivo inhibition model via abdominal surgery and testicular injection, that could be a powerful technique to study male meiosis. We also demonstrate that CENP-E inhibition results in chromosome misalignment and metaphase arrest in primary spermatocytes during meiosis I. Our in vivo inhibition method will facilitate mechanistic studies of meiosis, serve as a useful method for genetic modifications of male germ lines, and shed a light on future clinical applications.

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry

This article reports an in vivo inhibition of CENP-E through abdominal surgery and testicular injection of GSK923295, a valuable model for male meiotic division. Using the immunofluorescence, flow cytometry and transmission electron microscopy assays, we show that CENP-E inhibition results in chromosome misalignment and genome instability in mouse spermatocytes.

In eukaryotes, meiosis is essential for genome stability and genetic diversity in sexual reproduction. Experimental analyses of spermatocytes in testes ...

In Vivo Microinjection and Electroporation of Mouse Testis

Summary

Explore More Videos

Chapters in this video

In Utero Intraventricular Injection and Electroporation of E15 Mouse Embryos

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

In vivo Electroporation of Developing Mouse Retina

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

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

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Novel Method of Plasmid DNA Delivery to Mouse Bladder Urothelium by Electroporation

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry