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 border="0" cellpadding="0" cellspacing="3">
	<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 &#216; 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 00</td>
			<td>Catgut 
			Markenkirchen</td>
			<td></td>
		</tr>
		<tr>
			<td>syringe, with 30G 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 ddH2O</td>
		</tr>
		<tr>
			<td>10x PBS, pH 7.4</td>
			<td>Carl Roth</td>
			<td>3957.1</td>
			<td>NaCl [1.37 M]</td>
		</tr>
		<tr>
			<td>6781.1</td>
			<td>KCl [27 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>T106.2</td>
			<td>Na2HPO4+2H2O [100 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>3904.1</td>
			<td>KH2PO4 [18 mM]</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10% Ketamin</td>
			<td>Serum Werk 
			Bernburg</td>
			<td>Urotamin</td>
			<td>mix in a rate 1:1</td>
		</tr>
		<tr>
			<td>2% Xylazin</td>
			<td>Serum Werk 
			Bernburg</td>
			<td>Xylazin</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterilium</td>
			<td>Bode Chemie</td>
			<td>Sterillium</td>
			<td>disinfection</td>
		</tr>
		<tr>
			<td>vet ointment</td>
			<td>S&#38;K Pharma 
			GmbH</td>
			<td>Kerato Biciron 5% 
			Augensalbe</td>
			<td>or similar to prevent 
			dryness of eyes</td>
		</tr>
		<tr>
			<td>To-Pro-3 iodide</td>
			<td>Invitrogen</td>
			<td>T3605</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

Research

JoVE Journal

Biology

27.6K 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.