This work was supported by grants from the Ministry of Education, Culture, Sports, Science and Technology to M.I. (21229004 and 25221304).

The use of rats for research was in accordance with guidelines established by the Animal Welfare Committee of The University of Tokyo.
1.Preparation of Plasmid DNA
<ol>
	<li>Generate DNA plasmids for in vivo electroporation by subcloning the cDNA or shRNA sequence into an expression plasmid for mammalian cells12. Use a cytomegalovirus immediate early enhancer and chicken &#946;-actin promoter fusion (CAG) promoter-driven plasmid13 because it allows strong and stable expression. For expression of shRNAs under the control of a CAG promoter, use a mir30-based shRNA cassette system for subcloning the shRNA14.</li>
	<li>Purify plasmid DNA with a maxi-prep kit according to manufacturer&#39;s instructions, and resuspend the DNA with HEPES-buffered saline (140 mM NaCl, 0.75 mM Na2HPO4, 25 mM HEPES; pH 7.40). Adjust the concentration of DNA to &#8805;4 &#181;g/&#181;l.</li>
	<li>Prepare the plasmid DNA solution to a concentration of 4 &#181;g/&#181;l, and add a minimal amount of fast green dye (final concentration of 0.01%) to label the injection site. When simultaneous electroporation of multiple plasmids is required, adjust the total concentration of the plasmid DNA solution to 4 &#181;g/&#181;l. 
	Note: The optimal composition of plasmid DNAs should be determined according to the transfection efficiency of each plasmid.</li>
</ol>
2. Sterilization of Surgical Instruments and Saline
<ol>
	<li>Autoclave surgical instruments and the 0.9% NaCl solution.</li>
</ol>
3. Preparation of the Glass Micropipette
<ol>
	<li>Pull glass pipettes using a pipette puller. Cut the tip of the pipette to a diameter of 30-50 &#181;m. Use the following parameters: Heat, 600; Velocity, 50; Time, 75.</li>
</ol>
4. Animal Surgery, DNA Injection and Electroporation
Note: An overview of this step is described in Figure 1. Although the procedure for rat pups are described here, the method is also applicable to more mature animals using the same procedure.
<ol>
	<li>Anesthetize the rat with isoflurane in the induction box until the animal becomes immobile by adjusting the oxygen flow to 0.4 L/min and the isoflurane concentration to 4% (vol/vol). Perform toe pinching to confirm proper anesthetization.</li>
	<li>Put the rat on the preheated warmer under a binocular microscope, and maintain anesthesia by continuously administering isoflurane through the face mask. Adjust the oxygen flow to 0.2 L/min and the isoflurane concentration to 2% (vol/vol). Use eye drops to prevent dryness of eyes if the eyes of the animal are open.</li>
	<li>Fix the legs with surgical tape.</li>
	<li>Clean the skin on the posterior thigh with povidone-iodine, and make an incision with a scalpel. 
	Note: Shave surgical areas if the surgical areas are covered with hair.</li>
	<li>Expose the sciatic nerve by creating an opening between the quadriceps femoris muscle and biceps femoris muscle with sewing needles.</li>
	<li>Wet the nerve with 0.9% NaCl solution. Absorb excess water with lint-free paper.</li>
	<li>Insert the base of a glass micropipette onto flexible tubing, and fill the adequate amount of DNA solution (at least one microliter) into the micropipette by gently aspirating.</li>
	<li>Lift the exposed nerve by gently pulling the distal side of the nerve using a needle. 
	Note: Do not apply tension to the nerve to minimize mechanical stress.</li>
	<li>Insert the glass micropipette into the distal site on the nerve, and inject the DNA solution by applying pressure (i.e. by blowing into the open end of the flexible tube). Inject DNA solution until the nerve appears green (1 &#181;l maximum). Because frequent insertion of the micropipette can damage the nerve, do not insert the micropipette more than twice.</li>
	<li>Place a tweezer-type platinum electrode about 1-2 mm apart from the nerve. Fill the gap between the electrode and the nerve with 0.9% NaCl solution. 
	Note: Do not hold the nerve with the electrode to avoid mechanical stress on the nerve.</li>
	<li>Apply electric pulses to the injection site using an electroporator with the electrode. After the first pulse set, invert the electrode and apply another pulse set. Use the following parameters: voltage, 50 V; pulse duration, 5 msec; pulse interval, 100 msec; pulse number, 4 times.</li>
	<li>Clean the electroporation site with 0.9% NaCl solution.</li>
	<li>Repeat steps 4.4-4.11 on the contralateral sciatic nerve.</li>
</ol>
5. Post-electroporation
<ol>
	<li>Close the incisions with cyanoacrylate glue.</li>
	<li>After drying the glue, clean the wound with povidone-iodine.</li>
	<li>Release the pup from the face mask. Warm the pup on a&#160;warmer at least for an hour in order to allow it to fully recover from anesthesia. Do not leave the pup unattended until it has regained sufficient consciousness.</li>
	<li>After recovery from anesthesia, return the pup to the mother rat. Do not return the pup until fully recovered.</li>
</ol>
6. Post-surgery
<ol>
	<li>House the rat pups in the cage until conducting the experiments11 (see examples in Figure 3). Administer carprofen (5 mg/kg; i.p.), a non-steroidal anti-inflammatory drug, or buprenorphine (0.1 mg/kg, s.c.), an opioid analgesic, if required. 
	Note: If the rat pup does not grow well or inflammation is observed around the surgery site, exclude the animal from the experiments.</li>
</ol>

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The authors declare that they have no competing financial interests.

In this paper, we describe a simple and efficient method that allows in vivo gene transfer to myelinating SCs in the rat sciatic nerve using electroporation. This method allows highly selective gene expression in myelinating SCs by simply applying electric pulses to the plasmid DNA-injected sciatic nerve. Because the molecular mechanisms of myelination and demyelination in the peripheral nervous system remain unclear, the present in vivo electroporation method will be a powerful tool to clarify the roles of multiple genes of interest in living animals.
A critical requirement of this method is to keep damage to the nerve during surgery to a minimal level. Should surgical damage cause excessive inflammation, the sciatic nerve may degenerate. To avoid this, one must conduct surgery with extreme care, so as to not damage the blood vessels around the nerve. Mechanical stress to the nerve during the surgery can also be a cause of nerve damage. To minimize mechanical stress, lifting the exposed nerve should be done as gently as possible, and the tweezer-type electrode should be placed close to the nerve without contact. Furthermore, electrical pulses that are too strong can cause undesirable large leg movement, which leads to mechanical stress, or can burn the nerve. If significant damages are observed in the nerves, we recommend reducing the electrical pulse intensities or placing the electrode further away from the nerve.
In our present protocol, CAG promoter-driven plasmids were used as expression vectors. CAG promoter-driven plasmids allow high levels of gene expression in myelinating SCs in vivo. We also have tried a CMV promoter, another widely used universal promoter for mammalian gene expression, but expression of the gene product was very weak. This is consistent with previous results, in which electroporation-mediated transfection was conducted in the embryonic brain20. Therefore, we recommend using CAG promoter-driven plasmids for the in vivo electroporation method.
Because axonal signaling is a key factor in myelination/demyelination21, gene modification in neurons is also important. However, delivery of transgenes using our in vivo electroporation method is limited to SCs. It has been reported that gene delivery into sciatic nerve axons can be achieved when in vivo electroporation is applied to dorsal root ganglion (DRG) neurons in adult rats22. This suggests that delivery of plasmid DNA to the cell body is likely to be critical for in vivo transfection of peripheral axons. Thus, to examine the involvement of axonal molecules in myelination/demyelination, researchers should use neuron-specific genetic methods such as genetically modified animals, neuron-specific viral vectors, or in vivo electroporation to DRG neurons.
Compared with current methods, such as the generation of genetically modified animal lines23 and delivery of transgenes by viral vectors4-6, gene modification of SCs by in vivo electroporation is simpler. This method only requires several days for plasmid DNA construction and one day for electroporation surgery. Plasmid DNA construction does not require a biohazard room that is usually essential for viral vector handling. In addition, one of the advantages of the electroporation method is the capacity for simultaneous expression of multiple gene products using a simple protocol. Our novel technique will be useful for analyzing the interaction of a variety of signaling molecules involved in myelination and demyelination. In particular, by permitting the cotransfection of a number of different intracellular fluorescent probes, our method should be a powerful tool for investigating intracellular signaling dynamics in SCs using live imaging experiments.

The rapid transmission of sensory and motor information in the peripheral nervous system is permitted by the myelin sheath, which is formed by myelinating Schwann cells (SCs)1. Insulation of axons by the myelin sheath enables saltatory conduction, which increases the speed of nerve impulses. In disorders in which the development or maintenance of the myelin sheath is impaired, nerve conduction speed is reduced. This results in neuropathy involving motor and sensory dysfunction. Although there are many studies on the molecular mechanisms of myelination and demyelination in the peripheral nervous system, the roles of the numerous proteins involved in these processes remain unclear.
To study the molecular mechanisms of SC myelination/demyelination in vivo, genetic approaches have been used to modify gene expression in animals. A powerful approach is the use of knockout/knockin or transgenic animals. However, the generation of these animals is expensive and time consuming. For SC-specific gene manipulation, crossing floxed strains with Cre mice or other conditional gene expression methods are necessary. This again is laborious and time intensive. In recent years, a cutting-edge genetic technology, the CRISPR-Cas9 system, has made the generation of genetically modified mice much quicker (about 4 weeks)2,3, but this method is hindered by target sequence limitations, and suffers from off-target effects. As an alternative method, viral vector-mediated gene transfer is a faster and easier method of achieving gene transfer into SCs in vivo4-6. Indeed, the generation of viral vectors is less expensive, and takes a shorter time (within a few weeks), and gene manipulation of SCs can be achieved by simply injecting engineered viral vectors, such as adenoviral vectors, adeno-associated viral (AAV) vectors, and lentiviral vectors, into the sciatic nerve. Because these viral vectors have different characteristics, users have to choose the one best suited for their purpose. Adenoviral vectors infect axons and SCs in both young and mature sciatic nerves. In particular, adenoviral vectors have higher selectively for non-myelinating SCs than myelinating SCs. Adenoviruses can cause immune responses, and accordingly, immunodeficient strains should be used5. AAV vectors are currently the most widely used viral vectors, and allow in vivo gene transfer with lower toxicity7. AAV can transduce both axons and SCs by direct injection into the nerve fibers8,9. However, AAV-mediated protein expression usually requires 3 weeks or longer to reach maximum levels7,9. Therefore, it is difficult to analyze myelination, which actively progresses during the two week postnatal period. Lentiviral vectors have higher selectively for myelinating SCs than non-myelinating SCs, and do not have toxic effects on sciatic nerves. However, lentiviral vectors do not infect SCs in more mature nerves5, and therefore are unsuitable for analyzing events such as the demyelination process.
Electroporation is another faster and easier approach to achieve in vivo gene transfer. It has been reported that in vivo transfection of SCs can be achieved when electroporation is applied to transected rat sciatic nerves10. However, because this method requires nerve transection for gene delivery, the application is limited to the analysis of the damaged nerves. Here, we describe an alternative method that allows the delivery of transgenes into myelinating SCs in intact rat sciatic nerves using electroporation11. This method requires plasmid construction, which can usually be completed within a week. Then, by simply delivering electric pulses to the site on the sciatic nerve where the plasmid DNA was injected, highly selective transfection of myelinating SCs can be achieved in neonatal as well as in more mature animals. By electroporating multiple plasmids, simultaneous expression of a variety of genes can be easily achieved. The ability to simultaneous express multiple molecules, such as signaling proteins, short-hairpin RNAs (shRNAs) and functional probes, is crucial for investigating complex processes such as myelination and demyelination. The novel in vivo electroporation method described in this paper will be a powerful tool, allowing researchers to analyze the function of a multitude of molecules and their interactions in myelinating SCs.

<table border="0" cellpadding="0" cellspacing="0" width="573">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Genopure Plasmid Maxi Kit</td>
			<td style="width:71px;">Roche</td>
			<td style="width:119px;">03 143 422 001</td>
			<td style="width:167px;">Plasmid DNA purification kit</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fast Green CFC</td>
			<td>WAKO</td>
			<td>069-00032</td>
			<td>Dye for DNA injection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">GC 150T-10</td>
			<td>HARVARD APPARATUS</td>
			<td>30-0062</td>
			<td>Glass capillary</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Suction tubing</td>
			<td>Drummond</td>
			<td>05-2000-00</td>
			<td>Suction tubing for micro injection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MODEL P-97</td>
			<td colspan="2">SUTTER INSTRUMENT CO.</td>
			<td>Micropipette puller</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">CUY21 Single Cell</td>
			<td>BEX</td>
			<td>Electroporator CUY21 Single Cell</td>
			<td>Pulse generator</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Electric warmer</td>
			<td>KODEN</td>
			<td style="width:119px;">CAH-6A</td>
			<td>Warmer during the surgery</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Isofluolane</td>
			<td style="width:71px;">Mylan</td>
			<td style="width:119px;">1119701G1076</td>
			<td>Anesthetic</td>
		</tr>
	</tbody>
</table>

in vivo gene transfer to schwann cells in the rodent sciatic nerve by electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

An example of a sciatic nerve transfected with red fluorescent protein (RFP)-expressing plasmid is shown in Figure 2A. Cells showing bipolar morphology, a characteristic of SCs, were sparsely transfected with RFP. No RFP fluorescence was detected in axons. We usually find ~100 transfected SCs in every nerve. This transfection efficiency seems similar to the in vivo SC infection efficiency using lentiviral vectors4.
Immunostaining experiments showed that most (~96%) RFP-positive cells at P7 co-labeled for S100, a SC marker (Figure 2B), and 91% of RFP-positive cells at P14 co-labeled for MBP, a myelinating SC marker (Figure 2C), suggesting that gene transfer by electroporation is highly selective for myelinating SCs.
Introduction of multiple genes into SCs in vivo will be extremely useful for investigating the mechanisms of myelination/demyelination. A major advantage of the in vivo electroporation method described here is the capacity to transfer multiple genes with a simple procedure. Figure 2D shows a representative image of a sciatic nerve transfected with a mixture of GFP and RFP-expressing plasmids using in vivo electroporation. About 97% of SCs were GFP and RFP double positive, suggesting that highly efficient delivery of multiple genes can be achieved by simply electroporating mixtures of multiple plasmids.
In rodents, myelination initiates around birth, dramatically increasing during the first two weeks postnatally, and then gradually diminishes. Thus, by genetically manipulating SCs during these developmental time windows, the mechanisms underlying these different stages of myelination can be clarified. Lentiviral vectors are a good tool for analyzing myelination, particularly as they have minimal toxicity, but lentiviruses only infect neonatal sciatic nerves5,6. In comparison, electroporation-mediated gene transfer works well when transfection is conducted on P3 (Figure 2E, top) or on P14 (Figure 2E, bottom).
The applications of the novel in vivo electroporation method are described here. Figure 3A shows light microscopic images of GFP-expressing myelinating SCs at various developmental stages (P7, P14, P21 and P31). By light microscopic analysis, changes in morphological parameters, such as length and diameter, can be assessed. Note that these parameters have similar values compared to intact rat peripheral nerves15,16, suggesting that the electroporated nerves develop without significant damaging effects. Figure 3B shows an electron microscopic image of LacZ-expressing myelinating SCs. In this case, LacZ was used as an expression marker. &#946;-galactosidase staining using bluo-gal, an ethanol-insoluble substrate, enables the analysis of myelin structure of transfected SCs by electron microscopy11,17. In these experiments, the role of signaling molecules can be examined by silencing or augmenting their expression, thereby allowing the analysis of loss-of-function or gain-of-function effects. In addition to the analysis of fixed tissue, in vivo electroporation-mediated gene transfer can be also applied to live imaging experiments. For example, Figure 3C shows a myelinating SC co-expressing G-GECO1.118, a green fluorescent cytosolic Ca2+ indicator, and R-GECO1mt19, a red fluorescent mitochondrial Ca2+ indicator. By expressing these indicators, we identified a signaling pathway that controls cytosolic and mitochondrial Ca2+ concentrations in myelinating SCs. Thus, the present method can be used to study a variety of signaling mechanisms, particularly when genetically-encoded fluorescent probes are available to detect the signals of interest.
<img alt="Figure 1" src="/files/ftp_upload/54567/54567fig1.jpg" /> 
Figure 1: Schematic of the in vivo Electroporation Method. First, the sciatic nerve of the anesthetized rat is exposed. Second, plasmid DNA is injected into the sciatic nerve. Third, electric pulses are delivered to the injection site through the forceps-shaped electrode. Finally, the wound is closed with glue. This procedure can be repeated on the contralateral nerve. <a href="https://www.jove.com/files/ftp_upload/54567/54567fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54567/54567fig2.jpg" /> 
Figure 2: Representative Results on Transfected Sciatic Nerves. (A) A representative image of a transfected sciatic nerve. The nerve was transfected with RFP-expressing plasmid at P3, and fixed at P7. (B) A representative image of a RFP-transfected cell at P7 showing colocalization with S100, a SC marker. (C) A representative image of a RFP-transfected sciatic nerve at P14 showing colocalization with MBP, a myelinating SC marker. (D) A representative image of a sciatic nerve cotransfected with GFP and RFP-expressing plasmids. Transfected SCs simultaneously expressed GFP and RFP. (E) An image of myelinating SCs at P31 transfected at P3, when myelination begins (top), and an image of myelinating SCs at P31 transfected at P14, when most large axons become myelinated (bottom), suggesting that transfection of myelinating SCs can be achieved not only in neonatal nerves, but also in more mature nerves. Scale bars = 200 &#181;m (A); 50 &#181;m (B-E). This figure was modified from our previous publication11. <a href="https://www.jove.com/files/ftp_upload/54567/54567fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54567/54567fig3.jpg" /> 
Figure 3: Application of in vivo Electroporation. (A) A light microscopic analysis of the development of myelinating SCs. Sciatic nerves were electroporated with GFP-expressing plasmid at P3, and were fixed at various developmental stages (P7, P14, P21 and P31). Representative images of GFP-positive SCs are shown on the left. The average length and diameter are summarized as mean &#177; SEM (n = 30 - 47 from 3 nerves) on the right. The length and diameter of myelinating SCs increases as development proceeds. (B) An electron microscopic image of a sciatic nerve transfected with a plasmid encoding LacZ. A transfected SC (white asterisk, left) was finely labeled with precipitates of the &#946;-galactosidase reaction product. (C) An image of a SC cotransfected with G-GECO1.1, a green fluorescent cytosolic Ca2+ indicator, and R-GECO1mt, a red fluorescent mitochondrial Ca2+ indicator. The regions within the white dotted rectangles are shown enlarged in the panels on the right. Scale bars = 50 &#181;m (A and C); 1 &#181;m (B). This figure was modified from our previous publication11. <a href="https://www.jove.com/files/ftp_upload/54567/54567fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation at JoVE.com

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

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Nanyang Technological University

Research

JoVE Journal

Neuroscience

8.8K Views.  The University of Tokyo. Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

Video: In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Axoplasm Isolation from Rat Sciatic Nerve

Isolation of pure axonal cytoplasm (axoplasm) from peripheral nerve is crucial for biochemical studies of many biological processes. In this article, we demonstrate and describe a protocol for axoplasm isolation from adult rat sciatic nerve based on the following steps: (1) dissection of nerve fascicles and separation of connective tissue; (2) incubation of short segments of nerve fascicles in hypotonic medium to release myelin and lyse non-axonal structures; and (3) extraction of the remaining axon-enriched material. Proteomic and biochemical characterization of this preparation has confirmed a high degree of enrichment for axonal components.

We demonstrate a protocol for axoplasm isolation from adult rat sciatic nerve based on dissection of nerve fascicles and incubation in hypotonic medium to release myelin and lyse non-axonal structures, followed by extraction of the remaining axon-enriched material.

Isolation of pure axonal cytoplasm (axoplasm) from peripheral nerve is crucial for biochemical studies of many biological processes. In this article, we ...

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 ...

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 ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for nerve fibers in the central and peripheral nervous systems. This method is particularly applicable to dorsal root ganglia (DRG), which are primary sensory neurons that exhibit a pseudo-unipolar structure of nervous processes. The patterns and features of the action potentials passed along axons are recordable in these neurons. The present study uses in vivo single-fiber recordings to observe the conduction failure of sciatic nerves in complete Freund&#8217;s adjuvant (CFA)-treated rats. As the underlying mechanism cannot be studied using in vivo single-fiber recordings, patch-clamp-recordings of DRG neurons are performed on preparations of intact DRG with the attached sciatic nerve. These recordings reveal a positive correlation between conduction failure and the rising slope of the after-hyperpolarization potential (AHP) of DRG neurons in CFA-treated animals. The protocol for in vivo single fiber-recordings allows the classification of nerve fibers via the measurement of conduction velocity and monitoring of abnormal conditions in nerve fibers in certain diseases. Intact DRG with attached peripheral nerve allows observation of the activity of DRG neurons in most physiological conditions. Conclusively, single-fiber recording combined with electrophysiological recording of intact DRGs is an effective method to examine the role of conduction failure during the analgesic process.

Single-fiber recording is an effective electrophysiological technique that is applicable to the central and peripheral nervous systems. Along with the preparation of intact DRG with the attached sciatic nerve, the mechanism of conduction failure is examined. Both protocols improve the understanding of the peripheral nervous system&#39;s relationship with pain.

Single-fiber recording has been a classical and effective electrophysiological technique over the last few decades because of its specific application for ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Summary

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Chapters in this video

Axoplasm Isolation from Rat Sciatic Nerve

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

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

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Use of In Vivo Single-fiber Recording and Intact Dorsal Root Ganglion with Attached Sciatic Nerve to Examine the Mechanism of Conduction Failure

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury