The authors thank Katie Teng and Dr. Matthew J. Ford for providing plasmids used to generate representative results and protocol optimization. K.H. was supported by Fonds de recherche du Qu&#233;bec - Sant&#233; (FRQS) doctoral grant, Donnor Foundation, Delta Kappa Gamma World Fellowship, Centre for Research in Reproduction and Development (CRRD), Hugh E. Burke, Rolande &#38; Marcel Gosselin, and Alexander McFee graduate studentships.

Animals were housed in static microisolation cages with filter tops, located in a dedicated room containing a type II biosafety cabinet. All animal work was performed in accordance with institutional guidelines and was approved by McGill University&#39;s Faculty of Medicine and Health Sciences Animal Care Committee (AUP #7843).
1. Microinjection needle preparation
<ol>
	<li>Pull a glass capillary tube into a sharp point using a micropipette puller with the following program: P = 500, Heat = 576, PULL = 50, VEL = 80, DEL = 70.</li>
	<li>Using a pair of tapered ultrafine tip forceps, snip the pointed end of the pulled capillary tube under a dissecting microscope to create an opening, as shown in Figure 1A,B. Make sure that the opening is not too large, as this will damage the oviduct and prevent slow injection.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65131/65131fig01.jpg" /> 
Figure 1: Microinjection needle preparation. (A,B) Pulled capillary tube with a pointed end (A) that was snipped to create an opening (B). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65131/65131fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Preparing the anesthetic for intraperitoneal injection
<ol>
	<li>Prepare filtered, diluted avertin on the day of surgery. Avertin (2,2,2-tribromoethanol) stock solution is prepared at a concentration of 1.6 g/mL in tert-amyl alcohol by vigorous stirring and stored in the dark.</li>
	<li>Dilute the avertin stock solution with 1.25% v/v isotonic saline to a final concentration of 20 mg/mL inside a fume hood. Vortex the diluted avertin and filter through a 0.22 &#181;m sterile filter. Keep in the dark.</li>
</ol>
3. Preparing the surgery area
<ol>
	<li>The surgery area should be a dedicated sterile environment, preferably a clean bench or biosafety cabinet. Arrange the sterile surgery equipment, microscope, micromanipulator, and electroporator as needed. An example of this is shown in Figure 2.</li>
	<li>Heat up the heating pad/disc and place it underneath a clean, fully equipped cage. This cage will be used to house the mice following the surgery.</li>
	<li>Pour sterile 1x phosphate buffered saline (PBS) into a sterile petri dish. Use a pair of clean scissors to cut absorbent paper into squares, with approximate dimensions of 1 cm x 1 cm. Drop these pieces of absorbent paper into the 1x PBS to soak them.</li>
</ol>
4. Preparing the solution and needle for injection
<ol>
	<li>Prepare the injection solution on the day of the surgery. Mix the DNA/RNA/RNPs with filtered trypan blue to a minimum final concentration of 200 ng/&#181;L each and 5%, respectively. Store at room temperature prior to injection. 
	&#8203;NOTE: Use a 1 mL syringe to take up 100% trypan blue and filter through a 0.22 &#181;m sterile filter prior to use.</li>
	<li>Attach the pulled capillary needle, henceforth called a microinjection needle, to the clamp mount micromanipulator stabilized by a magnetic mount (setup shown in Figure 2B).</li>
	<li>Pipette 2 &#181;L of the injection solution onto a sterile Petri dish. While observing under the microscope, slowly take up 1-2 &#181;L of this solution into the microinjection needle using the air pressured syringe attached to the micromanipulator. Avoid bubbles/taking up bubbles into the microinjection needle.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65131/65131fig02.jpg" /> 
Figure 2: Surgery area preparation. (A) Surgery area setup inside a clean bench. (B) Dissection microscope and micromanipulator setup for a right-handed individual. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65131/65131fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Exposing the female reproductive tract
<ol>
	<li>Anesthetize a 6-8-week-old female mouse by administering filtered, diluted avertin (anesthetic) intraperitoneally at a dose of 0.25-0.5 mg/g. Following this, administer carprofen (preemptive analgesic) subcutaneously at a dose of 5 mg/kg. 
	NOTE: Other alternatives that can be used as an anesthetic are isoflurane or ketamine/xylazine.</li>
	<li>Place absorbent paper on a clean, heated pad/disc. Then, place the anesthetized mouse on this heated absorbent paper with the dorsal side facing up. Apply eye lubricant to each eye to prevent drying during the surgery. 
	NOTE: Confirm that the heating pad/disc is slightly warm to the touch by testing with the back of the hand.</li>
	<li>Test for deep anesthetic arrest by pinching the toe of the anesthetized mouse with a pair of forceps. Remove dorsal fur around the prospective incision site (location shown in Figure 3A) and wipe the bare skin with antiseptic.</li>
	<li>Use a pair of straight blunt forceps to pinch the bare skin and create a 1 cm long incision along the body midline using a pair of sterile sharp blunt scissors. Clean the area around the incision with antiseptic.</li>
	<li>Pinch and hold up one side of the cut site using a pair of sterile Adson forceps. Then, using a pair of sterile curved serrated forceps, gently separate the skin from the body wall, starting at the midline incision and moving laterally.</li>
	<li>Locate the fat pad below the kidney. Using a pair of sterile straight blunt forceps, pinch the body wall directly above the fat pad. Create a small incision in the body wall using a pair of sterile sharp-pointed dissecting scissors, taking care to avoid blood vessels.</li>
	<li>While still pinching the body wall with straight blunt forceps, insert a pair of sterile blunt curved forceps into the incision and widen the incision created in the body wall.</li>
	<li>Grab the visible fat pad with the blunt curved forceps and pull it out of the hole to expose the ovary, oviduct, and uterus.</li>
	<li>To keep the reproductive tract exposed, clamp the fat pad with a sterile bulldog clamp, as shown in Figure 3B.</li>
</ol>
6. In vivo oviduct injection and electroporation
<ol>
	<li>Carefully place the heated absorbent paper and anesthetized mouse with the reproductive tract exposed on the stage of a dissecting microscope, such that the tract can be observed. An example of the view as observed under the microscope is shown in Figure 3B.</li>
	<li>Adjust the micromanipulator so that the injection needle with solution can also be observed under the microscope.</li>
	<li>Using a pair of sterile tapered ultrafine tip forceps, hold the region of the oviduct that is to be injected steady. The region to be injected must be in line with the direction of the microinjection needle. 
	NOTE: Use the bulldog clamp anchoring the fat pad and reproductive tract to gently turn/move the tract to an optimal position, if needed.</li>
	<li>Adjust the micromanipulator to puncture the oviduct with the microinjection needle, while simultaneously feeding the oviduct onto the needle using the tapered ultrafine tip forceps. Gently move the microinjection needle to confirm that it has been inserted into the oviduct. 
	NOTE: Insert the microinjection needle into a straight segment instead of into a turning point of the coiled oviduct to prevent multiple punctures and leakage of injection solution.</li>
	<li>Slowly inject up to 1 &#181;L of solution into the oviduct, while observing the movement of the blue solution and expansion of the oviduct lumen under the microscope. Take care not to introduce any bubbles into the lumen. Representative images of oviducts injected with 100% trypan blue are shown in Figure 3C,D.</li>
	<li>Remove the needle from the oviduct. Cover the area to be targeted with a piece of absorbent paper presoaked in sterile 1x PBS (from step 3.3).</li>
	<li>Grasp the presoaked paper and the area/region to be targeted with a pair of electrode tweezers (1 mm, 3 mm, or 5 mm, depending on the size of the region to be targeted) and pull away from the body, before electroporating using these settings on the pulse generator/electroporator: 30 V, three pulses, 1 s interval, 50 ms pulse length, unipolar. 
	NOTE: The distance between the electrodes should be set such that the electrodes can clamp onto the targeted region without deforming the oviduct. The recommended distance is around 1 mm.</li>
	<li>After electroporation, remove the absorbent paper and place the mouse (with the heated absorbent paper underneath) back onto a heat pad. Unclamp the bulldog clamp and carefully push the exposed reproductive tract back under the body wall. Repeat section 5 to expose the reproductive tract on the other side. 
	NOTE: Electroporation is successful regardless of the mouse estrous stage as long as the lumen is filled with the solution.</li>
	<li>Cover the exposed tract with absorbent paper presoaked in sterile 1x PBS to prevent it from drying out, then prepare the microinjection needle for the second injection by repeating steps 4.2 and 4.3. 
	NOTE: Replace the microinjection needle as needed. Prevent drying of the exposed tissue by applying sterile 1x PBS when required.</li>
	<li>Repeat the steps above to inject and electroporate the oviduct on the opposing side. 
	&#8203;NOTE: Reapply the eye lubricant as needed during and after the surgery.</li>
	<li>After both oviducts have been electroporated and placed back under the body wall, suture or staple the dorsal incision site. For suturing, use a hemostat to grasp the needle of the silk braided sutures (3/8 circle; gauge: 5-0; needle size: 18 mm; thread length: 75 cm). Pinch and hold up one side of the cut site using a pair of sterile curved serrated forceps, then use the hemostat to manipulate the suture to close the wound.</li>
	<li>Move the mouse into a clean cage that is placed on top of a heated pad/disc (from step 3.2) and monitor until the mouse is active.</li>
	<li>Administer 5 mg/kg carprofen subcutaneously every 24 h for the&#160;next 3 days and monitor over the next 10 days.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65131/65131fig03.jpg" /> 
Figure 3: Female reproductive tract exposure and microinjection. (A) Location of midline incision (shown as a yellow line) on the dorsal side of a Rosa-LSLtdTomato mouse. (B) Female reproductive tract exposure. The fat pad was clamped using a sterile bulldog clamp to anchor the tract and keep it exposed. (C,D) Representative images of in vivo oviduct injection. A microinjection needle was inserted into the distal ampulla (labelled AMP), and filtered 100% trypan blue was injected into the oviduct lumen while the tract was exposed (C). Representative image of a dissected oviduct, demonstrating that injected trypan blue solution distributes throughout the distal and proximal oviduct lumen (D). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65131/65131fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

Crucial steps in this detailed protocol are the microinjection of DNA/RNA/RNP solution into the oviduct lumen and control of the electroporation strength and area. DNA/RNA/RNP solution leakage during microinjection may cause transfection of undesired areas/cells. For consistent and efficient electroporation, it is preferable to fill the oviduct lumen with the solution (Figure 3C,D). This is because the area of electroporation is mainly controlled by electrode size and placement. Weak electroporation reduces the number of electroporated cells, and harsh electroporation may cause the electroporation of undesired cells or disrupt tissue structure. The number of electroporated cells can be varied by adjusting the DNA/RNA/RNP solution concentration or electroporation parameters. However, since high voltages/heat production during electroporation could damage the tissue, these parameters should be tested prior to use on live/anesthetized mice. Finally, due to the demanding nature of this procedure, it is recommended to practice tract exposure and microinjection on dead mice prior to performing this surgery on live/anesthetized mice.
It is recognized that multiple genes are involved in cancer initiation, thus, recapitulating this event requires multi-allelic modification of several genes. Additionally, it is also known that not all cells are equally susceptible to oncogenic mutations9. Cancer modeling, therefore, requires specific Cre mouse lines to achieve tight control of oncogenic insults. However, their availability and specificity causes various challenges, including effects in non-targeted tissues and lethality19. Further, traditional G-GEMMs incur high costs for maintenance and require time for mouse line generation. The development of CRISPR/Cas9 genome editing technology and improvement of gene delivery into specific somatic cells has allowed us to overcome these issues. In this protocol, we present a DNA/RNA/RNP delivery method for somatic genome manipulation that can be used for cancer modeling, without the absolute need for specific mouse lines12. By injecting DNA/RNA/RNP solutions into the lumen and using different sizes of tweezer-type electrodes for electroporation-based delivery, restricted areas of the mouse oviduct mucosal epithelium are targeted (Figure 4A-F). This is especially useful in modeling the initiation of HGSCs that originate from the distal end of the fallopian tube.
The presented microinjection and in vivo electroporation method is highly versatile for targeting tissues/organs with a lumen. Organ parenchyma is also targetable using this method14. Viral delivery approaches, like lentiviral and adenoviral systems, can also be used for similar purposes. However, the advantages of electroporation over viral delivery approaches are: 1) multiple plasmid delivery into single cells, 2) no restriction on the insert size15, and 3) easy control of the area and timing of delivery. In addition, targeting specificity can be improved by using lineage specific promoters. This is sometimes difficult in viral systems due to the limit on viral packaging.
As is the nature of electroporation, electroporated epithelial cells are randomly interspersed with healthy, unedited cells (Figure 4G). However, this mosaicism in genetically modified and unmodified cells could be advantageous to study cancer initiation, as it recapitulates the sporadic nature of early cancer initiation within an immunocompetent microenvironment. The frequency of electroporated cells can be adjusted by varying the DNA/RNA/RNP solution concentrations and/or electroporation parameters. Using CRISPR-Cas-mediated gene editing in combination with the presented protocol, it is easy to screen multiple genes and generate heterogeneous patterns of mutations/allelic combinations in targeted genes; this is particularly useful in modeling cancers that present with a considerable number of genomic alterations like HGSCs20,21. Further, by sequencing targeted genes during cancer progression, it is possible to follow the in vivo clonal evolution of tumors and metastases in immunocompetent mice12.

The fallopian tube, called the oviduct in mice, is a tubular structure that connects the uterus to the ovary. It plays an essential role in mammalian reproduction, providing the environment for internal fertilization and preimplantation development1,2. Despite its importance, little is known about its function and homeostasis, partly due to the development of in vitro fertilization techniques circumventing any infertility issue related to this organ3. However, it has been recognized that precancerous lesions of high-grade serous ovarian carcinoma (HGSC), an aggressive histotype of ovarian cancer that accounts for around 75% of ovarian carcinomas and 85% of related deaths4, are restricted to the distal fallopian tube epithelium5,6,7,8. This indicates that not all cells in our body are equally susceptible to oncogenic insults, but rather only unique/susceptible cells in each tissue/organ become the cell-of-origin in cancer &#8212; termed cellular pliancy9. Along these lines, it has been shown that the epithelial cells of the distal fallopian tube, located adjacent to the ovary, are distinct from the rest of the tube10,11. Thus, traditional mouse models of HGSC, that target all cells in the oviduct, do not recapitulate the human condition. In a recent study, we used a combination of CRISPR-mediated genome editing, in vivo oviduct electroporation, and Cre-based lineage tracing to successfully induce HGSC&#160;by mutating four tumor suppressor genes in the distal mouse oviduct12,13. This manuscript presents a step-by-step protocol describing this procedure of microinjection and in vivo electroporation to target the mouse oviduct mucosal epithelium.
This method has several advantages. It can be adapted to target other tissues/organs, including organ parenchyma14. Although other in vivo gene delivery approaches like lentiviral and adenoviral systems can be used to achieve similar tissue/organ-specific targeting, the area of targeting is more easily adjusted using different sizes of tweezer-type electrodes for electroporation-based delivery. Depending on the concentration of DNA/RNA/ribonucleoproteins (RNPs), electroporation parameters, and size of electrodes, the number of electroporated cells can be altered. Further, specific cell types can be targeted when used in combination with promotors for Cas9 expression, without the absolute need for Germline genetically engineered mouse models (G-GEMMs). In addition, unlike viral delivery systems, electroporation allows for multiple plasmid delivery into single cells and less constraint in the insert DNA size15. In vivo screening of gene mutations can also be performed with relative ease due to this high flexibility. Further, electroporated cells can be tracked or traced when this method is used in combination with Cre reporter lines such as Tdtomato or Confetti16,17.

<table><tbody><tr><td>0.22 &amp;micro;m sterile filter unit&amp;nbsp;</td><td>LifeGene</td><td>SF0.22PES</td><td></td></tr><tr><td>1 mL syringe</td><td>Terumo Medical Corportation</td><td>SS-01T</td><td></td></tr><tr><td>1 mm tweezer-type electrodes</td><td>BEX CO., LTD</td><td>LF650P1</td><td></td></tr><tr><td>3 mm tweezer-type electrodes</td><td>BEX CO., LTD</td><td>LF650P3</td><td></td></tr><tr><td>30G x 1/2 Needle</td><td>BD</td><td>305106</td><td>Needle is attached to the 1 mL syringe when using injectable anesthetic or analgesic.</td></tr><tr><td>Adson forceps</td><td>Fisher Scientific</td><td>10-000-451</td><td></td></tr><tr><td>Air-pressured syringe</td><td>BD</td><td>B302995</td><td>Attached to the micromanipulator; as shown in Figure 2B</td></tr><tr><td>Analgesic</td><td>-</td><td>-</td><td>Carprofen, dose: 5mg/kg.</td></tr><tr><td>Antiseptic</td><td>Cardinal Health</td><td>OMEL0000017</td><td>Baxedin: 2% chlorhexidine in 70% isopropyl alcohol</td></tr><tr><td>Avertin (2,2,2-tribromoethanol) - Anesthetic</td><td>Sigma Aldrich</td><td>T48402-25G</td><td></td></tr><tr><td>Clamp mount micromanipulator</td><td>AD Instruments</td><td>MM-33</td><td></td></tr><tr><td>Curved serrated forceps</td><td>Fisher Scientific</td><td>NC0696845</td><td></td></tr><tr><td>Dissecting microscope</td><td>Zeiss</td><td>SteREO Discovery.V8</td><td>Apochromat S, 0.63X, FWD 107mm. Attached KL 200 LED for top light, labelled as light source in Figure 2B.</td></tr><tr><td>Eye lubricant</td><td>Alcon</td><td>-</td><td>Systane ointment (Lubricant eye ointment)</td></tr><tr><td>Glass capillary needles</td><td>Sutter Instrument Co.</td><td>B100&amp;ndash;50-10</td><td>Outside diameter: 1 mm, inside diameter: 0.5 mm, length: 10 cm</td></tr><tr><td>Hartman hemostats</td><td>Fisher Scientific</td><td>50-822-711</td><td></td></tr><tr><td>Heating pad/disc&amp;nbsp;</td><td>-</td><td>-</td><td>SnuggleSafe Pet Bed Microwave Heating Pad was used in this protocol. Microwave for 2-5min, and test with back of gloved hand.</td></tr><tr><td>Magnetic Mount for micromanipulator</td><td>AD Instruments</td><td>MB-B</td><td>Used to keep the clamp mount micromanipulator stable and upright during injection.</td></tr><tr><td>Micro bulldog clamp</td><td>Fisher Scientific</td><td>50-822-230</td><td>3 cm</td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument Co.</td><td>P-97</td><td>P-97 Flaming/Brown type micropipette puller. Program used: P = 500, Heat = 576, PULL = 50, VEL = 80, DEL = 70</td></tr><tr><td>Petri dish</td><td>Fisher Scientific</td><td>263991</td><td>Autoclaved/sterile glass petridishes can also be used.</td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Bio Basic Inc.</td><td>PD8117</td><td>10X PBS was diluted to 1X with DI water. Autoclave before use.&amp;nbsp;</td></tr><tr><td>Pulse generator/Electroporator</td><td>BEX CO., LTD</td><td>CUY21 EDIT II</td><td>Electroporation settings: 30 V, 3 pulses, 1 s interval, P. length = 50 ms, unipolar</td></tr><tr><td>Rosa-LSLtdTomato mice</td><td>The Jackson Laboratory</td><td>7914</td><td>Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J</td></tr><tr><td>Sharp-blunt dissection scissors</td><td>Fisher Scientific</td><td>28251</td><td></td></tr><tr><td>Sharp-pointed dissecting scissors</td><td>Fisher Scientific</td><td>SDI130</td><td></td></tr><tr><td>Shaver</td><td>-</td><td>-</td><td>Hair removal cream can also be used.</td></tr><tr><td>Silk braided sutures</td><td>Ethicon</td><td>682G</td><td>3/8 circle, gauge: 5-0, needle size: 18 mm, thread length: 75 cm. Staples can also be used.</td></tr><tr><td>Tapered ultrafine tip forceps&amp;nbsp;</td><td>Fisher Scientific</td><td>12-000-122</td><td></td></tr><tr><td>Thin absorbent paper</td><td>Kimberly-Clark Professional</td><td>34120</td><td>Kimwipes</td></tr><tr><td>Trypan blue</td><td>STEMCELL Technologies</td><td>7050</td><td>Filtered using 0.22 &amp;micro;m sterile filter before use for microinjection.</td></tr></tbody></table>

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.

Figure 1 and Figure 2 depict the microinjection needle preparation and surgery area setup, respectively, for in vivo microinjection and electroporation of the mouse oviduct. During the surgery, the female reproductive tract was exposed through incisions made in the dorsal skin (Figure 3A) and body wall of an anesthetized Rosa-LSLtdTomato (Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J)17 mouse. A bulldog clamp was clamped onto the fat pad above the tract to keep it exposed and anchored (Figure 3B). Injection solution containing PCS2 CreNLS plasmids18 at 400 ng/&#181;L concentration was injected into the ampulla (labelled AMP) of the coiled oviduct and allowed to disperse throughout the oviduct lumen (Figure 3C,D). Although PCS2 CreNLS plasmids were used to generate representative results in this manuscript, the described method can be used with easily interchangeable DNA/RNA/RNPs in the injection solution for CRISPR/Cas-mediated gene editing in the mouse oviduct. Then, 1 mm or 3 mm tweezer type electrodes were positioned, such that the distal oviduct was in between the electrodes for electroporation-based delivery.
Oviducts were harvested 4 days following the surgery and stretched out by removing the mesosalpinx to visualize electroporation specificity. Using either 1 mm or 3 mm tweezer-type electrodes, the targeted area, labelled with TdTomato, was restricted to the distal oviduct epithelium (Figure 4A,D). The size of this targeted area was further controlled by using electrodes of different sizes. Using 3 mm tweezer-type electrodes, we targeted a much larger area of the AMP (Figure 4B,C), as compared to targeting only the distal-most tip, the INF (Figure 4E,F), using 1 mm tweezer-type electrodes. These harvested oviducts were fixed, treated with a sucrose gradient, and sectioned using a cryostat for detailed analyses. Sections were counterstained with Hoescht 33342 and Phalloidin to stain the nuclei and cytoskeleton respectively, then they were imaged using a point-scanning confocal microscope. In the targeted region, electroporated cells, marked by TdTomato, were randomly distributed among non-electroporated (TdTomato-ve) cells (Figure 4G). Additionally, electroporated cells were restricted to the mucosal epithelium and not found in the underlying stromal or muscle layer (Figure 4G,H). Finally, to confirm CRISPR-Cas-mediated gene editing, TdTomato+ve cells can be isolated by fluorescence-activated cell sorting (FACS) for DNA isolation and MiSeq sequencing12.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65131/65131fig04.jpg" /> 
Figure 4: Validation of successful in vivo injection and electroporation of oviduct epithelial cells, 4 days post-surgery. (A-C) Electroporation using 3 mm sized tweezer-type electrodes. The oviduct was uncoiled by removing the mesosalpinx for better visualization of electroporation specificity (A). Electroporation area, identified by TdTomato expression, was restricted to the distal oviduct, labelled AMP (B,C). (D-F) Electroporation using 1 mm sized tweezer-type electrodes. The oviduct was uncoiled by removing the mesosalpinx for better visualization of electroporation specificity (D). The electroporation area was restricted to the infundibulum (labelled INF), the distal-most tip of the oviduct (E,F). (G,H) Transverse section of the distal oviduct 4 days following electroporation with PCS2 CreNLS plasmids. Electroporated cells labelled with TdTomato (G) were restricted to the epithelial monolayer (H). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65131/65131fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Somatic Genome-Engineered Mouse Models Using In Vivo Microinjection and Electroporation at JoVE.com

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.

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. ...

somatic-genome-engineered-mouse-models-using-vivo-microinjection

Nanyang Technological University

Research

JoVE Journal

Cancer Research

1.5K Views.  McGill University. This protocol describes microinjection and in vivo electroporation for regionally restricted CRISPR-mediated genome editing in the mouse oviduct.

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

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 ...

Neuroscience

Generation of Genetically Modified Mice through the Microinjection of Oocytes

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear injection of DNA expression constructs into fertilized oocytes remains the most commonly used technique to generate transgenic mice for overexpression. With the introduction of CRISPR technology for gene targeting, pronuclear injection into fertilized oocytes has been extended to the generation of both knockout and knockin mice. This work describes the preparation of DNA for injection and the generation of CRISPR guides for gene targeting, with a particular emphasis on quality control. The genotyping procedures required for the identification of potential founders are critical. Innovative genotyping strategies that take advantage of the "multiplexing" capabilities of CRISPR are presented herein. Surgical procedures are also outlined. Together, the steps of the protocol will allow for the generation of genetically modified mice and for the subsequent establishment of mouse colonies for a plethora of research fields, including immunology, neuroscience, cancer, physiology, development, and others.

The microinjection of mouse oocytes is commonly used for both classic transgenesis (i.e., the random integration of transgenes) and CRISPR-mediated gene targeting. This protocol reviews the latest developments in microinjection, with a particular emphasis on quality control and genotyping strategies.

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear ...

Developmental Biology

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Genetic modification of specific regions of the developing mammalian brain is a very powerful experimental approach. However, generating novel mouse mutants is often frustratingly slow. It has been shown that access to the mouse brain developing in utero with reasonable post-operatory survival is possible. Still, results with this procedure have been reported almost exclusively for the most superficial and easily accessible part of the developing brain, i.e. the cortex. The thalamus, a narrower and more medial region, has proven more difficult to target. Transfection into deeper nuclei, especially those of the hypothalamus, is perhaps the most challenging and therefore very few results have been reported. Here we demonstrate a procedure to target the entire hypothalamic neuroepithelium or part of it (hypothalamic regions) for transfection through electroporation. The keys to our approach are longer narcosis times, injection in the third ventricle, and appropriate kind and positioning of the electrodes. Additionally, we show results of targeting and subsequent histological analysis of the most recessed hypothalamic nucleus, the mammillary body.

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Despite the functional and medical importance of the hypothalamus, in utero genetic manipulation of its development has rarely been attempted. We show a detailed procedure for in utero electroporation into the mouse hypothalamus and show representative results of total and partial (regional) hypothalamic transfection.

Genetic  modification of specific regions of the developing mammalian brain is a very  powerful experimental approach. However, generating novel mouse ...

In Vivo Brain Tumor Modeling via Electroporation-Based Delivery of Plasmid DNA

Modeling Brain Tumors In Vivo Using Electroporation-Based Delivery of Plasmid DNA Representing Patient Mutation Signatures

Tumor models are critical for the preclinical testing of brain tumors in terms of exploring new, more efficacious treatments. With significant interest in immunotherapy, it is even more critical to have a consistent, clinically pertinent, immunocompetent mouse model to examine the tumor and immune cell populations in the brain and their response to treatment. While most preclinical models utilize orthotopic transplantation of established tumor cell lines, the modeling system presented here allows for a "personalized" representation of patient-specific tumor mutations in a gradual, yet effective development from DNA constructs inserted into dividing neural precursor cells (NPCs) in vivo. DNA constructs feature the mosaic analysis with the dual-recombinase-mediated cassette exchange (MADR) method, allowing for single-copy, somatic mutagenesis of driver mutations. Using newborn mouse pups between birth and 3 days old, NPCs are targeted by taking advantage of these dividing cells lining the lateral ventricles. Microinjection of DNA plasmids (e.g., MADR-derived, transposons, CRISPR-directed sgRNA) into the ventricles is followed by electroporation using paddles that surround the rostral region of the head. Upon electrical stimulation, the DNA is taken up into the dividing cells, with the potential of integrating into the genome. The use of this method has successfully been demonstrated in developing both pediatric and adult brain tumors, including the most common malignant brain tumor, glioblastoma. This article discusses and demonstrates the different steps of developing a brain tumor model using this technique, including the procedure of anesthetizing young mouse pups, to microinjection of the plasmid mix, followed by electroporation. With this autochthonous, immunocompetent mouse model, researchers will have the ability to expand preclinical modeling approaches, in efforts to improve and examine efficacious cancer treatment.

Author Spotlight: Modeling Brain Tumors In Vivo Using Electroporation-Based Delivery of Plasmid DNA Representing Patient Mutation Signatures  

Utilizing an immunocompetent, autochthonous tumor model driven by common patient mutations for preclinical testing is critical for immunotherapeutic testing. This protocol describes a method to generate brain tumor mouse models using electroporation-based delivery of plasmid DNA that represent common patient mutations, thus providing an accurate, reproducible, and consistent mouse model.

Tumor models are critical for the preclinical testing of brain tumors in terms of exploring new, more efficacious treatments. With significant interest in ...

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (&#62;1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily ...

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the efficiency in developing gene-knockout mice by inducing small indel mutation would be sufficient enough, the efficiency of embryo genome editing for making large-size DNA knock-in (KI) is still low. Therefore, in contrast to the direct KI method in embryos, gene targeting using embryonic stem cells (ESCs) followed by embryo injection to develop chimera mice still has several advantages (e.g., high throughput targeting in vitro, multi-allele manipulation, and Cre and flox gene manipulation can be carried out in a short period). In addition, strains with difficult-to-handle embryos in vitro, such as BALB/c, can also be used for ESC targeting. This protocol describes the optimized method for large-size DNA (several kb) KI in ESCs by applying CRISPR/Cas9-mediated genome editing followed by chimera mice production to develop gene-manipulated mouse models.

Here we present a protocol for developing genetically modified mouse models using embryonic stem cells, especially for large DNA knock-in (KI). This protocol is tuned up using CRISPR/Cas9 genome editing, resulting in significantly improved KI efficiency compared with the conventional homologous recombination-mediated linearized DNA targeting method.

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the ...

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal experiments. When generating animal models, the electroporation of zygotes offers higher efficiency, simplicity, cost, and throughput as an alternative to the gold standard method of microinjection. Electroporation is also gentler, with higher viability, and reliably delivers Cas9/single-guide RNA (sgRNA) ribonucleoproteins (RNPs) into the zygotes of common laboratory mouse strains (e.g., C57BL/6J and C57BL/6N) that approaches 100% delivery efficiency. This technique enables insertion/deletion (indels) mutations, point mutations, the deletion of whole genes or exons, and small insertions in the range of 100-200 bp to insert LoxP or short tags like FLAG, HA, or V5. While constantly being improved, here we present the current state of CRISPR-EZ in a protocol that includes sgRNA production through in vitro transcription, embryo processing, RNP assembly, electroporation, and the genotyping of preimplantation embryos. A graduate-level researcher with minimal experience manipulating embryos can obtain genetically edited embryos in less than 1 week using this protocol. Here, we offer a straightforward, low-cost, efficient, high-capacity method that could be used with mouse embryos.

Here, we describe a simple technique intended for the efficient generation of genetically modified mice called CRISPR RNP Electroporation of Zygotes (CRISPR-EZ). This method delivers editing reagents by electroporation into embryos at an efficiency approaching 100%. This protocol is effective for point mutations, small genomic insertions, and deletions in mammalian embryos.

With exceptional efficiency, accuracy, and ease, the CRISPR/Cas9 system has significantly improved genome editing in cell culture and lab animal ...

Mouse Genome Engineering Using Designer Nucleases

Transgenic mice carrying site-specific genome modifications (knockout, knock-in) are of vital importance for dissecting complex biological systems as well as for modeling human diseases and testing therapeutic strategies. Recent advances in the use of designer nucleases such as zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and the clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 system for site-specific genome engineering open the possibility to perform rapid targeted genome modification in virtually any laboratory species without the need to rely on embryonic stem (ES) cell technology. A genome editing experiment typically starts with identification of designer nuclease target sites within a gene of interest followed by construction of custom DNA-binding domains to direct nuclease activity to the investigator-defined genomic locus. Designer nuclease plasmids are in vitro transcribed to generate mRNA for microinjection of fertilized mouse oocytes. Here, we provide a protocol for achieving targeted genome modification by direct injection of TALEN mRNA into fertilized mouse oocytes.

Designer nucleases such as zinc finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs) can be used to modify the genome of mouse preimplantation embryos by triggering both the nonhomologous end joining (NHEJ) and homologous recombination (HR) pathways. These advances enable the rapid generation of mice with precise genetic modifications.

Transgenic mice carrying site-specific genome modifications (knockout, knock-in) are of vital importance for dissecting complex biological systems as well ...

Biology

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.

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.

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

Electroporation of Plasmid DNA into Mouse Skeletal Muscle

Transient gene expression modulation in murine skeletal muscle by plasmid electroporation is a useful tool for assessing normal and pathological physiology. Overexpression or knockdown of target genes enables investigators to manipulate individual molecular events and, thus, better understand the mechanisms that impact muscle mass, muscle metabolism, and contractility. In addition, electroporation of DNA plasmids that encode fluorescent tags allows investigators to measure changes in subcellular localization of proteins in skeletal muscle in vivo. A key functional assessment of skeletal muscle includes the measurement of muscle contractility. In this protocol, we demonstrate that whole muscle contractility studies are still possible after plasmid DNA injection, electroporation, and gene expression modulation. The goal of this instructional procedure is to demonstrate the step-by-step method of DNA plasmid electroporation into mouse skeletal muscle to facilitate uptake and expression in the myofibers of the tibialis anterior and extensor digitorum longus muscles, as well as to demonstrate that skeletal muscle contractility is not compromised by injection and electroporation.

Electroporation of plasmid DNA into skeletal muscle is a viable method to modulate gene expression without compromising muscle contractility in mice.

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

Summary

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Chapters in this video

In vivo Electroporation of Developing Mouse Retina

Generation of Genetically Modified Mice through the Microinjection of Oocytes

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Modeling Brain Tumors In Vivo Using Electroporation-Based Delivery of Plasmid DNA Representing Patient Mutation Signatures

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Mouse Genome Engineering Using Designer Nucleases

In Vivo Microinjection and Electroporation of Mouse Testis

Electroporation of Plasmid DNA into Mouse Skeletal Muscle