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/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/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/files/ftp_upload/65131/65131fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High+grade+serous+ovarian+carcinomas+originate+in+the+fallopian+tube.">High grade serous ovarian carcinomas originate in the fallopian tube.</a> Nature Communications. 8 (1), 1093 (2017).</li><li>Shaw, P. A., Rouzbahman, M., Pizer, E. S., Pintilie, M., Begley, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Candidate+serous+cancer+precursors+in+fallopian+tube+epithelium+of+BRCA1/2+mutation+carriers.">Candidate serous cancer precursors in fallopian tube epithelium of BRCA1/2 mutation carriers.</a> Modern Pathology. 22 (9), 1133-1138 (2009).</li><li>Soong, T. 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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oviduct+epithelial+cells+constitute+two+developmentally+distinct+lineages+that+are+spatially+separated+along+the+distal-proximal+axis.">Oviduct epithelial cells constitute two developmentally distinct lineages that are spatially separated along the distal-proximal axis.</a> Cell Reports. 36 (10), 109677 (2021).</li><li>Teng, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+high-grade+serous+ovarian+carcinoma+using+a+combination+of+in+vivo+fallopian+tube+electroporation+and+CRISPR-Cas9-mediated+genome+editing.">Modeling high-grade serous ovarian carcinoma using a combination of in vivo fallopian tube electroporation and CRISPR-Cas9-mediated genome editing.</a> Cancer Research. 81 (20), 5147-5160 (2021).</li><li>Ford, M. 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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.22 &#181;m sterile filter unit&#160;</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&#8211;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&#160;</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.&#160;</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&#160;</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 &#181;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/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

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JoVE Journal

Cancer Research

1.5K Views.  McGill University. Het gepresenteerde protocol is een zeer flexibele methode voor genoommanipulatie in de eileider van de muis. Het maakt de vorming mogelijk van sporadisch gemuteerde cellen omringd door onbewerkte cellen in een immunocompetente omgeving. Dit is voordelig om kankerinitiatie te bestuderen.Het belangrijkste voordeel van deze techniek is het hoge aanpassingsvermogen bij het richten van een orgaan, gebied, regio of celtype met gemakkelijk uitwisselbare sets mutatiecombinaties, allemaal zonde...

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

Using RNA-sequencing to Detect Novel Splice Variants Related to Drug Resistance in In Vitro Cancer Models

Drug resistance remains a major problem in the treatment of cancer for both hematological malignancies and solid tumors. Intrinsic or acquired resistance can be caused by a range of mechanisms, including increased drug elimination, decreased drug uptake, drug inactivation and alterations of drug targets. Recent data showed that other than by well-known genetic (mutation, amplification) and epigenetic (DNA hypermethylation, histone post-translational modification) modifications, drug resistance mechanisms might also be regulated by splicing aberrations. This is a rapidly growing field of investigation that deserves future attention in order to plan more effective therapeutic approaches. The protocol described in this paper is aimed at investigating the impact of aberrant splicing on drug resistance in solid tumors and hematological malignancies. To this goal, we analyzed the transcriptomic profiles of several in vitro models through RNA-seq and established a qRT-PCR based method to validate candidate genes. In particular, we evaluated the differential splicing of DDX5 and PKM transcripts. The aberrant splicing detected by the computational tool MATS was validated in leukemic cells, showing that different DDX5 splice variants are expressed in the parental vs. resistant cells. In these cells, we also observed a higher PKM2/PKM1 ratio, which was not detected in the Panc-1 gemcitabine-resistant counterpart compared to parental Panc-1 cells, suggesting a different mechanism of drug-resistance induced by gemcitabine exposure.

Using RNA-sequencing to Detect Novel Splice Variants Related to Drug Resistance in In Vitro Cancer Models

Here we describe a protocol aimed at investigating the impact of aberrant splicing on drug resistance in solid tumors and hematological malignancies. To this goal, we analyzed the transcriptomic profiles of parental and resistant in vitro models through RNA-seq and established a qRT-PCR based method to validate candidate genes.

Drug resistance remains a major problem in the treatment of cancer for both hematological malignancies and solid tumors. Intrinsic or acquired resistance ...

Intracarotid Cancer Cell Injection to Produce Mouse Models of Brain Metastasis

Metastasis, the spread and growth of malignant cells at secondary sites within a patient&#39;s body, accounts for &#62; 90% of cancer-related mortality. Recently, impressive advances in novel therapies have dramatically prolonged survival and improved quality of life for many cancer patients. Sadly, incidence of brain metastatic recurrences is fast rising, and all current therapies are merely palliative. Hence, good experimental animal models are urgently needed to facilitate in-depth studies of the disease biology and to assess novel therapeutic regimens for preclinical evaluation. However, the standard in vivo metastasis assay via tail vein injection of cancer cells produces predominantly lung metastatic lesions; animals usually succumb to the lung tumor burden before any meaningful outgrowth of brain metastasis. Intracardiac injection of tumor cells produces metastatic lesions to multiple organ sites including the brain; however, the variability of tumor growth produced with this model is large, dampening its utility in evaluating therapeutic efficacy. To generate reliable and consistent animal models for brain metastasis study, here we describe a procedure for producing experimental brain metastasis in the house mouse (Mus musculus) via intracarotid injection of tumor cells. This approach allows one to produce large number of brain metastasis-bearing mice with similar growth and mortality characteristics, thus facilitating research efforts to study basic biological mechanisms and to assess novel therapeutic agents.

Brain metastasis has become an urgent unmet medical need as its incidence has increased while therapeutic options have remained palliative. Creating experimental animal models of brain metastasis via intracarotid arterial injection of cancer cells facilitates mechanistic studies of the disease biology and evaluation of novel intervention regimens.

Metastasis, the spread and growth of malignant cells at secondary sites within a patient&#39;s body, accounts for &#62; 90% of cancer-related mortality. ...

In Vivo EPR Assessment of pH, pO2, Redox Status, and Concentrations of Phosphate and Glutathione in the Tumor Microenvironment

This protocol demonstrates the capability of low-field electron paramagnetic resonance (EPR)-based techniques in combination with functional paramagnetic probes to provide quantitative information on the chemical tumor microenvironment (TME), including pO2, pH, redox status, concentrations of interstitial inorganic phosphate (Pi), and intracellular glutathione (GSH). In particular, an application of a recently developed soluble multifunctional trityl probe provides unsurpassed opportunity for in vivo concurrent measurements of pH, pO2 and Pi in Extracellular space (HOPE probe). The measurements of three parameters using a single probe allow for their correlation analyses independent of probe distribution and time of the measurements.

In Vivo EPR Assessment of pH, pO2, Redox Status, and Concentrations of Phosphate and Glutathione in the Tumor Microenvironment

Low-field (L-band, 1.2 GHz) electron paramagnetic resonance using soluble nitroxyl and trityl probes is demonstrated for assessment of physiologically important parameters in the tumor microenvironment in mouse models of breast cancer.

This protocol demonstrates the capability of low-field electron paramagnetic resonance (EPR)-based techniques in combination with functional paramagnetic ...

A Syngeneic Pancreatic Cancer Mouse Model to Study the Effects of Irreversible Electroporation

Pancreatic cancer (PC), a disease which kills approximately 40,000 patients each year in the US, has successfully evaded several therapeutic approaches including the promising immunotherapeutic strategies. Irreversible electroporation (IRE) is a non-thermal ablation technique that induces tumor cell death without destruction of adjacent collagenous structures, thus enabling the procedure to be performed in tumors very close to blood vessels. Unlike thermal ablation techniques, IRE results in gradual apoptotic cell death, along with immediate ablation induced necrosis, and is currently in clinical use for selected patients with locally advanced PC. An ablative, non-target specific procedure like IRE can induce a myriad of responses in the tumor microenvironment. A few studies have addressed the effects of IRE on tumor growth in other tumor types, but none have focused on PC. We have developed a syngeneic mouse model of PC in which subcutaneous (SQ) and orthotopic tumors can be successfully treated with IRE in a highly controlled setting, facilitating various longitudinal studies post procedure. This animal model serves as a robust system to study the effects of IRE and ways to improve the clinical efficacy of IRE.

Irreversible electroporation (IRE) is a non-thermal ablation technique used for the treatment of locally advanced pancreatic cancer. Being a relatively new technique, the effects of IRE on the tumor growth are poorly understood. We have developed a syngeneic mouse model that facilitates studying the effects of IRE on pancreatic cancer.

Pancreatic cancer (PC), a disease which kills approximately 40,000 patients each year in the US, has successfully evaded several therapeutic approaches ...

Identification, Histological Characterization, and Dissection of Mouse Prostate Lobes for In Vitro 3D Spheroid Culture Models

Genetically engineered mouse models (GEMMs) serve as effective pre-clinical models for investigating most types of human cancers, including prostate cancer (PCa). Understanding the anatomy and histology of the mouse prostate is important for the efficient use and proper characterization of such animal models. The mouse prostate has four distinct pairs of lobes, each with their own characteristics. This article demonstrates the proper method of dissection and identification of mouse prostate lobes for disease analysis. Post-dissection, the prostate cells can be further cultured in vitro for mechanistic understanding. Since mouse prostate primary cells tend to lose their normal characteristics when cultured in vitro, we outline here a method for isolating the cells and growing them as 3D spheroid cultures, which is effective for preserving the physiological characteristics of the cells. These 3D cultures can be used for analyzing cell morphology and behavior in near-physiological conditions, investigating altered levels and localizations of key proteins and pathways involved in the development and progression of a disease, and looking at responses to drug treatments.

Genetically engineered mice are useful models for investigating prostate cancer mechanisms. Here we present a protocol to identify and dissect prostate lobes from a mouse urogenital system, differentiate them based on histology, and isolate and culture the primary prostate cells in vitro as spheroids for downstream analyses.

Genetically engineered mouse models (GEMMs) serve as effective pre-clinical models for investigating most types of human cancers, including prostate ...

Establishment of Gastric Cancer Patient-derived Xenograft Models and Primary Cell Lines

The use of preclinical models to advance our understanding of tumor biology and investigate the efficacy of therapeutic agents is key to cancer research. Although there are many established gastric cancer cell lines and many conventional transgenic mouse models for preclinical research, the disadvantages of these in vitro and in vivo models limit their applications. Because the characteristics of these models have changed in culture, they no longer model tumor heterogeneity, and their responses have not been able to predict responses in humans. Thus, alternative models that better represent tumor heterogeneity are being developed. Patient-derived xenograft (PDX) models preserve the histologic appearance of cancer cells, retain intratumoral heterogeneity, and better reflect the relevant human components of the tumor microenvironment. However, it usually takes 4-8 months to develop a PDX model, which is longer than the expected survival of many gastric patients. For this reason, establishing primary cancer cell lines may be an effective complementary method for drug response studies. The current protocol describes methods to establish PDX models and primary cancer cell lines from surgical gastric cancer samples. These methods provide a useful tool for drug development and cancer biology research.

The current protocol describes methods to establish patient-derived xenograft (PDX) models and primary cancer cell lines from surgical gastric cancer samples. The methods provide a useful tool for drug development and cancer biology research.

The use of preclinical models to advance our understanding of tumor biology and investigate the efficacy of therapeutic agents is key to cancer research. ...

Using the Chicken Chorioallantoic Membrane In Vivo Model to Study Gynecological and Urological Cancers

Mouse models are the benchmark tests for in vivo cancer studies. However, cost, time, and ethical considerations have led to calls for alternative in vivo cancer models. The chicken chorioallantoic membrane (CAM) model provides an inexpensive, rapid alternative that permits direct visualization of tumor development and is suitable for in vivo imaging. As such, we sought to develop an optimized protocol for engrafting gynecological and urological tumors into this model, which we present here. Approximately 7 days postfertilization, the air cell is moved to the vascularized side of the egg, where an opening is created in the shell. Tumors from murine and human cell lines and primary tissues can then be engrafted. These are typically seeded in a mixture of extracellular matrix and medium to avoid cellular dispersal and provide nutrient support until the cells recruit a vascular supply. Tumors may then grow for up to an additional 14 days prior to the eggs hatching. By implanting cells stably transduced with firefly luciferase, bioluminescence imaging can be used for the sensitive detection of tumor growth on the membrane and cancer cell spread throughout the embryo. This model can potentially be used to study tumorigenicity, invasion, metastasis, and therapeutic effectiveness. The chicken CAM model requires significantly less time and financial resources compared to traditional murine models. Because the eggs are immunocompromised and immune tolerant, tissues from any organism can potentially be implanted without costly transgenic animals (e.g., mice) required for implantation of human tissues. However, many of the advantages of this model could potentially also be limitations, including the short tumor generation time and immunocompromised/immune tolerant status. Additionally, although all tumor types presented here engraft in the chicken chorioallantoic membrane model, they do so with varying degrees of tumor growth.

We present the chicken chorioallantoic membrane model as an alternative, transplantable, in vivo model for the engraftment of gynecological and urological cancer cell lines and patient-derived tumors.

Mouse models are the benchmark tests for in vivo cancer studies. However, cost, time, and ethical considerations have led to calls for alternative in vivo ...

Intratibial Osteosarcoma Cell Injection to Generate Orthotopic Osteosarcoma and Lung Metastasis Mouse Models

Osteosarcoma is the most common primary bone cancer in children and adolescents, with lungs as the most common metastatic site. The five-year survival rate of osteosarcoma patients with pulmonary metastasis is less than 30%.Therefore, the utilization of mouse models mimicking the osteosarcoma development in humans is of great significance for understanding the fundamental mechanism of osteosarcoma carcinogenesis and pulmonary metastasis to develop novel therapeutics. Here, detailed procedures are reported to generate the primary osteosarcoma and pulmonary metastasis mouse models via&#160;intratibia injection of osteosarcoma cells. Combined with the bioluminescence or X-ray live imaging system, these living mouse models are utilized to monitor and quantify osteosarcoma growth and metastasis. To establish this model, a basement membrane matrix containing osteosarcoma cells was loaded in a micro-volume syringe and injected into one tibia of each athymic mouse after being anesthetized. The mice were sacrificed when the primary osteosarcoma reached the size limitation in the IACUC-approved protocol. The legs bearing osteosarcoma and the lungs with metastasis lesions were separated. These models are characterized by a short incubation period, rapid growth, severe lesions, and sensitivity in monitoring the development of primary and pulmonary metastatic lesions. Therefore, these are ideal models for exploring the functions and mechanisms of specific factors in osteosarcoma carcinogenesis and pulmonary metastasis, the tumor microenvironment, and evaluating the therapeutic efficacy in vivo.

The present protocol describes intratibia osteosarcoma cell injection to generate mouse models bearing orthotopic osteosarcoma and pulmonary metastasis lesions.

Osteosarcoma is the most common primary bone cancer in children and adolescents, with lungs as the most common metastatic site. The five-year survival ...

Intraductal Delivery and X-ray Visualization of Ethanol-Based Ablative Solution for Prevention and Local Treatment of Breast Cancer in Mouse Models

Breast cancer is the most prevalent cancer and the second-leading cause of cancer-related death for women in the USA. For high-risk women, prophylactic mastectomy is the most effective primary prevention strategy. Prophylactic mastectomy is an aggressive surgical procedure that completely removes the mammary epithelial cells from which breast cancer arises along with the surrounding tissue. We seek to develop a minimally invasive intraductal procedure as an alternative to prophylactic mastectomy to locally ablate the mammary epithelial cells before they can become malignant. We and others have developed an intraductal delivery procedure to reach and treat these epithelial cells in rodent models of breast cancer. While the mouse mammary gland with a single non-anastomosed ductal tree opening at the nipple has a much less complex and tortuous architecture than the human breast, chemically induced and genetically engineered mouse models of breast cancer are valuable to produce proof-of-concept studies of new preventative strategies. Here, we describe a procedure for intraductal delivery of an ethanol-based ablative solution containing micro-CT/X-ray tantalum-based contrast agent within the mouse mammary ductal tree for the therapeutic purpose of primary prevention of breast cancer. Intraductal delivery of aqueous reagents (e.g., cytotoxic compounds, siRNAs, AdCre) has been previously described in mouse models. Thus, we focus our protocol description on methodological modifications and unique experimental considerations for optimizing delivery of ethanol, for minimizing local and systemic side effects of ethanol administration, and for in vivo visualization of ductal tree filling via micro-CT/fluoroscopy imaging. Visualization of the ductal tree immediately after injection of a contrast-containing solution allows for confirmation of complete filling or unsuccessful outcomes such as underfilling or overfilling. This procedure can be applied for delivery and imaging of other ablative compounds aimed at either preventing tumor formation or locally treating early-stage tumors accessible via the ductal tree.

A method of intraductal injection of reagents for an ethanol-based ablative solution to the mouse mammary ductal tree for in vivo imaging and breast cancer prevention is described. Injection directly into the nipple opening allows for targeting mammary epithelial cells with minimal collateral tissue damage.

Breast cancer is the most prevalent cancer and the second-leading cause of cancer-related death for women in the USA. For high-risk women, prophylactic ...

Ex Vivo Organoid Model of Adenovirus-Cre Mediated Gene Deletions in Mouse Urothelial Cells

Bladder cancer is an understudied area, particularly in genetically engineered mouse models (GEMMs). Inbred GEMMs with tissue-specific Cre and loxP sites have been the gold standards for conditional or inducible gene targeting. To provide faster and more efficient experimental models, an ex vivo organoid culture system is developed using adenovirus Cre and normal urothelial cells carrying multiple loxP alleles of the tumor suppressors Trp53, Pten, and Rb1. Normal urothelial cells are enzymatically disassociated from four bladders of triple floxed mice (Trp53f/f: Ptenf/f: Rb1f/f). The urothelial cells are transduced ex vivo with adenovirus-Cre driven by a CMV promoter (Ad5CMVCre). The transduced bladder organoids are cultured, propagated, and characterized in vitro and in vivo. PCR is used to confirm gene deletions in Trp53, Pten, and Rb1. Immunofluorescence (IF) staining of organoids demonstrates positive expression of urothelial lineage markers (CK5 and p63). The organoids are injected subcutaneously into host mice for tumor expansion and serial passages. The immunohistochemistry (IHC) of xenografts exhibits positive expression of CK7, CK5, and p63 and negative expression of CK8 and Uroplakin 3. In summary, adenovirus-mediated gene deletion from mouse urothelial cells engineered with loxP sites is an efficient method to rapidly test the tumorigenic potential of defined genetic alterations.

Ex Vivo Organoid Model of Adenovirus-Cre Mediated Gene Deletions in Mouse Urothelial Cells

This protocol describes the process of the generation and characterization of mouse urothelial organoids harboring deletions in genes of interest. The methods include harvesting mouse urothelial cells, ex vivo transduction with adenovirus driving Cre expression with a CMV promoter, and in vitro as well as in vivo characterization.

Bladder cancer is an understudied area, particularly in genetically engineered mouse models (GEMMs). Inbred GEMMs with tissue-specific Cre and loxP sites ...

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

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