Research reported in this publication was supported by the National Institutes of Allergy and Infectious Diseases (NIAID) of the National Institutes of Health under award number 1R41OD021979 (to ZW) and by a research grant from the Next-Generation BioGreen 21 Program, Republic of Korea, grant no. PJ01107704 (to ZW) and grant no. PJ01107703 (to IK). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or BioGreen 21. We thank Dr. Nikolas Robl for editing the manuscript.

The procedures described in this protocol were approved by the Institutional Animal Care and Use Committee (IACUC) of Utah State University (IACUC protocol: 2484). Hamsters used in this protocol are adult (6 - 10 weeks of age) LVG strain golden Syrian hamsters. All hamsters are housed in the vivarium at the Bioinnovation center, Utah State University. Room temperature is set at 23 &#176;C, humidity is set at 40 - 50%, and light cycle is set 14L:10D (light:dark). Whenever possible, embryo manipulation and surugical procedures should be performed with sterile techniques.
1. sgRNA and Cas9/sgRNA Ribonucleoproteins (RNP) Preparation
<ol>
	<li>Synthesize sgRNAs following the in vitro transcription procedures described in the manual of synthesis kit (Materials Table, Supplement 1).</li>
	<li>To assemble ribonucleoproteins, mix 2 &#181;g of Cas9 with 1 &#181;g of sgRNA and incubate the mix at room temperature for 10 - 15 min. For PN injection, make a working solution at the concentration of 100 ng/&#181;L Cas9 and 50 ng/&#181;L sgRNA with 10 mM TE buffer (RNase free, Materials Table).</li>
</ol>
2. Vasectomy Preparation
NOTE: Vasectomy is performed on male hamsters at 6 - 8 weeks of age. The surgery should be performed 10-14 days ahead of the first mating. Sterility is confirmed by failed pregnancies from mating with fertile females. Vasectomized males can normally be used for a year before they become less sexually active.
<ol>
	<li>Anesthetize males by intraperitoneal (i.p.) injection of ketamine/xylazine with a dose of 40 mg/kg (ketamine) and 10 mg/kg (xylazine). Confirm anesthesia by lack of a pedal reflex (toe pinch).</li>
	<li>Lay the sedated hamster in dorsal recumbency on a dry tissue paper. Rinse the abdomen with three rounds of antiseptic treatments by alternating surgical scrub solutions between 70% alcohol and povidone-iodine.</li>
	<li>Make a vertical incision with surgical scissors starting 1.5 cm cranial to the prepuce extending 1 cm to each side from mid line (Figure 1a). Incise the skin, subcutaneous tissue, and linea alba to access the peritoneal space (Figure 1b).</li>
	<li>Apply gentle pressure to the caudal aspect of the scrotum to force the testes and fat tissue out. Grasp the vas deferens gently with forceps and carefully separate the blood vessel from the vas with a second set of forceps (Figure 1c).</li>
	<li>Hold the vas deferens with forceps to form a loop. Heat the tip of another forceps in flame until it is red, and then use it to remove the loop of the vas deferens. Insert fat tissue and testes back to abdomen. Remove the vas deferens on the opposing side with the same procedures.</li>
	<li>Suture the musculature first followed by suturing the skin. Place the animal on a warm pad in a cage for 30 min until the animal recovers. Observe the animal until it resumes normal activity.</li>
</ol>
3. Donor/Recipient Hamster Preparation Schedule
<ol>
	<li>Monitor and record estrous cycles. 
	NOTE: Female hamsters have a stable 4-days estrous cycle that is maintained immediately after parturition. Females on the first day of estrous can be identified by the opaque, yellowish and sticky vaginal discharge (Figure 2a). Females on Day 4 of estrous can be identified by the transparent, sticky mucus (Figure 2b).</li>
	<li>For superovulation of donor hamster, superovulate the sexually mature females (&#62;6 weeks) on the first day of the estrous cycle by IP injection of pregnant mare&#39;s serum gonadotropin (PMSG; dissolved in PBS as 50 IU/mL) (Table 1) between 9 - 12 AM. 
	NOTE: PMSG is for inducing superovulation but not for estrous cycle synchronization.</li>
	<li>80 - 84 h after injection of PMSG (Day 4, 6 - 8 PM), place the females in cages with males for mating. As hamster matings do not result in copulation plugs, ensure successful matings by watching the matings.</li>
	<li>Prepare pseudopregnant females by mating sexually mature females on Day 4 of estrous with vasectomized males. The time schedule for preparing donor and recipient females are shown in Table 2. 
	NOTE: True pregnant females can also be used as recipients, i.e., females with a coat color distinguishable from the golden color matted with the same color fertile males. Pups derived from embryo transfer can be identified based on coat colors. A potential advantage in using true pregnant females is that endogenously produced embryos would ensure successful pregnancies and help PN injected embryos to implant; a potential drawback in using true pregnant females as recipients is that PN injected embryos need to compete with endogenously produced embryos for implantation.</li>
</ol>
4. Zygote Isolation
<ol>
	<li>Prepare culture medium (HECM-9; recipe is described in Supplement 2), as described previously in McKiernan and Bavister12.</li>
	<li>One day prior to PN injection, prepare zygote handling dishes (25 &#181;L/drop) and a PN injection drop (100 &#181;L) in the lid of a 35 mm dish with HECM-9. Then cover the medium drops with mineral oil. Balance medium in an incubator overnight under the following conditions: 37.5 &#176;C, 10% CO2, 5% O2, 85% N2 and 100% humidity.</li>
	<li>On the day scheduled for PN injection, prepare embryo flushing dishes (1 dish/donor hamster) with HECM-9 and store all dishes in the incubator until use.</li>
	<li>Euthanize superovulated hamsters with CO2.</li>
	<li>Oviduct Isolation
	<ol>
		<li>Place a euthanized female hamster in dorsal recumbency on a surgical drape or tissue paper and prepare the abdomen with a 70% ethanol spray.</li>
		<li>Firmly hold skin and abdominal muscle layer at the midline and make an incision. Incise the peritoneum to expose the abdominal organs.</li>
		<li>Grasp one of the uterine horns with a fine forceps and retract the tissue from the abdomen. Separate the uterus from the mesometrium and fat tissue. Make a cut between the oviduct and ovary and a second cut adjacent to the junction of the oviduct and uterus (include a small part of uterus). Place the two oviducts in the same flush dish.</li>
	</ol>
	</li>
	<li>Flush and collect zygotes
	<ol>
		<li>Under the dissection microscope, grasp the side of uterine horn with a #5 Dumont forceps and insert a homemade needle (Figure 3) into the lumen of the oviduct from the infundibular end and flush the oviduct with 300 - 400 &#181;L of HECM-9 medium.</li>
		<li>Collect the zygotes immediately and wash them twice with HECM-9 medium in the handling dish. At this stage of development, all of the zygotes should be denuded from cumulus cells.</li>
		<li>Place the zygotes in an incubator (37.5 &#176;C, 10% CO2, 5% O2, 85% N2 and 100% humidity) immediately after collection to minimize exposure to light .</li>
	</ol>
	</li>
</ol>
5. PN Injection
<ol>
	<li>Thaw the Cas9/sgRNA RNP on ice and maintain the complex on ice during the entire PN injection process.</li>
	<li>Load the injection needles with Cas9/sgRNA RNP solution immediately before the injection experiments. Place the rear end of the injection needles into the tube containing the Cas9/sgRNA RNP solution and allow the Cas9/sgRNA RNP solution to fill the needle by capillary action.</li>
	<li>Prepare the injection dish and needles. Fill the holder pipette with mineral oil to within ~3 mm of the bend of the holder needle. Place the holder in the microinjector with the tip of the needle horizontal to the bottom of the dish and fill the tip of the needle with medium by suction. Set the injection needle at a 10 - 15&#176; angle opposite to holder.</li>
	<li>PN injection
	<ol>
		<li>Identify a zygote and apply a fine suction with the holder needle. Ideally, the male and female pronuclei are within the same focal range and aligned parallel to holder.</li>
		<li>Penetrate the zona and male pronuclei (3 o&#39;clock position) with the injection needle.</li>
		<li>Withdraw the injection needle quickly once the pronucleus swells to prevent the nucleus from adhering to the needle and to avoid damaging the pronucleus.</li>
	</ol>
	</li>
</ol>
6. Zygotes Transfer to Pseudopregnant Hamsters
<ol>
	<li>Anesthetize a pseudopregnant hamster (the same way as described in the vasectomy section above) and lay it on a surgical drape or dry tissue paper in ventral recumbency.</li>
	<li>Apply eye lubricants to the eyes of the animal to prevent eye dryness and cover the head with a cloth or paper to protect the eyes from light. Prepare the lateral aspect of the body by shaving a about 4 cm x 4 cm skin area on each sides of the body followed by applying 3 times of povidone-iodine scrubs and 3 times&#160; 70% ethanol scrubs. Make a 2 cm vertical incision 2 cm caudal to the last rib (as shown in Figures 4a, 4b) on each side of the back of the animal.</li>
	<li>Grasp the fat pad with forceps and pull gently until the oviduct and uterus are visible. Clamp the fat pad with a hemostats and reflect the fat pad dorsally over the spine (Figure 4c). Position the reproductive tract such that the uterine tube can be penetrated with a 30 gauge needle for embryo transfer (Figure 4d).</li>
	<li>Wash the injected zygotes with HECM-9 in a new dish. Use a new glass pipette (diameter: ~150 - 200 &#181;m) to load 10 - 15 zygotes. Arrange the zygotes as a chain of pearls followed by several air bubbles. Insert the pipette into the open penetrated tube from step 6.3 and gently blow into the pipette to transfer the zygotes to the oviduct. Continue blowing until the first air bubble is released into the oviduct tract (Figure 4e).</li>
	<li>Release the fat pad and return the tissue to the abdomen. Transfer approximately the same number (10 - 15) of injected embryos to another oviduct tract by the same processes.</li>
	<li>Suture the musculature and the skin in 2 layers. Administer Buprenorphine-HCL ( 0.5 mg/kg) subcutaneously as post-op analgesia. Return the hamster to a cage and place the cage on a warm pad for recovery. Return the cage to the animal room when the animal is fully recovered from anesthesia. Post-op analgesics are given again six hours later.</li>
</ol>

<ol>
	<li>Takahashi, M., Hori, M., Mutoh, M., Wakabayashi, K., Nakagama, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+animal+models+of+pancreatic+carcinogenesis+for+prevention+studies+and+their+relevance+to+human+disease.">Experimental animal models of pancreatic carcinogenesis for prevention studies and their relevance to human disease.</a> Cancers (Basel). 3 (1), 582-602 (2011).</li><li>Wold, W. S., Toth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+three--Syrian+hamster+as+an+animal+model+to+study+oncolytic+adenoviruses+and+to+evaluate+the+efficacy+of+antiviral+compounds.">Chapter three--Syrian hamster as an animal model to study oncolytic adenoviruses and to evaluate the efficacy of antiviral compounds.</a> Adv Cancer Res. , 69-92 (2012).</li><li>Thomas, M. A., 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=Syrian+hamster+as+a+permissive+immunocompetent+animal+model+for+the+study+of+oncolytic+adenovirus+vectors.">Syrian hamster as a permissive immunocompetent animal model for the study of oncolytic adenovirus vectors.</a> Cancer Res. 66 (3), 1270-1276 (2006).</li><li>Thomas, M. A., Spencer, J. F., Wold, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+the+Syrian+hamster+as+an+animal+model+for+oncolytic+adenovirus+vectors.">Use of the Syrian hamster as an animal model for oncolytic adenovirus vectors.</a> Methods Mol Med. , 169-183 (2007).</li><li>Hogarth, C. A., Roy, A., Ebert, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genomic+evidence+for+the+absence+of+a+functional+cholesteryl+ester+transfer+protein+gene+in+mice+and+rats.">Genomic evidence for the absence of a functional cholesteryl ester transfer protein gene in mice and rats.</a> Comp Biochem Physiol B Biochem Mol Biol. 135 (2), 219-229 (2003).</li><li>Ebihara, H., 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=A+Syrian+golden+hamster+model+recapitulating+ebola+hemorrhagic+fever.">A Syrian golden hamster model recapitulating ebola hemorrhagic fever.</a> J Infect Dis. 207 (2), 306-318 (2013).</li><li>Jove, M., 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=Lipidomic+and+metabolomic+analyses+reveal+potential+plasma+biomarkers+of+early+atheromatous+plaque+formation+in+hamsters.">Lipidomic and metabolomic analyses reveal potential plasma biomarkers of early atheromatous plaque formation in hamsters.</a> Cardiovasc Res. 97 (4), 642-652 (2013).</li><li>Vairaktaris, E., 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=The+hamster+model+of+sequential+oral+oncogenesis.">The hamster model of sequential oral oncogenesis.</a> Oral Oncol. 44 (4), 315-324 (2008).</li><li>Paciello, O., 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=Syrian+hamster+infected+with+Leishmania+infantum:+a+new+experimental+model+for+inflammatory+myopathies.">Syrian hamster infected with Leishmania infantum: a new experimental model for inflammatory myopathies.</a> Muscle Nerve. 41 (3), 355-361 (2010).</li><li>Safronetz, D., Ebihara, H., Feldmann, H., Hooper, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Syrian+hamster+model+of+hantavirus+pulmonary+syndrome.">The Syrian hamster model of hantavirus pulmonary syndrome.</a> Antiviral Res. 95 (3), 282-292 (2012).</li><li>Fan, Z., 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=Efficient+gene+targeting+in+golden+Syrian+hamsters+by+the+CRISPR/Cas9+system.">Efficient gene targeting in golden Syrian hamsters by the CRISPR/Cas9 system.</a> PLoS One. 9 (10), e109755 (2014).</li><li>McKiernan, S. H., Bavister, B. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culture+of+one-cell+hamster+embryos+with+water+soluble+vitamins:+pantothenate+stimulates+blastocyst+production.">Culture of one-cell hamster embryos with water soluble vitamins: pantothenate stimulates blastocyst production.</a> Hum Reprod. 15 (1), 157-164 (2000).</li><li>Takenaka, M., Horiuchi, T., Yanagimachi, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+light+on+development+of+mammalian+zygotes.">Effects of light on development of mammalian zygotes.</a> Proc Natl Acad Sci U S A. 104 (36), 14289-14293 (2007).</li></ol>

ZW has financial interests in Auratus Bio, LLC., a biotechnology company specialized in creating genetically engineered animals for biomedical research and agricultural applications.

To better exploit the potential of golden Syrian hamsters as models of human disease, we developed a PN injection protocol for delivering a CRISPR/Cas9 complex to target the hamster genome. The PN injection protocol optimizes several key variables including the embryo culture medium, temperature, and wavelengths of light13. There are also several hamster-specific animal handling procedures that need to follow for successfully conducting gene targeting in the hamster. For example, sexually matured ...

The golden Syrian hamster (Mesocricetus auratus) is one of the most widely used rodents for biomedical research. According to the U.S. Department of Agriculture, approximately 100,000 hamsters were used in the United States in 2015, representing 13% of total laboratory animal usage among the species covered by the Animal Welfare Act (http://www.aphis.usda.gov; accessed March 10, 2017).
The hamster offers several advantages over other rodents in the study of a number of human diseases. For example, the histopathology of N-nitrosobis(2-oxopropyl)amine (BOP) induced pancreatic ductal adenocarcinomas in hamster is similar to human pancreatic tumors, while BOP treatment mainly induces thyroid gland tumors in rats and lung and liver tumors in mice1. Because hamsters are the only small rodent found to support the replication of adenoviruses, they are also the model of choice for testing adenovirus-based oncolytic vectors and anti-adenovirus drugs2,3,4. Another example wherein the hamster model offers an advantage over mice and rats is in the study of hyperlipidemia. Humans and hamsters exhibit great similarities in lipid metabolic pathways and both species carry the gene encoding cholesteryl ester transfer protein (CETP), which plays a central role in lipid metabolisms, while CETP is absent in mice and rats5. Additionally, hamsters develop hemorrhagic disease more representative of the human manifestation following exposure to Ebola virus6. Hamsters are also the models of choice for studying atherosclerosis7, oral carcinomas8, and inflammatory myopathies9. Recently, it has also been demonstrated that hamsters are highly susceptible to Andes virus infection and develop hantavirus pulmonary syndrome-like disease, providing the only rodent model of Andes virus infection10.
To address the unmet need for novel genetic animal models to study the human diseases where no reliable small rodent model is available, we recently have succeeded in applying the CRISPR/Cas9 system to the hamster and have produced several lines of genetically engineered hamsters11. Hamster zygotes are highly sensitive to environmental milieus such that the PN injection protocols developed in other species are unsuitable. Therefore, we developed a PN injection protocol for the hamster that accommodates the special requirements for handling hamster embryos in vitro. Here, we describe the detailed PN injection procedure using the CRISPR/Cas9 system and the accompanying steps, from the preparation of single guide RNA (sgRNA) to the transfer of injected embryos into recipient females.

<table><tbody><tr><td>Cas9</td><td>Invitrogen</td><td>B25640</td><td>1 &amp;mu;g/&amp;mu;L (~6.1 &amp;mu;M)</td></tr><tr><td>GeneArtTM Precision Synthesis Kit</td><td>Invitrogen</td><td>A29377</td><td>For sgRNA synthesis</td></tr><tr><td>Albumin from human serum</td><td>Sigma</td><td>A1653</td><td>For cultivation medium</td></tr><tr><td>Illuminator</td><td>Nikon</td><td>NI-150</td><td>For embryo transfer</td></tr><tr><td>Incubator</td><td>New Brunswick</td><td>Galaxy 14S</td><td>For embryo cultivation</td></tr><tr><td>Microforge</td><td>Narishige</td><td>PB-7</td><td>For making injection needles</td></tr><tr><td>Microscope</td><td>Nikon</td><td>ECLIPSE Ti-S</td><td>For microinjection</td></tr><tr><td>Microscope</td><td>invitrogen</td><td>SMZ745T</td><td>For embryo transfer</td></tr><tr><td>Mineral oil</td><td>Sigma</td><td>M1840</td><td>Keep in dark</td></tr><tr><td>PMSG</td><td>Sigma</td><td>G4877-2000IU</td><td>For superovulation</td></tr></tbody></table>

production of genetically engineered golden syrian hamsters by pronuclear injection of the crispr/cas9 complex

The pronuclear (PN) injection technique was first established in mice to introduce foreign genetic materials into the pronuclei of one-cell stage embryos. The introduced genetic material may integrate into the embryonic genome and generate transgenic animals with foreign genetic information following transfer of the injected embryos to foster mothers. Following the success in mice, PN injection has been applied successfully in many other animal species. Recently, PN injection has been successfully employed to introduce reagents with gene-modifying activities, such as the CRISPR/Cas9 system, to achieve site-specific genetic modifications in several laboratory and farm animal species. In addition to mastering the special set of microinjection skills to produce genetically modified animals by PN injection, researchers must understand the reproduction physiology and behavior of the target species, because each species presents unique challenges. For example, golden Syrian hamster embryos have unique handling requirements in vitro such that PN injection techniques were not possible in this species until recent breakthroughs by our group. With our species-modified PN injection protocol, we have succeeded in producing several gene knockout (KO) and knockin (KI) hamsters, which have been used successfully to model human diseases. Here we describe the PN injection procedure for delivering the CRISPR/Cas9 complex to the zygotes of the hamster, the embryo handling conditions, embryo transfer procedures, and husbandry required to produce genetically modified hamsters.

The efficiency of the described protocol in producing genetically modified hamsters depends on the outcomes of the following two critical steps: the live birth rate of recipient females and the number of live pups with the intended genetic modifications. The live birth rate is a direct results of the embryo quality and the skill of the individual performing the PN injection and embryo transfer procedures. To ensure that the developmental potential of manipulated embryos is not compromised...

Watch this Scientific Journal Video about Production of Genetically Engineered Golden Syrian Hamsters by Pronuclear Injection of the CRISPR/Cas9 Complex at JoVE.com

Production of Genetically Engineered Golden Syrian Hamsters by Pronuclear Injection of the CRISPR/Cas9 Complex

Pronuclear (PN) injection of the clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated protein-9 nuclease (CRISPR/Cas9) system is a highly efficient method for producing genetically engineered golden Syrian hamsters. Herein, we describe the detailed PN injection protocol for the production of gene knockout hamsters with the CRISPR/Cas9 system.

The pronuclear (PN) injection technique was first established in mice to introduce foreign genetic materials into the pronuclei of one-cell stage embryos. ...

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Bioengineering

12.9K Views.  Utah State University. Pronuclear (PN) injectie van de geclusterde interspaced regelmatig korte palindromische herhaalt (CRISPR) en CRISPR-geassocieerde proteïne-9 nuclease (CRISPR/Cas9)-systeem is een zeer efficiënte methode voor de productie van genetisch gemanipuleerde gouden Syrische hamsters. Hierin beschrijven we het gedetailleerde PN injectie protocol voor de productie van gene knockout hamsters met het systeem CRISPR/Cas9.

Video: Production of Genetically Engineered Golden Syrian Hamsters by Pronuclear Injection of the CRISPR/Cas9 Complex

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

Genetics

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex determination, venom synthesis, and host-symbiont interactions, among others. A major limitation of working with this organism is the lack of effective protocols to perform directed genome modifications. An important part of genome modification is delivery of editing reagents, including CRISPR/Cas9 molecules, into embryos through microinjection. While microinjection is well established in many model organisms, this technique is particularly challenging to perform in N. vitripennis primarily due to its small embryo size, and the fact that embryonic development occurs entirely within a parasitized blowfly pupa. The following procedure overcomes these significant challenges while demonstrating a streamlined, visual procedure for effectively removing wasp embryos from parasitized host pupae, microinjecting them, and carefully transplanting them back into the host for continuation and completion of development. This protocol will strongly enhance the capability of research groups to perform advanced genome modifications in this organism.

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Microinjection of Nasonia vitripennis embryos is an essential method for generating heritable genome modifications. Described here is a detailed procedure for microinjection and transplantation of Nasonia vitripennis embryos, which will greatly facilitate future genome manipulation in this organism.

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex ...

Lentiviral Mediated Production of Transgenic Mice: A Simple and Highly Efficient Method for Direct Study of Founders

For almost 40 years, pronuclear DNA injection represents the standard method to generate transgenic mice with random integration of transgenes. Such a routine procedure is widely utilized throughout the world and its main limitation resides in the poor efficacy of transgene integration, resulting in a low yield of founder animals. Only few percent of animals born after implantation of injected fertilized oocytes have integrated the transgene. In contrast, lentiviral vectors are powerful tools for integrative gene transfer and their use to transduce fertilized oocytes allows highly efficient production of founder transgenic mice with an average yield above 70%. Furthermore, any mouse strain can be used to produce transgenic animal and the penetrance of transgene expression is extremely high, above 80% with lentiviral mediated transgenesis compared to DNA microinjection. The size of the DNA fragment that can be cargo by the lentiviral vector is restricted to 10 kb and represents the major limitation of this method. Using a simple and easy to perform injection procedure beneath the zona pellucida of fertilized oocytes, more than 50 founder animals can be produced in a single session of microinjection. Such a method is highly adapted to perform, directly in founder animals, rapid gain and loss of function studies or to screen genomic DNA regions for their ability to control and regulate gene expression in vivo.

Here, we present a protocol to promote transgene integration and production of founder transgenic mice with high efficacy by a simple injection of a lentiviral vector in the perivitelline space of a fertilized oocyte.

For almost 40 years, pronuclear DNA injection represents the standard method to generate transgenic mice with random integration of transgenes. Such a ...

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

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how genes regulate social traits, we have developed mutant ants via delivery of CRISPR complex into young embryos during their syncytial stage. Here, we provide a protocol of CRISPR-mediated mutagenesis in Harpegnathos saltator, a ponerine ant species that displays striking phenotypic plasticity. H. saltator ants are readily reared in a laboratory setting. Embryos are collected for microinjection with Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs) using home-made quartz needles. Post-injection embryos are reared outside the colony. Following emergence of the first larva, all embryos and larvae are transported to a nest box with a few nursing workers for further development. This protocol is suitable for inducing mutagenesis for analysis of caste-specific physiology and social behavior in ants, but may also be applied to a broader spectrum of hymenopterans and other insects.

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Many characteristics of insect eusociality rely on within-colony communication and division of labor. Genetic manipulation of key regulatory genes in ant embryos via microinjection and CRISPR-mediated mutagenesis provides insights into the nature of altruistic behavior in eusocial insects.

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how ...

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 ...

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 ...

High-throughput Parallel Sequencing to Measure Fitness of Leptospira interrogans Transposon Insertion Mutants During Golden Syrian Hamster Infection

In this manuscript, we describe a transposon sequencing (Tn-Seq) technique to identify and quantify Leptospira interrogans mutants altered in fitness during infection of Golden Syrian hamsters. Tn-Seq combines random transposon mutagenesis with the power of high-throughput sequencing technology. Animals are challenged with a pool of transposon mutants (input pool), followed by harvesting of blood and tissues a few days later to identify and quantify the number of mutants in each organ (output pools). The output pools are compared to the input pool to evaluate the in vivo fitness of each mutant. This approach enables screening of a large pool of mutants in a limited number of animals. With minor modifications, this protocol can be performed with any animal model of leptospirosis, reservoir host models such as rats and acute infection models such as hamsters, as well as in vitro studies. Tn-Seq provides a powerful tool to screen for mutants with in vivo and in vitro fitness defects.

High-throughput Parallel Sequencing to Measure Fitness of Leptospira interrogans Transposon Insertion Mutants During Golden Syrian Hamster Infection

We describe here a technique that combines transposon mutagenesis with high-throughput sequencing to identify and quantify transposon leptospiral mutants in tissues after a challenge of hamsters. This protocol can be used to screen mutants for survival and dissemination in animals and can also be applied to in vitro studies.

In this manuscript, we describe a transposon sequencing (Tn-Seq) technique to identify and quantify Leptospira interrogans mutants altered in fitness ...

Immunology and Infection

Removal of an Internal Translational Start Site from mRNA While Retaining Expression of the Full-Length Protein

The CRISPR-Cas9 gene-editing system, based on genome repair mechanisms, enables the generation of gene-modified mouse models more quickly and easily relative to traditional homologous recombination. The CRISPR-Cas9 system is particularly attractive when a single-point mutation is desired. The gap junction protein, Connexin 43 (Cx43), is encoded by gene Gja1, which has a single coding exon and cannot be spliced. However, Gja1 produces not only full-length Cx43 protein but up to six N-terminus truncated isoforms by a process known as internal translation, the result of ribosomal translation initiation at internal AUG (Methionine) start sites. GJA1-20k is the most commonly generated truncated isoform of Cx43 initiated at the AUG codon at position 213 of Gja1 mRNA. Because residue 213 occurs at the end of the last transmembrane domain of Cx43, GJA1-20k is effectively the 20 kDa C-terminus tail of Cx43 as an independent protein. Previous investigators identified, in cells, that a critical role of GJA1-20k is to facilitate trafficking of full-length Cx43 gap junction hemichannels to the plasma membrane. To examine this phenomenon in vivo, a mutant mouse with a Gja1 point-mutation was generated that replaces the ATG (Methionine) at residue 213 with TTA (Leucine, M213L mutation). The result of M213L is that Gja1 mRNA and full-length Cx43 are still generated, yet the translation of Gja1-20k is significantly reduced. This report focuses on choosing the restriction enzyme site to develop a one amino acid mutated (Gja1M213L/M213L) mouse model. This protocol describes genetically modified mice by the CRISPR-Cas9 system and rapid genotyping by combining PCR and restriction enzyme treatments.

The present protocol describes a single M213L mutation in Gja1 that retains full-length Connexin43 generation but prevents translation of the smaller GJA1-20k internally translated isoform.

The CRISPR-Cas9 gene-editing system, based on genome repair mechanisms, enables the generation of gene-modified mouse models more quickly and easily ...

Production of Genetically Engineered Golden Syrian Hamsters by Pronuclear Injection of the CRISPR/Cas9 Complex

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