Name: microinjection of crispr/cas9 protein into channel catfish, ictalurus punctatus, embryos for gene editing
Uploaded: 2018-01-20T13:00:00.000+00:00
Duration: 16 min 8 s
Description: Watch this Scientific Journal Video about Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing at JoVE.com

This research was funded by USDA-NIFA award 2015-67015-23488 to Roger Cone. The authors thank Dr. Ronald Phelps for the description of brood stock selection criteria in the video. Ahmed Elaswad and Karim Khalil would like to thank the Egyptian Cultural and Educational Bureau in Washington DC for funding their Ph.D. research.

This experiment was conducted at the Fish Genetics Research Unit, E. W. Shell Fisheries Research Center, Auburn University, AL. All experimental protocols used in this experiment were approved by the Auburn University Institutional Animal Care and Use Committee (AU-IACUC) before the experiment was initiated. A list of the equipment and supplies used in this experiment can be found in the Materials Table. The following are the steps and procedures for preparation and microinjection of channel catfish one-cell embryos as illustrated in Figure 1.
1. Brood Stock Selection and Spawning
<ol>
	<li>Choose healthy channel catfish brood stock from the strain or genetic line of interest. Channel catfish males and females should exhibit good secondary sex characteristics.</li>
	<li>Select males that have a broad muscular head that is wider than the rest of their bodies with well-developed genital papilla. Dark color, scarring and wounds from territorial fighting are also signs of sexual readiness. Select females that have heads that are narrower than the rest of their bodies, and have soft, palpable, well-rounded abdomen. For accuracy and to avoid stressing the fish during initial handling, stop feeding the females 2-3 days before examining their readiness for spawning.</li>
	<li>Perform fish handling quickly, but carefully. To reduce handling stress, perform all procedures that require handling females while holding fish in spawning bags (32 mm size mesh).</li>
	<li>Just before hormone injection to induce ovulation, measure the body weight of the females.</li>
	<li>Hang the spawning bags with females inside in a flow through tank with continuous water flow and adequate aeration.</li>
	<li>Aseptically, load the implanter that has a 14 G needle with 85 &#181;g/kg of luteinizing hormone releasing hormone analog (LHRHa) implant. Rinse the injection site (dorsal musculature) with 0.9% sterile saline solution before the injection. Insert the implanter at a 45&#176; angle and release the implant. Withdraw the needle and swipe the injection site with a cotton ball previously soaked in 70% ethanol.</li>
	<li>Place the spawning bag in the holding tank so that the fish is completely submerged in water 15-20 cm below the water surface. Adequate aeration (above 5 ppm dissolved oxygen) and good water quality are important for ovulation of high quality eggs.</li>
	<li>Predict the ovulation time based on the water temperature using the degree-hour (h) relationship35. Degree-h response time = time between hormone injection and ovulation (h) * water temperature (&#176;C). For example, for a degree-h response time of 900-1,000, a female is expected to spawn at 36-40 h from hormone injection time at 25 &#176;C. Usually, the first check for eggs is done at about 36 h after hormone injection and then at 4 h interval.</li>
	<li>To check for ovulation, gently lift the spawning bag above the water surface while you are looking for the eggs that are attached inside the spawning bag. Seeing at least 10 eggs indicates this female is ovulating and ready for hand stripping.</li>
</ol>
2. Sperm Preparation
NOTE: Sperm can be prepared a few h before the expected ovulation time.
<ol>
	<li>Euthanize males by a percussive blow to the head without anesthesia, since anesthesia might have a negative effect on sperm motility.</li>
	<li>Collect the testes in a small weighing pan, remove any tissue, and wash them with 50-mL of 0.9% saline to remove blood, if necessary. Well-developed testes are white with many villiform projections ( Figure 2).</li>
	<li>Determine the weight of the testes.</li>
	<li>To prepare the sperm solution for egg fertilization, transfer the testes to the center of a 100 cm2 100 &#181;m mesh using forceps. Fold the mesh with the testes inside, crush the testes and filter the sperm into a 50 mL collection tube. This can be done by applying pressure to testes inside the mesh against the rim of the collection tube via the thumb. Use 0.9% saline solution to wash the sperm from the mesh into the collection tube. Saline solution can be added up to 10 mL/g of testes.</li>
	<li>Determine the concentration of the sperm, check the motility and viability. 
	NOTE: Concentration can be determined by counting the sperm cells in a known diluted volume of milt using a hemocytometer. Alternatively, the sperm density can be estimated by evaluating the optical density of milt using a spectrophotometer. In this method, the absorption at 505 nm wavelength is compared to a control chart that has been previously prepared for the absorption of different concentrations of sperm36. Sperm motility can be checked under the microscope as the time from activation of the sperm until cessation of sperm movement (called motility duration)37. The viability of sperm can be determined by staining the sperm with selective dyes such as trypan blue, which will stain only non-viable sperm cells, while viable sperm will remain unstained. However, for these microinjection procedures, this is not necessary if the testes are well developed.</li>
	<li>Store sperm in 0.9% saline from step 2.5 at 4 &#176;C and use within 24 h after preparation.</li>
</ol>
3. Egg Collection and Fertilization
<ol>
	<li>Apply a very thin layer of vegetable shortening to a 20 cm diameter clean dry spawning pan.</li>
	<li>Place the female in the anesthetic solution containing 100 ppm of buffered tricaine methane sulfonate with sodium bicarbonate (pH of final solution should be 7.0) until the fish is completely anesthetized38. 
	CAUTION: Tricaine methane sulfonate may be irritating if inhaled, ingested or absorbed through the skin.</li>
	<li>Carefully, remove the fish from the spawning bag and dip it in 0.9% saline to wash off the anesthetic solution. Dry the fish off completely using a clean towel, avoiding the removal of the mucus layer on the fish body surface.</li>
	<li>Apply vegetable shortening to the area around the vent, including the pelvic fins, to prevent attachment of the eggs during hand stripping.</li>
	<li>Hand strip the eggs into the greased spawning pan by applying gentle pressure to the ovaries. Good eggs should flow freely, be golden yellow in color, and have minimal or no clumps or blood clots (Figure 2). Avoid contact between eggs and water before fertilization, as water can stimulate micropyle closure.</li>
	<li>To fertilize the eggs for microinjection, transfer 200&#8211;300 eggs to a greased spawning pan. Greased plastic plant labels can be used as a spoon to transfer eggs to the greased spawning pan. Add 1&#8211;2 mL of the sperm solution (from step 2.5) to the eggs and mix gently. The ratio of sperm to egg should be approximately 11,000 sperm/egg.</li>
	<li>Add freshwater to the eggs to activate the sperm and eggs. Add enough water to slightly cover the eggs. Gently swirl the eggs for 30 s. Fertilization should occur in 1&#8211;2 min. It is important to fertilize eggs which are arranged in a single layer. Catfish eggs adhere to each other making it difficult to microinject multiple layers of embryos.</li>
	<li>Add more freshwater to the fertilized eggs and allow the eggs to harden for 10&#8211;15 min.</li>
	<li>Cover the spawning pan containing the remaining unfertilized eggs by a wet towel to prevent drying and fertilize over a few hours. Fertilization can be done in a staggered manner, for example, one batch of 200&#8211;300 eggs can be fertilized every 30&#8211;60 min to ensure a continuous supply of embryos at the one cell stage for microinjection and control non-injected embryos. Avoid contact between the wet towel and the eggs as the eggs may adhere to the wet towel making it difficult to remove them. In this protocol, eggs fertilized within 2&#8211;3 h after stripping were efficiently microinjected and had similar success as eggs fertilized immediately after stripping.</li>
</ol>
4. Needle Pulling and Loading
<ol>
	<li>Pull a 1.0 mm OD borosilicate glass capillary into 2 needles (Figure 3). Store the needles in a paper box with a piece of sponge in which several incisions have been made. To avoid breaking the needle, the diameter of the sponge should be smaller than the length of the needle stem.</li>
	<li>Open the needle tip by breaking a small piece of the needle&#39;s thinnest region using a sharp object. Alternatively, open the needle by pinching off the thinnest region of the tip with forceps under the microscope.</li>
	<li>Prepare the injection solution by mixing gRNA(s) with Cas9 protein. Other types of RNA or protein can also be injected with the same procedure. In this protocol, gRNAs were designed to target two of the channel catfish immune related genes, toll/interleukin 1 receptor domain-containing adapter molecule (TICAM1) gene and rhamnose binding lectin (RBL) gene and prepared according to Shah et al. with the use of a high fidelity Taq polymerase32,39. The genomic targets for gRNAs were 5&#39; GGA GGT GAA GCA CGT CGA GGA 3&#39; for TICAM 1 gene (Figure 4) and 5&#39; GGA CTT TGA GTC GGA GAA GTG G 3&#39; for exon 1 of RBL gene with protospacer adjacent motif (PAM) sequence underlined (Figure 6).
	<ol>
		<li>BLAST guide RNA targets against the channel catfish genome sequence and select those with no potential off-target sites.</li>
		<li>Add phenol red to color the gRNA/Cas9 protein mix up to one third of the total volume.</li>
		<li>Adjust the final concentration of gRNA/Cas9 protein mix with nuclease free water so that each embryo is injected with 50 nL containing approximately 2.5 ng gRNA and 7.5 ng Cas9 protein. Incubate the mixture for 10 min on ice before use.</li>
	</ol>
	</li>
	<li>With a microloader, load 5&#8211;10 &#181;L of the gRNA/Cas9 mixture into the injection needle by inserting the microloader into the needle stem and expelling the mixture slowly while retracting the microloader tip (Figure 3). Avoid trapping air bubbles inside.</li>
	<li>Attach the needle to the micropipette holder and ensure a tight connection, and then attach the holder to the micromanipulator. Ensure free stable movement. Apply pressure for microinjection by opening the nitrogen cylinder and adjusting the pressure regulator. 
	NOTE: The volume of injection can be affected by the pressure, diameter of the needle aperture, and duration of injection. To determine the volume to inject, inject into a drop of mineral oil placed on a hemocytometer. If needed, the pressure, duration of injection, and needle diameter can be adjusted to increase or decrease the volume of the injection. Inject multiple times into the mineral oil to ensure consistent volume.</li>
</ol>
5. Microinjection of Catfish Embryos
<ol>
	<li>Apply a very thin layer of vegetable shortening to a 100 mm clean Petri dish.</li>
	<li>Using a greased plastic plant label, transfer 50&#8211;100 eggs from the fertilization pan to the Petri dish and cover them with Holtfreter&#39;s solution (Materials Table). Align the eggs against each other in a single layer. This alignment should hold the eggs in place during the microinjection process.</li>
	<li>Place the Petri dish with the eggs on the stage of the microscope and hold it with one hand. Lower the needle with the other hand (Figure 3) until it pierces the chorion and the yolk in a single smooth movement. The tip of the needle should be as close as possible to the blastodisc before delivering the injection material.
	<ol>
		<li>Depress the pedal to deliver the injection material into the yolk. If the blastodisc is not clearly visible, then the injection material can be spread into different locations in the yolk. To do that, the needle is inserted to the far end of the yolk, then depressing the pedal and withdrawing the needle are done simultaneously to spread the injection material through different areas of the yolk. Up to 50 nanoliters can be injected into the yolk of channel catfish embryos, however, the amount of gRNA/Cas9 protein needs to be optimized for each gene to achieve the best results for mutation rate and embryo survival32.</li>
	</ol>
	</li>
	<li>Retract the needle smoothly so as not to damage the egg. Move the Petri dish to inject another egg with the same procedure. Avoid movement of the Petri dish during the injection process as this may damage the egg or break the needle. Remove eggs that are ruptured or damaged due to microinjection. Injection can be started at 15&#8211;20 min after fertilization and continue as long as the embryos are still in the one cell stage (until about 60-90 min after fertilization).</li>
	<li>Place injected embryos back in Holtfreter&#39;s solution with 10 ppm of doxycycline and continuous gentle aeration for 6&#8211;7 days until hatching40. Change the solution daily and remove dead embryos.</li>
</ol>
6. Mutation Detection
<ol>
	<li>Randomly, collect several injected embryos at 72 h post fertilization, remove the yolk and extract genomic DNA. Include 3&#8211;5 non-injected embryos as a control. 
	NOTE: Genomic DNA can be extracted from embryos after removal of egg shells and yolk material using proteinase K digestion and ethanol precipitation32.</li>
	<li>Amplify a DNA segment flanking the genomic targets for gRNA of each gene, where mutations are expected to occur, using a high fidelity Taq polymerase32. For TICAM 1, the primers 5&#39;GCTGCTGAATGTCTGATTATG 3&#39; (forward) and 5&#39;GTCCTCCACACTCCTGAAG 3&#39; (reverse) were used to amplify a 750 bp while for RBL, the primers 5&#39;TCATTATTTCCTGGTACTCAGTA 3&#39; (forward) and 5&#39;AGAGATTAGTCACACATCATTATT 3&#39; (reverse) were used to amplify a 375 bp segment from the RBL gene.
	<ol>
		<li>Use the following PCR reaction: up to 20 &#181;L PCR grade water; 1x high fidelity Taq enzyme reaction buffer with MgCl2, 0.2 mM dNTP mix, 0.4 &#181;M for each of the forward and reverse primer of the same set, 100-300 ng genomic DNA and 1.25 units of high fidelity Taq enzyme blend. Perform PCR cycling conditions: initial denaturation at 94 &#176;C for 3 min; 35 cycles of denaturation at 94 &#176;C for 30 s, annealing at 60 &#176;C for 55 s for TICAM, and 40 s for RBL, extension at 72 &#176;C for 1 min/kb; and final extension at 72 &#176;C for 10 min.</li>
	</ol>
	</li>
	<li>Detect the mutant embryos using a mutation detection method such as sequencing of purified PCR products, surveyor mutation detection assay, heteroduplex mobility assay, loss of restriction enzyme recognition site, or any other mutation detection method that is appropriate for the target. In this protocol, detect mutant embryos using surveyor mutation detection assay for TICAM 1 and loss of restriction enzyme SapI cut site as well as sequencing of purified PCR products for RBL gene.</li>
	<li>To identify mutant alleles, clone PCR products and sequence vectors from individual colonies. The mutant alleles presented in Figure 4 and Figure 6 came from injected embryos. PCR products from six microinjected and 3 control embryos for each gene were pooled in two separate tubes, one for microinjected embryos and the other for control embryos, rapidly cloned and 86 vectors for TICAM 1 and 48 vectors for RBL gene sequenced, including 8 sequencing reactions from 3 control non-injected embryos for each gene.</li>
	<li>To identify different mutant alleles, compare the sequences from mutant embryos to the wild type sequence using any DNA sequence alignment tool such as BLAST or T-COFFEE.</li>
</ol>

The authors declare that they have no conflict of interest.

A detailed protocol for microinjection of channel catfish embryos to achieve gene knockout was presented. Injection of CRISPR/Cas9 protein in the yolk is much simpler, saves time, and does not require extensive training when compared to microinjecting the blastodisc of the one-cell embryo, which is the common technique for most gene transfer and gene editing with fish30,42. The current protocol proved to be successful in inducing indels in channel catfish TICAM 1...

Microinjection is a common laboratory technique used to deliver a small amount of a substance such as DNA, RNA, protein, and other macromolecules into cells or embryos through a glass capillary1. Microinjection is performed using special equipment setup including a microinjector, micromanipulator, and a microscope2. The technique has been used by researchers to genetically modify many organisms through the generation of transgenics, gene knockouts, and gene therapy, with the aim of understanding the dynamics of intracellular components3,4,5.
The channel catfish (Ictalurus punctatus) is the most popular catfish species in aquaculture and recreational fishing activities in the United States. There is an increasing need to study functional genomics in channel catfish, and sequencing of the complete genome of channel catfish enhances the utility of recent advances in genome editing tools6,7. Understanding gene function would not only enrich the research that is being conducted on catfish, but could also lead to more effective genetic improvement programs to enhance the catfish industry. Once critical genes for a given trait of interest are identified, they can be used to genetically improve catfish production through genome editing to generate beneficial alleles, selection for these alleles, suppressing the detrimental alleles, transferring the beneficial alleles through transgenesis, or some combination of these options. Combining the best genes for different commercially beneficial traits from different species in one fish would greatly improve productivity and profitability of fish production operations8.
Gene knockout is a direct method to study gene functions in vivo. Mutations at the DNA level will be inherited by future generations which will facilitate the study of their effects in different generations. Different genome editing tools have been developed including zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) system9,10,11,12.
The CRISPR/Cas9 system is a powerful, efficient tool that has been used to edit genomic DNA sequences including gene knockout in fish through RNA-guided site-specific DNA cleavage5,13. The system consists of a guide RNA (gRNA), which determines the targeted sequence in the genome and a DNA endonuclease enzyme, Cas9. The CRISPR/Cas9 system can be designed to target any sequence in the genome with several advantages over ZFNs and TALENs: (1) lower cost (2) easier engineering (3) more specific binding of guide RNA to the target sequence, and reduced off-target mutations (4) multiple sequences can be targeted with different gRNA at the same time (5) high mutagenesis rate in genes that could not be mutated by TALENs, and (6) improved germline transmission rate of mutations for up to 6-fold when compared to ZFNs and TALENs14,15,16,17,18.
The main alternative method for microinjection is electroporation, in which electric impulses are applied to embryos or cells to increase membrane permeability and the uptake of biological molecules19,20. Several transgenic fish were generated using electroporation such as medaka (Oryzias latipes), zebrafish (Danio rerio), chinook salmon (Oncorhynchus tshawytscha), channel catfish, sea bream (Sparus sarba), and common carp (Cyprinus carpio)21,22,23,24,25,26.
Electroporation has been used to deliver plasmid DNA, RNA, and Cas9 protein for gene knockout. In mammalian cells, Cas9/gRNA plasmid DNA, Cas9 mRNA/gRNA, and Cas9 protein/gRNA complexes were delivered using electroporation and the mutagenesis rates were highest using Cas9 protein/gRNA complexes for most electroporation conditions tested27. In the ascidian chordate (Ciona intestinalis), TALENs expressing constructs were electroporated into fertilized eggs to induce knockout of multiple genes28. ZFNs expressing plasmid constructs were electroporated to knock out luteinizing hormone in channel catfish29. However, once introduced into the cell, plasmid constructs will need to be transcribed to RNA and translated to functional proteins before they can target the desired DNA sequence, which would likely delay the time of mutagenesis compared to microinjection of RNA/protein. As an embryo is developing, delayed mutagenesis increases the mosaicism of injected founders. In addition, transgenic expression is unlikely to achieve the expression level possible with microinjection, lowering the mutagenesis efficiency.
As a means for introducing genome editing tools into cells, microinjection has several advantages over electroporation. Genome editing molecules can be reliably introduced into cells or embryos through microinjection30. Less injection material is needed. It is easy to determine the amounts of material injected. Higher mutation rates with low mosaicism in the founder fish can be achieved which would improve germline transmission of mutations to F1. Fewer founder fish need to be analyzed to produce the first generation, due to the higher mutation rate. In electroporation, more founder fish need to be analyzed which will increase costs. However, the survival of channel catfish microinjected embryos is lower than electroporated ones, though this can be overcome by injecting more embryos31,32. In channel catfish, survival of yolk-microinjected embryos ranged from 16 to 55%, depending on the genes being targeted and the dosage of gRNA/Cas9 protein32.
Regular microinjection of catfish embryos is technically demanding and time consuming, however, a rapid and efficient CRISPR/Cas9 protein microinjection protocol is presented for channel catfish embryos. This protocol requires less time and expertise since the CRISPR/Cas9 protein solution is injected into the yolk of one cell embryos. Hundreds of fertilized eggs can be injected in 1 h (approximately the time needed for the first cell division to occur). To validate the protocol, two disease susceptibility-related genes were knocked out in channel catfish, the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM1) gene and the rhamnose binding lectin (RBL) gene. TICAM 1 is involved in the signaling pathway initiated by toll-like receptor (TLR) 3. In channel catfish, TICAM 1 was dramatically upregulated following bacterial challenge with Edwardsiella ictaluri, while it was downregulated in blue catfish, a species resistant to E. ictaluri33. RBL plays an important role in early infection with Flavobacterium columnare, the causative agent of columnaris disease, where acute and robust upregulation was recorded in a columnaris-susceptible channel catfish strain compared to a columnaris-resistant strain34.

<table><tbody><tr><td>Reproboost implant</td><td>Center of Marine Biotechnology</td><td></td><td>Luteinizing hormone releasing hormone analog (LHRHa) for induction of ovulation in channel catfish females</td></tr><tr><td>TRICAINE-S</td><td>Western Chemical. Inc.</td><td></td><td>For sedation of brood stock fish during hormone injection and egg stripping.&amp;nbsp;</td></tr><tr><td>Phenol red</td><td>Sigma-Aldrich</td><td>P0290</td><td>0.5%, sterile filtered</td></tr><tr><td>Stereo microscope</td><td>Olympus</td><td>213709</td><td>For visualizing the eggs during microinjection</td></tr><tr><td>Microinjector</td><td>ASI-Applied Scientific Instrumentation&amp;nbsp;</td><td>Model MPPI-3</td><td>For the delivery of the injection material into the embryos</td></tr><tr><td>Micromanipulator</td><td>ASI-Applied Scientific Instrumentation&amp;nbsp;</td><td>Model MM33&amp;nbsp;</td><td>For holding and controlling the movement of the injection needle.&amp;nbsp;</td></tr><tr><td>Eppendorf Microloader</td><td>Eppendorf</td><td>5242956.003</td><td>For loading injection solution into microinjection needles.</td></tr><tr><td>Vertical needle puller</td><td>David Kopf Instruments</td><td>Model 720</td><td>For pulling microinjection needles</td></tr><tr><td>Cas9 protein</td><td>PNA Bio Inc.&amp;nbsp;</td><td>CP01</td><td>Recombinant Cas9 protein from Streptococcus pyrogenes.</td></tr><tr><td>Expand High FidelityPLUS PCR System</td><td>&amp;nbsp;Roche Diagnostics, USA</td><td></td><td>For PCR and amplification of DNA templates to be used in gRNA preparation</td></tr><tr><td>Borosilicate glass capillaries</td><td>Fisher Scientific</td><td></td><td>1 mm outer diameter (OD), for making microinjection needles.</td></tr><tr><td>Petri dish</td><td>VWR</td><td>25384-302</td><td>For holding the embryos during the microinjection.</td></tr><tr><td>Crisco</td><td>The J.M. Smucker Company</td><td></td><td>Vegetable shortening for coating spawning pans and petri dishes.&amp;nbsp;</td></tr><tr><td>Holtfreter`s solution</td><td>Home Made</td><td></td><td>59 mM NaCl, 0.67 mM KCl, 2.4 mM NaHCO3, 0.76 mM CaCl2, 1.67 mM MgSO4 to incubate the microinjected embryos till hatch.&amp;nbsp;</td></tr><tr><td>Doxycycline hyclate USP (monohydrate)</td><td>Letco Medical</td><td>690904</td><td>Antibiotic added to Holtfreter&#039;s solution to 10 ppm to prevent bacterial infections.</td></tr></tbody></table>

microinjection of crispr/cas9 protein into channel catfish, ictalurus punctatus, embryos for gene editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene function. Gene knockout has been used to study these gene functions in vivo. The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system is a powerful tool used to edit genomic DNA sequences to alter gene function. While the traditional approach has been to introduce CRISPR/Cas9 mRNA into the single cell embryos through microinjection, this can be a slow and inefficient process in catfish. Here, a detailed protocol for microinjection of channel catfish embryos with CRISPR/Cas9 protein is described. Briefly, eggs and sperm were collected and then artificial fertilization performed. Fertilized eggs were transferred to a Petri dish containing Holtfreter's solution. Injection volume was calibrated and then guide RNAs/Cas9 targeting the toll/interleukin 1 receptor domain-containing adapter molecule (TICAM 1) gene and rhamnose binding lectin (RBL) gene were microinjected into the yolk of one-cell embryos. The gene knockout was successful as indels were confirmed by DNA sequencing. The predicted protein sequence alterations due to these mutations included frameshift and truncated protein due to premature stop codons.

To demonstrate the efficiency of the microinjection protocol, gRNAs designed to target the channel catfish toll/interleukin 1 receptor domain-containing adapter molecule (TICAM1) gene and rhamnose binding lectin (RBL) gene were microinjected.
TICAM 1 
DNA sequencing of vectors from individual colonies from pooled, cloned PCR products revealed the indel mutations induced in TICAM 1 gene. The mutation rate wa...

Watch this Scientific Journal Video about Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing at JoVE.com

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes.

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

The complete genome of the channel catfish, Ictalurus punctatus, has been sequenced, leading to greater opportunities for studying channel catfish gene ...

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14.9K Views.  Auburn University. A simple and efficient microinjection protocol for gene editing in channel catfish embryos using the CRISPR/Cas9 system is presented. In this protocol, guide RNAs and Cas9 protein were microinjected into the yolk of one-cell embryos. This protocol has been validated by knocking out two channel catfish immune-related genes. 

Video: Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and behavioral phenotypes. The remarkably diverse phenotypes that have evolved as marine populations adapt to countless freshwater environments, combined with the ability to cross marine and freshwater forms, provide a rare vertebrate system in which genetics can be used to map genomic regions controlling evolved traits. Excellent genomic resources are now available, facilitating molecular genetic dissection of evolved changes. While mapping experiments generate lists of interesting candidate genes, functional genetic manipulations are required to test the roles of these genes. Gene regulation can be studied with transgenic reporter plasmids and BACs integrated into the genome using the Tol2 transposase system. Functions of specific candidate genes and cis-regulatory elements can be assessed by inducing targeted mutations with TALEN and CRISPR/Cas9 genome editing reagents. All methods require introducing nucleic acids into fertilized one-cell stickleback embryos, a task made challenging by the thick chorion of stickleback embryos and the relatively small and thin blastomere. Here, a detailed protocol for microinjection of nucleic acids into stickleback embryos is described for transgenic and genome editing applications to study gene expression and function, as well as techniques to assess the success of transgenesis and recover stable lines.

Transgenic manipulations and genome editing are critical for functionally testing the roles of genes and cis-regulatory elements. Here a detailed microinjection protocol for the generation of genomic modifications (including Tol2-mediated fluorescent reporter transgene constructs, TALENs, and CRISPRs) is presented for the emergent model fish, the threespine stickleback.

The threespine stickleback fish has emerged as a powerful system to study the genetic basis of a wide variety of morphological, physiological, and ...

Developmental Biology

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

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. ...

Determining the Role of Maternally-Expressed Genes in Early Development with Maternal Crispants

Early development depends on a pool of maternal factors incorporated into the mature oocyte during oogenesis that perform all cellular functions necessary for development until zygotic genome activation. Typically, genetic targeting of these maternal factors requires an additional generation to identify maternal-effect phenotypes, hindering the ability to determine the role of maternally-expressed genes during development. The discovery of the biallelic editing capabilities of CRISPR-Cas9 has allowed screening of embryonic phenotypes in somatic tissues of injected embryos or "crispants," augmenting the understanding of the role zygotically-expressed genes play in developmental programs. This article describes a protocol that is an extension of the crispant method. In this method, the biallelic editing of germ cells allows for the isolation of a maternal-effect phenotype in a single generation, or "maternal crispants." Multiplexing guide RNAs to a single target promotes the efficient production of maternal crispants, while sequence analysis of maternal crispant haploids provides a simple method to corroborate genetic lesions that produce a maternal-effect phenotype. The use of maternal crispants supports the rapid identification of essential maternally-expressed genes, thus facilitating the understanding of early development.

Early development is dependent on maternally-inherited products, and the role of many of these products is currently unknown. Herein, we described a protocol that uses CRISPR-Cas9 to identify maternal-effect phenotypes in a single generation.

Early development depends on a pool of maternal factors incorporated into the mature oocyte during oogenesis that perform all cellular functions necessary ...

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair templates and CRISPR-Cas reagents is the most common delivery method into embryos. However, this type of micromanipulation necessitates access to specialized equipment, is laborious, and requires a certain level of technical skill. Moreover, microinjection techniques often result in lower embryo survival due to the mechanical stress on the embryo. In this protocol, we developed an optimized method to deliver large DNA repair templates to work in conjunction with CRISPR-Cas9 genome editing without the need for microinjection. This protocol combines AAV-mediated DNA delivery of single-stranded DNA donor templates along with the delivery of CRISPR-Cas9 ribonucleoprotein (RNP) by electroporation to modify 2-cell embryos. Using this novel strategy, we have successfully produced targeted knock-in rat models carrying insertion of DNA sequences from 1.2 to 3.0 kb in size with efficiencies between 42% and 90%.

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus AAV and 2-Cell Embryo Electroporation

This protocol presents an optimized approach for producing genetically modified rat models. Adeno-associated virus (AAV) is used to deliver a DNA repair template, and electroporation is used to deliver CRISPR-Cas9 reagents to complete the genome editing process in 2-cell embryo.

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair ...

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 ...

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

Biology

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological phenomena. Zebrafish have been used as an effective genetic model to study numerous questions related to heritable disease, development, and toxicology at the whole-organ and -organism level. Due to the well-annotated and mapped zebrafish genome, numerous tools for gene editing have been developed. However, the efficacy of generating and ease of detecting precise knock-in edits using CRISPR is a limiting factor. Described here is a CRISPR-Cas9-based knock-in approach with the simple detection of precise edits in a gene responsible for cardiac repolarization and associated with the electrical disorder, Long QT Syndrome (LQTS). This two-single-guide RNA (sgRNA) approach excises and replaces the target sequence and links a genetically encoded reporter gene. The utility of this approach is demonstrated by describing non-invasive phenotypic measurements of cardiac electrical function in wild-type and gene-edited zebrafish larvae. This approach enables the efficient study of disease-associated variants in a whole organism. Furthermore, this strategy offers possibilities for the insertion of exogenous sequences of choice, such as reporter genes, orthologs, or gene editors.

This protocol describes an approach to facilitate precise knock-in edits in zebrafish embryos using CRISPR-Cas9 technology. A phenotyping pipeline is presented to demonstrate the applicability of these techniques to model a Long QT Syndrome-associated gene variant.

Clustered regularly interspaced short palindromic repeats (CRISPR) in animal models enable precise genetic manipulation for the study of physiological ...

Microinjection of CRISPR/Cas9 Protein into Channel Catfish, Ictalurus punctatus, Embryos for Gene Editing

Summary

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Chapters in this video

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Microinjection for Transgenesis and Genome Editing in Threespine Sticklebacks

Manipulation of Gene Function in Mexican Cavefish

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

Determining the Role of Maternally-Expressed Genes in Early Development with Maternal Crispants

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation

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

Efficient Genome Editing of Mice by CRISPR Electroporation of Zygotes

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

CRISPR-Cas9-Mediated Precise Knock-In Edits in Zebrafish Hearts