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 border="0" cellpadding="0" cellspacing="0" width="520">
	<tbody>
		<tr height="100">
			<td height="100" style="height:100px;width:136px;">Reproboost implant</td>
			<td style="width:115px;">Center of Marine Biotechnology</td>
			<td style="width:99px;"></td>
			<td style="width:171px;">Luteinizing hormone releasing hormone analog (LHRHa) for induction of ovulation in channel catfish females</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:136px;">TRICAINE-S</td>
			<td style="width:115px;">Western Chemical. Inc.</td>
			<td style="width:99px;"></td>
			<td style="width:171px;">For sedation of brood stock fish during hormone injection and egg stripping.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:136px;">Phenol red</td>
			<td style="width:115px;">Sigma-Aldrich</td>
			<td style="width:99px;">P0290</td>
			<td style="width:171px;">0.5%, sterile filtered</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:136px;">Stereo microscope</td>
			<td style="width:115px;">Olympus</td>
			<td style="width:99px;">213709</td>
			<td style="width:171px;">For visualizing the eggs during microinjection</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:136px;">Microinjector</td>
			<td style="width:115px;">ASI-Applied Scientific Instrumentation&#160;</td>
			<td style="width:99px;">Model MPPI-3</td>
			<td style="width:171px;">For the delivery of the injection material into the embryos</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:136px;">Micromanipulator</td>
			<td style="width:115px;">ASI-Applied Scientific Instrumentation&#160;</td>
			<td style="width:99px;">Model MM33&#160;</td>
			<td style="width:171px;">For holding and controlling the movement of the injection needle.&#160;</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:136px;">Eppendorf Microloader</td>
			<td style="width:115px;">Eppendorf</td>
			<td style="width:99px;">5242956.003</td>
			<td style="width:171px;">For loading injection solution into microinjection needles.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:136px;">Vertical needle puller</td>
			<td style="width:115px;">David Kopf Instruments</td>
			<td style="width:99px;">Model 720</td>
			<td style="width:171px;">For pulling microinjection needles</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:136px;">Cas9 protein</td>
			<td style="width:115px;">PNA Bio Inc.&#160;</td>
			<td style="width:99px;">CP01</td>
			<td style="width:171px;">Recombinant Cas9 protein from Streptococcus pyrogenes.</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:136px;">Expand High FidelityPLUS PCR System</td>
			<td style="width:115px;">&#160;Roche Diagnostics, USA</td>
			<td style="width:99px;"></td>
			<td style="width:171px;">For PCR and amplification of DNA templates to be used in gRNA preparation</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:136px;">Borosilicate glass capillaries</td>
			<td style="width:115px;">Fisher Scientific</td>
			<td style="width:99px;"></td>
			<td style="width:171px;">1 mm outer diameter (OD), for making microinjection needles.</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:136px;">Petri dish</td>
			<td style="width:115px;">VWR</td>
			<td style="width:99px;">25384-302</td>
			<td style="width:171px;">For holding the embryos during the microinjection.</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:136px;">Crisco</td>
			<td style="width:115px;">The J.M. Smucker Company</td>
			<td style="width:99px;"></td>
			<td style="width:171px;">Vegetable shortening for coating spawning pans and petri dishes.&#160;</td>
		</tr>
		<tr height="120">
			<td height="120" style="height:120px;width:136px;">Holtfreter`s solution</td>
			<td style="width:115px;">Home Made</td>
			<td style="width:99px;"></td>
			<td style="width:171px;">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.&#160;</td>
		</tr>
		<tr height="80">
			<td height="80" style="height:80px;width:136px;">Doxycycline hyclate USP (monohydrate)</td>
			<td style="width:115px;">Letco Medical</td>
			<td style="width:99px;">690904</td>
			<td style="width:171px;">Antibiotic added to Holtfreter&#39;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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14.8K 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

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the identification and verification of a drug resistance gene in the Apicomplexan parasite Toxoplasma gondii, the method of Quantitative Trait Locus (QTL) mapping using a Whole Genome Sequence (WGS) -based genetic map and the method of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9 -based gene editing. The approach of QTL mapping allows one to test if there is a correlation between a genomic region(s) and a phenotype. Two datasets are required to run a QTL scan, a genetic map based on the progeny of a recombinant cross and a quantifiable phenotype assessed in each of the progeny of that cross. These datasets are then formatted to be compatible with R/qtl software that generates a QTL scan to identify significant loci correlated with the phenotype. Although this can greatly narrow the search window of possible candidates, QTLs span regions containing a number of genes from which the causal gene needs to be identified. Having WGS of the progeny was critical to identify the causal drug resistance mutation at the gene level. Once identified, the candidate mutation can be verified by genetic manipulation of drug sensitive parasites. The most facile and efficient method to genetically modify T. gondii is the CRISPR/Cas9 system. This system comprised of just 2 components both encoded on a single plasmid, a single guide RNA (gRNA) containing a 20 bp sequence complementary to the genomic target and the Cas9 endonuclease that generates a double-strand DNA break (DSB) at the target, repair of which allows for insertion or deletion of sequences around the break site. This article provides detailed protocols to use CRISPR/Cas9 based genome editing tools to verify the gene responsible for sinefungin resistance and to construct transgenic parasites.

QTL Mapping and CRISPR/Cas9 Editing to Identify a Drug Resistance Gene in Toxoplasma gondii

Details are presented on how QTL mapping with a whole genome sequence based genetic map can be used to identify a drug resistance gene in Toxoplasma gondii and how this can be verified with the CRISPR/Cas9 system that efficiently edits a genomic target, in this case the drug resistance gene.

Scientific knowledge is intrinsically linked to available technologies and methods. This article will present two methods that allowed for the ...

Selection-dependent and Independent Generation of CRISPR/Cas9-mediated Gene Knockouts in Mammalian Cells

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide RNA (sgRNA) that confers specificity, the Cas9 protein cleaves both DNA strands at the targeted locus. The DNA break can trigger either non-homologous end joining (NHEJ) or homology directed repair (HDR). NHEJ can introduce small deletions or insertions which lead to frame-shift mutations, while HDR allows for larger and more precise perturbations. Here, we present protocols for generating knockout cell lines by coupling established CRISPR/Cas9 methods with two options for downstream selection/screening. The NHEJ approach uses a single sgRNA cut site and selection-independent screening, where protein production is assessed by dot immunoblot in a high-throughput manner. The HDR approach uses two sgRNA cut sites that span the gene of interest. Together with a provided HDR template, this method can achieve deletion of tens of kb, aided by the inserted selectable resistance marker. The appropriate applications and advantages of each method are discussed.

Recent advances in the ability to genetically manipulate somatic cell lines hold great potential for basic and applied research. Here, we present two approaches for CRISPR/Cas9 generated knockout production and screening in mammalian cell lines, with and without the use of selectable markers.

The CRISPR/Cas9 genome engineering system has revolutionized biology by allowing for precise genome editing with little effort. Guided by a single guide ...

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

Highly Efficient Gene Disruption of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. Complete gene ablation in HSCs required the generation of knockout mice from which HSCs could be isolated, and gene ablation in primary human HSCs was not possible. Viral transduction could be used for knock-down approaches, but these suffered from variable efficacy. In general, genetic manipulation of human and mouse hematopoietic cells was hampered by low efficiencies and extensive time and cost commitments. Recently, CRISPR/Cas9 has dramatically expanded the ability to engineer the DNA of mammalian cells. However, the application of CRISPR/Cas9 to hematopoietic cells has been challenging, mainly due to their low transfection efficiencies, the toxicity of plasmid-based approaches and the slow turnaround time of virus-based protocols.
A rapid method to perform CRISPR/Cas9-mediated gene editing in murine and human hematopoietic stem and progenitor cells with knockout efficiencies of up to 90% is provided in this article. This approach utilizes a ribonucleoprotein (RNP) delivery strategy with a streamlined three-day workflow. The use of Cas9-sgRNA RNP allows for a hit-and-run approach, introducing no exogenous DNA sequences in the genome of edited cells and reducing off-target effects. The RNP-based method is fast and straightforward: it does not require cloning of sgRNAs, virus preparation or specific sgRNA chemical modification. With this protocol, scientists should be able to successfully generate knockouts of a gene of interest in primary hematopoietic cells within a week, including downtimes for oligonucleotide synthesis. This approach will allow a much broader group of users to adapt this protocol for their needs.

A protocol for fast CRISPR/Cas9-mediated gene disruption in mouse and human primary hematopoietic cells is described in this article. Cas9-sgRNA ribonucleoproteins are introduced via electroporation with sgRNAs generated through in vitro transcription and commercial Cas9. High editing efficiencies are achieved with limited time and financial cost.

Advances in the hematopoietic stem cell (HSCs) field have been aided by methods to genetically engineer primary progenitor cells as well as animal models. ...

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical regions, is caused by infection with Plasmodium parasites. The development of more effective drugs to combat malaria can be accelerated by improving our understanding of the biology of this complex parasite. Genetic manipulation of these parasites is key to understanding their biology; however, historically the genome of P. falciparum has been difficult to manipulate. Recently, CRISPR/Cas9 genome editing has been utilized in malaria parasites, allowing for easier protein tagging, generation of conditional protein knockdowns, and deletion of genes. CRISPR/Cas9 genome editing has proven to be a powerful tool for advancing the field of malaria research. Here, we describe a CRISPR/Cas9 method for generating glmS-based conditional knockdown mutants in P. falciparum. This method is highly adaptable to other types of genetic manipulations, including protein tagging and gene knockouts.

CRISPR/Cas9 Gene Editing to Make Conditional Mutants of Human Malaria Parasite P. falciparum

We describe a method for generating glmS-based conditional knockdown mutants in Plasmodium falciparum using CRISPR/Cas9 genome editing.

Malaria is a significant cause of morbidity and mortality worldwide. This disease, which primarily affects those living in tropical and subtropical ...

Endogenous Protein Tagging in Human Induced Pluripotent Stem Cells Using CRISPR/Cas9

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or C-terminal fluorescent tags. The prokaryotic CRISPR/Cas9 system (clustered regularly interspaced short palindromic repeats/CRISPR-associated 9) may be used to introduce large exogenous sequences into genomic loci via homology directed repair (HDR). To achieve the desired knock-in, this protocol employs the ribonucleoprotein (RNP)-based approach where wild type Streptococcus pyogenes Cas9 protein, synthetic 2-part guide RNA (gRNA), and a donor template plasmid are delivered to the cells via electroporation. Putatively edited cells expressing the fluorescently tagged proteins are enriched by fluorescence activated cell sorting (FACS). Clonal lines are then generated and can be analyzed for precise editing outcomes. By introducing the fluorescent tag at the genomic locus of the gene of interest, the resulting subcellular localization and dynamics of the fusion protein can be studied under endogenous regulatory control, a key improvement over conventional overexpression systems. The use of hiPSCs as a model system for gene tagging provides the opportunity to study the tagged proteins in diploid, nontransformed cells. Since hiPSCs can be differentiated into multiple cell types, this approach provides the opportunity to create and study tagged proteins in a variety of isogenic cellular contexts.

Described here is a protocol for tagging endogenously expressed proteins with fluorescent tags in human induced pluripotent stem cells using CRISPR/Cas9. Putatively edited cells are enriched by fluorescence activated cell sorting and clonal cell lines are generated.

A protocol is presented for generating human induced pluripotent stem cells (hiPSCs) that express endogenous proteins fused to in-frame&#160;N- or ...

CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.

Presented here is a protocol for efficient CRISPR/Cas9 ribonucleoprotein-mediated gene editing in mammalian cells using tube electroporation.

Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of ...

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target against T. cruzi, it is important to validate the essentiality of the target gene in the mammalian stage of the parasite, the amastigote. Amastigotes of T. cruzi replicate inside the host cell; thus, it is difficult to conduct a knockout experiment without going through other developmental stages. Recently, our group reported a growth condition in which the amastigote can replicate axenically for up to 10 days without losing its amastigote-like properties. By using this temporal axenic amastigote culture, we successfully introduced gRNAs directly into the Cas9-expressing amastigote to cause gene knockouts and analyzed their phenotypes exclusively in the amastigote stage. In this report, we describe a detailed protocol to produce in vitro derived extracellular amastigotes, and to utilize the axenic culture in a CRISPR/Cas9-mediated knockout experiment. The growth phenotype of knockout amastigotes can be evaluated either by cell counts of the axenic culture, or by replication of intracellular amastigote after host cell invasion. This method bypasses the parasite stage differentiation normally involved in producing a transgenic or a knockout amastigote. Utilization of the temporal axenic amastigote culture has the potential to expand the experimental freedom of stage-specific studies in T. cruzi.

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Here, we describe a protocol to introduce a gene knockout into the extracellular amastigote of Trypanosoma cruzi, using the CRISPR/Cas9 system. The growth phenotype can be followed up either by cell counting of axenic amastigote culture or by proliferation of intracellular amastigotes after host cell invasion.

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target ...

CRISPR-Cas9-Mediated Genome Editing in the Filamentous Ascomycete Huntiella omanensis

The CRISPR-Cas9 genome editing system is a molecular tool that can be used to introduce precise changes into the genomes of model and non-model species alike. This technology can be used for a variety of genome editing approaches, from gene knockouts and knockins to more specific changes like the introduction of a few nucleotides at a targeted location. Genome editing can be used for a multitude of applications, including the partial functional characterization of genes, the production of transgenic organisms and the development of diagnostic tools. Compared to previously available gene editing strategies, the CRISPR-Cas9 system has been shown to be easy to establish in new species and boasts high efficiency and specificity. The primary reason for this is that the editing tool uses an RNA molecule to target the gene or sequence of interest, making target molecule design straightforward, given that standard base pairing rules can be exploited. Similar to other genome editing systems, CRISPR-Cas9-based methods also require efficient and effective transformation protocols as well as access to good quality sequence data for the design of the targeting RNA and DNA molecules. Since the introduction of this system in 2013, it has been used to genetically engineer a variety of model species, including Saccharomyces cerevisiae, Arabidopsis thaliana, Drosophila melanogaster and Mus musculus. Subsequently, researchers working on non-model species have taken advantage of the system and used it for the study of genes involved in processes as diverse as secondary metabolism in fungi, nematode growth and disease resistance in plants, among many others. This protocol detailed below describes the use of the CRISPR-Cas9 genome editing protocol for the truncation of a gene involved in the sexual cycle of Huntiella omanensis, a filamentous ascomycete fungus belonging to the Ceratocystidaceae family.

CRISPR-Cas9-Mediated Genome Editing in the Filamentous Ascomycete Huntiella omanensis

The CRISPR-Cas9 genome editing system is an easy-to-use genome editor that has been used in model and non-model species. Here we present a protein-based version of this system that was used to introduce a premature stop codon into a mating gene of a non-model filamentous ascomycete fungus.

The CRISPR-Cas9 genome editing system is a molecular tool that can be used to introduce precise changes into the genomes of model and non-model species ...

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to visceral pathology. Deciphering the nature of the vector/parasite interactions is of primary importance for better understanding of Leishmania transmission to their hosts. Among the parameters controlling the sand fly vector competence (i.e. their ability to carry and transmit pathogens), parameters intrinsic to these insects were shown to play a key role. Insect immune response, for example, impacts sand fly vector competence to Leishmania. The study of such parameters has been limited by the lack of methods of gene expression modification adapted for use in these non-model organisms. Gene downregulation by small interfering RNA (siRNA) is possible, but in addition to being technically challenging, the silencing leads to only a partial loss of function, which cannot be transmitted from generation to generation. Targeted mutagenesis by CRISPR/Cas9 technology was recently adapted to the Phlebotomus papatasi sand fly. This technique leads to the generation of transmissible mutations in a specifically chosen locus, allowing to study the genes of interest. The CRISPR/Cas9 system relies on the induction of targeted double-strand DNA breaks, later repaired by either Non-Homologous End Joining (NHEJ) or by Homology Driven Repair (HDR). NHEJ consists of a simple closure of the break and frequently leads to small insertion/deletion events. In contrast, HDR uses the presence of a donor DNA molecule sharing homology with the target DNA as a template for repair. Here, we present a sand fly embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 using NHEJ, which is the only genome modification technique adapted to sand fly vectors to date.

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

This protocol details the steps of CRISPR/Cas9 targeted mutagenesis in sand flies: embryo collection, injection, insect rearing, and identification as well as selection of mutations of interest.

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to ...