We thank Prof. Elly M. Tanaka for her continuous and long-term support. This work was supported by a National Natural Science Foundation of China (NSFC) Grant (317716), Research Starting Grants from South China Normal University (S82111 and 8S0109), and a China Postdoctoral Science Foundation Grant (2018M633067).

All animal experiments must be carried out in accordance with local and national regulations on animal experimentation and with approval of the relevant institutional review board.
1. Preparing the CAS9-gRNA RNP mix
<ol>
	<li>Design and synthesize gRNAs. 
	NOTE: Refer to other publications for designing and synthesizing gRNAs, including one exclusively concerning axolotls15,20,21,22.</li>
	<li>Obtain CAS9-NLS protein by preparing in-house or purchasing it commercially.</li>
	<li>Prepare CAS9-gRNA mix by mixing 5 &#181;g of CAS9-NLS protein, 4 &#181;g of gRNA, 0.9 &#181;L of 10x CAS9 buffer, and fill up the volume to 10 &#181;L with nuclease-free H2O. Aliquot the mixture and store at -80 &#176;C if not used immediately.</li>
</ol>
2. Preparing agarose plates for electroporation
<ol>
	<li>Prepare 200 mL of 2% (wt/vol) agarose solution in 1x DPBS. Dissolve by heating the solution in a microwave and pour it onto 10 cm Petri dishes to a depth of about 10 mm (deep enough to cover the whole animal).</li>
	<li>Allow the plates to solidify at room temperature. Store plates at 4 &#176;C if not used immediately.</li>
	<li>Using surgical scalpels, cut the agarose plate to make a slit for holding the tail straight, a well for fitting in the body of the animal, and two extra wells for placing the electrodes (Figure 1E).</li>
	<li>Place the plate on ice and fill it up to the brim with ice-cold 1x DPBS. Use the same DPBS solution for making the plates to ensure consistent electrical conductibility in the set-up.</li>
</ol>
3. Configuring the electroporator
<ol>
	<li>Configure the electroporator. Set up the poring pulse to consist of one bipolar pulse of 70 V, with a duration of 5 ms, interval of 50 ms, and no voltage decay. Set up the transfer pulse to have four bipolar pulses of 40 V, with a duration of 50 ms, interval of 999 ms, and 10% voltage decay.</li>
	<li>Connect the electrodes to the electroporator and adjust the tweezers to be 7 mm apart. Submerge the electrodes in a beaker of DPBS before and in between electroporation.</li>
</ol>
4. Preparing and loading microinjection glass capillaries
<ol>
	<li>Perform a ramp test on a flaming/brown micropipette puller as per the manufacturer&#8217;s instructions to determine the heat value.</li>
	<li>Pull injection capillaries with tapered ends using the following parameters: heat = ramp test value, pull = 100, velocity = 100, time = 100.</li>
	<li>Observe the needle under a stereomicroscope. Using fine forceps, break the capillary at an angle so that it has a slanted tip. Break at a position where it is thin enough to target the central canal, while being strong enough to pierce the skin. 
	NOTE: The point at which the diameter is at about 20 &#181;m or about 50% of the diameter of the spinal cord is a good starting point. Multiple tries may be needed to obtain optimal breaking.</li>
	<li>Add Fast Green FCF solution to the RNP mix at a ratio of 1:30 and load about 5 &#181;L of injection mix into the capillary with a Microloader pipette tip.</li>
	<li>Mount the capillary on the pneumatic pump, with the slanted tip facing downwards.</li>
</ol>
5. Configuring the pneumatic pump
<ol>
	<li>Configure the pneumatic pump. Set the hold to 0.5 pounds per square inch (psi), eject to 2 psi hold, and duration to gated.</li>
	<li>Test capillary and pneumatic pump settings by dipping the capillary into a drop of water and pressing the foot pedal to inject. Ensure that the injection mix comes out in a slow but steady stream.</li>
	<li>Adjust the hold pressure if the injection mix start leaking out spontaneously or water is sucked up the capillary. Increase the ejection pressure or break more of the capillary if the injection mix is coming out too slow. Decrease the ejection pressure if the injection is coming out too fast.</li>
</ol>
6. Injecting RNP mix into the spinal cord
<ol>
	<li>Anesthetize axolotls in 0.03% benzocaine solution for at least 10 min. Check that the animals are not responsive by flipping them upside down in the water.</li>
	<li>Pick up an animal with ring forceps and lay it on a bed of silicon, with the left side of the animal facing up and the tail pointing towards the right. Position the injection site in the middle of the field of view of the stereomicroscope (Figure 1A).</li>
	<li>Adjust the micromanipulator so that the capillary is coming in at 60&#176; from the right side of the field of view (Figure 1B).</li>
	<li>Identify the spinal cord and central canal (Figure 1C). 
	NOTE: The spinal cord is the tubular structure above the notochord.</li>
	<li>Aiming at the central canal, move the capillary forward to gently pierce the skin and muscle into the central canal of the spinal cord. Limit the movement of the capillary to small steps, as the spinal cord is right beneath the tail muscles.</li>
	<li>Press and hold down the foot pedal to inject the mix into the central canal. Observe whether the RNP mix spreads along the central canal as a blue line in both directions, which shows that the capillary has been correctly positioned (Figure 1D).</li>
	<li>If this does not occur, keep the foot pedal pressed while moving the capillary slightly in and out until the RNP mix starts going in. If the capillary becomes clogged by tissue, clear it by ejecting into water at higher pressures.</li>
	<li>Adjust the ejection pressure so that it is just enough to cause the RNP mix to spread in the central canal, but not so much that it blows up the structure.</li>
	<li>Continue to hold down the foot pedal so the injection mix spreads further along the central canal, until it reaches the terminal vesicle at the end of the spinal cord and ventricles in the brain. 
	NOTE: This may take up to 2 min depending on the size of the animal. It may be necessary to increase the pressure after some time to cause the RNP mix to enter the brain ventricles.</li>
	<li>Proceed as soon as possible to electroporation after successful injection.</li>
</ol>
7. Electroporation
<ol>
	<li>Transfer the animal with ring forceps onto the prepared agarose electroporation plate (Figure 1E) on ice. Place the tail inside the slit so it is sandwiched by agarose (Figure 1F).</li>
	<li>Place the electrodes into the wells on both sides of the tail. Ensure that the entire animal and electrodes are covered by ice-cold PBS.</li>
	<li>Press the foot switch to start the electroporation.</li>
	<li>After electroporation is complete, return the animal to water.</li>
	<li>Proceed to inject more animals if necessary.</li>
	<li>Check that the electroporated animals are healthy after anesthesia wears off and that there is no damage done to the tail.</li>
</ol>
8. Assessing knock-out efficiency
<ol>
	<li>Fix the tail of the electroporated animal. Make cross-sections of the tail followed by immunohistochemical staining to assess the efficiency of the knock-out at the protein level. 
	NOTE: If a tail amputation is to be performed after electroporation, the cut off tail could be used for this purpose. Animals electroporated with gRNAs against Gfp or Tyrosinase, for example, can be used as a negative control4.</li>
	<li>Alternatively, the electroporated spinal cord could be extracted and lysed for DNA. Perform genotyping PCR followed by Sanger sequencing or next-generation sequencing to assess the percentage of edited genetic loci15. 
	NOTE: The resulting editing efficiency will underestimate the actual editing efficiency for NSCs, due to the inclusion of non-electroporated cells and neurons that lack apical contact to the central lumen and thus will not receive the RNP mix.</li>
</ol>

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The authors declare no conflicts of interest.

The described protocol allows time and space restricted gene knock-out in the NSCs in the axolotl spinal cord. The current protocol allows specific targeting of NSCs at a defined time and location with high penetrance. It avoids potential undesired effects originating from gene knock-out in NSCs in other regions such as the brain that occur when using the Cre-LoxP system. It also avoids developmental effects originating from a persistent knock-out, allowing the study of gene function focused on regeneration. In addition,...

The spinal cord of most vertebrates is unable to regenerate following injury, leading to permanent disability. Several salamanders, such as the axolotl, are notable exceptions. The axolotl can fully regenerate a structurally identical spinal cord and completely restore spinal cord function. Much of the regenerative capability of the axolotl spinal cord is due to ependymal cells. These cells line the central canal, and unlike those in mammals, axolotl ependymal cells remain as neural stem cells (NSCs) post-embryonic development. After spinal cord injury (e.g., from a tail amputation), these NSCs proliferate to regrow the ependymal tube and differentiate to replace lost neurons1,2,3. Uncovering how axolotl spinal cord NSCs remain pluripotent and become activated after injury can provide valuable information on developing new therapeutic strategies for human patients.
Due to advances in the CRISPR-Cas9 gene knock-out technique, performing knock-outs to decipher gene function has become easier and has been shown to have broad applicability in various species, including axolotls4,5,6,7,8. The recent release of the full axolotl genome and transcriptome now allows any genomic locus to be targeted and for better assess to off-target effects9,10,11,12,13,14. Optimized protocols have been developed for knock-out and knock-in in axolotls using the CRISPR-Cas9 system15. Delivery of the CRISPR-Cas9 machinery in the form of CAS9 protein-gRNA ribonucleoprotein (RNP) has been shown to be more efficient than using Cas9 and gRNA-encoding plasmids4. This is likely due to the RNP being smaller in size than plasmid vectors, its ability to create DNA breaks immediately, and protecting of the gRNA from RNA degradation. In addition, using RNPs bypasses transcription and translation; thus, it avoids issues such as promoter strength and optimal codon usage when plasmid elements are derived from a different species.
Loss-of-function studies are one of the general approaches to investigating potential functions of genes of interest. In order to study gene function during regeneration, a knock-out should ideally be performed just before an injury to avoid effects on development. In addition, the knock-out should be restricted to both the NSCs and region of regeneration. A knock-out of the target gene in all NSCs (including those in the brain, which is the case in Cre-LoxP systems), may produce effects not related to regeneration that can confound the interpretation of results. Fortunately, the structure of the axolotl spinal cord provides a unique opportunity for time- and space-restricted knock-out in NSCs. Most of the spinal cord NSCs are in contact with the central canal and constitute the vast majority of the cells in contact with the central canal16,17. Therefore, an injection of the CAS9-gRNA complex into the central canal, followed by electroporation, allows delivery to spinal cord NSCs in a desired region at a specific time4,18,19. This protocol demonstrates how this is performed, leading to highly penetrating knock-out in the targeted spinal cord NSCs. Subsequent analysis are then performed to study the effects on regeneration and NSC behavior.

<table><tbody><tr><td>Agarose</td><td>Sigma-Aldrich</td><td>A9539</td><td></td></tr><tr><td>Benzocaine</td><td>Sigma-Aldrich</td><td>E1501-100G</td><td></td></tr><tr><td>Benzocaine 0.03 % (wt/vol)</td><td>Mix 500 ml of 10&amp;times; TBS, 500 ml of 400% (wt/vol) Holtfreter&amp;rsquo;s solution and 30 ml of 10% (wt/vol) benzocaine stock solution. Fill up the volume to 10 L with dH2O. The solution can be stored at room temperature for up to 6 months.</td><td></td><td></td></tr><tr><td>Benzocaine 10 % (wt/vol)</td><td>Mix 50 g of benzocaine in 500 ml of 100% (vol/vol) ethanol. The solution can be stored at room temperature for up to 12 months.</td><td></td><td></td></tr><tr><td>Borosilicate glass capillaries 1.2 mm O.D., 0.94 mm I.D.</td><td>Stutter Instrument&amp;nbsp;</td><td>BF120-94-8</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Merck</td><td>102382</td><td></td></tr><tr><td>CAS9 buffer, 10x</td><td>Mix 200 mM HEPES and 1.5 M KCl in RNase-free water. Adjust pH to 7.5. Filter sterilize, aliquot and store at &amp;minus;20 &amp;deg;C for up to 24 months</td><td></td><td></td></tr><tr><td>CAS9-NLS protein&amp;nbsp;&amp;nbsp;</td><td>PNA Bio</td><td>CP03</td><td></td></tr><tr><td>Cell culture dishes, 10cm</td><td>Falcon</td><td>351029</td><td></td></tr><tr><td>Dumont #5 - Fine Forceps</td><td>Fine Scientific Instruments</td><td>11254-20</td><td></td></tr><tr><td>Electroporator</td><td>Nepa Gene&amp;nbsp;</td><td>NEPA21</td><td></td></tr><tr><td></td><td>BEX</td><td>Pulse Generator CUY21EDIT II</td><td></td></tr><tr><td>Fast Green FCF</td><td>Sigma-Aldrich</td><td>F7252-5G</td><td></td></tr><tr><td>Fast Green FCF Solution, 5x</td><td>Dissolve 12.5 mg of Fast Green FCF powder in 10 mL of 1&amp;times; PBS.</td><td></td><td></td></tr><tr><td>Flaming/Brown Micropipette Puller&amp;nbsp;</td><td>Stutter Instrument&amp;nbsp;</td><td>P-97</td><td></td></tr><tr><td>Holtfreter&amp;rsquo;s solution 400% (wt/vol)&amp;nbsp;</td><td>Dissolve 11.125 g of MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H2O, 5.36 g of CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H2O, 158.4 g of NaCl and 2.875 g of KCl in 10 L of dH2O. The solution can be stored at room temperature for up to 6 months.</td><td></td><td></td></tr><tr><td>KCl</td><td>Merck</td><td>104936</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Merck</td><td>105886</td><td></td></tr><tr><td>Microloader pipette tips</td><td>Eppendorf</td><td>5242956003</td><td></td></tr><tr><td>Micromanipulator&amp;nbsp;</td><td>Narishige</td><td>MN-153&amp;nbsp;</td><td></td></tr><tr><td>NaCl</td><td>Merck</td><td>106404</td><td></td></tr><tr><td>Pneumatic PicoPump</td><td>World Precision Instruments&amp;nbsp;</td><td>SYS-PV830</td><td></td></tr><tr><td>Ring Forceps</td><td>Fine Scientific Instruments</td><td>11103-09</td><td></td></tr><tr><td>Stereomicroscope</td><td>Olympus</td><td>SZX10&amp;nbsp;</td><td></td></tr><tr><td>Tris base</td><td>Sigma-Aldrich</td><td>T6066</td><td></td></tr><tr><td>Tris-buffered saline, 10x</td><td>Dissolve 24.2 g of Tris base and 90 g of NaCl in 990 ml of dH2O. Adjust pH to 8.0 by adding 10 ml of 37% (vol/vol) HCl. The solution can be stored at room temperature for up to 6 months.</td><td></td><td></td></tr><tr><td>Tweezers w/Variable Gap 2 Round Platinum Plate Electrode, 10mm diameter</td><td>Nepa Gene&amp;nbsp;</td><td>CUY650P10</td><td></td></tr></tbody></table>

direct gene knock-out of axolotl spinal cord neural stem cells via electroporation of cas9 protein-grna complexes

The axolotl has the unique ability to fully regenerate its spinal cord. This is largely due to the ependymal cells remaining as neural stem cells (NSCs) throughout life, which proliferate to reform the ependymal tube and differentiate into lost neurons after spinal cord injury. Deciphering how these NSCs retain pluripotency post-development and proliferate upon spinal cord injury to reform the exact pre-injury structure can provide valuable insight into how mammalian spinal cords may regenerate as well as potential treatment options. Performing gene knock-outs in specific subsets of NSCs within a restricted time period will allow study of the molecular mechanisms behind these regenerative processes, without being confounded by development perturbing effects. Described here is a method to perform gene knock-out in axolotl spinal cord NSCs using the CRISPR-Cas9 system. By injecting the CAS9-gRNA complex into the spinal cord central canal followed by electroporation, target genes are knocked out in NSCs within specific regions of the spinal cord at a desired timepoint, allowing for molecular studies of spinal cord NSCs during regeneration.

Injection and electroporation of CAS9-gRNA complex against Sox2 into the axolotl spinal cord central canal led to a massive loss of SOX2 immunoreactivity in a majority of spinal cord NSCs, with gRNA against Tyrosinase (Tyr) as a control (Figure 2A). B3-tubulin (stained with TUJ1)&#160;is a marker for neurons and was not expressed in NSCs, and SOX2- TUJ1- cells surrounding the central canal were considered to be cells harboring Sox2 deletions. Quantification...

Watch this Scientific Journal Video about Direct Gene Knock-out of Axolotl Spinal Cord Neural Stem Cells via Electroporation of CAS9 Protein-gRNA Complexes at JoVE.com

Direct Gene Knock-out of Axolotl Spinal Cord Neural Stem Cells via Electroporation of CAS9 Protein-gRNA Complexes

Presented here is a protocol to perform time- and space-restricted gene knock-out in axolotl spinal cords by injecting CAS9-gRNA complex into the spinal cord central canal followed by electroporation.

The axolotl has the unique ability to fully regenerate its spinal cord. This is largely due to the ependymal cells remaining as neural stem cells (NSCs) ...

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Research

JoVE Journal

Genetics

7.9K Views.  South China Normal University. Presented here is a protocol to perform time- and space-restricted gene knock-out in axolotl spinal cords by injecting CAS9-gRNA complex into the spinal cord central canal followed by electroporation.

Video: Direct Gene Knock-out of Axolotl Spinal Cord Neural Stem Cells via Electroporation of CAS9 Protein-gRNA Complexes

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.

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.

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

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. ...

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 ...

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as axolotls, to regenerate entire limbs as adults. Here, we outline techniques used to generate chimeric axolotls with Cas9-mutagenized haploid forelimbs that can be used for exploring gene function and the fidelity of limb regeneration. We combine several embryological and genetic techniques, including haploid generation via in vitro activation, CRISPR/Cas9 mutagenesis, and tissue grafting into one protocol to produce a unique system for haploid genetic screening in a model organism of regeneration. This strategy reduces the number of animals, space, and time required for the functional analysis of genes in limb regeneration. This also permits the investigation of regeneration-specific functions of genes that may be required for other essential processes, such as organogenesis, tissue morphogenesis, and other essential embryonic processes. The method described here is a unique platform for conducting haploid genetic screening in a vertebrate model system.

This goal of this protocol is to produce chimeric axolotls with haploid forelimbs derived from Cas9-mutagenized donor tissue using embryonic tissue grafting techniques.

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as ...

A Guide to Precise Gene Disruption in Primary Bone Marrow-Derived Macrophages

High-Efficiency Gene Disruption in Primary Bone Marrow-Derived Macrophages Using Electroporated Cas9-sgRNA Complexes

Bone marrow-derived macrophages (BMDMs) from mice are a key tool for studying the complex biology of tissue macrophages. As primary cells, they model the physiology of macrophages in vivo more closely than immortalized macrophage cell lines and can be derived from mice already carrying defined genetic changes. However, disrupting gene function in BMDMs remains technically challenging. Here, we provide a protocol for efficient CRISPR/Cas9 genome editing in BMDMs, which allows for the introduction of small insertions and deletions (indels) that result in frameshift mutations that disrupt gene function. The protocol describes how to synthesize single-guide RNAs (sgRNA-Cas9) and form purified sgRNA-Cas9 ribonucleoprotein complexes (RNPs) that can be delivered by electroporation. It also provides an efficient method for monitoring editing efficiency using routine Sanger sequencing and a freely available online analysis program. The protocol can be performed within 1 week and does not require plasmid construction; it typically results in 85% to 95% editing efficiency.

Author Spotlight: Efficient CRISPR/Cas9 Genome Editing in Bone Marrow-Derived Macrophages for Precise Gene Disruption

This protocol describes the procedure for genome editing in mouse bone marrow-derived macrophages using Cas9-sgRNA ribonucleoprotein complexes assembled in vitro and delivered by electroporation.

Bone marrow-derived macrophages (BMDMs) from mice are a key tool for studying the complex biology of tissue macrophages. As primary cells, they model the ...

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 ...

Genetic Modification of Primate Cerebral Organoids

Targeted Microinjection and Electroporation of Primate Cerebral Organoids for Genetic Modification

The cerebral cortex is the outermost brain structure and is responsible for the processing of sensory input and motor output; it is seen as the seat of higher-order cognitive abilities in mammals, in particular, primates. Studying gene functions in primate brains is challenging due to technical and ethical reasons, but the establishment of the brain organoid technology has enabled the study of brain development in traditional primate models (e.g., rhesus macaque and common marmoset), as well as in previously experimentally inaccessible primate species (e.g., great apes), in an ethically justifiable and less technically demanding system. Moreover, human brain organoids allow the advanced investigation of neurodevelopmental and neurological disorders.
As brain organoids recapitulate many processes of brain development, they also represent a powerful tool to identify differences in, and to functionally compare, the genetic determinants underlying the brain development of various species in an evolutionary context. A great advantage of using organoids is the possibility to introduce genetic modifications, which permits the testing of gene functions. However, the introduction of such modifications is laborious and expensive. This paper describes a fast and cost-efficient approach to genetically modify cell populations within the ventricle-like structures of primate cerebral organoids, a subtype of brain organoids. This method combines a modified protocol for the reliable generation of cerebral organoids from human-, chimpanzee-, rhesus macaque-, and common marmoset-derived induced pluripotent stem cells (iPSCs) with a microinjection and electroporation approach. This provides an effective tool for the study of neurodevelopmental and evolutionary processes that can also be applied for disease modeling.

Author Spotlight: Targeted Microinjection and Electroporation of Primate Cerebral Organoids for Genetic Modification  

The electroporation of primate cerebral organoids provides a precise and efficient approach to introduce transient genetic modification(s) into different progenitor types and neurons in a model system close to primate (patho)physiological neocortex development. This allows the study of neurodevelopmental and evolutionary processes and can also be applied for disease modeling.

The cerebral cortex is the outermost brain structure and is responsible for the processing of sensory input and motor output; it is seen as the seat of ...

Developmental Biology

CRISPR/Cas9 Gene Knockout in Sliced Human Cortical Organoids via Electroporation

Electroporation of Sliced Human Cortical Organoids for Studies of Gene Function

Human cortical organoids have become important tools for studying human brain development, neurodevelopmental disorders, and human brain evolution. Studies analyzing gene function by overexpression or knockout have been instrumental in animal models to provide mechanistic insights into the regulation of neocortex development. Here, we present a detailed protocol for CRISPR/Cas9-mediated acute gene knockout by electroporation of sliced human cortical organoids. The slicing of cortical organoids aids the identification of ventricle-like structures for injection and subsequent electroporation, making this a particularly well-suited model for acute genetic manipulation during human cortical development. We describe the design of guide RNAs and the validation of targeting efficiency in vitro and in cortical organoids. Electroporation of cortical organoids is performed at mid-neurogenic stages, enabling the targeting of most major cell classes in the developing neocortex, including apical radial glia, basal progenitor cells, and neurons. Taken together, the electroporation of sliced human cortical organoids represents a powerful technique to investigate gene function, gene regulation, and cell morphology during cortical development.

Here, we present the acute genetic manipulation of sliced human cortical organoids by electroporation. These cortical organoid models are particularly amenable to injection as ventricle-like structures can be readily identified after slicing, enabling the functional investigation of human cortical development, neurodevelopmental disorders, and cortical evolution.

Human cortical organoids have become important tools for studying human brain development, neurodevelopmental disorders, and human brain evolution. ...

Generation of Knock-out Primary and Expanded Human NK Cells Using Cas9 Ribonucleoproteins

CRISPR/Cas9 technology is accelerating genome engineering in many cell types, but so far, gene delivery and stable gene modification have been challenging in primary NK cells. For example, transgene delivery using lentiviral or retroviral transduction resulted in a limited yield of genetically-engineered NK cells due to substantial procedure-associated NK cell apoptosis. We describe here a DNA-free method for genome editing of human primary and expanded NK cells using Cas9 ribonucleoprotein complexes (Cas9/RNPs). This method allowed efficient knockout of the TGFBR2 and HPRT1 genes in NK cells. RT-PCR data showed a significant decrease in gene expression level, and a cytotoxicity assay of a representative cell product suggested that the RNP-modified NK cells became less sensitive to TGF&#946;. Genetically modified cells could be expanded post-electroporation by stimulation with irradiated mbIL21-expressing feeder cells.

Here, we present a protocol to genetically modify primary or expanded human natural killer (NK) cells using Cas9 Ribonucleoproteins (Cas9/RNPs). By using this protocol, we generated human NK cells deficient for transforming growth factor&#8211;b receptor 2 (TGFBR2) and hypoxanthine phosphoribosyltransferase 1 (HPRT1).

CRISPR/Cas9 technology is accelerating genome engineering in many cell types, but so far, gene delivery and stable gene modification have been challenging ...

Immunology and Infection

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the ...

Direct Gene Knock-out of Axolotl Spinal Cord Neural Stem Cells via Electroporation of CAS9 Protein-gRNA Complexes

Summary

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Chapters in this video

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

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

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

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

High-Efficiency Gene Disruption in Primary Bone Marrow-Derived Macrophages Using Electroporated Cas9-sgRNA Complexes

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

Targeted Microinjection and Electroporation of Primate Cerebral Organoids for Genetic Modification

Electroporation of Sliced Human Cortical Organoids for Studies of Gene Function

Generation of Knock-out Primary and Expanded Human NK Cells Using Cas9 Ribonucleoproteins

Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion