Name: high-efficiency gene disruption in primary bone marrow-derived macrophages using electroporated cas9-sgrna complexes
Uploaded: 2023-08-04T14:50:00.000+00:00
Duration: 4 min 51 s
Description: Watch this Scientific Journal Video about Author Spotlight: Efficient CRISPR/Cas9 Genome Editing in Bone Marrow-Derived Macrophages for Precise Gene Disruption at JoVE.com

This work was funded by the NIH grant 5R01AI144149. The schematic figures were created with BioRender.

1. sgRNA design
NOTE: This step describes selection of the target sequences and design of the sgRNAs. It is helpful to design guides that are in the first large coding exon, so that any translated protein is disrupted early in the open reading frame. It is also helpful to select target sequences that lie within the same exon, as this will streamline the analysis of the editing efficiency (step 6). The examples of genome editing provided with this protocol used sgRNAs targeting the first exon of the Src gene and the Cblb gene, as well as in the non-coding Rosa26 locus of the mouse genome.
<ol>
	<li>Identify 20 nucleotide genomic sequences to target, using one of several free online designing tools (Table 1). Select four to five guides that are non-overlapping within each gene, as Cas9 activity varies by the specific guide chosen, and a priori prediction of a highly active guide is not possible. 
	NOTE: It is critical to ensure the proper orientation of the guide relative to the sequence of the targeted locus. Design tools typically provide the guide sequence in a 5&#39; to 3&#39; orientation, irrespective of which chromosomal strand is targeted. To verify the strand orientation, check that there is a protospacer adjacent motif (PAM) sequence for S. pyogenes Cas9 (nGG) immediately 3&#39; of the targeted genomic sequence. The sgRNA sequence does not include the PAM.</li>
</ol>
2. sgRNA synthesis
NOTE: This step describes how to synthesize sgRNAs using PCR to generate a template for in vitro transcription (IVT), and then purify the sgRNA using spin columns (Figure 1A). Custom synthetic sgRNAs are commercially available through several vendors as an alternative to PCR/IVT.
<ol>
	<li>Perform PCR to generate the IVT template for T7 RNA polymerase. The PCR reaction uses universal primers that contain a T7 RNA polymerase promoter and trans-activating CRISPR RNA (tracrRNA) sequence (Table 2), in addition to short individual primers with the gene-specific guide sequence.
	<ol>
		<li>This is the non-amplifying overlap extension step. Dilute the PCR buffer to 1x and add an additional 1 mM MgCl2. Then, add 0.25 &#181;L of high-fidelity DNA polymerase, 0.5 mM deoxynucleotide triphosphate (dNTP), 0.02 mM guide-specific primer, and 0.02 mM T7 reverse long universal primer (Table 2) to a total volume of 46 &#181;L. Then, place the tube in the thermocycler and run two cycles of: 95 &#176;C for 15 s, 37 &#176;C for 15 s, and 72 &#176;C for 15 s. Cool the reactions on ice.</li>
		<li>This is the PCR amplification step. To each reaction, add 2 &#181;L of 25 mM forward amplification primer and 2 &#181;L of 25 mM reverse amplification primer. The final reaction volume is 50 &#181;L. Denature for 30 s at 95 &#176;C. Then, run 35 cycles of: 95 &#176;C for 15 s, 65 &#176;C for 20 s, and 72 &#176;C for 15 s. Perform an additional extension at 72 &#176;C for 2 min.</li>
		<li>Check the PCR reaction product on a 2% agarose gel. Overlap-extension PCR results in a 127 bp dsDNA molecule with the T7 promoter upstream of the gene-specific 20 nucleotide guide and the tracrRNA immediately downstream (Figure 1B).</li>
	</ol>
	</li>
	<li>IVT template purification: There are numerous protocols and kits that work well for this. This protocol uses solid-phase reversible immobilization (SPRI) beads. Mix 50 &#181;L of the PCR product with 90 &#181;L of beads and follow the manufacturer&#39;s instructions. Elute the DNA by adding 40 &#181;L of buffer (5 mM Tris, 0.1 mM ethylenediaminetetraacetic acid [EDTA]) and heat at 65 &#176;C for 5 min.</li>
	<li>Measure the DNA concentration of the IVT template using absorption at a wavelength of 260 nm. A DNA concentration of at least 50 ng/&#181;L is needed for IVT. The IVT template can be stored at -20 &#176;C.</li>
	<li>IVT reaction
	<ol>
		<li>Use a commercial kit designed to produce high yields of IVT (see Table of Materials). Follow the manufacturer&#39;s recommendations to perform the IVT reaction. Using 250 ng of IVT template in a 20 &#181;L reaction, incubate at 37 &#176;C for 4-16 h.</li>
		<li>Check the IVT sgRNA product by running 1 &#181;L of the reaction on a 2% agarose gel. A bright RNA band should be observed, running similarly to 70 bp dsDNA (Figure 1C). Though not mandatory, to obtain sharp and more resolved bands, use an RNA sample buffer with urea, and denature the sample at 65 &#176;C for 5 min prior to loading. 
		NOTE: Faint higher-molecular weight bands and slight migration differences may be seen with some guide RNAs due to differences in the RNA secondary structure.</li>
	</ol>
	</li>
	<li>Dephosphorylate the sgRNA IVT product to minimize the activation of RIG-I (a foreign RNA sensor) and subsequent cell death following electroporation. For each 20 &#181;L IVT reaction, add 69 &#181;L of molecular-grade water and 15 U of calf intestinal phosphatase (CIP) in 1x CIP buffer. Incubate for 2 h at 37 &#176;C.</li>
	<li>Degrade the IVT template to minimize the activation of cellular DNA sensors, which trigger cell death following electroporation. To each IVT reaction, add 2 U of RNase-free DNase. Incubate for 15 min at 37 &#176;C. The IVT product can be stored at -20 &#176;C for 1 day or -80 &#176;C for several months.</li>
	<li>Purify the IVT product using spin columns. For this step, SPRI bead purification kits are inferior.
	<ol>
		<li>Bind the RNA to the column by adding 220 &#181;L of binding buffer to the IVT product, followed by 1 mL of 100% molecular biology-grade ethanol. Mix well and load the column. Thereafter, follow the manufacturer&#39;s instructions. Perform the final wash in a laminar flow biosafety cabinet to avoid contamination of the sgRNA.</li>
		<li>Perform the elution in the laminar flow hood to avoid contamination. Elute the RNA using 30 &#181;L of sterile 10 mM Tris buffer (pH 8.0).</li>
	</ol>
	</li>
	<li>Measure the concentration of sgRNA by measuring the absorbance at a wavelength of 260 nm. 
	NOTE: Typical sgRNA yields are at least 100 &#181;g.</li>
	<li>Store the sgRNA at -80 &#176;C. Make aliquots to avoid more than three freeze-thaw cycles.</li>
</ol>
3. Preparation for electroporation
NOTE: All steps should be performed in a laminar flow hood to avoid contamination. This protocol uses a commercially available electroporation system (see Table of Materials) with 10 &#181;L tips.
<ol>
	<li>Thaw the BMDMs 48-72 h before use and expand them on non- tissue culture (TC) treated plates. 
	NOTE: Per reaction, 4 x 105 cells are needed.</li>
	<li>Sterilize PCR tubes for reaction assembly. Prepare one PCR tube per reaction, and heat them in a thermocycler for 10 min at 98 &#176;C. Prepare the tubes in batches and store them closed. Place the sterilized PCR tubes on ice when ready to use.</li>
	<li>Clean the pipette station with 70% ethanol and place it in the laminar flow hood. Set the parameters to 1,900 V, 20 ms, and one pulse.</li>
	<li>Remove a transfection tube from the package with care to keep it sterile and fill it with 3 mL of &#34;E&#34; buffer. Ensure that the E buffer is at room temperature prior to electroporation. Insert the tube in the station and push it down until it clicks.</li>
	<li>For each reaction, aliquot 2.5 &#181;L of sgRNA at a concentration of 1,100 ng/&#181;L. Dilute the sgRNA in cold phosphate-buffered saline (PBS) with 0.9 mM CaCl2 and 0.5 mM MgCl2 (PBS + Ca/Mg). Leave it on ice.</li>
	<li>Prepare two plates: a 12-well uncoated plate for the cells and an additional 24-well plate to hold the culture media. Aliquot 1.5 mL of antibiotic-free medium per reaction into the wells of the 24-well plate.</li>
	<li>Prepare Cas9 protein. There are several commercial and academic suppliers of sterile-purified Cas9 protein. Dilute Cas9 to a final concentration of 20 &#181;M with cold PBS + Ca/Mg. For each electroporation reaction, aliquot 1 &#181;L of 20 &#181;M Cas9 into sterile PCR tubes. Leave on ice.</li>
	<li>Prepare cells for electroporation.
	<ol>
		<li>Remove the growth medium. Wash the cells using PBS without CaCl2 and MgCl2 (PBS-Ca/Mg) to remove the serum and divalent cations that promote adhesion. Then, add PBS + 1 mM EDTA and incubate at room temperature for about 5 min until the cells begin to detach.</li>
		<li>Pipette gently up and down to detach the cells. Transfer the cells to a low protein-binding 15 mL tube. Pellet the cells at 500 x g for 5 min. 
		NOTE: Macrophages are adherent to plasticware. The use of low protein-binding centrifuge tubes significantly improves the cell yield.</li>
		<li>Work quickly to avoid prolonged incubation of cells in PBS + EDTA. Remove most of the supernatant, leaving about 1 mL of supernatant in the tube. Resuspend the cells in the remaining supernatant, and then transfer to a low protein-binding 1.5 ml microfuge tube.</li>
		<li>Remove a small aliquot of cells to count. Pellet the remaining cells by centrifuging at 500 x g at room temperature for 5 min.</li>
		<li>During centrifugation, count the cells and warm the Cas9 and sgRNA tubes to room temperature.</li>
		<li>Remove all the supernatant from the cell pellet. Resuspend the cells in PBS + Ca/Mg at a concentration of 4 x 107 cells/mL (4 x 105 cells per 10 &#181;L reaction). 
		NOTE: Each kit is supplied with a limited amount of electroporation buffer (Buffer R, Buffer T). However, we find that the electroporation efficiency using PBS + Ca/Mg is equivalent to the proprietary buffers.</li>
		<li>When electroporating sgRNAs for different genes, aliquot the cells into different low protein-binding tubes for each gene to avoid cross-contamination between the sgRNAs. Keep the cells at room temperature until ready for electroporation.</li>
	</ol>
	</li>
</ol>
4. RNP assembly
<ol>
	<li>Keep the sgRNA and Cas9 at room temperature in order to avoid precipitation. Add 2.5 &#181;L of sgRNA to 1 &#181;L of the diluted Cas9 in sterilized PCR tubes. Dispense the sgRNA slowly over 15 s with mixing to prevent precipitation.</li>
	<li>Incubate for 5 min at room temperature to allow RNP complex formation.</li>
</ol>
5. RNP delivery by electroporation
<ol>
	<li>Prepare the electroporation tip (cuvette). Depress the plunger to extend the stem. Insert the stem into the tip. 
	NOTE: Ensure that the tip is secure and that the plunger slides smoothly and extends beyond the tip sheath. If the tip is not properly positioned, air bubbles may be introduced into the tip, causing tip failure.</li>
	<li>Resuspend the cells by pipetting up and down, and then load the electroporation tip with the cells. Withdraw the full 10 &#181;L volume capacity of the tip, avoiding bubbles.</li>
	<li>Starting with the negative control sgRNA, transfer 10 &#181;L of cells in the electroporation tip to the sample tube containing 3.5 &#181;L of RNP from step 4. Pipette up and down three times to mix well. Draw 10 &#181;L of the mixture back into the tip, ensuring no bubbles are present.</li>
	<li>Place the tip into the electroporation station, lowering it into the buffer. Avoid contacting plastic surfaces with tip to maintain sterility. Push Start on the touchscreen display.</li>
	<li>After completion, the display will indicate a successful pulse. Remove the pipette from the device and place the cells in a dry well of the labeled, uncoated 12-well plate.</li>
	<li>Rinse the tip by pipetting up and down twice in 15 mL of PBS + Ca/Mg. Set aside the pipette with the tip still attached. Ensure that the tip does not touch anything and remains sterile. 
	NOTE: In many experiments, it is possible to reuse the tip for multiple samples, such as after the inactive negative control sgRNA sample, or during the initial evaluations of the guide activity when four to five sgRNAs are tested per gene and the sole read-out is the editing efficiency. However, during functional analysis on edited cells, a new tip is recommended for each gene, as small amounts of sgRNA or cell carryover could impact the experimental results.</li>
	<li>Immediately add 1 mL of antibiotic-free medium (aliquoted in step 3.6) to the cells and gently shake the plate to mix.</li>
	<li>Repeat steps 5.2-5.6 for the remaining reactions.</li>
	<li>Return the cells to the incubator.</li>
	<li>Check the viability of the cells 1-2 h post-electroporation. The cells should be &#62;90% adherent. Optional: Add gentamicin (5 &#181;g/mL) to the cells 1-2 h post-electroporation to help prevent contamination if this will not interfere with the downstream experiments.</li>
</ol>
6. Assessing the editing efficiency
NOTE: Most editing is complete after 48 h.
<ol>
	<li>Check the cell monolayer 48 h after electroporation. If the cells are over 50% confluent, then proceed further. If the cells are &#60;50% confluent, wait an additional 1-2 days. The genomic DNA analysis to determine the editing efficiency requires 1 x 105 cells, in addition to the cells needed for the downstream experiment.</li>
	<li>Prepare genomic DNA (gDNA).
	<ol>
		<li>Wash the cells with PBS(-Ca/Mg). Add 1 mL of PBS + 1 mM EDTA. Incubate at room temperature for about 5 min until the cells begin to detach from the plate.</li>
		<li>Detach the cells by pipetting. Transfer to a low protein-binding microfuge tube.</li>
		<li>Count the cells and remove an aliquot of 1 x 105 cells. The remaining cells can be re-plated for use in further experiments. Generally, the experiments are conducted 5 days after electroporation in order to allow decay of the protein.</li>
		<li>Pellet the cells for gDNA analysis for 15 s at maximum speed in a microfuge tube.</li>
		<li>Resuspend the pellet in 50 &#181;L of lysis buffer. Vortex to detach any cells stuck to the walls of the tube, and heat at 98 &#176;C for 10 min to lyse the cells and inactivate endogenous DNase.</li>
		<li>Place the tubes on ice.</li>
		<li>Add 1 &#181;L&#160;proteinase K (20 mg/mL). Incubate for at least 1 h at 37 &#176;C; it can be left overnight for convenience.</li>
		<li>Heat to 98 &#176;C for 10 min to inactivate the proteinase K.</li>
		<li>Centrifuge at room temperature for 5 min at maximum speed. Remove the supernatant, which contains the DNA. This crude preparation without further purification works well for most PCR reactions.</li>
	</ol>
	</li>
	<li>Evaluate the editing efficiency with Sanger sequencing.
	<ol>
		<li>Design PCR primers 200-300 bp upstream of the most 5&#39; guide sites and 200-300 bp downstream of the most 3&#39; guide sites. The typical amplicon size is ~500 bp. Design a nested sequencing primer at least 125 bp away from the nearest guide site (Figure 2A).</li>
		<li>Perform PCR using 2 &#181;L of gDNA.</li>
	</ol>
	</li>
	<li>Prepare the PCR product for sequencing. This protocol uses an exonuclease I/shrimp alkaline phosphatase (ExoI/SAP) treatment. Commercial DNA purification kits also work but require more hands-on time.
	<ol>
		<li>Prepare 15 &#181;L of the PCR product. Add 7.5 &#181;L of water, 1 &#181;L of 10x ExoI buffer, 10 U of ExoI, and 1 U of SAP to degrade the dNTPs and primers that interfere with Sanger sequencing.</li>
		<li>Incubate at 37 &#176;C for 30 min, followed by 85 &#176;C for 20 min to deactivate the enzymes.</li>
	</ol>
	</li>
	<li>Carry out Sanger sequencing on the Exo/SAP-treated PCR product; use a known amount of DNA size marker to estimate the size of the PCR product present in the sample. The sequencing reactions may require optimization, as the TIDE software needs high-quality sequence chromatograms to accurately estimate the editing efficiency. The sequence chromatogram for the un-edited locus should provide clear peaks with minimal background (Figure 2B, C).</li>
	<li>To estimate the editing efficiency, use TIDE to analyze the Sanger sequencing chromatogram files using the edited gDNA and the unedited control gDNA. (Table 1, Figure 2). Upload the Sanger sequencing files and run the software with the default settings.</li>
</ol>

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

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The authors have nothing to disclose.

Genome editing using electroporated Cas9-sgRNA complexes allows effective disruption of gene function in BMDMs. The editing efficiency varies by the target sequence and gene. Typically, four to five sgRNAs are generally screened to identify one that is highly active. Some loci have lower editing efficiencies, most likely due the chromatin structure. In these cases, several modifications can be made to increase the editing efficiency. Co-delivery of two active sgRNAs to the same exon results in improved editing for some g...

Macrophages are innate immune cells that play critical roles in tissue repair and immunity1,2. Immortalized macrophage cell lines, such as mouse RAW 264.7 cells or human THP-1 cells, have several beneficial characteristics, including robust growth and ease of gene disruption by delivering vectors for RNA interference or CRISPR/Cas93,4. However, oncogenic transformation dramatically alters their physiology, which results in the aberrant activation of some pathways and muted responses of others5,6. Primary bone marrow-derived macrophages (BMDMs) more closely recapitulate in vivo macrophage physiology, but remain challenging to genetically manipulate due to the low efficiency of both plasmid transfection and viral transduction in these primary immune cells7,8. Thus, more efficient methods for disrupting gene function are needed.
CRISPR/Cas9 genome editing is a powerful tool for genetic manipulation across a range of biological systems, including mammalian cells9,10,11,12. The Streptococcus pyogenes Cas9 protein efficiently and specifically cleaves double-stranded DNA when complexed with a sequence-specific guide RNA. DNA repair through the non-homologous end joining (NHEJ) of the cleaved DNA results in small insertions or deletions (indels) that create frameshift mutations. In early studies, Cas9 and sgRNAs were delivered through plasmid or lentiviral vectors, which are effective delivery methods for many cell lines9,10. However, primary cells and, in particular, primary immune cells are often refractory to these methods due to the low efficiency of vector delivery by transfection or transduction. Subsequently, methods have been developed to generate sgRNA-Cas9 complexes in vitro and to deliver them via electroporation, and these methods have achieved high efficiency in a variety of cell types13,14. The results have suggested the possibility of using this approach to carry out genome editing in primary macrophages.
Here, we provide a protocol for using sgRNA-Cas9 ribonucleoprotein complexes (RNPs) to carry out genome editing in primary BMDMs. It contains steps to mitigate the activation of the immune sensors present in primary immune cells and results in up to 95% editing at targeted loci with minimal toxicity. This protocol also includes workflows to evaluate editing efficiency using routine polymerase chain reaction (PCR) and Sanger sequencing, followed by in silico analysis by Tracking of Indels by Decomposition (TIDE)15, a well-validated online software tool.

<table><tbody><tr><td>3T3-MCSF Cell Line</td><td>Gift from Russell Vance</td><td>not applicable</td><td></td></tr><tr><td>Alt-R Cas9 Electroporation Enhancer</td><td>IDT</td><td>1075915</td><td></td></tr><tr><td>Ampure XP Reagent Beads</td><td>Beckman Coulter</td><td>A63880</td><td></td></tr><tr><td>Calf intestinal alkaline phosphatase</td><td>NEB</td><td>M0525S</td><td></td></tr><tr><td>DNase</td><td>NEB</td><td>M0303S</td><td></td></tr><tr><td>DPBS +Ca/Mg (0.9mM CaCl2 and 0.5mM MgCl2)</td><td>Thermo Fisher</td><td>14040-133</td><td></td></tr><tr><td>DPBS -Ca/Mg</td><td>Thermo Fisher</td><td>14190-144</td><td></td></tr><tr><td>ExoI</td><td>NEB</td><td>M0293S</td><td></td></tr><tr><td>Fetal Calf Serum (FCS)</td><td>Corning</td><td>35-015-CV</td><td></td></tr><tr><td>Herculase DNA polymerase &amp;amp; buffer</td><td>Agilent</td><td>600677</td><td></td></tr><tr><td>HiScribe T7 High Yield RNA Synthesis Kit</td><td>NEB</td><td>E2040S</td><td></td></tr><tr><td>LoBind conical tubes 15 mL</td><td>Eppendorf</td><td>30122216</td><td></td></tr><tr><td>LoBind Eppendorf tubes 2 mL</td><td>Eppendorf</td><td>22431102</td><td></td></tr><tr><td>NEBuffer r2.1</td><td>NEB</td><td>B6002S</td><td></td></tr><tr><td>Neon Transfection System</td><td>Thermo Fisher</td><td>MPK5000, MPP100, MPS100</td><td></td></tr><tr><td>Neon Transfection System 10 uL Tips</td><td>Thermo Fisher</td><td>MPK1025 or MPK1096</td><td></td></tr><tr><td>PBS + 1mM EDTA</td><td>Lonza</td><td>BE02017F</td><td></td></tr><tr><td>Proteinase K</td><td>Thermo Fisher</td><td>EO0491</td><td></td></tr><tr><td>rCutSmart Buffer for ExoI</td><td>NEB</td><td>B6004S</td><td></td></tr><tr><td>Ribolock</td><td>Thermo Fisher</td><td>EO0384</td><td></td></tr><tr><td>RNA loading dye</td><td>NEB</td><td>B0363S</td><td></td></tr><tr><td>RNeasy Mini Kit</td><td>Qiagen</td><td>74104</td><td></td></tr><tr><td>S. pyogenes Cas9-NLS</td><td>University of California Macro Lab</td><td>not applicable</td><td>Available to non-UC investigators through&amp;nbsp; https://qb3.berkeley.edu</td></tr><tr><td>S. pyogenes Cas9-NLS, modified 3rd Generation</td><td>IDT</td><td>1081059</td><td></td></tr><tr><td>SAP</td><td>NEB</td><td>M0371S</td><td></td></tr></tbody></table>

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.

The IVT template is a 127 bp PCR product (Figure 1B). The full-length IVT product is a 98 nt RNA, which migrates similarly to a 70 bp double-stranded DNA fragment (Figure 1C).
After electroporation, the cells should be &#62;90% viable, with a total cell count of &#62;70% of the starting cell number. The resulting pool of mutant cells should have a diverse set of indels, starting near the Cas9 cleavage site. The analysis of the targete...

Watch this Scientific Journal Video about Author Spotlight: Efficient CRISPR/Cas9 Genome Editing in Bone Marrow-Derived Macrophages for Precise Gene Disruption at JoVE.com

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.

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 ...

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JoVE Journal

Genetics

1.4K Views.  University of California, Davis. 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.

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

Parallel Measurement of Circadian Clock Gene Expression and Hormone Secretion in Human Primary Cell Cultures

Circadian clocks are functional in all light-sensitive organisms, allowing for an adaptation to the external world by anticipating daily environmental changes. Considerable progress in our understanding of the tight connection between the circadian clock and most aspects of physiology has been made in the field over the last decade. However, unraveling the molecular basis that underlies the function of the circadian oscillator in humans stays of highest technical challenge. Here, we provide a detailed description of an experimental approach for long-term (2-5 days) bioluminescence recording and outflow medium collection in cultured human primary cells. For this purpose, we have transduced primary cells with a lentiviral luciferase reporter that is under control of a core clock gene promoter, which allows for the parallel assessment of hormone secretion and circadian bioluminescence. Furthermore, we describe the conditions for disrupting the circadian clock in primary human cells by transfecting siRNA targeting CLOCK. Our results on the circadian regulation of insulin secretion by human pancreatic islets, and myokine secretion by human skeletal muscle cells, are presented here to illustrate the application of this methodology. These settings can be used to study the molecular makeup of human peripheral clocks and to analyze their functional impact on primary cells under physiological or pathophysiological conditions.

Here, we describe settings to monitor in parallel circadian bioluminescence and the secretory activity of human islet cells and primary myotubes. For this, we employed lentiviral gene delivery of a luciferase core clock reporter, followed by in vitro synchronization and collection of outflow medium by continuous cell perifusion.

Circadian clocks are functional in all light-sensitive organisms, allowing for an adaptation to the external world by anticipating daily environmental ...

Perturbations of Circulating miRNAs in Irritable Bowel Syndrome Detected Using a Multiplexed High-throughput Gene Expression Platform

The gene expression platform assay allows for robust and highly reproducible quantification of the expression of up to 800 transcripts (mRNA or miRNAs) in a single reaction. The miRNA assay counts transcripts by directly imaging and digitally counting miRNA molecules that are labeled with color-coded fluorescent barcoded probe sets (a reporter probe and a capture probe). Barcodes are hybridized directly to mature miRNAs that have been elongated by ligating a unique oligonucleotide tag (miRtag) to the 3&#39; end. Reverse transcription and amplification of the transcripts are not required. Reporter probes contain a sequence of six color positions populated using a combination of four fluorescent colors. The four colors over six positions are used to construct a gene-specific color barcode sequence. Post-hybridization processing is automated on a robotic prep station. After hybridization, the excess probes are washed away, and the tripartite structures (capture probe-miRNA-reporter probe) are fixed to a streptavidin-coated slide via the biotin-labeled capture probe. Imaging and barcode counting is done using a digital analyzer. The immobilized barcoded miRNAs are visualized and imaged using a microscope and camera, and the unique barcodes are decoded and counted. Data quality control (QC), normalization, and analysis are facilitated by a custom-designed data processing and analysis software that accompanies the assay software. The assay demonstrates high linearity over a broad range of expression, as well as high sensitivity. Sample and assay preparation does not involve enzymatic reactions, reverse transcription, or amplification; has few steps; and is largely automated, reducing investigator effects and resulting in high consistency and technical reproducibility. Here, we describe the application of this technology to identifying circulating miRNA perturbations in irritable bowel syndrome.

We describe the use of a multiplexed high-throughput gene expression platform that quantitates gene expression by barcoding and counting molecules in biological substrates without the need for amplification. We used the platform to quantitate microRNA (miRNA) expression in whole blood in subjects with and without irritable bowel syndrome.

The gene expression platform assay allows for robust and highly reproducible quantification of the expression of up to 800 transcripts (mRNA or miRNAs) in ...

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most mutagenesis-based screens, researchers have relied on the use of toxic chemicals, carcinogens, or irradiation, which requires designated equipment, safety setup, and/or disposal of hazardous materials. We have developed a simple approach to induce heritable mutations in C. elegans using germline-expressed histone-miniSOG, a light-inducible potent generator of reactive oxygen species. This mutagenesis method is free of toxic chemicals and requires minimal laboratory safety and waste management. The induced DNA modifications include single-nucleotide changes and small deletions, and complement those caused by classical chemical mutagenesis. This methodology can also be used to induce integration of extrachromosomal transgenes. Here, we provide the details of the LED setup and protocols for standard mutagenesis and transgene integration.

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Genetically-encoded histone-miniSOG induces genome-wide heritable mutations in a blue&#160;light-dependent manner. This mutagenesis method is simple, fast, free of toxic chemicals, and well-suited for forward genetic screening and transgene integration.

Forward genetic screening in model organisms is the workhorse to discover functionally important genes and pathways in many biological processes. In most ...

Using CRISPR/Cas9 Gene Editing to Investigate the Oncogenic Activity of Mutant Calreticulin in Cytokine Dependent Hematopoietic Cells

Clustered regularly interspaced short palindromic repeats (CRISPR) is an adaptive immunity system in prokaryotes that has been repurposed by scientists to generate RNA-guided nucleases, such as CRISPR-associated (Cas) 9 for site-specific eukaryotic genome editing. Genome engineering by Cas9 is used to efficiently, easily and robustly modify endogenous genes in many biomedically-relevant mammalian cell lines and organisms. Here we show an example of how to utilize the CRISPR/Cas9 methodology to understand the biological function of specific genetic mutations. We model calreticulin (CALR) mutations in murine interleukin-3 (mIL-3) dependent pro-B (Ba/F3) cells by delivery of single guide RNAs (sgRNAs) targeting the endogenous Calr locus in the specific region where insertion and/or deletion (indel) CALR mutations occur in patients with myeloproliferative neoplasms (MPN), a type of blood cancer. The sgRNAs create double strand breaks (DSBs) in the targeted region that are repaired by non-homologous end joining (NHEJ) to give indels of various sizes. We then employ the standard Ba/F3 cellular transformation assay to understand the effect of physiological level expression of Calr mutations on hematopoietic cellular transformation. This approach can be applied to other genes to study their biological function in various mammalian cell lines.

Targeted gene editing using CRISPR/Cas9 has greatly facilitated the understanding of the biological functions of genes. Here, we utilize the CRISPR/Cas9 methodology to model calreticulin mutations in cytokine-dependent hematopoietic cells in order to study their oncogenic activity.

Clustered regularly interspaced short palindromic repeats (CRISPR) is an adaptive immunity system in prokaryotes that has been repurposed by scientists to ...

Cancer Research

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. ...

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

Lentiviral CRISPR/Cas9-Mediated Genome Editing for the Study of Hematopoietic Cells in Disease Models

Manipulating genes in hematopoietic stem cells using conventional transgenesis approaches can be time-consuming, expensive, and challenging. Benefiting from advances in genome editing technology and lentivirus-mediated transgene delivery systems, an efficient and economical method is described here that establishes mice in which genes are manipulated specifically in hematopoietic stem cells. Lentiviruses are used to transduce Cas9-expressing lineage-negative bone marrow cells with a guide RNA (gRNA) targeting specific genes and a red fluorescence reporter gene (RFP), then these cells are transplanted into lethally-irradiated C57BL/6 mice. Mice transplanted with lentivirus expressing non-targeting gRNA are used as controls. Engraftment of transduced hematopoietic stem cells are evaluated by flow cytometric analysis of RFP-positive leukocytes of peripheral blood. Using this method, ~90% transduction of myeloid cells and ~70% of lymphoid cells at 4 weeks after transplantation can be achieved. Genomic DNA is isolated from RFP-positive blood cells, and portions of the targeted site DNA are amplified by PCR to validate the genome editing. This protocol provides a high-throughput evaluation of hematopoiesis-regulatory genes and can be extended to a variety of mouse disease models with hematopoietic cell involvement.

Described are protocols for the highly efficient genome editing of murine hematopoietic stem and progenitor cells (HSPC) by the CRISPR/Cas9 system to rapidly develop mouse model systems with hematopoietic system-specific gene modifications.

Manipulating genes in hematopoietic stem cells using conventional transgenesis approaches can be time-consuming, expensive, and challenging. Benefiting ...

Gene Knock-in by CRISPR/Cas9 and Cell Sorting in Macrophage and T Cell Lines

Functional genomics studies of the immune system require genetic manipulations that involve both deletion of target genes and addition of elements to proteins of interest. Identification of gene functions in cell line models is important for gene discovery and exploration of cell-intrinsic mechanisms. However, genetic manipulations of immune cells such as T cells and macrophage cell lines using CRISPR/Cas9-mediated knock-in are difficult because of the low transfection efficiency of these cells, especially in a quiescent state. To modify genes in immune cells, drug-resistance selection and viral vectors are typically used to enrich for cells expressing the CRIPSR/Cas9 system, which inevitably results in undesirable intervention of the cells. In a previous study, we designed dual fluorescent reporters coupled to CRISPR/Cas9 that were transiently expressed after electroporation. This technical solution leads to rapid gene deletion in immune cells; however, gene knock-in in immune cells such as T cells and macrophages without the use of drug-resistance selection or viral vectors is even more challenging. In this article, we show that by using cell sorting to aid selection of cells transiently expressing CRISPR/Cas9 constructs targeting the Rosa26 locus in combination with a donor plasmid, gene knock-in can be achieved in both T cells and macrophages without drug-resistance enrichment. As an example, we show how to express human ACE2, a receptor of SARS-Cov-2, which is responsible for the current Covid-19 pandemic, in RAW264.7 macrophages by performing knock-in experiments. Such gene knock-in cells can be widely used for mechanistic studies.

This protocol uses fluorescent reporters and cell sorting to simplify knock-in experiments in macrophage and T cell lines. Two plasmids are used for these simplified knock-in experiments, namely a CRISPR/Cas9- and DsRed2-expressing plasmid and a homologous recombination donor plasmid expressing EBFP2, which is permanently integrated at the Rosa26 locus in immune cells.

Functional genomics studies of the immune system require genetic manipulations that involve both deletion of target genes and addition of elements to ...

Electroporation-Based CRISPR-Cas9-Mediated Gene Knockout in THP-1 Cells and Single-Cell Clone Isolation

The human acute monocytic leukemia (AML) THP-1 cell line is widely used as a model to study the functions of human monocyte-derived macrophages, including their interplay with significant human pathogens such as the human immunodeficiency virus (HIV). Compared to other immortalized cell lines of myeloid origin, THP-1 cells retain many intact inflammatory signaling pathways and display phenotypic characteristics that more closely resemble those of primary monocytes, including the ability to differentiate into macrophages when treated with phorbol-12-myristate 13-acetate (PMA). The use of CRISPR-Cas9 technology to engineer THP-1 cells through targeted gene knockout (KO) provides a powerful approach to better characterize immune-related mechanisms, including virus-host interactions. This article describes a protocol for efficient CRISPR-Cas9-based engineering using electroporation to deliver pre-assembled Cas9:sgRNA ribonucleoproteins into the cell nucleus. Using multiple sgRNAs targeting the same locus at slightly different positions results in the deletion of large DNA fragments, thereby increasing editing efficiency, as assessed by the T7 endonuclease I assay. CRISPR-Cas9-mediated editing at the genetic level was validated by Sanger sequencing followed by Inference of CRISPR Edits (ICE) analysis. Protein depletion was confirmed by immunoblotting coupled with a functional assay. Using this protocol, up to 100% indels in the targeted locus and a decrease of over 95% in protein expression were achieved. The high editing efficiency makes it convenient to isolate single-cell clones by limiting dilution.

The THP-1 cell line is widely used as a model to investigate the functions of human monocytes/macrophages across various biology-related research areas. This article describes a protocol for efficient CRISPR-Cas9-based engineering and single-cell clone isolation, enabling the production of robust and reproducible phenotypic data.

The human acute monocytic leukemia (AML) THP-1 cell line is widely used as a model to study the functions of human monocyte-derived macrophages, including ...

CRISPR-Cas9 Mediated Gene Deletion in Human Pluripotent Stem Cells Cultured Under Feeder-Free Conditions

The CRISPR-Cas9 system for genome editing has revolutionized gene function studies in mammalian cells, including stem cells. However, the practical application of this technique, particularly in pluripotent stem cells, presents certain challenges, such as being time- and labor-intensive and having low editing efficiency. Here, we describe the generation of a CRISPR-mediated gene knockout in a human embryonic stem cell (hESC) line stably expressing sgRNAs for the L2HGDH gene, using a highly efficient and stable lentiviral-mediated gene delivery system. The sgRNAs targeting exon 1 of the L2HGDH gene were chemically synthesized and cloned into the lentiCRISPR v2-puro vector, which combines the constitutive expression of sgRNAs with Cas9 in a highly efficient single-vector system to achieve higher lentiviral titers for hESC infection and stable selection using puromycin. Puromycin-selected cells were further expanded, and single-cell clones were obtained using the limited dilution method. The single clones were expanded, and several homozygous knockout clones for the L2HGDH gene were obtained, as confirmed by a 100% reduction in L2HGDH expression using Western blot analysis. Furthermore, using MSBSP-PCR, the CRISPR mutation site was mapped upstream of the PAM recognition sequence of Cas9 in the selected homozygous clones. Sanger sequencing was performed to analyze the exact insertions/deletions, and functional characterization of the clones was conducted. This method produced a significantly higher percentage of homozygous deletions compared to previously reported non-viral gene delivery methods. Although this report focuses on the L2HGDH gene, this robust and cost-effective approach can be used to create homozygous knockouts for other genes in pluripotent stem cells for gene function studies.

The presented method describes the generation of a CRISPR-mediated gene knockout in the human embryonic stem cell (hESC) line H9, which stably expresses sgRNAs targeting the L2HGDH gene using a highly efficient lentiviral-mediated gene delivery system.

The CRISPR-Cas9 system for genome editing has revolutionized gene function studies in mammalian cells, including stem cells. However, the practical ...

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

Summary

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Chapters in this video

Parallel Measurement of Circadian Clock Gene Expression and Hormone Secretion in Human Primary Cell Cultures

Perturbations of Circulating miRNAs in Irritable Bowel Syndrome Detected Using a Multiplexed High-throughput Gene Expression Platform

Optogenetic Random Mutagenesis Using Histone-miniSOG in C. elegans

Using CRISPR/Cas9 Gene Editing to Investigate the Oncogenic Activity of Mutant Calreticulin in Cytokine Dependent Hematopoietic Cells

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

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

Lentiviral CRISPR/Cas9-Mediated Genome Editing for the Study of Hematopoietic Cells in Disease Models

Gene Knock-in by CRISPR/Cas9 and Cell Sorting in Macrophage and T Cell Lines

Electroporation-Based CRISPR-Cas9-Mediated Gene Knockout in THP-1 Cells and Single-Cell Clone Isolation

CRISPR-Cas9 Mediated Gene Deletion in Human Pluripotent Stem Cells Cultured Under Feeder-Free Conditions