Generation of a RIP1 Knockout U937 Cell Line Using the CRISPR-Cas9 System

As CRISPR-related protocols become increasingly useful and accessible, complications and obstacles can still arise under specific experimental conditions. This protocol outlines the creation of a receptor-interacting serine/threonine-protein kinase 1 (RIPK1/RIP1) knockout human cell line using CRISPR/Cas9 and highlights potential challenges encountered during this process.

This protocol outlines a procedure for knocking out the RIP1 gene using CRISPR/Cas9 in the human monocyte U937 cell line. The method utilizes designated guide RNA plasmids and lentiviral packaging plasmids to achieve RIP1 gene knockout. The protocol addresses challenges and improvements to traditional CRISPR methods, enabling its replication for future cell death studies. The resulting mutant cells can be used to investigate mechanistic changes in cell death, where functional RIP1 proteins would otherwise play a role. Viability assays demonstrated a significant reduction in cell death in knockout cells following necroptosis induction. Fluorescence microscopy revealed a marked decrease in mitochondrial reactive oxygen species (ROS) in knockout cells under the same conditions. Together, these functional assays confirm the loss of RIP1 protein. Optimized for use with U937 human monocytes, this procedure may also be adapted to target other key cell death regulators, yielding functional, non-lethal mutants. Potential pitfalls are addressed throughout to provide insights into challenges that may arise during mutant generation.

The use of the CRISPR/Cas9 gene-editing technology has been rapidly evolving since its discovery^1,2,3. The ability to knock-in or knockout genes within cell lines or bacteria is invaluable to forwarding research and the understanding of intracellular mechanisms^1,2,3,4,5,6. The CRISPR-Cas9 system improves upon previous gene editing methods, such as transcription activator-like effector nuclease (TALEN), by simplifying the engineering of gene specificity. This procedure includes two fundamental components: guide RNA (gRNA) used to locate the intended gene target, and Cas9, which is an endonuclease that modifies the intended genome location with a double-stranded DNA break^3,4. The gRNA will act as a guide for the Cas9 endonuclease to locate and initiate the double-stranded break at the intended genetic sequence through Watson-Crick base pairing. The full process of genome editing by the CRISPR/Cas9 system involves cellular machinery repairing these double-stranded breaks in DNA through non-homologous end joining (NHEJ) or homologous recombination^3,7. It is more likely that NHEJ will occur, effectively creating a mutation in the genome that results in a loss of expression for the target gene^3,4.

Commercial sources have been able to create libraries of gRNA targets that can be expressed through bacterial growth and isolation, which significantly improves their ease of use. However, the main limitation of the CRISPR/Cas9 system is the difficulty in delivering the gRNA and Cas9 complex into target cell lines. These limitations arise in suspension cell lines, as they are generally referred to as hard-to-transfect⁸. Typical transfection methods are not generally efficient in delivering the CRISPR/Cas9 system into suspension cells, which is why viral delivery methods like lentiviral transfection and transduction are better suited for this type of cell line^8,9.

This type of transfection requires a lentiviral vector that encodes the gRNA and Cas9 endonuclease along with added lentiviral packaging plasmids, which are transfected into a cell line that is capable of manufacturing lentiviral particles. A typically chosen cell line for this process is HEK293T cells, as they are easier to transfect and work very efficiently in the assembly of gRNA and Cas9^9,10. These particles are then released as lentiviruses into the supernatant, which can be used to transduce the gRNA and Cas9 into the intended suspension cell line, such as U937 human monocytes. As such, the procedure described here has the following changes compared to established methods: (1) Alternate transfection method for hard-to-transfect cell lines; (2) No need to concentrate CRISPR plasmid DNA or use ultracentrifuge; and (3) It eliminates the need for single-cell cloning.

The direct focus of this article was to knockout the RIP1 gene in U937 human monocytes. The canonical form of the highly inflammatory cell death pathway necroptosis is controlled by RIP1, which serves as a pivotal target for cell death studies.^11,12,13,14 As RIP1 becomes active through autophosphorylation, it then recruits and causes direct phosphorylation and activation of receptor-interacting serine/threonine-protein kinase 3 (RIPK3/RIP3) and mixed lineage kinase domain-like (MLKL) pseudokinase to form the necrosome. Following this formation, the necrosome is free to move throughout the cell to interact with organelles such as the mitochondria^12,13. At the mitochondria, RIP1 potentiates a positive feedback loop with cellular metabolism, directly impacting the production of mitochondrial ROS, which in turn promotes further autophosphorylation of RIP1, necrosome formation, and the downstream execution of necroptosis^11,12,13,14.

While the focus of the current research group is the role of RIP1 in cell death, other reasons to study RIP1 include its roles in inflammation and infection. Upon activation by death receptors such as TNF receptors, RIP1 promotes the activation of the NF-κB signaling pathway, which triggers the transcription of pro-inflammatory cytokines, chemokines, and other molecules essential for immune cell recruitment and the amplification of the inflammatory response¹⁵. In addition to NF-κB activation, RIPK1 can also engage MAPK signaling pathways, further enhancing inflammation^15,16. Regarding its role in responses to infection, RIP1 acts as a pivotal mediator of the host inflammatory response, particularly in response to pathogen-associated molecular patterns (PAMPs) recognized by pattern recognition receptors (PRRs) such as Toll-like receptors (TLRs)¹⁷. Moreover, during sepsis, RIP1 is activated by signaling through death receptors such as the TNF receptor, leading to the initiation of pro-inflammatory cascades. RIPK1 mediates the activation of the NF-κB and MAPK pathways, promoting the production of pro-inflammatory cytokines such as TNF-α, IL-1β, and IL-6, which are key drivers of the systemic inflammatory response characteristic of sepsis¹⁸.

A schematic representation of the procedure is provided in Figure 1. Guide RNA (gRNA) and the target sequence are given in Table 1. Details of the reagents and the equipment used are listed in the Table of Materials.

1. Harvesting of RIP1 -targeting CRISPR gRNA lentiviral expression vector containing Cas9 endonuclease and puromycin resistance from Escherichia coli

1. Quadrant-streak E. coli harboring lentiviral vector gRNAonto LB agar plates supplemented with 100 µg/mL Ampicillin.
2. Incubate plates at 37°C for 1-2 days until individual colonies are visible in the most dilute area on the plate.
3. Select a single colony with a sterile loop and individually add each colony to 5 mL of LB broth supplemented with 100 µg/mL Ampicillin into a 50 mL conical tube. Ensure to pipette thoroughly up and down after adding the single colony to ensure homogenization.
4. Vent and tape down the cap of the 50 mL conical tube and incubate in an orbital shaker at 37°C and 225 rpm for 8 h.
5. During the 8 h incubation at 37°C, add 40 mL of LB broth supplemented with 100 µg/mL Ampicillin to each of four separate 50 mL conical tubes.
6. Following the 8 h incubation at 37°C, take 40 µL of grown E. coli culture and add it to all four 50 mL conical tubes.
7. Vent and tape down the caps of the 50 mL conical tubes and incubate them in an orbital shaker at 37°C and 225 rpm for 12-16 h.
8. Following this incubation at 37°C, spin down all tubes at 3220 x g in a swinging bucket rotor for 20 min to pellet the grown E. coli bacteria.
9. Decant supernatants to a waste beaker, then combine all four pellets in 10 mL of LB broth supplemented with 100 µg/mL Ampicillin.
10. Spin down the combined pellets again at max speed in a swinging bucket rotor for 20 min to get a singular pellet.
11. Discard as much of the supernatants as possible into the waste beaker, and proceed with one of the following options: (1) Freeze wet pellet as is at -80° for up to 1 month. (2) Proceed with lentiviral vector plasmid purification.
    NOTE: If option 1 is chosen, the protocol may be paused here.

2. Transfection of HEK293T cells with purified CRISPR gRNA lentiviral expression vector targeting RIP1

1. Seed HEK293T cells overnight at a concentration of 3 × 10⁶ to 5 × 10⁶ cells into a 10 cm plate containing DMEM with 4.5 g/L D-glucose, L-glutamine, 110 mg/L sodium pyruvate, and 10 % heat-inactivated FBS.
2. The following day, ensure plates have approximately 70%-90% confluency, then proceed with the protocol.
3. Setup the transfection of the cells with a 1:1:1:1 ratio of plasmids (experimental CRISPR plasmid: pLP1: pLP2: pLP/VSVG) using the transfection reagent in a 3:1 ratio (3 parts reagent: 1 part plasmid DNA), along with reduced serum medium.
   NOTE: The pLP1: pLP2: pLP/VSVG portion of the ratio is a lentiviral packaging mix.
   1. Equilibrate the transfection reagent, reduced serum medium, CRISPR plasmid DNA, and lentiviral packaging mix to room temperature prior to setup of transfection.
   2. Mix together 75 µL of the transfection reagent, 2500 µL of reduced serum medium, and 25 µg total DNA (calculation based on spectrophotometer analysis for CRISPR plasmid DNA and 1:1:1:1 ratio with lentiviral packaging mix).
   3. Allow this mixture to incubate at room temperature for 15-30 min. Carefully pipette the entire volume of this mixture to the confluent HEK293T cell plate right into the media (no need to change media).
4. Gently swirl the plate to ensure adequate homogenization of the transfection mix and plate media. Incubate the plate at 37 °C for 48 h.

3. Lentiviral transduction of U937 cells or target cells

1. Once the 48-h incubation at 37^°C is complete, transfer the media from the producer HEK293T cell plate into a 15 mL conical tube.
2. Centrifuge the volume at 800 x g for 5 min at room temperature to pellet any remaining HEK293T cells. Remove all the virus-containing supernatant, taking care not to disturb the pelleted HEK293T cells, and keep it in a separate 15 mL conical tube. Then, decontaminate and discard the pellet-containing tube in the appropriate biohazard waste container.
   NOTE: HIV-based lentivirus supernatants can be stored at -80°C; however, doing so carries a risk of up to 55% loss of virus stability after the first freeze/thaw cycle¹⁹.
3. Count and obtain a pellet of 2 × 10⁶ U937 cells in a 15 mL conical tube by centrifuging an appropriate volume at 400 x g for 10 min at room temperature and decant the supernatant, leaving the cell pellet in the tube.
4. Resuspend the U937 cell pellet with all the virus-containing supernatant and then centrifuge the tube at 290 x g for 60 min.
5. Following centrifugation, use a pipette to resuspend the pellet with the virus-containing supernatant already in the tube.
6. Place the tube on an end-over-end rotator (or similar rotating device) for 60 min.
   1. Following this incubation, centrifuge the tubes at 400 x g for 10 min at room temperature to pellet the cells.
   2. Resuspend the cell pellet with a 1:1 ratio media mixture of both virus-containing supernatants and complete RPMI-1640 media supplemented with 10% heat-inactivated FBS.
7. Transfer the cell mixture to a 10 cm tissue culture plate and incubate at 37°C for 48 h.
8. Following this incubation, centrifuge the U937 cells at 400 x g for 10 min and remove the supernatant.
   1. Resuspend the pellet with complete RPMI-1640 media supplemented with 10% heat-inactivated FBS and 5 µg/mL of puromycin and transfer the cells to a T25 flask.
   2. Incubate cells at 37 °C untouched for 2-3 weeks, ensuring to check on them every 1-2 days for signs of cell growth.
      NOTE: Once cells become confluent and have a healthy turnover time following an initial passage, they can be tested for the effectiveness of the completed protocol.

4. Testing effectiveness of the completed protocol in the creation of RIP1 CRISPR mutant cells using Western blot analysis

1. Generate cellular protein lysates of wild-type (WT) U937 monocytes, non-targeting control (NTC) CRISPR cells, and the RIP1 CRISPR mutant cells^11,12,13.
2. Run these lysates on an SDS-PAGE gel and proceed with Western blot analysis¹⁴.
   1. Normalize samples to a housekeeping protein first.
   2. Following normalization, analyze samples for RIP1 protein expression levels to adequately determine the success of creating a RIP1 CRISPR mutant cell line¹⁴.
3. Following western blot analysis conformation, cells can be used for experimentation purposes.

Following the production of a confluent population of RIP1 CRISPR mutant U937 cells, SDS-PAGE and Western blot analysis were performed. The Western blot analysis was used to determine the successful creation of a RIP1 CRISPR mutant cell line by assessing the loss of RIP1 protein expression levels. This determination was made based on the comparative result of WT U937 monocytes and NTC cells. In Figure 2, the expression of RIP1 was not detected for the 5 µg/mL puromycin RIP1 CRISPR mutant cells but was detected for both the WT U937 and NTC cells. This result demonstrates that this protocol is a reliable and effective method for selecting a viable RIP1 CRISPR mutant.

With the successful loss of RIP1 protein expression in the RIP1 CRISPR mutant cells, we then determined if this equated to a functional loss of RIP1 as well. In previously established studies, a combination of tumor necrosis factor-alpha (TNF-α), cycloheximide, and pan-caspase inhibitor zVAD-fmk is a direct stimulus of necroptosis that is abbreviated as TCZ^11,14. As seen in Figure 3, induction of necroptotic cell death with TCZ treatment is reverted following treatment with the known RIP1 inhibitor, necrostatin-1s (nec-1s), in both the WT U937 monocytes and NTC cell line. For the RIP1 CRISPR mutant cells, however, TCZ treatment resulted in a significant loss of cell death compared to WT and NTC cells. Subsequent treatment with nec-1s had no effect on RIP1 CRISPR mutant cells, indicating the absence of necroptosis and RIP1 function.

To further evaluate the loss of RIP1 functions within these RIP1 CRISPR mutant cells, live cell fluorescence microscopy was performed with a DNA Hoecsht stain (blue fluorescence) and a mitochondrial superoxide stain (red fluorescence). These experiments allowed us to determine if the previously established increase in mitochondrial superoxide production during necroptosis is lost in these cells. RIP1 is known to directly interact with cellular metabolism and significantly increase mitochondrial superoxide production. This abundance of superoxide within the cell, in turn, creates a positive feedback loop with the activation of RIP1 and downstream execution of necroptosis. As seen in Figure 4, treatment with TCZ showed an abundance of mitochondrial superoxide production in both the WT U937 monocytes and NTC cells, but not in the RIP1 CRISPR mutant cells. Subsequent treatment with nec-1s led to a significant decrease in red fluorescence staining for both the WT U937 monocytes and NTC cells, with no change being observed in the RIP1 CRISPR mutant cells.

Figure 1: Schematic model of CRISPR/Cas9 protocol. Visual representation of the various steps provided within this protocol for ease of understanding. Please click here to view a larger version of this figure.

Figure 2: Western blot analysis confirms loss of RIP1 protein expression in RIP1 CRISPR mutant cells. Expression of RIP1 protein was lost at a 5 µg/mL puromycin selection in RIP1 CRISPR mutant U937 cells when compared to WT U937 cells and NTC cells. Please click here to view a larger version of this figure.

Figure 3: Cell viability assay shows loss of function of RIP1 in RIP1 CRISPR mutant cells. Necroptotic cell death was induced via treatment with TCZ and analyzed with subsequent treatment with nec-1s. Both WT U937 monocytes and NTC cells showed a significant decrease in cell death following treatment with nec-1s. The RIP1 CRISPR mutant cells exhibited much less cell death when treated with TCZ and showed no change when subsequently treated with nec-1s. The results shown are from 3 independent experiments. Error bars represent standard deviation. Two-way ANOVA with Bonferroni post-test ***p < 0.001. ns = not statistically significant. Please click here to view a larger version of this figure.

Figure 4: Downstream functional analysis of RIP1 in RIP1 CRISPR mutant cells using fluorescence microscopy. (A) Measurement of mitochondrial superoxide staining (red fluorescence) showed a decrease in superoxide production in both WT U937 monocytes and NTC cells following nec-1s treatment. The RIP1 CRISPR mutant cells did not show production of mitochondrial superoxide when treated with TCZ and showed no change when subsequently treated with nec-1s. Scale bars: 100 µm. (B) Mean fluorescence intensity quantification of results represented in (A). The results shown are from 3 independent experiments. Error bars represent standard deviation. Two-way ANOVA with Bonferroni post-test, ***p < 0.001. Please click here to view a larger version of this figure.

Table 1: Guide RNA (gRNA) and target sequence.

This protocol aims to provide detailed instruction and analysis of potential pitfalls in the efficiency and reliability of lentiviral transfection and transduction to create an RIP1 knockout U937 cell line. Although this method of transfection and transduction is labor and time-intensive, it is generally considered to be an efficient way to incorporate the chosen gRNA and Cas9 endonuclease into hard-to-transfect cell lines^8,9,20. As is the case with monocytes like U937 cells and other immune cells, there is an inherent difficulty in performing transfection as they harbor innate sensors that recognize and respond to foreign nucleic acids. This can lead to nucleic acid destruction/hypermutation, transcriptional repression, and/or cell death in response to the foreign nucleic acids²¹.

This method of delivery is only possible using a cell line capable of manufacturing lentiviral particles, HEK293T cells, which are an adherent human embryonic kidney cell line that is widely accepted as easy to transfect^9,10. With this information, the most critical steps in this procedure are outlined that must be handled with care. Cell culturing and monitoring are of the utmost importance. This study used U937 cells as the target cell line for the gRNA and Cas9, which are fast-growing and easy-to-use suspension human monocytes²². As with most fast-growing cells, regular maintenance and passage of the cells is important to prevent them from becoming overgrown and not performing in experimentation as expected. The same concept applies to HEK293T cells in this protocol. Ensuring that these cell lines are properly maintained and kept at a healthy, confluent state during their use in the protocol is essential for efficient lentiviral production. Another critical step in this procedure is using the appropriate concentrations of transfection reagent and ratios of CRISPR plasmid DNA to lentiviral packaging mix. These mathematical proportions have been tested empirically under several different concentrations in this method, and the values listed are critical for the transfection of HEK293T cells.

As with most devised methods and procedures, there is a fair amount of troubleshooting that goes into the established working experimentation process. One issue determined here with other accepted protocols is the immediate challenge of puromycin following a shorter transduction step for the U937 cells. As the transduction step only utilizes 2 × 10⁶ U937 cells, a full 48 h of incubation with half of the media being complete RPMI-1640 supplemented with 10% heat-inactivated FBS was empirically determined to facilitate the generation of this RIP1 CRISPR mutant cell line. When the cells were continuously challenged with 5 µg/mL of puromycin without intervention, live and healthy U937 cells began to appear in the culture after 2-3 weeks. As this protocol is time-intensive, multiple iterations of this transfection and transduction step have several possible recoverable cultures. Once the cultures become confluent, one can check the effectiveness of the protocol through SDS-PAGE and Western blotting techniques. Even after successful isolation and confirmation of a RIP1 CRISPR mutant cell line, one would do well to perform routine monitoring of protein lysate-based SDS-PAGE and western blot analysis for cellular loss of RIP1 protein expression for continual use of these cells in experimentation.

Another reason for empirical troubleshooting is the lack of straightforward information available for the use of transfection reagents with immune cell lines. Between available information from companies and protocols, there is conflicting information regarding a concentration-based ratio for the transfection reagent used in this protocol to DNA. While attempting ranges from 5:1 to 3:1, respectively, it determined that 3:1 is the maximum ratio to be used, as otherwise, the transfection is not efficiently completed with the HEK293T cells. Along with this choice, it was determined that a singular iteration of CRISPR plasmid DNA at lower volumes was insufficient to fully transfect a 10 cm plate of HEK293T cells. As the plasmid purification kit yields a large volume of CRISPR plasmid DNA from E. coli, increasing the volumes of all transfection components, 5-fold yielded enhanced transfection efficiency for recovering the U937 cells after transduction. The volumes listed for these components in the protocol represent a 5-fold increase from a single iteration from accepted protocols. Between these changes in the transfection reagent ratio and the total volume used for these components, the protocol was devised without the need to concentrate CRISPR plasmid DNA or require the use of an ultracentrifuge. However, if these materials are available, the isolated CRISPR plasmid DNA can be concentrated using established methods, thereby decreasing the required volumes of other components of this protocol^23,24.

When using lentiviral transfection methods, limitations can be noted for their use in CRISPR/Cas9 gene editing. The most notable limitation is the time and labor requirements for this protocol^9,20. The procedure includes only two steps where it can be paused but is optimized for hard-to-transfect cell lines. For hard-to-transfect cell lines such as U937 monocytes, typical transfection methods, including chemical-mediated introduction of plasmids, are not as effective as lentiviral-based delivery²⁵.

The broader research interest focuses on the hyperglycemic shift from apoptosis to necroptosis in U937 monocytes, emphasizing the necessity of a functional knockout of RIP1 for mechanistic understanding during this process. As RIP1 is a critical protein in the execution of necroptosis, understanding all its functions and protein interactions within the cell is essential^11,12,13,14. Future directions of this protocol include applying these methods to further investigations and experiments regarding the hyperglycemic shift from apoptosis to necroptosis. This protocol is anticipated to be broadly applicable to lentiviral transfection and transduction methods with suspension cells for various gRNA targets using the CRISPR/Cas9 system, extending beyond U937 cells and the functional knockout of RIP1.

None.

This research was funded by the National Heart, Lung, and Blood Institute (NHLBI) of the National Institutes of Health (NIH), grant number NIH 2R15-HL135675-02 to T.J.L.