CRISPR/Cas9 gene editing in hematopoietic stem and progenitor cells to accelerate the translation of cellular therapies and immunotherapies

Abstract

Hematopoietic stem and progenitor cells (HSPCs) play a critical role in immune system regeneration and have therapeutic potential in hematological disorders and immunotherapy. However, gene editing in HSPCs remains challenging, particularly in murine HSPCs, limiting preclinical studies and translational advancements.

Here, we present a CRISPR/Cas9 mRNA-based protocol for efficient gene editing of human CD34+ HSPCs and murine HSPCs. Using electroporation of Cas9 mRNA and sgRNAs, we achieved up to 80% gene knock-down in HSPCs while retaining cell viability. Edited murine HSPCs successfully reconstituted bone marrow in lethally irradiated mice, demonstrating functional engraftment. We also demonstrated that knocking-out IL-6R in murine HSPCs and CD34+ cells skewed myeloid differentiation toward less immunosuppressive phenotypes, and combining IL-6R-knock-out murine HSPCs with immune checkpoint inhibitors enhanced glioma treatment efficacy in preclinical models, showcasing the potential of this method to facilitate translation of cellular therapies.

This method offers a scalable and cost-effective approach to genetic modification of HSPCs, supporting advancements in hematopoietic stem cell transplants (HSCTs) and next-generation immunotherapies. By streamlining gene editing for both preclinical and translational applications, this protocol has the potential to accelerate the development of personalised cellular therapies and improve clinical outcomes in oncology and immune-related disorders.

Introduction

Hematopoietic stem and progenitor cells (HSPCs) are the only cells in the immune system capable of long-term self-renewal and multipotency, and can therefore differentiate into all cell lineages (1), making them an attractive treatment for multiple diseases (2, 3).

In mammals, HSPCs are rare, with only approximately 10,000 long-term HSPCs in the entire marrow compartment present at one time, and only 1,000 of them simultaneously undergo hematopoiesis in humans (4, 5).

Gene engineering of HSPCs is currently at the forefront of the development of improved HSPC therapies. Even though successful HSPC editing has been achieved (6–8) and large numbers of CD34+ cells (>2 × 10⁶ CD34⁺ cells per kg of body weight) can be mobilised for collection for transplantations (9, 10), engineering of large numbers of HSPCs can be challenging, costly, and time consuming, leaving an urgent need to develop simpler methods for HSPC editing to accelerate the development of cellular therapies.

In human CD34+ cells, HSPC editing has been evaluated for diseases such as sickle cell anemia, β-thalassemia, storage and metabolic disorders, and HIV (11–16). Although some gene therapy methods using retroviruses and lentiviruses have been used to modify human CD34+ cells in multiple disease settings, safety is still a concern, specifically for their ability to insert genes in a semi-random manner (17). On the other hand, novel CRISPR-based gene editing approaches have shown promise in the field as they are site-specific, can overcome random insertion of sequences, have higher editing efficiency, are easy to design, and relatively inexpensive compared to other methods (18, 19).

Despite advances in the human setting, methods to successfully edit murine HSPCs (mHSPCs) to assess the efficacy of these therapies in animal models remain limited (20–22). mHSPCs have been more challenging to engineer because of their scarcity in the murine bone marrow, requiring larger amounts of starting material and more time to harvest, potentially resulting in higher cell death during experimental procedures (5, 23–25). Other factors that contribute to this difficulty are immunological barriers, such as innate immunity of HSPCs to gene therapy reagents (lentiviruses, gene-editing enzymes, etc.), or immunity to HSPC neoantigens introduced as part of the gene therapy (17, 22). Creating a reliable method to successfully engineer mHSPCs would allow for the contemporary development of therapies in the preclinical stage, while allowing for mechanistic laboratory experiments.

The utility of mRNA technologies has been highlighted by the efficacy of COVID-19 mRNA vaccines (26, 27). In this study, we propose a highly efficient and scalable RNA-based method for the genetic modification of mHSPCs and human CD34+ HSPCs using Cas9 mRNA to facilitate the development of novel HSPC therapies. As HSPCs are naturally capable of differentiating into any immune cell in the body, we leveraged this to drive cell fate differentiation of HSPCs using Cas9 mRNA coupled with guide RNAs (gRNA) for select genes. Here, we demonstrate this concept by gene-editing IL-6R from murine bone marrow-derived HSPCs to prevent their differentiation into immune regulatory myeloid-derived suppressor cells (MDSC). As expected, IL-6R^KO HSPCs led to a decreased proportion of MDSCs and macrophages. Ultimately, the use of IL-6R^KO HSPCs provided a therapeutic benefit when combined with anti-PD-1 blockade, as there was a significant increase in survival relative to anti-PD-1 blockade alone.

Materials and methods

Experimental design

Ethical approval for this study was granted by the University of Florida Institutional Review Board (IRB #201300482) and the University of Florida Institutional Animal Care and Use Committee (IACUC #202200000409 and #202100000053). This study was also conducted in adherence to the Declaration of Helsinki and to the ARRIVE guidelines 2.0. A complete checklist is provided in the supplementary materials.

Isolation and culture of Lin– murine HSPCs

Mouse hematopoietic stem and progenitor cells were obtained from the bone marrow of tibias and femurs of 5–9-week-old C57BL/6J mice after red blood cell lysis and depletion of lineage-positive populations using a magnetic bead isolation kit (Miltenyi Biotec, cat. 130–090–858), as previously described (28). Cells were plated at 0.5 × 10⁶–1 × 10⁶ cells/ml in SFEMII+10x media (SFEMII: StemCell Technologies, cat. 9655; CD34+ Expansion Supplement (10X): StemCell Technologies, cat. 02691) in 6-well or 24-well plates. Cells were split by dilution to 0.5 × 10⁶ cells/ml when the concentration surpassed 1.5 × 10⁶ cells/ml.

Isolation and culture of human CD34+ HSPCs (hCD34+ cells)

CD34+ HSPCs were obtained by processing cord blood obtained from healthy donors commercially available from LifeSouth Community Blood Centers (Gainesville, FL, USA). Permission to use cord blood in this study was granted by the University of Florida Institutional Review Board under permit number 201300482. Mononuclear cells were obtained from whole cord blood. The cells were then subjected to CD34+ selection using the human CD34 MicroBead Kit UltraPure (Miltenyi Biotec, cat. 130–100–453). Cells were plated at 0.8 × 10⁶–1 × 10⁶ cells/ml in SFEMII+10x media in 6-well, 24-well, or 48-well plates, depending on the starting cell number. Cells were split by dilution to 0.5 × 10⁶–0.8 × 10⁶ cells/ml when the concentration surpassed ∼1.5 × 10⁶ cells/ml.

Mice

Female 5–10 week-old C57BL/6J mice (Jackson Laboratories, 000664) and C57BL/6-Tg(UBC-GFP)30Scha/J (referred to as GFP mice, Jackson Laboratories, 004353) were used for the experiments. For survival studies, mice were closely monitored and euthanized at endpoint, according to IACUC (Institutional Animal Care and Use Committee) guidelines. Animals subjected to this surgical implantation show some post-operative pain and distress, for which they will be administered buprenorphine (analgesic) preoperatively as well as post-operatively as needed (mice: 0.05–0.1 mg/kg, s.c.). These are standard analgesic doses for rodents and have a duration of action of approximately 12 hrs. All animals were euthanized using inhalant CO2 as per IACUC protocol. Secondary method is conducted in all animals using either cervical dislocation or exsanguination.

The investigators adhered to the ‘Guide for the Care and Use of Laboratory Animals’ as proposed by the Committee on Care of Laboratory Animal Resources Commission on Life Sciences, National Research Council. The facilities at the University of Florida Animal Care Services are fully accredited by the American Association for the Accreditation of Laboratory Animal Care.

sgRNA design

eGFP sgRNA (GGGCGAGGAGCTGTTCACCG) was copied from lentiCRISPR – EGFP sgRNA 1 plasmid from Feng Zhang (Addgene plasmid #51760; http://n2t.net/addgene:51760; RRID:Addgene 51760) (29) and purchased from ThermoFisher Scientific ‘Custom gRNA order’ option. Mouse and human IL-6RA sgRNAs were purchased from the predesigned TrueGuide Synthetic gRNA database from ThermoFisher Scientific (cat. #A35533; CRISPR591728_SGM, CRISPR591748_SGM, CRISPR882880_SGM, and CRISPR882881_SGM).

Electroporation conditions

Electroporation was performed at a concentration of 10 × 10⁶ cells/ml in a 1 mm cuvette (BTX, cat. 45–0134) in Opti-MEM^™ (Gibco, cat. 31985070) or BTXPRESS electroporation solution (BTX, cat. 45–0801), using a BTX electroporator (ECM 830, Harvard Apparatus, cat. 45–0052). The optimised conditions for electroporation of HSPCs were 1 pulse of 125 V and 5 ms (field strength = 1250 V/cm), and all further experiments were performed under these conditions. For gene editing, Cas9 mRNA and sgRNA were combined at a ratio of 0.76:0.24 (Cas9 mRNA:sgRNA) per 0.5 × 10⁶ cells. The total amount of RNA per electroporation ranged from 2.5 µg to 10 µg, with comparable editing efficiency across this range. Note: The upper end of the range is included as a practical option: when performing only a single electroporation (0.5 × 10⁶ cells) at the lower RNA amounts (2.5 µg total RNA), the required volume of sgRNA (e.g. 0.6µg = 0.6 µL at 1 µg/µL) can be difficult to pipette accurately. Using a higher total RNA amount while maintaining the same ratio facilitates easier handling without compromising efficiency or cell viability. These RNA amounts and ratios are also detailed in Table 1.

Table 1.

Preparation of RNA mix (Cas9 mRNA and sgRNAs) for electroporation in HSPCs.

electroporation	RNA/Electroporations (μg)	Volume/ electroporation (μL)	Number of cells	RNA ID	RNA ratio	RNA concentration (e.g. for a 2.5 μg total RNA electroporation)
1	2.5–10	50	0.5 × 106	sgRNA	0.24	0.6μg
				Cas9	0.76	1.9μg

Glioma conditioned media

Tumor cell supernatants were collected two days after plating 2 × 10⁶ murine or human glioma cells (KR158B-luciferase or U87, respectively) in a 10 cm dish with 10 ml of DMEM (ThermoFisher Scientific, cat. 11965118) supplemented with 10% fetal bovine serum (FBS) (Avantor Seradigm cat. 89510–186), and 1% Penicillin–Streptomycin (5,000 U/mL) (ThermoFisher Scientific, cat. 15070063). After that, supernatants were filtered using a 0.22μm syringe filter (Millipore Sigma, cat. SLGVR33RS).

Flow cytometry

Flow cytometry was performed on FSC/SSC gating on the BD Biosciences FACSCanto-II and FACSymphony-A3. Cells were collected and washed with phosphate-buffered (PBS) (Gibco, cat. 10010023) before staining in 2% FBS (Avantor Seradigm cat. 89510–186) in PBS according to the panels provided below (Tables 2–4). TruStain FcX Antibody (BioLegend, cat. 101320 or cat. 422301) was used to prevent nonspecific binding when possible. Analysis was performed using FlowJo V.10 (FlowJo, LLC). LIVE/DEAD Fixable Blue Dead Cell Stain Kit (ThermoFisher, cat. L23105). LIVE/DEAD Fixable Yellow Dead Cell Stain Kit (ThermoFisher, cat. L34959). TruStain FcX Antibody (Biolegend, cat. 101320). APC anti-mouse CD126 (IL-6Rα chain) Antibody (Biolegend cat. 115811). PE anti-human CD126 (IL-6Rα) Antibody (Biolegend cat. 352803).

Table 2.

Murine HSPC panel for flow cytometry.

Antibody	Fluorophore
Lineage	FITC
Il-7Ra (CD127)	PE-Dazzle
CD117 (c-kit)	PE
Sca1	PE-Cy7
CD34	APC
CD16/32 (FcyR)	APC-Cy7
CD115 (M-CSFR)	PerCP-Cy5.5
CD135 (Flt3)	BV421
Blue Viability Dye	DAPI (BUV496)

Table 4.

Human HSPC and myeloid panel for flow cytometry.

Antibody	Fluorophore
CD34	PE-Cy7
HLA-DR	APC-Cy7
CD11c	BV711
CD11b	PercP-Cy5.5
CD15	FITC
CD14	BV785
IL-6R	PE
CD38	BV650
CD90	APC
CD45RA	Pacific Blue
Yellow Viability Dye	Qdot605/BV605

Bone marrow rescue

For bone marrow rescue experiments, C57/BL6J mice were lethally irradiated with a dose of 9Gy (X-RAD 320, Precision X-RAY Irradiation). The day after, mice received a dose of 0.25 × 10⁶–0.5 × 10⁶ HSPCs. For survival studies, mice were closely monitored and euthanized at endpoint according to IACUC guidelines.

Statistical analysis

All experiments were analyzed using Prism 5–10 (GraphPad). One-way analysis of variance (One-way ANOVA) with Dunnett’s, Tukey’s, or Šídák’s multiple comparisons test was used for experiments with multiple groups with one independent variable. Mantel–Cox log-rank test was used for survival experiments. Statistical significance was determined at p < 0.05. All experiments were repeated at least twice unless otherwise indicated. Animal studies were powered to include 5 randomised mice per group, unless otherwise noted. No animals or samples were excluded from analysis.

Limitations

This protocol is only suitable for gene knock-out of both murine and human hematopoietic stem and progenitor cells but has not been tested for the introduction of sequences into the genome of HSPCs. This will be addressed in future studies.

Procedure

This section describes a detailed protocol for genetically modifying murine and human hematopoietic stem and progenitor cells. For murine HSPCs, we recommend performing gene modification on day 1 post-isolation to avoid their differentiation. If differentiation is not a concern, cells can be cultured at 0.5–1 × 10⁶ cells/ml to allow cells to expand before modification. For human CD34+ HSPCs, we recommend expanding cells for 3–8 days in vitro at 0.8–1 × 10⁶ cells/ml to obtain higher numbers of CD34+ cells before modification.

Notes: All centrifugation steps are performed at 500rcf for 5 min at room temperature (∼22°C). Manipulation of cells and RNAs are to be done in a sterile tissue culture hood. RNA must be kept on ice throughout the process to avoid RNA degradation.

Preparation of hematopoietic stem and progenitor cells

Timing: allow 3–4 h to isolate HSPCs first; after that, 15–20 min of preparation.

Pause point

If stemness is not a critical factor, cells can be isolated and cultured in vitro for up to 8–10 days before gene modification.

• 1After successful isolation of HSPCs, centrifuge the cells to obtain a cell pellet.
• 2Resuspend cells in PBS to wash out the remaining media and perform manual counting of viable cells using a hemocytometer and trypan blue dye to exclude dead cells.
• 3After determining the cell count, centrifuge cells again and resuspend in Opti-MEM at a concentration of 10 × 10⁶ cells/ml.
• 4Determine the number of electroporations (0.5 × 10⁶ cells/electroporation = 50 µl of cell solution) to perform and the amount of RNA mix needed before proceeding.

Preparation of gene editing reagents and equipment

Timing: thaw Cas9 mRNA and sgRNAs on ice for at least 15 min before electroporation, 15–30 min for preparation, and 30 min for media warm-up at 37°C.

• 5
  Add SFEMII+10x media to the corresponding number of wells in either a 24-well plate or a 6-well plate, depending on the total number of cells being electroporated. Do not exceed 1 × 10⁶ cells/ml concentration. Allow for at least 30 min for media to warm up at 37°C.

  1. Example: When electroporating 1 × 10⁶ HSPCs, add 0.5 ml of media to each of the two wells in a 24-well plate. After electroporation, each well will receive 0.5 × 10⁶ HSPCs, for a final concentration of 1 × 10⁶ cells/ml.
• 6Unpack 1 mm cuvettes and open their caps.
• 7
  Turn on BTX electroporator, configure the settings before proceeding to next steps:

  1. Murine HSPCs and human CD34+ cells: 125 V, 5 ms, LV mode, 1 pulse, 1250 V/cm
• 8Once the cells are resuspended in Opti-MEM, prepare the RNA mix according to Table 1.

Critical

Steps 8–16 need to be performed quickly because RNA is sensitive and can be degraded easily. It is recommended to work quickly in the following steps until electroporation is completed.

Electroporation of Cas9 mRNA and sgRNAs in HSPCs

Timing: 5–15 min, depending on number of electroporations.

• 9Remove tissue culture plate with warmed-up media from the 37°C incubator and transfer into the sterile tissue culture hood.
• 10Add RNA mix to HSPCs resuspended in Opti-MEM. Carefully pipette up and down a couple of times and quickly transfer 50μL of cell suspension (50μL of cells = 0.5 × 10⁶ cells at 10 × 10⁶ cells/ml) to each cuvette.
• 11Work fast: Close the cap on the cuvette and transfer no more than 2 cuvettes at a time to the electroporator area. One at a time, introduce a cuvette in the machine and pulse cells with the pre-arranged settings.
• 12Immediately, remove the cuvette and repeat with the second cuvette. Quickly thereafter, transfer cuvettes back into the sterile tissue culture hood.
• 13Using a transfer pipette, remove cell suspension from the cuvette and transfer into tissue culture plate wells containing media.
• 14Repeat with remaining electroporations.
• 15Optional: to increase cell recovery from the cuvette, you can add ∼50μL of media back into the cuvette to collect any remaining cells. Transfer back to the wells with media.
• 16After all cells have been electroporated, transfer cells to an incubator at 37°C to allow recovery.

Critical

It is recommended to work in batches if the number of electroporations is greater than 10, which is equal to 5 × 10⁶ cells. Prepare different aliquots of cells and RNA mixes to avoid RNA degradation and increase editing efficiency. Keep RNA mixes on ice until ready to be added to the cell solution.

After electroporation, aim for 0.5–1 × 10⁶ cells/ml concentration to allow for successful recovery. If cells are too diluted or too concentrated, it can affect their viability and expansion after electroporation.

Recovery of modified HSPCs and downstream procedures

Timing: 1–24 h to allow for recovery before performing in vivo downstream experiments; or 2–3 days for in vitro downstream purposes and for assessing gene-editing efficiency.

1. For in vitro downstream purposes, culture cells at 37°C for at least two days to allow for successful cell recovery and editing efficiency.

2. If cells are to be used for in vivo experiments (e.g. transplantation of HSPCs into mice) on the same day, culture the cells at 37°C for 1–2 h before collecting them for in vivo downstream procedures.

Critical

Since electroporation causes stress to HSPCs, it is recommended to wait until the following day for collection, as this increases cell recovery and ensures more accurate viability counts before proceeding to downstream procedures.

Assessment of efficiency of gene modification in modified HSPCs

To determine whether gene modification is successful, we can assess efficiency using flow cytometry to easily and quickly determine knock-down of protein expression, and/or Sanger sequencing to determine gene disruption at the DNA level.

Note: if antibodies against a specific gene of interest are not available for flow cytometry, western blotting is another option for determining protein expression.

• 17After at least two days in culture, collect cells by pipetting up and down a few times. Wash and collect the remaining cells by adding PBS and pipetting up and down again.
• 18Transfer cells to FACS tubes. Collect at least 100,000–200,000 cells per condition to run the samples (see ‘Flow cytometry’ section).

Pause point

Cell pellets can be directly stored at −20°C or immediately processed for DNA extraction. Make sure to remove the supernatant completely without disturbing the cell pellet before storing at −20°C.

Results

In the last few decades, gene engineering of murine and human HSPCs for the development of cellular therapies has been a challenge. Despite recent advances in the modification of human CD34+ cells in preclinical and clinical settings, protocols for the successful modification of mHSPCs remain limited. Here, we established a reliable protocol to genetically modify both murine and human HSPCs.

Cas9 mRNA-based CRISPR achieves 80% decrease in gene expression in murine HSPCs

We first optimised the electroporation conditions to deliver mRNA to Lineage-negative (Lin-)mHSPCs and observed 125 V for 5 ms (field strength = 1250 V/cm) to be optimal (Figure S1a), but mRNA expression was transient after electroporation (Figure S1b).

Next, we tested the efficiency of Cas9 mRNA KO in mHSPCs. To do so, we isolated Lin– HSPCs from GFP+ mice and targeted them by delivery of a sgRNA against GFP in addition to Cas9 mRNA by electroporation (Figure 1a). After two or three days, we performed flow cytometry and fluorescence imaging of the mHSPCs to determine GFP expression. The results indicated a 5-fold (80%) decrease in GFP expression out of the total frequency of live cells and a >13-fold decrease in mean fluorescence intensity (MFI) (Figure 1b,c).

Figure 1. Cas9 mRNA KO yields an 80% reduction in GFP expression of mHSPCs in vitro.

a) Schematic of Cas9 mRNA and sgRNA electroporation for gene KO in Lin – mHSPCs. b) GFP expression and viability after GFP KO measured by flow cytometry and gated on live HSPCs. Associated plots are shown in (c). Expression was measured after two days in culture. c) Fluorescence imaging of WT, Cas9, and GFP^KO groups. GFP^KO group shows significant reduction in GFP expression compared to controls. n = 3 per condition. Images on the left pane: bright field; images on the right pane: GFP fluorescence. Statistics: b) One-way ANOVA with Dunnett’s multiple comparisons test was used to establish statistical significance. *p < 0.05; ****p < 0.0001; ns = not significant. Nomenclature: WT = wild-type control; Cas9 = Cas9 control; KO = knock-out. Experiments were repeated twice.

We also proved that this technique is effective for KO of genes that have a lower baseline expression in mHSPCs, as shown by the significant decrease in the expression of the interleukin 6 receptor (IL-6RA) gene in mHSPCs, with individual and combination of two different sgRNAs (Figure 2a,b).

Figure 2. Cas9 mRNA IL-6R KO achieves 4-fold decrease in gene expression and shifts myeloid phenotype in murine HSPC-derived cells.

a) Schematic of knock-out (KO) and in vitro culture of Lin – mHSPCs. b) Cas9 mRNA IL-6R KO decreases IL-6R expression by 4-fold compared to controls while maintaining high viability levels. c) In vitro expansion rates of unmodified WT mHSPCs for 7 days are variable (n = 2; n = 1, respectively). b-c) HSPCs were cultured in SFEMII +10x media as indicated in the ‘Experimental Design’ section. d) Modified mHSPCs expand at similar rates to unmodified mHSPCs after a 7-day in vitro expansion. e) Frequencies of Lin+ DCs, MDSCs, and macrophages after 3-day immunosuppressive culture. Associated flow plots are shown in (g). f) Schematic of knock-out and in vitro differentiation assay of Lin – HSPCs. g) Flow plots for the frequencies of DCs, MDSCs, and macrophages at each of the conditions: WT HSPCs (control), Cas9 HSPCs (control), and IL-6R^KO HSPCs (only IL-6R^KO gRNAs1 + 2 plots shown). d-g) HSPCs were cultured in SFEMII+10x for two days, then transferred to 50% SFEMII+10x and 50% KR158B-luc conditioned media for the remainder of the experiment. b,d,e,g) KO was performed with two different sgRNAs targeting the IL-6RA gene, individually, and in combination. b-g) mHSPCs for each experiment were isolated from the same batch of mice and split into the different shown conditions. Statistics: b,e) One-way ANOVA with Dunnett’s multiple comparisons test was used to establish statistical significance.*p < 0.05;**p < 0.005;***p < 0.0005;****p < 0.0001. Only significant values showcased for brevity, comparison to Cas9 only control. n = 2 for WT and Cas9 control groups; n = 3 for IL-6R^KO sg1, sg2, and sg1 + 2 groups. Nomenclature: sg = single-guide RNA; WT = wild-type control; Cas9 = Cas9 control; KO = knock-out. Experiments were repeated three times.

These findings suggest that delivery of Cas9 in the form of mRNA is a successful method for gene knock-out in mHSPCs, as it provides a balance of viability, effectiveness, and simplicity, which is extremely important when engineering HSPCs, as these cells are resistant and fragile to gene modification.

Gene-modified mHSPCs expand equally as WT-mHSPCs over a 7-day period

In vitro culture of mHSPCs routinely yields 7–10-fold expansion with unmodified mHSPCs over a period of 7 days when cultured in SFEMII+10x media. Murine HSPC and human CD34+ HSPC culture media (referred to as SFEMII+10x): Combine 9 ml of SFEMII with 1 ml of CD34+ Expansion Supplement (10X) to dilute to 1x. It is recommended to count the cells every 2 or 3 days. Occasionally, it can yield >50-fold expansion depending on factors such as the starting cell number of the culture, concentration, and frequency of passaging (Figure 2c). Obtaining these rates of expansion is critical to accurately assess KO efficiencies after gene modification of mHSPCs, as large cell numbers are needed for analysis via flow cytometry, western blotting, or Sanger sequencing. Therefore, it is recommended to start mHSPC cultures at a concentration of 1 × 10⁶ cells/ml and reduce to 0.5 × 10⁶ cells/ml the following day. We also recommend diluting the cells to 0.5 × 10⁶ cells/ml with fresh media when the concentration is higher than 1–1.5 × 10⁶ cells/ml, which usually aligns with every two or three days. We also wanted to determine whether gene editing affected the rate of expansion of viable mHSPCs. After comparing unmodified versus modified mHSPCs, we observed no differences in fold expansion after 7 days in vitro (Figure 2d).

Cas9 mRNA IL-6R KO skews HSPC myeloid differentiation under immunosuppressive culture conditions

After the successful modification of mHSPCs, we wanted to determine whether IL-6R KO skewed HSPC myeloid differentiation. To do so, we used Cas9 mRNA and two different gRNAs against IL-6Rα, individually and in combination, to KO IL-6R in freshly isolated HSPCs from the bone marrow of naïve C57BL/6 mice. After two days of culture, we measured KO efficiency by flow cytometry using a murine IL-6R antibody. The results indicated a ∼4-fold decrease in IL-6R expression (Figure 2a, b).

To determine whether IL-6R KO induced changes in HSPC myeloid differentiation, we performed phenotypic analysis by flow cytometry at different time points after KO. The results showed a significant increase in CD11c+ MHCII+ DCs after 2 days of culture in SFEMII+10x. Interestingly, we also observed a significant reduction in CD11b+ GR-1+ MDSCs and CD11b+ F4/80+ macrophages in IL-6R^KO HSPCs compared to WT control, but not when compared to Cas9 control (Figure S2a,b). We also observed a delay in early differentiation after IL-6R^KO, as indicated by the reduced proportion of Lin+ HSPCs in the IL-6R^KO groups compared to WT (Figure S2c,d). We inferred that this could be due to the longer time needed for recovery of electroporated vs. non-electroporated cells, since the differences with Cas9 control were not as significant.

Our laboratory and others have previously shown that glioma-conditioned media (GCM) induces MDSC and macrophage proliferation while suppressing DC proliferation (Figure S3) (28, 30). Therefore, we wanted to test whether IL-6R^KO would overcome GCM immunosuppression. To test this, we performed an in vitro differentiation assay of mHSPCs in 50% KR158B-luciferase-tumor-conditioned media and 50% SFEMII+10x, and phenotyped myeloid populations by flow cytometry (Figure 2f). After 3 days, we observed a significant increase in DCs after IL-6R^KO, as well as a significant decrease in MDSCs and macrophages (Figure 2e,g). Altogether, we identified IL-6R as a target for altering HSPC myeloid differentiation.

Gene-modified mHSPCs retain their reconstitution capabilities

Next, we wanted to determine whether modified mHSPCs by Cas9 mRNA electroporation maintained their bone marrow reconstitution functions. We first irradiated naïve healthy C57BL/6J mice with a lethal dose (9Gy) of total body irradiation to ablate the bone marrow. The following day, we delivered a single dose of 0.2 × 10⁶–0.3 × 10⁶ mHSPCs isolated from naïve healthy mice to each irradiated mouse to allow for rescue and reconstitution of the ablated bone marrow (Figure 3a). The results indicated that modified HSPCs remain functional and can repopulate the murine bone marrow compartment. 100% of mice survived for at least 75 days post-lethal irradiation in both modified, and WT and Cas9-only controls, indicating that electroporated modified mHSPCs retained their ability to reconstitute the bone marrow compartment (Figure 3b).

Figure 3. IL-6R^KO HSPCs retain bone marrow reconstitution abilities and provide superior efficacy in preclinical glioma.

a) Schematic of bone marrow rescue experiments in naïve C57BL/6J mice. b) IL6R^KO mHSPCs rescue lethally ablated C57BL/6J mice and 100% of the mice survive past 5 months. All mice received a lethal dose of irradiation (9Gy) on day 0, and received 200,000–300,000 mHSPCs (WT, Cas9, or IL6R^KO) the following day. This experiment was performed twice, once with Cas9 HSPCs only, and the representative experiment shown here with all groups. c) IL6R^KO mHSPCs provided superior efficacy in GL261 glioma. C57BL/6J mice were implanted with 10,000 GL261 cells, and treated with 250,000 HSPCs and 0.2 mg of anti-PD-1 drug each 5 days later.*p < 0.05,ns = not significant by Mantel-Cox Log-rank test (n = 5). Nomenclature: WT = wild type control; Cas9 = Cas9 control; KO = knock-out; aPD-1 = anti-PD-1 blockade; Gy = Gray.

IL-6R^KO HSPCs increase overall survival in preclinical glioma

We previously showed that the addition of HSPCs to two different immunotherapies significantly improved the survival of mice with aggressive gliomas (31–33). We then wanted to explore whether IL-6R^KO HSPCs provided a therapeutic benefit in preclinical glioma; therefore, we implanted C57BL/6J mice with GL261 glioma cells and treated them with either WT HSPCs + anti-PD-1 blockade or IL-6R^KO HSPCs + anti-PD-1 blockade. These results indicated that IL-6R^KO HSPCs + anti-PD-1 provided superior efficacy compared to WT HSPCs + anti-PD-1, demonstrating the therapeutic potential of gene-engineered HSPCs against tumors (Figure 3c).

Cas9 mRNA is efficacious to modify human CD34+ HSPCs

Next, we tested whether our Cas9 mRNA protocol was effective in modifying human cord blood CD34+ HSPCs. We first isolated and expanded CD34+ cells in vitro in SFEMII+10x for >4 days to obtain a minimum of 0.1 × 10⁶ cells per replicate for modification. Unmodified CD34+ cell expansion rates reached up to 79-fold after 7–8 days (Figure 4d). The addition of 50% U87-GCM to CD34+ cells resulted in >300-fold expansion (Figure S4a).

Figure 4. Cas9 mRNA IL-6R^KO achieves >7-fold decrease in gene expression in human CD34+ cells.

KO was performed with the combination of two different sgRNAs targeting the human IL-6RA gene. a) Viability after IL-6R^KO in CD34+ cells (n = 3). b) Frequency of CD34+ cells after IL-6R^KO (n = 3). c) Cas9 mRNA KO significantly decreases IL-6R expression compared to controls (n = 3). d) In vitro expansion rates of unmodified CD34+ cells. e) In vitro expansion rates of WT, Sham, Cas9, and IL-6R^KO CD34+ cells for 3 days. f) IL-6R^KO reduced monocytic MDSC frequency after a 4-day 50% GCM in vitro differentiation culture (n = 2). Statistics: One-way ANOVA with Dunnett’s, Šídák’s, or Tukey’s multiple comparisons test was used to establish statistical significance. *p < 0.05; **p < 0.01; ***p < 0.0005; ****p < 0.0001. Nomenclature: WT = wild type control; Sham = sham electroporation; Cas9 = Cas9 control; KO = knock-out; HD = Healthy Donor. Experiments were repeated >3 times using CD34+ cells isolated from 4 different healthy donors’ cord blood.

Our results indicated that Cas9 mRNA and sgRNAs against the human IL-6R gene (hIL-6R) mediated a highly efficacious KO in hCD34+ HSPCs, resulting in a >7.5-fold reduction in hIL-6R expression compared to controls (Figure 4c). Notably, viability remained >55% after electroporation and knock-out in CD34+ cells (Figure 4a). The frequency of CD34 expression varied among different attempts (Figure 4b). Contrary to our observations in mHSPCs, the expansion rates of Sham and Cas9 controls, along with IL-6R^KO were lower than those of WT (Figure 4e).

Finally, we wanted to determine whether IL-6R^KO affected the differentiation of human CD34+ cells. After IL-6R KO, CD34+ cells were placed in 50% U87 glioma-conditioned media. After 4 days, we performed flow cytometry to assess phenotypic differences between WT and IL-6R^KO HSPCs. The results indicated that IL-6R^KO skewed the differentiation of myeloid populations, as shown by the reduction in CD14+ HLA-DR^lo monocytic MDSCs (Figure 4f). However, IL-6R^KO effect on the frequencies of DCs, macrophages, and granulocytic MDSCs was inconclusive, as it varied in different attempts (Figure S4).

Discussion

Studies evaluating different methods for CRISPR/Cas9 gene editing of CD34+ HSPCs have been performed in human cell settings. However, limited protocols are available to efficiently and easily perform gene knock-outs in murine HSPCs. Here, we provide a comprehensive protocol using Cas9 mRNA and sgRNA electroporation to successfully edit both mHPSCs and human CD34+ cells that can also be used to drive changes in HSPC differentiation. Additionally, the brevity of this protocol allows for a reduction in experimental time after HSPC isolation. This is essential for successfully creating therapy delivery protocols, as it allows for larger numbers of engineered HSPCs for use in in vivo settings.

We also showed that the viability and expansion of the modified mHSPCs were comparable to those of unmodified mHSPCs. Furthermore, modified mHSPCs are capable of bone marrow reconstitution and can be utilized for multiple therapeutic purposes. This method can also be used to genetically modify CD34+ HSPCs. In this case, viability can be affected by gene modification, and expansion can be affected by electroporation. In both human CD34+ cells and mHSPCs, we showed that IL-6R^KO skewed myeloid differentiation to generally less immunosuppressive phenotypes, highlighting IL-6R modifications as potential therapeutic targets in HSPCs to reduce immunosuppressive populations.

HSPCs are inherently sensitive to ex vivo culture and gradually lose stem and progenitor characteristics over time, making it unrealistic to maintain a fully undifferentiated Lin⁻Sca-1⁺c-Kit⁺ (LSK) population throughout an 8-day protocol. This is consistent with recent studies showing that both murine and human HSPCs progressively downregulate stemness markers and activate differentiation programmes during in vitro expansion, even under cytokine-enriched, serum-free conditions (34, 35). For this reason, our approach was not designed to exclusively edit long-term HSCs but rather to target the broader hematopoietic stem/progenitor compartment. Importantly, functional validation confirmed that the edited population contained bona fide HSPCs: edited cells successfully rescued lethally irradiated mice, demonstrating the presence of long-term repopulating stem cells. Although some differentiation is expected with extended culture, the edited populations remained capable of lineage redirection, such as toward dendritic cells, consistent with the purpose of the protocol to modulate HSPC-derived cell fates. When lower cell numbers are sufficient, editing can also be performed earlier (e.g. days 1–3), when the Lin⁻ compartment is even more enriched (36), thereby minimising ex vivo differentiation.

The ultimate goal of this research was to establish a protocol for successful gene engineering of HSPCs for the improvement of immunotherapies for cancer. Our laboratory has previously shown that the addition of HSPCs to two different immunotherapies significantly improves survival in mice with aggressive gliomas (31–33). Here, we showed that IL-6R^KO HSPCs led to superior efficacy against preclinical gliomas. Our goal is to use edited HSPCs to alter immunoregulatory mechanisms and improve immunotherapy efficacy for brain tumors. In future studies, we plan to further test modified HSPCs in combination with immunotherapies (adoptive T-cell therapy and anti-PD-1 checkpoint blockade) to increase their efficacy in solid malignancies. This method can also be applied to other genes of interest for the development of additional therapeutic modalities, and we encourage researchers to use this detailed and easy-to-follow protocol to do so.

Lineage bias induced by HSPC editing poses potential risks, including aberrant myelopoiesis or exhaustion of long-term repopulating cells. These concerns underscore the need for controlled ex vivo expansion and in vivo lineage tracking in preclinical models prior to translation. The Cas9 mRNA-based approach presented here enables stable gene editing while limiting nuclease exposure, which reduces off-target and integration-related risks. However, because the edits are stable, thorough validation, such as long-term engraftment studies and clonal dynamics analyses, will be essential to ensure that lineage-skewing strategies do not compromise hematopoietic balance in future clinical applications.

This study aimed to establish a method for efficient gene knock-out of HSPCs; however, it excluded the introduction of donor sequences into the genome. In future studies, we aim to optimise a technique for the efficient knock-in of genes in HSPCs.

Table 3.

Murine myeloid panel for flow cytometry.

Antibody	Fluorophore
Lineage	FITC
CD45.1	AF700
GR-1	PE-Cy7
CD11c	PE-Dazzle
CD11b	PerCP-Cy5.5
F4/80	APC-Cy7
MHCII	BV421
IL-6R	APC
Blue Viability Dye	DAPI (BUV496)

Data availability statement

Data are available in repository Zenodo by accessing the following links. These materials include open CC-BY licenses: