What this is
- Optimized protocols were developed for delivery in primary mouse hematopoietic cells.
- The study focused on OT-I CD8T cells, bone marrow-derived macrophages (BMDMs), and hematopoietic stem/progenitor cells (HSPCs).
- These protocols aimed to improve gene editing efficiency while maintaining cell viability and function.
Essence
- protocols were optimized for efficient delivery across primary mouse hematopoietic cells, achieving high viability and functionality. The MaxCyte ExPERT platform demonstrated robust gene editing capabilities in OT-I CD8T cells, BMDMs, and HSPCs.
Key takeaways
- protocols Exp T3 and Exp T4 achieved over 95% viability in OT-I T cells after CRISPR delivery. Both protocols supported high-efficiency mRNA and RNP delivery without compromising cell viability.
- BMDMs maintained 88.7–94.4% viability post-, with the HSC-6 protocol achieving over 95% RNP delivery efficiency. This demonstrates the platform's suitability for gene editing in sensitive myeloid cells.
- HSPCs showed over 90% viability and near-uniform RNP delivery with the HSC-6 protocol. This supports the potential for efficient gene editing in stem/progenitor cells, crucial for therapeutic applications.
Caveats
- Editing efficiency was inferred from protein loss rather than direct sequencing, which may not capture all off-target effects. Future studies should incorporate DNA sequencing for precise quantification.
- The absence of detectable eGFP expression in BMDMs and HSPCs suggests limitations in mRNA stability or translation, indicating a need for further optimization.
Definitions
- Electroporation: A technique that uses electrical pulses to introduce nucleic acids into cells, allowing for genetic modification without viral vectors.
- CRISPR-Cas9: A genome editing tool that allows for precise modifications in DNA, utilizing a guide RNA to target specific sequences.
Simplified
Introduction
Primary hematopoietic cells are central to both mechanistic discovery and the development of cell-based therapies, yet genetic manipulation of these cells remains constrained by limited cell viability post-editing and reliance on resource-intensive viral delivery methods (1, 2). Establishing scalable, electroporation-based genome editing strategies that preserve cellular function is therefore essential for advancing both basic and translational research. Key hematopoietic populations of interest include CD8+ T cells, macrophages, and hematopoietic stem/progenitor cells (HSPCs), which collectively underpin disease modeling, therapeutic engineering, and immune function.
CD8+ T cells play a central role in adaptive immunity by contributing to host defense and anti-tumor responses (3, 4). Harnessing these CD8+ T cells has become a cornerstone of modern cancer immunotherapy. Among the most promising strategies is adoptive cell therapy (ACT), which involves the isolation, ex vivo expansion and/or engineering, and reinfusion of autologous T cells with enhanced specificity and effector capacity. ACT, including chimeric antigen receptor (CAR) T cell therapies, harnesses genetically modified T cells to recognize and eliminate tumor cells (5–7). The first two CAR-T products approved by FDA were produced by gamma-retroviral (Yescarta) (8) or lentiviral (Kymria) (9) gene transduction where synthetic receptor constructs are packaged using double-stranded viral elements in mammalian cells, followed by concentration of the viral particles which can be used to effectively deliver the CAR encoding DNA into T cells (10). Similarly, viral constructs can be used to encode specific short hairpin RNA (shRNA) molecules to induce RNA interference, leading to mRNA degradation and translational repression (11). However, application of viral gene or shRNA transduction is limited by high manufacturing costs, complex production pipelines, and off-target editing safety concerns, including semi-random genomic integration and insertional mutagenesis (12, 13). These challenges present opportunities for optimization and improved scalability in both basic biological studies and the application of findings to improve immune cell therapies.
While T cells are central to cancer immunotherapy, macrophages are increasingly recognized as promising effector cells. Playing an important role in innate immune activation and phagocytic clearance (14, 15), macrophages also possess protective and pathogenic functions in bone marrow transplantation (16). Bone marrow-derived macrophages (BMDMs) from mice are widely used to study macrophage biology owing to well-established differentiation protocols that yield large cell numbers (17). However, significant technical challenges create difficulty in achieving effective gene disruption in BMDMs and hamper their applications. This difficulty stems from the heightened sensitivity to exogenous nucleic acids, including viral vectors, which can induce innate immune activation (18, 19).
Beyond differentiated myeloid cells like BMDMs, HSPCs represent another key hematopoietic population in which precise genetic manipulation is both highly desirable and technically challenging (20, 21). HSPCs serve as a valuable source for the generation of various myeloid lineage immune cells and as such broadly used in basic research. Clinically, HSPCs are central to many therapeutic strategies including autologous transplantation of genetically corrected cells for inherited disorders and allogeneic transplantation to reconstitute the immune system in patients with treatment-resistant malignancies (22), inborn errors of immunity (23), and inborn errors of metabolism (24).
Genetic engineering of primary cells has historically relied on viral overexpression, shRNA-mediated knockdown or nuclease-based platforms such as transcription activator-like effector nucleases (TALENs). These approaches are limited by scalability, genomic integration, and cell-type-specific toxicity (25–27). The CRISPR (clustered regularly interspaced short palindromic repeats)–Cas (CRISPR-associated) system has vastly improved the possibilities of targeted genetic engineering strategies (28, 29). This advance was enabled by the identification of its core functional components: the Cas9 nuclease (30), a CRISPR RNA (crRNA) that specifies target DNA recognition (31), and a trans-activating CRISPR RNA (tracrRNA) which can be combined with crRNA into a single guide RNA (sgRNA) to simplify delivery and editing workflow (32, 33). Compared with shRNA-based knockdown, CRISPR-mediated gene deletion has emerged as a powerful and precise approach for dissecting function of genes and the encoded proteins (34, 35). The recent clinical approval of Casgevy, which uses CRISPR–Cas9 editing of human HSPCs by using MaxCyte ExPERT platform, demonstrates that scalable RNP delivery systems are clinically feasible (36). Consistent with this, electroporation-based gene editing in human macrophages has shown strong translational promise (37). However, adopting human cell engineering protocols to primary mouse immune cells remains challenging, limiting cross-species mechanistic and preclinical studies. Moreover, despite the molecular precision of CRISPR, viral delivery in sensitive primary cells is often constrained by limited payload capacity and prolonged nuclease expression, which can increase the risk of off-target editing. Together, these limitations underscore the need for optimized, electroporation-based delivery strategies that preserve cellular viability and function.
To address these limitations, non-viral gene delivery methods such as electroporation (EP) have emerged as promising alternatives for introducing CRISPR-Cas9 ribonucleoproteins (RNPs) and mRNA into primary cells. EP introduces editing machinery transiently, avoiding genomic integration and unintended off-target effects, and is compatible with good manufacturing practice (GMP) standards, making it an attractive platform for clinical translation (38, 39). Although the MaxCyte ExPERT platform offers a scalable and GMP-compatible electroporation system, systematically optimized protocols for primary mouse hematopoietic cell types have not been established. Primary cells are sensitive to alterations in culture media components such as serum, cytokines, growth factors, and editing efficiency is strongly influenced by cellular activation/differentiation states (40). Effective gene editing therefore requires careful optimization of EP timing and parameters to balance high editing efficiency and cell viability, and preservation of the cell’s original and the capacity for differentiation into effector T cells and macrophages.
In this study, we optimized MaxCyte ExPERT protocols for CRISPR-Cas9 RNP-mediated gene editing across multiple primary mouse cell types, OT-I transgenic CD8+ T cells, BMDMs and HSPCs. In CD8+ T cells, we evaluated and compared two protocols, originally designed for activated mouse T cells, and further demonstrated combinational workflows in which retroviral transduction can be applied post-EP to enable introduction of biosensors or genetic rescue constructs. For BMDMs and HSPCs, we evaluated multiple pre-programmed EP conditions and identified high-performing programs that support robust RNP delivery and efficient gene knockout. Across all cell types, these optimized workflows achieved high electroporation efficiency, cell viability, and reproducibility, with minimal disruption to proliferation, phenotype or the ability to differentiate. Together, these findings demonstrate that MaxCyte ExPERT electroporation is a reproducible, scalable, and functionally robust platform for genome editing in primary mouse hematopoietic lineage cells.
Materials and methods
Detailed procedures
C57BL/6-Tg(TcraTcrb)1100Mjb/J (OT-I) mice were used for primary CD8+ T cell activation experiments, with spleen and lymph nodes harvested for cell isolation. Primary bone marrow–derived monocytes/macrophages (BMDMs) and hematopoietic stem/progenitor cells (HSPCs) were isolated from femurs and tibias of C57BL/6J wild-type (WT) mice. All founder mice were obtained from The Jackson Laboratory and bred at the Centre for Molecular Medicine and Therapeutics (CMMT) Animal Facility in accordance with institutional guidelines (approval protocols #A23–0194 and A23-0282). A list of reagents, consumables, and equipment used throughout this protocol is provided in Table 1.
| Reagents | Source | Identifier |
|---|---|---|
| Alt-RCRISPR-Cas9 tracrRNA, ATTO™ 550® | IDT | Cat# 1075928 |
| Alt-RCRISPR-Cas9 tracrRNA, ATTO™ 647® | IDT | Cat# 10007853 |
| Super Bright 702 Anti-F4/80; Clone BM8 | eBioscience | Cat# 67-4801-82 |
| PE-Cy5 Anti-B220; Clone RA3-6B2 | BioLegend | Cat# 103210 |
| PE-Cy5 Anti-CD3; Clone 17A2 | BioLegend | Cat# 100274 |
| PE-Cy5 Anti-CD5; Clone 53-7.3 | BioLegend | Cat# 100610 |
| PE-Cy5 Anti-CD8; Clone 53-6.7 | BioLegend | Cat# 100710 |
| PE-Cy5 Anti-Gr-1; Clone RB6-8C5 | BioLegend | Cat# 108410 |
| FITC Anti-c-Kit (CD117); Clone QA17A09 | BioLegend | Cat# 155104 |
| BV421 Anti-Sca-1; Clone D7 | BioLegend | Cat# 108128 |
| AlexaFlour 700 Anti-CD48; Clone HM48-1 | BioLegend | Cat# 103426 |
| PE Anti-CD150; Clone TC15-12F12.2 | BioLegend | Cat# 115904 |
| Cas9-NLS purified protein | UC Berkeley QB3 MacroLab | N/A |
| CleanCap EGFP mRNA | TriLink | Cat# L-7201 |
| DPBS (1X) | Corning | Cat# 21-031-CV |
| Easysep™ Mouse CD8T Cell Isolation Kit+ | STEMCELL Technologies | Cat# 19853 |
| EasySep™ Mouse Hematopoietic Progenitor Cell Isolation Kit | STEMCELL Technologies | Cat# 19856A |
| Fixable Viability Dye eFluor 780 | eBioscience | Cat# 65-0865-18 |
| HyClone HyPure Water, Cell Culture Grade | Cytiva | Cat# SH30529.LS |
| HyClone™ Water, Molecular Biology Grade | Cytiva | Cat# SH3053802 |
| InVivoMAb anti-mouse CD28 | BioXCell | Cat# BE0015-1 |
| InVivoMAb anti-mouse CD3 | BioXCell | Cat# BE0002 |
| MaxCyteElectroporation Buffer® | MaxCyte | Cat# EPB-1 |
| Recombinant human IL-2 | PeproTech | Cat# 200-02 |
| Recombinant murine SCF | Peprotech | Cat# 250-03 |
| Recombinant murine IL-3 | Peprotech | Cat# 213-13 |
| Recombinant murine IL-6 | Peprotech | Cat# 216-16 |
| RPMI 1640 | Corning | Cat# 10-040-CV |
| Iscove’s Modification of DMEM (IMDM) | Corning | Cat# CA45000-366 |
| Trizmahydrochloride solution (10X)® | Sigma | Cat# T2819-1L |
| Trypan Blue Stain | Invitrogen | Cat# 15250061 |
Section 1: procedures for mouse primary CD8T cells +
Day 0 mouse primary CD8T cell isolation and culture +
Timing: 3.5 hours
Day 1 or 2 CRISPR-Cas9 ribonucleoprotein and mRNA electroporation
Timing: 2 hours
NOTE: Electroporation can be conducted 1 or 2 days post-activation depending on the experimental needs. CD8+ T cells activated using this protocol do not divide before 36 hours (41).
| crRNAs | Sequence (5’-3’) | Cell type used |
|---|---|---|
| Nfe2l2 | TTGGGGCAGCACCTGCT | CD8T cells+ |
| Uqcrfs1 | CAGAACAGGTGGCTCCGAGG | CD8T cells+ |
| Xdh | ATCTTCTTTATTCAACCCGG | BMDMs |
| G6pdx | TGCCCGCTCACGACTCACAG | HSPCs |
| Rosa26 | ACTCCAGTCTTTCTAGAAGA | All |
| Type of Processing Assembly | Samples: | Buffer ctrl | RNP | mRNA |
|---|---|---|---|---|
| Loading agent | R-50×3 or R-50×8 Processing Assembly | |||
| Stock concentration | – | 22.2 µM | 1 µg/µL | |
| Volume | 5.0 µL | 4.5 µL | 5.0 µL | |
| Final concentration | – | 2.0 µM | 100 ng/µL | |
| CD8T cells+ | ||||
| Cell concentration | 1.1×10cells/mL8 | |||
| Volume | 45.0 µL | 45.5 µL | 45.0 µL | |
| Cell number | 5.0×106 | |||
| BMDMs | ||||
| Cell concentration | 1.1×10cells/mL8 | |||
| Volume | 45.0 µL | 45.5 µL | 45.0 µL | |
| Cell number | 5.0×106 | |||
| HSPCs | ||||
| Cell concentration | 1.1×10cells/mL8 | |||
| Volume | 45.0 µL | 45.5 µL | 45.0 µL | |
| Cell number | 5.0×106 | |||
| Total volume | 50.0 µL | |||
| A | Parameter | Formula | Example |
|---|---|---|---|
| B | # of samples | N | 8 |
| C | Electroporation volume | V | 50.0 µL (0.050 mL) |
| D | Min. # of cells required | Total= (N + 2) × 1.0×10cells/mL × Vmin8 | Total= 5.0×10cellsmin7 |
| E | Total cells to collect | Total× 120%min | 6.0×10cells7 |
| F | Resuspension volume | (Total× 120%) ÷ 1.1×10cells/mLmin8 | 545 µL |
Day 2 or 3 (1 day post-electroporation)
Timing: 1.5 hours
Day 4–5
Continue expansion and culture of CD8+ T cells for functional or phenotyping experiments including Western blotting analysis (42). In this study, transient culturing in low-glucose media was used as an example application for phenotyping (43).
Section 2: procedures for mouse primary BMDMs
Day 0 mouse primary BMDMs isolation and electroporation
Day 1 post-electroporation
Section 3: procedures for mouse primary HSPCs
Day 0 mouse primary HSPCs isolation and culture
Timing: 3.5 hours
Day 1 HSPCs expansion
Timing: 30 minutes
1. Double the culture media volume by adding cIMDM.
Day 2 continue with HSPCs expansion
Timing: 1 hour
Day 3 eGFP mRNA and RNP electroporation
Timing: 2 hours
Day 4 (1 day post-electroporation)
Timing: 1.5 hours
1. Count and resuspend the electroporated HSPCs in cIMDM at 1.0×106 cells/mL in a 6-well non-TC treated plate.
NOTE: If positive control included, take 50,000 – 100,000 cells for flow cytometry as indicated in Section 1, Day 2 or 3, Step 3.
Day 5
The electroporated HSPCs can be used for downstream experiments, including colony assay and BM transplantation in irradiated mice (45, 46).
Results
Optimized electroporation of OT-I T cells enables high-efficiency mRNA and RNP delivery without compromising viability
When working with primary cells, factors such as cell viability and transfection efficiency are essential for establishing a reliable workflow for gene editing (47). OT-I TCR-transgenic (Tg) mice are widely used to investigate how CD8+ T cells respond to their cognate antigens (48). In this study, we used OT-I T cells that were 24 hours (Day 1) and 48 hours (Day 2) post-initial activation. Electroporation of OT-I T cells was performed using either the “Expanded T cell 3” (Exp T3, low energy) or “Expanded T cell 4” (Exp T4, high energy) protocol. As cargo, we delivered either buffer control, eGFP mRNA, or two independent RNP complexes (Figure 1a). Electroporation outcomes were evaluated by flow cytometric analysis of viability and efficiency. Cell viability was assessed 24 hours after the electroporation. Notably, when OT-I cells on Day 1 or Day 2 after activation were electroporated with either the Exp T3 or Exp T4 protocol, more than 95% of cells remained viable, as demonstrated in the representative dot plots (Figures 2a, b, Supplementary Figure 1a) and quantified in the corresponding bar graphs (Figures 2c, d). These findings indicate that neither protocol compromised T cell viability under the conditions tested. We next quantified the proportion of eGFP-expressing OT-I cells 24 hours after electroporation. Using either the Exp T3 or Exp T4 protocol, transfection efficiency exceeded 95% for both Day 1 and Day 2 OT-I cells, as shown by the representative histograms (Figure 2e) and summarized in the bar graphs (Figure 2f). Day 2 OT-I cells exhibited more than a twofold increase in eGFP geometric mean fluorescence intensity (gMFI) compared to Day 1 cells (Figure 2g). The elevated expression in Day 2 cells is consistent with the continuous increase in translational capacity during the activation of T cells, likely driven by increased mTORC1 signaling at 48 hours post-activation (49). Exp T3 and Exp T4 exhibited no statistically significant difference in % eGFP+, indicating equivalent mRNA delivery efficiency between the two protocols. Lastly, we assessed RNP delivery efficiency. Over 98% of OT-I Day 1 and Day 2 cells were positive for ATTO-550-labeled tracrRNA following electroporation with either RNP1 or RNP2, as shown in the histograms (Figure 2h) and bar graph quantification (Figure 2i). Together, these results demonstrate that both Exp T3 and Exp T4 protocols achieve robust mRNA and RNP delivery into primary OT-I T cells while preserving viability, establishing a reliable and reproducible platform for transient expression or gene-editing applications in primary mouse T cells.
Schematic illustration depicting mouse primary hematopoietic cells and electroporation procedures.Schematic illustration depicting OT-I T cell activation/expansionbone marrow-derived macrophages (BMDMs) isolationhematopoietic stem cells (HSPCs) isolation. Electroporation procedures are shown for each cell type. (a-c) (a) (b) (c)
Comparison of Expanded T cell 3 and Expanded T cell 4 electroporation programs demonstrates efficient cargo delivery and high viability in OT-I T cells 24 hr post-electroporation.100,000 OT-I T cells were harvested and assessed by flow cytometry 24 hr post-electroporation.Representative flow cytometry plots showing the cell viability following electroporation using Expanded T cell 3 (Exp T3) protocoland Expanded T cell 4 (Exp T4) protocol. Viability was measured using a fixable Live/Dead dye (eFlour 780).Quantification of the frequency of viable cells following electroporation using the Exp T3or Exp T4protocols.Day 1 or Day 2 activated OT-I T cells were electroporated with eGFP mRNA using either Exp T3 or Exp T4 protocols, and eGFP expression was measured 24 hr post-electroporation. Representative histograms show the distribution of eGFP intensity. Bar graphs summarize the frequency of eGFPcellsand the geometric mean fluorescence intensity (gMFI) of eGFPcells.Day 1 or Day 2 activated OT-I T cells were electroporated with either RNP1 or RNP2 using Exp T3 or Exp T4 protocols. Representative histograms show the ATTO-550-labeled tracrRNA expression, and bar graphs quantify the frequency of ATTO-550cells. (a-i) (a, b) (a) (b) (c, d) (c) (d) (e-g) (e) (f) (g) (h, i) (h) (i) + + +
Primary OT-I T cells retain growth capacity and show efficient gene-editing following electroporation
Following electroporation, we cultured OT-I T cells and monitored their expansion while confirming gene-editing efficiency by Western blotting (Figure 3a). Under the Exp T3 protocol, Day 1 cells displayed reduced early expansion but nonetheless increased cell number by approximately 3-fold over 72 hours (Figure 3b), whereas Day 2 cells expanded around 4-fold during the same period (Figure 3c). By the end of culture, fold expansion was only minimally different from buffer control T cells in both Day 1- and Day 2-electroporated groups (Figures 3b, c). Comparable results were observed with the Exp T4 protocol (Figures 3d, e); however, expansion of Day 2 cells was noticeably lower than that of Day 2 cells electroporated with Exp T3, suggesting that the expansion of more activated, proliferating T cells may be more sensitive to the higher-energy Exp T4 protocol.
OT-I cells electroporated on Day 1 were expanded for 3 days, after which cell pellets were collected and lysed for Western blot analysis. A protein normally upregulated by the gene-product targeted by RNP1 was efficiently knocked out using both the Exp T3 and Exp T4 protocols (Figure 3f, top blots). In contrast, the protein encoded by the gene targeted by RNP2 showed only a modest reduction in expression (Figure 3f, middle blots). Given that RNP2 electroporation efficiency exceeded 95% (Figures 2h, i), the limited knockout likely reflects suboptimal sgRNA performance rather than delivery inefficiency, highlighting the importance of testing multiple independent gRNA to ensure efficient gene editing and loss of protein expression. Similar patterns were observed in Day 2 OT-I cells, with robust knockout of the RNP1 target and minimal reduction of the RNP2 target across both electroporation protocols (Figure 3g), indicating that gene editing outcomes were consistent regardless of activation state at the time of electroporation. Together, these results show that electroporated OT-I T cells retain their doubling capacity across protocols and activation states at the time of electroporation, with only minor sensitivity of Day 2 cells to the higher-energy Exp T4 condition. Moreover, robust knockout of the RNP 1 target demonstrates that both protocols support efficient gene editing, while the limited effect of RNP 2 highlights sgRNA quality as the principal determinant of editing outcomes using these protocols. These data also highlight the need for an efficient, reproducible transfection method in order to avoid uncertainty with whether low knock-out efficiency is due to a target sequence issue or transfection issue.
Electroporated OT-I T cells retain proliferative capacity and support efficient gene knockout.Schematic of the experimental workflow used to quantify cell expansion following electroporation.Day 1or Day 2activated OT-I T cells were cultured for 72 hr following electroporation using the Expanded T cell 3 protocol. Relative proliferation was assessed by cell counting at 24 hr intervals.Day 1 or Day 2 activated OT-I T cells were cultured for 72 hr following electroporation using the Expanded T cell 4 protocol. Relative proliferation was assessed by cell counting at 24 hr intervals.Following electroporation, OT-I T cells were harvested and analyzed by immunoblotting for NQO1 and Risp expression. α-tubulin was used as a loading control. (a) (b, c) (b) (c) (d, e) (f, g)
Electroporation does not exacerbate ER stress–related mRNA expression in low-glucose CD8T cells +
Glucose availability is critical for N-linked protein glycosylation and limiting glucose can provoke ER stress. In addition, electroporation has also been reported to trigger ER stress (50). To assess whether reduced glucose and/or electroporation alter the nutrient-mediated ER stress response in CD8+ T cells, we measured mRNA expression of Atf4 and Ddit3 (encoding Chop), canonical markers of nutrient and ER stress, in CD8+ T cells under nutrient-restricted conditions (51). Both transcripts were consistently upregulated under low-glucose conditions (Supplementary Figure 2a, b). However, electroporation did not further increase Atf4 or Ddit3 mRNA expression, indicating that this potentially stress-inducing procedure does not exacerbate ER stress in primary CD8+ T cells. These findings demonstrate that while low-glucose CD8+ T cells mount a stress-responsive transcriptional program, electroporation does not amplify ER stress–related mRNA expression, underscoring the robustness of this workflow for functional studies under metabolic perturbation.
Sequential electroporation and retroviral transduction enable efficient co-expression of CRISPR-Cas9 RNP and a reporter construct in OT-I T cells
Gene knockout studies are frequently complemented by gene rescue experiments to validate the specific functional contribution of target genes. However, knock-in approaches often exhibit limited efficiency and typically require isolation of single integration clones for downstream analysis, posing a barrier for short-lived in vivo mouse T cell studies. To establish knockout-rescue workflow, we tested a sequential platform combining electroporation-mediated RNP delivery (knockout) with retroviral gene expression (rescue) in activated OT-I T cells. Day 1 activated OT-I T cells were electroporated using the Exp T3 protocol to deliver RNP complexes. Following a 24 hr recovery period, cells were retrovirally transduced on Day 2 by spin-fection with a GFP reporter construct and expanded for an additional 48 hr (Supplementary Figure 3a). Flow cytometric analysis revealed robust co-expression of GFP and ATTO-550–labeled tracrRNA, indicating successful sequential delivery of viral and non-viral cargos within the same cell population (Supplementary Figure 3b). The majority of GFP+ cells retained detectable ATTO-550–labeled tracrRNA fluorescence, demonstrating that electroporation does not impair subsequent retroviral transduction efficiency and that RNP delivery and editing can proceed alongside viral transduction and cell expansion. These findings establish a modular workflow in which rapid, non-viral genome editing can be coupled with optional viral delivery of reporter or rescue constructs, enabling flexible gene perturbation and functional complementation strategies in primary T cells while minimizing the downstream need for single clone isolation steps often used with knock-in strategies.
High-efficiency electroporation preserves viability and differentiation of mouse primary BMDMs
Given the robust gene-editing efficiency achieved in mouse primary OT-I T cells using the MaxCyte ExPERT platform, we next assessed its performance in additional primary cell types known to be exquisitely sensitive and difficult to genetically modify, including BMDMs and HSPCs (52, 53). Bone marrow was harvested from C57BL/6J mice and electroporated using pre-programmed protocols “Opt 0–2”, “HSC-4”, and “HSC-6”, followed by differentiation into BMDMs over 7 days (Figure 1b). Viability was assessed 24 and 48 hours after electroporation. Across buffer control (ctrl), eGFP mRNA, and RNP cargo conditions, BMDMs displayed 88.7–94.4% viability, as shown in representative dot plots (Figures 4a, b;Supplementary Figure 1b). Quantification confirmed no significant difference between electroporated groups and either non-electroporated or buffer-only ctrl at both time points (Figures 4c, d). These results demonstrate that BMDMs maintain high viability following electroporation, underscoring the suitability of this platform for gene-editing applications in difficult to genetically modify primary myeloid lineages.
To further evaluate the platform performance, we examined cargo delivery and gene-editing performance in BMDMs. eGFP expression was not detected following electroporation (Supplementary Figure 3a), which may reflect intrinsic RNase activity or reduced translational capacity in macrophages (54, 55). To block endogenous RNase activity, we electroporated BMDMs with eGFP mRNA in combination with an RNase inhibitor, which failed to induce eGFP expression, despite efficient expression of the same eGFP mRNA in T cells. To assess RNP delivery efficiency, we measured uptake of ATTO-550-labeled tracrRNA pre-complexed with XOR-targeting guide RNA and Cas9 protein by flow cytometry. Approximately 20% of BMDMs electroporated with the Opt 0–2 protocol was ATTO-550 positive at both 24 and 48 hours, whereas the HSC-4 and HSC-6 protocols yielded > 95% ATTO-550+ cells (Figures 4e, f). Among these, HSC-6 provided the most robust performance as indicated by the highest ATTO-550 signal. Consistent with the observed delivery efficiency, Western blot analysis confirmed that RNP electroporation with the HSC-6 protocol effectively reduced XOR protein expression in BMDMs (Figure 4g). The percentage of ATTO-550+ cells showed a strong negative correlation with optical density measured from Western blots assessing the expression of the XOR protein, suggesting that higher RNP delivery leads to more efficient gene knockout (Figure 4h). Importantly, electroporation with the HSC-6 protocol did not impair macrophage differentiation, as Day 7 BMDMs showed comparable F4/80 expression across all conditions (Supplementary Figure 3b). Together, these results identify HSC-6 as an optimal electroporation protocol for efficient and reproducible RNP delivery and gene editing in mouse primary BMDMs.
Optimized electroporation supports efficient RNP delivery and gene editing in primary BMDMs. Isolated bone marrow cells were electroporated with buffer control (ctrl), eGFP mRNA, or RNP using three electroporation protocols: Opt 0–2, HSC-4, and HSC-6.Representative flow cytometry dot plots showing the frequency of viable cells at 24 hrand 48 hrpost-electroporation.Quantification of viable cell frequencies in non-electroporated (No EP) and electroporated (EP) groups at 24 hrand 48 hrpost-electroporation.Representative histograms showing electroporation efficiency based on ATTO-550–labeled tracrRNA fluorescence at 24 hr and 48 hr post-electroporation.Quantification of ATTO-550cells at 24 hr and 48 hr post-electroporation.Following electroporation, bone marrow cells were differentiated into macrophages for 7 days as described in the Methods and analyzed by immunoblotting for XOR expression. β-actin was used as a loading control.Quantification of ATTO-550cells (48 hr post-EP) and correlation with XOR protein expression levels measured by Western blot optical density (O.D.). (a, b) (a) (b) (c, d) (c) (d) (e) (f) (g) (h) + +
Optimized electroporation supports high-efficiency gene editing in mouse primary HSPCs
Having confirmed efficient delivery in macrophages, we next tested HSPCs, a distinct BM-resident cell type used for therapeutic genome engineering (22). HSPCs were isolated from C57BL/6J bone marrow and expanded for 3 days prior to electroporation (56). Day 3 HSPCs were electroporated using the pre-programmed protocols “Opt 0–2”, “HSC-4”, and “HSC-6” (Figure 1c). Viability remained consistently above 90% at 24 hours post-electroporation and was similarly maintained at 48 hours across all cargo conditions, including buffer ctrl, eGFP mRNA, and G6PD-targeting RNP complex (Figures 5a-d). Similar to BMDMs, eGFP expression was not detected following electroporation in HSPCs with or without addition of RNAse inhibitors in the electroporation cargo (Supplementary Figure 4c). To assess RNP delivery efficiency, we measured uptake of ATTO-550-labeled tracrRNA pre-complexed with G6PD-targeting guide RNA and CAS9 protein by flow cytometry. Approximately 80% of HSPCs electroporated with Opt 0–2 or HSC-4 were ATTO-550+ at 24 hours, with a modest decrease of 5–10% observed at 48 hours post-electroporation (Figures 5e, f). By contrast, HSC-6 yielded near-uniform delivery, with ~99% ATTO-550+ cells maintained stably at both 24 and 48 hours (Figures 5e, f). Cells were expanded until day 5 and day 7, and pellets were collected for Western blot analysis. Consistent with the delivery results, HSC-6 demonstrated the most robust loss of G6PD protein expression, an effect that was preserved in day 7 cells (Figure 5g). The proportion of ATTO-550+ cells, exhibited a strong inverse correlation with the optical density quantification of Western blots assessing the expression of the G6PD protein, indicating that increased RNP delivery is associated with enhanced gene knockout efficiency (Figure 5h).
During extended in vitro culture, a proportion of HSPCs undergo spontaneous differentiation by Day 7 (57). To control for this effect and focus analysis on the stem/progenitor compartment, we compared the abundance of HSPCs without lineage markers in cultures with and without electroporation. No significant differences were observed in the frequencies of Lin– or cKit+ between non-electroporated (No EP) and electroporated (EP; HSC-6 protocol) conditions. Similarly, the frequency of LSK (Lin– Sca1+ cKit+) cells was also preserved, indicating that electroporation does not promote spontaneous HSC differentiation and preserves key stem-cell phenotypic markers (Supplementary Figure 4d). Together, these findings identify HSC-6 as the optimal protocol for efficient and durable RNP-mediated gene editing in primary mouse HSPCs.
Optimized electroporation achieves efficient gene editing in primary mouse HSPCs. HSPCs were electroporated with buffer control (ctrl), eGFP mRNA, or RNP using three electroporation protocols: Opt 0–2, HSC-4, and HSC-6.Representative flow cytometry dot plots showing the frequency of viable cells at 24 hrand 48 hrpost-electroporation.Quantification of viable cell frequencies in non-electroporated (No EP) and electroporated (EP) groups across the three protocols at 24 hrand 48 hrpost-electroporation.Representative histograms showing RNP delivery efficiency based on ATTO-550–labeled tracrRNA fluorescence at 24 hr and 48 hr post-electroporation.Quantification of ATTO-550cells in buffer ctrl and EP groups at 24 hr and 48 hr post-electroporation.Immunoblot analysis of HSPCs harvested on Days 5 and 7 post-electroporation assessing G6PD protein expression.Quantification of ATTO-550cells (48 hr post-EP) and correlation with G6PD protein expression levels measured by Western blot optical density (O.D.). (a, b) (a) (b) (c, d) (c) (d) (e) (f) (g) (h) + +
Discussion
Primary mouse hematopoietic cells are foundational to both mechanistic immunology and translational cell engineering, yet their genetic manipulation remains heavily dependent on viral delivery platforms that are costly, time-intensive, and biologically confounding due to genomic integration, innate sensing of exogenous nucleic acid, and prolonged transgene expression (58). Non-viral approaches provide a scalable alternative, but in sensitive primary cells, achieving efficient delivery without perturbing phenotype or function requires careful optimization. In this study, we address this bottleneck by optimizing electroporation workflows using the MaxCyte ExPERT platform across three distinct primary mouse hematopoietic lineages, CD8+ T cells, BMDMs, HSPCs, demonstrating robust delivery while preserving viability, differentiation capacity, and functionality. To our knowledge, this is the first study and comprehensive workflow for highly efficient gene editing across these primary mouse hematopoietic lineages.
In activated OT-I T cells, both the Exp T3 and Exp T4 programs supported high-efficiency mRNA and RNP delivery with minimal impact on viability, establishing electroporation as a reliable platform for transient expression and genome editing in primary T cells. Notably, Exp T3 and Exp T4 yielded equivalent delivery efficiency, but subtle differences in proliferative capacity emerged under the higher-energy Exp T4 condition. These findings suggest a practical balance between delivery energy and proliferative robustness that may depend on activation state rather than intrinsic delivery efficiency. Because starting cell density and viability were carefully controlled prior to electroporation, these differences are unlikely to reflect confounding input variability. From an operational perspective, Exp T3 therefore represents a suitable default when preservation of proliferative capacity is critical, whereas Exp T4 may be preferable for experiment-dependent applications requiring higher energy with acceptable reductions in cell growth.
Electroporation is frequently assumed to impose substantial physiological stress on primary cells (59), raising concern that it may confound downstream readouts, particularly in metabolic or stress-response studies central to the field of immunometabolism. Consistent with this concern, both nutrient limitation and electroporation have been reported to activate ER stress pathways in other systems (50, 51). Here, we demonstrate that although CD8+ T cells appropriately induce canonical stress-responsive transcripts such as Atf4 and Ddit3 under glucose-limited conditions, electroporation following our workflow does not further induce this response in glucose-replete cultures. These findings establish an important biological guardrail for applying electroporation-based workflows in studies focusing on stress pathways as mechanistic mediators or experimental readouts. Whether higher energy programs such as Exp T4 subtly engage additional stress pathways that could contribute to reduced proliferation remains an important area for future investigation.
In contrast to T cells, eGFP mRNA expression was not detectable in BMDMs and HSPCs under the tested conditions, despite robust RNP delivery and efficient gene knockout. Several non-mutually exclusive mechanisms may explain this lineage-specific difference. Myeloid cells are enriched for innate nucleic acid sensing pathways and RNase activity, which may limit stability or translation of exogenous mRNA (54, 55). However, co-electroporating with RNase inhibitor failed to induce the expression of eGFP in either BMDMs or HSPCs, while the same sample of eGFP mRNA induced robust expression which worked well in T cells. Translational capacity and expression kinetics may differ across lineages, and the 24–48-hour sampling window used here may not capture transient expression in these cell populations. The successful delivery and functional activity of RNP complexes indicate that cargo entry per se is not the limiting factor. Future optimization strategies may include incorporation of RNA stabilizing modifications (60) or alternative timing of readout, although such approaches fall beyond the scope of this study. Importantly, the absence of detectable eGFP signal does not imply that mRNA cannot be delivered into these cells. It may indicate that under the tested conditions, functional expression was not achieved at detectable levels.
Among the electroporation programs evaluated for BMDMs and HSPCs, HSC-6 consistently yielded near-uniform RNP uptake and robust loss of target protein expression while preserving viability and differentiation capacity. We speculate that HSC-6 performed the best amongst the chosen electroporation programs because it delivers the highest amount of energy. Because mouse cells are smaller than their human counterparts, higher electroporation energy is required to open pores in the cell membrane. Typically, empirical testing of several electroporation conditions is required to find the most efficient electroporation program for a given cell type. With the MaxCyte ExPERT platform, the guesswork of testing dozens of combinations of voltage, pulse width, and other factors is taken out in favor of pre-programmed protocols tailored to a specific cell type. In this case, although no protocols for BMDMs existed, we predicted that because BMDMs are similar to HSPCs, the electroporation protocols designed for human HSPCs (HSC-4, HSC-6) would be effective for mouse HSPCs and BMDMs.
It should be noted that in our experiments we use HSPCs as a source of gene-edited progenitors for downstream myeloid cell differentiation in vitro, rather than to preserve gene-edited HSPCs with high long-term repopulation capacity. Polyvinyl alcohol (PVA)-based culture systems are now considered maintaining mouse HSPCs function ex vivo (61). Future work should determine whether alternative isolation strategies to obtain enriched HSCs combined with PVA-based culture conditions can be incorporated into the MaxCyte ExPERT workflow without reducing editing performance, particularly for applications in which long-term engraftment of gene-edited HSCs is required.
Electroporation can provide a gene editing strategy that supports scalability and avoids unwanted genomic integration. The MaxCyte platform has been implemented in GMP-compliant clinical-scale electroporation using the ExPERT GTx flow electroporation system (62), suggesting that the workflows described here can be readily adapted for large-scale translational applications. In addition, the workflows established here provide a modular framework in which rapid non-viral genome editing can be coupled with optional downstream viral delivery for reporter or rescue constructs, enabling flexible sequential gene disruption and re-expression while minimizing viral exposure.
Several limitations should be acknowledged. We inferred editing efficiency through protein loss and tracrRNA uptake rather than direct locus-level sequencing, and future studies incorporating DNA sequencing will enable more precise quantification of on-target efficiency and account for potential off-target outcomes. Additional validation using extended functional assays, including T cell effector activity, macrophage functional responses, and myeloid lineage HSPC differentiation, could further strengthen translational relevance. Nonetheless, the strong preservation of viability, expansion capacity, and stress-response fidelity across lineages supports the robustness of the platform under the conditions examined in this study and open the door to functional studies downstream using primary mouse cells.
In summary, we establish the MaxCyte ExPERT electroporation as a scalable, non-viral platform for efficient CRISPR RNP delivery across multiple primary mouse hematopoietic cells while preserving viability and lineage integrity. In activated OT-I T cells, both Exp T3 and Exp T4 programs supported robust delivery, with subtle activation-state–dependent tradeoffs in expansion using the higher energy T4 electroporation protocol. In BMDMs and HSPCs, we identify HSC-6 as a high-performing program for consistent high-level RNP delivery and target gene knockout. Together, these optimized workflows lower the barrier to rapid, reproducible genome engineering in primary hematopoietic cells for both discovery and translational applications.
Funding Statement
The author(s) declared that financial support was received for this work and/or its publication. This work was supported by the following funding: a University of British Columbia 4-Year Fellowship (to JO), BCCHR Canucks for Kids Fund Childhood Diabetes Laboratory Master’s Studentship (to DW), BCCHR Jan Freidman Masters and Doctoral Studentships (to ET), Breakthrough T1D—formerly JDRF—postdoctoral fellowship (3-PDF-2024-1504-A-N to LM), Michael Smith Health Research BC Trainee award (RT-2022-2619 to A-SA), Banting postdoctoral fellowship (509775 to A-SA), Canadian Institutes for Health Research (DT4-179512 to RK), Natural Sciences and Engineering Research Council of Canada (RGPIN-2020-05390 to RK), Canadian Cancer Society (Early Scholar Award to RK), Michael Smith Health Research BC Scholar award (SCH-2022-2767 to RK).
Footnotes
Data availability statement
The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found in the article/. 1
Ethics statement
The animal study was approved by Centre for Molecular Medicine and Therapeutics (CMMT) (approval protocols #A23-0194 and #A23-0282). The study was conducted in accordance with the local legislation and institutional requirements.
Author contributions
RKG: Conceptualization, Formal analysis, Funding acquisition, Supervision, Writing – original draft, Writing – review & editing. JO: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Visualization, Writing – original draft, Writing – review & editing. LY: Conceptualization, Formal analysis, Investigation, Writing – original draft, Writing – review & editing, Methodology. ET: Conceptualization, Formal analysis, Investigation, Writing – original draft, Writing – review & editing. LdBM: Formal analysis, Investigation, Writing – original draft, Writing – review & editing, Methodology. DW: Formal analysis, Investigation, Writing – original draft, Writing – review & editing, Methodology. A-SA: Investigation, Writing – original draft, Writing – review & editing.
Conflict of interest
Author LY was employed by the company MaxCyte, Inc.
The remaining author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Generative AI statement
The author(s) declared that generative AI was not used in the creation of this manuscript.
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Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fimmu.2026.1820963/full#supplementary-material↗
References
Associated Data
Supplementary Materials
Data Availability Statement
The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found in the article/. 1