CRISPR-Cas9–based gene editing as a proof-of-concept approach in an inborn error of immunity caused by a DCLRE1C variant
Tugce Duran, Mehmet Ali Karaselek, Burak Dagdelen, Serkan Kuccukturk, Sükrü Nail Guner, Sevgi Keles, Ismail Reisli
TL;DR
Researchers used CRISPR-Cas9 to correct a genetic mutation in immune cells, showing it's possible to fix the mutation but not fully restore normal function.
Contribution
This is the first proof-of-concept study using CRISPR-Cas9 to correct a DCLRE1C variant in CD4+ T cells.
Findings
CRISPR-Cas9 successfully restored the DCLRE1C gene to its wild-type sequence in CD4+ T cells.
Editing led to increased CD25 activation and Artemis protein expression, though mRNA levels remained unchanged.
The observed functional changes were statistically significant but did not reach levels seen in healthy controls.
Abstract
Hypomorphic DCLRE1C variants impair T and B cell development, leading to combined immunodeficiency (CID) or leaky severe combined immunodeficiency (SCID). Current treatment options, such as allogeneic hematopoietic stem cell transplantation (aHSCT), are associated with significant risks, highlighting the need for alternative therapeutic strategies. In this study, we report the first a proof-of-concept CRISPR-Cas9–mediated correction of a hypomorphic DCLRE1C variant (c.194 C > T; p.T65I) in CD4 + helper T (Th) cells using CRISPR-Cas9 gene-editing technology. CD4 + Th cells were isolated, and the variant region was edited with sgRNA and donor DNA. Gene editing efficiency was confirmed by Sanger sequencing, revealing successful restoration of the target region to its wild-type sequence. Functional analyses showed a significant increase in CD25 activation and Artemis protein expression…
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Taxonomy
TopicsImmunodeficiency and Autoimmune Disorders · CRISPR and Genetic Engineering · Genomics and Rare Diseases
Introduction
The DCLRE1C gene (DNA Cross Link Repair 1 C; 47.2 kb and consisting of 14 exons) encodes a protein called Artemis. Artemis is a complex protein with endonuclease and exonuclease activities and play a key role in V (D) J recombination, which is an important mechanism in immunoglobulin (Ig) and T cell receptor (TCR) diversity, as well as in the repair of double-strand DNA breaks [1, 2]. Some pathogenic variants of the DCLRE1C gene significantly impair T and B cell development, leading to severe combined immunodeficiency (SCID), while hypomorphic variants retain residual enzymatic activity and typically manifest as combined immunodeficiency (CID) or leaky SCID [3, 4]. Clinically, patients with hypomorphic DCLRE1C gene variants frequently present with recurrent respiratory infections, lymphoproliferation, autoimmunity, and malignancy, while lymphopenia and hypogammaglobulinemia represent the most common immunological features [5–7].
In the treatment of patients affected by pathogenic DCLRE1C gene variants, supportive therapies such as intravenous (IVIg) or subcutaneous (SCIg) immunoglobulin replacement therapy (IgRT) and antibiotic prophylaxis are commonly employed, while the only currently curative treatment option remains allogeneic hematopoietic stem cell transplantation (aHSCT) [5, 8–10]. However, aHSCT presents notable risks, such as difficulties in identifying a compatible donor, the possibility of graft rejection, and the occurrence of graft-versus-host disease (GvHD) [11, 12]. Additionally, mortality rates following aHSCT in patients with DCLRE1C deficiency have been reported to be generally high, although they vary depending on the timing of the treatment [13, 14]. Therefore, there is a clear unmet need for alternative therapeutic strategies, and recent studies provide strong evidence that gene therapy may offer a curative treatment option for individuals with inborn errors of immunity (IEIs) [15].
The first gene therapies were applied to SCID patients with ADA (Adenosine deaminase) and X-linked IL2RG (IL-2 receptor γ-subunit) variants using gamma-retroviral and lentiviral vectors [16, 17]. Moreover, there are studies suggesting that gene therapy using lentiviral vectors could be a viable approach for treating gene variants responsible for IEIs [18–20]. Although gene therapy methods utilizing retroviral and lentiviral vectors have dramatically advanced in recent years, the associated risk of oncogenesis remains an undeniable concern [21, 22]. In contrast, gene therapy based on the CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)-Cas9 (CRISPR-associated protein 9) system appears to be a promising approach for patients with IEI, offering a potential treatment option without the immunological complications observed in aHSCT [15, 23–25].
In our study, we present a proof-of-concept application of CRISPR-Cas9–mediated gene editing targeting a hypomorphic DCLRE1C variant in CD4⁺ helper T (Th) cells from a patient with IEI.
Materials and methods
Study design
The study was conducted in the Department of Pediatric Immunology and Allergy involved a female patient (two years-old) with the hypomorphic DCLRE1C c.194 C > T; p.T65I) gene variant who had not undergone aHSCT. A total of 10 mL peripheral blood was collected from patient into K3-EDTA tubes in the study and analyses were performed in triplicate. The study included two experimental conditions: a gene-edited group and a non-gene-edited group. The patient participating in the study and her parents were informed about the study and written informed consent from parents. This study was performed in line with the principles of the Declaration of Helsinki. Approval was granted by the Ethics Committee (approval number: 2024/5066).
CD4 + Th cell isolation and culture
Peripheral blood mononuclear cells (PBMCs) were isolated from the patient’s blood sample using the density-gradient centrifugation with Histopaque (Sigma-Aldrich, ABD). Subsequently, CD4 + Th cells were sorted using the human CD4 + T Cell Isolation Kit (Miltenyi Biotec., Germany) according to the manufacturer’s instructions, and the cell count was determined using the Countess 3 Automated Cell Counter (Thermo Scientific Invitrogen). The cells were then plated in culture at a concentration of 2 × 10^5^ cells/ml, and were cultured in RPMI 1640 medium [supplemented with 10% fetal bovine serum (FBS), sodium pyruvate, non-essential amino acids and 1% penicillin/streptomycin] and stimulated with 50 U/ml IL-2, CD2-Biotin (100 µg/ml), CD3-Biotin (100 µg/ml), and CD28-Biotin (100 µg/ml) for 72 h at 37 °C and 5% CO_2_. Uncultured cells were cryopreserved in FBS containing 10% dimethyl sulfoxide (DMSO) and stored at −80 °C until further use.
Single guide RNA (sgRNA) and donor DNA (dDNA) design and validation
The sgRNAs were designed using the online Benchling platform (benchling.com/crispr/). First, -NGG protospacer adjacent motif (PAM) sequences within the target region for gene editing were identified for CRISPR-Cas9–mediated gene editing. Candidate sgRNA sequences were evaluated in silico for predicted on-target efficiency and off-target specificity using established prediction algorithms implemented in both the Benchling platform and the CRISPOR tool, and sgRNAs with high predicted on-target efficiency and favorable off-target specificity profiles were selected for synthesis. Also, sgRNAs were chosen to target the hotspot mutation region of the DCLRE1C gene. The dDNA sequence was designed using a Benchling platform with a length of 200 bp. Two sgRNA (sgRNA1 and sgRNA2) were designed according to these criteria (Fig. 1A).
Fig. 1sgRNAs designed strategy and flowchart of study. (A) DCLRE1C gene structure, sgRNAs and donor template DNA design strategy. (B) Flowchart of study. B1. sgRNAs and dDNA design, B2. Transfer of plasmids to bacteria, multiplication and plasmid DNA extraction, B3. Transformation and obtaining recombinant plasmids, B4. Transfer to recombinant plasmid to bacteria, multiplication and plasmid DNA extraction, B5. Electroporation, B6. Th cell culture after electroporation, B7. Structure and functional analysis after gene modification. (sgRNA, single-guide RNA; dDNA, donor template DNA; Th, helper T cell)
Transformation
The transformation process involved the cloning of sgRNAs into CRISPR plasmid vector pX459 v2.0-eSpCas9) (Addgene). The plasmid vector obtained commercially, was resuspended in SOC (Super Optimal Broth with Catabolite Repression) medium and cultured in LB (Lysogeny broth). The plasmids were then transformed into DH5α Escherichia coli cells using the heat-shock transformation method.
Plasmid DNA was isolated at a concentration of 100 ng/µl using NucleoSpin^®^ Plasmid Mini Kit (Macherey-Nagel). The plasmid was cleaved using the BbsI restriction enzyme (New England Biolabs, England) to linearize the vector at the sgRNA scaffold region (1 µg pX459V2.0-eSpCas9(1.1) plasmid DNA, 7 µl 10x NEBuffer, 2 µl BbsI, total volume 50 µl, 37 °C 1 h).
Subsequently, each sgRNAs oligonucleotide pair was annealed to form an oligoduplex structure using T4 Polynucleotide Kinase (New England Biolabs, England) (37 °C for 30 min followed by 95 °C for 5 min). The resulting oligoduplex sgRNAs were ligated into linearized plasmids using T4 DNA Ligase (Promega, USA) (total volume 10 µl reaction components: 100 ng linear plasmid, 1 µl gRNA oligodupleks, 2 µl 10X T4 DNA Ligase buffer, 1 µl T4 DNA Ligase, 16 °C overnight).
The recombinant plasmid was transferred back to DH5α E. coli using the heat-shock method, and bacteria under ampicillin selection to generate a recombinant plasmid library. Plasmids were subsequently isolated from DH5α bacterial cells using the Wizard^®^ Plus SV Minipreps DNA Purification System kit according to the manufacturer’s protocol (Promega, USA). Finally, ligation was confirmed by using vector-specific primers (forward: 5’TTTGTGATGCTCGTCAGGGG3’; reverse: 5’CCAAGTGGGCAGTTTACCGT3’).
Transfection
Electroporation method was used for transfection. For this purpose, CD4 + Th cells previously in the culture medium were taken and centrifuged at 330 g for 6 min, and 1 × 10^5^ cell was transferred to Opti-MEM (Sigma-Aldrich, ABD). Electroporation was performed with 200 V, 2 m/s, 2 pulses using 1 × 10^5^ cell, 2 µg plasmid and 1 µg dDNA with Gene Pulser Xcell Electroporation Systems (Bio-Rad Laboratories, Inc.).
Flow cytometry
Flow cytometry was used to detect GFP-positive cells after transfection. Cell viability and apoptosis rates were determined using Apoptosis Detection Kit (BioLegend, BioLegend Inc., San Diego). Cell counts were performed with a Beckman Coulter (Beckman Coulter, USA) flow cytometer and the results were analyzed with Navios EX software (Beckman Coulter, USA).
Confocal microscope analysis
Live and dead Th cells were stained (Calsein, CYTOX) using a commercial kit (for mammalian cells). Images were acquired and analyzed with a Nikon A1R1 confocal microscope system (Nikon, Japan).
DNA sequence analysis
The edited DCLRE1C region was amplified using specific primers (forward: 5’-TTCCACCCCCACCTTGCT-3’; reverse: 5’-AGGCCGGCGGATGGATC-3’) and, the PCR reaction was carried out in a MiniAmp Plus Thermal Cycler (Thermo Fisher Scientific, ABD) with the following conditions: initial denaturation of 94 °C for 6 min followed by 32 cycles of 94 °C for 30 s, annealing beginning at 65 °C for 29 s and 72 °C 30 s. A final extension of 72 °C for 10 min. Amplified PCR products were purified using QIAquick PCR Purification Kit (Qiagen, Germany) and were performed to Sanger sequencing according to the manufacturer’s instructions. The obtained sequence results were analyzed with the FinchTV software (version 1.4.0, Geospiza Inc., USA) program.
qPCR and melting curve analysis
Cells were placed in TRIzol (ThermoFisher, USA) for RNA extraction, which was performed using an QIAwave RNA Mini Kit (Qiagen, Germany). cDNA synthesis was carried out according to the protocol provided with the cDNA synthesis Kit (Qiagen, Germany). To assess Artemis expression, qPCR was performed on the LightCycler^®^ 96 system (Roche, Germany) using specific primers for DCLRE1C (forward: 5’-GTTGGAGTGCAGACACACCT-3’; reverse: 5’-CTCCAACAGTGGTCCCTCAC-3’), cDNA, and 2X SYBR Green Master Mix (Hibrigen, Turkey). The β-actin gene served as the reference for normalization, and the 2^−ΔΔCT^ method described by Livak was applied to determine relative gene expression levels (Livak and Schmittgen 2001) [26].
Melting curve analysis was conducted immediately following the amplification protocol under the following conditions with QuantStudio 3 qPCR system (Thermo Fisher Scientific Inc., Waltham, MA, USA): no hold time at 95 °C, 15 s at 60 °C, and no hold time at 95 °C. The temperature change rate was set at 20 °C/s, except during the final step, where it was reduced to 0.1 °C/s. The resulting melt peak indicated the presence of the specific amplified product. Melting analysis was performed with one primer pair covering the target region and two primer pairs that did not (Table S1). The crossing point (Cp) was determined as the maximum value of the second derivative of the fluorescence curve.
CD25 activation analysis
Cells cultured after transfection were incubated with IL-2 and T cell activation kits (Miltenyi Biotec. Germany) for 72 h at 37 °C in 5% CO_2_. After the incubation, CD25 expression percentage was compared to the basal CD25 expression. The cells were stained with CD3 (APC), CD4 (PE), and CD25 (FITCH) mAb according to the surface staining protocol. Samples were analyzed with Beckman Coulter (Beckman Coulter, USA) and the results were analyzed with Navios EX software (Beckman Coulter, USA). CD25 activation was determined by comparing the percentage of CD25 after activation to the baseline.
In-silico analysis
The degree of stabilization of Artemis protein resulting from hypomorphic variant in DCLRE1C gene was determined. Firstly, the tertiary structure of wild type Artemis protein was obtained from Protein Data Bank (6WO0) and, the degree of stabilization of the protein was determined as PremPs tool [27].
Results
Modification of DCLRE1C gene variant with CRISPR-Cas9
Using the CRISPR-Cas9 system, we established a gene-editing approach targeting the c.194 C > T region of the DCLRE1C gene in human Th cells. This approach involved the design of two 20-nucleotide sgRNAs flanking the variant site. The PAM sequences were 2 and 4 nucleotides away from the target region for the sgRNAs, respectively, and the predicted Cas9 cleavage site for both sgRNAs coincided with the position of the variant. In silico analyses of the designed sgRNAs revealed high predicted on-target efficiency and favorable off-target specificity, as summarized in Table 1.
Table 1. In silico on-target and off-target efficacy results of sgRNAssgRNAOrientationSequence (5’–3’)PAMOn-target scoreSpecificity score (CFD)Predicted off-target profilesgRNA1ForwardATACTGTTCACCTGTGATTAAGG7483Predominantly intergenic/intronicsgRNA1ReverseAACAACAACTCCTTAATCACAGG7477Predominantly intergenic/intronic
Flow cytometry
Transfection efficiency and post-transfection cell viability were assessed by flow cytometry. Following electroporation, the mean percentage of GFP-positive cells and cell viability were 15%, 64.99% respectively (Fig. 2B).
Fig. 2. Result of confocal microscopy and flow cytometry. (A) Confocal microscopy result. (B) Determination of GFP-positive cell with flow cytometry after transfection. (C) Determination of cell viability and apoptotic with flow cytometry after transfection
Confocal microscope analysis
The effect of the electroporation on cell viability was assessed using Calcein and Cytox dyes by confocal microscopy. Calcein staining was used to identify viable cells, whereas Cytox staining was used non-viable cells. As shown in Fig. 2A, the electroporated groups exhibited predominant green fluorescence (Calcein), with no red fluorescence (Cytox), indicating that the cell viability was preserved following electroporation. Similary, in the non-electroporated control group, green fluorescence predominated, and no Cytox-positive cells were observed. These findings demonstrate that the electroporation process did not result in overt cytotoxicity and was compatible with short-term cell viability under the experimental conditions used.
DNA sequence and melting curve analysis
Sanger sequencing results demonstrate that the CRISPR-Cas9 system successfully targeted the hypomorphic DCLRE1C variant. Genomic editing was confirmed by Sanger sequencing, which revealed restoration of the target region to the wild-type sequence (Fig. 3A).
Fig. 3DNA sequence analysis, qPCR and CD25 activation analysis results after CRISPR-Cas9 application. (A) Sanger sequence analysis results (upper: pre-treatment, lower: post-treatment). (B) Melting curve analysis. (C) DCLRE1C gene expression analysis by qPCR. (D) CD25 activation analysis results. (E) In-slico analysis (*: p < 0.05. arrow: arrow: Region where the DCLRE1C variant is located)
Melting curve analysis revealed a distinct melting temperature (Tm) peak in the edited DCLRE1C gene region (Fig. 3B upper panel). In contrast, no specific melting peak was observed in the unedited samples (Fig. 3B middle and lower panels). This shift in Tm supports the specificity of the targeted gene editing achieved using CRISPR-Cas9. Furthermore, there is no evidence of the amplification and editing process (Figure S1-S3).
qPCR analysis
The expression of DCLRE1C gene was evaluated via qPCR following CRISPR-Cas9 application. A comparison between controls and patient revealed no significant difference in DCLRE1C gene expression (1.41-fold; p = 0.128) (Fig. 3C).
CD25 activation test
To evaluate T-cell functionality responsiveness following CRISPR-Cas9–mediated modification of the DCLRE1C gene, a CD25 activation assay was performed. The analysis included three experimental groups: control, pre-treatment (un-edited), and post-treatment (gene-edited). In healthy controls, CD25 expression increased by 63.25-fold, 59.02-fold, and 85.4-fold in CD3+, CD4+, and CD8 + T cells, respectively. In the pre- and post-treatment groups, CD3+, CD4+, and CD8 + T cell activation showed increases of 1.4- and 8.85-fold, 1.72- and 6.57-fold, and 1.21-fold and 8.15-fold, respectively (Fig. 3D).
In-silico analysis of hypomorphic DCLRE1C variant
We determined the ΔΔG value to be 1.02 kcal/mol, indicating that the Artemis protein became destabilized because of the amino acid substitution in the post-variant protein structure. This alteration disrupted intramolecular hydrogen bonding patterns. In the wild-type structure, the Thr65 residue established a hydrogen bond with Gly118 (2.98 Å) and two distinct hydrogen bonds with Ser62 (2.931 Å and 3.064 Å). However, following the variant, the bond with Gly118 and one of the bonds with Ser62 were disrupted. Instead, the Ile65 residue formed a novel hydrogen bond with Cys64 (3.165 Å) in the mutated structure (Figure S4).
Discussion
Many hereditary diseases, including IEI, currently have only one treatment option: aHSCT. This method involves the transplantation of healthy hematopoietic stem/progenitor cells (HSPCs) from an allogeneic donor. However, its clinical use is constrained by the availability of suitable donors and the risk of life-threatening complications [28, 29]. Recently, autologous gene therapy, based on the ex vivo transduction of HSCs), has emerged as a promising alternative, with growing evidence supporting its potential application in patients with IEIs [15, 28, 30–33]. In our study, we present a proof-of-concept CRISPR-Cas9–mediated correction of a hypomorphic DCLRE1C variant in the CD4 + Th cells at the cellular level, highlighting the feasibility of this approach at the cellular level and its potential relevance for future translational studies.
For certain IEIs such as ADA deficiency and Wiskott-Aldrich Syndrome (WAS), gene therapy has emerged as an established therapeutic option; however, aHSCT remains the current standard of care, and each treatment approach has distinct advantages and limitations [33]. A major advantage of gene therapy is the avoid of graft-versus-host disease (GvHD), a serious complication associated with aHSCT [34]. In FDA-approved gene therapy for ADA deficiency, overall survival rate following aHSCT have been reported to be approximately 95%, with GvHD-free survival rate of around is 87%, whereas gene therapy achieves comparable survival outcomes without the associated risk of GvHD [34, 35]. In contrast, survival outcomes in patients with hypomorphic DCLRE1C variants are not well defined, although untreated DCLRE1C deficiency has been associated with an increased risk of autoimmunity and malignancy [7, 10]. In this context and given the risks associated with aHSCT, the conceptual development of gene therapy approaches for DCLRE1C deficiency warrants further investigation, particularly with respect to their potential to reduce disease-associated comorbidities, rather than as an immediately applicable therapeutic alternative.CRISPR-Cas9 technology offers conceptual and technical advantages over traditional gene therapy strategies that rely on lentiviral, retroviral, adenoviral, or adeno-associated viral (AAV) vectors. Unlike viral systems that rely on random genomic integration, CRISPR-Cas9 enables site-specific, targeted genome editing, thereby theoretically reducing the risk of insertional mutagenesis and associated oncogenesis [36–38]. In contrast to adenoviral and AAV vectors, which may elicit strong immune responses or induce neutralizing antibodies, CRISPR-Cas9 can be delivered using non-viral approaches, such as electroporation or lipid nanoparticles, depending on the experimental context, potentially reducing immunogenicity and improving safety profiles. Additionally, the transient expression or activity of CRISPR-Cas9 may limit prolonged nuclease exposure, thereby reducing likelihood of long-term off-target effects, unlike integrative viral vectors [39]. Among patients with IEIs, gene therapies are currently approved for ADA-SCID and X-linked SCID, whereas CRISPR-Cas9-based gene-editing therapies have yet entered routine clinical practice for IEIs [17, 31, 40]. Nevertheless, promising preclinical and early translational studies have been reported in conditions associated with mutations in CTLA-4, hereditary angioedema, WAS, RAG2, FoxP3, BTK, CD40L etc [18, 19, 24, 25, 41–43]. In our study, we provide the first evidence that the hypomorphic DCLRE1C variant can be corrected using CRISPR-Cas9 at the cellular level. In this context, the present study provides a proof-of-concept demonstration of CRISPR-Cas9–mediated correction of a hypomorphic DCLRE1C variant at the cellular level. Although gene editing in mature CD4⁺ T cells cannot reverse V(D)J recombination defects established during early lymphocyte development, this approach offers valuable insight into the feasibility of genomic correction and its potential downstream cellular effects.
Transfection represents a critical step in CRISPR-Cas9-based applications, particularly with respect to cell viability and delivery efficiency. In human Th cells, a GFP-positive cell rate of approximately 10–30% following electroporation is generally acceptable in experimental settings [44, 45]. Consistent with this range, a previous study employing CRISPR-Cas9-mediated gene editing in CD4 + Th cells reported a GFP+ cell proportion of 9.1% [25]. In another study focusing on BTK gene editing, electroporation of K562 cells at 255 V, 5 ms using a single pulse achieved an integration efficiency approximately 15%; however, K562 cells are known to be substantially more permissive to electroporation than primary CD4 + Th cells [24]. Similarly, in patients with FoxP3 variants, chemical transduction of HSCs resulted in a plasmid integration rate of 31%, reflecting the higher transducibility of stem cell populations compared with mature lymphocytes [18]. Conversely, in a study involving CRISPR-Cas9-mediated correction of the CD40L variant, high-current electroporation (3 pulses of 1400 V, 10-millisecond pulse width) of T cells yielded a low efficiency of approximately 3%, highlighting the technical challenges associated with gene delivery into primary T cells [19]. In the present study, the GFP+ cell rate following electroporation was 15.6%, accompanied by a cell viability of 64.99%, which falls within the reported experimental range for primary human CD4⁺ T cells. These findings indicate that the electroporation conditions used were technically feasible and comparable to those reported in the literature, and they further supported by confocal microscopy findings, which corroborated the observed transfection efficiency and cell viability. The verification of gene editing at both genomic and functional levels is essential for demonstrating the biological relevance of CRISPR-Cas9-mediated gene editing. In previous studies, the edited genomic regions have been validated using Sanger sequencing [24, 25]. In present study, DNA Sanger sequencing confirmed correction of the hypomorphic DCLRE1C variant at genomic level. In addition to genomic confirmation, it is also crucial to assess the downstream functional consequences of gene editing. In patients with hypomorphic DCLRE1C variant, major laboratory findings include residual enzymatic activity, T and B cell lymphopenia, hypogammaglobulinemia, and impaired CD25 upregulation following T-cell activation [4, 7, 10]. Previous studies have reported that at CD25 expression fails to increase upon activation in patients with hypomorphic DCLRE1C mutations [4, 7]. In our study, a significant increase in CD25 activation and Artemis protein expression was observed following CRISPR-Cas9-mediated gene editing compared with pre-editing levels. Although CD25 expression did not reach the levels observed in healthy controls, the approximately 6–8% increase observed after gene editing was statistically significant, supporting a partial functional response. Notably, there was no significant difference in DCLRE1C mRNA expression before and after gene editing. This finding is consistent with the report by Volk et al., who similarly observed no significant difference in DCLRE1C mRNA expression between patients and healthy controls [4], suggesting that functional changes may occur independently of detectable mRNA-level alterations. Melting curve analysis is a widely used method for evaluating the efficiency of targeted genome editing using CRISPR-Cas9 [46–48]. In this context, distinct differences in melting temperatures between edited and un-edited samples support the presence of targeted genomic modification, consistent with the sequencing-based findings in the present study.
Importantly, the findings of this study should be interpreted in the context of biological feasibility rather than therapeutic readiness. The successful CRISPR-Cas9–mediated correction of the DCLRE1C variant in mature T cells demonstrates that targeted gene editing is technically achievable at the cellular level. However, this proof-of-concept evidence does not indicate immediate clinical applicability. Therapeutic readiness would require efficient and safe gene correction in hematopoietic stem cells, durable immune reconstitution, and comprehensive preclinical safety assessments, including in vivo validation and long-term off-target evaluation.
Despite major advances in genome-editing technologies, achieving functional correction in primary immune cells remains highly context dependent. Although plasmid-based Cas9 delivery may induce innate immune activation and cellular stress in primary T cells, multiple studies using state-of-the-art editing platforms demonstrate that delivery modality alone does not overcome intrinsic biological constraints. In immunodeficiency settings involving somatic mosaicism, precise genome editing failed to increase wild-type allele frequency in patient-derived T cells due to limited proliferative capacity and clonal dominance of revertant populations [49]. Similarly, CRISPR-engineered human HSPC models of GATA2 deficiency revealed a marked intrinsic fitness disadvantage of mutant cells, resulting in consistent out competition by wild-type counterparts despite accurate editing [50]. In contrast, functional benefit in CXCR4-associated WHIM syndrome required additional selective pressures, such as hematopoietic conditioning, underscoring that genome editing alone is often insufficient for functional rescue [51]. Together, these findings indicate that the lack of functional rescue observed in this study reflects disease-specific biological limitations rather than solely methodological shortcomings, consistent with the current translational landscape of genome-editing approaches for immunodeficiency [52].
The main limitation of this study is its single-patient design and the restriction of gene editing experiments to peripheral CD4⁺ T cells. Therefore, definitive conclusions regarding therapeutic efficacy or clinical safety cannot be drawn. In addition, gene editing experiments were performed in mature peripheral CD4⁺ T cells rather than in developing lymphocytes or hematopoietic stem cells; therefore, this study does not address correction of V(D)J recombination defects occurring during early immune development. In addition, functional outcomes of gene editing in mature peripheral T cells are strongly influenced by cell-intrinsic biological constraints, including limited proliferative capacity and absence of selective advantage, irrespective of the delivery platform used. Recent studies employing state-of-the-art genome-editing strategies have similarly demonstrated that optimization of editing modalities alone is insufficient to achieve functional rescue in immunodeficiency contexts. Accordingly, the functional assays used in this study were not intended to demonstrate immune reconstitution but to assess feasibility and downstream biological responsiveness following genomic correction in mature peripheral CD4⁺ T cells. Consequently, gene editing of mature CD4⁺ T cells cannot reverse previously established V(D)J recombination events but serves to assess the feasibility of genomic correction and its potential downstream cellular effects. Accordingly, the findings should be interpreted as a proof-of-concept experience demonstrating feasibility and biological plausibility rather than clinical effectiveness. Further studies involving hematopoietic stem cells and in vivo models are required to evaluate the translational potential of CRISPR-Cas9–based gene editing for DCLRE1C-associated IEI.
Conclusion
In conclusion, this study presents a single-patient proof-of-concept experience demonstrating CRISPR-Cas9–mediated correction of a hypomorphic DCLRE1C variant in CD4⁺ helper T cells. Limited functional analyses, indicated measurable cellular changes following gene editing; however, these findings do not allow definitive conclusions regarding therapeutic efficacy or clinical applicability. Importantly, the present work provides a biological rationale and technical feasibility framework for future studies focusing on hematopoietic stem cells and in vivo models. Further investigations are required to determine the translational potential of CRISPR-Cas9–based gene editing approaches for DCLRE1C-associated IEI.
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