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Article

Precise CRISPR-Mediated Editing of the TGFBI R555W Mutation in Patient-Derived Peripheral Blood Mononuclear Cells

by
Burak Dagdelen
1,*,
Hilal Arikoglu
1,
Dudu Erkoc-Kaya
1 and
Banu Bozkurt
2
1
Department of Medical Biology, Faculty of Medicine, Selçuk University, 42130 Konya, Türkiye
2
Department of Ophthalmology, Faculty of Medicine, Selçuk University, 42130 Konya, Türkiye
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2026, 27(5), 2418; https://doi.org/10.3390/ijms27052418
Submission received: 2 February 2026 / Revised: 25 February 2026 / Accepted: 28 February 2026 / Published: 6 March 2026
(This article belongs to the Topic Advances in Gene Therapy of Human Diseases)
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Abstract

Over 70 mutations in the transforming growth factor beta-induced (TGFBI) gene are associated with corneal dystrophies that impair vision. The R555W hotspot mutation is a major cause of granular corneal dystrophy type 1 (GCD1). Here, we evaluated the technical feasibility of CRISPR/Cas9-mediated editing of the R555W mutation in peripheral blood mononuclear cells (PBMCs) obtained from a patient with GCD1. Three single guide RNAs (sgRNA1–3) and matched single-stranded oligodeoxynucleotide donors (ssODN1–3) were designed and co-transfected into PBMCs. Transfected cells were enriched by flow cytometric sorting, with GFP-positive cells representing approximately 2–4% of the total electroporated population. Editing outcomes were initially screened using high-resolution melting (HRM) analysis, and the sgRNA3–ssODN3 combination identified as the most promising candidate was subsequently validated by next-generation sequencing (NGS). Sequencing revealed a homology-directed repair efficiency of 98.2% among GFP-positive sorted cells, demonstrating efficient and precise genome editing within the enriched population. Because PBMCs are not disease-relevant corneal epithelial cells and only genomic endpoints were assessed, the clinical applicability of this study is limited and the work should be considered a technical proof-of-concept. This framework supports optimization of CRISPR-based strategies prior to studies in biologically relevant corneal models.

Graphical Abstract

1. Introduction

Corneal dystrophies are a group of non-inflammatory, genetically and epigenetically driven disorders characterized by progressive loss of corneal transparency and visual impairment, affecting approximately 0.09% of the population [1]. Different corneal dystrophies are caused by mutations in specific genes, including the transforming growth factor beta-induced (TGFBI) gene located on chromosome 5q31 [2]. The transforming growth factor-β-induced protein (TGFBIp), encoded by the TGFBI gene and predominantly expressed in corneal epithelial cells [3], is a corneal stromal matrix protein essential for establishing and maintaining corneal transparency [4].
Currently, more than 70 distinct mutations have been identified in the TGFBI gene [5]. These mutations lead to the accumulation of insoluble protein deposits in the cornea, ultimately resulting in visual impairment [6]. Among them, the R555W (c.1663C>T) hotspot mutation is most commonly associated with granular corneal dystrophy type 1 (GCD1) [7]. GCD1 is characterized by round, irregular, gray-white deposits that accumulate within various layers of the cornea [8,9]. Although the pathogenesis of GCD1 is not fully understood, TGFBI mutations are thought to cause the accumulation of mutant TGFBIp aggregates by impairing protein secretion or folding [10].
Several clinical approaches targeting the corneal epithelium, such as keratectomy and corneal transplantation, are currently available for GCD1. However, current therapeutic approaches remain insufficient to slow, halt, or eliminate aggregate formation [11]. Consequently, gene-based therapeutic strategies aimed at providing long-term or permanent solutions have gained increasing attention. According to the Human Gene Mutation Database, more than 50% of disease-associated variants are point mutations, also referred to as single-nucleotide changes [12]. In recent years, programmable nucleases have been increasingly applied as gene-editing tools to enable the repair or precise editing of disease-associated point mutations [13]. Among the programmable nucleases, the clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 system has become the most widely used powerful genome-editing platform due to its simplicity, versatility and high efficiency [14].
Cas9, guided by a single guide RNA (sgRNA), functions as an endonuclease that introduces double-stranded breaks (DSBs) at specific genomic loci [15]. The sgRNA contains a 20 nucleotide protospacer sequence complementary to the target DNA, while a 3′ protospacer-adjacent motif (PAM) is required for Cas9 binding. The commonly used Streptococcus pyogenes Cas9 (SpCas9) recognizes the 5′-NGG-3′ PAM sequence and introduces DSBs three base pairs upstream of the PAM site [16]. These DSBs can be repaired through homology-directed repair (HDR) using single-stranded oligodeoxynucleotides (ssODNs) as donor templates, enabling precise genome editing [17,18].
Previous studies have demonstrated the feasibility of CRISPR/Cas9-mediated targeting of TGFBI mutations and other corneal dystrophy-associated models in experimental systems [19]. However, challenges related to delivery efficiency, cell-type specificity, and long-term therapeutic efficacy remain significant barriers to clinical translation [20]. In this context, accessible primary cells such as peripheral blood mononuclear cells (PBMCs) offer a practical platform for evaluating the technical performance of genome-editing strategies in patient-derived cells [21]. Although PBMCs do not represent disease-relevant corneal cell types and do not recapitulate corneal pathology, they provide a useful platform for assessing editing efficiency and feasibility in primary human cells.
In this study, we evaluated the technical feasibility of CRISPR/Cas9-mediated editing of the TGFBI R555W mutation in PBMCs obtained from a patient with GCD1.

2. Results

2.1. Primary Culture of PBMCs from a GCD1 Patient

Peripheral blood mononuclear cells (PBMCs) were isolated from a granular corneal dystrophy type 1 (GCD1) patient carrying the heterozygous R555W (c.1663C>T) mutation in exon 12 of the TGFBI gene, which has been previously reported [22], as demonstrated by the Sanger electropherogram in Figure 1b. Following isolation, PBMCs were stimulated with PHA for 72 h and subsequently expanded under standard culture conditions until CRISPR/Cas9-mediated gene-editing experiments were performed. The cultures maintained high cell viability and normal morphology throughout the pre-transfection period, as confirmed by microscopic imaging (Figure 2a).

2.2. Ligation of sgRNAs into the pCAG-eCas9-GFP-U6-gRNA Plasmid

Three single guide RNAs (sgRNAs) and their corresponding single-stranded oligodeoxynucleotide donors (ssODNs) were designed to target the TGFBI R555W mutation. sgRNA1 and sgRNA2 targeted the sense strand, whereas sgRNA3 targeted the antisense strand of the TGFBI gene, as illustrated in Figure 3. All sgRNAs were successfully cloned into the scaffold region of the pCAG-eCas9-GFP-U6-gRNA plasmid. The cloning workflow, including BbsI digestion, ligation of sgRNAs, bacterial transformation, antibiotic selection, and Sanger sequencing verification, is summarized in Figure 4.

2.3. Electroporation of PBMCs

After three days of culture, PBMCs were collected and resuspended in electroporation buffer. For each reaction, 1 × 106 cells were mixed with 2 µg sgRNA-cloned plasmid DNA and 1 µg of the corresponding ssODN donor template, followed by electroporation using a single square-wave pulse (250 V, 5 ms) (Figure 2a,b). Plasmid delivery was confirmed by GFP expression 24 h post-electroporation using fluorescence microscopy (Figure 2c). GFP-positive cells were subsequently sorted by fluorescence-activated cell sorting (FACS). Transfection efficiencies were 3.7%, 2.7%, and 2.4% for the sgRNA1–ssODN1, sgRNA2–ssODN2, and sgRNA3–ssODN3 groups, respectively (Figure 2d). Sorted PBMCs were cultured under standard conditions and used for downstream genome-editing analyses.

2.4. Genome Editing Analyses

High-resolution melting (HRM) analysis, commonly used for the detection of single-nucleotide polymorphisms, has also been applied as a preliminary screening tool to identify nucleotide changes introduced by CRISPR/Cas9-mediated genome editing [23,24]. HRM analysis was performed on DNA amplicons obtained from sorted PBMCs to screen for genome-editing events. DNA from non-CRISPR-treated PBMCs displayed heteroduplex melting profiles, whereas CRISPR-treated samples showed homoduplex melting curves, consistent with sequence modification at the target locus (Figure 5). Among the tested sgRNA–ssODN combinations, the sgRNA3–ssODN3 group demonstrated a clearly distinguishable melting curve profile compared with both mock-treated cells and other sgRNA–ssODN groups, consistent with successful editing at the target locus. Based on this distinct HRM profile, the sgRNA3–ssODN3 combination was selected for subsequent next-generation sequencing (NGS) analysis to enable precise quantification of editing outcomes. The selection of sgRNA3–ssODN3 for NGS analysis was based on its clearly distinguishable HRM profile, which reflects altered DNA duplex stability consistent with successful HDR-mediated sequence correction.
NGS analysis confirmed CRISPR/Cas9-mediated editing of the c.1663C>T (R555W) mutation in the TGFBI gene, consistent with the HRM screening results. Detailed sequence-level editing outcomes are shown in Figure 6a. Analysis of sorted GFP-positive PBMCs (Figure 6b) demonstrated that the intended T>C substitution was the predominant editing outcome, accounting for 98.2% of sequencing reads, while the non-edited mutant allele and small deletion events were observed at frequencies of 1.2% and 0.6%, respectively.

3. Discussion

In this study, we investigated CRISPR/Cas9-mediated editing of the R555W hotspot mutation in the TGFBI gene in peripheral blood mononuclear cells (PBMCs) derived from a patient with granular corneal dystrophy type 1 (GCD1) as a technical feasibility study in patient-derived primary cells. Importantly, this work does not aim to compete with or replace studies performed in corneal cell types, but rather complements the existing literature by establishing a technically robust, patient-derived platform for evaluating precise editing of a clinically relevant TGFBI mutation.
Although PBMCs are not the primary disease-relevant corneal epithelial cells affected in GCD1, they provide an accessible patient-derived cell-based in vitro platform for rapid optimization and validation of CRISPR/Cas9 genome editing strategies. The use of PBMCs enables evaluation of editing efficiency, donor design, and workflow feasibility within the patient’s native genetic background before transitioning to more biologically relevant corneal cell models. Moreover, as technical validation typically requires multiple rounds of optimization, peripheral blood collection—being minimally invasive—minimizes patient burden and circumvents the need for more invasive tissue sampling procedures. Therefore, the primary objective of this study was technical validation of precise HDR-mediated editing rather than direct therapeutic modeling. This stepwise approach may facilitate subsequent translation to corneal epithelial or stem cell-based systems.
The substitution of arginine with tryptophan at position 555 of TGFBIp is responsible for the formation of deposits in the corneal stromal layer in GCD1. GCD1, which follows an autosomal dominant pattern of inheritance, is the most common form of corneal dystrophy, with patients presenting blurred vision, light sensitivity, photophobia, and progressive vision loss in later life [2,25]. Current clinical treatments for GCD1 target corneal epithelial cells to remove deposits; however, because corneal epithelial cells are continuously replenished from limbal stem cells located at the corneal periphery, these improvements are typically temporary [5,11]. Consequently, gene-based therapeutic approaches aimed at addressing the genetic defect underlying GCD1 have gained increasing scientific interest.
Gene therapy approaches for corneal dystrophies include gene replacement, gene silencing and genome editing [26]. Gene replacement involves delivering a functional wild-type gene to target cells, whereas gene silencing aims to inactivate a disease-causing gene. In contrast, genome editing enables precise, targeted modifications of gene sequences [27].
Gene replacement therapy has previously been applied for autosomal recessive retinal dystrophy [28]. For example, Leber’s congenital amaurosis type 2, caused by a loss-of-function mutation in the RPE65 gene, has been successfully treated with gene replacement therapy, with patients maintaining improved vision for at least three years [29,30,31]. However, gene replacement is not suitable for autosomal dominant diseases, such as TGFBI-associated corneal dystrophies, due to continuous production of mutant protein from the existing allele. Therefore, gene silencing or gene editing approaches aimed at repairing the mutant allele can be used as therapies for autosomal dominant corneal diseases.
CRISPR/Cas9-mediated genome editing relies on the induction of double-stranded breaks (DSBs) in the target DNA sequence, which are then repaired via homology-directed repair (HDR) using an exogenous donor DNA template [18]. In recent years, CRISPR/Cas9 technology has been increasingly applied in both in vivo and in vitro studies for the treatment of corneal dystrophies. Taketani et al. [19] achieved over 60% repair of the disease-causing R124H mutation in the TGFBI gene in primary corneal keratocytes derived from a GCD2 patient using CRISPR/Cas9-mediated HDR. Different CRISPR/Cas9 strategies are adopted depending on the nature of the genetic defect. Frameshift mutations in coding regions can lead to permanent gene silencing. In autosomal dominant disorders, allele-specific gene inactivation via non-homologous end joining (NHEJ) has been successfully applied as a CRISPR/Cas9-based strategy. In TGFBI-associated corneal dystrophies, selective silencing of the mutant allele without affecting the wild-type allele has been shown to alleviate the disease phenotype [32].
For instance, Kim et al. [33] introduced frameshift mutations in exon 4 of TGFBI in limbal stem cells, effectively knocking out the mutant allele through CRISPR/Cas9-mediated NHEJ. Similarly, Christie et al. [34] applied a mutation-independent, allele-specific strategy by designing sgRNAs targeting PAM sites unique to the mutant R124H allele, achieving efficient knockout in PBMCs from GCD2 patients. This study highlights that mutation-independent, allele-specific targeting provides a permanent and broadly applicable strategy for autosomal dominant diseases, particularly when multiple pathogenic mutations exist within a single gene, circumventing the need to target each mutation individually.
Since GCD1 is among the most common corneal dystrophies worldwide, and is also prevalent in our population, we aimed to develop a strategy to target the causative mutation. Using a mutation-dependent CRISPR/Cas9 approach, we successfully edited the R555W mutation in PBMCs from a patient with GCD1.
The design of the sgRNA is a critical factor affecting the success of CRISPR/Cas9 gene editing at the target site. To induce DSBs near the R555W mutation (c.1663C>T) in the TGFBI gene, we designed three different sgRNAs: sgRNA1 and sgRNA2 targeting the sense strand, and sgRNA3 targeting the antisense strand of DNA. sgRNA1, sgRNA2, and sgRNA3 were designed to induce DSBs 64 nt, 68 nt, and 6 nt away from the R555W mutation site, respectively. Cas9-mediated DSB generation is a major source of undesired off-target effects [35]. Therefore, off-target scores of the designed sgRNAs were calculated using the high-confidence Benchling CRISPR Tool, and sgRNAs with the highest on-target and lowest predicted off-target scores were selected for further experiments. The specificity of the designed sgRNAs was additionally assessed using NCBI BLAST (Basic Local Alignment Search Tool; National Center for Biotechnology Information, Bethesda, MD, USA; https://blast.ncbi.nlm.nih.gov/Blast.cgi; accessed on 15 June 2021), and no significant homology was detected outside the targeted TGFBI gene region. However, experimental validation of off-target activity was not performed in this study, and therefore off-target effects cannot be fully excluded.
Several factors can enhance HDR efficiency, such as using exogenous donor DNA as a repair template. Both Song et al. [36] and Li et al. [37] reported that longer homology arms in exogenous donor DNA improve HDR efficiency. Moreover, it is recommended to use single-stranded oligonucleotides rather than double-stranded oligonucleotides as exogenous donor DNA to prevent integration into the host genome [38]. Considering these factors, we designed three 200 nt-long ssODNs as repair templates. Each ssODN contains a thymine at nucleotide position 1663th to repair the R555W mutation in exon 12 of the TGFBI gene, and silent mutations were also introduced to prevent re-binding of the sgRNAs after editing.
In our study, we used PBMCs, which are well-suited for genome-editing research because they can be obtained non-invasively from peripheral blood and are easier to culture than many other primary cell types. However, efficient delivery of CRISPR/Cas9 components into PBMCs remains challenging due to their inherently low transfection capacity. The reduced CRISPR/Cas9 efficiency frequently observed in PBMCs is largely attributed to limited plasmid uptake and, in particular, the strong innate immune response of PBMCs against foreign nucleic acids [39]. Electroporation has been recommended as the most effective method to enhance transfection efficiency in PBMCs [40]. Therefore, we employed electroporation to achieve efficient delivery of CRISPR/Cas9 components and ssODN repair templates into PBMCs.
In our study, the sorting efficiency of PBMCs was higher than that reported in the CRISPR/Cas9 study on lymphocytes conducted by Johnston et al. [41], who used pCAG-eCas9-GFP-U6-gRNA plasmids.
High-resolution melting (HRM) analysis has been widely and successfully applied in CRISPR/Cas9 genome-editing studies. Amplicon size is one of the most critical factors affecting HRM sensitivity. In accordance with previous reports indicating that amplicons larger than 150 bp reduce HRM resolution [42,43], we designed primers that generated 143 bp amplicons encompassing the R555W mutation site of TGFBI gene. The GC content of the amplicon was 48%, which is within the optimal range for reliable HRM performance, as higher GC content (~65%) has been reported to compromise melting curve specificity and reproducibility [44]. Consistent with these design considerations, HRM analysis revealed a clearly distinguishable melting profile in PBMCs transfected with the sgRNA3–ssODN3 guided CRISPR/Cas9 system, consistent with sequence modification at the target locus.
We consider that this outcome is due to the DSB induced by sgRNA3 being much closer to the targeted R555W mutation site compared with the other sgRNAs. Specifically, the DSB induced by sgRNA3 is only 6 nucleotides away from the mutation, whereas the DSBs generated by sgRNA1 and sgRNA2 are 64 and 68 nucleotides away, respectively. Therefore, sgRNA3-guided Cas9 editing was the most effective. Consistent with this, Yang et al. [45] and Elliott et al. [46] reported that genome editing efficiency increases as the CRISPR/Cas9-induced DSB is positioned closer to the target site. This observation aligns with the HRM screening results, which identified sgRNA3–ssODN3 as the most promising candidate for subsequent NGS validation.
According to the next-generation sequencing (NGS) results of ssODN3/CRISPR-edited PBMCs, the HDR rate was remarkably high. We consider several factors that may explain this outcome. First, the most likely reason for the high HDR ratio is that genome-editing efficiency was evaluated using sorted PBMCs, which enriches the population of successfully transfected cells. Second, Bialk et al. [47] demonstrated that using a double-strand-cleaving nuclease, rather than a nickase, increases genome editing efficiency; in accordance with this, we also used a double-strand-cleaving Cas9 nuclease. Finally, the ssODNs used in our study contained silent blocking mutations, which prevent hybridization of the ssODNs with the sgRNAs, thereby protecting them from Cas9-mediated cleavage and preserving a larger proportion of ssODNs for HDR. These blocking mutations also protect the edited genomic region by preventing re-recognition and re-cutting by the sgRNAs.
The cornea is an ideal target for both in vivo and ex vivo gene therapy studies because it is avascular, immune-privileged, easily accessible, and compatible with well-established surgical and advanced imaging techniques [48]. Clinical CRISPR-based gene therapy approaches remain highly needed to repair mutations underlying ocular disorders across different layers of the eye. CRISPR-mediated genome engineering holds potential to contribute to the development of future personalized therapeutic strategies.
Gene-based therapeutics must be widely investigated to achieve effective clinical outcomes in corneal dystrophies. Successful therapeutic application of CRISPR requires the elimination of off-target activity and possible cytotoxic effects at the preclinical stage. Although the development and application of CRISPR/Cas systems continue to progress rapidly, this also underscores the need for appropriate regulatory oversight to ensure responsible clinical translation.
Despite the strengths of this study, several limitations should be acknowledged. First, genome-editing experiments were performed in PBMCs, which do not represent disease-relevant corneal cell types and do not recapitulate corneal-specific TGFBI expression or pathology. Accordingly, this study was designed as a technical feasibility study rather than a disease-model or therapeutic study. Second, editing efficiency was evaluated in GFP-positive, sorted PBMCs, resulting in a highly enriched cell population; therefore, the reported HDR efficiency does not reflect bulk editing efficiency or allele-specific outcomes. Third, experimental validation of off-target activity and allele-specific editing was not performed, and off-target assessments were limited to in silico predictions. Finally, additional control experiments, including wild-type donor cells or non-targeting constructs, were not included in the present study, which was designed as a technical proof-of-concept. These limitations should be considered when interpreting the results and underscore the need for future studies in disease-relevant corneal cell models with comprehensive safety assessments.
Beyond demonstrating efficient locus-specific editing, the ultimate goal of this work is to establish a technically optimized CRISPR/Cas9 workflow that can be applied to disease-relevant corneal epithelial or stem cell-based models. Our following studies will focus on transferring this strategy to corneal cell types, evaluating functional restoration of TGFBI expression, and performing comprehensive safety assessments, including off-target and phenotypic analyses, to support potential translational applications. The design principles identified here may further guide optimization in these disease-relevant models.

4. Materials and Methods

4.1. Ethics Statement and Patient Recruitment

Peripheral blood mononuclear cells (PBMCs) were obtained after written informed consent in accordance with the Declaration of Helsinki. The study was approved by the Ethics Committee of the Faculty of Medicine, Selçuk University (Approval No. 2021/327). Written informed consent was obtained from the participant for sample collection and publication of clinical and genetic data. Genome editing experiments were conducted using PBMCs obtained from a single female patient (n = 1) clinically diagnosed with granular corneal dystrophy type 1 (GCD1) following a detailed ophthalmological examination at the Department of Ophthalmology, Faculty of Medicine, Selcuk University. The heterozygous R555W mutation had been previously confirmed by Sanger sequencing in an earlier report (Figure 1).

4.2. Isolation and In Vitro Culture of Primary Human PBMCs

A peripheral blood sample (5 mL) was obtained from the GCD1 patient and immediately transferred into vials containing EDTA as an anticoagulant. PBMCs were isolated using Lymphopure™ (BioLegend, San Diego, CA, USA) according to the manufacturer’s instructions and washed twice with ice-cold phosphate-buffered saline (PBS). The cells were then resuspended in RPMI 1640 medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (all Gibco, Thermo Fisher Scientific, Waltham, MA, USA). For mitogenic stimulation, 1 μg/mL phytohemagglutinin (PHA; Gibco, Thermo Fisher Scientific, Waltham, MA, USA) was added to the culture, and the cells were incubated for three days at 37 °C in a humidified incubator with 5% CO2.

4.3. Design of sgRNAs and CRISPR/Cas9 Plasmid Construction

sgRNAs targeting the R555W mutation site of the human TGFBI gene were designed using the online platform Benchling (https://www.benchling.com/, accessed on 15 June 2021). On-target and off-target scores were calculated with Benchling’s high-confidence CRISPR tool, and the three sgRNAs with the highest on-target and lowest off-target scores were selected for use in this study. Detailed information on the sgRNAs is provided in Figure 3.
The sgRNA sequences were synthesized as forward and reverse oligonucleotides, flanked by the desired restriction sites and beginning with a guanine (G) nucleotide to enhance transcription efficiency, as summarized in Table 1.
For ligation of sgRNAs into the plasmid, the complementary forward and reverse oligonucleotides were first phosphorylated using T4 Polynucleotide Kinase (T4PNK; New England Biolabs, Ipswich, MA, USA), followed by annealing at 95 °C for 5 min. Annealing was performed by gradually decreasing the temperature from 95 °C to 25 °C at a rate of 0.1 °C/sec using a thermocycler. The annealed sgRNAs were then ligated into the linearized pCAG-eCas9-GFP-U6-gRNA vector (Addgene plasmid #79145), a human codon-optimized SpCas9 and chimeric guide RNA expression plasmid with a 2A-eGFP, using BbsI (New England Biolabs, Ipswich, MA, USA) and T4 DNA ligase (Promega Corporation, Madison, WI, USA). The resulting constructs were transformed into chemically competent Escherichia coli DH5α cells (New England Biolabs, Ipswich, MA, USA) by heat-shock.

4.4. Confirmation of sgRNA Ligation

Plasmid DNA was extracted from bacterial cultures using the Wizard® Plus SV Minipreps DNA Purification System (Promega Corporation, Madison, WI, USA) according to the manufacturer’s instructions. Ligation of sgRNA1, sgRNA2, and sgRNA3 into the pCAG-eCas9-GFP-U6-gRNA backbone was confirmed by PCR and Sanger sequencing.
The universal hU6 forward primer and the custom-designed Cmv reverse (Cmv-R) primer, flanking the sgRNA scaffold region of the vector, were used for PCR amplification. The Cmv-R primer was designed using Primer3 (Primer3 web interface, https://primer3.ut.ee/; accessed on 15 June 2021), and both primers (hU6-F and Cmv-R) were synthesized by Sentebiolab (Ankara, Turkey) as listed in Table 1.
Target regions were amplified using hU6-F and Cmv-R primers with 2× At Taq Master Mix (Vivantis Technologies, Selangor, Malaysia). PCR conditions were as follows: initial denaturation at 94 °C for 5 min; 35 cycles of 95 °C for 60 s, 52.4 °C for 60 s, and 72 °C for 60 s; followed by a final extension at 72 °C for 5 min. PCR amplicons were subjected to Sanger sequencing by Letgen Biotechnology (Ankara, Turkey). Sequence data were analyzed using SnapGene software (version 8.2.2; GSL Biotech LLC, Chicago, IL, USA) and displayed as electropherogram peaks.

4.5. Design of ssODNs

The 200 nucleotide ssODNs were used as donor templates to mediate the editing of the R555W (c.1663C>T) hotspot mutation in exon 12 of the human TGFBI gene. Each ssODN was designed in the sense orientation with 100 nucleotide left and right homology arms and contained no chemical modifications. In addition to the intended nucleotide substitution, silent mutations were introduced to disrupt the sgRNA recognition sequence and prevent Cas9 re-cutting following successful HDR. The ssODNs were designed using Benchling’s online software (https://www.benchling.com/, accessed on 15 June 2021) and synthesized by Integrated DNA Technologies (IDT; Coralville, IA, USA), as detailed in Table 2.

4.6. Transfection of PBMCs

Electroporation was used to transfect ssODNs and pCAG-eCas9-GFP-U6-gRNA plasmid constructs into PBMCs. PHA-stimulated PBMCs were harvested, centrifuged at 300× g for 7 min, and resuspended in Opti-MEM (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) at a concentration of 1 × 107 cells/mL. For each electroporation, 2 μg of plasmid and 1 μg of the corresponding ssODN were added to 100 μL of PBMC suspension. The mixture was transferred into 0.2 cm-gap cuvettes (Bio-Rad Laboratories, Hercules, CA, USA) and incubated on ice for 10 min. Cells were electroporated using a Gene Pulser Xcell Electroporator (Bio-Rad Laboratories, Hercules, CA, USA) with a single square-wave pulse at 250 V for 5 ms. Co-electroporations were performed for the following pairs: sgRNA1 + ssODN1, sgRNA2 + ssODN2, and sgRNA3 + ssODN3. For each sgRNA–ssODN combination, three independent electroporation experiments were performed (1 × 107 cells per electroporation). Following electroporation, cells from the same group were pooled prior to flow cytometric analysis. After electroporation, PBMCs were incubated in the cuvettes at room temperature for 10 min, followed by the addition of 1 mL pre-warmed RPMI 1640 medium supplemented with 10% FBS. Electroporated PBMCs were seeded into 12-well plates and incubated overnight at 37 °C with 5% CO2. Mock-electroporated PBMCs (electroporation without plasmid DNA or ssODN, water only) were processed in parallel and served as negative controls.

4.7. Flow Cytometry

Green fluorescent protein (GFP) expression in electroporated PBMCs was analyzed 24 h post-electroporation. Briefly, PBMCs were centrifuged at 300× g for 7 min, resuspended in 300 µL 1× HBSS (Gibco, Thermo Fisher Scientific, Waltham, MA, USA), and transferred to flow cytometry tubes (BD Pharmingen, San Diego, CA, USA). After a second centrifugation at 300× g for 7 min, cells were resuspended in 200 µL 2% FBS/HBSS and sorted using a fluorescence-activated cell sorting (FACS) instrument (FACS Aria III, BD Biosciences, San Jose, CA, USA) through the FITC channel to isolate GFP-positive cells. Sorted PBMCs were collected into tubes containing RPMI 1640 medium supplemented with 10% FBS, 1% penicillin/streptomycin, and 1 mg/mL PHA. GFP expression images were also captured using fluorescence microscopy (Olympus BX51 microscope, Olympus Corporation, Tokyo, Japan). Sorted PBMCs were seeded into 12-well plates and maintained under standard culture conditions (37 °C, 5% CO2) until further analyses.

4.8. Genomic DNA Extraction

Genomic DNA was extracted from PBMCs using the Wizard® Genomic DNA Purification System (Promega Corporation, Madison, WI, USA) following the manufacturer’s instructions. Extracted DNA was diluted with ddH2O to a final concentration of 100 ng/μL. Genome editing efficiency was evaluated using high-resolution melting (HRM) analyses and next-generation sequencing (NGS). For all analyses, DNA extracted from mock-electroporated PBMCs (electroporated without plasmid or ssODN) was used as a negative control.

4.9. HRM Analysis

Primers targeting the TGFBI c.1663C>T region (TGFBI_E12_F and TGFBI_E12_R; see Table 1) were designed using Primer3 (Primer3 web interface, https://primer3.ut.ee/; accessed on 15 June 2021) and synthesized by Sentebiolab (Ankara, Türkiye). HRM analysis was used as an initial screening method to identify editing-positive samples. HRM analysis was performed on a LightCycler® 480 II system (Roche Diagnostics GmbH, Mannheim, Germany) using FastStart Essential DNA Green Master (Roche Diagnostics GmbH, Mannheim, Germany). Each 20 μL reaction contained 2× Master Mix, 0.5 μM of each primer, and 100 ng of genomic DNA. The qPCR program consisted of an initial denaturation at 95 °C for 10 min, followed by 45 cycles of 95 °C for 10 s, 61 °C for 10 s, and 72 °C for 15 s. Melting analysis was conducted immediately after amplification, with an initial step at 95 °C for 10 s, 65 °C for 1 min, 97 °C for 1 s, followed by a temperature ramp from 65 °C to 95 °C at 0.1 °C/s. Peak areas under the derivative melting curves (−dF/dT) were measured using ImageJ software for descriptive, semi-quantitative evaluation as part of the HRM screening workflow. Because electroporated cells from independent experiments were pooled prior to analysis, HRM measurements were interpreted descriptively, and no inferential statistical tests were performed.

4.10. NGS Analysis

Genomic regions surrounding the target sites were PCR-amplified using specific primers as listed in Table 1. Each 20 μL reaction contained 10× Taq buffer, 0.5 μM of each primer, 2 mM dNTPs, 25 mM MgCl2, 0.1 U Taq DNA polymerase, 100 ng of template DNA, and nuclease-free water. PCR cycling conditions were as follows: initial denaturation at 94 °C for 5 min; 35 cycles of 94 °C for 30 s, 61 °C for 45 s, and 72 °C for 30 s; followed by a final extension at 72 °C for 5 min. PCR amplicons were then sequenced by Macrogen Europe B.V. (Amsterdam, The Netherlands) using NGS. Variant frequencies at the c.1663 position in the TGFBI gene were automatically calculated by Golden Helix GenomeBrowse (version 3.1.0; Golden Helix, Inc., Bozeman, MT, USA) based on the proportion of sequencing reads containing each nucleotide.

4.11. Statistical Analysis

Statistical analyses were limited to descriptive evaluation of HRM peak area measurements as part of the screening workflow. Because electroporated cells from independent experiments were pooled prior to analysis, each condition represents a single pooled sample (n = 1), and no inferential statistical tests were performed. Next-generation sequencing (NGS) data were analyzed using Golden Helix GenomeBrowse software, and editing efficiency was calculated as the proportion of sequencing reads carrying the intended nucleotide substitution relative to the total number of aligned reads (allele frequency analysis). Because the study was designed primarily as a technical proof-of-concept, no additional comparative or inferential statistical analyses were performed.

5. Conclusions

This study demonstrates the technical feasibility of CRISPR/Cas9-mediated editing of the TGFBI R555W mutation in patient-derived primary cells. The high editing efficiency observed with the sgRNA3–ssODN3 combination highlights the importance of guide RNA positioning and donor template design for precise genome editing. Although peripheral blood mononuclear cells (PBMCs) do not represent disease-relevant corneal cell types, this work provides a methodological framework for future studies in corneal epithelial cells, keratocytes, or limbal stem cells. Further investigations incorporating appropriate control experiments, comprehensive off-target analyses, and optimized delivery strategies will be required before considering translational applications. Overall, this study establishes a practical and reproducible strategy for CRISPR-mediated editing of TGFBI mutations and supports future translation to disease-relevant corneal models.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/ijms27052418/s1.

Author Contributions

Conceptualization, B.D. and H.A.; Methodology, B.D.; Investigation, B.D.; Formal analysis, B.D.; Resources, B.B.; Supervision, H.A.; Writing—original draft preparation, B.D.; Writing—review and editing, H.A. and D.E.-K.; Funding acquisition, H.A. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Selçuk University Scientific Research Projects Coordination Unit, grant number 19102037.

Institutional Review Board Statement

This research was conducted in accordance with the Declaration of Helsinki and approved by the Ethics Committee of the Faculty of Medicine, Selçuk University (Approval No. 2021/327, approval date on 9 June 2021).

Informed Consent Statement

Written informed consent was obtained from the participant for participation in the study and for publication of clinical images and genetic data.

Data Availability Statement

The raw sequencing data generated in this research are publicly available in the NCBI Sequence Read Archive (SRA) under BioProject accession number PRJNA1415318. The dataset includes targeted amplicon sequencing FASTQ files and contains no identifiable personal or whole-genome information.

Acknowledgments

The authors acknowledge the administrative support provided by the Selçuk University Scientific Research Projects Coordination Unit.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Clinical and genetic characterization of the patient with GCD1. (a) Anterior segment photograph showing multiple irregular gray-white corneal stromal deposits characteristic of granular corneal dystrophy type 1. (b) Sanger sequencing electropherogram demonstrating the heterozygous C/T genotype at the R555W locus. Colored peaks represent nucleotide signals (A = green; C = blue; G = black; T = red). Images were obtained with written informed consent.
Figure 1. Clinical and genetic characterization of the patient with GCD1. (a) Anterior segment photograph showing multiple irregular gray-white corneal stromal deposits characteristic of granular corneal dystrophy type 1. (b) Sanger sequencing electropherogram demonstrating the heterozygous C/T genotype at the R555W locus. Colored peaks represent nucleotide signals (A = green; C = blue; G = black; T = red). Images were obtained with written informed consent.
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Figure 2. Delivery of CRISPR/Cas9 components and enrichment of PBMCs. (a) Pre-electroporation preparation of PBMCs. Representative bright-field microscopy images showing PBMC morphology at 4×, 10×, 20×, and 40× magnification. The schematic also illustrates the CRISPR/Cas9 plasmid construct and ssODN donor used for transfection. (b) Electroporation conditions using a single square-wave pulse. (c) Fluorescence microscopy images showing GFP expression 24 h post-electroporation, confirming successful plasmid delivery. (d) Flow cytometric analysis and sorting of GFP-positive PBMCs, indicating transfection efficiencies for each sgRNA–ssODN combination.
Figure 2. Delivery of CRISPR/Cas9 components and enrichment of PBMCs. (a) Pre-electroporation preparation of PBMCs. Representative bright-field microscopy images showing PBMC morphology at 4×, 10×, 20×, and 40× magnification. The schematic also illustrates the CRISPR/Cas9 plasmid construct and ssODN donor used for transfection. (b) Electroporation conditions using a single square-wave pulse. (c) Fluorescence microscopy images showing GFP expression 24 h post-electroporation, confirming successful plasmid delivery. (d) Flow cytometric analysis and sorting of GFP-positive PBMCs, indicating transfection efficiencies for each sgRNA–ssODN combination.
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Figure 3. Design and genomic location of sgRNAs targeting exon 12 of the TGFBI gene. sgRNA1, sgRNA2, and sgRNA3 are color coded, and their corresponding PAM sequences are shown in matching colors. Predicted on-target and off-target scores are indicated for each guide (100 denotes the optimal score). The c.1663C>T nucleotide and the R555W mutation site are highlighted in dark and light blue, respectively.
Figure 3. Design and genomic location of sgRNAs targeting exon 12 of the TGFBI gene. sgRNA1, sgRNA2, and sgRNA3 are color coded, and their corresponding PAM sequences are shown in matching colors. Predicted on-target and off-target scores are indicated for each guide (100 denotes the optimal score). The c.1663C>T nucleotide and the R555W mutation site are highlighted in dark and light blue, respectively.
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Figure 4. Cloning workflow for sgRNA construction. (a) Digestion of the pCAG-eCas9-GFP-U6-gRNA vector with BbsI and ligation of sgRNA oligonucleotides. (b) Transformation of ligated plasmids into E. coli DH5α competent cells. (c) Selection of transformed colonies on ampicillin-containing LB agar plates. (d) Verification of sgRNA insertion by Sanger sequencing. In the Sanger sequencing chromatograms, the blue-highlighted region represents the successfully ligated sgRNA insert sequence, and the dashed vertical reference lines point to position indicators within the sequence display.
Figure 4. Cloning workflow for sgRNA construction. (a) Digestion of the pCAG-eCas9-GFP-U6-gRNA vector with BbsI and ligation of sgRNA oligonucleotides. (b) Transformation of ligated plasmids into E. coli DH5α competent cells. (c) Selection of transformed colonies on ampicillin-containing LB agar plates. (d) Verification of sgRNA insertion by Sanger sequencing. In the Sanger sequencing chromatograms, the blue-highlighted region represents the successfully ligated sgRNA insert sequence, and the dashed vertical reference lines point to position indicators within the sequence display.
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Figure 5. High-resolution melting (HRM) screening of CRISPR/Cas9-edited PBMCs. Derivative melting curves (−dF/dT, y-axis, versus temperature, x-axis) are shown for PBMCs transfected with sgRNA1–ssODN1, sgRNA2–ssODN2, and sgRNA3–ssODN3, each compared with mock-electroporated controls. HRM was used as an initial screening step to identify candidate editing events based on differences in melting curve profiles. For semi-quantitative evaluation, peak areas under the melting curves were measured using ImageJ software (version 1.54g; National Institutes of Health, Bethesda, MD, USA). Bar graphs represent descriptive peak area measurements obtained from pooled samples (n = 1 per condition) and are presented for descriptive comparison only. No inferential statistical tests were performed.
Figure 5. High-resolution melting (HRM) screening of CRISPR/Cas9-edited PBMCs. Derivative melting curves (−dF/dT, y-axis, versus temperature, x-axis) are shown for PBMCs transfected with sgRNA1–ssODN1, sgRNA2–ssODN2, and sgRNA3–ssODN3, each compared with mock-electroporated controls. HRM was used as an initial screening step to identify candidate editing events based on differences in melting curve profiles. For semi-quantitative evaluation, peak areas under the melting curves were measured using ImageJ software (version 1.54g; National Institutes of Health, Bethesda, MD, USA). Bar graphs represent descriptive peak area measurements obtained from pooled samples (n = 1 per condition) and are presented for descriptive comparison only. No inferential statistical tests were performed.
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Figure 6. Next-generation sequencing (NGS) analysis of CRISPR/Cas9-edited TGFBI locus. (a) Representative sequencing reads from GFP-positive PBMCs transfected with the sgRNA3–ssODN3 combination showing the targeted nucleotide substitution and ssODN-introduced guide-blocking silent mutations (b) Schematic overview of the sgRNA3 target region and detected editing outcomes at the c.1663 site. A total of 811,358 sequencing reads were obtained for this amplicon (see Supplementary Table S1).
Figure 6. Next-generation sequencing (NGS) analysis of CRISPR/Cas9-edited TGFBI locus. (a) Representative sequencing reads from GFP-positive PBMCs transfected with the sgRNA3–ssODN3 combination showing the targeted nucleotide substitution and ssODN-introduced guide-blocking silent mutations (b) Schematic overview of the sgRNA3 target region and detected editing outcomes at the c.1663 site. A total of 811,358 sequencing reads were obtained for this amplicon (see Supplementary Table S1).
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Table 1. sgRNA and PCR primers used in this study.
Table 1. sgRNA and PCR primers used in this study.
NameTargetSequences (5′ → 3′)Amplicon Size (bp)
sgRNA1F
sgRNA1R		-caccGACGGAGACCCTCAACCGGGA
aaacTCCCGGTTGAGGGTCTCCGTC		-
sgRNA2F
sgRNA2R		-caccGACTGACGGAGACCCTCAACC
aaacGGTTGAGGGTCTCCGTCAGTC		-
sgRNA3F
sgRNA3R		-caccGAGTCTGCTCCATTCTCTTGG
aaacCCAAGAGAATGGAGCAGACTC		-
hU6-F
Cmv-R		sgRNA Scaffold GAGGGCCTATTTCCCATGATT
GGATAACCGCAATGATACCCT		600 bp
TGFBI_E12_FTGFBICAACCGGGAAGGAGTCTACA143 bp
TGFBI_E12_RGCCCTGAGGGATCACTACTTT
Table 2. Sequences of the designed ssODNs.
Table 2. Sequences of the designed ssODNs.
ssODNsSequence (5′ → 3′)
ssODN1CATCCAGTCTGCAGGACTGACGGAGACCCTCAATAGGGAAGGAGTCTACACAGTCTTTGCTCCCACAAATGAAGCCTTCCGAGCCCTGCCACCAAGAGAACGGAGCAGACTCTTGGGTAAAGACCAACTTAAGTACACGTCTCCATTTTTCTAAAGTAGTGATCCCTCAGGGCCCCAGCAGCAAACAGTTGGCACATCAA
ssODN2CATCCAGTCTGCAGGACTGACGGAGACCCTCAACCGCGAAGGAGTCTACACAGTCTTTGCTCCCACAAATGAAGCCTTCCGAGCCCTGCCACCAAGAGAACGGAGCAGACTCTTGGGTAAAGACCAACTTAAGTACACGTCTCCATTTTTCTAAAGTAGTGATCCCTCAGGGCCCCAGCAGCAAACAGTTGGCACATCAA
ssODN3CATCCAGTCTGCAGGACTGACGGAGACCCTCAACCGGGAAGGAGTCTACACAGTCTTTGCTCCCACAAATGAAGCCTTCCGAGCCCTGCCACCAAGGGAACGGAGCAGGCTCTTGGGTAAAGACCAACTTAAGTACACGTCTCCATTTTTCTAAAGTAGTGATCCCTCAGGGCCCCAGCAGCAAACAGTTGGCACATCAA
The ssODN sequences were designed to edit the R555W (c.1663C>T) mutation, with the thymine nucleotide (blue) centrally located within each ssODN. Additional nucleotides introducing guide-blocking silent mutations are shown in red.
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Dagdelen, B.; Arikoglu, H.; Erkoc-Kaya, D.; Bozkurt, B. Precise CRISPR-Mediated Editing of the TGFBI R555W Mutation in Patient-Derived Peripheral Blood Mononuclear Cells. Int. J. Mol. Sci. 2026, 27, 2418. https://doi.org/10.3390/ijms27052418

AMA Style

Dagdelen B, Arikoglu H, Erkoc-Kaya D, Bozkurt B. Precise CRISPR-Mediated Editing of the TGFBI R555W Mutation in Patient-Derived Peripheral Blood Mononuclear Cells. International Journal of Molecular Sciences. 2026; 27(5):2418. https://doi.org/10.3390/ijms27052418

Chicago/Turabian Style

Dagdelen, Burak, Hilal Arikoglu, Dudu Erkoc-Kaya, and Banu Bozkurt. 2026. "Precise CRISPR-Mediated Editing of the TGFBI R555W Mutation in Patient-Derived Peripheral Blood Mononuclear Cells" International Journal of Molecular Sciences 27, no. 5: 2418. https://doi.org/10.3390/ijms27052418

APA Style

Dagdelen, B., Arikoglu, H., Erkoc-Kaya, D., & Bozkurt, B. (2026). Precise CRISPR-Mediated Editing of the TGFBI R555W Mutation in Patient-Derived Peripheral Blood Mononuclear Cells. International Journal of Molecular Sciences, 27(5), 2418. https://doi.org/10.3390/ijms27052418

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