Advancing gene editing therapeutics: Clinical trials and innovative delivery systems across diverse diseases.

Introduction

The evolution of CRISPR (clustered regularly interspaced short palindromic repeats) technology offers a remarkable story, transitioning from its origins as a bacterial defense system to its current role as a groundbreaking tool in genetic engineering. The first discovery of CRISPR sequences dates back to 1987, when Ishino and colleagues 1 observed repetitive DNA elements within the genome of Escherichia coli . 2 This discovery began efforts to understand CRISPR as part of a microbial immune system that defends against invading viruses and plasmids by adapting to these threats. 3 Over time, research efforts, including those by Jansen and his team, revealed the complexity of CRISPR systems by identifying genes linked to CRISPR loci and noting their widespread presence across different bacterial species. 4 The concept of horizontal gene transfer emerged as a key factor in understanding how these defense mechanisms spread and evolve, as CRISPR systems were shown to transfer between bacterial species. 5 Godde and Bickerton later contributed to the field by categorizing the various families of CRISPR-associated (Cas) genes, providing further insight into their evolutionary history ( Figure 1 ). 6 A breakthrough occurred in the early 2010s when Doudna and Charpentier uncovered the mechanism by which CRISPR-Cas9 could be repurposed for precise genome editing. 7 This discovery demonstrated that CRISPR-Cas9 could be programmed to target specific DNA sequences, transforming the system from a bacterial defense mechanism into a versatile genetic editing tool. Their work laid the foundation for wide applications of CRISPR in organisms ranging from plants to animals and humans. 8 Since then, the versatility of CRISPR has been expanded further through the development of different CRISPR-Cas variants, such as Cas12 and Cas13, each offering unique applications and capabilities. 9 The repurposing of CRISPR-Cas9 for gene editing revolutionized genetic research. Doudna and Charpentier demonstrated that this system could introduce precise breaks in DNA, 7 enabling gene disruption or modification in eukaryotic cells. 10 This was a significant leap forward from earlier gene editing techniques like zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs), which were more complex and less efficient. 11 ZFNs were the first programmable nucleases to demonstrate targeted genome editing capabilities in mammalian systems and represent a significant milestone in the development of gene editing technologies. 12 ZFNs are engineered proteins consisting of two functional components: a DNA-binding domain made up of an array of zinc-finger motifs and a DNA-cleaving domain derived from the FokI endonuclease. 13 Each zinc-finger motif typically recognizes a triplet of DNA bases, allowing for modular assembly of arrays capable of targeting specific genomic sequences with high precision. 14 The FokI nuclease domain functions as a dimer, meaning that two ZFN monomers must bind to adjacent DNA sequences to induce a site-specific double-stranded break (DSB). 15 , 16 Once the DSB is created, the cell’s intrinsic DNA repair pathways are activated, primarily non-homologous end joining (NHEJ) or, in the presence of a donor template, homology-directed repair (HDR). 17 , 18 NHEJ often results in insertions or deletions (indels) at the break site, leading to gene disruption or knockout, while HDR enables precise gene correction or targeted insertion of new genetic material. 19 These mechanisms have made ZFNs an attractive tool for applications ranging from functional gene studies to therapeutic genome modification. 19 ZFNs have been evaluated in several early-phase clinical trials, demonstrating proof of concept for in vivo and ex vivo gene editing in human patients. 20 , 21 , 22 , 23 Notably, ZFNs have been investigated for the disruption of the CCR5 gene in T cells, aiming to confer resistance to HIV infection by mimicking the naturally occurring CCR5- Δ 32 mutation that provides protection against viral entry. 24 , 25 , 26 , 27 , 28 In addition, ZFNs have been explored for treating monogenic disorders such as hemophilia B, mucopolysaccharidosis (MPS), and other lysosomal storage diseases by enabling site-specific gene correction or transgene insertion in hepatocytes. 29 , 30 , 31 Despite these advances, the widespread adoption of ZFNs has been limited by several challenges. Designing highly specific zinc-finger arrays requires extensive protein engineering expertise, as each zinc finger interacts not only with its target triplet but also with neighboring motifs, complicating the development of truly modular systems. 32 , 33 Furthermore, concerns regarding off-target cleavage events, cytotoxicity, and variable editing efficiency have led to the pursuit of alternative, more accessible gene editing platforms. 34 Nevertheless, the development and clinical testing of ZFNs provided foundational knowledge that has informed subsequent genome editing technologies, including TALENs and CRISPR-based systems, while remaining a viable option for therapeutic applications where delivery constraints and regulatory considerations favor their use. 35 Figure 1 Evolution of CRISPR technology A timeline illustrating key milestones in the development of CRISPR, from the discovery of repetitive DNA sequences to genome editing breakthroughs, variant development, and ongoing ethical and regulatory considerations.

Base editors represent a groundbreaking advancement in genome editing technologies, offering the ability to introduce precise single-nucleotide substitutions without inducing DSBs or relying on donor DNA templates. 36 Developed as an alternative to conventional nuclease-based approaches, base editors enable highly efficient and predictable correction of point mutations, which account for approximately 60% of known pathogenic human genetic variants. 37 The core structure of a base editor typically combines a catalytically impaired or “nickase” version of Cas9 or Cas12 (which retains DNA-targeting capability but lacks complete nuclease activity) with a deaminase enzyme that catalyzes specific nucleotide conversions at the target site. 38 There are two primary classes of base editors. Cytosine base editor systems convert a cytosine (C) to a thymine (T) (or guanine to adenine on the complementary strand) by deamination of cytosine to uracil, followed by cellular repair mechanisms that interpret the uracil as thymine during DNA replication or repair. 39 This approach enables targeted C•G to T•A transitions. Adenine base editors (ABEs) facilitate the conversion of adenine (A) to guanine (G) (or thymine to cytosine on the complementary strand) through enzymatic deamination of adenine to inosine, which is read as guanine by the cell’s replication machinery, enabling A•T to G•C transitions. 39 , 40 , 41 These precision editing tools have several advantages over traditional nuclease-based systems. By avoiding the induction of DSBs, base editors significantly reduce the risk of large-scale genomic rearrangements, unintended indels, and activation of p53-mediated DNA damage responses, which are common concerns with standard CRISPR-Cas9 editing. 36 , 41 , 42 , 43 Furthermore, base editing achieves higher editing efficiency in non-dividing or slowly dividing cells, broadening its therapeutic potential for tissues such as the nervous system, retina, or muscle. 43 The therapeutic application of base editors is rapidly advancing, with preclinical studies demonstrating successful correction of pathogenic point mutations associated with a variety of genetic disorders, including Duchenne muscular dystrophy (DMD), β-thalassemia, sickle cell disease (SCD), and inherited retinal degenerations (IRDs). 44 , 45 , 46 , 47 Moreover, the relatively compact size of base editor components compared to full-length CRISPR systems facilitates their delivery using clinically relevant platforms such as adeno-associated viruses (AAVs) or lipid nanoparticles (LNPs). 48 Despite these significant advantages, challenges remain, including limitations in the types of nucleotide conversions achievable (restricted to transition mutations), the presence of bystander editing within the activity window, and potential off-target deaminase activity at unintended genomic or RNA sites. 49 , 50 , 51 , 52 Ongoing engineering efforts aim to expand the targeting scope, improve specificity, and develop next-generation base editors capable of broader mutation correction, such as prime editors, which offer the ability to introduce small insertions, deletions, and all types of point mutations with high precision. 49 , 53 While CRISPR-Cas9 remains the most widely recognized and versatile genome editing tool, capable of introducing targeted DSBs with relative ease, the complementary development of ZFNs and base editors has expanded the gene editing toolbox. One of CRISPR-Cas9’s most significant advantages is its ease of use. A CRISPR-Cas9 system can be designed to target almost any DNA sequence, and, by introducing multiple guide RNAs (single guide RNAs [sgRNAs] or dual guide RNA [dgRNAs]), it is possible to simultaneously edit multiple genes or genomic loci, a process known as multiplex gene editing ( Figure 2 ). This flexibility has greatly benefited fields like cancer research, where CRISPR is used to study the effects of gene mutations on tumor development. 54 Beyond medical research, CRISPR-Cas9 has also been employed in agricultural science. For example, Wada et al. highlighted its application in plant breeding, where CRISPR has been used to modify plant genomes to improve traits like disease resistance and crop yield. 11 This ability to make precise genetic changes offers significant advantages for improving food security and agricultural sustainability. CRISPR technology has also made significant strides in animal models, particularly mammals. For instance, Niu et al. successfully used CRISPR-Cas9 to modify the genomes of cynomolgus monkeys, highlighting its potential for creating models of human disease. 55 Such advancements have opened the door for translational research, where CRISPR can be used to test potential treatments for genetic disorders. However, despite the remarkable progress made with CRISPR-Cas9, several challenges remain. Off-target effects, where unintended genetic changes occur, are still a concern. Researchers are actively improving CRISPR accuracy and developing alternative systems, such as Cpf1 (also known as Cas12a), which offers some advantages in reducing off-target effects. 56 As CRISPR technology evolves, innovations such as these are crucial for addressing the remaining challenges. Figure 2 Mechanism of CRISPR-Cas9 gene editing The Cas9-sgRNA complex induces a double-stranded break at a target DNA site, which is repaired either by error-prone non-homologous end joining (NHEJ), causing insertions or deletions, or by precise homology-directed repair (HDR) using a donor DNA template.

CRISPR-Cas9 has reshaped the field of genetic engineering, providing a powerful and flexible tool for scientific research and biotechnology. Its influence spans various disciplines, from medicine and agriculture to basic biological research. As the technology continues to improve and diversify, new applications are constantly emerging. At the same time, discussions surrounding ethical considerations and regulatory frameworks are becoming increasingly important, particularly as CRISPR moves from research settings into therapeutic applications. 57 The ongoing refinement and responsible use of this technology hold great promise for the future of genetic science.

Delivery systems for CRISPR genome editing

Due to its precise gene editing capabilities, CRISPR offers significant promise for understanding and treating human genetic diseases. However, to be effective, CRISPR must reach the specific cells or tissues affected by the disease. 58 CRISPR faces challenges such as instability, susceptibility to degradation, and limited cellular uptake. Effective delivery strategies must address these limitations to ensure precise transport to target cells or tissues while minimizing off-target effects and reducing the immunogenicity associated with CRISPR components. 59 , 60 Maximizing the therapeutic benefits of CRISPR requires the development of reliable delivery platforms capable of safely and efficiently carrying various forms of CRISPR components to specific tissues and cells. 61 Delivery of the CRISPR-Cas9 system typically relies on three main approaches: physical techniques, viral vectors, and non-viral vectors each with specific benefits, drawbacks, and suitability for in vitro and in vivo applications. CRISPR-Cas components can be introduced in different formats, including plasmid DNA encoding Cas9 and guide RNA (gRNA), Cas9 mRNA with gRNA, or a ribonucleoprotein (RNP) complex composed of the Cas9 protein and gRNA. These delivery formats differ in structure, charge characteristics, and stability profiles.

Physical method

Physical methods like microinjection and electroporation have introduced CRISPR elements into target cells. Microinjection involves the direct delivery of Cas9 and sgRNAs into cells using a fine needle under microscopic guidance, allowing precise control over component placement. 62 This technique allows accurate plasmid DNA, mRNA, or RNP insertion straight into the nucleus, often achieving nearly 100% delivery efficiency. Bypassing barriers such as the extracellular matrix, cell membrane, and intracellular obstacles, microinjection provides precise control over the release of CRISPR components and helps reduce off-target activity. 63 Despite its precision, microinjection is limited by its time-consuming, manual process, requiring injecting cells one at a time. This lack of scalability confines its application primarily to in vitro or ex vivo cultured cells, making it unsuitable for in vivo use in human systems. 59 Electroporation uses brief high-voltage electrical pulses to create temporary pores in the cell membrane, allowing CRISPR components to enter the cell efficiently. 64 Electroporation has proven effective in editing mouse and rat embryos, initially developed for delivering Cas9 mRNA and later refined for the CRISPR-Cas9 RNP complex. In contrast to standard transfection methods, electroporation offers greater adaptability and is suitable for hard-to-transfect cell types. Delivering RNPs via this method enables rapid action, enhances gene editing efficiency, and reduces mosaicism in modified zygotes. Electroporation has also contributed to ex vivo gene editing advancements, particularly in stem cell therapies for disorders like hemoglobinopathies. Nevertheless, its application in human in vivo settings is limited due to the need for high-voltage pulses to permeabilize cell membranes.

Viral methods

Viruses have long been integral to medical research. In their natural form, wild-type viruses infect the human body and cause disease due to their pathogenic nature. However, through genetic engineering, viruses can be modified to eliminate their disease-causing potential while retaining their inherent ability to efficiently deliver genetic material into host cells. 65 , 66 This is typically achieved by removing or inactivating viral genes essential for replication and pathogenicity, while preserving or enhancing genes required for cell entry and gene transfer. For example, adenoviral vectors are rendered replication-deficient by deleting E1 and E3 genes, lentiviral vectors are modified by removing accessory genes and splitting essential components across multiple plasmids to enhance biosafety, and AAV vectors are engineered to package therapeutic transgenes while lacking the viral genome responsible for replication. 65 , 67 , 68 , 69 These modifications ensure that viral vectors can safely and efficiently deliver therapeutic genetic cargo without causing disease. 66 In recent years, AAVs, lentiviruses, and adenoviruses have become effective carriers for CRISPR-Cas9 delivery in both in vitro and in vivo applications. 70 AAVs, belonging to the Parvoviridae family, are small, non-enveloped viruses containing single-stranded DNA. Their non-pathogenic nature and low initial immunogenicity make them a relatively safe option for gene delivery in human cells. 71 Despite these advantages, AAVs face a significant constraint: their limited packaging capacity of approximately 4.5–5 kilobases makes it challenging to deliver the complete CRISPR-Cas9 system efficiently. 72 Lentiviruses, which can carry genetic material up to about 10 kb, can efficiently provide the complete CRISPR RNP complex in a single transfection. 73 However, their use carries the risk of off-target effects, as lentiviral vectors may cause insertional mutagenesis and lead to prolonged expression of the site-specific nucleases. 74 Adenovirus vectors can carry up to around 8 kb of foreign DNA. Unlike other viral vectors, they do not contain an endogenous integrase enzyme, which reduces the risk of off-target effects and insertional mutagenesis. 75 These vectors efficiently deliver CRISPR components into dividing and non-dividing cells, making them suitable for specific gene editing applications. However, their use is often complicated by the potential to trigger immune responses, cause tissue inflammation, and lead to the elimination of the vector. 76 , 77 To overcome these issues, approaches such as coating the vectors with polymers or utilizing non-human adenovirus variants have been explored, demonstrating reduced cross-reactive immunity associated with the CRISPR system. 78

Non-viral methods

Although physical and viral delivery methods have made significant progress, their limitations restrict widespread in vivo application of CRISPR. As an alternative, nanoparticles have gained attention as promising carriers, offering precise and safe delivery of CRISPR components to target cells to achieve therapeutic effects. 60 LNPs can form stable complexes with Cas9 plasmid DNA, mRNA, and gRNA, facilitating their entry across the cell membrane and promoting endosomal escape of CRISPR cargo within target cells. 79 However, the permanent positive charge of some cationic lipids in LNPs can cause problems such as toxicity, instability, and reduced transfection efficiency. 80 Researchers have focused on optimizing LNPs by incorporating ionizable cationic lipids and biodegradable lipid components to improve intracellular delivery and enhance transfection. 81 Ionizable lipids become positively charged in the acidic environment of late endosomes, allowing LNPs to interact with negatively charged endosomal membranes, which promotes membrane fusion and disruption. 82 This mechanism effectively releases the CRISPR payload into the cytoplasm. Moreover, biodegradable lipid-like nanoparticles carrying Cas9 protein and sgRNA have demonstrated high delivery efficiency, achieving gene knockouts in approximately 70%–90% of cultured human cells. 83 Polymer-based nanoparticles—including materials such as polyethylenimine (PEI), poly(L-lysine), poly[2-(dimethylamino)ethyl methacrylate], polyamidoamine, chitosan, and poly(amino-co-ester)s—show significant potential for in vivo CRISPR delivery due to their low immunogenicity, excellent biocompatibility, and ease of chemical modification. 60 , 84 , 85 , 86 , 87 , 88 , 89 These polymers have been applied to deliver CRISPR components in vitro and in vivo . However, their inherent toxicity remains challenging, necessitating further optimization strategies such as combining lipids to reduce toxicity while improving performance. Polymers like poly(lactic-co-glycolic acid) (PLGA), PEI, and chitosan can also be engineered within nanoparticle systems to enable controlled release and targeted delivery of CRISPR therapeutics. Exosomes are small extracellular vesicles, typically 30 to 150 nm in diameter, produced by almost all cell types and found throughout various bodily fluids. They naturally carry diverse molecules, including proteins, small nucleic acids, and lipids. 90 Exosomes offer several benefits when used as delivery vehicles: they can evade rapid clearance by the immune system, traverse biological barriers like the blood-brain and placental barriers, and selectively target specific cells or tissues due to their unique surface proteins. However, their limited size presents a challenge for efficiently packaging the relatively large CRISPR components. Various approaches have been explored, including engineering exosomes, creating hybrid delivery systems, and employing direct loading methods. 91

In vivo vs. ex vivo CRISPR delivery

Gene therapy delivery can be categorized into in vivo and ex vivo approaches. While currently approved gene therapy drugs typically do not modify human genetic material directly, emerging gene editing therapies aim to correct mutations within the human genome. Both in vivo and ex vivo strategies are actively being developed using advanced gene editing technologies to achieve these precise genetic corrections. 92 For in vivo therapy, genetic material is delivered directly into the patient’s target organ, either by using a vector or via the blood vessels supplying that organ. 93 Unlike ex vivo methods, in vivo therapy does not involve removing, modifying, and reintroducing the patient’s cells, simplifying the treatment process and reducing costs. However, this approach faces several challenges, including immune reactions against viral vectors, difficulties in delivering genetic material specifically to the correct cells and their nuclei, low efficiency of non-viral delivery systems, potential toxicity, and often only short-lived effects. In vivo gene editing is mainly explored for treating monogenic diseases affecting organs such as the liver, eyes, lungs, and muscles. Despite its potential, few in vivo gene editing therapies have advanced to clinical trials due to safety concerns and the risks associated with administering gene editing directly inside patients. 94 Ex vivo therapy consists of extracting cells from a patient, genetically correcting mutations in these cells outside the body and then transplanting the modified cells back into the patient. 95 Ex vivo gene therapy often targets stem cells because the corrected cells can rapidly replace those carrying faulty genes. This approach supports a wide range of delivery methods including viral vectors, liposomes, polymers, and physical techniques offering greater flexibility. 92 Additionally, toxicity is less of a concern since damaged or unedited cells can be excluded before reintroduction. The risk of insertional mutagenesis, which refers to the unintended integration of viral genetic material into the host genome, remains a concern in viral vector-based gene therapies, particularly with integrating vectors such as retroviruses and lentiviruses. Insertional events near oncogenes or tumor suppressor genes can disrupt normal gene regulation and potentially lead to malignant transformation, as was observed in early gene therapy trials for severe combined immunodeficiency (SCID), where some patients developed leukemia due to vector integration near proto-oncogenes. 96 , 97 However, in the context of immunotherapy such as ex vivo modification of T cells for chimeric antigen receptor (CAR)-T cell therapies, the risk is considered lower. This is because the modified cells can be extensively screened and expanded in vitro before reinfusion, and the transient lifespan of these immune cells reduces long-term risks compared to therapies targeting stem cells or dividing tissues. 98 Additionally, non-integrating viral vectors (e.g., AAVs) and improved vector designs have further minimized this risk in certain applications. However, ex vivo gene therapy has its limitations. This approach is primarily applicable to specific, easily accessible cell types that can be isolated, genetically modified, and reintroduced into the patient. 99 Common targets for ex vivo gene editing include hematopoietic stem cells (HSCs) for blood disorders, T lymphocytes for cancer immunotherapy, and, in experimental settings, mesenchymal stem cells or epithelial cells. 96 , 97 , 98 However, the ability to modify only a subset of the patient’s cells is limited by factors such as the efficiency of cell collection, the viability and expansion potential of modified cells, and the success of engraftment after reinfusion. 69 While HSC-based gene therapy can achieve long-lasting correction within the hematopoietic system, replacing large fractions of other tissue types remains challenging due to difficulties in harvesting sufficient numbers of progenitor cells and ensuring stable engraftment. 100 Moreover, ex vivo therapies require highly complex, personalized procedures involving specialized laboratory infrastructure, rigorous quality control, and high production costs, all of which restrict broad accessibility. Despite these limitations, ex vivo gene editing holds significant promise, particularly for blood disorders, infectious diseases such as HIV, and enhancing cancer immunotherapy. Numerous ongoing clinical trials are focused on refining delivery methods and improving the efficiency of gene editing platforms, primarily targeting blood cells or T lymphocytes to advance treatments for hematological and immune-related diseases. 100 , 101 , 102

CRISPR in clinical trials: Progress and early successes

The clinical translation of gene editing technologies is advancing rapidly, with numerous ongoing trials targeting a wide range of diseases. To enhance clarity, we have categorized the current clinical trial landscape according to major disease areas, including hematologic disorders (inherited blood diseases), cancer, chronic bacterial infections, hereditary amyloidosis, immunodeficiencies, cardiovascular diseases, viral infections, metabolic disorders, autoimmune diseases, muscular dystrophies, and inherited eye diseases. Although therapeutic targets and clinical endpoints differ across these indications, common technological patterns are evident in terms of editing platforms and delivery strategies. Ex vivo gene editing, primarily utilizing CRISPR-Cas9 or base editing technologies in combination with viral vectors such as lentiviruses, remains dominant for hematologic and immunologic disorders. In contrast, in vivo approaches often employ AAV or LNP-based delivery systems to enable targeted gene editing for metabolic, ocular, and other systemic diseases. In the following sections, we provide an updated overview of selected clinical trials for each disease category, highlighting key details including therapeutic targets, gene editing platforms, delivery methods, and available safety or efficacy outcomes.

Hematologic diseases (inherited blood disorders)

Gene editing therapies for inherited blood disorders are among the most clinically advanced, with ex vivo editing of hematopoietic stem and progenitor cells (HSPCs) using CRISPR-Cas9 or ZFN platforms showing promising results. Current trials primarily target β-globin gene mutations or fetal hemoglobin regulatory pathways in diseases such as β-thalassemia and SCD, with most studies in phase 1/2 or recently approved therapies entering clinical use. 103 , 104 Genetic blood disorders encompass a range of inherited conditions caused by mutations affecting blood cells, such as red blood cells, platelets, and white blood cells. 105 Key inherited blood diseases targeted by CRISPR technology include β-thalassemia, SCD, hemophilia B, and SCID. 106 These disorders rank among the most prevalent single-gene diseases globally. 107 Clinical trials using CRISPR-Cas9 or Cas12 systems in hematology primarily focus on β-thalassemia and SCD as part of interventional treatments. β-thalassemia and SCD are inherited disorders caused by mutations in the β-globin (HBB) gene (hemoglobin subunit beta), which encodes hemoglobin. Severe forms of β-thalassemia, such as β0/β+ or β+/β+, can be life threatening due to a significant reduction (β+) or complete absence (β0) of β-globin chains. 108 One therapeutic strategy involves using lentiviral vectors to deliver functional β-globin genes into HSCs. Another gene editing approach targets the BCL11A gene with ZFNs to modify patients’ HSPCs, thereby restoring fetal hemoglobin production. 109 Clinical trials using CRISPR-edited CD34+ HSPCs have demonstrated increased fetal hemoglobin levels, reducing disease symptoms and eliminating the need for blood transfusions in patients with β-thalassemia and SCD. 103 Notably, Casgevy—a CRISPR-Cas9-based therapy targeting fetal hemoglobin genes in stem cells—has been approved by the Food and Drug Administration (FDA) as the first gene editing treatment worldwide. 110 Most ongoing CRISPR clinical trials for transfusion-dependent β-thalassemia (covering genotypes such as β0/β0, β+/β0, βE/β0, and β+/β+) focus on disrupting the erythroid-specific enhancer region of the BCL11A gene or the HBG1/2 promoter to reactivate γ-globin expression in the patient’s own hematopoietic stem or progenitor cells ( Table 1 ). Table 1 Overview of ongoing clinical trials utilizing CRISPR-Cas9 gene editing for β-thalassemia treatment, detailing trial identifiers, phases, targeted disease conditions, gene targets, intervention types, editing methods, and sponsoring organizations Trial ID Phase Disease condition Target gene Intervention Gene editing method Sponsor Reference NCT04211480 ; NCT05577312 1/2 β-thalassemia major (β 0 /β 0 , β + /β 0 , β E /β 0 ) BCL11A gene disruption CRISPR-Cas9 BRL Medicine Fu et al. 111 NCT05356195 ; NCT03655678 – TDT-homozygous or compound heterozygous β-thalassemia including β-thalassemia/hemoglobin E BCL11A gene disruption; CTX001 CRISPR-Cas9 Vertex Pharmaceuticals Frangoul et al. 103 NCT04925206 1 TDT defined by protocol BCL11A enhancer gene disruption; ET-01 CRISPR-Cas9 EdiGene (GuangZhou) – NCT06041620 – TDT defined by protocol HBG1/2 promoter gene disruption; VGB-Ex01 CRISPR-Cas12b Institute of Hematology & Blood Diseases Hospital – NCT05444894 1/2 TDT-homozygous or compound heterozygous β-thalassemia including β-thalassemia/hemoglobin E HBG1/2 promoter gene disruption; EDIT-301 CRISPR-Cas12a Editas Medicine – ChiCTR2100052858 1 TDT (β 0 /β 0 , β + /β 0 , β E /β 0 , β + /β + ) HBG1/2 promoter gene disruption; RM-001 CRISPR-Cas9 Guangzhou Reforgene Medicine Wang et al. 112 ChiCTR2100053406 1 TDT (genotype not indicated) HBG1/2 promoter gene disruption; RM-001 CRISPR-Cas9 Guangzhou Reforgene Medicine Liu et al. 113 NCT05477563 3 TDT- same as above; severe SCD BCL11A gene disruption; CTX001 CRISPR-Cas9 Vertex Pharmaceuticals – NCT03745287 ; NCT05329649 2/3; 3 severe SCD as defined BCL11A gene disruption; CTX001 CRISPR-Cas9 Vertex Pharmaceuticals Frangoul et al. 103 NCT04443907 1/2 SCD (β S /β S , β S /β C , β S /β 0 , or others) BCL11A gene disruption; OTQ923 CRISPR-Cas9 Novartis Pharmaceuticals – NCT05951205 3 β S /β C BCL11A gene disruption; Exa-cel CRISPR-Cas9 Vertex Pharmaceuticals – NCT04774536 1/2 severe SCD as defined HBB gene correction; CRISPR-SCD001 CRISPR-Cas9 Mark Walters, MDUCSF – NCT05456880 1/2 SCD with β S /β S , β S /β 0 , or β S /β + genotypes & severe SCD as defined HBG1/2 promoter gene correction; BEAM-101: an ABE base-edited CD34 + hematopoietic stem base editing Beam Therapeutics – NCT04853576 1/2 severe SCD (β S /β S , β S /β 0 , β S /β + ) HBG1/2 promoter gene disruption; EDIT-301 CRISPR-Cas12a Editas Medicine –

Cancer

Gene editing clinical trials in cancer predominantly focus on adoptive cell transfer (ACT) therapies, especially CRISPR-Cas9-edited T cells including CAR-T, T cell receptor (TCR)-T, and tumor-infiltrating lymphocytes (TILs), with most studies currently in early-phase trials (phase 1/2) aimed at improving anti-tumor efficacy and safety through precise gene disruption. 114 , 115 , 116 , 117 Electroporation of Cas9 RNP complexes into patient-derived T cells remains the leading delivery approach for ex vivo gene editing in these immunotherapies. 114 , 118 Cancer represents a highly heterogeneous group of diseases where gene editing is chiefly being explored to enhance immune-based therapies. Most CRISPR-based clinical trials in cancer are focused on developing therapeutic interventions for both hematologic malignancies and solid tumors, with a particular emphasis on enhancing ACT strategies. 116 ACT involves the isolation, genetic modification, and reinfusion of tumor-specific T lymphocytes to enhance their anti-tumor activity. 116 The three main types of ACT are CAR-T cells, TCR-T cells, and TILs, all of which have been modified using CRISPR-Cas9 technology, primarily through electroporation of Cas9 RNP complexes. 119 CAR-T cells are created by combining human T cells with synthetic CARs that enable the cells to specifically recognize and bind to tumor-associated antigens independent of the major histocompatibility complex (MHC). 120 CARs are engineered receptors typically composed of three key regions: an extracellular single-chain variable fragment (scFv) that recognizes the tumor antigen, a spacer and transmembrane domain, and intracellular signaling domains such as CD28-CD3ζ or 4-1BB-CD3ζ, which activate the T cell upon antigen binding. 121 This design combines the high specificity of antibody-mediated antigen recognition with the cytotoxic capacity of T cells, enabling CAR-T cells to destroy cancer cells without the need for antigen presentation through MHC molecules, particularly human leukocyte antigens (HLA). As a result, CAR-T therapies offer broader applicability compared to TCR-T cells, which remain restricted by MHC compatibility 122 ( Table 2 ). In addition to CAR-T technology, CRISPR-Cas9 gene editing is increasingly used to enhance the safety, specificity, and efficacy of ACT-based cancer therapies. CRISPR is applied to precisely disrupt genes that limit T cell function or contribute to immune evasion by tumors. For example, programmed cell death protein 1 (PD-1) is a critical immune checkpoint receptor that suppresses T cell activity within the immunosuppressive tumor microenvironment. 134 CRISPR-mediated knockout of the PDCD1 gene encoding PD-1 has been shown to restore T cell effector function and improve anti-tumor responses. 134 , 135 Moreover, CRISPR-Cas9 is used to delete the endogenous TCR α and β chains, which prevents graft-versus-host disease (GVHD) in allogeneic T cell therapies by eliminating native TCRs and replacing them with tumor-specific receptors. 136 , 137 , 138 This strategy enhances safety in “off-the-shelf” allogeneic CAR-T cell products. Furthermore, multiplex gene editing using CRISPR enables simultaneous modification of multiple genes within T cells, allowing researchers to address complex mechanisms of tumor resistance, reduce immune exhaustion, and improve T cell persistence and function. 114 Several early-phase clinical trials have demonstrated the feasibility and safety of CRISPR-engineered T cells in cancer treatment. 114 , 139 A landmark first-in-human clinical trial showed that CRISPR-mediated PD-1 disruption, along with deletion of endogenous TCR chains, in T cells infused into patients with refractory cancers was technically feasible and well tolerated. 114 Building on this, ongoing research is exploring CRISPR-based strategies to insert CAR constructs into defined genomic loci, such as the TRAC locus, which has been shown to improve T cell fitness, enhance anti-tumor responses, and reduce tonic signaling that can otherwise impair CAR-T cell efficacy. 140 , 141 Thus, CRISPR technology has become an essential tool in advancing T cell-based cancer immunotherapies. By enabling precise, efficient, and multiplex gene editing, CRISPR is paving the way for safer, more potent, and more personalized therapeutic strategies for both hematologic cancers and solid tumors. 115 Table 2 Overview of CRISPR-Cas9 clinical trials targeting cancer Trial ID Phase Disease Target Cell type Intervention Gene editing method Country Reference NCT04035434 1/2 relapsed/refractory B cell malignancies TRAC and B2M allogeneic T cells CTX110: CD19-directed allogeneic T cells genetically modified ex vivo using CRISPR-Cas9 gene editing components – United States, Australia, Canada, France, Germany, Spain McGuirk et al. 123 NCT05643742 1/2 relapsed/refractory B cell malignancies Regnase-1, TGFβR2, TRAC, B2M, and CD70 allogeneic T cells CTX112: CTX110 + disruption of Regnase-1 and TGFβR2 CRISPR-Cas9 United States Terrett et al. 124 NCT03166878 1/2 relapsed/refractory CD19+ leukemia or lymphoma (multiple myeloma) TCR and B2M allogeneic T cells engineered allogeneic anti-BCMA CAR-T – China – NCT04037566 1 relapsed/refractory CD19+ leukemia or lymphoma HPK1 autologous T cells engineered autologous anti-CD19 CAR-T (XYF19 CAR-T cells) – China Zhang et al. 125 NCT04637763 1 relapsed/refractory B cell non-Hodgkin lymphoma TRAC and PD-1 allogeneic T cells CB-010: a CRISPR-edited allogeneic CAR-T cell therapy targeting CD19 – United States – NCT03398967 1/2 B cell leukemia/B cell lymphoma CD52 and TRAC allogeneic T cells engineered allogeneic anti-CD19 and CD20 or CD22 CAR-T – China – NCT04557436 1 B cell acute lymphoblastic leukemia CD52 and TRAC allogeneic T cells engineered allogeneic anti-CD19 CAR-T – United Kingdom Ottaviano et al. 126 NCT05631912 1/2 non-Hodgkin lymphoma, B cell TRAC autologous T cells engineered autologous CD19-STAR-T cell CRISPR-Cas9 China – NCT06014073 1/2 non-Hodgkin lymphoma TRAC and Power3 allogeneic T cells TRAC and Power3 genes knockout allogeneic CD19-targeting CAR-T cell (ATHENA CAR-T) – China – NCT04213469 1 relapse/refractory B cell lymphoma PD-1 autologous T cells gene editing of autologous T cells with anti-CD19 ScFv expression and knockout of PD-1 – China Zhang et al. 127 NCT04767308 1 relapsed/refractory hematopoietic malignancies CD5 autologous T cells the endogenous CD5 in CT125A cells is knocked out to prevent fratricide during CAR-T cell manufacturing CRISPR-Cas9 China Dai et al. 128 NCT05722418 1 relapsed/refractory multiple myeloma B2M and insertion of a B2M–HLA-E fusion protein allogeneic T cells CB-011: an allogeneic CAR-T cell therapy targeting BCMA; cyclophosphamide chemotherapy for lymphodepletion and fludarabine chemotherapy for lymphodepletion – United States – NCT04244656 1 relapsed/refractory multiple myeloma TRAC and B2M allogeneic T cells CTX120: BCMA-directed engineered allogeneic anti-CD19 CAR-T cells CRISPR-Cas9 United States, Australia, Canada, Spain Dar et al. 129 NCT04502446 1 T cell lymphoma CD70, TRAC, and B2M allogeneic T cells CTX130: engineered allogeneic anti-CD70 CAR-T cells CRISPR-Cas9 United States, Australia, Canada Iyer et al. 130 NCT05397184 1 relapsed/refractory T cell acute lymphoid leukemia CD52, CD7, and TCR allogeneic T cells cryopreserved BE-CAR7 T cells (BE752TBCCLCAR7PBL); single-dose intravenous infusion (weight-based dosing) of a banded dose of CAR7+ T cells/kg BE-CAR7 – United Kingdom – ISRCTN15323014 1 T cell acute lymphoid leukemia TRBC, CD7, and CD52 allogeneic T cells single-dose intravenous infusion (weight-based dosing) of a banded dose of CAR7+ T cells/kg BE-CAR7 – United Kingdom Chiesa et al. 131 NCT03545815 1 mesothelin-positive solid tumors PD-1 and TCR autologous T cells engineered autologous anti-mesothelin CAR-T cells CRISPR-Cas9 China Wang et al. 132 NCT03747965 1 mesothelin-positive solid tumors PD-1 T cells infusions of mesothelin-directed CAR-T cells 1 day after the completion of conditioning regimen of paclitaxel and cyclophosphamide CRISPR-Cas9 China – NCT04976218 1 advanced EGFR-positive solid tumors TGFβR2 T cells TGFβR-KO CAR-EGFR T cells – China – NCT05812326 1/2 advanced breast cancer PD-1 autologous T cells PD-1 knockout anti-MUC1 CAR-T cells – China – NCT04438083 1 renal cell carcinoma CD70, TRAC, and B2M allogeneic T cells CTX130: engineered allogeneic anti-CD70 CAR-T cells CRISPR-Cas9 United States, Australia, Canada, Netherlands Pal et al. 133 NCT05795595 1/2 relapsed or refractory solid tumors CD70, TRAC, B2M, Regnase-1, and TGFβR2 allogeneic T cells CTX131: CTX130 + disruption of Regnase-1 and TGFβR2 CRISPR-Cas9 United States Terrett et al. 124

Chronic bacterial infection

Gene editing approaches targeting bacterial virulence factors, such as quorum sensing genes involved in biofilm formation, are emerging as innovative strategies to combat persistent infections. 142 , 143 Current research, mostly at the preclinical or early exploratory stage, utilizes CRISPR-Cas9 to disrupt key bacterial genes and impair biofilm development, with potential implications for improving treatment of device-associated infections. 144 , 145 , 146 , 147 Chronic bacterial infections caused by Escherichia coli in urinary tract infections (UTIs) represent a significant clinical challenge, particularly due to the bacteria’s ability to form biofilms on indwelling urinary catheters. 148 These biofilms protect bacteria from host defenses and antibiotic treatments, contributing to persistent infections. Kang et al. (2017) applied CRISPR-Cas9 genome editing to clinical E. coli SE15 isolates, precisely deleting the luxS gene, which plays a critical role in quorum sensing and biofilm formation. Their study demonstrated that disruption of luxS significantly reduced biofilm formation and downregulated related biofilm-associated genes, suggesting that targeted genome editing can elucidate molecular mechanisms underlying chronic infections and may provide a foundation for developing novel therapeutic strategies against catheter-associated UTIs. 149 Traditional antibiotics often fail to eliminate these pathogens, leading to recurrent infections and long-term tissue damage. CRISPR-based technologies offer a novel approach to combating chronic bacterial infections by enabling precise targeting and disruption of bacterial genomes. 143 Engineered bacteriophage viruses that infect bacteria can be modified to deliver CRISPR-Cas systems (such as Cas9 or Cas3) into pathogenic bacteria. These systems can selectively cut essential genes or resistance determinants, leading to bacterial death or antibiotic sensitization. 150 This targeted approach holds promise for overcoming antibiotic resistance and providing durable solutions for persistent bacterial infections. Moreover, the modular nature of CRISPR systems allows for rapid reprogramming to combat emerging bacterial threats or outbreaks ( Table 3 ). Table 3 Ongoing clinical trials using CRISPR-based approaches for targeting infectious diseases Trial ID Phase Disease Target Cell type Intervention Gene editing method Country Reference NCT05277350 1 E. coli infections bacteria bacteriophages SNIPR001: engineered phages targeted bacteria CRISPR-Cas9 SNIPR BIOME ApS, Denmark Gencay et al. 151 NCT06474416 1 herpes simplex virus type I stromal keratitis (HSK) HSV-1 virus gene HSV-1-erasing lentiviral particles (HELPs) BD111: lentivirus-like particle delivering gRNA-expressing cassettes and SpCas9 mRNA CRISPR-Cas9 Shanghai BDgene Co., Ltd, China – NCT06474442 2 herpes simplex virus type I stromal keratitis (HSK) HSV-1 virus gene HSV-1-erasing lentiviral particles (HELP) BD111: lentivirus-like particle delivering gRNA-expressing cassettes and SpCas9 mRNA CRISPR-Cas9 Shanghai BDgene Co., Ltd, China – NCT04191148 1 urinary tract infections, UTI ( E. coli ) bacteria causing urinary tract infections bacteriophages LBP-EC01: engineered bacteriophage cocktail with a CRISPR-Cas3 construct targeting the E. coli genome CRISPR-Cas3 Locus Biosciences, Inc, United States – NCT05488340 2 urinary tract infections, UTI ( E. coli ) bacteria causing urinary tract infections bacteriophages LBP-EC01: engineered bacteriophage cocktail with a CRISPR-Cas3 construct targeting the E. coli genome CRISPR-Cas3 Locus Biosciences, Inc, United States Kim et al. 152

Hereditary amyloidosis

CRISPR-based therapies for hereditary amyloidosis are advancing rapidly, with a primary focus on reducing circulating transthyretin (TTR) protein levels to prevent progressive organ damage. 153 Current clinical strategies employ LNP delivery systems for in vivo administration of CRISPR-Cas9 components, with early-phase trials demonstrating promising safety and efficacy profiles. 154 Transthyretin amyloidosis (ATTR) is a life-threatening multisystem disorder caused by point mutations in the TTR gene, leading to the misfolding and pathological accumulation of TTR protein in tissues, particularly in peripheral nerves and the heart. 155 , 156 This protein aggregation results in progressive polyneuropathy, cardiomyopathy, and, ultimately, organ failure. 157 ATTR shares mechanistic features with other amyloid-related neurodegenerative diseases such as Alzheimer’s and Parkinson’s disease. 158 A first-in-human gene editing approach for ATTR is exemplified by Intellia Therapeutics, Inc. (NTLA),2001, an investigational in vivo CRISPR-Cas9 therapy designed to achieve permanent reduction of pathogenic TTR protein levels. 159 NTLA-2001 employs LNPs to encapsulate mRNA encoding the Cas9 endonuclease, along with a sgRNA precisely targeting the TTR gene in hepatocytes, the primary source of circulating TTR. 153 Preclinical studies demonstrated a durable knockout of the TTR gene and sustained reduction of serum TTR levels following a single dose. 153 In an ongoing phase 1 clinical trial ( NCT04601051 ), NTLA-2001 has been evaluated for safety and pharmacodynamic effects in patients with hereditary ATTR amyloidosis presenting with polyneuropathy. Initial cohorts received escalating single doses of either 0.1 mg/kg or 0.3 mg/kg. The treatment was generally well tolerated, with only mild adverse events reported. Notably, a dose-dependent reduction in serum TTR levels was observed: by day 28, patients in the lower-dose group exhibited an average 52% reduction in TTR (range 47%–56%), while those in the higher-dose group experienced an impressive 87% reduction (range 80%–96%). 153 These findings mark a significant milestone as NTLA-2001 represents the first systemically administered, in vivo CRISPR-Cas9 therapeutic to achieve targeted gene editing in humans. By permanently disrupting the TTR gene, this one-time treatment has the potential to halt disease progression and reduce the burden of lifelong therapies for patients with hereditary ATTR amyloidosis 153 ( Table 4 ). Table 4 Ongoing clinical trials investigating CRISPR-based therapies for transthyretin amyloidosis Trial ID Phase Disease Target Delivery system Intervention Gene editing method Country Reference NCT06082050 ; NCT06539208 1; 1/2 transthyretin amyloidosis cardiomyopathy (ATTR) – – YOLT-201 – YolTech Therapeutics, china – NCT04601051 ; NCT06128629 1; 3 transthyretin amyloidosis cardiomyopathy (ATTR-CM) accumulated misfolded TTR protein LNP encapsulating mRNA for Cas9 protein and an sgRNA targeting TTR NTLA-2001 engineered LNP CRISPR-Cas9 Intellia Therapeutics, EU, UK, and New Zealand Gillmore et al. 153

Immunodeficiency

Gene editing approaches for immunodeficiency disorders are advancing toward precise, durable therapies, with ex vivo base editing strategies being explored for primary immunodeficiencies like X-linked chronic granulomatous disease (X-CGD), while in vivo CRISPR-based treatments delivered via LNPs are under investigation for conditions such as hereditary angioedema (HAE), with early-phase clinical trials demonstrating feasibility. 160 , 161 X-CGD is a primary immunodeficiency disorder caused by mutations in the CYBB gene, which encodes the cytochrome b-245 beta chain. Traditional gene therapy approaches for X-CGD involve modifying a patient’s HSPCs using lentiviral vectors carrying a corrected CYBB sequence. However, these methods face limitations, such as reduced expression due to the use of weak promoters to minimize the risk of activating oncogenes and the absence of natural gene regulation. Base editing offers a potential alternative by enabling precise correction of point mutations without introducing DSBs. A recent study by Bzhilyanskaya et al. demonstrated using an optimized protospacer adjacent motif (PAM)-flexible Cas9 variant, streptococcus pyogenes Cas9-RY variant (SpRY), fused to the adenine base editor ABE8e. This system successfully corrected two disease-causing mutations in CYBB with high precision and efficiency in patient-derived HSPCs (clinical trial ID: NCT06325709 ). 160 HAE is a rare inherited disorder characterized by recurrent episodes of severe swelling in various parts of the body. NTLA-2002 is a gene editing treatment currently under investigation, utilizing CRISPR-Cas9 technology for in vivo application. This therapy aims to provide a single-dose solution by permanently disrupting the KLKB1 gene, which encodes plasma prekallikrein a protein that triggers swelling episodes. To achieve this, NTLA-2002 employs liver-targeted LNPs to deliver two essential components: mRNA coding for the Cas9 nuclease and an sgRNA that directs the editing machinery to the KLKB1 gene. This approach lowers plasma kallikrein levels by editing the gene directly in hepatocytes. The treatment is being evaluated in a clinical study registered as NCT05120830 162 ( Table 5 ). Table 5 Ongoing clinical trials employing CRISPR-Cas9 and base editing for the treatment of chronic granulomatous disease and hereditary angioedema Trial ID Phase Disease Target Delivery system Intervention Gene editing method Country Reference NCT06325709 1/2 chronic granulomatous disease (CGD) autologous hematopoietic stem and progenitor cells (HSPCs) lentiviruses containing the sequence of the corrected cytochrome b-245 beta chain (CYBB) gene base-edited stem cells to improve the white cells’ functioning and fewer CGD-related infections base editing National Institute of Allergy and Infectious Diseases (NIAID), United States Bzhilyanskaya et al. 160 NCT05120830 ; NCT06634420 1/2; 3 hereditary angioedema (HAE) expressed KLKB1 in the liver LNP encapsulating mRNA for Cas9 protein and a 20-nucleotide sgRNA targeting KLKB1 NTLA-2002; engineered gene encoding kallikrein B1 (KLKB1) to reduce its plasma level CRISPR-Cas9 Intellia Therapeutics, US, UK, and New Zealand Longhurst et al. 162

Cardiovascular disease

Gene editing therapies are rapidly emerging as innovative solutions for reducing cardiovascular risk by targeting genetic contributors to elevated lipoproteins and cholesterol levels. Several CRISPR-based candidates employ precise genome and base editing technologies delivered via LNPs to achieve durable suppression of key pathogenic genes in liver cells. 163 , 164 Genetic and epidemiological evidence have established that high levels of lipoprotein (a) (Lp(a)) are a causal factor in atherosclerosis and associated cardiovascular conditions. CTX320 is a novel in vivo CRISPR-Cas9-based gene editing therapy developed to silence the gene encoding apolipoprotein (a), a key component of Lp(a), specifically within liver cells. The therapeutic formulation includes Cas9 mRNA and a gRNA, encapsulated in LNPs for effective systemic delivery. In this study, we evaluated the performance, safety profile, and lasting impact of CTX320 in both primary human cell models and non-human primate subjects. 165 Low-density lipoprotein cholesterol (LDL-C) is critical in developing atherosclerotic cardiovascular disease (ASCVD). Although lipid-lowering treatments like statins have significantly improved patient outcomes, individuals with heterozygous familial hypercholesterolemia (HeFH) often struggle to achieve optimal LDL-C reduction. PCSK9, an enzyme recently discovered, contributes to elevated LDL-C levels by binding to low-density lipoprotein receptors (LDLRs) and promoting their breakdown. Verve Therapeutics, Inc. (VERVE)-101 is an innovative CRISPR-based gene editing approach designed to silence the PCSK9 gene in the liver permanently. This therapy utilizes a LNP system to deliver messenger RNA encoding an adenine base editor and a guide RNA directed at the PCSK9 gene. The targeted A•T to G•C substitution disrupts a splice donor site, effectively inactivating PCSK9. Preclinical studies in mice and non-human primates have demonstrated long-term suppression of PCSK9 and sustained LDL-C reduction, with effects in primates lasting over 12 months following a single intravenous dose. 166 Familial hypercholesterolemia (FH) is an inherited condition that impairs the body’s ability to clear LDL-C, commonly known as “bad” cholesterol, resulting in elevated blood levels that increase the risk of premature cardiovascular disease. YOLT-101, developed by YolTech, targets specific genetic mutations linked to FH using cutting-edge gene editing techniques combined with LNP delivery systems. Preclinical studies have shown that a single administration of YOLT-101 significantly lowers LDL-C levels for nearly 2 years in non-human primate models. VT-10201 is an open-label phase 1b trial involving a single ascending dose aimed at evaluating the safety of VERVE-102 in patients diagnosed with HeFH or premature coronary artery disease (CAD) who need additional LDL-C reduction. VERVE-102 employs base editing to disrupt PCSK9 gene expression in the liver, thereby reducing circulating PCSK9 and LDL-C levels. This study primarily focuses on assessing the safety and pharmacodynamic effects of VERVE-102 in the target patient group. 167 VERVE-201 is an experimental therapy consisting of messenger RNA encoding an adenine base editor alongside a guide RNA targeting the ANGPTL3 gene, all encapsulated within a N-Acetylgalactosamine (GalNAc)-conjugated LNP. Administered as a single intravenous infusion, this treatment aims to silence the ANGPTL3 gene in the liver, thereby reducing LDL-C levels to combat ASCVD. The initial focus of VERVE-201 is on treating HeFH, a rare inherited condition characterized by severe or complete deficiency of the LDLR, which results in extremely high LDL-C levels and early onset of ASCVD 168 ( Table 6 ). Table 6 Clinical trials of gene editing therapies for cardiovascular diseases Trial ID Phase Disease Target Delivery system Intervention Gene editing method Country Reference ACTRN12623001095651p 1 cardiovascular disease or calcific aortic valve stenosis apolipoprotein (a) component of Lp(a) in the liver LNP encapsulating Cas9 mRNA and gRNA targeting lipoprotein (a) (Lp(a)) CTX320 designed to knock out the apo-Lp(a) component of Lp(a) in the liver CRISPR-Cas9 CRISPR Therapeutics, Boston, MA Morrow et al. 165 NCT05398029 1 familial hypercholesterolemia (FH) the proprotein convertase subtilisin kexin type 9 (PCSK9) gene in the liver LNP encapsulating Cas9 mRNA and gRNA targeting PCSK9 gene VERVE-101 designed to inactivate the PCSK9 gene in the liver base editing Verve Therapeutics, Inc., Boston, MA Horie and Ono 166 NCT06461702 1 familial hypercholesterolemia (FH) the PCSK9 gene in the liver LNP encapsulating Cas9 mRNA and gRNA targeting PCSK9 gene YOLT-101 engineered to decrease the PCSK9 gene CRISPR-Cas9 First Affiliated Hospital Bengbu Medical College, YolTech Therapeutics, china – NCT06164730 1 familial hypercholesterolemia, HeFH, or premature coronary artery disease (CAD) the PCSK9 gene in the liver N-acetylgalactosamine (GalNAc)-LNP to target the liver VERVE-102 designed to disrupt the expression of the PCSK9 gene in the liver and lower circulating PCSK9 and LDL-C base editing Verve Therapeutics, Inc., Boston, MA Hooper et al. 167 NCT06451770 1 refractory hypercholesterolemia the ANGPTL3 gene GalNAc LNP packaging an adenine base editor mRNA and gRNA targeting ANGPTL3 gene VERVE-201 designed to inactivate ANGPTL3 in the liver and decrease LDL-C concentrations base editing VERVE Therapeutics, Inc., Boston, MA Lee et al. 168

Viral diseases

Gene editing therapeutics are being actively explored to target persistent viral infections and chronic bacterial diseases, with innovative strategies aimed at directly eradicating viral reservoirs or enhancing host resistance. 169 , 170 Current clinical trials employ diverse gene editing platforms, including ARCUS nucleases, CRISPR-Cas systems, and ZFNs, delivered via LNPs or viral vectors to achieve precise and durable modification of infected or susceptible cells. 169 , 170 , 171 Precision BioGene-Hepatitis B virus(PBGENE-HBV) represents a pioneering therapeutic candidate in clinical trials aimed at eradicating the underlying cause of chronic hepatitis B infection: the persistent covalently closed circular DNA (cccDNA) residing in liver cells. 172 Unlike existing antiviral treatments, which require daily dosing and achieve only a 1%–3% rate of functional cure, PBGENE-HBV employs the proprietary ARCUS gene editing technology. 173 This system delivers a specifically engineered nuclease encapsulated in LNPs directly to hepatocytes, where it selectively targets and destroys both the cccDNA and integrated viral DNA sequences. 171 The clinical study ( NCT04560790 ) is an investigator-led, open-label, single-arm, non-randomized trial conducted at a single institution. Separately, an investigational CRISPR-based treatment for herpes simplex virus type 1 (HSV-1) stromal keratitis, known as HELP, was administered as a single dose to three patients with severe refractory disease during corneal transplant procedures. 174 HELP’s therapeutic agent is a lentiviral vector suspended in phosphate-buffered saline, which carries two types of RNA within the same particle: SpCas9 mRNA and a gRNA cassette. The vector includes an integrase mutation (D64V) to prevent integration into the host genome. Instead, the gRNA cassette forms circular episomal DNA in infected cells, enabling expression of two gRNAs targeting HSV-1 genes UL8 and UL29. 175 This study aims to explore a novel approach for treating HIV using a protein called ZFN. ZFNs are engineered proteins designed to remove the CCR5 protein, which certain common strains of HIV (CCR5-tropic) use to gain entry into T cells. T cells, especially the subset known as CD4 T cells, play a crucial role in the immune system’s defense against HIV infection. By targeting and eliminating CCR5, this method seeks to prevent the virus from infecting these vital immune cells. 169 Some individuals naturally lack the CCR5 protein on their T cells, and these people remain healthy while showing strong resistance to HIV infection. Others have lower amounts of CCR5 on their T cells, which is linked to a milder progression of HIV infection and slower development of AIDS. 176 This study aims to remove the CCR5 protein from T cells by collecting large quantities of these cells from participants and introducing ZFNs using a delivery method called an adenoviral vector. The vector is applied early in the cell preparation process, allowing the ZFNs to eliminate CCR5. By the time the modified T cells are given back to the participants, very little of the adenovirus or ZFN remains, but the CCR5 removal is permanent. 176 The treatment involves extracting white blood cells containing CD4+ T cells from the patient’s blood. These CD4+ T cells are then genetically altered using ZFNs to remove the CCR5 gene, which HIV uses to enter the cells. After modification, a larger batch of these resistant T cells is produced and infused back into the patient. The goal is for these edited cells to resist HIV infection and to generate new HIV-resistant CD4+ T cells within the body. Laboratory experiments have demonstrated that CD4+ T cells modified with ZFNs can withstand HIV infection, preventing the virus from destroying them. These promising findings suggest that ZFN therapy might help HIV-infected individuals by preserving their CD4+ T cells and strengthening their immune response. 176 SB-728-T is a therapeutic product created by applying ZFNs delivered either through an adenoviral vector or as mRNA to modify a patient’s own CD4+ T cells by targeting the CCR5 gene. The engineered ZFNs are designed to induce a precise DSB within the coding region of the CCR5 gene. This gene editing approach aims to generate a stable population of CD4+ T cells that are permanently resistant to HIV infection. As a result, SB-728-T therapy has the potential to enhance immune recovery and contribute to sustained functional control of HIV in affected individuals. 177

Gene therapy presents a promising alternative to lifelong antiretroviral therapy for treating HIV-1 infection. The chemokine receptor CCR5 serves as a critical co-receptor that allows R5-tropic HIV-1 to enter human cells. 26 Removing CCR5 makes cells resistant to this form of HIV-1 infection, a fact highlighted by cases where allogeneic stem cell transplants from donors naturally lacking CCR5 led to potential cures. However, the necessity of finding HLA-matched donors with homozygous CCR5 deletions limits the broad use of this approach. 26 To overcome this, researchers have developed a method to disrupt the CCR5 gene directly in HSPCs using ZFNs. After consultations with regulatory bodies, preclinical studies were conducted both in vitro and in vivo to assess the feasibility and safety of this ZFN-based CCR5 editing in HSPCs. This report details the clinical-scale manufacturing process for delivering CCR5-targeted ZFN mRNA into HSPCs via electroporation, along with supporting preclinical safety findings. The data show efficient bi-allelic disruption of CCR5 in up to 72.9% of modified colony-forming units derived from mobilized adult HSPCs, with preserved hematopoietic function demonstrated both in laboratory and animal models. 26

The safety profile of CRISPR-based genome editing for human gene therapy remains largely unestablished. Targeting CCR5 is a plausible strategy for curing HIV-1 infection, as cells lacking CCR5 exhibit significant resistance to HIV-1 entry, though it is not entirely protective. In one case, HSPCs with CRISPR-mediated CCR5 disruption were transplanted into a patient suffering from both HIV-1 infection and acute lymphoblastic leukemia (ALL). 178 Following transplantation, the patient’s ALL reached complete remission with full donor chimerism, and the edited donor cells lacking CCR5 were detectable for over 19 months without any adverse effects related to the gene editing. During a monitored interruption of antiretroviral therapy, a modest increase in the proportion of CD4+ cells with CCR5 disruption was observed. 178 While this case demonstrated the feasibility of successful transplantation and sustained engraftment of CRISPR-edited HSPCs, the level of CCR5 disruption in lymphocytes remained low around 5% highlighting the necessity for continued investigation to improve editing efficiency in this therapeutic approach. 178

Metabolic disorders

Gene editing therapies for metabolic disorders are predominantly focused on in vivo strategies using AAV vectors or LNPs to deliver nucleases or gene templates directly to the liver, the key metabolic organ. Current clinical trials target a spectrum of rare, often life-threatening, inherited diseases, with early-phase studies demonstrating encouraging safety and evidence of durable therapeutic effects. Several gene editing therapies are currently being evaluated in clinical trials for the treatment of metabolic disorders, with promising developments targeting both rare and severe inherited conditions. Sangamo Therapeutics has pioneered in vivo gene editing using ZFNs to treat MPS types I and II through phase 1/2 trials ( NCT02702115 and NCT03041324 ), demonstrating favorable safety profiles and evidence of targeted genome editing in the liver, although long-term enzyme expression remains a challenge. 29 In parallel, iECURE, Inc. is conducting the ornithine transcarbamylase-HBB on-target precision editing (OTC-HOPE) trial ( NCT06255782 ) for neonatal-onset ornithine transcarbamylase deficiency (OTCD), employing a dual-vector AAV system that integrates a functional OTC gene into the hepatocyte genome. 179 Initial results from the first treated infant showed encouraging safety and biochemical improvements, supporting the potential of durable, genome-integrated therapies for OTCD. Homology Medicines has initiated the phenylketonuria editing (pheEDIT) phase 1 trial ( NCT05222178 ) for phenylketonuria, utilizing a novel AAV-mediated, non-nuclease gene editing approach to integrate functional phenylalanine hydroxylase (PAH) genes into the genome. 180 Preclinical studies demonstrated reduced phenylalanine levels and integration specificity, and early human trials are underway to assess safety and efficacy, with plans to eventually expand into pediatric populations. A remarkable recent example of patient-specific gene editing was reported by Musunuru et al. (2025), who developed a customized adenine base editing therapy (k-abe) for a neonate with carbamoyl phosphate synthetase 1 deficiency, a life-threatening urea cycle disorder characterized by hyperammonemia. Remarkably, within 6 months of the patient’s birth, the team designed, preclinically tested, and administered k-abe using LNP delivery. The therapy improved the patient’s metabolic stability, enabling increased dietary protein intake and a reduced dose of nitrogen-scavenger medication, even during episodes of infection-induced metabolic stress. Although long-term follow-up is required to fully assess safety and efficacy, this first-in-human application underscores the potential of rapid, individualized gene editing approaches for hepatic metabolic disorders. Furthermore, the use of LNPs—offering the possibility of repeat dosing—presents a significant advantage over viral vector-based therapies and may pave the way for broader clinical translation of customizable in vivo gene editing. 181

In the area of dyslipidemia, CRISPR Therapeutics has advanced gene editing therapies targeting ANGPTL3, a key regulator of plasma lipoproteins implicated in familial combined hypolipidemia and atherosclerotic cardiovascular risk, aiming to achieve durable reductions in triglyceride and cholesterol levels through genetic inactivation strategies. Furthermore, CRISPR Therapeutics is exploring gene editing for type 1 diabetes ( NCT05210530 and NCT05565248 ) and refractory dyslipidemias (ACTRN12623000809639), 182 while RenJi Hospital in China is evaluating CRISPR-based approaches for primary hyperoxaluria type 1 (PH1) ( NCT06511349 ). For PH1, recent preclinical studies using LNP-mediated CRISPR-Cas9 delivery targeting the HAO1 gene demonstrated efficient editing, significant reductions in oxalate accumulation, and sustained therapeutic effects without major off-target risks or immune responses, positioning LNP-CRISPR as a safer, one-time treatment alternative for this life-threatening disorder. 183 Collectively, these advances highlight how genome editing technologies, particularly those leveraging AAV or LNP platforms, are reshaping the therapeutic landscape for metabolic diseases, offering the potential for durable, single-administration cures where traditional treatments fall short.

Autoimmune diseases

Gene editing for autoimmune diseases is primarily centered on engineering immune cells especially B cell-targeted CAR-T cells using CRISPR-Cas9 and other platforms to achieve durable immunomodulation in treatment-resistant conditions. Early clinical trials leverage allogeneic and off-the-shelf cell products, with delivery strategies designed to overcome immune rejection and enhance long-term persistence. One promising strategy involves engineering immune cells to target key mediators of autoimmune pathology. Bioray Laboratories, for instance, is conducting phase 1 clinical trials evaluating gene-edited allogeneic CD19-targeted CAR-T cell therapy for neuromyelitis optica spectrum disorder, refractory autoimmune diseases, and systemic lupus erythematosus (SLE) ( NCT06633042 , NCT05859997 , and NCT05988216 ). 184 , 185 In one study, healthy-donor-derived CD19 CAR-T cells were genetically modified using CRISPR-Cas9 to overcome immune rejection barriers and infused into patients with severe refractory autoimmune diseases, including necrotizing myopathy and systemic sclerosis. 184 These gene-edited cells achieved complete B cell depletion within 2 weeks, persisted for over 3 months, and resulted in significant clinical improvements, such as reversal of inflammation and fibrosis, without severe adverse events like cytokine release syndrome. Similarly, an investigator-initiated trial ( NCT05988216 ) assessed allogeneic CD19 CAR-T cells (TyU19) in patients with refractory SLE. 185 Despite participants having a history of multi-organ involvement and prior neuropsychiatric lupus, treatment with gene-edited CAR-T cells led to disease remission without reactivation of central nervous system symptoms, suggesting both efficacy and a favorable safety profile. Beyond Bioray, other companies are advancing gene editing-based immune therapies for autoimmune conditions. Century Therapeutics and Fate Therapeutics are investigating induced pluripotent stem cell-derived, off-the-shelf CAR-T cell products ( NCT06255028 and NCT06308978 ). 186 , 187 These next-generation CAR-T cells, exemplified by FT819, integrate the CAR transgene into the T cell receptor alpha constant (TRAC) locus for consistent expression and include complete bi-allelic disruption of the endogenous TCR to prevent GVHD. Preclinical models demonstrated their antigen-specific cytolytic activity and sustained tumor clearance, with early clinical data supporting safety and feasibility in heavily pre-treated patients. Xuanwu Hospital in Beijing is also investigating gene-edited immune cell therapies for refractory autoimmune diseases ( NCT06485232 ), reflecting global efforts to translate gene editing tools into durable, targeted treatments for immune dysregulation. Collectively, these studies underscore the growing potential of gene editing technologies, particularly CRISPR-based engineering of CAR-T cells, to offer transformative therapies for autoimmune diseases by precisely modulating B cell-mediated pathology while addressing critical challenges such as immune rejection and treatment durability.

Muscular dystrophies

Gene editing trials for muscular dystrophies, particularly DMD, are primarily utilizing CRISPR-based in vivo strategies to restore or upregulate dystrophin expression. These approaches aim to provide durable, potentially one-time interventions, though recent studies highlight the need for cautious safety assessments due to immune-related risks. DMD is a severe X-linked recessive disorder caused by mutations in the dystrophin gene, leading to progressive muscle degeneration and premature loss of ambulation. DMD remains the most common childhood-onset muscular disease, and despite the approval of antisense oligonucleotide therapies based on exon skipping, their efficacy is limited, fueling the demand for more durable genetic interventions. 188 Cure Rare Disease, Inc. initiated a phase 1 trial ( NCT05514249 ) employing CRISPR-based gene activation technology to treat a rare dystrophin mutation in a patient with DMD. 189 The therapeutic strategy involved intravenous administration of CRD-TMH-001, a recombinant AAV9 vector encoding a CRISPR-transactivator system designed to upregulate dystrophin expression. However, the trial highlighted serious safety challenges. The sole participant developed an acute cytokine-mediated immune response, manifesting as capillary leak syndrome and progressing to severe acute respiratory distress syndrome, ultimately leading to death. Investigators attributed this severe reaction to host-specific immune factors rather than vector-specific properties, underscoring the critical need for cautious dose escalation, immune monitoring, and individualized risk assessment in gene therapy for DMD. HuidaGene Therapeutics Co., Ltd. is similarly pursuing a phase 1 clinical trial ( NCT06594094 ) investigating CRISPR-based approaches for DMD. 92 While specific outcomes from this trial are pending, gene editing is widely regarded as a promising next-generation strategy capable of correcting the underlying genetic defect, potentially offering long-term therapeutic benefits with a single administration. Nonetheless, recent analyses of gene therapy development for rare diseases highlight ongoing concerns regarding accelerated regulatory approvals, limited long-term efficacy data, and the occurrence of serious adverse events, including fatalities. 92 These findings emphasize that, while gene editing holds transformative potential, its clinical application requires robust safety monitoring, standardized trial protocols, and transparent risk communication with patients and families. Beyond DMD, Simone Spuler, MD, is leading a phase 1/2 trial for limb-girdle muscular dystrophy ( NCT05588401 ), which is similarly characterized by progressive skeletal muscle weakness. This trial reflects broader efforts to extend gene editing strategies to diverse forms of muscular dystrophy, although comprehensive biological and clinical outcomes are still forthcoming. Together, these studies illustrate both the promise and complexity of applying gene editing technologies to muscular dystrophies. While significant progress is being made toward targeting the genetic root causes, recent adverse events underscore the importance of rigorous safety evaluations, patient-specific considerations, and collaborative approaches among stakeholders to maximize therapeutic benefit while minimizing harm.

Inherited eye diseases

Gene editing and RNA-targeted therapies are rapidly expanding into ophthalmology, with early clinical trials exploring in vivo gene correction and RNA modulation to treat inherited retinal disorders and other vision-threatening diseases. These approaches focus on overcoming challenges such as large gene size, precise targeting of disease-causing mutations, and providing long-lasting alternatives to current treatments for retinal degeneration and neovascularization. Several early-phase clinical trials are underway exploring innovative approaches to address genetic defects in retinal disorders. Ascidian Therapeutics is evaluating RNA exon editing to treat ABCA4 retinopathy ( NCT06467344 ), a condition associated with mutations in large genes that exceed traditional AAV vector capacity. 190 Their approach employs trans-splicing to selectively replace mutated exons at the RNA level while preserving endogenous gene expression. This strategy allows correction of disease-causing mutations within large genes using a single-vector system, offering a potential therapeutic option for conditions previously considered inaccessible to conventional gene therapy. Editas Medicine is conducting phase 1/2 trials targeting CEP290 -associated IRDs using CRISPR-Cas9-mediated gene editing ( NCT03872479 ). EDIT-101 is delivered via subretinal injection and designed to correct the pathogenic IVS26 intronic variant of CEP290 , a major cause of Leber congenital amaurosis. Interim results from 14 participants demonstrated that EDIT-101 was well tolerated with no dose-limiting toxicities. Meaningful improvements in photoreceptor function were observed, including enhanced retinal sensitivity, best-corrected visual acuity (BCVA), mobility test performance, and vision-related quality of life in a subset of treated patients. These findings support the therapeutic potential of in vivo CRISPR-based interventions for IRD. Complementary to this, natural history studies have characterized the progression and clinical endpoints in CEP290 -related IRDs, validating BCVA, full-field stimulus testing, and mobility assessments as reliable outcome measures for future interventional trials. 191 , 192 HuidaGene Therapeutics is advancing CRISPR-based RNA-targeting strategies for neovascular age-related macular degeneration ( NCT06031727 and NCT06623279 ). 193 Their candidate HG202 employs an AAV-delivered, high-fidelity CRISPR-Cas13 system to reduce vascular endothelial growth factor A ( VEGFA ) mRNA levels and inhibit choroidal neovascularization. Preliminary data from the specific i n vivo genome-editing for hematopoiesis therapy-1 (SIGHT-I) study demonstrated promising biological and functional responses, with significant reductions in retinal fluid, improved central retinal thickness, and nearly 15-letter gains in BCVA within 8 weeks following a single subretinal injection in patients unresponsive to conventional anti-VEGF therapies. HG202 was well tolerated with no serious adverse events or dose-limiting toxicities reported to date, highlighting its potential as a long-acting, mutation-independent therapeutic alternative to current repetitive anti-VEGF injections. Other early-phase trials include investigations of gene editing for primary open-angle glaucoma by Shanghai BDgene Co., Ltd. ( NCT06465537 ) and for retinitis pigmentosa by Peking University Third Hospital ( NCT05805007 ), although specific outcome data remain pending. Collectively, these studies underscore significant advancements in ocular gene and RNA therapies, demonstrating both biological target engagement and functional improvements in select patient populations. Continued rigorous clinical evaluation is essential to validate long-term safety, durability, and efficacy, particularly given the irreversible nature of vision loss in these diseases.

Future perspectives

The future of gene editing therapeutics is marked by rapid advancements that are reshaping the treatment landscape for genetic and acquired diseases. Central to this progress is the continuous development of safe and efficient delivery platforms, including LNPs, viral vectors, and innovative non-viral technologies, which are essential for enhancing target specificity, minimizing off-target effects, and improving the overall safety profile of both ex vivo and in vivo gene editing approaches. In addition to well-established tools such as CRISPR-Cas9, ZFNs, and base editors, emerging technologies like prime editing and epigenome editors are rapidly gaining traction in preclinical and translational studies. Prime editing offers a highly versatile and precise method capable of making targeted insertions, deletions, and all twelve possible base substitutions without introducing DSBs, which may reduce off-target mutations and improve safety. Epigenome editors provide reversible control over gene expression through targeted modification of DNA methylation or histone marks, enabling therapeutic modulation of disease-associated genes without permanently altering the DNA sequence. These innovations expand the gene editing toolbox and hold promise for addressing a wider array of genetic and epigenetic disorders. Expanding the clinical applications of gene editing beyond current focuses such as hematologic disorders and cancer immunotherapy is critical to unlocking its full therapeutic potential. Ongoing and future trials are expected to increasingly target autoimmune, muscular, ophthalmic, and other genetic diseases with significant unmet medical needs. However, addressing safety concerns remains a fundamental challenge, particularly for in vivo interventions where risks such as immune reactions, insertional mutagenesis, and unintended genomic alterations must be carefully mitigated through technological improvements, optimized delivery strategies, and robust patient monitoring. Furthermore, as gene editing technologies advance toward widespread clinical use, the establishment of clear ethical frameworks and regulatory pathways is imperative to guide responsible application, ensuring both patient safety and public trust especially in the context of irreversible or germline modifications. The integration of gene editing with precision medicine strategies holds the potential to further personalize therapeutic interventions by tailoring treatments based on individual genetic profiles, thereby enhancing efficacy and minimizing adverse effects. Achieving these advancements will require sustained investment in fundamental research; refinement of gene editing tools; collaborative efforts between academia, industry, and regulatory agencies; and a commitment to long-term clinical validation. Together, these efforts are expected to drive the safe, ethical, and effective translation of gene editing into transformative, durable treatments for a broad spectrum of currently intractable diseases.

Conclusion

Gene editing has emerged as a revolutionary approach with the potential to fundamentally alter the treatment paradigm for a wide spectrum of diseases, ranging from metabolic and autoimmune disorders to muscular dystrophies and inherited eye diseases. The clinical trials currently underway reflect significant progress in translating laboratory discoveries into real-world therapies. Technologies such as CRISPR-Cas9, ZFNs, and base editors, combined with advanced delivery systems like LNPs and viral vectors, have demonstrated promising efficacy and safety profiles in early-phase studies. Despite these advances, several challenges remain before gene editing can become a routine clinical option. These include improving the precision and efficiency of gene modification; minimizing off-target effects; ensuring durable and controlled gene expression; and developing scalable, cost-effective manufacturing processes. Furthermore, long-term monitoring of treated patients will be crucial to understand the durability of therapeutic benefits and any unforeseen adverse effects.

Looking forward, future research should focus on optimizing delivery platforms to target specific tissues with higher specificity and lower immunogenicity. The integration of novel editing tools such as prime editing and epigenetic modulators could expand the scope of treatable conditions and improve safety. Additionally, combination therapies that integrate gene editing with existing treatments may enhance clinical outcomes. As the field advances, regulatory frameworks will need to evolve in parallel to ensure patient safety while facilitating rapid innovation. Collaborative efforts between academia, industry, regulatory bodies, and patient communities will be essential to overcome existing barriers.

In conclusion, gene editing holds tremendous promise to deliver durable cures and improve the quality of life for patients suffering from a range of currently intractable diseases. Continued investment in research, clinical development, and ethical considerations will be critical to unlocking the full potential of this transformative technology in the years to come.

Acknowledgments

The authors gratefully acknowledge 10.13039/501100010679 Pasteur Institute of Iran . They also thank Grammarly for assisting with language editing and improving the readability of the manuscript.

Author contributions

M.R.: preparation of the first draft of the manuscript and writing – review and editing. Z.E.: figure preparation and writing – review and editing. A.A.: writing – review and editing. F.K.-L.: conceptualization, supervision, and writing – review and editing.

Declaration of interests

The authors declare no competing interests.

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