Genome Editing Therapy for the Blood: Ex Vivo Success and In Vivo Prospects
Abstract
Hematopoietic stem cells (HSCs) provide the body with a continuous supply of healthy, functional blood cells. In patients with hematopoietic malignancies, immunodeficiencies, lysosomal storage disorders, and hemoglobinopathies, therapeutic genome editing offers hope for corrective intervention, with even modest editing efficiencies likely to provide clinical benefit. Engineered white blood cells, such as T cells, can be applied therapeutically to address monogenic disorders of the immune system, HIV infection, or cancer. The versatility of CRISPR-based tools allows countless new medical interventions for diseases of the blood, and rapid ex vivo success has been demonstrated in hemoglobinopathies via transplantation of the patient’s HSCs following genome editing in a laboratory setting. Here we review recent advances in therapeutic genome editing of HSCs and T cells, focusing on the progress in ex vivo contexts, the promise of improved access via in vivo delivery, as well as the ongoing preclinical efforts that may enable the transition from ex vivo to in vivo administration. We discuss the challenges, limitations, and future prospects of this rapidly developing field, which may one day establish CRISPR as the standard of care for some diseases affecting the blood.
Introduction
The emergence of CRISPR technology has signaled a significant shift in the approach to treating genetic diseases.1 In the past 10 years, rapid advancements have moved CRISPR from a laboratory technique to a promising clinical tool.2 This clinical translation has been accelerated, especially in the field of hematopoietic stem cells (HSCs) and T cells, primarily due to the feasibility of their manipulation outside of the body in an ex vivo context (Fig. 1). The pipeline for a typical ex vivo therapy involves harvesting cells, engineering them in a laboratory setting using an appropriate intracellular delivery technology, and transplanting edited cells into the patient (Fig. 2).
A major milestone in this journey was achieved in 2019 when CRISPR was employed to address severe sickle cell disease (SCD) or transfusion-dependent β-thalassemia (TDT) by targeting the erythroid-specific BCL11A enhancer in patient-derived HSCs, followed by autologous transplantation.3 As BCL11A is a transcriptional repressor of fetal hemoglobin, disruption of the BCL11A enhancer in HSCs causes increased fetal hemoglobin expression, which can in turn compensate for defective adult hemoglobin and alleviate the symptoms of diseases such as SCD and TDT.4 This therapy has been safe and effective, having been administered to over 70 patients.5 In a world first, the resulting product, marketed as Casgevy, was recently approved by the United Kingdom Medicines and Healthcare Products Regulatory Agency for the treatment of TDT and SCD, which was followed by the United States Food and Drug Administration (FDA) and the European Medicine Agency (EMA).6,7
CRISPR has also been used to engineer therapeutic T cells in the ex vivo setting. In the field of cancer immunotherapy, chimeric antigen receptor T cells (CAR-T cells) are now being augmented using CRISPR engineering, such as by removing endogenous T cell receptors (TCRs) and immune checkpoint molecules.8–10 In a first of its kind, multiplex CRISPR engineered CAR-T cells were dosed in patients with refractory cancer. Harvested patient cells were edited to knockout the TCR α- and β-chain (TRAC and TRBC) and PDCD-1, and a CAR transgene was inserted at the TCR locus. The cells were then infused back into the patient and were found to have successfully persisted for up to 9 months.8 Caribou Biosciences launched an “off-the-shelf” allogeneic CRISPR edited CAR-T cell therapy that is currently in Phase 1 clinical trials.9 Base-edited T cells have also been employed to treat patients with acute lymphoblastic leukemia. In a landmark study, two patients achieved remission after receiving base-edited, anti-CD7 CAR-T cell therapy, marking a significant advancement in cancer treatment strategies using CRISPR.10 Although beyond the scope of this review, it is noteworthy that precise engineering of B cells has shown immense promise in the deliberate manipulation of humoral immunity and other applications,11–17 offering a landscape for therapeutic intervention that is complementary to that of T cells.
Ex vivo genome editing has been clinically beneficial in several instances, but it isn’t without its challenges. In the case of therapies that rely on engineered HSCs, one primary concern is the conditioning regimen, which requires chemotherapy using harsh drugs such as busulfan or radiotherapy to deplete the patient’s HSCs to facilitate engraftment of transplanted cells. While this step is critical, it is associated with serious consequences, including leukemia risks, myelodysplastic syndrome, and infertility, emphasizing the need for thorough evaluation of patient risks and each therapy’s long-term impact.18 Recently, a trial performed by Graphite Bio, which utilized an ex vivo CRISPR homology-directed repair (HDR, or “knock-in”) approach with adeno-associated virus (AAV) serotype 6 delivery of the template to correct the SCD mutation, was paused due to a serious adverse event in the first patient dosed, which presented as prolonged pancytopenia, likely reflecting poor engraftment of the engineered cells.19–21 The same editing strategy is being pursued by Kamau therapeutics, which aims to incorporate improvements to the manufacturing pipeline, hopefully improving the likelihood of robust engraftment.22 As another example of potential complications of transplant, the lymphodepletion protocols required before administration of ex vivo manufactured CAR-T cells can lead to a risk of infection. Recently, a death due to fungal infection in a child was reported following administration of a base-edited CAR-T cell product; this tragic outcome was ascribed to the immunosuppression protocol.23
Beyond the medical complexities, there are also economic hurdles to consider. The combined costs of ex vivo genome editing, hospital stays, and associated treatments are currently priced at $2.2–3.6 million.24 This is an extreme and unattainable price for most individuals suffering from SCD, especially in countries where insurance would not cover the cost of this life-saving therapy. Furthermore, there is concern about the ex vivo culture of HSCs, which can lead to transcriptional changes with unknown impact on engraftment, stemness, and possible immune rejection. Finally, whether it is autologous or allogeneic, cell manufacture of ex vivo genome editing products is expensive, slow, and complicated, relying on sophisticated logistics, transport, cryopreservation, highly trained personnel, and dedicated facilities.25
Considering the aforementioned challenges, the field is gravitating toward in vivo delivery vectors that can preferentially edit certain cell types following systemic administration (Fig. 2), bypassing the need for ex vivo manufacture. Notable efforts include Intellia Therapeutics’ in vivo CRISPR trials for conditions such as transthyretin amyloidosis (TTR) and hereditary angioedema (HAE), both targeting the liver.26,27 Initiatives in in vivo genome editing targeting T cells include a trial by Excision BioTherapeutics, which employs an AAV vector with multiplex CRISPR targeting to facilitate the excision of integrated proviral DNA within HIV-infected reservoirs.28 Umoja Biopharma is creating CAR-T cells in vivo using lentiviral vectors (LVs), currently at the preclinical stage,29 and it has been shown that virus-like particles (VLP) could be augmented with CRISPR cargo to create edited CAR-T cells in vivo.30 In addition, in the field of HSC editing, progress has been made in vivo using lipid nanoparticle (LNP)-based delivery to HSCs31 and VLP platforms, both showing promise.30
However, with the advance of these innovative avenues, safety remains paramount. In vivo techniques, while addressing some of the challenges of ex vivo methods, introduce their own set of concerns, such as the potential for genetic changes in nontarget tissues, with the germline receiving special attention owing to risks of heritable editing.32 In particular, the lack of unique cell-type specific surface markers on stem cells heightens the challenges associated with HSC-specific targeting in vivo, and may pose an increased risk of germline cell targeting owing to the expression of stem cell markers on both cell types. Therefore, achieving precise in vivo targeting remains a major goal. In this review, we describe clinical development of genome editing in HSCs and T cells, as well as the delivery technology development that may allow the transition from ex vivo to in vivo CRISPR therapies.
HSCs
Background
Editing of HSCs is currently a major area of research for the treatment of various hematological diseases. Monogenic diseases, such as SCD, are at the forefront of this research, given their often straightforward etiology and compatibility with genetic intervention. In theory, just a single locus needs to be modified for patients to be phenotypically cured. As early as 1949, SCD gained recognition when it was categorized by Linus Pauling as the first molecular disease.33 The condition itself was first clinically described in 1910, in a patient named Walter Clement Noel, marking the initial identification of SCD in medical literature.34 It was later discovered that a single base pair mutation in the HBB (β-globin) gene causes the substitution of a negatively charged amino acid, glutamic acid, with a hydrophobic amino acid, valine. This change alters the rheological properties of hemoglobin, consequently transforming the shape and function of red blood cells, with significant physiological repercussions. It was later discovered that a single base pair mutation in the HBB (β-globin) gene causes the substitution of the negatively charged amino acid, glutamic acid, with the hydrophobic amino acid, valine.35 This mutation alters the physical characteristics of red blood cells, affecting their shape and function, impacting the rheological properties of whole blood, and leading to abnormal blood flow and increased risk of vaso-occlusive events. In β-thalassemia, a mutation at a different site within the HBB gene leads to a significant reduction or absence of the β-globin subunit of hemoglobin. This deficit disrupts hemoglobin assembly and the stability of red blood cells, resulting in ineffective erythropoiesis and increased red blood cell destruction, known as hemolysis. These processes not only lead to anemia but also cause a complex array of secondary complications. These include iron overload owing to increased gastrointestinal absorption and repeated blood transfusions, bone deformities from marrow expansion, and increased risk of infections owing to splenic dysfunction. Engineered cell therapies for these two diseases will be discussed in addition to progress being made for the treatment of other diseases of the blood.
Ex vivo genetic therapy in HSCs
Drawing on decades of expertise in bone marrow and HSC transplantation, ex vivo gene therapy has rapidly advanced during the past decade.36 A series of specialized and complex procedures, however, is required for an ex vivo HSC gene therapy protocol. First, pharmaceutical agents such as plerixafor and granulocyte colony-stimulating factor are used to induce the mobilization of HSCs and granulocytes from the bone marrow into the peripheral bloodstream, which are then collected by leukapheresis. The next step is to purify the HSCs and perform the ex vivo genetic modification in the laboratory. Before transplanting the edited HSCs, the patient undergoes a substantial conditioning regimen involving myeloablative chemotherapy to eradicate bone marrow cells, creating space for repopulation with engineered cells. Following this, the patient’s HSCs, which have been edited ex vivo, are transplanted into the patient. Because autologous ex vivo transplantation of HSCs had already been established by hematologists (including prior use for administration of engineered cell therapies), incorporation of CRISPR technology has been relatively straightforward. Hastened by this synergy, a substantial number of clinical trials have already been conducted using ex vivo HSC editing (Table 1).3,37,38
Study identifier | Status | Study title | Sponsor | Condition | Intervention | Study type |
---|---|---|---|---|---|---|
HSCs | ||||||
NCT06325709 | Recruiting | Base Editing for Mutation Repair in Hematopoietic Stem & Progenitor Cells for X-Linked Chronic Granulomatous Disease | National Institute of Allergy and Infectious Diseases (NIAID) | X-linked chronic granulomatous disease | Single infusion of base-edited (BE) autologous HSCs | Phase 1/2 |
NCT05444894 | Recruiting | EDIT-301 for Autologous Hematopoietic Stem Cell Transplant (HSCT) in Participants With Transfusion-Dependent Beta Thalassemia (TDT) | Editas Medicine | TDT | EDIT-301: autologous genome-edited HSCs-(CRISPR-Cas12a BCL11A KO) | Phase 1/2 |
NCT05662904 | Not yet recruiting | Genetic Ablation of CD33 in HSC to Broaden the Therapeutic Index of CD33-directed Immunotherapy in Patients with AML (GALAXY33) | German Cancer Research Center | Relapsed AML | GALAXY33: Donor-derived CD34+ HSC with CRISPR-Cas9-mediated CD33 deletion | Phase 1 |
NCT05951205 | Not yet recruiting | Evaluation of Efficacy and Safety of a Single Dose of Exa-cel in Participants with Severe Sickle Cell Disease, βS/βC Genotype | Vertex Pharmaceuticals | Severe SCD, βS/βC genotype (HbSC). | CTX001 (exa-cel): autologous genome-edited HSCs (CRISPR-Cas9 BCL11A KO) | Phase 3 |
NCT05397184 | Recruiting | Base Edited CAR7 T Cells to Treat T Cell M alignancies (TvT CAR7) (TvT CAR7) | Great Ormond Street Hospital for Children NHS Foundation Trust | Relapsed/Refractory CD7-positive T-ALL | Base edited allogeneic CAR T cells against CD7, KO of TCR, CD7, and CD52 | Phase 1 |
NCT04443907 | Active, Not Recruiting | Study of Safety and Efficacy of Genome-edited Hematopoietic Stem and Progenitor Cells in Sickle Cell Disease (SCD) | Novartis Pharmaceuticals | SCD | OTQ923: autologous HSC, CRISPR-Cas9-edited (BCL11A KO) | Phase 1 |
NCT04819841 | Recruiting | Gene Correction in Autologous CD34+ Hematopoietic Stem Cells (HbS to HbA) to Treat Severe Sickle Cell Disease (CEDAR) | Kamau Therapeutics (previously Graphite Bio) | SCD | Nula-cel: Autologous CD34+ Hematopoietic Stem Cells to Convert HbS to HbA for Treating Severe Sickle Cell Disease-CRISPR-Cas9 system + AAV6-based HDR template | Phase 1/2 |
NCT05456880 | Recruiting | BEACON: A Study Evaluating the Safety and Efficacy of BEAM-101 in Patients with Severe Sickle Cell Disease (BEACON) | Beam Therapeutics | SCD | BEAM-101: autologous base edited HSCs (HbF induction) | Phase 1/2 |
T cells | ||||||
NCT04502446 | Active, not recruiting | A Safety and Efficacy Study Evaluating CTX130 in Subjects with Relapsed or Refractory T or B Cell M alignancies (COBALT-LYM) | CRISPR Therapeutics AG | Refractory T or B Cell M alignancies | CTX130: anti-CD70 allogeneic CRISPR-Cas9-engineered T cells | Phase 1 |
NCT04438083 | Active, Not Recruiting | A Safety and Efficacy Study Evaluating CTX130 in Subjects with Relapsed or Refractory Renal Cell Carcinoma (COBALT-RCC) | CRISPR Therapeutics AG | Advanced, Relapsed or Refractory Renal Cell Carcinoma with Clear Cell Differentiation | CTX130: anti-CD70 allogeneic CRISPR-Cas9-engineered T cells | Phase 1 |
NCT05795595 | Recruiting | A Safety and Efficacy Study Evaluating CTX131 in Adult Subjects with Relapsed or Refractory Solid Tumors | CRISPR Therapeutics AG | Relapsed or Refractory Solid Tumors | CTX131: anti-CD70 allogeneic CRISPR-Cas9-Engineered T Cells (CTX131) | Phase 1/2 |
NCT05643742 | Recruiting | A Safety and Efficacy Study Evaluating CTX112 in Subjects with Relapsed or Refractory B-Cell M alignancies | CRISPR Therapeutics AG | Relapsed or refractory B-cell m alignancies | CTX112: allogeneic CD19-directed chimeric antigen receptor (CAR) T cell; genetically modified ex vivo using CRISPR-Cas9 | Phase 1/2 |
NCT04035434 | Active, not recruiting | A Safety and Efficacy Study Evaluating CTX110 in Subjects with Relapsed or Refractory B-Cell M alignancies (CARBON) | CRISPR Therapeutics AG | Relapsed or Refractory B-Cell M alignancies | CTX110: anti-CD19 allogeneic CRISPR-Cas9-engineered T cells | Phase 1/2 |
NCT04426669 | Recruiting | A Study of Metastatic Gastrointestinal Cancers Treated with Tumor Infiltrating Lymphocytes in Which the Gene Encoding the Intracellular Immune Checkpoint CISH Is Inhibited Using CRISPR Genetic Engineering | Intima Bioscience | Metastatic Gastrointestinal Epithelial Cancer | TILs: CISH gene inactivated w/CRISPR-Cas9 | Phase 1/2 |
NCT05722418 | Recruiting | CRISPR-Edited Allogeneic Anti-BCMA CAR-T Cell Therapy in Patients with Relapsed/Refractory Multiple Myeloma (CaMMouflage) | Caribou Biosciences | Relapsed/Refractory Multiple Myeloma | CB-011: a CRISPR-edited allogeneic anti-BCMA CAR-T Cell | Phase 1 |
NCT05566223 | Not yet recruiting | CISH Inactivated TILs in the Treatment of NSCLC (CheckCell-2) | Intima Bioscience | Metastatic NSCLC | TIL: CISH gene inactivated with CRISPR-Cas9 | Phase 1/2 |
NCT04637763 | Recruiting | CRISPR-Edited Allogeneic Anti-CD19 CAR-T Cell Therapy for Relapsed/Refractory B Cell Non-Hodgkin Lymphoma (ANTLER) | Caribou Biosciences | Relapsed/Refractory B Cell Non-Hodgkin Lymphoma | CB-010: a CRISPR-edited allogeneic anti-CD19 CAR-T cell therapy | Phase 1 |
NCT03655678 | Active, Not Recruiting | A Safety and Efficacy Study Evaluating CTX001 in Subjects with Transfusion-Dependent β-Thalassemia | Vertex Pharmaceuticals | TDT | CTX001: autologous CRISPR-Cas9-modified HSCs (BCL11A KO) | Phase 2/3 |
NCT05356195 | Recruiting | Evaluation of Safety and Efficacy of CTX001 in Pediatric Participants with Transfusion-Dependent β-Thalassemia (TDT) | Vertex Pharmaceuticals | TDT | CTX001: autologous CRISPR-Cas9-modified HSCs (BCL11A KO) | Phase 3 |
NCT05477563 | Recruiting | Evaluation of Efficacy and Safety of a Single Dose of CTX001 in Participants with Transfusion-Dependent β-Thalassemia and Severe Sickle Cell Disease | Vertex Pharmaceuticals | TDT and SCD | CTX001: autologous CRISPR-Cas9-modified HSCs (BCL11A KO) | Phase 3 |
NCT05329649 | Recruiting | Evaluation of Safety and Efficacy of CTX001 in Pediatric Participants with Severe SCD | Vertex Pharmaceuticals | SCD | CTX001: autologous CRISPR-Cas9-modified HSCs (BCL11A KO) | Phase 3 |
NCT03745287 | Active, Not Recruiting | A Safety and Efficacy Study Evaluating CTX001 in Subjects with Severe SCD | Vertex Pharmaceuticals | SCD | CTX001: autologous CRISPR-Cas9-Modified HSCs (BCL11A KO) | Phase 2/3 |
NCT04774536 | Not yet recruiting | Transplantation of Clustered Regularly Interspaced Short Palindromic Repeats Modified HSCs (CRISPR_SCD001) in Patients with Severe SCD | University of California, University of San Francisco | SCD | CRISPR_SCD00: autologous HSCs-CRISPR-Cas9 system + HDR template | Phase 1/2 |
NCT04244656 | Active, not recruiting | A Safety and Efficacy Study Evaluating CTX120 in Subjects with Relapsed or Refractory Multiple Myeloma | CRISPR Therapeutics AG | Relapsing or refractory multiple myeloma | CTX120: anti-BCMA allogeneic CRISPR-Cas9-engineered T Cells | Phase 1 |
NCT05144386 | Active, not Recruiting | Study of EBT-101 in Aviremic HIV-1 Infected Adults on Stable ART | Excision BioTherapeutics | HIV infection | EBT-101-in vivo gene therapy product intended to excise the replication-competent proviral HIV from latently infected cells | Phase 1 |
NCT05942599 | Recruiting | Base Edited CAR T Cells Against AML: Deep Conditioning Ahead of Allogeneic Stem Cell Transplantation (CARAML) | Great Ormond Street Hospital for Children NHS Foundation Trust | AML | BE CAR33 T cells | Phase 1 |
NCT05885464 | Recruiting | A Study Evaluating the Safety and Efficacy of BEAM-201 in Relapsed/Refractory T-Cell Acute Lymphoblastic Leukemia (T-ALL) or T-Cell Lymphoblastic Lymphoma (T-LL) | Beam Therapeutics | T-ALL and T-LL | BEAM-201: base-edited, allogeneic anti-CD7 CAR-T Cells | Phase 1/2 |
AML, acute myeloid leukemia; B-ALL, B cell acute lymphoblastic leukemia; HSCs, Hematopoietic stem cells; NSCLC, nonsmall cell lung cancer; SCD, sickle cell disease; T-ALL, T cell acute lymphoblastic lymphoma; TDT, transfusion-dependent beta thalassemia; TILs, tumor infiltrating lymphocytes; T-LL, T cell lymphoblastic lymphoma.
To make genetic modifications in HSCs, viral and nonviral delivery methods have been utilized (Fig. 2). Viral vectors such as AAVs and LVs have been extensively researched in the traditional gene therapy field owing to their ability to efficiently transfer genetic cargo to HSCs (Fig. 1). AAVs gained traction in the early days of gene therapy, however their use in vivo has raised concerns owing to serious and even fatal adverse effects following their in vivo administration at high doses in human patients.39,40 LVs offer an appealing option for the ex vivo delivery of functional genes to HSC, as exemplified by gene addition therapy. Bluebird Bio’s LV platform, designed to introduce the HBB gene into HSCs for treating transfusion-dependent thalassemia (TDT), received FDA approval for commercialization in 2022, and a similar product aimed at treating SCD was approved in 2023.41 Despite successful use in non-CRISPR gene therapy protocols, the application of LV vectors in the HSC genome editing field has not achieved widespread use. The ability of LV vectors to integrate into the genome could lead to persistent expression of CRISPR-Cas9 components, potentially increasing genotoxicity.42 Moreover, the risk of LV-induced genotoxicity and persistent expression is further enhanced by the inherent characteristics of the targeted cell types, which require lifelong proliferation and maintenance. This adds a layer of complexity and risk, particularly in the context of their long-term presence and activity within the body. The challenges associated with viral-based CRISPR delivery have spurred increased interest in nonviral delivery methods. These nonviral methods allow direct delivery of CRISPR Cas9 mRNA and synthetically produced guide RNA (gRNA), or CRISPR ribonucleoproteins (RNPs), usually delivered ex vivo by electroporation. This strategy allows for enhanced control over the duration of component presence in cells while also minimizing off-target impacts and toxicity.43
The development of current gene therapies for conditions such as SCD and β-thalassemia has benefited from an understanding of natural compensatory mechanisms, initially uncovered through the study of families exhibiting hereditary persistence of fetal hemoglobin (HPFH), first reported in a case study by Phaedon Fessas and George Stamatoyannopoulos in 1964.44 Several genome editing approaches have been explored with the aim of treating β-hemoglobinopathies by boosting fetal hemoglobin (HbF) expression.45 One of the most mature strategies relies on modifying regulation of the BCL11A gene via an erythroid-specific enhancer region. The BCL11A gene encodes a repressor of γ-globin, a subunit of fetal hemoglobin that fulfills a role similar to that of β-globin in adult hemoglobin (HbA). Following birth, the switch from HbF to HbA is associated with the clinical onset of SCD and β-thalassemia symptoms46 and those with SCD experience less severe symptoms if they also carry a gene for HPFH.47 By downregulating BCL11A via CRISPR-mediated editing of the gene’s erythroid-specific enhancer region, γ-globin (and thus HbF) can in turn be elevated, alleviating the symptoms of β-hemoglobinopathies.
Vertex and CRISPR Therapeutics codeveloped an ex vivo Cas9-enabled therapy known as exagamglogene autotemcel, commonly referred to as exa-cel and marketed as Casgevy. This therapy employs preassembled RNP complexes that specifically target the erythroid-specific enhancer of BCL11A, ex vivo delivered to HSCs by electroporation. Cas9 activity introduces indels (small insertions and/or deletions), leading to a disruption of the GATA1-binding site at the +58 BCL11A erythroid enhancer, increasing γ-globin expression and elevating fetal hemoglobin levels in erythroid cells.3 As BCL11A plays a role in different physiological processes in the hematopoietic system, the specificity of targeting and erythroid-specific BCL11A enhancer ensures that HSC differentiation and the function of nonerythroid lineages remain unaffected.48
During its clinical development phase, this therapy was successfully administered to 47 patients with β-thalassemia and 35 patients with SCD (both adults and adolescents). The treatment’s safety and efficacy were shown in a Phase 3, open-label study including patients aged 12–35 years who had experienced at least two severe vaso-occlusive crises (VOCs) annually in the 2 years before their participation.34 Casgevy treatment led to significant clinical improvements: the median total hemoglobin levels increased to 11.9 g/dL by 3 months post-treatment and stabilized around 12.5 g/dL by 6 months—levels that were maintained through the study period. Fetal hemoglobin levels showed a marked increase, with a mean of 36.9% at 3 months and 43.9% at 6 months post-treatment. Remarkably, 97% of the patients evaluated were free from VOCs for at least 12 consecutive months, and 100% avoided hospitalizations for such crises over the same period. These outcomes signify a substantial alleviation of the disease’s symptoms, including severe pain and related complications that typically lead to hospital admissions. The treatment was well tolerated, with a safety profile consistent with the myeloablative conditioning used. Importantly, no treatment-related malignancies or vector-related adverse events were reported. This impressive clinical result led to the approval of Casgevy, making it the first CRISPR-based therapy to receive market authorization in the United Kingdom, United States, Europe, Saudi Arabia, and Bahrain, offering a significant advancement in the treatment of SCD and β-thalassemia.7,49,50 It is important to highlight that although Casgevy received regulatory approval in the United Kingdom, it was not recommended by the National Institute for Health and Care Excellence (NICE) for routine use within the NHS without additional supportive data. NICE expressed concerns regarding the long-term efficacy and cost-effectiveness of Casgevy, highlighting the need for more comprehensive data to better understand its clinical and economic impacts.51
CRISPR-mediated introduction of indels for induction of HbF has also been explored by other studies and therapeutic candidates. In a pilot study conducted in China, two pediatric patients with β-thalassemia received treatment with autologous HSCs that were genetically modified using a CRISPR-based genome editing strategy analogous to the one employed by Vertex/CRISPR Therapeutics. Outcomes were comparable, with patients experiencing marked amelioration of symptoms, attaining transfusion independence post-treatment. Furthermore, there were no reported instances of severe adverse events attributable to the therapy and the patients were considered functionally cured.37
Other therapeutic candidates that utilize CRISPR to promote modifications by non-homologous end joining (NHEJ) to induce HbF expression in erythroid cells include Novartis/Intellia’s OTQ923, which leads to a 5 kb deletion that generates a functional HBG1–HBG2 fusion gene, and Editas Medicine’s EDIT-301 (or reni-cel), that targets the promoters of HBG1 and HBG2 using a unique variant of the Cas12a nuclease.52 Administration of the autologous therapy OTQ923 to three individuals suffering from severe SCD led to a persistent increase in HbF levels within red blood cells, ranging from 19% to 27%, along with a notable amelioration in the clinical symptoms.38 Despite the results showing that OTQ923 can successfully result in expression of HbF at levels that surpass the protective threshold of 20%, Novartis discontinued the development program for OTQ923 in February 2023, citing strategic reasons.53 Reni-cel was tested in two trials including 6 patients with SCD and 11 patients with β-thalassemia.
A second approach using CRISPR does not act on HbF induction, instead aiming to perform HDR to correct the disease-causing mutation, thus restoring functional adult hemoglobin expression. These HDR-based trials include one by the University of California (NCT04774536) and another by Graphite Bio (NCT04819841) (summarized in Table 1). Graphite’s trial consisted of a protocol that used CRISPR to target the SCD mutation, and AAV serotype 6 for delivery of the DNA template for HDR. In separate work, HDR edited cells have been shown to engraft at extremely low rates, which may be due to inefficient HDR editing in long-term HSC populations in comparison to edited progenitor cells.54 The trial was interrupted due to serious adverse events after transplantation of the edited cells. The first treated patient experienced prolonged pancytopenia which requires ongoing transfusion and growth factor supplementation, leading to speculation regarding the potential impact of AAV6 on HSC function.19,21 It has long been understood that pluripotent stem cells, such as the small population of HSCs that exists in the bone marrow, are highly resistant to viral infection.55 Differentiation and manipulation of cells has allowed for ex vivo editing strategies,56 however genome editing technologies must overcome this innate protective mechanism to efficiently alter HSCs in vivo.57 It is possible that the decrease observed in HSC function and/or capacity for engraftment after treatment with Graphite’s gene therapy product might have been a result of the proprietary conditions used to enhance HDR rates. In another unexpected outcome of this trial, the patient ultimately began producing extremely high levels of fetal hemoglobin (nearly 80% of the total hemoglobin), although the intention was to restore expression of adult hemoglobin.58 Finally, next-generation editors that do not rely on double stranded breaks—such as base editors (BEs) and prime editors—are showing promise at the preclinical stages.59 Beam Therapeutics’ clinical programs using BEs for ex vivo T cell (NCT05885464) and HSC editing (NCT05456880) are currently ongoing (Table 1).
Ex vivo genome editing has been a remarkable case of success, but the overarching procedure and associated risks remain consistent. These include adverse events that may occur during HSC mobilization in SCD patients, such as VOCs and increased pain, as well as more serious complications potentially linked to the myeloablative procedure. Long-term data on allogeneic HSC transplantation to SCD patients suggest that the conditioning process itself, often involving agents such as busulfan, could be associated with increased risk of secondary neoplasms in this population.18 In addition, the potential for off-target effects of gene-editing tools and the expansion of pre-existing clonal hematopoiesis must also be considered as potential contributors to malignancy risk. Even though ex vivo HSC editing has been successful in clinical trials for the treatment and functional cure of SCD and β-thalassemia, the risks of developing cancer and infertility owing to the conditioning process motivate the development of treatments that avoid this step. Moreover, the financial and logistical burdens of ex vivo genome editing significantly impede access for many patients.24
In vivo genome editing of HSCs
Direct in vivo genome editing of HSCs is still in its infancy: only a handful of published articles have attempted this feat, attaining limited success thus far (Table 2). It has proven to be a significant challenge, as targeting, delivery, distribution, editing efficiency, and cell survival are all critical factors that must be optimized for in vivo editing. It remains a major obstacle to target the HSCs, a rare population within the bone marrow estimated to consist of only 50,000–200,000 cells that are active in human hematopoiesis at any given time.60 Achieving a targeted, safe, and effective in vivo delivery of genome editing technologies to HSCs could pave the way for treating several hematological disorders and expand access to genetic therapies by lowering both economic and practical barriers. Delivery platforms utilized for in vivo delivery of editing cargo into HSCs can be broadly classified as either viral or nonviral methods (Fig. 2).
Cell type | Target | Delivery platform | Main findings | Ref |
---|---|---|---|---|
HSCs | β-globin gene repair via nucleotide excision or HDR (homology-dependent repair) | Triplex-forming γPNA carried by nanoparticles with SCF treatment | Up to 7% β-globin gene correction in HSCs. | 80 |
HSCs | CRISPR-Cas9-mediated disruption of a BCL11A binding site | HSC mobilization followed by IV injection of CD46-targeting HDAd virus | Target site cleavage in HSCs resulted in reactivation of γ-globin expression in up to 13% of RBCs | 61 |
HSCs | Erythroid BCL11A enhancer (AncBE4max platform) | Transduction of mobilized HSCs with adenovirus HDAd5/35++ | Generation of the −113A>G HPFH mutation at a conversion rate of 20% in HSCs of β-YAC mice, leading to >40% γ-globin+ peripheral RBCs. This marking rate was maintained in secondary recipients. | 61 |
HSCs | ABE8e to the BCL11A binding site in the HBG1/2 promoters | HDAd5/35++ vector expressing ABE8e to install a −113A >G HPFH mutation in the γ-globin promoters | >60% −113A >G conversion, with 30% γ-globin of β-globin expressed in 70% of erythrocytes. | 62 |
HSCs | β-globin gene repair via prime editing | In vivo non integrating, prime editor—expressing HDAd5/35++ vectors into mobilized CD46/Townes mice | On average, 43% of sickle hemoglobin was replaced by adult hemoglobin, thereby greatly mitigating SCD phenotypes | 114 |
HDAd, helper-dependent adenovirus; HPFH, hereditary persistence of fetal hemoglobin; HSCs, Hematopoietic stem cells; RBCs, red blood cells; SCF, stem-cell factor; γPNA, gamma-substituted peptide nucleic acid.
Viral delivery to HSCs in vivo
Lieber and colleagues have researched helper-dependent adenovirus (HDAd) for the delivery of DNA cassettes encoding cytosine and adenine BEs.61,62 The aim is for BEs to be expressed in HSCs and introduce edits that downregulate BCL11A. This two-component platform delivers a BE expressing cassette in the first HDAd, plus a gene to enhance cell proliferation of the edited cells in the second HDAd.62 Vector capacity is a limitation that must be overcome when delivering Cas9 variants to cells. In this case, two vectors were employed to accommodate the desired payload.63 Although it has proven to be safe in nonhuman primates, there is controversy regarding the safety of adenoviral vectors in humans, and it is unclear whether this approach will be advanced to clinical trials.64 Recently, another study utilized capsid-engineered adenovirus vector to target cells that express CD46 or DSG2 receptors and laminin-interacting α6β1, α6β4, α3β1, and α7β1 integrin classes, and showed that the vector could transduce HSC-enriched bone marrow populations 1,900-fold more efficiently than to CD34-negative bone marrow cells, which suggests a degree of selectivity.65
Nonviral delivery to HSCs in vivo
The remaining vectors for in vivo delivery are nonviral. Although they are in relatively early stages of development, they suggest a path toward safe delivery to HSCs within the body, broadening access to genome edited HSC therapies.
Lipid nanoparticles
Of the several nonviral platforms being developed for cell-targeted delivery of CRISPR reagents in vivo, LNPs have been the focus of much attention and development. LNPs were deployed globally for delivery of the spike protein mRNA in the Moderna and Pfizer COVID-19 vaccines, a success story that has inspired widespread optimism for the platform’s potential to enable genetic therapies.66 LNPs are composed of ionizable lipids whose chemical properties change depending on the pH of their environment. At acidic pH, ionizable (cationic) lipids associate with the anionic RNA cargo, and upon neutralization the hydrophobic domains assemble to form RNA-bearing LNPs. Once the LNPs have entered cells via endocytosis, the low pH environment of the late endosome promotes ionization of the lipids, causing LNP destabilization as well as endosomal escape, the key bottleneck in cytosolic delivery of macromolecular cargo.67
The LNP platform has shown substantial promise for the delivery of CRISPR components (e.g., mRNA encoding Cas9 protein and a synthetic gRNA), and underlies the encouraging trials conducted by Intellia. In these two trials, CRISPR-loaded LNPs have been used in vivo for delivery to the liver targeting either the transthyretin locus to treat TTR26 or the kallikrein B1 locus to treat HAE.68 Not only has one company been able to repurpose the same delivery platform rapidly to enable a second trial but LNP compositions have been used with BEs targeting the liver suggesting versatility in enabling hepatic genome editing.69,70 The strength of liver tropism may also hinder broader use of LNPs: their propensity to accumulate in the liver has posed a challenge when attempting to target other parts of the body, including HSCs within the bone marrow. There has been encouraging progress using LNPs for delivery of CRISPR reagents to cultured HSCs, resulting in efficient knockout or knock-in with low toxicity and minimal transcriptomic impact.71 However, using the same reagents for HSC editing in vivo may prove difficult because nontargeted and targeted LNPs alike have the tendency to accumulate in the liver72 and LNPs may be toxic at the doses required for editing other organs.73 It has been shown that the rate of protein synthesis in HSCs is lower than in other stem cells, which suggests the necessity of greater doses of mRNA—and thus LNP—to meet editing thresholds.74 An important issue to be considered for the development of LNP-based in vivo genome editing products is the emerging evidence of antiplatform immunity, which was observed in the case of PEG antibodies that arose following repeated administration of COVID-19 vaccines.75
In vivo editing of HSCs using LNPs is limited by the physical barrier of the bone marrow endothelium, necessitating high levels of LNP in the blood and in turn increasing risks of toxicity. It may be possible to attain clinically beneficial levels of LNP-mediated HSC-targeted genome editing in vivo—while avoiding toxicity—by altering particle composition, increasing dosage, and/or implementing novel strategies to alter particle tropism.76 This latter aspect has been explored using “endogenous” targeting that relies on serum proteins to influence the fate of the IV-administered LNP,77 whereas “active” targeting involves decorating the particles with functional groups promoting homing to HSCs. A promising recent advance implemented the latter strategy to enable genetic modification of HSCs using LNPs.76 In this work, LNPs were decorated with an anti-CD117 antibody that promotes targeting of bone marrow HSCs, successfully delivering Cre recombinase to HSCs. After intravenous administration in a fluorescent reporter mouse, LNPs encoding Cre recombinase altered the genome of >50% of murine HSCs in vivo. In a separate nonhuman primate study, CD90-targeted LNPs for HbF reactivation showed great promise for long-term repopulation and engraftment of HSCs.78,79 HSC-targeted LNPs were also shown to target human HSCs ex vivo, enabling highly efficient CRISPR-mediated base editing in SCD donor HSCs, resulting in near-complete reduction of sickling.
Peptide nucleic acids
One noteworthy approach involves nuclease-free editing via peptide nucleic acids (PNA). PNAs are synthetic oligonucleotides with a similar structure to DNA and they possess the ability to bind complementary sequences in the genome. However, instead of a sugar-phosphate backbone, the nucleotide bases are linked with a pseudopeptide polymer, rendering them uncharged and more flexible than their natural counterpart. In a recent application addressing β-thalassemia, PNAs complementary to the mutated β-globin gene were delivered via LNPs. Upon reaching the nucleus, the PNAs promote formation of a nucleic acid triplex, with the IVS2-654 (C > T) mutated region sandwiched between two single-stranded PNAs.80 This high-affinity DNA hybridization event leads to excision and repair of the single-nucleotide polymorphism via the cells’ innate homology-dependent repair (HDR). Unlike nucleases that may cut at off-target sites and/or induce chromosomal damage, PNAs rely on merely strand invasion, resulting in specific genome modification with minimal risk of genotoxicity. Unfortunately, low editing rates associated with PNAs are a hurdle that must be overcome to achieve clinically relevant levels of genome editing.
T Cells
Ex vivo gene therapy in T cells
Decades of effort have enabled cell-based therapies for the treatment of cancer via immunotherapy, including recent trials featuring CRISPR-mediated engineering (Table 1). In an increasingly common immuno-oncology strategy, a patient’s T cells are engineered ex vivo to introduce genes encoding CAR that preferentially target proteins present on the surface of cancer cells, releasing cytokines to promote an anti-cancer immune response.81 Such CAR-T cell therapies have revolutionized the treatment of certain cancers by equipping a patient’s own immune cells to attack cancerous cells.48 Engineered CAR-T cells were originally produced by delivering exogenous receptor genes via γ-retroviral vectors or LVs.82 Initial trials using γ-retroviral vectors for T cell manufacture showed promise but were ultimately sidelined owing to the oncogenic nature of some vector insertions.83 More recent efforts rely on LVs for the integration of exogenous CAR genes into T cells. The resulting cell therapies have been widely successful for immuno-oncology, especially for cancers of the blood, but there is a substantial practical limitation: the transgenes can be integrated at random sites in the genome, including oncogenes.84 The location and effects of these unpredictable insertions cannot be robustly characterized and pose risks to patients.83 Indeed, in November 2023 the FDA announced the investigation of potential risks of secondary cancer linked to CAR-T cell therapy.85 In January 2024, the FDA issued a directive to producers of CAR-T therapies, mandating the incorporation of warnings on the packaging of these products to alert consumers to the potential for increased cancer risks linked to the therapies.
CRISPR genome editing technology can address the practical limitations of virally-engineered T cell therapies, which typically feature an exogenous CAR or TCR transgene that imparts anti-cancer activity. Semi-random integration of the exogenous CAR or TCR typically results in unnatural and/or highly heterogeneous expression, which underlies inferior function as compared to precise, CRISPR-mediated insertion of the transgene into the site of its endogenous analog.86,87 Targeting the TRAC locus to insert the CAR transgene has also been shown to enhance tumor rejection.86 Furthermore, the targeted insertion of a CAR transgene into the programmed cell death protein 1 (PD1) genomic locus via a non-viral delivery system exemplifies a strategic application of CRISPR/Cas9 genome editing. This approach not only ensures site-specific transgene integration but also simultaneously disrupts the PD1 gene, a modification that is hypothesized to enhance the therapeutic efficacy of the resultant CAR-modified T cells by potentially mitigating inhibitory signals during immune responses.88 This strategy reached the clinic in a Phase I study for the treatment of 21 non-Hodgkin’s lymphoma patients, with promising results.89
In addition to improving the transgene incorporation step, there is value in augmenting a LV-engineered T cell therapy with the knock-out of other genes, a process that can be facilitated by versatile genome editing technology. CRISPR-Cas9 was also used to disrupt three genes (TRAC, TRBC, and PDCD1), combined with LV-mediated insertion of a tumor-specific TCR to improve antitumor immunity in patients with refractory cancer.8 Targeting immune checkpoint molecules such as PD1 has been shown to improve CAR-T cell persistence, whereas elimination of endogenous TCR is a necessary step for TCR-engineered T cell therapies of to generate allogeneic CAR-T Cells while preventing alloreactivity.87 A first-in-human Phase I clinical trial demonstrates the feasibility and safety of nonviral precision genome engineering of a personalized adoptive T cell transfer anticancer therapeutic, where the two endogenous TRAC genes were knocked out while two chains of a neoantigen-specific TCR (neoTCR) isolated from circulating T cells of patients were knocked in.90 Companies such as Caribou Biosciences are combining both types of CRISPR engineering, pairing knock-out and knock-in edits to produce cells with extensive modification such as precise transgene incorporation and immune stealthing. Such cells are deployed as “off the shelf” allogeneic cell therapies—manufactured from one donor but administered to distinct recipients—and have shown encouraging potency in initial clinical trials. Allogeneic base-edited anti-CD7 CAR-T cells incorporating TRBC, CD7, and CD52 knockouts were recently shown to induce remission in a patient with T-Cell acute lymphoblastic leukemia.10
Established methods of genome editing have revolutionized ex vivo immuno-oncology cell therapies. Some of the earliest progress made in T cell engineering was done using zinc-finger nucleases (ZFNs) to knockout the CCR5 gene in T cells and HSCs to make them HIV resistant.91 HIV-resistant T cells were autologously administered in a clinical trial (NCT00842634) as a form of preventative treatment against HIV infection. This trailblazing success story has been followed by advances in genome editing, together paving the way for more than a dozen anti-HIV cell therapies.92
Precise manipulation of the genome using the CRISPR-Cas9 programmable nuclease system has made it possible to integrate necessary DNA templates efficiently, as well as disrupt unneeded genes for both research and therapeutic uses.8 CRISPR is a promising method for improved CAR-T cell engineering and has recently been used to directly knock out innate TCR genes.8 This approach eliminates redundant endogenous TCR gene expression and addresses the long-standing complications associated with endogenous and exogenous TCR chain mispairing.63 CRISPR-Cas9 RNPs targeting genes of commonly mispaired receptor chains (and/or other undesirable genes) are typically delivered to T cells ex vivo via electroporation.8,86 In addition to TRAC/TRBC gene knockout to prevent protein mispairing, precision editing can also knock out the PDCD1 gene encoding programmed cell death protein (PD-1), thereby preventing suppression of the T cell inflammatory response and increasing efficiency of tumor elimination,8 or the beta-2-microglobulin (B2M) gene, which is increasingly being used as a reporter site or to disrupt the major histocompatibility complex.93–95 Following genome editing, LV was used to introduce an exogenous CAR transgene. This early demonstration of complex, multistep editing resulted in an efficient—but highly heterogeneous—multiplex knockout in T cells. A similar study has performed multiplex editing paired with lentiviral transgene incorporation, but supplanted nucleases with BEs, which do not rely on double-strand breaks (DSBs) in the genome and minimize the genotoxicity risks associated with nuclease-mediated DSBs at multiple genomic loci.93
Issues caused by semi-random lentiviral-mediated transgene delivery (discussed above) can be resolved by using CRISPR-Cas9 to induce site-specific gene knock-ins. Electroporation of CRISPR RNP, paired with a DNA-based transgene template to facilitate HDR, has enabled precise non-viral genome editing of T cells.96 Genome editing provides an alternate route for CAR-T cell manufacture, improving cell function and homogeneity of engineered cells. CRISPR RNP delivery was also used to repair a loss-of-function mutation in the IL-2 gene, demonstrating proof-of-concept for correction of a T cell-based disease.96 This marked a step forward for non-viral genome editing, as it was previously thought that delivery of large (>1 kb) dsDNA was toxic to cells. Marson and team’s technique works by delivering a preformed Cas9 RNP alongside a large dsDNA template to T cells ex vivo via electroporation, resulting in the integration of the functional genes of interest. In both cases, transplantation of the edited T cells resulted in clinically relevant levels of T cell response in vivo.96
Genome editing enables the creation of highly sophisticated immuno-oncology products, but autologous cell therapy strategies still face a substantial manufacturing bottleneck because a patient’s own cells must be used. Several teams and biotech companies are working to circumvent this issue by developing batches of “off the shelf” CAR-T cells for allogeneic immuno-oncology therapies.97 Early work on this idea relied on transcription activator-like effector nucleases (TALENs) to edit T cells, which are administered to nongenetically similar patients as a cell therapy.97,98 This method of T cell engineering was done by lentiviral transduction of a CAR gene targeting the CD19 antigen and TALEN-mediated disruption of the CD52 and TRAC genes. This resulted in anti-CD19 CAR-T cells resistant to graft-versus-host-disease,97 rendering them capable of allogeneic administration.
As we look toward the future direction of cell-derived cancer treatments, developing allogeneic CAR-T cells presents a more cost-effective, efficient, and streamlined manufacturing process than the individualized autologous treatments currently being administered. However, allogeneic CAR-T cells must be deliberately engineered via genome editing to render them tolerable to recipients. This can be done using multiple edits in the genome using CRISPR-Cas9. CRISPR-derived BEs could offer advantages in scenarios calling for multiplex editing, since they do not rely on double-stranded DNA breaks and thus minimize the risk of chromosomal translocations.99 Feasible therapeutic allogeneic CAR-T cell therapy may very likely be on the horizon, as biotechnology company, Caribou Biosciences, has developed an allogeneic T cell therapy that is showing promise in human patients.9 The ability to select from readily available CAR-T cells, based on a patient’s therapeutic needs—and without enduring a substantial manufacturing delay—may present a valuable option in tackling cancer.
In vivo genome editing of T cells
Ex vivo genome engineering of CAR-T cells is widely successful, but efforts are underway to manipulate cells entirely in vivo, potentially opening the door for in situ generation of CAR-T cells, the treatment of immune diseases, or protection of T cells from infections such as HIV. Early experiments with genome editing in animal models utilized viral vectors for the delivery of CRISPR components, enabling gene transduction via self-replication, extrachromosomal duplication, or integration into the host’s genome.100 However, any traditional viral delivery approach is associated with persisting enzyme expression and activity, heightening the risk of unintended genetic alterations.101
Viral delivery for T cells in vivo
Vectors based on viruses, such as AAV, adenoviruses, and lentiviruses (LVs), are frequently utilized for the delivery of CRISPR systems (Fig. 2). There are three steps to ex vivo gene transduction into T cells: 1) isolation, 2) activation (a necessary step owing to the paucity of low-density lipoprotein receptors needed for LV targeting102), and 3) transduction via a LV pseudotyped with the glycoprotein G of the vesicular stomatitis virus (VSV). The activation step poses cellular risks and complications but can be eliminated if in vivo editing is made possible.103 Developments have been made to modify the surface proteins on LVs to increase specificity to T cells to enable in vivo targeting.103 In vivo genome editing could bypass procedures related to cell isolation, manufacturing, and transplantation, potentially making the technology more accessible and less costly. Buchholz and colleagues delivered the anti-CD19 CAR gene to lymphocyte cells via an engineered LV containing CD8 or CD4 cell-surface glycoproteins within the viral envelope, allowing them to preferentially target and enter CD8+ or CD4+ T cells, respectively. This platform was administered via intravenous injection to generate anti-CD19 CAR-T cells in vivo,104 which led to the elimination of CD19+ tumor cells in preclinical models. Interestingly, by incorporating CD3-specific single-chain variable fragments (scFvs) on NiV-based LVs, it was shown that it’s possible to simultaneously activate T cells and achieve targeted gene delivery in vivo. These CD3-tagged vectors facilitate gene transfer into nonactivated T lymphocytes efficiently in vitro, including within human whole blood, without the need for any external activation signals, and also generated CAR-T cells in vivo in humanized mice.105
CRISPR genome editing machinery has often been delivered in vivo using AAV vectors, thanks in part to the in vivo clinical track record established in traditional gene therapy. AAV versatility also has been demonstrated with the delivery of CRISPR components targeting different organs, including the brain.106 A clinical trial led by Excision BioTherapeutics aims to assess the safety and efficacy of in vivo AAV-mediated CRISPR delivery to target HIV reservoirs.28 These reservoirs are typically T cells that express CD4 among other markers. CRISPR enzymes can recognize sequences unique to HIV within these cells, targeting the virus for removal while leaving the host genome intact. In a preclinical study in nonhuman primates, a single intravenous dose of the AAV9-delivered CRISPR cargo demonstrated proof-of-concept for the anti-HIV therapy, and is currently under clinical investigation.28 As part of a distinct study, it was recently shown that utilizing AAV2 vectors that display both mono- and bispecific designed ankyrin repeat proteins (DARPins) targeting CD4 and CD32a can enhance the precision of genome editing in targeting HIV reservoir T cells in vivo, presenting a new approach that could significantly improve the specificity and safety of in vivo genome editing.107
Nonviral delivery to T cells in vivo
Nonviral delivery methods that have the potential to be utilized for in vivo applications include LNPs and VLPs, sometimes referred to as enveloped delivery vehicles (EDVs). VLPs that encapsulate CRISPR RNP complexes offer a novel approach to cell-specific genome editing. This innovation leverages the fusion of a Cas protein to the Gag polyprotein during the production of VLPs, enhancing the packaging of these complexes. VLP production typically relies on mammalian cells for production and assembly, and subsequent purification for use in research or clinical trials, which may be subject to the same bottlenecks associated with lentiviral manufacture.108,109 A significant advancement in targeting specificity is achieved by incorporating an scFv derived from a CD19-targeting antibody, fused to the CD8a transmembrane domain, into the VLPs. This method aims to exploit the natural budding process of VLPs from plasma membranes for the delivery of CRISPR-Cas9 components. By coexpressing this scFv fusion with a modified VSV-G protein and essential lentiviral elements, the resulting Cas9-VLPs recognize specific T cell-surface markers, enabling generation of genome-edited CAR-T cells in vivo, in humanized mice.30
LNPs have been employed to deliver modified mRNA directly to T cells, facilitating in vivo creation of antifibrotic CAR-T cells that persisted transiently and led to a reduction in fibrosis and improved cardiac function in a mouse heart failure model.110 LNPs can be used to deliver the CRISPR cargo as either RNP or mRNA with sgRNA. By changing the LNP molar composition with supplemental molecules to alter its internal charge, it is possible to influence which cells are targeted in vivo. This technology, named selective organ targeting (SORT), enables the engineering of nanoparticles for precise delivery of various cargoes such as mRNA and CRISPR components to specific organs.111 A similar approach using noncationic LNPs carrying RNPs has also been employed in vivo, facilitating genome editing in the lungs and liver, marking a significant step toward systemic in vivo genome editing applications.112
Trends, perspectives, and future directions
With the first CRISPR therapies reaching the market, it is natural to speculate on the ultimate impact of this emerging clinical modality. How broadly can these medicines be deployed in terms of both clinical need and practical patient access?
CRISPR-based therapies have provided immense benefit to dozens of patients, producing robust, predictable results in cases where delivery is not a major barrier. This is especially apparent in the context of ex vivo manufacture of CRISPR-enabled cell therapies wherein laboratory hardware is used for efficient and predictable intracellular delivery and clinicians can employ well-established routes of administration to transplant the engineered cells. Ex vivo manufacture is especially well-suited for CRISPR since laboratory culture supports cell division, a requisite for efficient HDR or prime editing that may not be attainable in vivo. Indeed, ex vivo cell engineering allows researchers and clinicians to leverage the entire CRISPR toolbox, potentially enabling curative therapies for countless genetic diseases, many of which are debilitating or fatal and completely devoid of treatment options. Although CRISPR technology is inherently “plug & play,” therapies for rare (or ultra-rare) diseases will only be possible with breakthroughs in regulatory policy and strategies to cover the associated costs. Ex vivo manufacturing of a given CRISPR therapy deftly sidesteps certain delivery barriers, but the requisite cell transplantation will still limit access for logistical reasons. For rare diseases or sophisticated anticancer cell therapies requiring specific forms of editing that may not be feasible without cell culture, relatively accessible ex vivo therapy may eventually take place at centers of excellence that provide custom CRISPR-engineered cells via point-of-care manufacture.113 The significant manufacturing costs associated with the current state of the art in ex vivo genome editing reinforces the need for innovations that enhance accessibility. Exploring nonviral delivery methods, which offer substantial advantages over viral vectors, is particularly promising, with ample opportunities for the development of more accessible ex vivo products, setting the stage for eventual progression to in vivo applications.
Systemic, cell-targeted delivery of CRISPR reagents in vivo—ideally via a one-time intravenous infusion—represents an ideal solution for deployment of genetic therapies. This route of administration would eliminate the cell manufacturing costs, patient burden, and other practical challenges associated with ex vivo CRISPR therapies. Substantial progress is continuously being made toward in vivo delivery to T cells and HSCs, with the latter representing the greater challenge owing to their anatomical location: bone marrow is more difficult to access than the spleen. Identifying delivery platforms that can reach—and edit—the relevant cells is just one of the challenges that must be overcome before in vivo CRISPR therapies will be feasible for blood-associated cells. To date, the field has struggled to engineer a delivery platform with pronounced tropism for the targeted cell type. Even purportedly HSC-targeted reagents often result in predominant delivery to the liver, raising safety concerns. Furthermore, cells within the body are much more likely to be quiescent, narrowing the functional CRISPR toolkit to only approaches that work well in non-dividing cells (i.e., nucleases, BEs, and epigenome editors, as opposed to prime editors and many knock-in strategies). Finally, there are trade-offs when considering delivery platforms. Viral platforms are potent, but the capsids can be challenging to manufacture and the associated long-term expression of CRISPR cargo is associated with increased genotoxicity and immunogenicity. Nonviral platforms such as LNPs sidestep many of these problems, but to date predictable targeting has been elusive and LNPs may cause acute toxicity at the doses needed for delivery to non-liver tissues. A hybrid approach is embodied by VLPs, which can offer an appealing combination of cell-type tropism, potency, minimal toxicity, and transient expression of CRISPR cargo, although these particles face manufacturing challenges that are undeniably virus-like.
People living with β-hemoglobinopathies now have new therapeutic options thanks to CRISPR, and anticancers have similarly benefitted from genome editing, but deploying CRISPR therapy to millions is currently out of reach. Fortunately, a monumental and multifaceted global effort is underway to expand the impact of CRISPR therapy for diseases of the blood, and access is likely to improve in coming years with the development of improved in vivo delivery technology.
Acknowledgment
The authors thank Dr. Clarissa Lima e Moura de Souza, hematologist, for critically reviewing the article.
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Copyright 2024, Mary Ann Liebert, Inc., publishers.
History
Published in print: October 2024
Published online: 26 September 2024
Accepted: 15 July 2024
Received: 16 April 2024
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C.A.G., L.O., and R.C.W.: Conceived the article. C.A.G., S.U.S., L.O., B.S.F.S., and R.C.W.: Wrote and edited the article. C.A.G., S.U.S., B.S.F.S., and R.C.W.: Created the figures.
Author Disclosure Statement
S.U.S. and R.C.W. are named inventors on a patent application related to this field. R.C.W. is a co-founder of Editpep, Inc.
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This work was supported by the Innovative Genomics Institute, D’Or Institute and Pioneer Science Initiative. Figures were generated with the assistance of Sarah Pyle.
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