Patients with lipopolysaccharide-responsive beige-like anchor protein (LRBA) deficiency typically suffer from severe B cell dysfunction. However, the underlying mechanisms remain incompletely understood. In this study, we identify non-muscle myosin IIA (NMIIA) as an interaction partner of LRBA in B cells, and uncover a role for LRBA in regulating actin cytoskeleton dynamics during B cell activation. LRBA-deficient B cells exhibit abnormal migration, impaired F-actin polymerization, and reduced B cell receptor signalling and polarization upon activation. In addition, LRBA deficiency severely disrupts immune synapse formation as evidenced by diminished central SMAC formation, reduced microtubule organizing center translocation and disrupted BCR and lysosome polarization. Consistent with these defects, internalization of the BCR-antigen complex is also impaired. Mechanistically, NMIIA activation, assessed by myosin light chain (MLC) phosphorylation, is reduced in LRBA-deficient cells. In addition, LRBA co-localizes with active NMIIA during both migration and immune synapse formation. Collectively, our findings establish LRBA as an important regulator of cytoskeleton dynamics during B cell activation, which may contribute to the defective humoral immunity observed in LRBA-deficient patients.

Autosomal recessive congenital ichthyosis (ARCI) refers to a group of rare, highly debilitating skin disorders that significantly impair patients’ quality of life and lack any effective treatment options. Here, we report clinically relevant in situ correction of the most common ARCI-causing mutation, TGM1 c.877-2A>G, a splice-site aberration, in human disease models. Targeted skin barrier modulation followed by topical application of the cytosine base editor eTd packaged into lipid nanoparticles yielded functional restoration of ∼30% of wild-type transglutaminase 1 activity in skin tissue. Toxicity studies and comprehensive off-target analysis demonstrated an excellent safety profile even after repeated application, without systemic distribution of the lipid nanoparticles or the genetic cargo as determined via highly sensitive methods, including desorption electrospray ionization (DESI) metabolic imaging. This study presents comprehensive preclinical data on the feasibility of in situ gene correction of genodermatoses-causing mutations, showcasing its therapeutic potential and paving the way for curative next-generation treatments for severe genetic skin diseases.

Adoptive immunotherapies have emerged as a promising strategy in the treatment of hematological malignancies. To date, seven chimeric antigen receptor (CAR)-T cell products have obtained market authorization in Europe for B-cell malignancies, where they have revolutionized treatment for eligible patients. In addition, natural killer (NK) cells and γδ T cells are of particular interest for cell-based therapies due to their strong intrinsic cytotoxicity and favorable safety profile. Addressing challenges facing the clinical translation of NK cell therapies was one of the issues discussed in the NK & ILC Symposium that took place in Freiburg, Germany from March 11 to 13, 2026, as the annual meeting of the NK & ILC study group of the German Society for Immunology (DGfI). Topics ranging from basic immunological research to technological innovations and clinical trial results were discussed during the 3-day conference, spanning over 40 presentations and 100 posters. A highlight of the conference was a workshop featuring short presentations and a panel discussion that focused specifically on the current challenges and future prospects of the clinical implementation of NK cell therapies; the findings of this workshop are summarized in this opinion paper.

Background: Chimeric antigen receptor (CAR)-T cells are therapeutic breakthroughs against advanced non-Hodgkin lymphomas and myelomas. On the other hand, no CAR-T cell product has been so far clinically approved for therapy of Hodgkin Lymphoma (HL), T cell lymphoma (TCL), or Epstein-Barr-Virus (EBV)-associated lymphoproliferative diseases (EBV-LPDs). CD30 (TNFRSF8) is commonly expressed on HL and on subsets of TCL and EBV-LPDs. CD30CAR-T cells generated via transduction with viral vectors have been tested in clinical trials, showing overall good responses against HL. CAR-T cells produced entirely with locus-specific gene editing methods are emerging as attractive next-generation engineered cell products for ease of multiple seamless cell modifications. Methods: Using CRISPR/Cas9-mediated techniques, we optimized homology-directed repair templates (HDRTs) and performed all-in-one multiplex editing to knock-in (KI) CD30CAR within the TCRα constant ( TRAC ) locus and to simultaneously knock-out (KO) PD-1 or/and β2M. CD30CAR-T cells were tested in CD30 + cell models of HL, TCL, and EBV-LPDs. Results: We compared mouse versus human anti-CD30 scFv designs in HDRTs incorporating TRAC homology arms, FcIg spacer/detection domain, and CD28 / CD3z signaling domains. We obtained an average of 30% TRAC KI CD30CAR-T cells and efficient in vitro cytotoxicity with CD30 + cell targets. CARs incorporating the high-affinity humanized 5F11 scFv showed the highest CAR expression, and the editing templates were further modified to incorporate a truncated CD34 (tCD34) spacer/detection domain. 5F11-CD30CAR-tCD34-T cells showed high CAR-KI rates (approx. 50-80% 12-14 days after editing) and potency in vitro and in vivo . Subsequently, we tested all-in-one CAR KI with additional KOs by co-electroporation of guide RNAs (gRNAs) targeting the genes encoding PD-1 or /and β2M to improve function and allow for improved cell persistence in allogeneic recipients, respectively. Compared with CD30CAR-T cells, CD30CAR-β2M KO -T cells were similarly viable and functional and showed low risk of translocations. PD1 KO enabled CD30CAR-T cells to produce higher levels of cytotoxic features upon exposure to targets. However, simultaneous β2M KO and PD-1 KO compromised the expansion capacity of CD30CAR-T cells and resulted in detectable translocations. Conclusions: Non-virally engineered 5F11-CD30CAR-T cells represent a novel cell therapy modality against CD30 + lymphomas. Multiplex editing remains to be optimized to avoid unwanted genomic alterations and chromosomal translocations.

Clinical evidence demonstrates that ex vivo gene therapy and genome engineering of hematopoietic stem and progenitor cells (HSPCs) could represent one-time cures. However, while genome editing itself has become increasingly efficient and precise, the toxic conditioning required for hematopoietic stem cell transplantation remains a major barrier to broad clinical implementation of these otherwise curative therapies. In particular, the use of busulfan for myeloablative conditioning constitutes a major safety concern. While preclinical studies established CD117 as a promising target for antigen-specific therapy, clinical translation faced setbacks balancing efficacy and safety. To overcome current limitations, we generated a new CD117-blocking monoclonal antibody (CIM058) and demonstrate its potency to block wild-type HSPCs. To enable long-term blockade of host HSPCs even after transplantation, we used prime editing to engineer CIM058-resistant human CD34+ HSPCs. When combined, CIM058 and the epitope engineered CD34+ HSPCs ameliorated disease phenotype in a β-thalassemia model. Our results suggest that this approach may overcome the reliance on busulfan or other myeloablative conditioning regimens with their associated morbidities, and by enabling toxin-free conditioning and in vivo selection of edited cells, may facilitate clinical implementation of these highly valuable genetic therapies.

Background Pathological conditions often arise from dysregulation of complex gene networks. MicroRNAs (miRNAs) are central modulators of these networks, with upregulation of disease-promoting miRNAs suppressing beneficial pathways, and downregulation of protective/therapeutic miRNAs normally restraining pathological programs. Because individual miRNAs coordinately regulate multiple genes, they represent powerful therapeutic targets. We hypothesized that pathology-associated gene expression imbalances could be corrected by placing downregulated protective/therapeutic miRNAs under the control of promoters driving overexpression of disease-promoting miRNAs, thereby simultaneously disabling pathogenic programs and inducing therapeutic ones. We termed this concept castling, after the chess move. Methods Candidate miRNA pairs for castling were identified as inversely regulated in CAR T cells at early and late stages of the chronic antigen stimulation eventually leading to their dysfunction. Proof-of-concept experiments with castling of selected miRNA clusters (knock in of miR-17~92 into a miR15/16 locus) was then implemented in both primary T cells and in CAR T cells using a newly developed genome-editing procedure TRIPLE (Targeted Replacement Induced by Persistent Locus Editing) that enhances homology-directed repair via sequential cleavage. This was followed by differential expression of miRNA and mRNAs resulting from castling compared to similarly treated non-castled corresponding control cells. In addition, castled CAR T cells were evaluated functionally in the chronic antigen stimulation assay. Results In primary T-cells, swapped expression patterns of the castled miR17~92 and miR15-16 clusters elicited expected bidirectional changes of expression of their predicted target mRNA subsets. In CAR T cells under chronic antigen stimulation, castling of these miRNA clusters delayed dysfunction, enhanced cytokine production, and reshaped transcriptional programs consistent with restored T cell fitness. This was accompanied by up- and downregulation of multiple supporting genes. Conclusions Castling, implemented via TRIPLE or any other suitable gene editing technology, offers a broadly applicable strategy to reprogram disease-driven gene regulatory networks by converting pathological regulatory loops into self-correcting circuits. As a conceptual therapeutic approach, castling may be broadly applicable for improvement of multiple types of cell therapies in a variety of indications as well as for manipulation of not only miRNA but also of protein coding mRNAs. ### Competing Interest Statement H.K., S.A., S.A.N., E.S., D.Z. are employed at Lepton Pharmaceuticals. M.H., M.P. and A.D. are employed at TAmiRNA GmbH. E.F. and C.M. are advisors to Lepton Pharmaceuticals. T.C. is an advisor to AaviGen, AstraZeneca, Cimeio Therapeutics, Excision BioTherapeutics, GenCC, and Novo Nordisk. All other authors declare no conflicts of interest. Lepton Pharmaceuticals German Federal Ministry of Research, Technology and Space (BMFTR) CRACK IT Challenge

Base editors enable precise correction of point mutations without requiring DNA double-strand breaks, yet platform-and cell type-specific genotoxicities remain incompletely characterized. Here, we applied cytosine base editing (CBE) to disrupt a cryptic splice-site mutation in the Unc13d locus of Jinx mice, a model of familial hemophagocytic lymphohistiocytosis type 3 (FHL3). Efficient editing (62%-89%) in fibroblasts, T cells, and hematopoietic stem cells (HSCs) restored Unc13d splicing, reconstituted cytotoxic T cell function, and protected mice from virus-triggered hyperinflammation after transplantation of edited HSCs. Comparative genotoxicity profiling revealed distinct platform-and cell type-specific patterns: hyperactive CBE induced broader off-target activity and more structural variants than CRISPR-Cas9. Although off-target sequence edits persisted, the stability of CBE-induced chromosomal translocations differed between cell types. These findings establish base editing as a therapeutic strategy for a genetically predisposed hyperinflammatory syndrome and underscore the importance of context-specific safety profiling to guide the clinical translation of genome editors.

Liver-directed adeno-associated viral (AAV) vector-mediated homology-independent targeted integration (AAV-HITI) by CRISPR-Cas9 at the highly transcribed albumin locus is under investigation to provide sustained transgene expression following neonatal treatment. We show that targeting the 3' end of the albumin locus results in productive integration in about 15% of mouse hepatocytes achieving therapeutic levels of systemic proteins in two mouse models of inherited diseases. We demonstrate that full-length HITI donor DNA is preferentially integrated upon nuclease cleavage and that, despite partial AAV genome integrations in the target locus, no gross chromosomal rearrangements or insertions/deletions at off-target sites are found. In line with this, no evidence of hepatocellular carcinoma is observed within the 1-year follow-up. Finally, AAV-HITI is effective at vector doses considered safe if directly translated to humans providing therapeutic efficacy in the adult liver in addition to newborn. Overall, our data support the development of this liver-directed AAV-based knockin strategy.

SYNTAXIN-11 (STX11) is a SNARE protein that mediates the fusion of cytotoxic granules with the plasma membrane at the immunological synapses of CD8 T or NK cells. Autosomal recessive inheritance of deleterious STX11 variants impairs cytotoxic granule exocytosis, causing familial hemophagocytic lymphohistiocytosis type 4 (FHL-4). In several FHL-4 patients, we also observed hypogammaglobulinemia, elevated frequencies of naive B cells, and increased double-negative DN2:DN1 B cell ratios, indicating a hitherto unrecognized role of STX11 in humoral immunity. Detailed analysis of Stx11-deficient mice revealed impaired CD4 T cell help for B cells, associated with disrupted germinal center formation, reduced isotype class switching, and low antibody avidity. Mechanistically, Stx11-/- CD4 T cells exhibit impaired membrane fusion leading to reduced CD107a and CD40L surface mobilization and diminished IL-2 and IL-10 secretion. Our findings highlight a critical role of STX11 in SNARE-mediated membrane trafficking and vesicle exocytosis in CD4 T cells, important for successful CD4 T cell-B cell interactions. Deficiency in STX11 impairs CD4 T cell-dependent B cell differentiation and humoral responses.

Undesired on- and off-target effects of CRISPR-Cas nucleases remain a challenge in genome editing. While the use of Cas9 nickases has been shown to minimize off-target mutagenesis, their use in therapeutic genome editing has been hampered by a lack of efficacy. To overcome this limitation, we and others have developed double-nickase-based strategies to generate staggered DNA double-strand breaks to mediate gene disruption or gene correction with high efficiency. However, the impact of paired single-strand nicks on genome integrity has remained largely unexplored. Here, we developed a novel CAST-seq pipeline, dual CAST, to characterize chromosomal aberrations induced by paired CRISPR-Cas9 nickases at three different loci in primary keratinocytes derived from patients with epidermolysis bullosa. While targeting COL7A1, COL17A1, or LAMA3 with Cas9 nucleases caused previously undescribed chromosomal rearrangements, no chromosomal translocations were detected following paired-nickase editing. While the double-nicking strategy induced large deletions/inversions within a 10 kb region surrounding the target sites at all three loci, similar to the nucleases, the chromosomal on-target aberrations were qualitatively different and included a high proportion of insertions. Taken together, our data indicate that double-nickase approaches combine efficient editing with greatly reduced off-target effects but still leave substantial chromosomal aberrations at on-target sites.

The CRISPR-Cas12a platform has attracted interest in the genome editing community because the prototypical Acidaminococcus Cas12a generates a staggered DNA double-strand break upon binding to an AT-rich protospacer-adjacent motif (PAM, 5′-TTTV). The broad application of the platform in primary human cells was enabled by the development of an engineered version of the natural Cas12a protein, called Cas12a Ultra. In this study, we confirmed that CRISPR-Cas12a Ultra ribonucleoprotein complexes enabled allelic gene disruption frequencies of over 90% at multiple target sites in human T cells, hematopoietic stem and progenitor cells (HSPCs), and induced pluripotent stem cells (iPSCs). In addition, we demonstrated, for the first time, the efficient knock-in potential of the platform in human iPSCs and achieved targeted integration of a GFP marker gene into the AAVS1 safe harbor site and a CSF2RA super-exon into CSF2RA in up to 90% of alleles without selection. Clonal analysis revealed bi-allelic integration in >50% of the screened iPSC clones without compromising their pluripotency and genomic integrity. Thus, in combination with the adeno-associated virus vector system, CRISPR-Cas12a Ultra provides a highly efficient genome editing platform for performing targeted knock-ins in human iPSCs.

Precise genome editing requires the resolution of nuclease-induced DNA double strand breaks (DSBs) via the homology-directed repair (HDR) pathway. In mammals, this is typically outcompeted by non-homologous end-joining (NHEJ) that can generate potentially genotoxic insertion/deletion mutations at DSB sites. Because of higher efficacy, clinical genome editing has been restricted to imperfect but efficient NHEJ-based approaches. Hence, strategies that promote DSB resolution via HDR are essential to facilitate clinical transition of HDR-based editing strategies and increase safety. Here we describe a novel platform that consists of a Cas9 fused to DNA repair factors to synergistically inhibit NHEJ and favor HDR for precise repairing of Cas-induced DSBs. Compared to canonical CRISPR/Cas9, the increase in error-free editing ranges from 1.5-fold to 7-fold in multiple cell lines and in primary human cells. This novel CRISPR/Cas9 platform accepts clinically relevant repair templates, such as oligodeoxynucleotides (ODNs) and adeno-associated virus (AAV)-based vectors, and has a lower propensity to induce chromosomal translocations as compared to benchmark CRISPR/Cas9. The observed reduced mutational burden, resulting from diminished indel formation at on- and off-target sites, provides a remarkable gain in safety and advocates this novel CRISPR system as an attractive tool for therapeutic applications depending on precision genome editing.

Abstract Background The development of the CRISPR-Cas12a platform has generated considerable interest in the genome editing community. Due to its AT-rich protospacer-adjacent motif (PAM, 5’-TTTV), Cas12a increased the potential number of targetable sites for gene editing beyond that of the prototypical Streptococcus pyogenes CRISPR-Cas9 system. Moreover, evaluation of the off-target activity of CRISPR-Cas12a nucleases suggested high specificity of the platform. Broad application of the CRISPR-Cas12a platform in primary human cells was recently enabled by the development of a re-engineered version of the natural Acidaminococcus Cas12a, called Cas12a Ultra. Methods We transferred the CRISPR-Cas12a Ultra system in the form of ribonucleoprotein complexes into clinically relevant human cells, including T cells, multipotent hematopoietic stem and progenitor cells (HSPCs), and induced pluripotent stem cells (iPSCs). Allelic gene editing frequencies were determined at various target sites using standard genotyping and next-generation sequencing. Furthermore, we evaluated targeted integration of transgenes into the AAVS1 safe harbor site and the CSF2RA locus of patient-derived iPSCs. Results We achieved allelic gene disruption frequencies of over 90% at various target sites in multiple primary human cell types. In addition, we demonstrated efficient knock-in of a GFP marker gene into the AAVS1 locus, and achieved targeted integration of a therapeutic DNA template into 90% of CSF2RA alleles in iPSCs without selection. Clonal analysis revealed bi-allelic integration in > 50% of the screened iPSC clones without compromising their pluripotency and genome integrity. Conclusions Herein, we demonstrate that the CRISPR-Cas12a Ultra system provides a highly efficient genome editing platform for human stem cell applications, expanding the toolbox for clinical applications.

Transcription activator-like effector nucleases (TALENs) are programmable nucleases that have entered the clinical stage. Each subunit of the dimer consists of a DNA-binding domain composed of an array of TALE repeats fused to the catalytically active portion of the FokI endonuclease. Upon DNA-binding of both TALEN arms in close proximity, the FokI domains dimerize and induce a staggered-end DNA double strand break. In this present study, we describe the implementation and validation of TALEN-specific CAST-Seq (T-CAST), a pipeline based on CAST-Seq that identifies TALEN-mediated off-target effects, nominates off-target sites with high fidelity, and predicts the TALEN pairing conformation leading to off-target cleavage. We validated T-CAST by assessing off-target effects of two promiscuous TALENs designed to target the CCR5 and TRAC loci. Expression of these TALENs caused high levels of translocations between the target sites and various off-target sites in primary T cells. Introduction of amino acid substitutions to the FokI domains, which render TALENs obligate-heterodimeric (OH-TALEN), mitigated the aforementioned off-target effects without loss of on-target activity. Our findings highlight the significance of T-CAST to assess off-target effects of TALEN designer nucleases and to evaluate mitigation strategies, and advocate the use of obligate-heterodimeric TALEN scaffolds for therapeutic genome editing.

While chimeric antigen receptor (CAR) T cell therapy has shown promising outcomes among patients with hematologic malignancies, it has also been associated with undesirable side-effects such as cytokine release syndrome (CRS). CRS is triggered by CAR T-cell-based activation of monocytes, which are stimulated via the CD40L–CD40R axis or via uptake of GM-CSF to secrete proinflammatory cytokines. Mouse models have been used to model CRS, but working with them is labor-intensive and they are not amenable to screening approaches. To overcome this challenge, we established two simple cell-based CRS in vitro models that entail the co-culturing of leukemic B cells with CD19-targeting CAR T cells and primary monocytes from the same donor. Upon antigen encounter, CAR T cells upregulated CD40L and released GM-CSF which in turn stimulated the monocytes to secrete IL-6. To endorse these models, we demonstrated that neutralizing antibodies or genetic disruption of the CD40L and/or CSF2 loci in CAR T cells using CRISPR-Cas technology significantly reduced IL-6 secretion by bystander monocytes without affecting the cytolytic activity of the engineered lymphocytes in vitro. Overall, our cell-based models were able to recapitulate CRS in vitro, allowing us to validate mitigation strategies based on antibodies or genome editing.

Abstract Background The development of the CRISPR-Cas12a platform has generated considerable interest in the genome editing community. Due to its AT-rich protospacer-adjacent motif (PAM, 5’-TTTV), Cas12a increased the potential number of targetable sites for gene editing beyond that of the prototypical Streptococcus pyogenes CRISPR-Cas9 system. Moreover, evaluation of the off-target activity of CRISPR-Cas12a nucleases suggested high specificity of the platform. Broad application of the CRISPR-Cas12a platform in primary human cells was recently enabled by the development of a re-engineered version of the natural Acidaminococcus Cas12a, called Cas12a Ultra. Methods We transferred the CRISPR-Cas12a Ultra system in the form of ribonucleoprotein complexes into clinically relevant human cells, including T cells, multipotent hematopoietic stem and progenitor cells (HSPCs), and induced pluripotent stem cells (iPSCs). Allelic gene editing frequencies were determined at various target sites using standard genotyping and next-generation sequencing. Furthermore, we evaluated targeted integration of transgenes into the AAVS1 safe harbor site and the CSF2RA locus of patient-derived iPSCs. Results We achieved allelic gene disruption frequencies of over 90% at various target sites in multiple primary human cell types. In addition, we demonstrated efficient knock-in of a GFP marker gene into the AAVS1 locus, and achieved targeted integration of a therapeutic DNA template into 90% of CSF2RA alleles in iPSCs without selection. Clonal analysis revealed bi-allelic integration in > 50% of the screened iPSC clones without compromising their pluripotency and genome integrity. Conclusions Herein, we demonstrate that the CRISPR-Cas12a Ultra system provides a highly efficient genome editing platform for human stem cell applications, expanding the toolbox for clinical applications.

BACKGROUND:Hemophagocytic lymphohistiocytosis (HLH) is a hyperinflammatory disorder characterized by a life-threatening cytokine storm and immunopathology. Familial HLH type 3 (FHL3) accounts for approximately 30% of all inborn HLH cases worldwide. It is caused by mutations in the UNC13D gene that result in impaired degranulation of cytotoxic vesicles and hence compromised T-cell- and natural killer-cell-mediated killing. Current treatment protocols, including allogeneic hematopoietic stem cell (HSC) transplantation, still show high mortality. OBJECTIVE:We sought to develop and evaluate a curative genome editing strategy in the preclinical FHL3 Jinx mouse model. Jinx mice harbor a cryptic splice donor site in Unc13d intron 26 and develop clinical symptoms of human FHL3 upon infection with lymphocytic choriomeningitis virus (LCMV). METHODS:We employed clustered regularly interspaced short palindromic repeats (CRISPR)-Cas technology to delete the disease-causing mutation in HSCs and transplanted Unc13d-edited stem cells into busulfan-conditioned Jinx recipient mice. Safety studies included extensive genotyping and chromosomal aberrations analysis by single targeted linker-mediated PCR sequencing (CAST-Seq)-based off-target analyses. Cure from HLH predisposition was assessed by LCMV infection. RESULTS:Hematopoietic cells isolated from transplanted mice revealed efficient gene editing (>95%), polyclonality of the T-cell receptor repertoire, and neither signs of off-target effects nor leukemogenesis. Unc13d transcription levels of edited and wild-type cells were comparable. While LCMV challenge resulted in acute HLH in Jinx mice transplanted with mock-edited HSCs, Jinx mice grafted with Unc13d-edited cells showed rapid virus clearance and protection from HLH. CONCLUSIONS:Our study demonstrates that transplantation of CRISPR-Cas edited HSCs supports the development of a functional polyclonal T-cell response in the absence of genotoxicity-associated clonal outgrowth.