ABSTRACT Oncogenic fusion drivers are common in hematological cancers and are thus relevant targets of future CRISPR-Cas9-based treatment strategies. However, breakpoint-location variation

in patients pose a challenge to traditional breakpoint-targeting CRISPR-Cas9-mediated disruption strategies. Here we present a new dual intron-targeting CRISPR-Cas9 treatment strategy, for

targeting t(8;21) found in 5–10% of de novo acute myeloid leukemia (AML), which efficiently disrupts fusion genes without prior identification of breakpoint location. We show in vitro growth

rate and proliferation reduction by 69 and 94% in AML t(8;21) Kasumi-1 cells, following dual intron-targeted disruption of _RUNX1-RUNX1T1_ compared to a non t(8;21) AML control.

intron-targeting CRISPR-Cas9 treatment strategy in AML t(8;21) without need for precise knowledge of the breakpoint location. SIMILAR CONTENT BEING VIEWED BY OTHERS BIOLOGICAL SIGNIFICANCE

OF MONOALLELIC AND BIALLELIC _BIRC3_ LOSS IN DEL(11Q) CHRONIC LYMPHOCYTIC LEUKEMIA PROGRESSION Article Open access 09 July 2021 IN VIVO CRISPR/CAS9 TARGETING OF FUSION ONCOGENES FOR

SELECTIVE ELIMINATION OF CANCER CELLS Article Open access 08 October 2020 _RUNX1_ MUTATIONS IN BLAST-PHASE CHRONIC MYELOID LEUKEMIA ASSOCIATE WITH DISTINCT PHENOTYPES, TRANSCRIPTIONAL

PROFILES, AND DRUG RESPONSES Article Open access 11 August 2020 INTRODUCTION Acute myeloid leukemia (AML) is characterized by proliferation of undifferentiated myeloid cells. A sizeable

fraction of AML cases is characterized by recurrent cytogenetic aberrations as well as genetic mutations affecting genes involved in the hematopoiesis. Among these, core-binding factor (CBF)

AMLs are characterized by balanced translocations affecting the transcription factor CBF subunits and are divided into two subtypes; inv(16)(p13.1q22)/t(16;16)(p13.1;q22) and

t(8;21)(q22;q22.1) (AML t(8;21)), resulting in the fusion genes _CBFB-MYH11_ and _RUNX1-RUNX1T1_ [1]. These subtypes, accounting for an estimated 15% of de novo AML cases, are associated

with a favorable prognosis compared with other AML subtypes with an estimated 5-year overall survival of 60–70% [2,3,4]. While first-line treatment consisting of high-dose anthracycline and

cytarabine with the possibility of adding gemtuzumab ozogamicin, will usually result in more than 90% of patients achieving a complete remission [5, 6], there is an unmet for novel

strategies, as an estimated 30% of patients will experience a relapse [7, 8]. Disease monitoring is essential for identifying primary treatment response and early relapse, which is

accomplished by determining measurable residual disease (MRD) primarily by monitoring the disease-defining translocations in peripheral blood and bone marrow [9]. Furthermore, as recent

advances in the treatment of AML have primarily benefitted the younger patient population [10], there is a collective need for novel treatment strategies with low inherent toxicity that can

target MRD in order to prevent relapse and perhaps act as bridging to allogeneic hematopoietic stem cells transplantation. Just as important, such a venture might allow for more efficient

cytoreduction in the elderly patients without compromising safety [11]. In animal models, _RUNX1-RUNX1T1_ has been shown to be independently insufficient to instigate AML t(8;21)

leukemogenesis, and it is believed that additional genetic aberrations are necessary in order to cause AML [12]. However, as _RUNX1-RUNX1T1_ has been shown to both inhibit the

differentiation of the hematopoietic cells as well as aid in evasion of apoptotic cell death, the fusion gene is hypothesized to constitute a key factor in the maintenance of the leukemic

cell population [13,14,15], making _RUNX1-RUNX1T1_ an attractive putative target for medical intervention [16]. Novel therapies for AML are continuously being introduced to the treatment

regimens for AML patients, with Bcl-2, _FLT3_ and _IDH1/2_ inhibitors representing the new wave of available therapies. In addition, progress is being made in CAR-T cell based treatment

modalities. However, a major challenge yet to be overcome is identifying AML specific antigens that prevent CAR-T mediated damage of healthy cells [17]. In spite of these recent treatment

advances, AML remains a malignancy with few available treatment options and a high overall mortality [18]. Gene editing using the clustered regularly interspaced palindromic repeats

(CRISPR)-Cas9 offers the possibility to target pre-defined DNA sequences in human cells [19,20,21]. Recent advances in the CRISPR-Cas9 technique have demonstrated both feasibility and

efficaciousness in treatment of transfusion-dependent β-thalassemia, sickle cell disease, aromatic L-amino acid decarboxylase (AADC) deficiency and transthyretin amyloidosis

[22,23,24,25,26], demonstrating the potential power of CRISPR-Cas9 methodologies in precision medicine. We hypothesize that a novel dual intron-targeting CRISPR-Cas9 methodology [27] can be

used to disrupt _RUNX1-RUNX1T1_ without damaging wild type _RUNX1_ and _RUNX1T1_. By targeting two meticulously selected intron regions, flanking the fusion breakpoint, it is possible to

induce major CRISPR-Cas9-mediated gene-disrupting deletions, without knowing the precise location of the breakpoint. Double strand break repair by non-homologous end-joining (NHEJ) joins

together exons with out-of-frame codons following CRISPR-Cas9 cleavage causing a shift in the fusion gene reading frame resulting in a premature stop codon and thereby non-functional

oncogene. This dual intron-targeting approach negates the need for precise knowledge of the fusion breakpoint location in patients offering a gene therapy solution without a need for

preceding fusion breakpoint sequencing. The only genetic information needed to qualify for treatment would be the standard diagnostic t(8;21) identification, which can be available less than

72 h after the diagnosis is made. In this proof-of-principle study, we demonstrate that the _RUNX1-RUNX1T1_ can be targeted and disrupted utilizing a dual intron-targeting

CRISPR-Cas9-mediated strategy elucidating this novel methodology’s potential in future treatment of AML t(8;21) patients. METHODS CELL LINES AND PATIENT SAMPLES The human CBF AML cell line

Kasumi-1 (ATCC, Manassas, VA, USA), positive for t(8;21)(q22;q22.1) was used for _RUNX1-RUNX1T1_ disruption experiments, with AML cell line THP-1 (ATCC), which does not harbor the t(8;21)

translocation, as negative control and the immortalized human dermal fibroblast cell line, MJ26146 as non-malignant control. Cells were cultured in RPMI-1640 Medium, with L-glutamine and

sodium bicarbonate (MERCK, Burlington, MA, USA) supplemented with 20% fetal Bovine Serum (FBS) (GIBCO, Thermo Fisher Scientific, Waltham, MA, USA) and 1% Penicillin-Streptomycin (GIBCO) at

37 °C and 5% CO2. Archival mononuclear cells, preserved in DMSO in liquid nitrogen from a peripheral blood sample or bone marrow sample collected as part of routine diagnostic workup from

four patients diagnosed with CBF AML with t(8;21)(q22;q22.1) at the Department of Hematology at Aarhus University Hospital, Denmark were utilized (Supplementary Table 1). Informed consent

and ethical approval were waived due to the anonymous and proof-of-concept nature of this study (the Central Denmark Region Committees on Health Research Ethics, reference: 186/2017 and the

Danish Data Protection Agency, reference: 727067/1-16-02-173-21). CRISPR-CAS9 GENE EDITING Synthetic guide RNA molecules (sgRNAs) were designed to target intron regions in the

_RUNX1-RUNX1T1_ fusion gene (Synthego, Menlo Park, CA, USA) (Supplementary Table 2 and Fig. 1). Two sgRNAs were targeted against the _RUNX1_ intron region between exon 4 and 5 and further

two sgRNAs were targeted against the intron region between exon 1a and 2 in _RUNX1T1_. The sgRNA target sites did not contain common single nucleotide polymorphisms ensuring robust target

sites with minimal potential patient-to-patient Cas9 cleavage efficiency variation. Target sites were also placed in intron regions without functions in splicing or other regulatory

mechanisms. Ribonucleoprotein (RNP) complexes comprising sgRNAs and Streptococcus pyogenes Cas9 (spCas9) (Integrated DNA Technologies, Coralville, IA, USA) in a 2.6:1 ratio were preassembled

on ice. Kasumi-1 and THP-1 cells were adjusted to desired concentration in Opti-MEM (Thermo Fisher Scientific) prior to transfection. Transfection was performed in 25 µl reactions in

electroporation strip tubes (Lonza Basel, Switzerland) by electroporation (CM138, 4D-NucleofectorTM X, Lonza). Controls treated with Cas9 without sgRNAs were included. The same CRISPR-Cas9

approach was used to target the _RUNX1-RUNX1T1_ in mononuclear cells from the patient. The CRISPR-Cas9-mediated disruption of the _RUNX1-RUNX1T1_ fusion gene was validated using targeted PCR

and Sanger sequencing (Mix2Seq, Eurofins, Luxembourg). The NGS based off-target analysis was performed using a custom ampliseq panel designed to cover the target sites for the RX2 and RXT1

guides (on-target) as well as the top 48 and 42 off-target sites, respectively, predicted by CRISPRoff (v1.2beta) software [28,29,30]. The panel was designed for standard DNA (275 bp) to

maximize the coverage of the top off-target sites for each guide resulting in coverage of the two on-target sites as well as 90 off target sites. The libraries were made using the AmpliSeq™

Library PLUS for Illumina with AmpliSeq™ UD Indexes for Illumina® according to manufacturer’s protocol. The quality of the libraries was checked using a bioanalyzer and the concentration

determined using qubit and sequenced using an NextSeq 500 with the NextSeq 500/550 High Output Kit v2.5 (300 Cycles). The bioinformatic off-target analysis was performed using BWA MEM for

alignment and crispRvariants for mutation efficiency calculations [31]. POLYMERASE CHAIN REACTION DNA was purified from cells using the QIAamp DNA Mini Kit (QIAGEN, Hilden, Germany)

according to manufacturer’s instructions. Ten to 100 nanograms of input DNA and primers targeting intron regions surrounding sgRNAs in _RUNX1_ and _RUNX1T1_ (MERCK) (Supplementary Table 2)

were added to a 20 µl PCR reaction with DreamTaq DNA Polymerase and buffer (Thermo Fisher Scientific) according to manufacturer’s instructions. Targets were amplified on a thermal cycler as

following: (1) Initial denaturation at 95 °C for 3 min, (2) 40 cycles of denaturation at 95 °C for 30 s, (3) annealing at 58 °C for 30 s, (4) extension at 72 °C for 1 min, (5) final

extension at 72 °C for 7 min. The PCR products were visualized by capillary electrophoresis using the QIAxcel Advanced System (QIAGEN) and agarose gel electrophoresis. QX DNA Alignment

Marker 15 bp/1 kb (QIAGEN) was used for PCR product size estimation. CELL GROWTH MONITORING Following sgRNAs:Cas9 treatment, Kasumi-1 and THP-1 cells were transferred to 48 well-plates in

triplicates and cultured in RPMI-1640 (MERCK) supplemented with 20% FBS (GIBCO) and 1% Penicillin-Streptomycin (GIBCO) at 37 °C, 5% CO2. On experiment days 2, 4, 6, 8, 10 and 12 cells were

taken out, mixed with Solution 18, AO•DAP (Chemometec, Allerod, Denmark) and counted in NC slide A8 (Chemometec) using the NucleoCounter NC-250 (Chemometec). CELL PROLIFERATION Following

sgRNAs:Cas9 treatment, Kasumi-1 and THP-1 cells were split into triplicates and recovered for 24 h before staining with CellTrace Violet Cell (CTV) Proliferation Kit (7.5 µM, Invitrogen)

according to manufacturer’s protocol. On experiment days 2, 4, 6, 9 and 11, 50,000 cells were stained with Zombie NIR viability dye (BioLegend, San Diego, CA, USA) before flow cytometry

analysis (NovoCyte 3000RYB with 13 detectors (ACEA Biosciences Inc., San Diego, CA, USA. Agilent, Santa Clara, CA, USA)). Theoretically, the signal intensity will half by each proliferative

cycle. Median fluorescence intensity (MFi) values were compared for treatment/control samples to determine the change in proliferation following treatment. Data was acquired using

NovoExpress version 1.5 (ACEA Biosciences, Inc.) and analyzed using FlowJo version 10.7.1 (BD, Ashland, OR, USA). CELL SORTING Following sgRNAs:Cas9 treatment, Kasumi-1 cells were recovered

for 24 h before staining with CTV (7.5 µM, Invitrogen, Thermo Fisher Scientific) according to manufacturer’s protocol. On day 10 after staining, 8 µg/ml propidium iodide (PI, BD Bioscience,

San Jose, CA, USA) was added to the cell suspension. Immediately after, cells were sorted on a FACSAria III equipped with four lasers, 405, 488, 561 and 633 nm (BD Bioscience). The

definition of CTV high and CTV low subpopulations were based on peak MFi values (Supplementary Figs. 1–4). Area scaling was 0.6, a 100 µm nozzle and 20 psi was used. Cells were sorted and

collected at 4 °C. Data were collected using FACSDiva software version 8.0.2 (BD Bioscience). RT-QPCR _RUNX1-RUNX1T1_ gene expression was measured in triplicates in purified RNA from

Kasumi-1 cells, THP-1 cells, and cells from peripheral blood and bone marrow of two patients diagnosed with AML t(8;21), respectively. The assay was performed in accordance with current

clinical guidelines as previously described [32, 33]. ANIMAL STUDIES Six BALB/cAnNRj-Foxn1 nu/nu mice were injected with CRISPR-Cas9 _RUNX1-RUNX1T1_-disrupted Kasumi-1 cells or

Cas9-control-treated Kasumi-1 cells in the right and left flank, respectively. Animals were sacrificed 4 weeks after injection and tumor volume was evaluated using the caliper method and the

formula; _V_ = (_W_2 × _L_)/2. STATISTICS All estimates are reported with a 95% confidence interval (CI) and _p_ values < 0.05 were considered statistically significant. Two-means

comparisons for continuous variables were performed using Student’s _t_ test or the Wilcoxon–Mann–Whitney test depending the distribution of the data. Multiple-means comparisons for

continuous variables were performed using a two-way ANOVA with Geisser-Greenhouse correction. Prism version 8.2.1 (GraphPad Software, Inc. San Diego, CA, USA) was used for statistical

analysis as well as design of figures. Regression for analysis of covariance was conducted using Stata version 15.1 (StataCorp LLC, College Station, TX, USA). RESULTS DUAL INTRON-TARGETING

CRISPR-CAS9 DISRUPTION OF THE ONCOGENIC DRIVER _RUNX1-RUNX1T1_ LEADS TO EFFECTIVE INHIBITION OF AML T(8;21) CANCER CELL GROWTH AND PROLIFERATION IN VITRO We hypothesized that a

_RUNX1-RUNX1T1_ disruption in the AML t(8;21) cell line, Kasumi-1, would lead to reduced proliferation and cell population growth. Four sgRNAs were designed to target intron regions at

various sites flanking the _RUNX1-RUNX1T1_ breakpoint in pairs. By using this dual-guide CRISPR-Cas9-mediated approach in Kasumi-1, we were able to induce a major deletion in _RUNX1-RUNX1T1_

involving exons following exon 4 of _RUNX1_ and exons preceding exon 2 of _RUNX1T1_. By this, the function of the fusion gene was disrupted both by generation of frame shift in _RUNX1T1_

and through deletion of functional domains in _RUNX1_ (Fig. 1 and Supplementary Fig. 5). Different combinations of the four guides were tested and all sgRNA pairs were able to produce

_RUNX1-RUNX1T1_-disruption through major deletions (Fig. 2A). Potential off-target effects were examined by targeted sequencing of 48 and 42 possible off-target sites for RX2 and RXT1

sgRNAs, respectively. No major off-target events were observed (Fig. 3A–C and Supplementary Figs. 6–9). On-target efficiencies for RX2 sgRNA were 23%, 38%, 90%, and 83% and for RXT1 sgRNA

were 4%, 13%, 81%, and 47%, for Patient 1, Patient 3, THP-1, ad Kasumi-1, respectively (Fig. 3). To investigate whether the disruptions affected Kasumi-1 cell population growth, the number

of cells was determined at different time points through 12 days following the CRISPR-Cas9-mediated _RUNX1-RUNX1T1_ disruption for all sgRNA combinations. All sgRNA pairs caused a

significant reduction in cell population growth in Kasumi-1 by a median of 69.4% (range 61.0–74.5%) (_p_ < 0.0002 (range 0.0002–<0.0001)) (Fig. 2B). As expected, no difference in

growth was observed in the THP-1 control cell line (Fig. 2C) as well as no significant difference was observed between the reduced population growth among the different sgRNA pairs in

Kasumi-1 cells (Fig. 2B). Due to similar performance on _RUNX1-RUNX1T1_ disruption and cell-population growth reduction in Kasumi-1, sgRNA pair (RX2-RXT1) was used for all for further

investigations. To investigate whether the observed reduced cell growth was due to a reduced proliferation rate in the _RUNX1-RUNX1T1_-disrupted Kasumi-1 cells, the proliferation was

monitored, day 2 through day 11. Proliferation was observed as a reduction in CTV dye intensity over time. Two subpopulations in the Kasumi-1 population treated with RX2-RXT1 were observed

on day 9 and 11 (Fig. 4A). In silico sorting of the subpopulations on day 11, showed that the CTV low subpopulation had a 2.8% (_p_ = 0.993) decreased proliferation rate and the other CTV

high subpopulation a 94.6% (_p_ < 0.0001) decreased proliferation rate compared to the non-disrupted Kasumi-1 cells control (Fig. 4B). Flow cytometry mediated cell sorting on basis of CTV

signal intensity followed by PCR showed that the highly proliferative subpopulation (CTV low) had a smaller proportion of cells with effectively disrupted _RUNX1-RUNX1T1_ compared to the

less proliferative subpopulation (CTV high), explaining the difference in proliferative potential within the RX2-RXT1 treated Kasumi-1 cells (Fig. 4C). Overall, we observed a gradual

reduction in proliferation rate over the culture period following the _RUNX1-RUNX1T1_ disruption in the Kasumi-1 cells (Fig. 4D). THP-1 cells treated with the _RUNX1-RUNX1T1-_disrupting

sgRNAs RX2-RXT1 showed a non-significant 29% increase in proliferation rate on day 11, compared to untreated THP-1 cells (Fig. 4E) showing that dual intron-targeting CRISPR-Cas9

_RUNX1-RUNX1T1_ disruption only affects Kasumi-1, cells harboring _RUNX1-RUNX1T1_, and not THP-1 cells which do not have the targeted fusion gene. Together, these results show that dual

intron-targeting CRISPR-Cas9-mediated disruption of the oncogenic driver _RUNX1-RUNX1T1_ leads to a profound inhibition of AML t(8;21) cancer cell growth and proliferation in vitro. DUAL

INTRON-TARGETING OF _RUNX1-RUNX1T1_ CAN INDUCE A GENE FUSION IN CELL LINES NOT CARRYING THE FUSION GENE We detected trace amounts of the disrupted _RUNX1-RUNX1T1_ with a similar

_RUNX1-RUNX1T1_ sequence in THP-1 to the one induced in Kasumi-1 following treatment with RX2-RXT1 (Supplementary Fig. 10a) indicating, as expected, a CRISPR-Cas9 cleavage activity in both

_RUNX1_ and _RUNX1T1_ introns in THP-1 cells. We monitored the THP-1 cells for a detectable cleavage product via gel electrophoresis for 6 days following treatment. The translocation

continued to be detectable after 6 days (Supplementary Fig. 11). However, given that we detected no functional (i.e., non-disrupted) _RUNX1-RUNX1T1_ sequence in the RX2-RXT1-treated THP-1

cells, we infer from these data that the presence of the disrupted _RUNX1-RUNX1T1_ sequence could be due to an event caused by the CRISPR-Cas9-mediated disruption of _RUNX1_ and _RUNX1T1_ in

THP-1 followed by faulty NHEJ repair, commonly characterizing cancer cell lines. This was confirmed by sequencing showing a similar break point to the one induced in Kasumi-1 following

treatment with RX2-RXT1 (Supplementary Fig. 10b). We performed RT-qPCR on the RX2-RXT1-treated THP-1 cells and did not detect any _RUNX1-RUNX1T1_ mRNA (data not shown). In order to evaluate

the risk of generating this gene fusions in healthy cells, we then investigated RX2-RXT1 mediated disruption in a fibroblast cell line, MJ26146. We did indeed observe the presence of a

translocation between chromosomes 8 and 21 corresponding to the disrupted _RUNX1-RUNX1T1_ fusion. The fusion continued to be detectable after 10 days of observation (Supplementary Fig. 11).

These data suggest that induction of translocations in healthy cells when using dual sgRNAs targeting introns is a legitimate risk when using CRISPR-Cas9, and should be investigated further

in future research. _RUNX1-RUNX1T1_ DISRUPTION AND EXPRESSION DOWNREGULATION IS FEASIBLE IN PRIMARY AML CELLS FROM T(8;21) POSITIVE PATIENTS Next, we wanted to evaluate whether the

disruption observed in the cell model applies to patient AML cells. To this end, we performed CRISPR-Cas9-mediated _RUNX1-RUNX1T1_ disruption utilizing our in vitro setup with the four

sgRNAs (RX1, RX2, RXT1, RXT2) and showed the specific target site variations in a pre-therapeutic blood sample from one patient (Patient 4) diagnosed with CBF AML with t(8;21)(q22;q22.1).

The _RUNX1-RUNX1T1_ disruption was confirmed with PCR (Fig. 5A) and Sanger sequencing (Supplementary Fig. 12). The remaining three patients (Patient 1, 2, and 3) were _RUNX1-RUNX1T1_

disrupted using only RX2 and RXT1. We showed significant downregulation of _RUNX1-RUNX1T1_ expression following CRISPR editing in two of the patients (Fig. 5B) thus providing evidence that

the CRISPR-Cas9-mediated method for fusion gene disruption is feasible also in patient-derived AML t(8;21) cancer cells. These data thus suggest that this new approach is also feasible for

targeting primary human cancer cells. _RUNX1-RUNX1T1_ DISRUPTION DECREASES TUMOR VOLUME IN VIVO Lastly, we wanted to evaluate whether the Kasumi-1 population growth reduction following

_RUNX1-RUNX1T1_ disruption would also translate to reduced tumor growth in vivo. _RUNX1-RUNX1T1_-disrupted Kasumi-1 cells were injected into the right flanks of six immunodeficient nude

mice. Each mouse also received an injection of non-disrupted Kasumi-1 cells into the corresponding left flank as control. Over 4 weeks, two out of six mice developed tumors in both flanks.

Further two mice developed tumors, but only in the non-disrupted control flank. The last two mice did not develop tumors in either flank (Fig. 6A). In the two mice with tumor development in

both flanks, tumor volumes were decreased by 69 and 91% in the _RUNX1-RUNX1T1_-disrupted flank, as compared to the control flank (Fig. 6B). These results support our in vitro findings and

show that disruption of _RUNX1-RUNX1T1_ disruption leads to a decrease in tumor volume also in vivo. DISCUSSION There is a collective need for novel treatment strategies with low inherent

toxicity that can target MRD in AML t(8;21) in order to prevent relapse, offer efficient cytoreduction in the elderly patients without compromising safety and perhaps act as bridging to

allogeneic hematopoietic stem cells transplantation. In this proof-of-principle study, we have demonstrated that dual intron-targeting CRISPR-Cas9-mediated disruption of _RUNX1-RUNX1T1_

leads to a reduction of chimeric _RUNX1-RUNX1T1_ fusion transcripts and a significant decrease in AML t(8;21) leukemic tumor cell proliferation and growth both in vitro and in vivo. We have

shown the feasibility of using standardized sgRNAs to disrupt the _RUNX1-RUNX1T1_ in the Kasumi-1 cell line, in both an in vitro setting, to reduce tumor population size and proliferation,

and in an in vivo mouse model where _RUNX1-RUNX1T1_ disruption led to a tumor-volume reduction. We also proved able to induce disruption of _RUNX1-RUNX1T1_ in AML t(8;21) patient-derived

cells showing the feasibility of CRISPR-Cas9-mediated gene editing of primary leukemic cells. While the literature has described a second hit as necessary for development of leukemia in the

presence of _RUNX1-RUNX1T1_ [12, 34,35,36,37,38,39], our findings confirm the reported _RUNX1-RUNX1T1_ dependence of AML t(8;21) cells for sustaining the leukemic cell population [13,14,15,

40]. Collectively, our data suggests that dual intron-targeting CRISPR-Cas9-mediated _RUNX1-RUNX1T1_ disruption represents a potential new future therapy modality for AML patients with

t(8;21)(q22;q22.1), a gene therapy solution without a need for preceding fusion breakpoint sequencing as the only genetic information needed to qualify for treatment would be the standard

diagnostic t(8;21) identification. As described by Martinez-Lage et al. [27], a feasible CRISPR-Cas9-mediated genome editing strategy to specifically disrupt fusion gene driven cancers

should ensure that (1) the sgRNAs are designed to target only intron regions, avoiding disruption of exon regions and thereby affect expression of wild type alleles and (2) that the sgRNA

target sites encompasses all patient specific breakpoints and fusion gene isoforms. We adhered to these criteria by locating our sgRNAs in intron regions, without single nucleotide

polymorphisms, that encompass most known clinically relevant breakpoints in AML t(8;21). Furthermore, the location of our target sites ensured a disruption of the essential _RUNX1_ Runt

domain, responsible for DNA binding and protein-protein interactions of both the native _RUNX1_ and the fusion protein _RUNX1-RUNX1T1_, thus not relying solely on frameshift causing

deletions to disrupt _RUNX1-RUNX1T1_ activity, making the disruption strategy more robust. Initially, multiple sgRNAs were used to target the _RUNX1-RUNX1T1_ fusion gene in leukemic cells in

vitro and in vivo and showed equal efficacy. Given the individual efficacy of all sgRNAs used in this study, we expect this approach to offer robust targeting of the fusion gene as well the

potential variations in individual patient sequences, thus offering the possibility of a “one size fits all” approach to future gene therapy of AML t(8;21). In the flow cytometry-based

cell-proliferation study, we observed two subpopulations in Kasumi-1 cells treated with sgRNAs RX2-RXT1. Sorting of the two subpopulations on basis of proliferative signal followed by a PCR

specific for the CRISPR-Cas9-disrupted _RUNX1-RUNX1T1_ indicated that the two subpopulations were a result of imperfect delivery or low efficiency of ribonucleoprotein (RNP) complexes. One

subpopulation had a lower fraction of disrupted _RUNX1-RUNX1T1_ as compared to the other and showed similar proliferative potential as the Kasumi-1 cells treated without sgRNAs (Fig. 4C). At

least two factors could explain this observation, first, the efficiency of the individual sgRNA in inducing deletions at the target site would indeed affect how large a fraction of the

Kasumi-1 cells could be expected to harbor a disrupted _RUNX1-RUNX1T1_ fusion gene following electroporation. A second factor is the efficiency of the electroporation-based delivery utilized

in this study which could theoretically be amended by developing a more efficacious method for delivery of CRISPR-Cas9 to the target cells. We speculate that both factors contribute to the

two subpopulations observed in our study when analyzing with flow cytometry. Future research efforts should be allocated toward quantification of the efficiency of individual sgRNA as well

as optimizing delivery of CRISPR-Cas9 to leukemic cells. Even though delivery of CRISPR-Cas9 components was previously achieved through electroporation by Frangoul et al. [22] in a

transplant setting, this approach is not likely to be feasible in the setting of AML treatment as transplantations are allogeneic and not autologous. This clinical challenge necessitates

further investigation of in vivo delivery of CRISPR-Cas9-mediated treatments in a clinical setting. An alternative to electroporation-based delivery when utilized as patient treatment could

be an adeno-associated virus (AAV) derived vector with high affinity for the hematopoietic cells, such as the AAV serotype 6 [24, 41]. Generally, AAVs have been shown to have a high

transfection efficiency which makes them candidates for clinical CRISPR-Cas9 delivery [26, 42]. A challenge with AAVs could arise if patients require multiple administrations, as adaptive

mechanisms of the patients immune system, such as neutralizing antibodies or T-cells, can hinder multiple treatment cycles [43]. However, work is being done to engineer AAVs to evade immune

responses [44] to overcome this particular challenge. Another delivery strategy, lipid nanoparticles, have been shown to instigate a low immune response potentially allowing for repeated

treatment cycles with CRISPR-Cas9 [25, 45]. Speculatively, based on the clinical efficacy of anti-CD33 targeting in AML t(8;21) using gemtuzumab ozagamicin [46], a vector carrying the

CRISPR-Cas9 payload, e.g. an AAV or a lipid nanoparticle, targeted toward CD33+ cells could be pursued in order to ensure high fidelity toward leukemic cells while potentially sparing

healthy cells. Future research should aim to clarify the potential of different delivery approaches and their transduction efficacy in AML. Ultimately, the potential success of

CRISPR-Cas9-mediated fusion gene disruption as a treatment for AML will depend on the ability to deliver the technological components to the leukemic cells efficiently in vivo. We detected

an unexpected effect of Cas9-cleavage in the THP-1 cell line leading to the formation of a t(8;21) translocation identical to the CRISPR-Cas9-disrupted _RUNX1-RUNX1T1_ fusion in the Kasumi-1

cell line (Supplementary Fig. 10). However, no fusion transcript mRNA (data not shown) or growth inhibition could be detected in the in vitro studies (Fig. 2C) and only a non-significant

increase in proliferation was observed in the cell population harboring a subpopulation with the disrupted _RUNX1-RUNX1T1_ translocation (Fig. 4E). The findings were confirmed in a

fibroblast cell line demonstrating that the observed fusion was not due to an inherent deficiency of the DNA repair pathway in the leukemic cell line. We suggest that the disrupted fusion

gene in THP-1 cells was a product of error prone DNA repair mechanisms such as NHEJ and Microhomology-mediated end joining (MMEJ) which have been shown to be upregulated and implicated in

both primary and secondary development of fusion-driven leukemias increasing the risk of chromosomal translocations following CRISPR-Cas9 induced double strand breaks [47,48,49]. However,

nothing in our data suggested the formation of a neo-oncogene in the THP-1 cells as the gene fusion is similar to the disrupted (i.e., non-functional) _RUNX1-RUNX1T1_ gene fusion created in

the Kasumi-1 cells following treatment with CRISPR-Cas9 (Supplementary Fig. 10b) [50]. Our results clearly indicate that the technology is associated with the risk of generating gene fusions

in the non-targeted cells. As such, future research should carefully evaluate the implications of these fusions in murine models. In summary, the dual intron-targeting CRISPR-Cas9

technology can effectively inhibit proliferation and decrease tumor volume in AML t(8;21). However, potential side effects include induction of translocations whose potential effect on

healthy cells need to be rigorously investigated preclinically. While the current data are intriguing, future studies focusing on strategies for in vivo delivery of the technology,

benchmarking the efficacy in combination with currently approved treatments along with off-target effects studies will be necessary for this technology to reach AML t(8;21) patients. DATA

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PubMed  PubMed Central  Google Scholar  Download references ACKNOWLEDGEMENTS We thank Anne Laugaard Thielke and Trine Skov Petersen, Aarhus University and Henriette Rugholm Petersen, Aarhus

University Hospital for technical assistance and the FACS Core Facility, Aarhus University, for access to and assistance with the flow cytometry assays as well as the Hematoloy Laboratory at

Department of Hematology, Aarhus University Hospital, Denmark for access to archival BM/PB samples. Thanks to Marie Beck Iversen, Aarhus University, and the animal facilities at the

Department of Biomedicine, Aarhus University animal work assistance. Thanks to Christopher Nordentoft Vejgaard and Anita Kruse at Aarhus University Hospital for RT-qPCR technical assistance

and data interpretation. Finally, the group thanks Sebastian Haunstrup Reimick for aiding with experimental work as part of the review process. The work was supported by grants from AUFF

NOVA (Aarhus University) to CKH, the Karen Elise Jensen Foundation to PH and ML and the Danish Cancer Society to PH, Health Research Foundation of Central Denmark Region to MB and

Tømrermester Jørgen Holm og Hustrus Mindelegat to JFS. FUNDING Open access funding provided by Aarhus University Hospital. AUTHOR INFORMATION Author notes * These authors contributed

equally: Signe Neldeborg, Johannes Frasez Soerensen. * These authors jointly supervised this work: Maja Ludvigsen, Christian Kanstrup Holm. AUTHORS AND AFFILIATIONS * Department of

Pathology, Aarhus University Hospital, Aarhus, Denmark Signe Neldeborg, Kasper Holm & Magnus Stougaard * Department of Clinical Medicine, Aarhus University, Aarhus, Denmark Signe

Neldeborg, Johannes Frasez Soerensen, Boe Sorensen, Peter Hokland, Magnus Stougaard & Maja Ludvigsen * Department of Hematology, Aarhus University Hospital, Aarhus, Denmark Johannes

Frasez Soerensen, Marie Bill & Maja Ludvigsen * Department of Biomedicine, Aarhus University, Aarhus, Denmark Charlotte Thornild Møller, Zongliang Gao, Rasmus O. Bak, Mette Nyegaard, 

Yonglun Luo & Christian Kanstrup Holm * Department of Clinical Biochemistry, Aarhus University Hospital, Aarhus, Denmark Boe Sorensen * Department of Health Science and Technology,

Aalborg University, Aalborg, Denmark Mette Nyegaard * Steno Diabetes Center Aarhus, Aarhus University Hospital, Aarhus, Denmark Yonglun Luo * Danish Life Science Cluster, Copenhagen, Denmark

Magnus Stougaard Authors * Signe Neldeborg View author publications You can also search for this author inPubMed Google Scholar * Johannes Frasez Soerensen View author publications You can

also search for this author inPubMed Google Scholar * Charlotte Thornild Møller View author publications You can also search for this author inPubMed Google Scholar * Marie Bill View author

publications You can also search for this author inPubMed Google Scholar * Zongliang Gao View author publications You can also search for this author inPubMed Google Scholar * Rasmus O. Bak

View author publications You can also search for this author inPubMed Google Scholar * Kasper Holm View author publications You can also search for this author inPubMed Google Scholar * Boe

Sorensen View author publications You can also search for this author inPubMed Google Scholar * Mette Nyegaard View author publications You can also search for this author inPubMed Google

Scholar * Yonglun Luo View author publications You can also search for this author inPubMed Google Scholar * Peter Hokland View author publications You can also search for this author

inPubMed Google Scholar * Magnus Stougaard View author publications You can also search for this author inPubMed Google Scholar * Maja Ludvigsen View author publications You can also search

for this author inPubMed Google Scholar * Christian Kanstrup Holm View author publications You can also search for this author inPubMed Google Scholar CONTRIBUTIONS SN, JFS and ML drafted,

wrote and revised the manuscript. PH, MB, ML, MS and CKH provided input for revision of the manuscript. SN, CTM, MN, YL, PH, ML, MS and CKH conceived and designed the study. SN, CTM, YL, ZG,

BS and ML wrote and executed protocols for the experimental work. SN, JFS, CTM, YL, ZG, KH, ML, MS and CKH performed data analysis and interpretation. ML and MB acquired patient cells. SN

and JFS share first autorship. ML and CKH share last authorship. All authors have approved the final version and agreed to be accountable for all aspects of the work in ensuring that

questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. CORRESPONDING AUTHOR Correspondence to Maja Ludvigsen. ETHICS DECLARATIONS

COMPETING INTERESTS A patent has been filed relating to the data presented in this research study by Aarhus University with the following authors as inventers, CTM, MS, YL, MN and CKH

(PCT/EP2019/085957). The remaining authors declare no competing financial interests. ADDITIONAL INFORMATION PUBLISHER’S NOTE Springer Nature remains neutral with regard to jurisdictional

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http://creativecommons.org/licenses/by/4.0/. Reprints and permissions ABOUT THIS ARTICLE CITE THIS ARTICLE Neldeborg, S., Soerensen, J.F., Møller, C.T. _et al._ Dual intron-targeted

CRISPR-Cas9-mediated disruption of the AML _RUNX1-RUNX1T1_ fusion gene effectively inhibits proliferation and decreases tumor volume in vitro and in vivo. _Leukemia_ 37, 1792–1801 (2023).

https://doi.org/10.1038/s41375-023-01950-9 Download citation * Received: 01 April 2022 * Revised: 18 May 2023 * Accepted: 19 June 2023 * Published: 18 July 2023 * Issue Date: September 2023

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