Duchenne muscular dystrophy (DMD) is a debilitating and lethal neuromuscular disorder caused by mutations in the DMD gene located on the X chromosome (Hoffman et al., 1988). Diagnosis is made between the age of 2 and 5, loss of ambulation occurs around 12 and young adults die due to respiratory and/or cardiac failure (Koeks et al., 2017).

DMD is the largest human gene known (Tennyson et al., 1995). Three full-length transcripts encode 427 kDa proteins, while further intragenic promoters drive the expression of progressively truncated variants. Dp427 and the dystrophin-associated protein complex (DAPC) are important for the functional development of differentiating myotubes (Shoji et al., 2015) and subsequently prevent contraction-induced injury in the mature muscle (Rader et al., 2016).

Several studies showed that the ablation of dystrophin in fully differentiated myofibres did not trigger their degeneration (Rader et al., 2016; Ghahramani Seno et al., 2008), and even that myofibres can function without dystrophin (Waugh et al., 2014; Vieira et al., 2015).

In fact, DMD pathology is active prior to diagnosis: delays in the attainment of motor and non‐motor milestones are discernible in 2 months old DMD babies (van Dommelen et al., 2020) and transcriptomes of muscles from asymptomatic DMD patients revealed typical dystrophic abnormalities (Pescatori et al., 2007). Studies of human foetuses (Toop and Emery, 1977; Emery, 1977; Vassilopoulos and Emery, 1977) and various animal DMD models, including GRMD dogs (Nguyen et al., 2002), Sapje zebrafish (Bassett et al., 1977) and Dmd^mdx mouse embryos (Merrick et al., 2009) revealed that the pathology starts already in prenatal development.

Indeed, in skeletal muscle lineages modelled in human DMD pluripotent stem cells, we have recently demonstrated marked transcriptome and miRNA dysregulations identifiable even before muscle specialisation (Mournetas et al., 2021). These data, combined with the existence of the specific embryonic dystrophin Dp412e (Massouridès et al., 2015), substantiate the early disease manifestations in muscle precursor cells.

Importantly, there is increasing evidence that dysregulation of myogenic cells is behind muscle pathology and disease progression in adult DMD muscle. However, the mechanism(s) remain(s) to be elucidated and, given the interaction of myogenic and inflammatory cells in muscle regeneration (Tidball, 2017), distinguishing the primary and the secondary consequences of dystrophin deficiency is crucial for the development of effective therapies.

Helen Blau proposed that DMD is intrinsic to the undifferentiated myoblast (Blau et al., 1983). This hypothesis was initially discounted (Hurko et al., 1987), but there is new evidence that DMD mutations produce a range of cell-autonomous abnormalities in both human and mouse myogenic cells from adult muscles (Yablonka-Reuveni and Anderson, 2006; Yeung et al., 2006; Sacco et al., 2010; Górecki, 2016).

If DMD directly affects myoblasts, cells that are key to muscle regeneration, a better definition of the consequences of the loss of DMD gene expression could help identify early disease biomarkers and establish better therapeutic targets.

Herein, using a combination of RNA-Seq, molecular and functional approaches, we compared dystrophic and healthy myoblasts isolated from skeletal muscles of the commonly used Dmd^mdx mouse model. The Dmd^mdx has a stop codon mutation in exon 23 (Sicinski et al., 1989) resulting in the loss of full-length (Dp427) dystrophin expression, which reflects the molecular defect affecting the majority of DMD patients. Detailed histological and molecular analyses indicated that the clinical phenotype in 3–10 weeks old Dmd^mdx is as aggressive as in Duchenne patients (Duddy et al., 2015; DiMario et al., 1991; Massopust et al., 2020). Therefore, during this period, Dmd^mdx is a good model for the human pathology. Unlike human, after this phase of severe degeneration-regeneration, most Dmd^mdx muscles start to regenerate (DiMario et al., 1991) due to an unknown mechanism.

While Dp427 is expressed in satellite cells and myofibres and its loss in Dmd^mdx reflects the predominant molecular defect in DMD patients, dystrophin Dp71 is present in regenerating muscles and specifically in myoblasts (Howard et al., 1999). Yet, little attention has been given to the DMD pathology in the dystrophin-null individuals. Given that we have recently described exacerbated pathology in Dmd^mdx-βgeo mice lacking all dystrophins (Young et al., 2020), we also investigated the consequences of the total loss of Dmd expression in Dmd^mdx-βgeo myoblasts (Wertz and Füchtbauer, 1998).

Finally, we compared transcriptomic alterations in mouse and human dystrophic myoblasts to identify defects occurring across species. Given that skeletal muscle is a key metabolic site, accounting for around 90% of total oxygen uptake, muscle metabolism is a significant variable (Zurlo et al., 1990). Moreover, metabolic dysfunction can impair muscle regeneration, and the impact of DMD on cellular energetics in muscle progenitors has recently been identified (Matre et al., 2019). Therefore, using the RNA-Seq profiles, we reconstructed the human myoblast-specific enzyme-constrained genome-scale metabolic model.

We report that the absence of expression of the full-length dystrophin triggers major transcriptomic and functional abnormalities in myoblasts. These abnormalities are cell autonomous, as they persist in myogenic cells maintained long-term in culture. Importantly, key alterations are common between mouse and human myoblasts.

DMD presents in muscle stem cells, where the loss of dystrophin affects asymmetrical cell divisions (Dumont et al., 2015b). Subsequently, the absence of dystrophin during myotube differentiation causes typical dystrophic abnormalities such as altered calcium homeostasis and creatine kinase leakage (Shoji et al., 2015). We hypothesised that myoblasts originating from dystrophic stem cells, which give rise to myotubes, are also affected. Indeed, we demonstrate here for the first time that, in proliferating human and mouse myoblasts, the absence of DMD gene expression results in major abnormalities. Our data combining global RNA-Seq and functional analyses demonstrate that DMD directly affects myoblasts.

The key alterations concerned proliferation, chemotaxis, and differentiation. Importantly, synchronisation of these three processes, involving exit from the cell cycle, migration to the site of damage, activation of the skeletal muscle-specific genes and cell fusion, is essential for muscle development and regeneration. Ultimately, myoblasts are the effector cells of muscle growth and repair, and it is the failure of muscle regeneration that drives DMD progression.

These defects are cell autonomous rather than caused by the inflammatory environment of the dystrophic niche because they persist in the dystrophic cell line maintained long-term in culture: established cells reproduced the key transcriptomic anomalies and were described before to have functional alterations found in primary cells (Róg et al., 2019).

Moreover, alterations are shared between mouse and human DMD myoblasts, despite the significant heterogeneity within human samples (Mournetas et al., 2021; Choi et al., 2016). Therefore, these dystrophic anomalies manifest irrespective of genetic backgrounds and across species, which confirms their significance. Furthermore, altered proliferation, migration and differentiation in DMD myoblasts have been reported (Witkowski and Dubowitz, 1985; Melone et al., 2000; Nesmith et al., 2016; Sun et al., 2020) and our data provide the molecular underpinning for these abnormalities.

DMD pathology being intrinsic to the myoblast has been proposed previously (Blau et al., 1983) and there has been growing evidence that DMD mutations cause a range of cell-autonomous abnormalities in human and mouse myogenic cells and myoblasts, which affect cell division, differentiation, energy metabolism and signalling (Yablonka-Reuveni and Anderson, 2006; Yeung et al., 2006; Sacco et al., 2010; Sacco et al., 2010; Young et al., 2020; Young et al., 2020; Dumont et al., 2015b; Onopiuk et al., 2009; Onopiuk et al., 2015; Dumont et al., 2015a; Young et al., 2015). Interestingly, Dmd^mdx cell migration found reduced under physiological stimuli (this work) can be increased in the inflammatory environment (Young et al., 2020). In fact, altered proliferation and migration resulting in highly metastatic phenotypes, have been associated with the loss of Dp427 in tumours featuring myogenic differentiation (Wang et al., 2014).

Our team recently demonstrated that DMD mutations evoke marked transcriptome and miRNome dysregulations early in human muscle cell development (Mournetas et al., 2021). In that study, expression of key coordinators of muscle differentiation was dysregulated in proliferating dystrophic myoblasts, the differentiation of which was subsequently also found to be altered, in line with the mouse cells studied here (Figure 5c).

A significant number of downregulated transcripts in dystrophic myoblasts are controlled by the same TFs, most notably MyoD and MYOG. Given that MYOG itself is regulated by MyoD, the significant downregulation of Myod1 expression found here appears to be the key trigger mechanism of the dystrophic abnormality in myoblasts: the lower levels of Myod1 transcript strongly correlate with downregulated expression of genes controlled by this TF, and which are key coordinators of skeletal muscle repair. Consequently, altered expression of myogenic regulatory factors in dystrophic myoblasts could have multiple consequences. Indeed, complete Myod1 ablation in Dmd^mdx has been shown to exacerbate the disease due to a diminished capability for muscle regeneration (Megeney et al., 1999; Schuierer et al., 2005). Downregulation of MyoD, which inhibits the cell cycle, could explain the increased proliferation in dystrophic cells. The Myod1-Mymk-Mymx (Myomaker/ Myomixer) axis altered here was found to determine myotube formation (Zhang et al., 2020). Given that Pax3 inhibits myogenic differentiation of myoblasts (Epstein et al., 1995), its downregulation in dystrophic cells could also contribute to this accelerated differentiation pattern observed here.

GO categories concerning calcium homeostasis and signalling, known to be altered in dystrophic muscles across species, were also found altered in dystrophic myoblasts: calsequestrins 1 and 2 levels being significantly downregulated, in line with previous findings in the dystrophic muscle Waugh et al., 2014; Doran et al., 2004 and cell lines (Róg et al., 2019; Onopiuk et al., 2015).

Given that Dp71 dystrophin is found in undifferentiated myogenic cells (Howard et al., 1999), and that in development shorter dystrophins have been associated with proliferation and migration, and long isoforms with terminal commitment (Hildyard et al., 2020), we hypothesized that eliminating expression of Dp71 may exacerbate dysfunctions in myogenic cells. Surprisingly, we found that both Dmd^mdx and Dmd^mdx-βgeo cells had similar abnormalities and, contrary to Dp71 overexpression increasing myoblast proliferation (Farea et al., 2020), BrdU incorporation in dystrophin-null myoblasts was still significantly increased compared to WT. Interestingly, Dmd^mdx myoblasts, having a point mutation in exon 23, expressed significantly lower levels of Dp71 (Appendix 1—figure 1), whose expression is driven by a promoted located over 40 exons downstream of the mutation. But whether this downregulation is sufficient to mimic its total loss is unknown.

Combined, these data bring to light the significance of full-length dystrophin defects in undifferentiated muscle cells, the consequences of which correspond to the impaired muscle regeneration occurring in this disease.

Clearly, the important question is the mechanism altering the myoblast phenotype. Dp427 in satellite cells appears to control asymmetrical cell division (Dumont et al., 2015b), which might suggest an interaction problem also in myoblasts.

However, even healthy myoblasts do not express Dp427 protein at levels detectable by Western blotting: an observation that contributed to the belief that myoblasts are not affected by DMD mutations. Although our attempts at immunolocalization in unsynchronised primary myoblasts failed to detect Dp427 (Appendix 1—figure 2), a precise spatio-temporal requirement for small amounts of full-length dystrophin, analogous to its role in satellite cells Dumont et al., 2015b or at neuronal synapses Lidov et al., 1990 is a possibility that should be studied. Importantly, WT myoblasts express the full-length transcript, which is depleted in dystrophic cells. It is worth noting that this 14 kb mRNA, which transcription time may be as long as 16 hr Tennyson et al., 1995, is produced in rapidly dividing myoblasts. Its expression is therefore unlikely to be meaningless. Indeed, downregulation of full-length dystrophin occurs across various malignancies and is associated with alterations of molecular pathways that were identical with alterations found in Duchenne muscles. Moreover, low DMD gene expression in tumours was associated with poor patient survival (https://www.biorxiv.org/content/10.1101/2022.04.04.486990v3). Clealry, dystrophin mis-expression is far more consequenctial and can affect many more cells and tissues than previously understood.

Is it possible that loss of the full-length dystrophin expression may trigger a mechanism not involving its protein product? Considering the increasing number of human pathologies caused by RNA, including another dystrophy – dystrophia myotonica Kino et al., 2018, it is not inconceivable that loss of a large 14 kb transcript made up of 79 exons may trigger a novel RNA-mediated disease process Chan et al., 2018. Abnormalities in Dmd^mdx cells, that harbour just a point mutation, indicate that the patho-mechanism does not require large Dmd gene rearrangements Sudmant et al., 2015. Yet, improvements resulting from the expression of mini/micro dystrophins do not support the requirement for the entire 14 kb transcript or a toxicity of mRNA breakdown products generated via nonsense-mediated decay. However, the impact of premature dystrophin transcript termination on the downregulation of other genes Kamieniarz-Gdula and Proudfoot, 2019 cannot be excluded.

A more likely explanation involves alterations of epigenetic regulation, a mechanism of increasing importance in DMD: The absence of dystrophin/DAPC in satellite cells leads to aberrant epigenetic activation, which impairs functions of the newly generated cells. This dystrophic SC division defect has been linked to polarization problem due to MARK2 mis-interaction Chang et al., 2016 combined with β-syntrophin abnormalities Dumont and Rudnicki, 2016; Ribeiro et al., 2019. Downregulation of β-syntrophin, a DAPC member, results in impaired polarization of its interactor - p38γ. This, in turn, increases CARM1 phosphorylation and reduces MYF5 activation by PAX7 in in the daughter cell due for myogenic differentiation Chang et al., 2018. Furthermore, dystrophin anchors nNOS Brenman et al., 1995, which becomes mis-localised in dystrophic cells. This results in decreased nitric oxide-dependent S-nitrosylation of HDAC2 Colussi et al., 2008 and higher deacetylase activity that has been shown to be involved in the progression of muscular dystrophy Minetti et al., 2006.

Thus, loss of the full-length dystrophin in muscle stem cells does not abrogate myogenic cell divisions but causes significant abnormalities. The descendant myoblasts appear to be harbouring somatically heritable epigenetic changes analogous to genomic imprinting Tucci et al., 2019 and themselves might manifest significant epigenotype abnormalities Zoghbi et al., 2016.

Indeed, our DMD knockdown and rescue experiments in skeletal muscle progenitor cells Mournetas et al., 2021 showed that phenotypic changes at day 17 were the consequences of some earlier events that occurred at day 10 and that some abnormalities might not be reproduced nor alleviated after crossing a specific checkpoint.

The data here indicate that epigenetic changes in dystrophic primary myoblasts may be present, as amongst the significantly altered genes Hist2h2ac, Hist1h2ag, Hist1h2ah, Hist1h1a, Smyd1 and Hmga1 gene products are involved in chromatin modifications or belong to the GO category ‘negative regulation of gene expression, epigenetic’. Methylated histone H3 lysine 4 (H3K4) is a key epigenetic signal for gene transcription. H3K4 methylation is mediated by several methyltransferases, including muscle-active Smyd1 Tan et al., 2006, found among the top downregulated transcripts in dystrophic myoblasts. Histone methylation can be reversed by histone demethylases, of which Lysine-Specific Histone Demethylase 1 A (LSD1 encoded by Kdm1a) is an important muscle enzyme Tosic et al., 2018. The network of interactions between genes in this aforementioned GO category (Appendix 1—figure 3, Supplementary file 1-8) showed a clear interface between the Kdm1a node and significantly downregulated histone genes. LSD1 is involved in controlling Myod1 expression Scionti et al., 2017, which we found significantly downregulated in dystrophic myoblasts. Importantly, treatment of Dmd^mdx mice with histone deacetylase inhibitors (HDACi) promoted myogenesis Saccone et al., 2014. Epigenetically, HDACi upregulate Myod1 Mal et al., 2001 and therefore would counteract its decreased expression and the resulting downregulated expressions of genes controlled by this TF, which we described here.

We postulate that, in dystrophic myoblasts, epigenetic dysregulation of Myod1 expression causes a pathological cascade of downregulated transcriptions of a range of genes controlled by MyoD, with functional consequences for muscle regeneration.

If it is the absence of dystrophin in myogenic cells that determines their fate, developmental muscle abnormalities should occur. This is indeed the case: asymptomatic DMD patients already have transcriptomic alterations Pescatori et al., 2007 and myogenesis in Dmd^mdx embryos is severely disrupted, with myotube hypotrophy, reduced myotube numbers and displacement defects Merrick et al., 2009. This developmental abnormality continues in adult muscles during regeneration.

Altogether, our data identify a continuum, where mutations disrupting expression of full-length dystrophin cause SC division abnormalities impacting myoblast generation but also imprinting these cells with functionalities further reducing muscle regeneration. And as dystrophin is needed at the myotube stage for the initial assembly of the DAPC and subsequent formation of viable myofibres Shoji et al., 2015, dystrophinopathy results in contraction-induced injury in mature muscle (Rader et al., 2016). Thus, the vicious circle of disease is closed with both degenerative and regenerative abnormalities contributing to DMD progression (Figure 10).

Such a scenario explains the apparent contradiction, where removal of dystrophin and/or DAPC in fully developed myofibres does not result in dystrophic muscle damage (Rader et al., 2016; Ghahramani Seno et al., 2008). It would be the case if, once formed in a healthy muscle, DAPC remains stable even if dystrophin is subsequently lost, as indeed have been demonstrated (Ghahramani Seno et al., 2008).

Moreover, as the efficacy of exon skipping and gene targeting methods is much higher in proliferating cells than in myofibres, all outcomes of these treatments could arise from dystrophin re-expression in myoblasts, which later differentiate into myotubes with the functional DAPC.

Findings described here agree with the existing data that loss of dystrophin expression disrupts many downstream processes. These processes offer good targets for therapeutic interventions that are not constrained by the causative mutation Percival et al., 2012; Vidal et al., 2012; Sinadinos et al., 2015. Importantly, adjustments made in dystrophic myoblasts can reduce or even prevent occurrence of dystrophy. For example, upregulation of Jagged 1 can counteract damage in muscles without any dystrophin Vieira et al., 2015 and corrections to COUP-TFII Xie et al., 2016, EGFR-Aurka and Ghrelin expression Wang et al., 2019 can reduce muscle damage.

Given that these pathways are active in myoblasts and the proteins involved are structurally incompatible with substituting the scaffolding properties of dystrophin, these findings further challenge the role of dystrophin in myofibre sarcolemma stabilization as the key pathological alteration in DMD.

Interestingly, the metabolic analysis indicated a significant alteration in key metabolic pathways in DMD myoblasts, which were in line with alterations in the rate of glycolysis/gluconeogenesis, tricarboxylic acid cycle, fatty acid activation, and propanoate metabolism previously found in DMD patients (as previously reviewed Timpani et al., 2015). Clearly, the DMD impact on metabolism is an early and widespread disease manifestation, with abnormalities presenting already in the embryonic development Mournetas et al., 2021 and present in myoblasts Vallejo-Illarramendi et al., 2014 (and this work), in differentiated muscle and also in dystrophic brains Kreis et al., 2011. As an increase in energy demand is observed in DMD muscle, the upregulation of the glycolysis/gluconeogenesis metabolic pathway suggests the upregulation of glucose metabolism-related enzymes, including hexokinase (D-glucose:ATP) and pyruvate kinase, which compensates for the energy demand Filippelli and Chang, 2021. These changes are also compensated by other metabolic alterations in mitochondrial pathways and reactions, for example the downregulation of mitochondrial beta-oxidation of branched-chain fatty acids, odd-chain fatty acids, and di-unsaturated fatty acids (n-6) Nsiah-Sefaa and McKenzie, 2016. Notably, these metabolic adaptations are activated in dystrophic myoblasts and not just myofibres, which could be easier explained by increased energy demands.

The synthesis of ATP and phosphocreatine is catalysed by creatine kinase in a reversible mechanism Bradley, 2004; Scholte and Busch, 1980. Therefore, creatine kinase, significantly downregulated in Arginine and proline metabolism of mitochondrial centre (Figure 9b) signifies the reduction of ATP production in DMD myoblasts Kelly-Worden and Thomas, 2014. The ATP dysfunction at the mitochondrial level induces several patho-physiological events and should exacerbate dystrophinopathy Scholte and Busch, 1980.

The mitochondrial leukotriene metabolism is found to be significantly elevated in DMD myoblasts, consistently with the fact that muscle can produce leukotrienes, which are implicated in inflammation and contribute to muscle weakening Korotkova and Lundberg, 2014. Furthermore, the observed upregulation in propanoate metabolism leads to the accumulation of propionyl-CoA and associated metabolites in plasma, and increases the levels of propionyl-CoA and protein propionylation, which can impair myogenic differentiation in human myoblasts Lagerwaard et al., 2021.

Glucose-6-phosphate dehydrogenase is known to contribute to the regulation of glucose uptake in skeletal muscle Lee-Young et al., 2016. The downregulated fold change of glucose 6-phosphate dehydrogenase (G6PDH2cNo1) suggests that the concentration of glucose is significantly altered in DMD myoblasts. The reduction in the glucose level could impair the differentiation of skeletal myoblasts through the activation of a pathway that targets the enzymatic activity of SIRT1 Fulco et al., 2008.

Our findings here, combined with the other data, provide compelling foundation for the call to re-evaluate the established belief on the pathogenesis of DMD. Such reconsideration is important because most current therapeutic approaches aim at dystrophin restoration in differentiated myofibres and are often conducted in the advanced-stage disease, with very limited regenerative muscle potential. Despite decades of intensive research and numerous clinical trials, none of the candidate treatments delivered disease-modifying results Rodrigues and Yokota, 2018. Part of the problem is immune responses to re-expressed dystrophin Vila et al., 2019, which need to be tackled Sharma et al., 2017 or prevented. However, dystrophin re-expression using new technologies Mangeot et al., 2019 allowing re-targeting dystrophin to myoblasts and satellite cells in younger patients or, better still, in newborns able to develop neonatal tolerance to dystrophin, could produce effective therapies for this devastating disease.

All data generated or analysed during this study are included in the manuscript and supporting files. Supplementary tables uploaded with the manuscript provide the numerical data that are represented in graphical format in the manuscript figures or as summary tables. RNA-Seq data for the mouse myoblast samples have been deposited in the ArrayExpress database at EMBL-EBI under accession number E-MTAB-10322 and E-MTAB-7287 for primary myoblasts and established cell lines respectively (RNA Sequencing section), and under E-MTAB-8321 for human primary myoblasts (Human primary adult myoblast RNA-Seq data section).

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Author details

1. Maxime RF Gosselin

   School of Pharmacy and Biomedical Sciences, University of Portsmouth, Portsmouth, United Kingdom

   Contribution

   Investigation, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-8916-0953

2. Virginie Mournetas

   INSERM UEVE UMR861, I-STEM, AFM, Corbeil-Essonnes, France

   Contribution

   Formal analysis, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6557-4190

3. Malgorzata Borczyk

   Laboratory of Pharmacogenomics, Maj Institute of Pharmacology PAS, Krakow, Poland

   Contribution

   Formal analysis, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

4. Suraj Verma

   School of Computing, Engineering and Digital Technologies, Teesside University, Middlesbrough, United Kingdom

   Contribution

   Formal analysis, Investigation, Writing – original draft

   Competing interests

   No competing interests declared

5. Annalisa Occhipinti

   School of Computing, Engineering and Digital Technologies, Teesside University, Middlesbrough, United Kingdom

   Contribution

   Investigation, Formal analysis

   Competing interests

   No competing interests declared

6. Justyna Róg

   1. School of Pharmacy and Biomedical Sciences, University of Portsmouth, Portsmouth, United Kingdom
   2. Laboratory of Cellular Metabolism, Nencki Institute of Experimental Biology, Warsaw, Poland

   Contribution

   Investigation, Writing – original draft

   Competing interests

   No competing interests declared

7. Lukasz Bozycki

   1. School of Pharmacy and Biomedical Sciences, University of Portsmouth, Portsmouth, United Kingdom
   2. Laboratory of Cellular Metabolism, Nencki Institute of Experimental Biology, Warsaw, Poland

   Contribution

   Investigation, Visualization

   Competing interests

   No competing interests declared

8. Michal Korostynski

   Laboratory of Pharmacogenomics, Maj Institute of Pharmacology PAS, Krakow, Poland

   Contribution

   Software, Formal analysis, Supervision, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

9. Samuel C Robson

   1. School of Pharmacy and Biomedical Sciences, University of Portsmouth, Portsmouth, United Kingdom
   2. Centre for Enzyme Innovation, University of Portsmouth, Portsmouth, United Kingdom

   Contribution

   Data curation, Formal analysis, Supervision, Investigation, Writing – review and editing

   Competing interests

   SR received royalties from Cancer Research Technology Ltd through University of Cambridge Enterprise Ltd under agreement A13086 for "Chromatin-bound METTL3 m6A methyltransferase facilitates myeloid leukaemogenesis by modulating ribosome stalling"

   "This ORCID iD identifies the author of this article:" 0000-0001-5702-9160

10. Claudio Angione

    School of Computing, Engineering and Digital Technologies, Teesside University, Middlesbrough, United Kingdom

    Contribution

    Conceptualization, Software, Formal analysis, Supervision, Methodology

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-3140-7909

11. Christian Pinset

    CNRS, I-STEM, AFM, Corbeil-Essonnes, France

    Contribution

    Formal analysis, Supervision, Writing – review and editing

    Competing interests

    No competing interests declared

12. Dariusz C Gorecki

    School of Pharmacy and Biomedical Sciences, University of Portsmouth, Portsmouth, United Kingdom

    Contribution

    Conceptualization, Supervision, Funding acquisition, Investigation, Writing – original draft, Project administration, Writing – review and editing

    For correspondence

    darek.gorecki@port.ac.uk

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0003-3584-1654

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