In eukaryotes, mitotic entry is marked by the profound reorganization of chromatin into mitotic chromosomes driven by the condensin complex (Hirano, 2016). While mitotic chromosome assembly or condensation is essential for the accurate transmission of the genome to daughter cells, our understanding of the mechanisms by which condensin associates with chromatin, shapes mitotic chromosomes, and contributes to their accurate segregation in anaphase remains incomplete.

Condensin is a ring-shaped ATPase complex that belongs to the structural maintenance of chromosomes (SMC) family of genome organizers, which also includes the cohesin complex involved in chromatin folding during interphase and in sister-chromatid cohesion (Hirano, 2016; Davidson and Peters, 2021). Condensin is composed of a core ATPase heterodimer, made of the SMC2 and SMC4 proteins, associated with a kleisin and two HEAT-repeat subunits. Most multicellular eukaryotes possess two condensin variants, named condensin I and II, made of a same SMC2/4 core but associated with distinct sets of non-SMC subunits (Ono et al., 2003; Hirano, 2012). Budding and fission yeasts, in contrast, possess a single condensin complex, similar to condensin I. Thereafter, condensin complexes will be collectively referred to as condensin, unless otherwise stated. There is robust evidence that condensin shapes mitotic chromosomes by massively binding to DNA upon mitotic entry and by folding chromatin into arrays of loops (Gibcus et al., 2018; Kakui et al., 2017). Thereby, condensin conceivably reduces the length of chromosomes, confers to chromosome arms the stiffness to withstand the spindle traction forces (Sun et al., 2018), and promotes the removal of catenations between chromosomes and sister-chromatids by orientating the activity of topoisomerase II (Topo II) toward decatenation (Baxter et al., 2011; Charbin et al., 2014). Hence, when condensin is impaired, sister-centromeres often reach the opposite poles of the mitotic spindle in anaphase but chromosome arms fail to separate, forming stereotypical chromatin bridges. In vitro studies have shown that condensin anchors itself on naked DNA through sequence-independent electrostatic interactions and uses the energy of ATP hydrolysis to extrude adjacent DNA segments into a loop of increasing size (Kschonsak et al., 2017; Ganji et al., 2018; Kong et al., 2020). Although such a loop extrusion reaction convincingly explains the structural properties of mitotic chromosomes (Nasmyth, 2017; Davidson and Peters, 2021), we still ignore whether and how it could take place in the context of a chromatinized genome, crowded with potential hindrances.

There is robust evidence that chromatin microenvironments impinge upon the localization of condensin. ChIP-sequencing (ChIP-seq) studies performed on species ranging from yeasts to mammals have revealed a conserved condensin pattern along the genome, constituted of a broad and basal distribution punctuated by peaks of high occupancy at centromeres, rDNA repeats, and in the vicinity of highly expressed genes (D’Ambrosio et al., 2008; Kim et al., 2013; Kranz et al., 2013; Dowen et al., 2013; Sutani et al., 2015). Various factors such as the chromokinesin Kif4 (Samejima et al., 2012), the zinc-finger protein AKAP95 (Steen et al., 2000), transcription factors, and chromatin remodelers (for review see Robellet et al., 2017) have been involved in the binding of condensin to chromatin in yeasts or vertebrate cells. Additional cis-acting factors that increase condensin’s local concentration at centromeres and/or at rDNA repeats have been identified in budding or fission yeast (Tada et al., 2011; Johzuka and Horiuchi, 2009; Verzijlbergen et al., 2014). Such enrichments are likely to play a positive role since there is clear evidence that condensin contributes to the stiffness of centromeric chromatin and to the bilateral attachment of centromeres in early mitosis (Ono et al., 2004; Gerlich et al., 2006; Nakazawa et al., 2008; Ribeiro et al., 2009; Verzijlbergen et al., 2014; Piskadlo et al., 2017). Likewise, the segregation of the rDNA is acutely sensitive to condensin activity (Freeman et al., 2000; Nakazawa et al., 2008; Samoshkin et al., 2012). Highly expressed genes, in contrast, are thought to constitute a permeable barrier where active condensin complexes stall upon encounters with DNA-bound factors such as RNA polymerases (Brandão et al., 2019; Rivosecchi et al., 2021). Consistent with a local hindrance, in fission yeast, attenuating transcription that persists during mitosis improves chromosome segregation when condensin is impaired (Sutani et al., 2015). Further evidence in budding yeast indicates that dense arrays of protein tightly bound to DNA can constitute a barrier for DNA-translocating condensin (Guérin et al., 2019). Thus, depending on the context, condensin enrichment can reflect either positive or negative interplays.

Microscopy studies have clearly shown that condensin I is enriched at telomeres during mitosis and meiosis in mammalian cells (Walther et al., 2018; Viera et al., 2007), and ChIP-seq has further revealed that condensin I accumulates at telomere repeats in chicken DT40 cells, but the mechanisms underlying such enrichment as well as its functional significance have remained unknown. We and others previously showed that the separation of sister-telomeres in anaphase involves condensin regulators such as Cdc14 phosphatase in budding yeast (Clemente-Blanco et al., 2011), and Aurora-B kinase in fission yeast (Reyes et al., 2015; Berthezene et al., 2020), but whether and how condensin could play a role has remained unclear.

In the present study, we sought to determine how and why condensin is enriched at telomeres by using the fission yeast Schizosaccharomyces pombe as a model system. Telomeres contain G-rich repetitive sequences that are protected by a conserved protein complex called Shelterin (de Lange, 2018; Lim and Cech, 2021), which is composed, in fission yeast, of Taz1 (a myb-domain DNA-binding protein homologous to human TRF1 and TRF2), Rap1, Poz1 (a possible analog of TIN2), Tpz1 (an ortholog of TPP1), Pot1, and Ccq1. While Taz1 binds to double-stranded G-rich telomeric repeats, Pot1 binds to 3′ single-stranded overhang. Rap1, Poz1, and Tpz1 act as a molecular bridge connecting Taz1 and Pot1 through protein-protein interactions. Ccq1 contributes to the recruitment of the nucleosome remodeler Mit1 and of telomerase (for review on fission yeast shelterin see Moser and Nakamura, 2009; Dehé and Cooper, 2010). We found that Taz1 plays the role of a cis-acting enrichment factor for condensin at telomeres, while Mit1 antagonizes condensin’s accumulation. Thus, telomeres are a remarkable chromosomal environment where condensin is enriched by a shelterin-dependent cis-acting mechanism. Our results further indicate that the level of condensin at telomeres, set up by Taz1 and Mit1, is instrumental for their proper disjunction during anaphase, hence associating a key biological function to this local enrichment. Based on these data, we propose that condensin is enriched at telomeres via interplays with shelterin proteins to drive sister-telomere separation in anaphase.

With this work, we show that condensin is enriched at fission yeast telomeres during mitosis and that such enrichment results from the balancing acts of telomeric proteins. We also show that the separation of sister-telomeres is not the mere consequence of the separation of chromosome arms and that condensin acts in cis at telomeres to drive their disjunction during anaphase. We provide evidence that condensin might achieve this task by counteracting cohesin.

Previous work has shown that the kleisin subunit of condensin II (CAPH2) binds human TRF1, a counterpart of Taz1, in RPE-1 cells (Wallace et al., 2019), but since no physical or functional link has been described between TRF1 and other subunits of the condensin II holocomplex, it was unclear whether CAPH2 might act at telomeres independently of condensin II. Hence, the biological significance of the presence of condensin complexes at telomeres remained enigmatic. Here, we show that the kleisin subunit of fission yeast condensin is bound to the telomere repeats of TEL2R in metaphase and anaphase and that such binding relies on the Cut14^SMC2 and Cut3^SMC4 ATPases (Saka et al., 1994), arguing therefore that the condensin holocomplex is bound to TEL2R. Since southern blotting and FISH experiments have shown that chromosome I and II contains similar sub-telomeric elements (Funabiki et al., 1993; Oizumi et al., 2021), our observations made using TEL2R DNA are likely to be relevant to most fission yeast telomeres.

We further show that condensin occupancy at telomeres is controlled by the telomeric proteins Taz1 and Mit1. Taz1 being a core component of the shelterin complex, we cannot formally rule out that the reduced binding of condensin stems from a collapse of the overall telomeric structure in cells lacking Taz1. However, two observations argue against such scenario. First, the fact that deleting Rap1, another key component of shelterin, does not impair condensin localization and, second, our finding that condensin binding to telomeres is also controlled, negatively, by the nucleosome remodeler Mit1, which associates with telomeres via the Ccq1 subunit of shelterin (Sugiyama et al., 2007; van Emden et al., 2019). Such negative regulation by Mit1 not only strengthens our previous work suggesting that nucleosome arrays are an obstacle for condensin binding to DNA in vivo (Toselli-Mollereau et al., 2016), but also strongly suggests that condensin localization at telomeres relies on a dedicated pathway that involves interplays with telomeric components. However, and in sharp contrast with the human TRF1 and CAP-H2 (Wallace et al., 2019), we detected no protein-to-protein interactions between Taz1 and Cnd2/condensin. Together with our observation that Taz1 does not enrich condensin at discrete Taz1-DNA binding sites located outside telomeres, this suggests that Taz1 is not a cis-acting recruiter for condensin at telomeres. Rather, by analogy with the accumulation of condensin at highly expressed genes (Brandão et al., 2019; Rivosecchi et al., 2021), we speculate that arrays of Taz1 proteins tightly bound to telomere repeats might create a permeable barrier onto which condensin molecules accumulate. However, unlike highly expressed genes that most likely hinder condensin-mediated chromosome segregation in anaphase (Sutani et al., 2015), the Taz1 barrier would play a positive role in chromosome segregation by promoting sister-telomere disjunction in anaphase.

Using Hi-C and live cell imaging, we provide evidence that condensin takes part in telomere declustering during the early steps of mitosis and in sister-telomere disjunction in anaphase. It is tempting to speculate that condensin promotes the dissociation of telomeric clusters, inherited from the Rabl organization of chromosomes in interphase, by folding chromatin into mitotic chromosomes. As condensation proceeds, axial shortening and stiffening of chromosome arms would drive the movement of the pairs of telomeres located at the opposite ends of a chromosome away from each other. The separation of sister-telomeres during anaphase, in contrast, cannot be the passive consequence of the separation of sister-chromatids. Indeed, the striking observation that sister-telomere disjunction can be uncoupled from the separation of chromosome arms, as seen in the decatenation-defective topo-250 mutant, implies the existence of a mechanism independent of chromosome arms, driving sister-telomere disjunction. In fission yeast, condensin must play a key role within such telomere-disjunction pathway since modulating its occupancy at telomeres while leaving chromosome arms largely unchanged, using taz1Δ or mit1Δ mutations, is sufficient to change accordingly the efficiency of sister-telomeres disjunction. The fact that condensin occupancy at telomeres is a limiting parameter for their disjunction argues for a role played in cis. This finding is reminiscent of telomere separation in human cells that specifically relies on the activity of the poly(ADP-ribose) polymerase tankyrase 1 (Dynek and Smith, 2004), and suggest therefore that the existence of a dedicated pathway for sister-telomere disjunction is a conserved feature of eukaryotic cells. We therefore conclude that condensin enriched at telomeres via the balancing acts of Taz1 and Mit1 drives the separation of sister-telomeres in anaphase. The corollary is that failures to disjoin sister-telomeres most likely contribute to the stereotypical chromatin bridge phenotype exhibited by condensin-defective cells. Our results do not rule out the possibility that Topo II contributes to telomeres disentanglements, but nevertheless imply that Topo II catalytic activity is dispensable for telomere segregation provided that condensin is active. The close proximity of DNA ends could explain such a dispensability. It has been reported in budding yeast that the segregation of LacO repeats inserted in the vicinity of TelV is impaired by the top2-4 mutation (Bhalla et al., 2002). At first sight, this appears at odds with our observations made using the telomere protein Taz1 tagged with GFP. However, since LacO arrays tightly bound by LacI proteins constitute a barrier for the recoiling activity of budding yeast condensin in anaphase (Guérin et al., 2019), the insertion of such a construct might have created an experimental condition in which condensin activity was specifically impaired at TELV, hence revealing the contribution of Topo II. In addition, the telomere structure in budding and fission yeast is significantly different. Budding yeast protects its telomeres via two independent factors, Rap1 and the Cdc13-Stn1-Ten1 complex, whereas in fission yeast Taz1 and Pot1 are bridged by a complex protein interaction network (Rap1-Poz1-Tpz1). This is a remarkable conserved structural feature between the shelterin of S. pombe and the human shelterin. Notably, it was recently shown that the telomeric components of S. pombe can dimerize leading to a higher complex organization of the shelterin (Sun et al., 2022). It is thus likely that dimerization of Taz1, Poz1, and the Tpz1-Ccq1 subcomplex may also contribute to the clustering of sister and non-sister-chromatid telomeres. The architectural differences in telomere organization between budding and fission yeast may require different mechanisms to properly segregate telomeres during mitosis.

Understanding how condensin takes part in the disjunction of sister-telomeres will require identifying the ties that link them. Cohesin has been involved in telomere cohesion in budding yeast (Antoniacci and Skibbens, 2006; Renshaw et al., 2010), but in human cells the situation remains unclear. Although Scc3^SA1 is a likely target of the tankyrase 1 pathway for telomere disjunction (Canudas and Smith, 2009), telomeric cohesion appears independent of other cohesin subunits (Bisht et al., 2013). Our finding that rad21-K1, a loss-of-function mutation in the kleisin subunit of fission yeast cohesin, accelerates sister-telomere disjunction in an otherwise wild-type genetic background would be consistent with a role for cohesin in ensuring cohesion between sister-telomeres in fission yeast. Alternatively, rad21-K1 might indirectly increase the impact of condensin at chromosome ends, for instance by altering the structure of sub-telomeric heterochromatin (Dheur et al., 2011). However, such an indirect effect seems less likely because the kinetics of sister-telomere disjunction is not accelerated in cells lacking the core heterochromatin protein Swi6 (Reyes et al., 2015). Therefore, we favor the conclusion that condensin drives sister-telomere disjunction by counteracting cohesin at chromosome ends. Whether it could be cohesive- or loop-extruding cohesin remains to be determined, but we note that an antagonism between condensin and cohesin for the folding of interphase chromatin as well as for telomere segregation in anaphase has been reported in Drosophila and budding yeast, respectively (Rowley et al., 2019; Renshaw et al., 2010). Thus, unravelling the mechanism by which condensin drives telomere disjunction in anaphase will require further investigations not only on the interplays between condensin and cohesin at telomeres, but also on the role played or not by condensin loop extrusion activity and on the dynamics of shelterin. Because of its ability to organize telomeres into various structures (Lim and Cech, 2021), the shelterin complex may link sister-telomeres together and loop extrusion by condensin may provide the power stroke to disentangle such structures. Thus, as speculated in Figure 7, the accumulation of condensin against Taz1^TRF1-2 barriers, together with a possible up-regulation by Aurora-B kinase in anaphase (Reyes et al., 2015), might allow condensin-mediated DNA translocation to pass a threshold beyond which the ties between sister-telomeres would be eliminated, be it cohesin- and/or shelterin-mediated. Whatever the mechanism, given the conservation of shelterin, and the abundance of condensin complexes at telomeres during mitosis and meiosis in mammals (Viera et al., 2007; Walther et al., 2018), we speculate that condensin specifically drives the separation of telomeres in other living organisms.

Figure 7

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Scripts and pipelines are available in the git repository https://gitbio.ens-lyon.fr/LBMC/Bernard/chipseq (tag v0.1.0). Pipelines were executed with nextflow (version 23.02.1). In the subsequent section, genome refers to the S. pombe genome (ASM294v2, or its TEL2-R extended version available at Omnibus GEO GSE196149), while calibration genome refers to the genome of S. cerevisiae (sacCER3 release R64-1-1) used for internal calibration. To perform the analyses, we use a modified version of the nf-core/chipseq (version 2.0.0) pipeline (https://doi.org/10.1038/s41587-020-0439-x). We modified this pipeline as follows: We added an optional --fasta_calibration parameter to pass the calibration genome. We modified the subworkflow prepare_genome.nf to run GUNZIP_FASTA on both the --fasta and --fasta_calibration files and merge the two fasta file with the MIX_FASTA process, while adding a cali_ prefix to the names of the chromosome of the calibration genome. The workflow chipseq.nf was modified to run a new process, BAM_CALIB (replacing BEDTOOLS_GENOMECOV), on the IP bam file and their corresponding Total (INPUT) bam file to generate calibrated IP and INPUT bigwig files. The workflow chipseq.nf was modified to then run BIGWIG2BAM to generate synthetic bam files from the output of BAM_CALIB. These synthetic bam files are single-end and replace the output of the mapping step in the next parts of the pipeline. The BIGWIG2BAM outputs are indexed with a SAMTOOLS_INDEX process. The BAM_CALIB tool (https://gitbio.ens-lyon.fr/LBMC/Bernard/bamcalib v0.1.5) takes into inputs a sorted bam file for the IP data and a sorted bam file for the TOTAL data (mapped on the concatenation of the two genomes), and output a calibrated bigwig for the reference genome. For the normalization, we modified a previously described method (Hu et al., 2015) in order to account for biases in coverages in the TOTAL fractions. We introduce the following notation: ${IP}_x\left(t\right)$ is the coverage at position $t$ in the IP sample on the reference genome, ${IP}_c\left(t\right)$ is the coverage at position $t$ in the IP sample on the calibration genome, ${WCE}_x\left(t\right)$ is the coverage at position $t$ in the TOTAL (whole cell extract) sample on the reference genome, and ${WCE}_c\left(t\right)$ the coverage at position $t$ in the TOTAL sample on the calibration genome. In a reference genome of size $T_x$ (ignoring the chromosome segmentation), and in a calibration genome of size $T_c$ , Hu et al., 2015 compute the occupancy ratio (OR) as follows:

Instead, we used the following formula with β (default to 10³) an arbitrary scaling factor.

This formula can be described as follows:

The technical variations on the IP efficiencies are corrected by scaling ${\displaystyle I{P}_x(t)}$ by the calibration genome coverage:

To account for variations in cells proportion, we correct by a scaled WCE coverage:

To be able to analyze the coverage information at repetitive regions of the genome, we propose to normalize the signal nucleotide by nucleotide and introduce the OR ratio:

with:

We then find such that (not to distort the normIP signal on average):

which gives

With this method, we retain the internal calibration developed by Hu et al., 2015, and we account for variations in read density at each base in WCE samples.

In the BAM_CALIB tools, the coverage does not correspond to the number of read covering a given position like in classical tools outputting bigwig. Instead we compute the number of fragments for paired-end data. To compute the coverage density ${\displaystyle X_y(t)}$ with ${\displaystyle X\in [IP,WCE]}$ and ${\displaystyle y\in [c,x]}$ we count the number of reads r(t) overlapping with position ${\displaystyle t}$. For properly paired reads (with a mate read on the same chromosome and with a starting position ending after the end of the read) we also count a density of 1 between the end of the first reads and the start of his mate read g(t). ${\displaystyle X_y(t)=r(t)+g(t)}$. Some fragment can be artificially long (with reads mapping to repeated regions at the start and end of a chromosome), therefore, we compute a robust mean µ of the gap size, between two reads of a pair, by removing the 0.1 upper and lower quantile of the fragment length distribution. Fragments with a size higher than ${\displaystyle {\varphi }^{-1}(0.95,\mu ,1.0)}$ are set to end at the ${\displaystyle {\varphi }^{-1}(0.95,\mu ,1.0)}$ value, with ${\displaystyle \varphi ()}$ the Normal CDF function. For fragments shorter than the read length, we don't count the overlapping reads region as a coverage of two fragments but as the coverage of 1 fragment. The BIGWIG2BAM Tools (https://gitbio.ens-lyon.fr/LBMC/Bernard/bamcalib v0.1.1) generate a synthetic bam file from a bigwig file and a reference genome. The purpose of this tool is to create a bam file having the same coverage profile as the one described in the input bigwig file. Therefore, we can run any chip-seq tools working with bam file instead of bigwig file on our normalized data.

Datasets have been deposited in NCBI GEO under the accession code GSE196149.

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Author details

1. Léonard Colin

   1. CNRS - Laboratory of Biology and Modelling of the Cell, Lyon, France
   2. ENS de Lyon, Université Lyon, site Jacques Monod, Lyon, France

   Contribution

   Investigation, Visualization

   Contributed equally with

   Celine Reyes

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6333-2569

2. Celine Reyes

   MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France

   Contribution

   Investigation, Visualization

   Contributed equally with

   Léonard Colin

   Competing interests

   No competing interests declared

3. Julien Berthezene

   MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France

   Contribution

   Investigation, Visualization

   Competing interests

   No competing interests declared

4. Laetitia Maestroni

   CNRS, INSERM, Aix Marseille Université, Institut Paoli-Calmettes, CRCM, Equipe labellisée par la Ligue Nationale contre le Cancer, Marseille, France

   Contribution

   Investigation, Visualization

   Competing interests

   No competing interests declared

5. Laurent Modolo

   1. CNRS - Laboratory of Biology and Modelling of the Cell, Lyon, France
   2. ENS de Lyon, Université Lyon, site Jacques Monod, Lyon, France

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-7606-4110

6. Esther Toselli

   1. CNRS - Laboratory of Biology and Modelling of the Cell, Lyon, France
   2. ENS de Lyon, Université Lyon, site Jacques Monod, Lyon, France

   Contribution

   Investigation, Visualization

   Competing interests

   No competing interests declared

7. Nicolas Chanard

   1. MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France
   2. CBI, MCD-UMR5077, CNRS, Chromatin Dynamics Team, Toulouse, France

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

8. Stephane Schaak

   1. MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France
   2. CBI, MCD-UMR5077, CNRS, Chromatin Dynamics Team, Toulouse, France

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

9. Olivier Cuvier

   1. MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France
   2. CBI, MCD-UMR5077, CNRS, Chromatin Dynamics Team, Toulouse, France

   Contribution

   Supervision, Investigation, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0644-2734

10. Yannick Gachet

    MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France

    Contribution

    Conceptualization, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing – review and editing

    For correspondence

    yannick.gachet@univ-tlse3.fr

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0001-7548-7974

11. Stephane Coulon

    CNRS, INSERM, Aix Marseille Université, Institut Paoli-Calmettes, CRCM, Equipe labellisée par la Ligue Nationale contre le Cancer, Marseille, France

    Contribution

    Conceptualization, Supervision, Funding acquisition, Investigation, Visualization, Methodology, Writing – review and editing

    For correspondence

    stephane.coulon@inserm.fr

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0001-8090-914X

12. Pascal Bernard

    1. CNRS - Laboratory of Biology and Modelling of the Cell, Lyon, France
    2. ENS de Lyon, Université Lyon, site Jacques Monod, Lyon, France

    Contribution

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

    For correspondence

    pascal.bernard@ens-lyon.fr

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0003-2732-9685

13. Sylvie Tournier

    MCD, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, Toulouse, France

    Contribution

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

    For correspondence

    sylvie.tournier-gachet@univ-tlse3.fr

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-7869-6891

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