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The cryptic complex rearrangements involving the DMD gene: etiologic clues about phenotypical differences revealed by optical genome mapping

Human Genomics volume 18, Article number: 103 (2024) Cite this article

Abstract

Background

Deletion or duplication in the DMD gene is one of the most common causes of Duchenne and Becker muscular dystrophy (DMD/BMD). However, the pathogenicity of complex rearrangements involving DMD, especially segmental duplications with unknown breakpoints, is not well understood. This study aimed to evaluate the structure, pattern, and potential impact of rearrangements involving DMD duplication.

Methods

Two families with DMD segmental duplications exhibiting phenotypical differences were recruited. Optical genome mapping (OGM) was used to explore the cryptic pattern of the rearrangements. Breakpoints were validated using long-range polymerase chain reaction combined with next-generation sequencing and Sanger sequencing.

Results

A multi-copy duplication involving exons 64–79 of DMD was identified in Family A without obvious clinical symptoms. Family B exhibited typical DMD neuromuscular manifestations and presented a duplication involving exons 10–13 of DMD. The rearrangement in Family A involved complex in-cis tandem repeats shown by OGM but retained a complete copy (reading frame) of DMD inferred from breakpoint validation. A reversed insertion with a segmental repeat was identified in Family B by OGM, which was predicted to disrupt the normal structure and reading frame of DMD after confirming the breakpoints.

Conclusions

Validating breakpoint and rearrangement pattern is crucial for the functional annotation and pathogenic classification of genomic structural variations. OGM provides valuable insights into etiological analysis of DMD/BMD and enhances our understanding for cryptic effects of complex rearrangements.

Introduction

Duchenne muscular dystrophy (DMD; OMIM #310200) and Becker muscular dystrophy (BMD; OMIM #300376) are the most common neuromuscular disorders, characterized by progressive muscle loss with various severity [1]. One in every 3600–6000 live births of male infants is diagnosed with either DMD or BMD [2].

Both DMD and BMD are X-linked recessive diseases caused by variants in the DMD gene (OMIM #300377), encoding dystrophin [3]. The DMD gene is located at Xp21.1 and includes 79 exons spanning 2.4 Mb [4]. Among documented distinct DMD variants, deletions account for the majority (60–70%), followed by small insertions/deletions and SNV (20%), and finally duplications (5–15%) [5]. However, this distribution varies across different geographic regions [6]. Out-of-frame variants in DMD lead to loss of dystrophin protein expression, resulting in DMD, whereas in-frame variants partially retain functional dystrophin, causing BMD. The reading frame rule has been the main hypothesis to explain the phenotypic differences between DMD and BMD [7, 8]. However, nearly 9% of the cases violate the rule, mainly attributable to alternative splicing patterns and levels of the RNA creating in-frame or out-of-frame transcripts, or disturbance of the exonic splicing enhancers inducing skipping of partial or a certain exon(s), which may restore or disrupt the reading frame [9]. Loss or disturbance of intronic elements required for the proper inclusion of a certain exon(s) also accounts for exceptions in some cases [9]. These exceptions had a high rate in the BMD series [8]. Furthermore, duplications make up nearly one-third of the variants that break the reading frame restrictions [9].

Deletions or duplications in the DMD gene can be detected routinely using multiplex ligation-dependent probe amplification (MLPA) [10], or in some cases chromosomal microarray analysis or copy number variation sequencing (CNV-seq) can be adopted [11, 12]. One of the most common limitations of these methodologies is their inability to identify balanced genomic rearrangements and profile the pattern of balanced or unbalanced structural changes [13, 14]. As a result, most complex rearrangements, together with the related breakpoints and patterns, are frequently neglected in routine genetic testing. These structural rearrangements can result in severe disorders [15], presumably due to dosage or position effects arising from the loss or gain of genomic sequences or changes in the position of genomic segments [16]. Understanding these potential patterns and evaluating the phenotypic effects of these complex rearrangements require the precise mapping of the breakpoints involved. This is particularly true for deletions or duplications in the DMD gene, which may potentially disrupt the regulatory intronic elements or exonic enhancers related to alternative splicing of the DMD gene transcripts.

Optical genome mapping (OGM) is a novel technique for detecting various classes of SVs [17]. Its ultra-long read length (even more than 200 kb) with an average resolution of 500 bp, allows for the identification of precise location and orientation of aberrant structures across the whole genome [18, 19]. Mantere et al. used OGM to identify an inverted region on chromosome 13 and the breakpoint cutoff KLHL1 gene [17]. Dai et al. used OGM to directly quantify the number of repeats in the D4Z4 repeat sequence and the level of post-zygotic mosaicism in facioscapulohumeral muscular dystrophy, the results highly concordant with the Southern blot analysis [20]. Compared with short-read sequencing techniques, OGM simplifies the identification of complicated structural rearrangements [17].

The pathogenicity of complex rearrangements involving DMD with unknown breakpoints is not well understood. In this study, we identified different complex rearrangements in two families involving DMD duplications using OGM: an in-cis multi-copy tandem repeats and a reversed inversion with a segmental repeat. We investigated the possible patterns and breakpoints of these complex rearrangements and assessed their pathogenicity by combining the clinical phenotypes of the two families. Our findings revealed potential mechanisms generating these complex rearrangements, thus providing new insights into the classification and effects of structural variants.

Methods

Participants

Two families (A and B, as shown in Figs. 1 and 2, respectively) with DMD segmental duplications (duplication events occurring with a genomic segment as a rearrangement unit) exhibiting phenotypical differences were recruited. A comprehensive family history was investigated and detailed clinical manifestations were evaluated for all members. Peripheral blood samples were collected from all the participants. Written informed consent was obtained from all the participants. This study was approved by the Ethics Committee of the Second Affiliated Hospital of Guangxi Medical University (NO. 2022-KY0632).

Fig. 1
figure 1

Pedigree and duplication variation of Family A. A. The pedigree of Family A. Dup: DMD duplications. Dup hemi: hemizygous DMD. Black arrow represents proband. B. CNV-seq results showed the proband (II-3) and her mother (I-2) had approximately seven copies of the repeat, and her elder brothers (II-1 and II-2) had approximately six copies of the repeat. The region of the duplication at chrX: 30880000–31260000. The horizontal axis shows the position of the X chromosome, and the vertical axis shows the copy number. C. The results of copy number of duplication region for Family A (violet) and four samples with known copy number of chromosome X (45,X; 46,XX; 47,XXX; 46,XY) (blue) by qPCR. D. MLPA showed that the duplication in Family A involved exons 64–79 of the DMD gene

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Fig. 2
figure 2

Pedigree and duplication variation of Family B. A. The pedigree of Family B. Dup: DMD duplications. Dup hemi: hemizygous DMD variations. The black arrow represents proband. B. MLPA showed that the duplication in Family B involved exons 10–13 of the DMD gene

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CNV-seq

Genomic DNA was extracted from peripheral whole blood using the DNeasy Blood and Tissue Kit (Qiagen GmbH, Hilden, Germany). The CNV-seq was conducted to analyze the copy number variation in Family A. The sequencing libraries were prepared based on PCR-free methods with a input of 100ng genomic DNA as previously described [21], which were subsequently sequenced using the NextSeq CN500 platform (Illumina, San Diego, CA, USA) to produce at least 10 million 36-bp single-end raw reads. After filtration, only uniquely mapped reads were aligned to the human genome (GRCh37/hg19) using the Burrows–Wheeler Alignment tool (BWA) [22], and then were allocated to 20-kb bins across the whole genome. Binned reads were counted, and the ratio between the observed and expected average read counts in each of the bin was computed. The read count ratio of each bin for the test sample was then compared and normalized internally with that for the reference from in-house database, representing the relative copy number of the test sample compared to the reference, as previously described [23].

Quantitative real-time PCR (qPCR)

The qPCR was used to verify the specific copy number of the identified complex duplication in Family A. The primers for the target locus, Dup309 allocating within the duplication region (chrX: 30985277–30985418, GRCh38/hg38) and the reference locus, XP60 allocating on Xp11.3 outside the duplication region (chrX: 43949919–43950089, GRCh38/hg38) were designed as shown in Table 1. To assess the PCR efficiencies and consistencies of the target and reference loci, we set a sample with one copy of chromosome X (45,X) as control and used another four samples with known copy number of chromosome X (45,X; 46,XX; 47,XXX; 46,XY) as calibrators. The PCR reaction was conducted in triplicate for each samples according to the PowerUp SYBR Green Master Mix protocol (Thermo Fisher Scientific, Waltham, MA, USA) in a 20 µL volume with a input genomic DNA of 30ng. The CT values of the test, calibrator, and control samples were used to perform a comparative CT (ΔCT) relative quantitation analysis. The copy number of the target locus in the test/calibrator samples was calculated by comparing the difference of the threshold cycles between the test/calibrator and control samples after normalizing and adjusting the corresponding threshold cycles of the reference locus.

Table 1 Primer information
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MLPA

The MLPA analysis was performed using the SALSA MLPA Probemixes P034 DMD-1 and P035 DMD-2 kit (MRC Netherlands, Amsterdam, The Netherlands), according to the manufacturer’s instructions. Briefly, 100ng genomic DNA was denatured and hybridized overnight with SALSA probe mixes P034 and P035, followed by ligation with DNA ligase and amplification with specific fluorescent-labeled primers. The amplified products were scanned on the ABI 3500XL Dx Genetic Analyzer (Thermo Fisher Scientific, Waltham, MA, USA) and the data were analyzed using the Coffalyser.Net software, as previously described [24]. The following cutoff values for the final ratio (FR) were used to analyze the copy number of the duplications: normal 0.80 < FR < 1.20, duplication in males (two copies) 1.65 < FR < 2.25, heterozygous duplication in females (three copies) 1.30 < FR < 1.65 and heterozygous triplication/homozygous duplication in females (four copies) 1.75 < FR < 2.15, according to the manufacturer’s updated instructions. All other FR values represented ambiguous copy number of the duplication.

OGM

Ultra-long genomic DNA was extracted using the SP Blood and Cell Culture DNA Isolation Kit (SKU 80030, Bionano Genomics, San Diego, CA, USA), and the DLE-1 enzyme in the Direct Label and Stain DNA Labeling Kit (Bionano Genomics, San Diego, CA, USA) was used as a fluorescently labeled base. Labeled DNA molecules were linearized in nanochannel arrays on the Saphyr chip (Bionano Genomics) and imaged by the Saphyr System (Bionano Genomics), using a high-throughput and automated method. This software automatically converts image data into molecular data and assembles them into a genome-wide optical map. De novo assembly of single molecules into consensus genome maps was performed using Bionano Solve v3.7.1. The variant analysis of Bionano data was performed using two separate pipelines: “CNV pipeline” and “SV pipeline.” The CNV pipeline recovers large and unbalanced aberrations based on normalized coverage. The SV pipeline is used to detect a number of small SVs and compare them with the reference genome map (GRCh38/hg38). The filter settings have been optimized with data from Bionano control sample database following prior analysis of OGM results to reduce the number of variants. SVs were called against using the following confidence values: insertion/deletion ≥ 0, inversion ≥ 0.7, duplications ≥ -1, intra-translocation ≥ 0.05, inter-translocation ≥ 0.05, and CNV ≥ 0.99. For CNV calls, only segments > 500 kb was considered [25].

Long-range PCR and next generation sequencing (NGS)

Long-range PCR was performed to amplify 100ng genomic DNA using TaKaRa LA Taq DNA Polymerase protocol (Takara Bio, Beijing, China) with primers as shown in Table 1. The PCR amplification conditions were as follows: 94 °C for 1 min followed by 30 cycles of 98 °C for 10 s, 64 °C for 24 min, and 72 °C for 10 min and finally, stored at 10 °C. The PCR products were confirmed by 1% agarose gel electrophoresis and were then used to construct the sequencing libraries with Twist EF Library Preparation kit 1, 2.0 (Twist Bioscience, San Francisco, CA, USA), according to the manufacturer’s instructions. The libraries were sequenced on the Illumina NextSeq CN500 platform (Illumina, San Diego, CA, USA), and the sequencing data were analyzed as described previously [26]. GRCh38/hg38 was used as the reference genome.

Sanger sequencing

Specific breakpoints of the complex rearrangements detected by NGS were further confirmed using regular PCR and Sanger sequencing. The PCR was performed using 2 x Taq Plus Master Mix P211 protocol (Vazyme, Nanjing, China) with primers as shown in Table 1. The PCR amplification conditions were as follows: 93 °C for 3 min followed by 35 cycles of 95 °C for 30 s, 60 °C for 30 s, 72 °C for 1 min, and 72 °C for 5 min, and finally, stored at 4 °C. The PCR products were sequenced using the ABI 3500XL Dx Genetic Analyzer (Thermo Fisher Scientific, Waltham, MA, USA), and the data were analyzed using CodonCode Aligner software (CodonCode Corporation, Centerville, Utah, USA). GRCh38/hg38 was used as the reference genome.

Results

Phenotypical differences in the two families

In Family A (Fig. 1A), no history of neuromuscular disease was found. The couple (I-1 and I-2) had two sons (II-1 and II-2), aged 11 and 8 years, respectively. Both boys were normal in terms of movement, walking, and cognition, and no sign of Gower’s or bilateral pseudohypertrophy of the lower legs was observed. The woman (I-2) was pregnant at a gestational age of 24 weeks when she was referred to the clinic and underwent amniocentesis for routine prenatal genetic diagnosis because of advanced age (36 years). Based on comprehensive analysis and interpretation of the genetic findings, the woman decided to continue the pregnancy and delivered a healthy girl (proband II-3).

In Family B (Fig. 2A), the proband (II-2) was an 8-year-old boy with growing proximal limb weakness since the age of 3. The proband exhibited typical phenotype of DMD with signs of Gower’s and bilateral pseudohypertrophy of the calf muscles. Muscle strength examination revealed that his bilateral lower extremities had diminished muscular weakness (trunk muscles G2, hip flexors G3, hip extensors G3−, knee extensors, and knee flexors G3−). Biochemical tests revealed increased levels of lactate dehydrogenase (LDH), creatine kinase (CK), and creatine enzyme isoenzymes (CK-MB) (LDH, 910 U/L; CK, 8941 U/L; and CK-MB, 264 U/L). No abnormalities were noted on electrocardiography or cardiac echocardiography, and his intelligence was within the normal range. The patient had no family history of muscular dystrophy.

Copy-number gains involving DMD identified using CNV-seq, qPCR and/or MLPA

In Family A, a multi-copy duplication with a segmental size of 0.36 Mb involving exons 64–79 of DMD was identified using CNV-seq. The proband (female, II-3) and her mother (I-2) showed a copy number of seven for the segmental repeat (inferred from the estimated average copy number of 7.14 ± 0.86 and 7.03 ± 0.92, respectively), and her elder brothers (II-1 and II-2) had approximately six copies of the repeat (inferred from the estimated average copy number of 5.51 ± 0.88 and 5.63 ± 0.79, respectively), as shown in Fig. 1B. According to the American College of Medical Genetics and Genomics (ACMG) Guidelines based on the evidences (1 A, 2 J, 3 A, 4E, the final point value: 0.2) [27], the duplication can be classified as a variant of uncertain significance. To validate the copy number and the involved exons of the duplication, qPCR and MLPA were performed. In qPCR, when comparing with the control sample (45,X), the target locus Dup309 of the four calibrator samples (45,X; 46,XX; 47,XXX; 46,XY) presented highly consistent copy number as expected, 1.09 ± 0.24, 2.02 ± 0.05, 2.87 ± 0.06, 1.03 ± 0.11, respectively, after normalizing and adjusting the corresponding copy number of the reference locus XP60. For the test samples, the proband’s father (I-1) and mother (I-2) showed a relative copy number of 0.98 ± 0.08, and 6.89 ± 0.56, respectively, for the target locus, while that for her elder brothers was 5.37 ± 0.47 and 6.05 ± 0.22, respectively, as shown in Fig. 1C. MLPA confirmed that the duplication involved exons 64–79 of DMD, with the FR values of 2.88–3.12 in the proband (II-3) and 2.82–3.48 in her mother (I-2), and 4.47–5.58 and 4.43–5.56 in her elder brothers II-1 and II-2, respectively, all of which were far beyond the reference range provided by the manufacturer (Fig. 1D).

In the proband from Family B (II-2), who exhibited typical DMD neuromuscular manifestations, a duplication involving exons 10–13 of DMD with a copy number of two was identified using MLPA (FR value: 1.95–2.04). This variant was inherited from his unaffected mother (I-2), who had a copy number of three for the segment (FR value: 1.50–1.52). The duplication could be pathogenic and presumably disrupted the function of the dystrophin protein, causing the disease (Fig. 2B).

The cryptic complex rearrangements involving DMD determined using OGM

To further explore the potential patterns underlying the identified duplications, OGM was performed on II-2 from Family A and II-2 from Family B. Genomic mapping for II-2 from Family A enabled the construction of a map exhibiting a complex in-cis tandem repeat with the segment covering exon 64–79 of the DMD gene (chrX: 30889742–31236824) (Fig. 3A), and no other conflicting map was constructed. The inferred upstream breakpoint of the segmental repeat was allocated to the closely upstream region of the TAB3 gene (OMIM #300480) with unknown morbid effects, and the downstream breakpoint was located in intron 63 of the DMD gene. With the limitation in the length of the assembled optical reads form OGM, the rearranged haplotype map only presented four complete copies and partial segment of the repeat; However, CNV analysis using OGM showed an estimated copy number of six for the repeat (Fig. 3A), consistent with CNV-seq and qPCR results. As per the pattern of the rearrangement inferred from OGM, at least one complete copy (reading frame) of the DMD gene were most likely to be retained, regardless of the structural changes (Fig. 3B). Thus, the dosage and function of the gene were presumably maintained.

Fig. 3
figure 3

The rearrangement pattern and breakpoints of the complex in-cis tandem repeat in Family A. A. The OGM mapping revealed a complex in-cis tandem repeat involved in DMD. The numbers in the CNV call section indicate the copy number. The blue dashed box represents the partial segment of the repeat that OGM failed to show. The red arrows indicate the breakpoint junctions that will be used for breakpoint analysis. B. The pattern and breakpoint of the rearrangement. The blue line indicates the location of the upstream breakpoint was close to the upstream region of the TAB3 gene. The orange line indicates the location of the downstream breakpoint was in the intron 63 of the DMD gene. “I”: intron, “E”: exon

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The OGM rearranged map for II-2 from Family B showed that a genomic segment from a donor chromosome X (chrX: 32595757–37697377) approximately 5.10 Mb in size was inverted and most likely inserted to another chromosome X (accepter). A genomic segment on the accepter chromosome X (chrX: 32660901–37661984) was replaced by the inserted donor segment (Fig. 4A). The reversed insertion generated an unbalanced chromosome X with a segmental repeat involving exons 10–13 of the DMD gene, exons 1–2 of the XK gene (OMIM#314850), and exons 4–5 of the LANCL3 gene, as shown in Fig. 4B. The rearrangement was predicted to break the complete copy and estimated reading frame of DMD, thereby causing dosage insufficiency and gene dysfunction.

Fig. 4
figure 4

The rearrangement pattern and breakpoints of the unbalanced inversion-insertion rearrangement in Family B. A. The OGM showed an unbalanced inversion-insertion rearrangement involving DMD in Family B. The numbers in the CNV call section indicate the copy numbers. Red and purple arrow indicate the region of the breakpoint verification. B. The pattern of the rearrangement showed that the breakpionts of the complex rearrangement involved exons 10–13 of the DMD gene, exons 1–2 of the XK gene, and exons 4–5 of the LANCL3 gene. “F”: forward primer, “R”: reverse primer

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The specific breakpoints of the complex rearrangements validated using NGS and Sanger sequencing

To determine the potential mechanisms underlying the various functional effects of the complex rearrangements, we further validated the specific breakpoints and investigated the characteristics of breakpoint junctions using NGS combined with long-range PCR amplification and Sanger sequencing. The rearrangement involving a multi-copy in-cis tandem repeat in Family A formed an identical breakpoint junction, whose upstream and downstream sequences could be mapped to chrX: 30886535 and chrX: 31238575, respectively, as shown in Fig. 5A and B. Sanger sequencing confirmed the results, but the downstream breakpoint can be mapped to chrX: 31238573 or chrX: 31238575 (Fig. 5C and D) because the dinucleotide repeat “GT” surrounding the breakpoint was homologous between the DMD and TAB3 gene sequences. Based on these findings, further reassessment of the variation was performed according to the evidences (1 A, 2 J, 3 A, 4 K, the final point value: -0.6) from the ACMG guidelines. Although the variation was still reclassified as a variant of uncertain significance, the evidences applicable and the final point values had changed: the evidence (4E) was not appropriate and the evidence (4 K) was adopted, and the corresponding point values changed from 0.2 to -0.6.

Fig. 5
figure 5

Breakpoint analysis of the rearrangement of Family A. A. Gel analysis of the long-range PCR product; The red arrow indicates the long-range PCR product. Control: healthy control; kb: kilobase. B. The breakpoints were validated using NGS. The integrative genomics viewer (IGV) screenshot showed the locations of breakpoint junctions. C. Gel analysis of the regular PCR product. The red arrow indicates the regular PCR product. Control, healthy control; bp, base pair. D. The result of Sanger sequencing. The precise breakpoint junctions located in intron 63 of the DMD gene and intron 1 of TAB3 gene. The red dashed box represents homologous base

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The unbalanced inversion-insertion rearrangement in Family B generated two breakpoint junctions. According to the NGS results, the breakpoint junction 1 can be mapped to chrX: 32664240 and chrX: 37705288 (Fig. 6A and B), respectively, and the breakpoint junction 2 can be mapped to chrX: 32587504 and chrX: 37656582 (Fig. 7A and B), respectively. These were supported by the findings from Sanger sequencing, which further characterized the signatures of the breakpoint junctions. A 19-bp microhomology “AGTAGCTGGGACTACAGGC” was identified in the breakpoint junction 1 (chrX: 32664240 and chrX: 37705288) (Fig. 6C and D), and a nucleotide “T” was inserted in the breakpoint junction 2 (chrX: 32587504 and chrX: 37656582) (Fig. 7C and D).

Fig. 6
figure 6

Breakpoint analysis in breakpoint junction 1 of rearrangement in Family B. A. Gel analysis of the long-range PCR product. The red arrow indicates the PCR product. Control, healthy control; kb, kilobase. B. The breakpoint was validated using NGS. The IGV screenshot showed the locations of breakpoint junction 1. C. Gel analysis of the long-range PCR product. The red arrow indicates the regular PCR product. C, healthy control; bp, base pair. D. The results of Sanger sequencing showed that the precise locations of breakpoint junctions 1 were in DMD intron 9 and XK intron 1. The red dashed box represents microhomology sequences

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Fig. 7
figure 7

Breakpoint analysis in breakpoint junction 2 of the rearrangement in Family B. A. Gel analysis of the long-range PCR product. The red arrow indicates the long-range PCR product. Control, healthy controls; kb, kilobase. B. The breakpoint was validated using NGS. The IGV screenshot showed that the locations of breakpoint junction 2. C. Gel analysis of the regular PCR product. The red arrow indicates the regular PCR product. C, healthy control; bp, base pair. D. The results of Sanger sequencing showed that the precise locations of breakpoint junctions 2 were in DMD intron 13 and LANCL3 intron 2. The purple dashed box represents the inserted nucleotide

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Discussion

In this study, we explored the potential rearrangement patterns involving DMD duplications in two families presenting phenotypical differences and analyzed the relationship between the rearrangement patterns and phenotypes. We identified a multi-copy (estimated six copies in males) duplication involving exons 64–79 of DMD in Family A and a duplication involving exons 10–13 of DMD in Family B, using CNV-seq, qPCR and/or MLPA. However, Family B showed DMD phenotypes, whereas Family A exhibited neither DMD nor BMD manifestation. To the best of our knowledge, this is the first report of duplications with more than four copies involving DMD but unaffected by the disease.

Exon(s) duplication in the DMD gene is typically regarded as pathogenic due to its impact on the reading frame and subsequent production of pathogenic truncated proteins [28, 29]. Recent studies have shown that not every duplication involving DMD exons(s) is pathogenic [29, 30]. Three non-contiguous duplications in exons 51–53, exons 64–79, and intron 55 of the DMD gene were identified in a family with an asymptomatic phenotype, and further breakpoint analysis revealed the extra copies involved in the complex rearrangement were located outside the DMD gene region [29]. Individuals with duplications involving the same exons of the DMD gene could present variable phenotypes, which can be explained by the different breakpoint junctions of the rearrangements (extragenic or intragenic) [30]. In this study, we initially investigated the possible connection between DMD duplication and the phenotypic variations, using OGM. Our findings show that the duplication in Family A displayed an in-cis tandem repeat containing an intact DMD copy, whereas Family B exhibited a reversed insertion with two segmental repeats. Using long-range PCR and NGS for breakpoint analysis, we discovered that the rearrangement in Family A occurred outside the DMD gene, thereby presumably maintaining the integrity of the DMD reading frame. In contrast, in Family B, the rearrangement disrupted the DMD reading frame, thereby probably contributing to the loss of dystrophin function. The differences in rearrangement features and integrity of the reading frames further explained the phenotypic variations between the two families, suggesting that the effect of segmental duplications is not only due to changes in copy number (dosage effect of the gene) but also possibly due to its location or pattern (position effect of the gene), thus resulting in the disease. This is also supported by previous findings from patients with BMD, in which an inversion involving DMD disrupted the gene structure [31]. Therefore, analyzes of rearrangement patterns and breakpoint locations are crucial for considering gene dosage and position effects, which provide valuable evidence for pathogenic classification and vital suggestions for genetic diagnosis and counseling. This is particularly true for carriers without any family history of DMD or BMD and for fetuses who exhibit rearrangements involving DMD inferred from the prenatal testing. More importantly, the further phenotypical reassessments and experimental validations for the function and dosage of dystrophin in the two families are of great significance to illustrate the genotype-phenotype correlation. Unfortunately, the subjects and their parents refused further clinical evaluations and examinations, and extra blood resampling or muscle biopsy for RNA analysis or protein testing. Additionally, long-term follow-ups and further investigations on more similar cases are still required.

Genomic recombination has been associated with at least five different mechanisms: homologous recombination, non-homologous end joining (NHEJ), microhomology-mediated replication-dependent recombination (MMRDR), retrotransposition mediated by long interspersed element-1, and telomere repair [32, 33]. NHEJ and MMRDR are the main mechanisms involved in the formation of complex rearrangements [32, 34]. In this study, microhomology sequences were found at the breakpoints of the rearrangement identified in Family A and the breakpoint junction 1 of the rearrangement discovered in Family B, thereby implying that these complex rearrangements may be mediated via MMRDR. Furthermore, a single nucleotide “T” was inserted at the breakpoint junction 2 of the rearrangement found in Family B, which is consistent with the NHEJ-mediated pattern. Additionally, the novel structural rearrangements may have arisen during meiotic recombination in the oocyte.

Compared to the traditional assays, such as CNV-seq, MLPA, qPCR, etc., OGM has distinct advantage in identifying balanced genomic rearrangements and profiling the pattern of balanced or unbalanced structural changes. Owing to its genome-wide molecular labels with high resolution [19], the breakpoints or junctions of the complex rearrangements can be easily located and mapped, facilitating further interpretation of the functional effect driven by these structural changes. Regardless of this, validating the precise breakpoints and calling the detailed junction sequences by other methodologies is still required. For example, in Family A from this study, OGM indicated that one of the breakpoints was allocated to the closely upstream region of the TAB3 gene, however, NGS and Sanger sequencing revealed that the actual breakpoint was mapped to intron 1 of TAB3. This discrepancy arises because OGM relies on base-specific labeling rather than base sequencing to identify the location of the rearrangement breakpoint [35]. Furthermore, we discovered that the existing OGM’s capacity to construct and visualize the intact rearrangements maps could decrease when faced with complex structural changes involving oversize genomic segments, just as it does not display the entire pattern of the rearrangements in the Family A (estimated six copies of in-cis tandem repeats for a ~ 0.36-Mb segment).

Conclusions

We identified cryptic complex rearrangements involving the DMD gene causing phenotypical differences and investigated the possible patterns underlying the formation of these complex structures. Validating breakpoints and rearrangement patterns is crucial for the functional annotation and pathogenic classification of genomic structural variations. OGM provides valuable insights into etiological analysis of DMD/BMD and enhances our understanding for cryptic effects of complex rearrangements related to DMD/BMD or other similar disorders.

Data availability

No datasets were generated or analysed during the current study.

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Acknowledgements

We would like to thank the families for participating in this study.

Funding

This work was supported in part by the National Natural Science Foundation of China (82001531 and 81860272 to BG), Guangxi Major Research Programme (AB22035013 to BG), Guangxi Natural Science Foundation (2023GXNSFBA026124 to CG, 2018GXNSFAA281067 to BG), Initial Scientific Research Fund for Advanced Talents from the Second Affiliated Hospital of Guangxi Medical University (2019112 to BG), Special Scientific Research Fund of Guangxi Ten-Hundred-Thousand Talents Project (2021186 to BG), the Guangxi Medical University Training Program for Distinguished Young Scholars (to BG as team member), the Science Foundation for Young Scholars of Guangxi Medical University (GXMUYSF202115 to CG), and Innovation Project of Guangxi Graduate Education (YCSW2022221 to YM, YCSW2023239 to XZ).

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Author notes

  1. Yunting Ma, Chunrong Gui and Meizhen Shi contributed equally to this work.

Authors and Affiliations

  1. The Second School of Medicine, Guangxi Medical University, No. 166, Daxuedong Road, Xixiangtang District, Nanning, Guangxi Zhuang Autonomous Region, 530007, China

    Yunting Ma, Zifeng Cheng, Xu Zhou, Jiefeng Luo, Yan Huang & Baoheng Gui

  2. Center for Medical Genetics and Genomics, The Second Affiliated Hospital of Guangxi Medical University, No. 166, Daxuedong Road, Xixiangtang District, Nanning, Guangxi Zhuang Autonomous Region, 530007, China

    Chunrong Gui, Meizhen Shi, Bobo Xie, Haiyang Zheng, Xiaoyun Lei, Xianda Wei & Baoheng Gui

  3. The Guangxi Health Commission Key Laboratory of Medical Genetics and Genomics, The Second Affiliated Hospital of Guangxi Medical University, No. 166, Daxuedong Road, Xixiangtang District, Nanning, Guangxi Zhuang Autonomous Region, 530007, China

    Chunrong Gui, Meizhen Shi, Bobo Xie, Haiyang Zheng, Xiaoyun Lei, Xianda Wei, Jiefeng Luo, Yan Huang & Baoheng Gui

  4. Department of Obstetrics, The Second Affiliated Hospital of Guangxi Medical University, No. 166, Daxuedong Road, Xixiangtang District, Nanning, Guangxi Zhuang Autonomous Region, 530007, China

    Lilin Wei & Yan Huang

  5. Department of Rehabilitation Medicine, The Second Affiliated Hospital of Guilin Medical University, No. 212, Renmin Road, Lingui District, Guilin, Guangxi Zhuang Autonomous Region, 541100, China

    Junfang He

  6. Department of Pediatrics, The Second Affiliated Hospital of Guangxi Medical University, No. 166, Daxuedong Road, Xixiangtang District, Nanning, Guangxi Zhuang Autonomous Region, 530007, China

    Shaoke Chen

  7. Department of Neurology, The Second Affiliated Hospital of Guangxi Medical University, No. 166, Daxuedong Road, Xixiangtang District, Nanning, Guangxi Zhuang Autonomous Region, 530007, China

    Jiefeng Luo

Authors
  1. Yunting Ma
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Contributions

LW, JH and YM collected and refined the clinical data. YM and XZ extracted DNA. MS, CG and HZ analyzed CNV-seq data. YM, MS and ZC completed qPCR experiment and analysis. YM, BX and BG completed MLPA experiment and analyzed. YM, CG and BG analysis OGM data. MS, XL and XW analyzed NGS data. YM, CG and MS designed primers and completed PCR experiments. YM, CG and MS wrote the manuscript. SC, JL and YH supervised and validated the research progress. BG reviewed and revised the manuscript. All authors have checked and approved the final manuscript.

Corresponding authors

Correspondence to Jiefeng Luo, Yan Huang or Baoheng Gui.

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Ethics approval and consent to participate

The studies involving human participants were reviewed and approved by the Ethics Committee of the Second Affiliated Hospital of Guangxi Medical University (NO. 2022-KY0632). Written informed consent was obtained from the participants or their legal guardian.

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The authors declare no competing interests.

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Ma, Y., Gui, C., Shi, M. et al. The cryptic complex rearrangements involving the DMD gene: etiologic clues about phenotypical differences revealed by optical genome mapping. Hum Genomics 18, 103 (2024). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s40246-024-00653-1

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Keywords

Human Genomics

ISSN: 1479-7364