{"paper_id":"84008458-1104-4acf-adda-f6f6cddf6527","body_text":"Osteogenesis imperfecta (OI ) is a group of genetically related connective tissue disorders, diverse in terms of both genotypes and phenotypes, characterized by increased bone fragility. Clinically, five types of  OI  ranging from mild (type I) to moderate (types III and IV) and severe or lethal (type II) are distinguished [ 1 , 2 , 3 ]. So far, numerous mutations in genes encoding collagen type I (Col I) or involved in collagen modification, e.g., cartilage-associated protein (CRTAP) and leucine proline-enriched proteoglycan 1 (LEPRE1) [ 4 , 5 ]; folding, e.g., serpin peptidase inhibitor (SERPINH1) [ 6 ]; and processing, e.g., bone morphogenetic protein 1 (BMP1) [ 7 ], have been shown to cause various types of  OI . Nevertheless, a majority of  OI  cases inherited in an autosomal-dominant manner are caused by mutations in the collagen type I α1 or α2 encoding genes ( COL1A1  and  COL1A2 , respectively), resulting in a reduced amount or abnormal structure of collagen [ 8 , 9 , 10 ].\nThe earliest stage when an  OI  diagnosis can be made is the prenatal period. It depends on the severity of the  OI  and is based on family history; genetic tests, i.e., sequencing of genomic DNA and specific mutation identification; and prenatal tests, i.e., a decrease in the femoral length (FL) [ 11 ]. In the post-natal period, positive family history and pivotal clinical symptom (bone fragility, typical features of blue sclera) notification with bone density measurement (BMD) implementation are routinely used in  OI  diagnosis [ 12 , 13 ]. Additional diagnostic methods include magnetic resonance imaging (MRI), radiography of the uterus, and other genetic tests, such as multiplex ligation-dependent probe amplification (MLPA) or chorionic villus sampling (CVS) [ 12 , 14 ]. If the typical clinical symptoms and positive family history are noted by the clinician, routine diagnostic methods are sufficient to diagnose  OI ; however, about 10–15% of patients suffer from less common variants of the disease. Moreover, any diagnostic method used may be clear-cut and have drawbacks, e.g., employment of the FL carries the risk of other skeletal dysplasia’s recognition instead of  OI  [ 13 , 15 ]. In such circumstances, further procedures, mostly based on molecular diagnostics, are needed [ 11 ].\nMicroRNA (miRNA) constitutes short, about 20-nucleotide, single-stranded, and endogenous RNA molecules that have been observed to negatively regulate gene expression in two manners: i) by attaching to their target messenger RNA (mRNA) at different mRNA sites, most often 100% complementary, and cutting it; or ii) by binding to the 3′-untranslated region (3′UTR) of the target mRNA and inhibiting translation, an action that does not require 100% complementarity [ 16 ]. The principal role of miRNAs is the regulation of a number of biological processes, such as embryogenesis, apoptosis, hosting immune responses during infections, or cancerogenesis [ 16 ]. miRNAs fulfill a crucial role in the regulation of gene expression and are involved in bone formation and bone diseases [ 17 , 18 , 19 ]. For instance, miR-217 has been reported to promote cell proliferation and osteogenic differentiation during the development of steroid-associated osteonecrosis in bone marrow mesenchymal stem cells [ 20 , 21 ], whereas miR-200a-3p has been linked with the synthesis of ECM components in fibrotic diseases [ 22 ].\nOver the past two decades, the mechanisms of human extracellular matrix (ECM) regulation by miRNAs have been extensively studied, including the genes involved in collagen synthesis. For example, the negative influence of hsa-miR-133a-3p, which has been assigned a possible antifibrotic role [ 23 , 24 ]; hsa-miR-27b-3p, which increases collagen expression [ 25 ]; and hsa-miR-382-5p, which promotes epidural fibrosis [ 26 , 27 ], has been demonstrated. Furthermore, many miRNAs that can regulate genes involved in the enzymatic processing or degradation of natively synthesized collagen have been recently discovered. For example, hsa-miR-377-3p has been shown to inhibit the synthesis of matrix metalloproteinase-2 (MMP-2) and MMP-16, whereas hsa-miR-200b-3p has been proven to regulate tissue inhibition of metalloproteinase-2 (TIMP2) [ 26 ]. Moreover, miRNAs can affect signaling pathways involved in ECM synthesis and degradation; e.g., hsa-miR-216a-5p has been shown to regulate Janus kinase 2 (JAK2) [ 28 , 29 ], while hsa-miR-25-3p has been shown to control the transforming growth factor β receptor I (TGFβR1) and neurogenic locus notch homolog protein 1 (NOTCH1) signaling pathways [ 30 , 31 ]. To date, however, only a few miRNAs, such as let-7a-5p, miR-92a, and miR-29b, have been studied in  OI  [ 17 , 18 ].\nThe aim of this study was to identify and analyze the expression profiles of miRNAs in osteogenesis imperfecta ( OI ) type I caused by mutations in the  COL1A1  and  COL1A2  genes.\n\nOsteogenesis imperfecta  type I in the majority of cases (~95%) is caused by mutations in the  COL1A1  and  COL1A2  genes; hence, in this study, we focused on sequencing analyses of those genes [ 1 , 2 ]. Mutations in  COL1A1  were found in 8 study subjects and in  COL1A2  in 2 out of 10 study subjects diagnosed with  OI  type I. The missense mutations were detected in both  COL1A1  (donor nos. 1–4) and  COL1A2  (donor nos. 9 and 10). In  COL1A1 , other mutation types were also found, including a single-nucleotide deletion resulting in a frameshift in three donors (nos. 5–7) and an exon skipping in one patient (no. 8). The missense mutations detected in the  COL1A1  gene resulted in three different substitutions, namely glycine by alanine in exon 17 (patient no. 1), glycine by valine in exon 9 (patient no. 2), and a third one also found in exon 17 that resulted in valine-to-phenylalanine alteration (patient no. 3). Additionally, in two other patients (nos. 6 and 7), two deletions of T located in exon 30 and exon 17 of the  COL1A1  gene resulted in the production of an altered protein. In contrast, both missense mutations in the  COL1A2  gene resulted in the substitution of glycine by serine in exon 31 (patient no. 9) or exon 38 (patient no. 10).\nMoreover, we identified a series of new mutations in both genes: substitution of guanine for thymine in exon 9 in patient no. 2 and deletion of 10 nucleotides in exon 50 (frameshift) adjacent to a missense mutation (substitution of glutamine by valine) in  OI  patient no. 4, both in the  COL1A1  gene. Another alteration comprised a guanine deletion in exon 2 of the  COL1A1  gene and was identified in patient no. 5. A new mutation found in the  COL1A2  gene was represented by a substitution of guanine for adenine in exon 31 of patient no. 9.\nA brief clinical description of  OI  patients harboring previously reported mutations as well as the new mutations in  COL1A1  and  COL1A2  identified in this study are presented in  Table 1 .\nFirst, we performed identification analyses of the miRNAs in the  OI -derived cells and control cells. As shown in  Table 2 , the results were categorized into the three following parts: (A) miRNAs detected in  OI  but not in the control fibroblasts; (B) miRNAs detected in both  OI  and the control cells; (C) miRNAs not detected in  OI  nor the control cells.\nAs presented in  Table 2 , nine specific miRNAs, i.e., hsa-miR-19a-3p, hsa-miR-19b-3p, hsa-miR-133a-3p, hsa-miR-200b-3p, hsa-miR-204-5p, 216a-5p, hsa-miR-377-3p, hsa-miR-449a, and hsa-miR-590-5p, were identified in the fibroblasts of 30% to 70% of  OI  donors, but not in the control fibroblasts. Other 26 miRNA molecules were detected in both the  OI  and the control cells, with the percentage of  OI  donors with a specific miRNA detection ranging from 70% to 100%. Finally, five miRNAs selected for analysis, i.e., hsa-miR-1-3p, hsa-miR-141-3p, hsa-miR-142-3p, hsa-miR-200a-3p, and hsa-miR-217, were not detected in the  OI  or the control cells.\nBased on the miRNA identification, an expression analysis was performed for miRNAs detected in >50% of  OI  patients and compared with the controls. As depicted in  Figure 1 , increased expression was observed in the case of six miRNAs (hsa-miR-10b-5p, hsa-miR-19a-3p, hsa-miR-19b-3p, has-miR-204-5p, has-miR-216a-5p, and hsa-miR-449a), while decreased expression was observed in the case of four miRNAs (hsa-miR-129-5p, hsa-miR-199b-5p, hsa-miR-664a-5p, and hsa-miR-30a-5p) in  OI  as compared to the controls. Among the miRNAs analyzed, miR-449a’s expression increased about 20-fold, followed by an almost 5-fold increase in miR-19b-3p and a ~4-fold increase in miR-19a-3p. Meanwhile, an almost 6-fold decrease in expression in  OI  was found in the case of miR-664a-5p, whereas a slightly less decrease was observed for miR-129-5p (3.6-fold), miR-199b-5p (2.8-fold), miR-30a-5p (2.2-fold), and miR-25-3p (1.9-fold) ( Table 3 ).\nAmong the miRNAs that were overexpressed in the  OI  cells, two sets displaying moderate-to-strong positive correlations within each group were observed ( Table 4 ). The first group consisted of miR-10a-5p, miR-10b-5p, miR-199a-5p, and miR-382-5p, and the second consisted of miR-19a-3p, miR-19b-3p, miR-204-5p, miR-216a-5p, and miR-449a. miR-17-5p, on the other hand, did not show strong correlations with any of the overexpressed miRNAs.\nA significantly decreased expression in the  OI  group was noted for 13 miRNAs ( Table 5 ). In this group, an almost complete correlation was found for miR-21-5p, miR-25-3p, miR-26a-5p, miR-26b-5p, miR-27a-3p, miR-27b-3p, miR-29a-3p, and miR-29c-3p, which have been previously reported to regulate the expression of genes associated with fibrosis (cystic fibrosis in children or liver fibrosis) and cancerogenesis [ 24 , 25 , 48 , 49 , 56 ]. Further, miR-30a-5p showed a strong correlation with almost all miRNAs presented in  Table 5 .\nIn contrast, a moderately negative correlation of miR-129-5p with miR-21-5p, miR-26b-5p, miR-29a-3p, and miR-29c-3p was observed. This may indicate an antagonistic function of miR-129-5p in the regulation of collagen synthesis. Finally, the weakest correlations with other tested downregulated miRNAs were observed for miR-199b-5p and miR-29b-3p ( Table 5 ).\nIn the group of  OI  patients harboring mutations in  COL1A1 , upregulated expression of miR-17-5p, miR-21-5p, miR-26b-3p, miR27a-3p, miR27b-3p, miR29a-3p, mir29c-3p, miR-30a-5p, miR-34a-5p, miR-92a-3p, and miR143-3p was found. In contrast, in the  OI  caused by mutations in  COL1A2 , only miR-25-3p was found to be upregulated ( Table 6 ).\nInterestingly, the expression of miR-132-3p was comparable in the controls and the  COL1A1 -mutated fibroblasts but increased in the group of  OI  caused by mutations in  COL1A2  ( Table 6 ). These observations may suggest that miR-25-3p and miR-132-3p could play a particularly significant role in the pathogenetic processes of  OI  induced by mutations in the  COL1A2  gene.\nConsidering the type of mutation, it was noted that patients no. 1, 2, and 3 with missense mutations in the  COL1A1  gene revealed similar overexpression patterns for several miRNAs, indicating that these mutations may have similar effects on miRNA regulation. Patients no. 4, 6, and 7, with frameshift mutations in the  COL1A1  gene, displayed similar overexpression patterns for some miRNAs but also individually dependent differences among particular types of miRNAs, and the profiles of expression also varied from those found in  OI  with missense mutations. In turn, patient no. 5, with a frameshift mutation in the  COL1A1  gene, had a unique expression pattern of overexpressed and downregulated miRNAs. In summary, the miRNA expression patterns observed in the individually analyzed  OI  patients indicate that particular mutations may influence the regulation of specific miRNAs, potentially leading to downstream effects on genes involved in bone development and maintenance ( Table 6 ).\n\nDespite similar and uniform phenotypic symptoms, the molecular mechanisms of pathogenetic processes contributing to  OI  remain unclear. This disorder is difficult to treat as it can be caused not only by direct defects in the amount of type I collagen produced but also by the influence of various proteins and regulatory RNA molecules, like miRNAs, on the processing of synthesized collagen, including the pool of aberrant proteins.\nAround 5% of  OI  cases have an unknown genetic background. DNA sequencing analyses revealed the presence of mutations in both genes of all  OI  donors enrolled in this study, including four novel and previously unreported, of which three were identified in the  COL1A1  gene and one in the  COL1A2  gene. The missense mutations detected in the  COL1A1  and  COL1A2  genes resulted in the substitution of glycine and other amino acids, leading to disruptions in the collagen chain interactions and destabilization of the collagen triple helix [ 1 , 76 ]. Frameshift mutations, including single-nucleotide deletion, ten-nucleotide deletion, and exon skipping in the  COL1A1  gene, altered the reading frame, resulting in truncated or absent Col I chains. Additionally, a single-nucleotide deletion in exon 2 of  COL1A1  disrupted the formation of the collagen triple helix. These mutations likely contribute to the development of  OI  by affecting the stability and packing of the collagen molecule [ 77 ]. The mutations identified in this study align with previous reports on  OI , reaffirming that mutations in the  COL1A1  and  COL1A2  genes are responsible for the majority of  OI  cases [ 78 , 79 ].\nThis research focused on the investigation of the miRNAs that regulate the expression of the ECM, including Col I processing and trafficking in the context of  OI  pathogenesis. Nine miRNAs that were identified in  OI  but not in the controls, namely hsa-miR-19a-3p, hsa-miR-19b-3p, hsa-miR-133a-3p, hsa-miR-200b-3p, hsa-miR-204-5p, 216a-5p, hsa-miR-377-3p, hsa-miR-449a, and hsa-miR-590-5p, have been previously reported to regulate the TGF-β and Wnt signaling pathways, indicating their essential role in the regulation of genes involved in bone formation and turnover [ 28 , 29 , 32 , 33 , 34 ].\nFive out of forty analyzed miRNAs, i.e., hsa-miR-1-3p, hsa-miR-141-3p, hsa-miR-142-3p, hsa-miR-200a-3p, and hsa-miR-217, were absent in the  OI  group and the controls. These molecules affected suppressor genes, such as the phosphatase and tensin homolog ( PTEN)  and the dickkopf WNT signaling pathway inhibitor 1 ( DKK1) , as well as genes involved in ECM turnover, i.e., Rho-associated coiled-coil-containing protein kinase 2 ( ROCK2)  [ 20 , 21 , 22 , 71 , 72 , 73 , 74 , 75 ]. These miRNAs have not been directly linked to genes related to collagen synthesis and secretion or connective tissue diseases; however, they participate in the inhibition of the epithelial–mesenchymal transition in renal tubular epithelial cells (e.g., miR-141-3p) and the promotion of cell proliferation and osteogenic differentiation during the development of steroid-associated osteonecrosis in bone marrow mesenchymal stem cells (e.g., miR-217) [ 21 , 73 ] ( Table 2 ). Moreover, hsa-miR-200a-3p has been found to participate in the overproduction of ECM proteins in fibrotic diseases.\nThe involvement of miRNAs in ECM regulation has recently become an area of intense research, as their ability to modulate gene expression post-transcriptionally makes them potent regulators of crucial cellular processes [ 17 , 18 , 19 ]. Understanding how these miRNAs affect ECM components will be of great importance in the search for new therapeutic targets for therapies for diseases associated with ECM dysregulation.\nThe expression analysis performed herein showed altered expression of 10 miRNAs in the fibroblasts of  OI  patients compared to the controls. The expression of six, i.e., hsa-miR-10b-5p, hsa-miR-19a-3p, hsa-miR-19b-3p, has-miR-204-5p, has-miR-216a-5p, and hsa-miR-449a, was substantially increased, while the expression of four (hsa-miR-129-5p, hsa-miR-199b-5p, hsa-miR-664a-5p, and hsa-miR-30a-5p) decreased.\nFurther, the fold change analysis provided robust validation of the observed miRNA expression alterations in  OI  compared to the controls, displaying a significant multiplication in up- or downregulation. Remarkably, miR-449a’s expression increased about 20-fold, while miR-449a was exclusively detected in  OI  patients, which may suggest the potential relevance of both miRNAs in the pathogenesis of  OI .\nIn contrast, we noted a six-fold expression decrease in miR-664a-5p. Though this miRNA has been implicated in the modulation of tumor suppressor p53 and the expression of p53 target genes and has recently been associated with osteosarcoma [ 70 ], this finding adds an intriguing dimension to this study, hinting at potential links between  OI  and p53-related pathways that warrant further investigation.\nThe intricate relationships between miRNAs may offer a unique opportunity to establish connections between molecular processes, providing insights into the pathogenesis of diverse diseases. For better clarification of the impact of miRNAs on bone ECM changes in  OI , we explored their potential correlations, whether positive or negative. Correlation analyses were carried out within the groups with upregulated and downregulated expressions of miRNAs. The miRNAs overexpressed in  OI  that strongly correlate with each other are involved in the regulation of osteoblast differentiation and osteogenesis [ 26 , 27 , 44 , 45 , 46 , 66 , 67 ], while the miRNAs presented in the latter group have been previously associated with various biological processes and diseases, including cancer, inflammation, and neuronal differentiation [ 28 , 29 , 32 , 33 , 34 , 36 , 37 , 38 ]. With the exception of miR-129-5p, which showed a moderate negative correlation with four other miRNAs associated with various cancers, both groups of analyzed miRNAs revealed positive correlations with miR-29 family members [ 49 , 52 , 56 ]. The miR-29 family negatively regulates collagen expression by directly targeting  COL1A1 ’s and  COL1A2 ’s mRNA transcripts, while the other miRNAs are linked to fibrosis and cancers, suggesting their effect on the synthesis of the ECM [ 30 , 31 , 48 , 49 , 50 , 51 ].\nIn the present study, 40 miRNAs in total were analyzed in 10  OI  donors in whom mutations, including four unreported ones, were identified in  COL1A1 / COL1A2 . Overexpression of miR-10b-5p and miR-382-5p in some patients suggests their involvement in the pathogenesis of  OI . On the other hand, decreased expression of miR-129-5p, miR-199b-5p, miR-29b-3p, and miR-664a-5p in most patients suggests their importance in normal bone development and function ( Table 6 ). Kaneto et al. reported decreased expression of miR-29b during the osteoblastic differentiation of mesenchymal stem cells (MSCs). Likewise, in our research, we observed a modest decrease in the expression of miR-29b in skin fibroblasts from a group of patients diagnosed with  OI  type I. Upon conducting separate analyses for each  OI  patient, we observed an upregulation of miR-29b expression in certain individuals, which contrasts with the overall trend observed in the entire  OI  group. This variability may be attributed to the diverse range of patient demographics, including age and gender. miR-29 family members, including miR-29a, miR-29b, and miR-29c, have been implicated in the regulation of ECM proteins, such as collagen, which are important for bone development and homeostasis [ 52 , 53 , 54 , 55 , 56 ]. The aberrant regulation of those miRNAs in patients with  OI  type I induced by mutations in  COL1A1  may contribute to reduced collagen synthesis and impaired bone formation. Regarding  OI  patients harboring mutations in the  COL1A2  gene, we observed overexpression of miRNA-25-3p. Noticeably, this miRNA has been implicated in the regulation of bone mineralization and may play a role in osteoblast differentiation [ 30 , 31 ]. We also observed some discrepancies in the expression of certain miRNAs, such as let-7a, which was found to be overexpressed in some patients with missense mutations, but its expression was downregulated in patients with an exon-skipping mutation. Interestingly, one patient (no. 5) with a novel identified mutation that resulted in a frameshift in exon 2 of the  COL1A1  gene displayed a distinct miRNA expression pattern compared to other examined patients, which may suggest a unique background of  OI  in this individual.\nOne of the potential therapies for  OI  based on targeting the miRNAs involved in the regulation of bone formation and remodeling can be applied by either increasing the expression of beneficial miRNAs or decreasing the expression of harmful ones [ 80 , 81 , 82 ]. However, there are several challenges that need to be addressed before its implementation into clinical practice, including: (i) the identification of disease-specific and most effective targeted miRNAs (as discussed in the current work); (ii) the validation of relevant delivery methods that would not only effectively “reach” the targeted micromolecule but also efficiently deliver miRNAs to the bone; and (iii) the overcoming of the immune response to exogenous miRNAs [ 83 ].\n\nDespite similar and uniform phenotypic symptoms in individuals with  OI , the molecular basis of this disease seems to be varied and still unclear. Moreover, it is difficult to treat, as it can be caused not only by direct defects in the amount of type I collagen produced but also by the influence of various proteins and regulatory RNA molecules, including miRNAs, on the processing of synthesized collagen and the pool of aberrant proteins. Therefore, understanding the function of miRNAs that can affect collagen or signaling pathways that indirectly regulate the amount or quality of collagen is extremely important and necessary. Overall, the current study underscores the importance of the ECM in the development of  OI  and other genetic diseases of connective tissue. The identified mutations and miRNA expression profiles shed light on the intricate processes governing bone formation and ECM regulation, paving the way for further research and potential therapeutic advancements in  OI  and other genetic diseases related to bone abnormality management.\n\nThis study was carried out in accordance with the Declaration of Helsinki and was approved by the Bioethics Committee of the Jagiellonian University in Krakow, Poland (protocol no. DK/KB/CM/0031/689/2010). Clinical examination and medical histories of the subjects were taken by a clinician from the Chair of Pediatrics, Department of Medical Genetics, UJCM, Krakow, Poland. All subjects (or representing individuals) read and signed a written informed consent form prior to inclusion in this study.\nTen subjects (ranging in age from 0.5 to 29 years, of both sexes, and diagnosed with  OI  type I) were enrolled in this study. The diagnosis was confirmed on the basis of multiple (from ~9 to 20) bone fractures in the post-natal period and/or blue sclera or osteoporosis. The inclusion criterion was a clinically confirmed diagnosis of  OI  type I. Exclusion criteria included the clinical diagnosis of other types of  OI  and absence of mutations in the  COL1A1  and/or  COL1A2  genes. A clinical and molecular description of the  OI  type I subjects enrolled in this study is presented in  Table 1 . As a control, the human fibroblast cell line BJ (CRL-2522, ATCC, Manassas, VA, USA) was used.\nSkin biopsy samples were collected from 10 patients diagnosed with  OI  type I. To establish a fibroblast culture, skin biopsies of a size of about 3–5 mm 2  were cut into smaller pieces and placed in a Petri dish with 10 mL of Dulbecco’s Modified Eagle’s Medium (DMEM), supplemented with 4.5 g/L glucose, 10% fetal calf serum (FCS), 50 μg/mL streptomycin, 50 U/mL penicillin, and 25 µg/mL amphotericin B (PAA, Austria). After one week, the remaining parts of the skin explants were removed from the dish, and the fibroblast culture was kept until 90% confluence. Subsequently, the cells were harvested using trypsin/EDTA, plated in fresh DMEM with 4.5 g/L glucose, 10% FCS, streptomycin, penicillin, and amphotericin B in plastic flasks, and kept until homogeneous cultures were obtained. The homogeneity of the fibroblast culture was confirmed after first passage by immunofluorescent staining against vimentin. Cell viability amounted to 98% using Trypan blue.\nFor the experiments, 2 × 105 cells per well were plated in 6-well culture plates and maintained for 7 days in DMEM with 4% FBS supplemented every second day with a fresh L-ascorbic acid 2-phosphate sesquimagnesium salt hydrate (Sigma-Aldrich, St. Louis, MO, USA) at a concentration of 40 μg/mL. The experiments were carried out between the third and sixth passages. Control human fibroblast cell line BJ (CRL-2522, ATCC, Manassas, VA, USA) was cultured and treated identically as the primary skin fibroblasts recovered from the study group.\nLysis and DNA isolation from primary skin fibroblasts and the control cell line were conducted according to the manufacturer’s protocol using DNA purification kit Blood Mini (A&A Biotechnology, Gdansk, Poland). The DNA amplification reactions were conducted using FastStartTaq DNA polymerase kit (Roche, Basel, Switzerland). The composition of the reaction mixture and conditions used are described elsewhere [ 84 , 85 ]. Briefly, for each gene, 10 amplification reactions were conducted, and the DNA fragments were visualized in 1% agarose gel (Agarose, LE Analytical Grade, Promega, Madison, WI, USA) in TAE buffer.\nSequencing PCR for the modified Sanger’s enzymatic method was conducted according to the recommendation provided by the manufacturer of the ABI Prism 3130xl DNA Analyzer. In brief, 20 ng of each purified amplified PCR product was mixed with 5 pM of the appropriate sequencing primer and Big Dye Terminator v.3.1, as previously described [ 84 , 85 ]. Before sequencing analysis, DNA fragments were purified employing the BigDye XTerminator Purification Kit (Applied Biosystems, Waltham, MA, USA) according to the manufacturer’s manual. Bioinformatic analysis of the DNA sequences obtained was conducted by the Chromas Lite 2.01 and NCBI Nucleotide Blast software against the  COL1A1  ( NG_007400.1 ) and  COL1A2  ( NG_007405.1 ) genes as the reference sequences.\nPrimary skin fibroblasts and control cell line cultures were harvested with trypsin/EDTA (PAA Laboratories, Colbe, Germany) and washed twice with sterile phosphate buffered saline (PBS) (PAA Laboratories, Colbe, Germany). The cell amount was counted using Trypan Blue (BIO-RAD, Hercules, CA, USA) in the TC-20 Automated Cell Counter (BIO-RAD, Hercules, CA, USA). Cell lysis and purification of total RNA and miRNA were conducted using 2.5 × 106 cells in total. The isolation procedure was performed with the miRNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. The purity and concentration of the collected RNA samples were determined using a Nanodrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA).\nThe miScript II RT Kit (Qiagen, Hilden, Germany) and an RNA concentration of 250 ng were used to synthesize cDNA. The obtained cDNA was used to examine the defined miRNA expression ( Table 2 ). For this purpose, expression analysis of Custom miScript miRNA PCR Array, conf. no. CMIHS02722F (Qiagen, Hilden, Germany), was performed in a thermal cycler, the LightCycler 480 II (Roche, Basel, Switzerland). The quantitative polymerase chain reaction analysis was carried out in quadruplicate with miScript SYBR Green PCR Kit (Qiagen, Hilden, Germany) according to the manufacturer’s manual.\nThe expression analysis of miRNAs was performed by GenEx ver6 software (MultiD Analyses AB). Raw data were normalized to the reference genes ( SNORD61 ,  SNORD68 ,  SNORD72 ,  SNORD95 ,  SNORD96A , and  RNU6-2 ) according to the manufacturer’s protocol. The Kolmogorov–Smirnov test was employed to determine the distribution of the data ( Supplementary Table S1 ). For miRNA data that presented normal distribution, a parametric, unpaired, one-tailed t-test was conducted. In the case of miRNAs that showed non-normal distribution of data, a one-tailed Mann–Whitney test was used. The results were considered statistically significant at  p  < 0.05. The correlation analyses of miRNAs were performed using the Spearman method. Comparative expression of miRNAs in the study group and the control cell line was determined by scatter plot analysis.","source_license":"CC-BY-4.0","license_restricted":false}