The USCs were isolated from 250 to 300 mL urine samples collected from three healthy male donors aged 32–42 years by centrifugation at 500 g for 10 min. The cell pellets were washed with phosphate-buffered saline (PBS) supplemented with 100 μg/mL primocin (InvivoGen, San Diego, CA, United States) and then resuspended in primary medium comprising DMEM/F-12 (Thermo Fisher Scientific, Waltham, MA, United States), 10% fetal bovine serum (FBS, Welgene, Daejeon, Korea), renal epithelial growth medium (REGM) SingleQuots Kit (Lonza, Basel, Switzerland), and 100 μg/mL primocin. The cells were plated on 0.2% gelatin-coated six-well culture plates and incubated for 3 days. The cells were then cultured in growth medium, which comprised 1:1 REGM (basal media supplemented with 10% FBS, REGM SingleQuots Kit, and 100 μg/mL primocin) and mesenchymal cell proliferation medium (high glucose DMEM supplemented with 10% FBS, 1× GlutaMax, 1× non-essential amino acids [Thermo Fisher Scientific], 5 ng/mL basic fibroblast growth factor [bFGF; PeproTech, Seoul, Korea], 5 ng/mL platelet-derived growth factor-AB [PDGF-AB, PeproTech], and 5 ng/mL epithelial growth factor [EGF; R&D Systems, Minneapolis, MN, United States]). Upon reaching 80% confluence, the USCs were detached using 0.25% trypsin-EDTA (Thermo Fisher Scientific) and reseeded on gelatin-coated plates. Human osteoblasts (OBs) (#CC-2538; Lonza) were cultured in osteoblast growth medium (PromoCell, Heidelberg, Germany). Human dermal fibroblasts (HDFs) and MCF-7 cells, a human breast cancer cell line, were cultured in DMEM supplemented with 10% FBS and 1% penicillin–streptomycin (Thermo Fisher Scientific).

The cDNA sequences of Lin28A obtained from the pCXLE-hUL plasmid (Addgene plasmid #27080) and the 30Kc19α used in the previous study (Ryu et al., 2016) were cloned into the pET-23a vector to bind Lin28A to the C-terminus of 30Kc19α. Exponentially growing cultures of Escherichia coli transformed with the recombinant vector were treated with 0.5 mM isopropyl-β-D-thiogalactopyranoside at 25°C for 16 h. After cell lysis by sonication, the soluble fraction of the recombinant protein was purified using Ni-NTA chromatography (HisTrap HP, Cytiva, Uppsala, Sweden). After dialysis against DMEM, the purified protein samples were mixed with a reducing sample buffer containing sodium dodecyl sulfate (SDS) and β-mercaptoethanol and loaded on a 10% polyacrylamide gel for separation. Protein bands were visualized through Coomassie Brilliant Blue R-250 (Sigma-Aldrich, St. Louis, MI, United States) staining.

For immunostaining, the cells were fixed with 4% paraformaldehyde (PFA) and permeabilized with 0.25% Triton X-100 in PBS. After blocking, cells were sequentially incubated with anti-T7 tag polyclonal primary antibody (Abcam, Cambridge, United Kingdom) at 4°C overnight and Alexa Fluor 488-conjugated secondary antibody (Thermo Fisher Scientific) at room temperature for 1 h. After treatment with 4′,6-diamidino-2-phenylindole (DAPI, Sigma-Aldrich) to stain the cell nuclei, fluorescence images were obtained by confocal laser scanning microscopy (CarlZeiss, Oberkochen, Germany). To investigate the cytotoxicity of 30Kc19α–Lin28A protein treatment, USCs (2×10⁴ cells/cm²) were seeded on 0.2% gelatin-coated 96-well plates and incubated for 24 h in growth medium. After treatment with 30Kc19α–Lin28A in growth medium for 24 h, a water-soluble tetrazolium (WST)-8 solution (Quanti-MAX WST-8 Cell Viability Assay Kit reagent; Biomax, Seoul, Korea) was added to each well and incubated for another 2 h. Finally, absorbance was measured at 460 nm using a microplate reader.

To induce osteoblast differentiation, human USCs (2×10⁴ cells/cm²) were seeded in a 0.2% gelatin-coated 24-well plate and incubated for 24 h in growth medium. The growth medium was then replaced with DMEM supplemented with 10% FBS, 10 mM-glycerol phosphate, 50 μg/mL ascorbic acid, and 100 nM dexamethasone (osteogenic medium), and the cells were further cultured with medium changes every 2 days. During the first week of the osteoblast differentiation, the cells were treated two or three times with 30Kc19α–Lin28A, with a two-day interval between each treatment. Subsequently, the culture was continued in osteogenic medium.

For Alizarin Red S (ARS) staining, the differentiated cells were fixed with 4% PFA and washed thrice with PBS. Then, they were stained with 2% ARS solution (Sigma-Aldrich) as previously described (Lee J et al., 2020) and observed under a microscope. Briefly, each stained culture was incubated with 10% acetic acid and gently shaken for 10 min at room temperature. After heating at 85°C for 10 min and neutralizing with 10% ammonium hydroxide, the absorbance at 405 nm was measured using a microplate reader. The calcified matrices were visualized using the OsteoImage mineralization assay according to the manufacturer’s instructions (Lonza). After fixation with 70% ethanol for 20 min, a fluorescent staining reagent was added to each well and incubated for 30 min in dark. The bone-like calcified matrices were observed using a fluorescence microscope.

Total RNA was isolated using the Ribospin Total RNA Purification Kit (GeneAll Biotechnology Co., Ltd., Seoul, Korea) according to the manufacturer’s instructions. The isolated RNA was then reverse transcribed into cDNA using TOPScript RT DryMIX (Enzynomics Co., Ltd.) and the dT 18 plus primer. For reverse transcription polymerase chain reaction (RT-PCR) analysis, each cDNA was amplified using 2× TOPsimple DyeMIX-HOT (Enzynomics Co. Ltd.) and relevant primers in a T100 Thermal Cycler (Bio-Rad, Hercules, CA, United States) for 25 cycles. Quantitative real-time PCR (qPCR) was performed using TOPreal qPCR 2× PreMIX (SYBR Green with low ROX; Enzynomics) with glyceraldehyde 3-phosphate dehydrogenase serving as an internal control. All experiments were conducted in quadruplicate, and the relative comparison between cell groups was determined using the 2^−ΔΔCT method. The primer sequences used for both RT-PCR and qPCR analyses are summarized in Supplementary Table S1.

For RNA-sequencing (RNA-seq) analysis, total RNA was isolated using the TRI reagent (Thermo Fisher Scientific), and its quality was evaluated using the Agilent TapeStation 4000 system (Agilent Technologies, Amstelveen, Netherlands). The RNA library of the control and experimental groups were constructed by employing the QuantSeq 3′mRNA-Seq Library Prep Kit (Lexogen Inc., Vienna, Austria) according to the manufacturer’s instructions. After amplifying the library amplification to include the complete adapter sequences required for cluster generation, the finished library was purified from the PCR components. High-throughput single-end sequencing was performed using NextSeq 550 (Illumina, Inc., United States) as single-end 75 sequencing. Gene classification was based on searches conducted using DAVID (http://david.abcc.ncifcrf.gov/) and Medline (http://www.ncbi.nlm.nih.gov/). Raw data and graphic visualization were processed using ExDEGA graphic software (Ebiogen, Seoul, Korea). Genes with ≥1.5 fold change (upregulated/downregulated) were considered statistically significant. The STRING software (https://string-db.org/) was used to evaluate possible interactions, co-expressed genes, and protein networks.

All results are expressed as mean ± standard deviation. Statistical significance was confirmed using Student’s t-test and one-way analysis of variance (ANOVA). The results were considered significant at p < 0.05. All quantitative analyses presented in this study were obtained from replicate samples in a representative experiment conducted several times.

USCs were isolated from urine samples obtained from three donors. Initially, each isolated USC was cultured in the primary medium and then switched to the growth medium. Microscopic observation revealed that the cultured cells exhibited a fibroblast-like morphology, which is consistent with previously reported characteristics (Figure 1A). As previous studies have reported that human USCs share similar characteristics with MSCs (Bharadwaj et al., 2011; Bharadwaj et al., 2013; He et al., 2016), we investigated the expression of the typical MSC surface markers CD73, CD90, and CD105 in isolated human USCs (Supplementary Figure S1). Despite some variations in the expression levels, RT-PCR analysis demonstrated that all three USC lines expressed these positive MSC marker mRNAs. These markers were detected in human BM-MSCs, but not in MCF-7 cells, indicating successful USC isolation from the three different donors.

The 30Kc19α–Lin28A fusion protein was designed to link Lin28A to the C-terminus of 30Kc19α. All recombinant proteins used in this study had a T7 tag at the N-terminus for immunostaining, and a 6× histidine tag at the C-terminus for Ni-NTA affinity chromatography (Figure 1B). SDS–PAGE indicated that the molecular weights of purified proteins corresponded to their theoretical sizes (13, 26, and 39 kDa for 30Kc19α, Lin28A, and 30Kc19α–Lin28A, respectively; Supplementary Figure S2). We investigated whether 30Kc19α fusion could enhance the stability of Lin28A by comparing the degradation rate at high temperatures. After incubation at 37°C in serum-free and 10% FBS-containing DMEM for different time intervals, the remaining Lin28A and 30Kc19α–Lin28A were analyzed by SDS–PAGE (Figure 1C). The amount of non-degraded Lin28A significantly reduced after 48 h of incubation, whereas 30Kc19α–Lin28A remained more stable than intact Lin28A. These results indicate that 30Kc19α fusion can improve Lin28A stability in the culture environment.

To examine the effect of 30Kc19α fusion on intracellular delivery, we randomly selected a particular cell line, designated USC #1, from a group of three independent cell lines. Subsequently, the cells were treated with equimolar amounts of 30Kc19α, Lin29A, and 30Kc19α–Lin28A for 2 h. Confocal microscopy revealed that 30Kc19α and 30Kc19α–Lin28A were delivered intracellularly, whereas Lin28A did not penetrate the cell significantly (Figure 1D). Subsequently, the cytotoxicity of the 30Kc19α–Lin28A protein was analyzed by WST-8 assay. The USCs were treated with various 30Kc19α–Lin28A concentrations of for 24 h in proliferative medium. Even at the highest concentration (7 μM), the cell viability did not decrease significantly (Figure 1E). These results clearly indicate that 30Kc19α fusion confers a cell-penetrating property to the non-permeable Lin28A protein without any significant cytotoxicity.

A decline in the self-renewal and differentiation potential of SSCs is a well-known hallmark of aging (Roobrouck et al., 2008). Previous studies have demonstrated that forced Lin28A expression in various SSCs via lentiviruses can increase cell expansion and proliferative marker expression. Additionally, Lin28A overexpression improves bone and cartilage differentiation of aged MSCs, thereby enhancing the tissue repair capacity of SSCs after transplantation (Pieknell et al., 2022). Despite its benefits, the lentiviral delivery of Lin28A raises safety concerns for clinical applications, including tumorigenesis and alterations in cell fate due to non-specific chromosomal modifications in recipient SSCs. To overcome these limitations of lentivirus-mediated DNA delivery, various integration-free strategies, such as RNA and protein delivery, have been developed. However, direct protein delivery requires repeated protein transduction owing to low stability and limited intracellular protein delivery, which can lead to severe cytotoxicity. Based on the results presented in Figure 1, which confirmed the high stability, intracellular delivery, and low cytotoxicity of the 30Kc19α–Lin28A protein, we investigated its potential on the osteoblast differentiation of USCs.

Since Lin28A is associated with inhibiting the terminal differentiation of SSCs (Molenaar et al., 2012; Urbach et al., 2014), excessive exposure to Lin28A might interfere with USC differentiation. Therefore, we aimed to determine the optimal condition for 30Kc19α–Lin28A treatment. Human USCs (USC #1) were treated with 0–7 μM 30Kc19α–Lin28A protein two or three times during the early stage of osteogenesis (Supplementary Figure S3). ARS staining revealed that 30Kc19α–Lin28A dose-dependently enhanced USC osteogenesis. At the highest dose (7 μM, three times), ARS-stained calcium deposits covered most of the culture area, whereas no significant staining was observed in the non-treated control cells. In contrast to that with two treatments with 7 μM 30Kc19α–Lin28A, osteogenesis was significantly enhanced with the third treatment (Figure 2A). The OsteoImage mineralization assay also demonstrated the enhanced osteogenesis of USCs by 30Kc19α–Lin28A, showing that the calcium deposits increased than that in the non-treated control (Supplementary Figure S4). Accordingly, we subsequently treated USCs with 7 μM 30Kc19α–Lin28A three times during the first week of osteogenesis. Following three treatments with 30Kc19α-Lin28A, there were no notable differences in cell confluence during the continuous osteogenesis process, indicating that the proliferation of USCs was not significantly affected by the treatment with 30Kc19α-Lin28A. We also assessed the effects of 30Kc19α, Lin28A, and 30Kc19α–Lin28A proteins on USC osteogenesis. The cells were treated with 7 μM each protein three times, and calcified matrix formation was evaluated after osteogenic induction for 21 days. ARS staining revealed no significant effect of 30Kc19α and Lin28A on USC osteogenesis, whereas calcified matrices substantially covering the entire culture area were formed in only 30Kc19α–Lin28A-treated cells (Figure 2B). Quantitative analysis for the ARS staining also demonstrated that 30Kc19α–Lin28A treatments enhanced the osteoblastic differentiation of USCs, in contrast to 30Kc19α or Lin28A alone (Figure 2C). Considering the results of the cell penetration test, we postulated that this result could be attributed to the inability of Lin28A to penetrate the cells, whereas 30Kc19α penetrated the cells but did not enhance osteogenesis. These findings support our conclusion that Lin28A delivered intracellularly by fusion with the cell-penetrating 30Kc19α, enhances the differentiation potential of USCs into osteoblasts. While there is a possibility that the conjugated 30Kc19α could have a minor influence on the osteogenesis-promoting effect of 30Kc19α-Lin28A, it is more reasonable to attribute the primary functionality to the Lin28A itself.

To further investigate the impact of 30Kc19α–Lin28A on the osteogenesis of USCs, we examined the expression levels of osteoblast marker genes through qRT-PCR analysis. Specifically, the mRNA expression levels of the osteoblast-specific markers runt-related transcription factor 2 (Runx2), alkaline phosphatase (ALP), osteopontin (OPN), and osteocalcin (OCN) were compared on days 7 and 14 of osteogenesis. Our results indicated that 30Kc19α–Lin28A treatment significantly increased the mRNA expressions of both early (Runx2 and ALP) and late (OPN and OCN) osteogenic markers (Figure 2D). Although this increase in expression varied slightly for each marker with differentiation duration, 30Kc19α–Lin28A treatment clearly upregulated the expression of osteoblast-related genes, thereby enhancing the osteogenesis of human USCs.

To evaluate the changes in transcriptional profiles, the RNA of 30Kc19α–Lin28A-treated human USCs, non-treated control cells, and OBs were extracted after 14 days of osteogenesis and compared. The scatter plot analysis of the identified transcripts revealed 648 DEGs with 1.5-fold change in 30Kc19α–Lin28A-treated USCs compared to that in non-treated control cells (Figure 3A). A heat map obtained by hierarchical clustering of DEGs showed that the 30Kc19α–Lin28A-treated group and OBs clustered together and were separated from non-treated control cells, indicating that 30Kc19α–Lin28A treatment enhanced osteogenesis (Figure 3B).

Several genes, including Raf1 and Wnt5A involved in cell proliferation, differentiation, survival, and osteogenic differentiation, were upregulated. Previous studies have reported that Raf1 plays a pivotal role in preventing the apoptosis of osteocytes induced by fatigue loading (Li et al., 2011), and Wnt5A-induced noncanonical Wnt signaling promotes MSC osteogenesis and suppresses adipogenesis (Takada et al., 2007; Takada et al., 2009). Conversely, among the downregulated genes, PPP2R5B is a subunit of PP2A that regulates the cell cycle and growth factor signaling. The reduction of PP2A expression promotes osteogenic differentiation by regulating bone-forming transcription factors, such as osterix (Okamura et al., 2017).

Gene ontology analysis revealed a significantly altered expression of genes involved in transcriptional regulation by the RNA polymerase II promoter (Figure 3C). Previous studies have shown that phosphorylated extracellular signal-related kinase (p-ERK) phosphorylates Runx2, which is necessary for inducing osteoblast-specific transcription (Ge et al., 2009). This p-ERK-mediated Runx2 phosphorylation initiates a series of chromatin changes associated with RNA polymerase II recruitment and transcription. RNA polymerase II recruitment is also related to the functions of several homeodomain proteins, such as Dlx3 and Dlx5, which play crucial roles in the transcriptional regulation of osteoblast differentiation. These homeodomain proteins, along with Runx2 and OCN, are coordinately recruited with increased occupancy of RNA polymerase II (Hassan et al., 2004; Hassan et al., 2006). These findings suggest that 30Kc19α–Lin28A treatment is likely to promote osteogenesis in USCs by regulating transcriptional processes through RNA polymerase II. Furthermore, a STRING analysis of all DEGs revealed gene clusters involved in osteogenesis, indicating the transcriptomic responses of 30Kc19α–Lin28A-treated USCs toward osteogenesis (Figure 3D).

Despite the transcriptomic changes after 30Kc19α–Lin28A treatment, RNA-seq analysis has some limitations. The analysis of a single sample instead of multiple replicates for each experimental group made it challenging to minimize the variation in the measurement of the expression level of each transcript, even though the analysis of clusters with multiple genes was available. Moreover, there was a lack of detailed analysis regarding the impact of 30Kc19α–Lin28A on the transcriptional regulatory network and cellular signaling that enhanced the osteogenesis of USCs. Conducting large-scale analysis with an increased number of repeated samples in future studies would provide detailed insights into the molecular mechanism through which intracellularly delivered 30Kc19α–Lin28A proteins affect the transcriptional changes in USCs.

SSCs display notable variations in their self-renewal and differentiation potentials depending on the donor (Heathman et al., 2016; Zhang et al., 2021). To investigate such differences, we treated three independent USC lines derived from each donor with 30Kc19α–Lin28A protein. After osteoblast differentiation for 21 days, ARS staining revealed a significant variation in ossification levels between the cell lines, indicating donor-specific variation in osteogenic potential. Nevertheless, all 30Kc19α–Lin28A-treated USC lines exhibited excellent calcium deposition (Figure 4A). Interestingly, the osteogenesis of USC #3, which was at least partially differentiated into osteoblasts, in contrast to USC #1 used in previous experiments, was also significantly enhanced by 30Kc19α–Lin28A treatment (Figure 4B). These findings indicate the potential utility of 30Kc19α–Lin28A protein for the clinical applications of USCs obtained from various donors. However, it is widely acknowledged that a notable reduction in the expression of the tumor suppressor let-7 miRNA is a common occurrence in various types of human cancers (Nair et al., 2012). Since the maturation process of the primary let-7 transcript is impeded by the highly conserved RNA-binding protein, Lin28, the oncogenic potential of Lin28A is a major obstacle to clinical application. Nevertheless, in the present study, we administered the 30Kc19α-Lin28A protein to human USCs in three intermittent cycles, followed by osteogenic induction without any further treatment. Compared to previous studies that utilized viral gene delivery for continuous Lin28A expression (Shyh-Chang et al., 2013; Pieknell et al., 2022), the intermittent exposure to Lin28A through protein delivery may substantially reduce the risk of tumorigenicity by minimizing alterations in the transcriptional regulatory network. In addition, based on the results of RNA-seq analysis, when investigating the genes related to tumor cell response, there were no significant differences in expression levels between non-treated and 30Kc19α-Lin28A-treated USCs (Supplementary Figure S5). These results suggest that the administration of 30Kc19α-Lin28a protein had minimal impact on the transcriptional network involved in tumor cell response and tumorigenicity in human USCs.