Introduction
The exceptional capability of stem cells to self-renew and differentiate into multiple and distinct cell types has rendered them an invaluable resource for replacing damaged tissues, particularly in regenerative engineering research [1,2]. The discovery of human embryonic stem cells (hESCs), a type of pluripotent cell, by Thomson et al. [3] had a profound impact on stem cell research; however, it also sparked ethical concerns that prompted the hunt for new sources of pluripotent cells.
Induced pluripotent stem cells (iPSCs) are a compelling substitute for hESCs that is obtained by genetically reprogramming adult somatic cells via the introduction of exogenous pluripotent factors. iPSCs have pluripotent characteristics that resemble those of hESCs but are not ethically problematic [4]. Of note, since the first generation of iPSCs was achieved by the retroviral transduction of Yamanaka factors (Oct4, Sox2, Klf4, and c-Myc) into murine fibroblasts [5], the scientific community has witnessed remarkable advances in techniques for the isolation of stem cells from a variety of somatic cell sources, including bone marrow, blood, adipose tissues, and dermal fibroblasts [6,7]. However, the intrusiveness of methods that are used to extract these cells can make it challenging to obtain cells from these sources, because they frequently reduce the pool of possible donors. Conversely, urine cells (UCs) offer a potentially viable solution for this sample scarcity. UCs can be obtained from voided urine samples, thereby providing a virtually unlimited supply of cells that can be reprogrammed into iPSCs [8,9]. Compared with other adult somatic cells, UCs are more affordable and readily available, and their collection is painless, thus expanding the donor pool. This method is well-suited for infant and pediatric patients [10], especially those with neurogenetic disorders. UCs also exhibit crucial mesenchymal stem cell characteristics, such as the ability to differentiate into many lineages, and exhibit immunomodulatory properties [11,12]. These properties render UCs valuable for disease modeling, regenerative medicine, and stem cell research.
However, several obstacles prevent the use of urinederived iPSCs in therapeutic settings; these challenges must be addressed before their clinical application. First, there are no generally recognized and standardized protocols for creating iPSCs, which leads to variations in the dependability and quality of the cells [13]. Second, conventional techniques for producing stem cells often involve risk factors, such as animal feeder cells (e.g., mouse embryonic fibroblasts) [14], transduced viruses [15], or undefined serum supplements [5], which can introduce significant variability in the culture conditions and raise safety concerns regarding transgene insertion causing mutagenesis, leading to a potential of tumor development and genomic instability [16]. These challenges have triggered vigorous attempts to revise the existing protocols to remove xenobiotic components and viral factors from the reprogramming process. For instance, the generation of virus-free human iPSCs via nonintegrated strategies using Sendai virus delivery, transient episomal transfection, and mRNA or protein delivery has been widely reported [17-20]. However, the mRNA and protein transfection efficiencies are low and the process is laborious. In turn, Sendai virus transfection frequently triggers the host immune system and requires stringent virus-handling conditions, coupled with a high financial research burden and the risk of retention of any RNA virus trace after reprogramming. Therefore, episomal vectors have become the top choice for academic research or pilot experiments because of their suitability in terms of safety and simplicity. Episomal vectors have garnered interest regarding their affordability, ease of use, low hands-on time, and lower likelihood of inducing an immune response. To date, episomal vectors have been used in iPSC-based clinical cell-therapy trials [21].
In this study, we aimed to develop a protocol for the generation of iPSCs from UCs in a standardized manner and to assess the potential for the large-scale production of iPSCs across a diverse patient population, regardless of genetic background. We desire to generate a human stem cell bank that is devoid of serum, feeder cells, or viral vectors, which would increase the therapeutic utility of UCs.
Methods
All participants in this study provided written consent to donate urine samples for stem cell research. The experiments were carried out at the Seoul National University Biomedical Research Center and were reviewed and approved by its Institutional Review Board (IRB 1905-116-1035 and 2210-115-1372).
1. Isolation of UCs
Urine samples were collected from patients, stored on ice, and processed within 1 hour. UCs were isolated and cultured as described previously [22]. Briefly, the samples were centrifuged at 400×g for 10 minutes, and the cell pellet was washed with 10 mL of DPBS (Gibco, USA) containing 1% penicillin/streptomycin (P/S; Gibco). The pellets were resuspended in 0.5 mL of primary UC medium (DMEM/F12, Gibco) supplemented with 10% fetal bovine serum (FBS; Gibco), P/S, and SingleQuot Kit (Lonza, Switzerland), then plated onto a single well of a 12-well plate coated beforehand with 0.1% gelatin (Welgene, Korea). The cells were incubated under standard conditions (37°C, 5% CO2). For the following 3 days, 0.5 mL of primary UC medium was added daily to each well. From day 4, half of the medium was gradually replaced with RE/MC medium (proliferation UCs medium), which was prepared by mixing (at a 1:1 ratio) Renal Epithelial Basal Medium (Lonza; mixed with the SingleQuot Kit and P/S) and Mesenchymal Cell medium (which is composed of DMEM/high glucose, 10% FBS, GlutaMAX, NEAA [Gibco], and SingleQuot Kit). Small colonies appeared between days 5 and 10, followed by rapid proliferation. UCs were further passaged using TrypLE (Gibco) and maintained in RE/MC medium. Before freezing, healthy cells were tested for bacterial and fungal contamination (16S polymerase chain reaction [PCR]) and stored in a liquid nitrogen tank using a serum-free cryopreservation medium (Cell Banker 2, Zenogen Pharma, Japan).
2. Generation and culture of iPSCs from patient-derived UCs
UCs were reprogrammed into iPSCs using the Epi5 Episomal Reprogramming Kit (Invitrogen, USA) (Fig. 1A). On day 0, UCs at 70%–80% confluency were electroporated with plasmids containing the reprogramming factors OCT3/4, SOX2, KLF4, L-MYC, and LIN28. UCs were washed and detached using TrypLE and centrifuged at 1,200 rpm for 4 minutes, and the pellets were resuspended in Buffer R (Invitrogen) with the Epi5 vectors (1 μL each). Transfection was performed using a Neon system (1,200 V, 50 msec, 1 pulse). The transfected UCs were plated onto a Matrigel (Corning)-coated 6-well plate and incubated overnight in a 37°C/5% CO2 incubator in RE/MC medium supplemented with 10 μM ROCK inhibitor (StemCell Technologies, Canada).
On day 1, the medium was replaced with N2B27 medium, which consisted of DMEM/F12 with HEPES (Gibco), N2 (Gibco) (1×), B27 (Gibco) (1×), MEM-NEAA (1×), GlutaMAX (1×), β-mercaptoethanol (55 μM), and basic fibroblast growth factor (Gibco). The N2B27 medium was changed every 48 hours. On day 15, the medium was switched to mTeSR medium (StemCell Technologies), and iPSC-like colonies were picked and transferred to a new Matrigel-coated 6-well-plate. Expanded iPSCs were routinely propagated every 4 or 5 days using ReLeSR (StemCell Technologies).
3. iPSC validation
1) Sequencing analysis
DNA was extracted from iPSC lines by a QIAamp DNA Mini Kit (Qiagen, Germany) according to the manufacturer’s instructions. Mutation sites were confirmed through Sanger sequencing.
2) Gene expression analysis
The expression of pluripotent marker genes (OCT3/4, SOX2, LIN28, and NANOG) was assessed. Total RNA was extracted using the RNeasy Mini Kit (Qiagen) and transcribed to complementary DNA using a High-Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific, USA). PCR was performed on a PCR system (Applied Biosystems, USA) using the following primer sets: OCT3/4-F, 5´-CCCCAGGGCCCCATTTGGTACC and OCT3/4-R, 5´-AC CTCAGTTTGAATGCATGGGAGAGC; SOX2-F, 5´-TTCACAT GTCCCAGCACTACCAGA and SOX2-R, 5´-TCACATGTGTG AGAGGGGCAGTGTGC; LIN28-F, 5´-AGCCATATGGTAGCC TCATG TCCGC and LIN28-R, 5´-TCAATTCTGTGCCCCGGG AGCAGGGTAGG; and NANOG-F, 5´-TGAACCTCAGCTACA AACAG and NANOG-R, 5'-TGGTGGTAGGA AGAGTAAAG. The amplification program was as follows: initial denaturation at 95°C for 3 minutes; followed by 35 cycles of 95°C for 30 seconds, 60°C for 30 seconds, and 72°C for 1 minute; and a final extension step at 72°C for an additional 5 minutes. The samples were run in triplicate, and actin was used as the housekeeping gene. The PCR products were analyzed by 2% agarose gel electrophoresis and imaged with a ChemiDoc Imaging System (Bio-Rad, USA), followed by densitometric analysis, which was performed using ImageJ software (National Institute Health, USA).
3) Immunostaining
Attached cells were washed in PBS, fixed with 4% paraformaldehyde for 10 minutes, permeabilized with 1% Triton X-100 for 10 minutes, and blocked with 5% goat serum. The cells were stained overnight at 4°C with primary antibodies against OCT3/4 (dilution 1:200; Abcam, USA), NANOG (dilution 1:200; Thermo Fisher, USA), SSEA-4 (dilution 1:500; Thermo Fisher), and TRA-1-60 (dilution 1:200; Thermo Fisher), followed by 3 washes with PBS. The cells were then incubated with secondary antibodies (Alexa Fluor 488-conjugated species-specific antibodies; dilution 1:1000; Thermo Fisher) for 3 hours at room temperature. DAPI (4´,6-diamidino-2-phenylindole, 1:1000; Sigma-Aldrich, USA) was used to stain nuclei. Fluorescence images were acquired on a Leica Biosystem (Germany) microscope.
4) Embryoid body differentiation assay
iPSC colonies were initially plated in 6-well plates and treated on day 0 with 2-mg/mL collagenase IV in KnockOut DMEM (Gibco) for 30–45 minutes at 37°C, until a complete detachment of the colonies was observed. Subsequently, the cells were resuspended in embryoid body-KnockOut (EB-KO) medium, which consisted of 250 mL of KnockOut DMEM supplemented with 45 mL of KnockOut serum replacement (Gibco), MEM-NEAA (1×), GlutaMAX (1×), P/S (1×), and 540 μL of β-mercaptoethanol (55-mM stock). The cell suspension was then transferred to a 15-mL conical tube and centrifuged for 3 minutes at 1,200 rpm. The pellets were washed again with EB-KO and subsequently plated in low-adherence plates (Corning, USA) to facilitate EB formation. The EB-KO medium was routinely replaced every 48 hours. Starting from day 8, the EBs were carefully transferred to a 0.1% gelatin-coated 24-well plate and cultured in EB-fetal calf serum (FCS) medium containing components similar to EB-KO, albeit with the replacement of KnockOut serum with FCS. On day 16, the differentiated EBs were fixed and stained for smooth muscle actin (SMA; for the mesoderm), alpha-fetoprotein (AFP, dilution 1:200; for the endoderm; Thermo Fisher), and β3-tubulin III (dilution 1:200; for the ectoderm; BioLegend, USA) to assess the expression of specific markers corresponding to the 3 germ layers.
5) Chromosomal microarray
DNA was extracted using the QIAamp DNA Mini Kit (Qiagen) according to the manufacturer’s instructions. Chromosomal microarray (CMA) analysis was carried out using the SurePrint G3 Human CGH Microarray Kit (Agilent Technologies, USA). DNA was labeled with Cy3-dUTP (sample) and Cy5-dUTP (sex-matched reference), the slides were scanned, and the raw microarray data were analyzed using Feature Extraction Software (Agilent, USA). Data visualization and analysis were performed using GeneSpring GX software.
Results
1. Urine collection and UC isolation
Twelve urine specimens were collected from patients (7 males, 5 females) aged 1 month to 41 years and UCs were successfully isolated, with informed consent. Five specimens obtained from adult patients were included in the analysis, because all the diseases were known to have pediatric-age onset. For younger children, urine bags were used to assist the collection. The volumes of urine collected ranged from 8 to 97 mL. We observed that this pain-free and simple collection method afforded a more diverse and scalable sample-collection process.
The samples were stored at 4°C until UC isolation was performed. We found that expeditious sample processing greatly increased the success rate of UC isolation. This result is consistent with those of other studies indicating that UCs become less viable after exposure to urine for longer than 5 hours, because of a considerable decrease in available nutrients within the urine and alterations in pH [23]. According to previous reports, UCs can successfully be prepared from voided urine and can afford a sizable number of cells from a single clone [8]. Similarly, in our cases, adherent UCs were visible as early as days 5–7, and multiple UC colonies quickly expanded throughout the culture plate, displaying tight clusters of cells toward the center of the plate (Fig. 1B). A 16S PCR microbial and fungal contamination test was performed before the preparation of cryopreservation stocks. The frozen UC stocks were stored and could be thawed for unlimited proliferation and reprogramming.
These preliminary results revealed the successful and reproducible isolation of UCs from a diverse population of donors, regardless of sex and age. Moreover, they demonstrate that our UC isolation protocol can be universally applied to most individuals, ranging from healthy individuals to those with various diagnosed conditions.
2. iPSC characterization
To produce iPSCs using a non-integrating technique, the UCs were treated with the Epi5 Episomal iPSC Reprogramming Kit. The episomal plasmids included in this kit encode essential transcription factors, such as Oct4, Sox2, Klf4, L-Myc, and Lin28, which are necessary for converting somatic cells into iPSCs. Because in this approach episomal plasmids do not integrate into the host genome, insertional mutagenesis is reduced, and genomic stability is maintained, it is a safer method compared with conventional viral transfection. Once inside cells, the plasmids transiently express the necessary reprogramming factors, which convert the UCs into iPSCs over time, typically around 15–21 days [24]. To promote their proliferation and self-renewal, these iPSCs are cultivated under carefully controlled mTeSR and feeder-free environments. In fact, we observed the first iPSC colony at 25 days post-reprogramming, which exhibited a rounded shape, smooth borders, and a flatter appearance (Fig. 1C). This morphology indicates that the UCs were successfully reprogrammed into iPSCs. This distinct colony was selected to establish feeder-free and serum-free iPSCs through long-term culture on Matrigel and mTeSR.
Furthermore, pluripotency validation was carried out to investigate the PSC-like traits of the newly generated stem-cell-like colony. In the comparison of iPSCs with unprogrammed UCs, pluripotency was confirmed by the upregulated mRNA expression of important markers, such as NANOG, OCT4, SOX2, and LIN28 (Fig. 2A). Moreover, all colonies were positive for the OCT4, NANOG, and SSEA-4 pluripotency protein markers, as assessed via immunostaining (Fig. 2B). Using immunocytochemistry during the formation of EBs, we observed that iPSCs were capable of differentiating into all 3 embryonic germ layers: mesoderm (identified by SMA), endoderm (identified by AFP), and ectoderm (identified by β3-tubulin III) (Fig. 2C). This comprehensive analysis of pluripotency indicates that the iPSCs produced here exhibit all the potent traits needed for subsequent differentiation and scientific applications.
When encountering prolonged culture and a complicated reprogramming process, iPSCs tend to become more vulnerable and prone to genomic instability [25]. Therefore, their genomic integrity should be tracked. Of note, previous research frequently employed traditional karyotype analysis, which has several drawbacks, including limited resolution for identifying significant large genomic aberrations. To address this issue and enhance the quality of the iPSCs, we opted for CMA analysis, which offers a higher resolution, thus allowing the detection of chromosomal abnormalities with greater sensitivity [25].
The genomic profiles from CMA showed that the majority of the iPSCs created using the proposed approach did not display any irregularities (Table 1). However, 4 out of 12 lines exhibited notable chromosomal deletions or duplications. The size of the chromosomal aberration was around 5 Mb in 2 out of 4 iPSC lines, which could possibly have been missed by the conventional karyotyping method. A follow-up CMA analysis that was performed using lines from an earlier passage confirmed that 3 of the 4 lines retained the abnormalities, thus revealing their instability and inadequacy for further application. These findings highlight the importance of genetic screening throughout the entire culturing process, to guarantee the safety and dependability of iPSCs for future applications.
Overall, in the current study, we offered a workable procedure for generating virus-free iPSCs under harsh conditions, such as serum- and feeder-free settings, which is ideal for clinical applications. Furthermore, as urine collection is noninvasive and does not inconvenience patients, this procedure can be universally used by all contributors, thus facilitating the establishment of an iPSC bank.
Discussion
Regardless of an individual’s age, sex, or genetic origin, our goal is to create a large-scale, standardized procedure for producing iPSCs from UCs. UCs are the best adult somatic cells for this purpose, because they possess several benefits over other types of somatic cells. First, the noninvasive method that is used for their collection significantly boosts donor engagement. Because they do not experience the physical discomfort and pain associated with the collection of intrusive specimens, this practice greatly relieves all patients, especially pediatric patients. Compared with tissue biopsies, which are labor-intensive and require specialized equipment, the isolation of UCs from urine specimens is simple and can be carried out in a typical laboratory setting [26]. Second, UCs exhibit a high proliferative capacity compared with other adult somatic cells. Previous research has demonstrated that a single cell may create 5–7 clones from 100 mL of urine in 2–3 days; in addition, because of their long telomeres and high telomerase activity, these can grow into a huge number of UCs [12,27]. By the early-passage stage (passage <p8), UCs can reach millions of cells in a matter of weeks, which renders them a prolific and sustainable source of cells for therapeutic applications [28]. Because of their rapid expansivity and highly proliferative capacity, UCs are a desirable and scalable cell type for developing an effective and clinically relevant iPSC platform, which will accelerate applications in regenerative therapies, personalized medicine, and disease modeling [29]. Reprogramming UCs into iPSCs has been the subject of numerous investigations. Interestingly, UCs reprogram into iPSCs faster and more effectively than other somatic cells. This accelerated pace of reprogramming probably occurs because UCs are epithelial, which means that a mesenchymal-to-epithelial transition is not necessary [9,30]. Takahashi and colleagues first prepared reprogrammed iPSCs by introducing 4 key transcription factors—OCT4, SOX2, KLF4, and c-MYC—into UCs using retroviral transduction. Despite its success, this technique sparked worries regarding safety, especially in light of the carcinogenic potential of c-MYC [18]. Since then, efforts have been directed toward the creation of nonviral reprogramming methods that minimize tumorigenicity by lowering the likelihood of transgene insertion. Techniques such as purified proteins, synthetic mRNA, Sendai virus, and episomal vectors have emerged as safer substitutes; however, to date, only episomal vectors have been employed in clinical settings. Here, we employed the Epi5 episomal vector system, which introduces 5 episomal vectors carrying human genes—OCT4, SOX2, LIN28, KLF4, and L-MYC—into UCs via electroporation. This system offers a significant advantage over viral-based methods because it substitutes the tumorigenic c-MYC with L-MYC, which is a less oncogenic variant, thus rendering this approach more suitable for clinical applications [18]. Moreover, our protocol moves toward a defined, serum-free, and feeder-free culture system, using Matrigel as the matrix and a defined mTeSR medium, which promotes genetic stability and minimizes the risks associated with serum and feeder environments.
Furthermore, the preservation of genomic integrity during the ongoing processes of self-renewal and proliferation is one of the fundamental challenges in the creation of a workable and trustworthy methodology for the production of a consistent and high-quality iPSC bank. iPSCs are exceptionally prone to genetic changes, particularly when they are cultivated for long periods or come into contact with exogenous substances that trigger the production of variants during reprogramming. Therefore, the genomic validation of iPSCs is very important, because genomic instability raises the likelihood of cancer in stem-cell-derived therapies. Cytogenetic G-banding karyotyping is widely accepted as a “gold standard” for assessing genomic stability; however, the preparation and analysis of each sample require substantial time, financial resources, and effort, and this technique is not sufficiently sensitive to discern subtle genetic alterations. Higher resolution has been attained through the use of CMA; using this technology, submicroscopic chromosomal aberrations can be identified [31]. In turn, whole-genome sequencing (WGS) and RNA sequencing (RNA-seq) provide superior resolution for identifying a broad range of genomic abnormalities, including single-nucleotide variants, small insertions and deletions, and transcriptomic changes. For in-depth mechanistic investigations, these techniques provide complete genomic and transcriptomic landscapes at single-base resolution, which is highly advantageous. However, both WGS and RNA-seq require significant computational resources, bioinformatics expertise, and higher costs, which can constrain their routine application for quality control in iPSC cultures [32]. Conversely, CMA is an affordable, well-established method that accurately identifies copy number variations and large-scale chromosomal abnormalities, which are essential for preserving the genomic integrity of iPSC lines over extended culture. For these practical reasons—cost-effectiveness, ease of implementation, and our laboratory’s current expertise— we opted to use CMA for our routine screening. In our case, among the 12 iPSC lines generated, 8 passed a rigorous genomic CMA screening, whereas the remaining lines exhibited signs of chromosomal aberrations. Large deletions (>20 Mb) of chromosomes 20 and 8 were observed in iPSC_SNU23_05 and iPSC_SNU23_01, respectively. Moreover, microdeletions, which are frequently overlooked when using conventional karyotyping, were detected on chromosomes 9 (deletion size, ~11 Mb), 12 (deletion size, ~4.8–6.5 Mb), and 19 (deletion size, ~1.58 Mb). Reportedly, chromosomes 12 and 20 are frequently aberrant in iPSCs. Chromosomal 20 instabilities, including additions and losses in the 20q region, have been noted in numerous investigations. The long-arm 20q11.21 region has been found to be the most-often amplified chromosomal locus in both ESCs and iPSCs [33]. This particular area is likely necessary for the selection advantage during cell culture because it contains genes that enhance survival and proliferation (e.g., BCL-xL overexpression). However, although 12p short-arm amplification is linked to the increased expression of genes related to pluripotency and cell survival (e.g., NANOG and GDF3), 12q long-arm aberrations are infrequently observed in iPSCs [21]. Of note, the deletions detected on chromosomes 8, 9, and 19, which are just partially explored, are concerning because they impact the stability and differentiation capacity of these cells. In addition, early-passage iPSCs displayed normal genomic profiles in later trials (iPSC_SNU23_03), suggesting that earlier-stage cells may be more genetically stable. In fact, a recent extensive analysis of more than 100 hPSC lines from numerous laboratories revealed that late-passage iPSCs are twice as likely to contain genomic alterations as early-passage cells [34]. Therefore, CMA analysis should be performed on a regular basis to guarantee the consistency of iPSCs.
In conclusion, we successfully developed a viral-free, serum-free, and feeder-free pediatric neurogenetic disease iPSC bank derived from UCs. Compared with other protocols, the collection of urine samples is easy, and our protocol can be reliably expanded to the large number of patients necessary to build the iPSC bank program. High-resolution molecular genetic screening procedures, such as CMA, should be employed at every stage of the experiment to detect small chromosomal imbalances and ensure the production and maintenance of high-quality iPSCs. This approach will help maintain profound genomic stability, particularly regarding the cultivation of stem cells, which is crucial for enhancing their safety profile in the context of stem cell therapies for clinical purposes.