Direct reprogramming of urine-derived cells with inducible MyoD for modeling human muscle disease
© The Author(s). 2016
Received: 18 May 2016
Accepted: 23 August 2016
Published: 15 September 2016
Cellular models of muscle disease are taking on increasing importance with the large number of genes and mutations implicated in causing myopathies and the concomitant need to test personalized therapies. Developing cell models relies on having an easily obtained source of cells, and if the cells are not derived from muscle itself, a robust reprogramming process is needed. Fibroblasts are a human cell source that works well for the generation of induced pluripotent stem cells, which can then be differentiated into cardiomyocyte lineages, and with less efficiency, skeletal muscle-like lineages. Alternatively, direct reprogramming with the transcription factor MyoD has been used to generate myotubes from cultured human fibroblasts. Although useful, fibroblasts require a skin biopsy to obtain and this can limit their access, especially from pediatric populations.
We now demonstrate that direct reprogramming of urine-derived cells is a highly efficient and reproducible process that can be used to establish human myogenic cells. We show that this method can be applied to urine cells derived from normal individuals as well as those with muscle diseases. Furthermore, we show that urine-derived cells can be edited using CRISPR/Cas9 technology.
With progress in understanding the molecular etiology of human muscle diseases, having a readily available, noninvasive source of cells from which to generate muscle-like cells is highly useful.
KeywordsUrine cells MyoD Reprogramming Myotubes Muscular dystrophy CRISPR
The genetic diversity of muscle diseases, including the muscular dystrophies, is extensive [1, 2]. Defining underlying mechanisms and developing therapies to correct these defects require appropriate models of disease. Animal models offer in vivo insight and have proved highly useful, but animal models are time consuming and costly to develop and cannot represent the large number of alleles responsible for muscle diseases. Cellular models of disease have expanded in recent years with significant advances in reprogramming and the creation of induced pluripotent stem cells (iPSCs) for many disorders. For muscle diseases, the comparative ease of generating cardiomyocytes, rather than skeletal myocytes, from iPSCs has led to this cell type being used as a surrogate for skeletal muscle [3–5]. Although progress has been made in generating muscle-like lineages from iPSCs, this process is still time-consuming and has variable outcomes within and across culture systems [6, 7]. Other commonly used cell models of skeletal muscles include the mouse C2C12, rat L6, and human myoblast cell lines, which undergo differentiation into multinucleate myotubes under appropriate conditions [8, 9]. These myoblast models have been highly useful for the field but represent normal skeletal muscle and not disease-specific conditions. While myoblasts can be cultured from human muscle, biopsies from patients are invasive and at times problematic for those with muscle mass-limiting conditions. A noninvasive patient-specific in vitro skeletal muscle model is desirable for studying pathogenesis and testing potential drugs.
Direct reprogramming of fibroblasts with MyoD into myotubes was a central experiment used to demonstrate the role of transcription factors [10, 11]. Since that original observation, MyoD delivery into fibroblasts has been optimized using viral vectors and this method can be applied to make directly reprogrammed myogenic cells from humans and mice [12, 13]. A fusion between MyoD and the estrogen receptor was generated to create a tamoxifen-inducible MyoD, and this construct was used in mouse and human fibroblasts to induce myogenesis [14–16]. Thus, direct reprogramming of fibroblasts remains a highly useful method for modeling human muscle diseases. However, dermal fibroblasts are obtained through skin biopsy, and the invasive nature of skin biopsy, while considerably less than muscle biopsy, can be particularly problematic in children.
One potential noninvasive source of patient-specific cells is cells isolated from urine. It has been estimated that about 2000 to 7000 renal tubular epithelial cells are shed in urine everyday [17, 18]. Obtaining a urine sample is noninvasive, making urine a good source of cells, especially from children. Urine cells have been isolated from both healthy volunteers and patients of a wide age range [18–21]. Urine-derived cells (UDCs) are thought to be of a mixed population that originates from either the renal epithelium or the uroepithelium [18, 21]. Interestingly, a subset of UDCs has been observed to carry mesenchymal stem cell markers as well as low endogenous expression of Oct3/4, a pluripotency marker [4, 20, 22]. As such, UDCs demonstrate propensity for differentiation into several lineages, one of which is muscle-like [20, 23].
We show here that UDCs serve as a noninvasive source for MyoD-induced myogenesis. UDCs transduced with a tamoxifen-inducible MyoD lentiviral vector (iMyoD) formed multinucleate myotubes following induction with tamoxifen and culture in differentiation media for up to 35 days. Urine cell-derived myotubes expressed muscle marker transcripts, formed sarcomeres, and displayed contractile properties. Further, UDCs isolated from individuals with muscular dystrophies were also able to form myotube-like structures and reflected the disease mutations. Thus, UDCs are a ready source for direct reprogramming by MyoD and provide an easily obtainable, noninvasive source for modeling disease mechanisms.
Urine collection and cell isolation
Written and informed consent was obtained from all human subjects. All work was approved by and conducted under the Institutional Review Boards of the University of Chicago (IRB8249) and Northwestern University (IRB STU00104548), which serve as the ethics boards for each institution, respectively. All studies were conducted in compliance with the Helsinki Declaration. Urine samples were collected using midstream clean catch kits according to the directions provided in the kits (Parter Medical Products Inc, Carson, CA; 273516). Cells were isolated within 1 h of collection using a protocol previously published with minor modifications . Briefly, the entire urine samples were equally distributed into 50-mL conical tubes and centrifuged at 400×g for 10 min at room temperature. The supernatant was aspirated leaving ~1 mL of urine into which pellets were resuspended and combined into a single tube, if necessary. Ten milliliters of wash buffer was added per 100 mL of initial urine sample. Samples were centrifuged at 200×g for 10 min at room temperature. The supernatant was aspirated leaving ~0.2 mL, and the cell pellet was resuspended in 1 mL of primary media. All media formulations were obtained from a previously published protocol and are detailed below . Cells were plated in 24-well plates pre-coated with 0.1 % gelatin (Millipore, Billerica, MA; ES-006-B, Stemcell Technologies, Vancouver, Canada; 7903). Roughly one third of the cell suspension was plated in the first well, with the remaining two thirds equally divided into four additional wells. The final volume in each well was then brought to 500 μL with primary media. The plates were placed in a 37 °C incubator with 5 % CO2.
For 3 days, 500 μL of primary media was added to each well every 24 h. On day 4, 1.5 mL of primary media was removed and replaced with 500 μL of proliferation media. An aliquot of the primary media was added to a separate dish containing Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10 % FBS without antibiotics or antimycotics to test for potential contamination. On day 5, all media were removed from each well and replaced with 500 μL of proliferation media, which was changed daily until the isolated cells expanded and were replated in larger dishes. Antibiotics and antimycotics were removed from media once uncontaminated cultures were confirmed. Isolated cells were observed as early as 1 day after the addition of proliferation media. When the cells became confluent or when cell foci began to outgrow the monolayer, cells were trypsinized using 0.25 % trypsin-EDTA (Thermo Fisher Scientific, Waltham, MA; 25200-072), subcultured, and designated as passage 1 (p1). Modifications from  include plating of cells in five wells of a 24-well gelatin-coated plate (vs a single well of 12-well plate), increase of FBS content in the proliferation media to 15 %, and the removal of the antimycotics and antibiotics from the media after lack of contamination was observed.
All media were made following a previously published protocol with the following modifications . Wash buffer consisted of 1× phosphate-buffered saline (PBS) without Ca2+ and Mg2+ (Thermo Fisher Scientific, Waltham, MA; 14190-250) supplemented with 1 % penicillin/streptomycin (Thermo Fisher Scientific, Waltham, MA; 15070-063) and 0.5 μg/mL amphotericin B (Sigma Aldrich, St. Louis, MO; A2942). Primary media were composed of 1:1 mix of high glucose DMEM without sodium pyruvate (GE Healthcare, Logan, UT; SH30022.FS) and Ham’s F-12 Nutrient Mix (Thermo Fisher Scientific, Waltham MA; 11765-054) supplemented with Renal Epithelium Growth Medium SingleQuot Kit Supplements (Lonza, Basel, Switzerland; CC-4127), 10 % fetal bovine serum (Thermo Fisher Scientific, Waltham, MA; 16000-044), 1 % penicillin/streptomycin, and 0.5 μg/mL amphotericin B. Proliferation media were composed of 1:1 mix of Renal Epithelium Growth Medium Bullet Kit (Lonza, Basel, Switzerland; CC-3190) and high glucose DMEM supplemented with 15 % FBS, 0.5 % Glutamax (Thermo Fisher Scientific, Waltham, MA; 35050-061), 0.5 % nonessential amino acids (Thermo Fisher Scientific, Waltham, MA; 11140-050), and 2.5 ng/mL of bFGF (Peprotech, Rocky Hill, NJ; 100-18B, Miltenyi Biotec Inc, San Diego, CA; 130-093-842), PDGF-AB (Peprotech, Rocky Hill, NJ; 100-00AB), and EGF (Peprotech, Rocky Hill, NJ; AF-100-15). The Renal Epithelium Growth Medium (REGM) Bullet Kit was made according to the manufacturer’s instructions, with the omission of the amphotericin B/gentamycin supplement. Freeze media were composed of DMEM (Thermo Fisher Scientific, Waltham, MA; 11995-073) supplemented with 30 % FBS, 1x pen/strep, and 10 % DMSO (Sigma Aldrich, St. Louis, MO; D2650).
Lentiviral construct and transduction
The tamoxifen-inducible MyoD lentiviral construct (pLv-CMV-MyoD-ER(T)), referred to as iMyoD, was previously described  and kindly provided by Dr. Jeffrey Chamberlain (University of Washington; Addgene plasmid # 26809). The construct was packaged by the Skin Disease Research Center DNA/RNA Delivery Core at Northwestern University (Chicago, IL). Urine cells from healthy volunteers or patients were plated in 12-well plates at 75,000 cells/well and transduced within 24–36 h of plating with iMyoD at MOI = 50. Transductions were performed in proliferation media with reduced serum (5 %) and 7.5 μg/mL polybrene (Millipore, Billerica, MA; TR-1003-G) for 6–16 h in a 37 °C incubator.
Evaluation of transduction efficiency
After transduction, an aliquot of cells were plated in two wells of a 24-well plate at 2000–10000 cells/cm2. Cells were treated with 5 μM 4-hydroxytamoxifen (Sigma Aldrich, St. Louis, MO; H7904) for 24 to 48 h, and MyoD was detected using immunofluorescence. Methods and antibodies are as described below. A minimum of three fields and a minimum of 100 cells total were analyzed to determine transduction efficiency. MyoD-positive nuclei and total nuclei were counted per field, and efficiency was calculated by dividing the number of MyoD-positive nuclei by the total number of nuclei per field. Efficiencies were graphed as mean ± standard error of the mean (SEM) using GraphPad Prism (GraphPad, La Jolla, CA). All images were taken using the Floid Cell Imaging Station (Thermo Fisher Scientific, Waltham, MA; 4471136). The cell counts for MyoD-positive nuclei were performed manually using ImageJ (NIH, Bethesda, MD), while the cell counts for total nuclei were performed using the Particle Analyzer function of ImageJ (NIH, Bethesda, MD).
Transduced cells were plated in 24-well plates coated with 8 μg/cm2 collagen I (Sigma Aldrich, St. Louis, MO; C3867, Corning, Corning, NY; 354236) at a density of 37,500 to 50,000 cells/cm2 and cultured until confluent. Cells were then overlaid with Growth Factor Reduced Matrigel (Corning, Corning, NY; 354230) diluted 1:4 in cold DMEM/F-12 50:50 mix (Corning, Corning, NY; 10-092-CV). Matrigel was allowed to gel for 2 to 3 hours at 37 °C. The Matrigel-overlaid iMyoD-treated cells were induced with 4-hydroxytamoxifen (2.5 μM, 24–48 h) diluted in high glucose DMEM (Thermo Fisher Scientific, Waltham, MA; 11995-073) supplemented with 10 % FBS. Hydrocortisone dexamethasone (HD) differentiation media were based on previously published formulation [23, 25] and were composed of high glucose DMEM with sodium pyruvate supplemented with 10 % FBS, 5 % horse serum (Thermo Fisher Scientific, Waltham, MA; 16050-122), 50 μM hydrocortisone (Sigma Aldrich, St. Louis, MO; H0888), and 0.1 μM dexamethasone (Sigma Aldrich, St. Louis, MO; D4902). Two hundred and fifty microliters of differentiation media was replaced every 72 h. Additional coatings tested during plating optimization included collagen IV (Sigma Aldrich, St. Louis, MO; C5533), fibronectin (Sigma Aldrich, St. Louis, MO; F1141), laminin (Sigma Aldrich, St. Louis, MO; L2020), Growth Factor Reduced Matrigel, poly-l-lysine (Sigma Aldrich, St. Louis, MO; P4832), and poly-d-lysine (Sigma Aldrich, St. Louis, MO; P6407).
Primers for PCR
DMD (exons 1 to 3)
DMD (exons 42 to 50/51)
DMD (exons 44 to 48)
Primers for qPCR
Differentiated myotubes were subject to immunofluorescence microscopy at four different time points (7, 14, 28, and 35 days). Myotubes were gently washed with 1× PBS (Thermo Fisher Scientific, Waltham, MA; 14200-166) three times, or until most of the Matrigel was removed, and then fixed in either 4 % paraformaldehyde (15 min at RT) or ice cold 100 % methanol (2 min on ice). Myotubes fixed in paraformaldehyde were subsequently permeabilized in 0.25 % Triton-X (Sigma Aldrich, St. Louis, MO; T8787) for 20 min at RT. Cells were blocked in 10 % horse serum for 30 min to 1 h at 4 °C. Primary antibody incubations were done overnight at 4 °C. Cells were washed three times for 10 min each with PBS, where PBS was supplemented with 0.1 % Triton-X in the second wash. Cells were incubated with secondary antibodies (1 h at RT) and then washed three times for 10 min each as described above. Nuclei were stained with Hoechst 33342 (1:10,000, Thermo Fisher Scientific, Waltham, MA; H3570). Plates were imaged on the Zeiss Axio Observer Z.1 inverted microscope (Carl Zeiss Microscopy, Jena, Germany) and/or Floid Cell Imaging Station. All antibodies were diluted in 0.1 % Triton-X and 2 % horse serum, and all solutions were made with 1× PBS.
Primary antibodies were as follows: rabbit polyclonal anti-MyoD (1:1000, Santa Cruz, Dallas, TX; sc-304), mouse monoclonal anti-desmin (1: 1000, Sigma Aldrich, St. Louis, MO; D1033), mouse monoclonal anti-α-actinin (1: 1000, Sigma Aldrich, St. Louis, MO; A7811), rabbit polyclonal anti-dystrophin (1: 1000, Thermo Fisher Scientific, Waltham, MA; PA1-37587), mouse monoclonal anti-dystrophin (1: 100, Leica Biosystems, Newcastle, UK; NCL-DYSB), mouse monoclonal anti-MYH1 (1: 10 or 2 μg/mL, deposited to Developmental Studies Hybridoma Bank by Fischman, DA; MF 20), mouse monoclonal anti-fast myosin heavy chain (1: 1000, Sigma Aldrich, St. Louis, MO; M4276), mouse monoclonal anti-titin antibody (1: 10 or 2 μg/mL, deposited to Developmental Studies Hybridoma Bank by Greaser, ML; 9 D10), and rabbit polyclonal anti-γ-sarcoglycan (1: 300 to 1: 500 ). Secondary antibodies were as follows: Alexa Fluor 488 donkey anti-rabbit (1: 1,1000; A21206), Alexa Fluor 488 donkey anti-mouse (1: 1000; A21202), Alexa Fluor 594 donkey anti-rabbit (1: 1000; A21207), and Alexa Fluor 594 donkey anti-mouse (1: 1,1000; A21203, all Thermo Fisher Scientific, Waltham, MA).
DMD exons 42 through the junction of exons 50 and 51 were amplified by PCR, gel purified using QIAEXII Gel Extraction Kit (Qiagen, Valencia, CA; 20021). The purified products were amplified by PCR using nested primers for DMD exons 44 through 48 and then sequenced at the NUSeq Core at Northwestern University (Chicago, IL). Chromatograms and sequences were analyzed on SeqMan Pro (DNASTAR, Madison, WI) and FinchTV (Geospiza, Seattle, WA).
To generate a point mutation in control urine cells, we used the Cas9-GFP plasmid designed by the Zhang lab (Addgene plasmid #48138) . The CRISPR design tool (crispr.mit.edu) was used to identify and select guide RNA (gRNA) targets in human SGCG exon 6. The target on the anti-sense strand was gRNA 5′ CTAAGGTCTTGAAACGGGT 3′. This guide permitted a synonymous mutation of the protospacer adjacent motif (PAM) and preserved the 5′ SGCG exon 6 sequence. The gRNA was expressed from the U6 polymerase III promoter as previously described . The promoter, crRNA-tracrRNA, and target were synthesized as a gBlock by Integrated DNA Technologies (IDT, Coralville, IA), then PCR amplified (F: TGTACAAAAAAGCAGGCTTTAAAG; R: TAATGCCAACTTTGTACAAGAAAG), and purified using the Qiagen PCR purification kit (Qiagen, Valencia, CA; 28106). The single-stranded oligodeoxynucleotide (ssODN) was designed according to published guidelines . The ssODN, based on the anti-sense strand template, was centered in between the nuclease site and the desired thymine deletion with 71 bp flanking arms on either side (142 bp total). It contained a downstream G > A synonymous mutation that destroyed the PAM –NGG site (CGG > CAG). The ssODN ultramer ggagaggttgttaatatttgaacaaaaattcttacCTAAGGTCTTGAAACGGGTCaGCTCTGACAAGGGGTGTCTCCACTGAATGTTCAAAAGAGCCCCTTCAGGCCctaaacaaaaaacaaagatgtatcaggagcagaag was synthesized and PAGE purified by IDT. Uppercase letters indicate the SGCG exon 6 coding region, and lowercase letters indicate the flanking intronic regions. The single underlined region encodes the targeted mutation of the protospacer adjacent motif (PAM), and the italicized sequence is the 4 adenine (AAAA) sequence which encodes the single thymine (T) deletion in the exon 6 coding region.
Electroporation with CRISPR reagents
On the day of electroporation, urine cells were trypsinized, centrifuged at 400×g for 5 min (RT), and resuspended in PBS. An aliquot was removed for cell counting, and cells were centrifuged at 400×g for 5 min (RT). Cells were resuspended in electroporation buffer 1 × 106 cells/mL (Bio-Rad, Hercules, CA; 165-2677) and 2 × 105 cells were added to the Cas9 GFP plasmid (1 μg), U6 gRNA (500 ng), and ssODN (0.5 nmol) reagents. After gentle mixing, the cells were placed in a sterile cuvette with a 0.4-cm gap (Bio-Rad, Hercules, CA; 1652088) and electroporated using Gene Pulser Xcell Electroporation System (Bio-Rad, Hercules, CA; 1652660) with the following settings: (1) single square wave pulse, 200 V, 20 msec or (2) three square waves pulses, 150 V, 20 msec (0.1 s between pulses). The electroporated cells were added to gelatin-coated six-well plates and brought to a final volume of 2 mL with proliferation media. The media were replaced within 12 h and replaced every 24 h for 3–5 days until fluorescence-activated cells sorting (FACS).
FACS sorting and clonal expansion
Prior to cell sorting, 96-well plates were pre-coated with 0.1 % gelatin and placed in the incubator for 1 h. Gelatin was removed and replaced with 100 μL of proliferation media that were supplemented with 0.5× pen/strep. The electroporated cells were trypsinized, centrifuged, and resuspended in 1 mL of proliferation media supplemented with 0.5× pen/strep. Single cell sorting of GFP-positive cells into 96-well plates was conducted by the Northwestern Flow Cytometry Facility using the FACS AriaII (BD Biosciences, San Jose, CA). After sorting, 96-well plates were briefly centrifuged at 200×g for 1 min (RT) and placed in a 37 °C incubator. After 48 h, 100 μL of proliferation media without pen/strep was added to the 96-well plates. A complete media exchange was performed on days 4, 8, and 12 using proliferation media without pen/strep. Expansion of single cell clones was evident 7–10 days after FACS, with confluent wells observed about day 14. Confluent wells were subcultured onto gelatin-coated 24-well plates until confluent and then subcultured onto gelatin-coated six-well plates.
Genotype analysis of clones
Clones expanded to the six-well stage were analyzed for genome editing via the CRISPR/Cas9 system. Genomic DNA was isolated and PCR amplified using the GeneArt cleavage detection kit (Thermo Fisher Scientific, Waltham, MA; A24372) according to the manufacturers’ instructions. The primer set amplified a genomic region containing SGCG exon 6 along with a portion of the flanking introns (F: TAGGGAAAGAGTTGGGCCTATTT) (R: GAGGCAGATTGGAGAGGAAGAA). To test for clones edited through homology-dependent repair (HDR), PCR-amplified products were subjected to Alu1 digest and restriction fragment length polymorphism (RFLP) analysis. In addition, PCR products from individual clones were sequenced at the NUSeq Core at Northwestern University (Chicago, IL). Chromatograms and sequences were analyzed using SeqMan Pro (DNASTAR, Madison, WI).
Generation of urine-derived cells with inducible MyoD expression
Urine cells were transduced with the iMyoD lentivirus, regularly achieving >80 % efficiency with an MOI of 50, as determined by immunofluorescence microscopy and image analysis for MyoD expression (Fig. 1c, d). In the absence of tamoxifen, MyoD was seen in cytoplasm. After tamoxifen exposure, MyoD staining also became nuclear (Fig. 1c). Because lentiviral transduction was of such high efficiency, nuclear MyoD staining was routinely evident in 80 % of the cultures (Fig. 1d).
Morphological and molecular myogenic reprogramming of iMyoD urine-derived cells
Myogenic differentiation and maturation
MyoD controls temporal expression of early and late genes in muscle development through a feed-forward mechanism . To characterize myogenic marker expression, we tested tamoxifen-treated iMyoD cells by RT-PCR. Robust and sustained expression of MYOG (myogenin) and MYH3 (embryonic myosin) was observed (Fig. 3b). In addition, there was a temporal increase in expression of ACTN2 (α-actinin) and DMD (dystrophin) (Fig. 3b). Expression of MYH2, the adult form of skeletal myosin, was only detected when differentiation was extended to the 28d time point (Fig. 3b). Similarly, real-time PCR detected most transcripts after 7 days of differentiation, with sustained or moderately increased expression over time (Fig. 3c). However, DMD and MYH2 demonstrated greater temporal increases in transcript expression as myogenic maturation progressed (Fig. 3c). Evaluation of myotubes by immunofluorescence microscopy showed accumulation of α-actinin, dystrophin, and fast myosin heavy chain (MHC), concordant with the messenger RNA (mRNA) data. Expression of these muscle proteins was associated with increased myotube formation, organization, and elongation, indicative of myogenic differentiation and maturation over time (Fig. 3d).
Movie 1. Video showing spontaneous myotube twitching. (MOV 1366 kb)
Movie 2. Video showing spontaneous myotube twitching. (MOV 1869 kb)
Direct reprogramming of urine cells to model muscular dystrophy
CRISPR/Cas9 mediated genome editing in single urine cell clones
The genomic region including SGCG exon 6 was PCR amplified to analyze the results of gene editing. Restriction digest of the PCR products revealed the presence of an Alu1 site in a subset of clones, indicative of HDR repair (Fig. 7c). Sequence analysis demonstrated efficient genome editing in UDCs with the expected variety of both heterozygous and homozygous mutations, including insertions and deletions from nonhomologous end joining (NHEJ) repair and specific point mutations directed by HDR (Fig. 7c, d). Generation of clones with both heterozygous and homozygous 521ΔT mutations was evident in line 2 (Fig. 7c, d). Interestingly, a subset of HDR-edited clones contained a point mutation in the PAM region without the upstream thymine deletion, suggesting that this region is refractory to editing and/or required a gRNA target closer to the 5T region. A summary of the gene editing results is shown in Fig. 7b, Additional file 3: Table S1 and S2, and Figure S1 and S2. Collectively, these data demonstrated the ability to edit the genome of urine-derived cells with CRISPR/Cas9, generating single cell clones with a specific point mutation.
The limited availability of human skeletal muscle tissue encumbers the study of muscle development and the pathology of muscle disease. Many novel strategies have been taken to overcome this limitation, relying on genetic engineering with direct and indirect reprogramming of cells. Regardless of strategy, a key component to success is the initial cell source, specifically the ease of acquisition and ability to reprogram. Urine is an underutilized, noninvasive, and cost effective source of primary human cells that has served as a platform for urothelial-based culture and iPSC reprogramming [19–21]. Recent advances in urine cell isolation have generated clonal populations of autologous human cells with a high proliferative capacity, evident telomerase activity, and plasticity for reprogramming into multiple lineages [20, 21]. These characteristics make urine-derived cells an attractive source of primary human cells for in vitro models. We now extended these findings showing that inducible expression of MyoD efficiently reprogrammed UDCs into the myogenic lineage, providing a simple strategy to generate an in vitro model of skeletal muscle from virtually any human subject.
Urine cells demonstrate an inclination towards the mesodermal lineage. A subpopulation of cells, classified as urine stem cells, express pericyte and mesenchymal stem cell markers and were able to differentiate into podocytes, smooth muscle, and urothelial cells [20, 21, 32]. The ability of UDCs to differentiate into skeletal muscle is less defined. Studies showed growth on specific biomaterials, presence of specific growth factors, or defined media modifications promoted expression of myogenic markers such as MyoD, desmin, and myosin [23, 34]. However, these cells were not observed to form striations nor to display contractile properties. Given the known expression of these markers, it is possible that without sufficient MyoD, UDCs differentiate more towards smooth rather than striated muscle lineages . The strategy with an induced MyoD pulse combined with media manipulation drove urine cells through the initial stages of myogenic development into a more mature phenotype, complete with expression of dystrophin, adult myosin, titin, and γ–sarcoglycan, along with the presence of functional sarcomeres. Despite these advancements, this cellular model still lacks major components of mature muscle such as transverse tubules and resident stem cell niches.
An alternative strategy to direct reprogramming with MyoD would be to first reprogram UDCs into iPSCs and then use the iPSC lines for myogenic differentiation. This approach has the advantage of iPSC self-renewal, although immortalization of UDCs directly could obviate this need in vitro. DMD UDCs were previously successfully reprogrammed into iPSC and differentiated into cardiomyocytes, recapitulating cardiac phenotypes in vitro .
We also showed that UDCs can be directly gene edited using CRISPR/Cas9 technology. The use of gene editing for muscle disease has focused intently on genetic correction strategies with significant success on the study of muscular dystrophy, as recent studies showed the ability to correct DMD disease-causing mutations in animal models and human cells [35–43]. However, the ability to insert mutations of interest is also useful since it allows the study of disease models without the need to access patients directly. Nevertheless, gene editing in human iPSCs is challenged by the availability of patient cell lines, isolation of pure clonal populations, and low editing efficiency, especially when targeting precise mutations with homology-dependent repair [29, 44, 45]. While some technical aspects of the CRISPR/Cas9 system are still undergoing improvement, the relative ease of urine cell isolation, coupled with the ability to generate single cell clones, makes this an attractive primary cell platform for CRISPR/Cas9 gene editing.
Here, we showed that direct reprogramming of urine cells using MyoD is feasible from patients with muscle disease and can recapitulate the basic disease phenotype. Because urine cells are readily available and collected noninvasively, this approach provides a highly useful model of human muscle disease.
Fetal bovine serum
Dulbecco’s Modified Eagle Medium
Fibroblast growth factor
Platelet-derived growth factor
Epidermal growth factor
Reverse transcription polymerase chain reaction
Clustered regularly interspaced short palindromic repeats
CRISPR-associated protein 9
Protospacer adjacent motif
Nonhomologous end joining
Double strand DNA break
Fluorescence-activated cells sorting
Restriction fragment length polymorphism
Limb girdle muscular dystrophy
Duchenne muscular dystrophy
Myosin heavy chain
Induced pluripotent stem cells
Standard error of the mean
We thank the patients for their participation and the Kurt+Peter Foundation. We acknowledge the Skin Disease Research Center, Flow Cytometry Core, and NUSeq Core Facilities at Northwestern University.
This work was supported by NIH HL61322, NIH T32 HD009007, and the Kurt+Peter Foundation.
EYK conducted the experiments, analyzed the data, prepared the figures, and wrote the manuscript; PP conducted the experiments; LDC coordinated the collection of human materials; EMM analyzed the data and wrote and edited the manuscript; EW conducted the experiments, analyzed the data, prepared the figures, and wrote the manuscript. All authors read and approved the final manuscript.
The authors have no competing financial interests related to the conduct of the work presented in this manuscript.
Ethics approval and consent to participate
Written and informed consent was obtained from all human subjects. All work was approved by and conducted under the Institutional Review Boards of the University of Chicago (IRB8249) and Northwestern University (IRB STU00104548), which serve as the ethics boards for each institution, respectively. All studies were conducted in compliance with the Helsinki Declaration.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
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