Author contributions: E.B.R.: conception and design, data collection and analysis, and manuscript writing; S.A.G. and R.A.D.: conception and design and data collection and analysis; C.P., R.M.K.T., B.O., and D.A.C.: data collection; C.O.R.: conception and design; P.R.: data analysis. J.R.: manuscript writing; J.M.H.: conception and design and manuscript writing.
The presence of tissue specific precursor cells is an emerging concept in organ formation and tissue homeostasis. Several progenitors are described in the kidneys. However, their identity as a true stem cell remains elusive. Here, we identify a neonatal kidney-derived c-kit+ cell population that fulfills all of the criteria as a stem cell. These cells were found in the thick ascending limb of Henle's loop and exhibited clonogenicity, self-renewal, and multipotentiality with differentiation capacity into mesoderm and ectoderm progeny. Additionally, c-kit+ cells formed spheres in nonadherent conditions when plated at clonal density and expressed markers of stem cells, progenitors, and differentiated cells. Ex vivo expanded c-kit+ cells integrated into several compartments of the kidney, including tubules, vessels, and glomeruli, and contributed to functional and morphological improvement of the kidney following acute ischemia-reperfusion injury in rats. Together, these findings document a novel neonatal rat kidney c-kit+ stem cell population that can be isolated, expanded, cloned, differentiated, and used for kidney repair following acute kidney injury. These cells have important biological and therapeutic implications. STEM Cells 2013;31:1644–1656
The search for putative stem cells or precursors within the kidney has been the focus of extensive research. The identification of a kidney stem cell would provide important biological insights and could therapeutically be used to generate new tubular, glomerular, and vascular cells in the treatment of both acute and chronic kidney injuries.
To date several possible kidney stem cell candidates have been described. These include label retaining cells (LRCs) or slow-cycling cells identified by using bromodeoxyuridine pulse-chase analyses. LRCs are detected in proximal tubules, thick ascending limb (TAL) of Henle's loop distal tubules, and collecting ducts after a short (2-week) chase period  or in the interstitium and in the papillary tubules after a long (2-month) chase period [2, 3]. Other stem cell candidates include kidney cells expressing surface markers and found in different locations, such as the interstitium (Sca1) [4, 5], Bowman's capsule (CD24, CD133) [6–9], papilla (Nestin, CD133) , and proximal tubular compartment (CD24, CD133) [11, 12] (or only CD133) . These studies demonstrate multipotentiality in vitro and the capacity of these cells to integrate into the kidney during development or in response to injury.
However, kidney epithelial tubular regeneration has been the subject of intense debate generating multiple hypotheses. Cell-tracking studies using transgenic mice provide strong evidence in favor of an intratubular regeneration source, suggesting that differentiated epithelial cells that survive acute injury undergo proliferative expansion [14, 15]. More recently, a study involving two-step sequences of nucleotide analog pulses following murine ischemia-reperfusion injury (IRI) further suggests an absence of kidney stem cells in the adult kidney . Moreover, telomerase activity-expressing cells were reported in 5% of the LRCs but are not involved in kidney repair .
These studies generated controversy in the field, because they challenged the significance of work from many groups investigating the existence and the role of putative postnatal kidney stem cells. Notably, Lin et al.  and Humphreys et al. [15, 16] do not provide conclusive evidence for the absence of postnatal kidney stem cells and they do not eliminate the possibility of a tubular stem cell population, possibly of more limited potency. Those cells derived from the Six-2+ cap mesenchyma or expressing kidney specific-cadherin would be identically labeled in the regenerating tubules. Furthermore, there is evidence that in the renal papilla, LRCs or their immediate progeny are able to proliferate and migrate, as demonstrated in transgenic mice conditionally expressing GFP fused to histone 2B . Additionally, the SDF-1/CXCR4 axis is involved in the papillary LRC activation after acute kidney injury .
Studies of other organs have engendered similar controversy. In the pancreas, studies using a transgenic reporter mouse strain showed that the major source of new β-cells during adult life and after pancreatectomy arose from the proliferation of terminally differentiated β-cells rather than from pluripotent stem cells . However, more recently, rare pancreas-derived multipotent precursor cells that form spheres, express insulin and generate multiple pancreatic and neural cell types in vivo, were observed in adult human tissue . The presence of differentiation markers was also described in human neuronal stem cells that display morphologic and molecular characteristics of differentiated astrocytes .
Expression of c-kit receptor, a tyrosine kinase receptor, is detected in differentiated cells that do not exhibit stem cell properties, such as mast cells, germ cells, melanocytes, gastrointestinal Cajal cells, fetal endothelial cells, and epithelial cells, including breast ductal, sweat gland, some cells of skin adnexa, and cerebellar neurons . However, c-kit+ cells have been described as a marker of stem cells in many organs and tissues, such as bone marrow , liver , heart , amniotic fluid , and lungs .
C-kit+ cells have also been identified during metanephric mesenchyme (MM) development and the ligand for c-kit, stem cell factor (SCF), is abundantly expressed in the ureteric bud . Therefore, we hypothesized that c-kit+ cells isolated from neonatal rat kidney represent a population of stem cells. Here, we show that c-kit+ cells possess stem cell properties, including self-renewal capacity, clonogenicity, and multipotentiality. Furthermore, they exhibit the potential to treat renal failure by multicompartment engraftment, including tubular, vascular, and glomerular, following acute IRI.
Materials and Methods
Explant Culture of Neonatal Rat Kidney
Neonatal rat kidneys from Sprague-Dawley (SD; n = 6–8) were harvested, chopped, digested with collagenase II, and incubated in red blood cell lysing buffer (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com). Expansion medium included Dulbecco's modified Eagle medium (DMEM/F12), 20% fetal bovine serum (FBS), 100 U/mL penicillin, and 100 μg/mL streptomycin (Sigma-Aldrich).
Immunopanning and Fluorescence-Activated Cell Sorting
C-kit+ cells were isolated by immunopanning using rabbit polyclonal c-kit antibody (H300, Santa Cruz, Santa Cruz, CA, http://www.scbt.com) and further selected using fluorescence-activated cell sorting (FACS) (FACSAria, BD Biioscience, Sand Diego, CA, http://www.bdbioscience.com). Depletion of hematopoietic stem cells lineage was also performed to ensure that c-kit+ cells came from kidney and that bone marrow-derived cells were removed. Therefore, the APC lineage antibody cocktail was used, which depletes CD3e, CD11b, CD45R/B220, erythroid cells, and Ly-6G and Ly-6C (BD Pharmingen, San Diego, CA, http://www.bdbiosciences.com/index[lowen]us.shtml), and the anti-mouse CD117-PE conjugated (eBioscience, San Deigo, CA, http://us.ebioscience.com).
Expansion of C-kit+ Cells
Sorted c-kit+/Lin− cells were plated and cultured in DMEM/F12 supplemented with 10% FBS, 10 ng/mL basic fibroblast growth factor (bFGF), 20 ng/mL epidermal growth factor (EGF), 40 ng/mL SCF (PeproTech, Rocky Hill, NJ, http://www.peprotech.com), 10 ng/mL leukemia inhibitory factor (LIF; Millipore, Billerica, MA, http://www.millipore.com), insulin-transferrin-selenium A liquid media supplement (ITS; Invitrogen, Carlsbad, CA, http://www.invitrogen.com), and antibiotics.
Quantitative Real-Time Polymerase Chain Reaction and Immunofluorescence
All experiments for c-kit characterization were performed 5–10 passages after sorting. For quantitative polymerase chain reaction (qPCR), total RNA was extracted from cells using Pure-Link Micro-to-Midi Total RNA Purification System (Qiagen, Hilden, Germany, http://www1.qiagen.com) and reverse-transcribed using High capacity cDNA reverse transcription kit (Applied Biosystems, Foster City, CA, http://www.appliedbiosystems.com). All samples were treated with Turbo DNase (Ambion, Austin, TX, http://www.ambion.com). qPCR was performed in triplicate using a 20 μL reaction mixture containing 10 ng cDNA, TaqMan Universal PCR Master Mix (Roche, Basel, Switzerland, http://www.roche-applied-science.com) and primers/probes sets for specific genes (TaqMan Gene Expression Assays, Applied Biosystems). As an internal control glyceraldehyde 3-phosphate dehydrogenase or 18s was determined in each reaction. Reactions conditions were performed according to the manufacturer: one cycle of 50°C for 2 minutes, one cycle of 90°C for 10 minutes, and 40 cycles of 95°C for 15 seconds and 60°C for 1 minute. Software from iQ5 multicolor real-time PCR detection system (Bio-Rad, Hercules, CA, http://www.bio-rad.com) was used for PCR analyses. Relative fold change was calculated by 2ΔΔCt method and compared to baseline values (set at 1). For immunofluorescence, cells were fixed in paraformaldehyde 4%, blocked with bovine serum albumin (BSA) 1%, and Tween-20 0.5%, incubated with primary antibodies, and then incubated with 488- or 568-conjugated secondary antibodies (Invitrogen).
Fragments of kidney were fixed overnight in neutral buffered formalin 10%, dehydrated in alcohol, and embedded in paraffin. Sections of 4–5 μm thickness were stained with hematoxylin and eosin (H&E) and Periodic Acid Schiff reagent. For immunofluorescence, renal sections were deparaffinized with xylene and rehydrated in alcohol series and water. The sections were subsequently microwaved twice for 10 minutes in citrate buffer and then blocked for 1 hour in donkey serum. Primary antibodies were applied for 1 hour at room temperature or overnight at 4°C. Incubations for 1 hour using 488- or 568-conjugated secondary antibodies (Invitrogen) were then performed. For green fluorescent protein (GFP) staining, an antibody goat or rabbit polyclonal anti-GFP fluorescein isothiocyanate (FITC)-conjugated (Abcam, Cambridge, U.K., http://www.abcam.com) was applied for 1 hour at 37°C. Nuclei labeling was obtained with 4′,6-diamidino-2-phenylindole (DAPI). After that, slides were incubated with Sudan Black 0.1% (Sigma-Aldrich) at room temperature. In control experiments, the first antibody was omitted.
A total of 1–2 × 106 c-kit+ cells were used for FACS characterization. For intranuclear markers, cells were fixed in cold 80% ethanol, washed twice with 1 mL phosphate buffered saline (PBS) during centrifugation for 10 minutes at 200g, incubated 1 hour with FACS buffer (1% BSA and 5% FBS diluted in distilled water) on ice, and subsequently 1 hour with the primary and secondary antibodies. For intracellular markers, we used the BD Cytofix/Cytoperm Fixation/Permeabilization kit (BD Pharmigem); for surface markers, periods of incubation with FACS buffer for 1 hour, and primary and secondary antibodies incubation were also performed. Each analysis included at least 10,000 events and was performed on at least three separate cell preparations.
In Vitro Differentiation
For endothelial differentiation, c-kit+/Lin− cells were cultured in endothelial cell growth medium-2 (EGM-2; Lonza, Walkersville, MD, http://www.lonza.com) supplemented with 2% FBS, EGF, vascular endothelial growth factor (VEGF), insulin-like growth factor (IGF), bFGF, hydrocortisone, ascorbic acid, and heparin for 1–4 weeks. For in vitro Matrigel tube formation assay, cells were trypsinized and plated on 300 μL Matrigel, reduced growth factor (BD Pharmigem). Seeding density was 10 × 104 cells per well and endothelial cell basal medium without supplements was added in 0.1% BSA. Tube formation was analyzed at two time points; 6 hour and 24 hour. Number of tubes and tube length (μm) were calculated by using image J software (imagej.nih.gov). For epithelial differentiation, cells were incubated in DMEM containing 10% FBS, 50 ng/mL bFGF, 20 ng/mL LIF, and 5 ng/mL transforming growth factor beta (TGF-β) for 3 weeks. For adipogenic differentiation, cells were incubated in DMEM high glucose containing 10% FBS, 1 μM dexamethasone, 0.5 μM 1-methyl-3-isobuthylxanthine, 10 μg/mL insulin, and 100 μM indomethacin (Sigma-Aldrich) for 2 weeks. For osteogenic differentiation, cells were cultured in minimum essential medium alpha (MEM-α) and 10% FBS that contained 107 M dexamethasone, 0.2 mM ascorbic acid, and 10 mM β-glycero-phosphate (Sigma-Aldrich) for 4 weeks. Oil red O and Alizarin S staining quantification protocols are described in Supporting Information. We used GFP transgenic rat to isolate mesenchymal stem cells (MSCs) from bone marrow and perform mesodermic differentiation as a positive control. For neuronal differentiation, c-kit+ cells were plated on fibronectin-coated dishes at a seeding density of 5 × 103 cells per centimeter square in DMEM/F12 supplemented with 5% FBS and 100 ng/mL bFGF (PeproTech) for 2 weeks.
For cell tracking, c-kit+ cells were transfected with a lentiviral vector carrying the gene encoding GFP. The protocol is described in Supporting Information.
In Vivo Endothelial Differentiation
All procedures involving animals were approved by the respective Institutional Animal Care and Use Committees of the University of Miami. C-kit+/Lin− cells were labeled with GFP and cultured in endothelial medium for 1 week. They were then subcutaneously injected into non-obese diabetic-severe combined immunodeficiency (NOD-SCID) mice (n = 3; 2 × 106 cells) in a Matrigel plug. Negative control included the Matrigel plug containing only EGM-2 (n = 3). Positive control included human umbilical vein endothelial cells (HUVEC).
Calcium (Ca2+) Transient
Intracellular Ca2+ was measured using the Ca2+-sensitive dye Fura-2 and a dual-excitation spectrofluorometer (IonOptix LLC, Milton, MA, http://www.ionoptix.com) and is described in Supporting Information.
Dissociated single c-kit+ cells obtained from c-kit sheets and from clones obtained in 96-well plates were plated on ultra-low attachment six-well plates (Corning, Costar, Acton, MA, http://www.corning.com/lifesciences) at clonal density, 1 × 103 cells per well, in DMEM-F12 medium containing 20% knockout serum replacement, 10 mM MEM nonessential amino acids, 0.2 mM β-mercaptoethanol (Gibco, Grand Island, NY, http://www.invitrogen.com), l-glutamine (Sigma-Aldrich), and 20 ng/mL bFGF. Medium was changed every 3 days. After 12 days, cultures were assessed for nephrosphere number. Nephrospheres were defined as free-floating spheres of >40 μm diameter, and results were expressed as a percentage of the plated cells. To assess size and number, spheres were visualized with a Nikon Eclipse TS100 (Nikon, Melville, NY, http://www.nikoninstruments.com) inverted microscopic fitted with a Nikon digital camera image capture system and analyzed with image J software.
Acute Kidney IRI
Briefly, we applied vascular clamps across both renal pedicles for 35 minutes in female 2 month old SD rats weighing 200–250 g (Charles River). After removing the clamps, reperfusion was visually observed. Subsequently, 2 × 106 cells (GFP-labeled c-kit+ cells or MSCs from GFP-SD) or saline was injected directly into the abdominal aorta above the renal arteries, after application of a vascular clamp to the abdominal aorta below the renal arteries. The needle used for cell injection was 31G, length 8 mm (5/16″) insulin syringe needle (BD Biosciences). Cells were then resuspended in 300 μL of saline. The same volume of saline (300 μL) was injected into the aorta in the control group. Blood collection was performed at baseline, days 1, 2, 4, and 8 post-IRI for creatinine and blood urea nitrogen (BUN) measurement (Products Vitros Chemistry, Rochester, NY, http://www.cmmc.org). Kidneys were harvested after 8 days for histological analyses.
Morphologic studies, Immunofluorescence, and Proliferating Cell Nuclear Antigen Index on Kidney Tissue
Acute tubular necrosis (ATN) was assigned by semiquantitative analysis of each individual variable (casts, brush border loss, tubular dilation, necrosis, and calcification) to augment the ATN score (maximum 7). Scoring for proliferating cell nuclear antigen (PCNA)-positive cells was carried out by counting the number of positive nuclei in four randomly chosen kidney cortex and outer medulla sections using ×20 magnification and after applying a rabbit polyclonal PCNA antibody (Santa Cruz). Data from all fields and all kidneys were pooled to obtain PCNA score.
Error bars represent mean ± SEM or mean ± SD, as indicated. The means of two populations were compared by Student's t test or Mann-Whitney test. For multiple comparisons, ANOVA was used. Differences were considered significant at p < .05.
Neonatal Rat Kidneys Contain C-kit+ Stem Cells
Cells expressing the c-kit epitope on their cell surface were widely distributed in the neonatal rat kidney, localized not only to renal papilla (Fig. 1A) but also to the medulla and the nephrogenic zone. These cells expressed E-cadherin (Fig. 1B) and N-cadherin (Fig. 1C). C-kit+ cells were located primarily within a laminin-positive membrane, indicating that they are epithelial cells (Fig. 1D). In contrast, c-kit did not colocalize with Dolichos biflorus agglutinin (Fig. 1E), a marker of ureteric bud and its derivates, or with the Na-Cl cotransporter (NCCT/SLC12A3), a distal tubule marker (Fig. 1F). However, c-kit colocalized at the apical membrane of epithelial cells of the TAL of Henle's loop with the Na-K-2Cl cotransporter (NKCC2/SCL12A1) in both nephrogenic cortex (Fig. 1G) and medulla (Fig. 1H). Aquaporin 1 (AQP1) did not colocalize with c-kit (Fig. 1I). C-kit+ cells were not detected in vessels or glomeruli. Importantly, in the adult rat kidney, c-kit+ cells exhibited identical distribution as found in neonatal rat kidney, for example, colocalization with NKCC2 in the TAL (Supporting Information Fig. S1).
Next, we isolated c-kit+ cells and evaluated their stemness properties in vitro. C-kit+ cells were isolated from neonatal rat kidney explants (n = 6–8) by immunopanning and FACS (Fig. 1J). These cells were found to be Lin− (lineage cells depleted) and represented 1.1% of the cells (∼0.15% per kidney) (Fig. 1K). These cells exhibited the ability to self-replicate and grow in a monolayer on plastic (Fig. 1L). After sorting, the c-kit epitope remained detectable by immunofluorescence microscopy (Fig. 1M). By FACS, 88.6% ± 5.5% of the cells were positive for c-kit, and this high level of c-kit positivity continued up to 50 passages (76.2% ± 8.6%) (Supporting Information Fig. S2).
Characterization of c-Kit+/Lin− Cells
We observed that c-kit+/Lin− cells expressed proteins associated with early stem cells and reprogramming genes, such as Oct4, sex-determining-region Y-box 2 (Sox2), c-myc, and Kruppel-like factor 4 (Klf4) (Fig. 2A; Supporting Information Fig. S3A). Negative control was obtained by omitting the primary antibodies. All those markers were confirmed by immunofluorescence staining (Fig. 2B) and by qPCR (Fig. 2C; Supporting Information Table S1). Vascular (von Willebrand factor [vWF], isolectin, α-actin 2 [Acta2]), epithelial (ZO-1, NKCC2, NCCT, AQP1), neuronal (nestin and neurofilament-heavy chain [NF-H]), and mesenchymal (CD73, CD90, vimentin) markers were also detected (Fig. 2B, 2C; Supporting Information Fig. S3B). A low percentage of kidney-derived c-kit+/Lin− cells expressed CD24 (<10%), CD133 (∼30%), and Pax2 (∼30%) (Supporting Information Fig. S3B).
By qPCR, CD73, NF-H, AQP1, CD90, vimentin, and Klf4 expression was at least 2.5-fold higher than neonatal rat kidney (Fig. 2C). C-kit+/Lin− cells were negative for CD45, a marker of hematopoietic cells (Fig. 2B).
C-kit+/Lin− cells were subcultured for more than a year (>100 passages) without any evidence of senescence or growth arrest (Supporting Information Fig. S4A). Cells frozen at different passages and thawed 6 and 12 months later retained their original characteristics. Similar telomerase activity was detected at different passages of c-kit+/Lin− cells (Supporting Information Fig. S4B). Moreover, c-kit+/Lin− cells exhibited a normal karyotype (Supporting Information Fig. S4C).
Nonclonal C-kit+-Derived Cells Differentiate into Mesoderm and Neuroectoderm Lineages but not into Endoderm
To assess their plasticity, c-kit+ cell monolayers were treated for 1–4 weeks with differentiation media to promote adipogenic, osteogenic, neuronal, epithelial, or endothelial differentiation (Fig. 3A). The cells successfully differentiated and expressed markers for these cell types, as assessed by immunochemical and histochemical stainings and qPCR (Fig. 3B–3E).
C-kit+/Lin− cells grown in adipogenic medium for 2 weeks accumulated lipid droplets, that stained positive for Oil Red O, and upregulated peroxisome proliferator-activated receptor-gamma (PPAR-γ) and adiponectin (Fig. 3B). Later passage cells continued to show commitment to adipogenic differentiation, although it was less pronounced (Supporting Information Fig. S5A). MSCs, the positive control for mesoderm differentiation, significantly exhibited higher lipid accumulation than c-kit+ early and late passage cells (Supporting Information Fig. S5B). PPAR-γ upregulation was comparable between c-kit+ cells and MSCs, whereas adiponectin was higher in c-kit+ differentiated cells. (Supporting Information Fig. S5C).
Growing c-kit+/Lin− cells in osteogenic medium for 4 weeks resulted in Alizarin Red S positivity, indicative of mineralization, which correlated with a significant upregulation of Runx2 and alkaline phosphatase (AP) expression (Fig. 3C). Osteopontin expression was not significantly upregulated. Later passages also exhibited Alizarin Red S positivity (Supporting Information Fig. S5D). Similar to adipogenic differentiation, MSCs had more Alizarin Red S staining compared to c-kit+ cells and a greater upregulation of Runx2 and osteopontin (Supporting Information Fig. S5E). At baseline, AP was expressed at low levels in c-kit+ cells, as opposed to MSCs; after differentiation, however, AP upregulation was more pronounced in c-kit+ cells (Supporting Information Fig. S5F).
After 2 weeks in the neuronal medium, c-kit+/Lin− cells at low density decreased their proliferation and exhibited prolongations (Fig. 3D). Cells were positive for β-3 tubulin which colocalized with NF-H (Fig. 3D). β-3 Tubulin was significantly upregulated (Fig. 3D).
Epithelial differentiation was induced by growing c-kit+/Lin− cells in medium containing bFGF, TGF-β3 , and LIF  for 3 weeks. After 1 week in epithelial medium, the morphology of c-kit+/Lin− cells changed and they started to form packed clusters. These clusters detached after 3 weeks and acquired an embryoid body-like morphology (Fig. 3E). Even late passage (P50–P52) cells acquired this morphology (Supporting Information Fig. S5G). CD24, cytokeratin 18 (KRT18), Wnt4, Notch2, and AQP1 were all upregulated (Fig. 3E), suggesting mesenchymal–epithelial transition [32, 33]. In these epithelial spheres, E-cadherin colocalized with pan-cytokeratin (Fig. 3E). We did not detect albumin or alpha-fetoprotein after growing c-kit+/Lin− cells in the hepatocytic medium, suggesting that these cells have limited to no capacity for endodermal differentiation.
C-kit Vascular Differentiation Is Associated with Functional Activity In Vitro
Based on the presence of vascular markers, we performed in vitro endothelial differentiation by culturing c-kit+/Lin− cells in endothelial basal medium supplemented with growth factors (VEGF, bFGF, IGF-1, and EGF) for 1–4 weeks. Between the third and fourth week, myotube-like structures appeared (Fig. 4A), which stained for Acta2 and costained for vWF (Fig. 4B).
Endothelial tubes were examined at two time points (6 hour and 24 hour) by the in vitro Matrigel assay performed on c-kit+/Lin− cells (Fig. 4C). Tubes were more abundant and longer after 24 hours compared to 6 hours (Fig. 4D). Both early (P15-20) and late (P50-71) passage cells formed endothelial tubes on Matrigel (Supporting Information Fig. S6A). C-kit+/Lin− cells produced significantly more but shorter tubes compared to MSCs at 24 hours (Supporting Information Fig. S6B). In vivo endothelial differentiation demonstrated that GFP-labeled c-kit+/Lin– cells embedded in Matrigel formed network-like connections when injected into NOD-SCID mice (Fig. 4E). These connections were positive for Acta2 and platelet endothelial cell adhesion molecule-1 (PECAM-1) (Fig. 4E). HUVEC, a positive control, exhibited pronounced connections in the Matrigel plug, while no connections were seen when Matrigel plug containing only EGM-2 was injected (Supporting Information Figs. S6C–S6F).
qPCR data of in vitro endothelial differentiation showed a time-dependent upregulation of vWF, VEGFa, and desmin (p < .05), a marginal upregulation of PECAM-1 (p = .055), and no significant change in the expression of Acta2 (Fig. 4F). Notch2 and WT-1 genes were also time-dependently regulated. Podocyte markers were not expressed.
After growing c-kit+/Lin− cells in EGM-2 for 1 week, they began to express angiotensin II (Ang II) type 1a (AT1a) receptor and its expression increased significantly with time (Fig. 4G). In contrast, Ang II type 2 receptor was not detected after differentiation (Supporting Information Table S1). Calcium (Ca2+) gradient analysis demonstrated higher intracellular Ca2+ concentration in differentiated cells at baseline and their response to extracellular Ca2+ was more pronounced compared to undifferentiated cells (Fig. 4H). Therefore, responsiveness to Ang II was assessed. Differentiated cells exhibited higher depolarization following Ang II administration (100 nM), a response that was selectively blocked by Losartan, but not by PD123319 (Fig. 4I), confirming the involvement of the Ang II type 1a and not Ang II type 2 receptor. Antagonism of inositol-1,4,5-triphosphate receptor by 2-aminoethoxydiphenylborane (2-APB; 60 μM) decreased Ca2+ dependent influx from the sarcoplasmic reticulum after Ang II administration  (Fig. 4I). The dose-response to Ang II is described in Supporting Information Figure S7A. The response to endothelin (ET) via ETA and ETB receptors and to prostaglandin F-2α (PGF2α) was more intense in differentiated compared to undifferentiated cells, a response that was attenuated by 2-APB, as well as by the specific antagonists (BQ-123, BQ-788, and SQ 29548, respectively) (Supporting Information Fig. S7B–S7J). Notably, extracellular Ca2+ absence attenuated endothelin response in both differentiated and undifferentiated cells (Supporting Information Fig. S7D, S7E). However, that effect was more pronounced in the undifferentiated cells. The response to bradykinin was more intense in undifferentiated cells and was attenuated by 2-APB (Supporting Information Fig. S7K, S7L). That response was specifically mediated by B2 receptor, since HOE 140 decreased the Ca2+ influx (Supporting Information Fig. S7M–S7O). ETA, PGF2α, B2 bradykinin receptors were upregulated with time in endothelial medium (Supporting Information Fig. S7P). Together these results support the differences in the intracellular Ca2+ and in the responses to extracellular Ca2+ and to vasoactive agents between undifferentiated and differentiated cells.
C-kit+/Lin− Cells Are Clonogenic and C-kit-Derived Clones Exhibit the Capacity for Multipotent Differentiation
To further substantiate the stemness of these cells, we documented clonogenicity. We obtained c-kit+/Lin− single cells by carrying out two serial dilutions in 96-well plates (Fig. 5A). After obtaining one cell per well, we picked three faster proliferating clones and expanded them under nonadherent conditions. All three clones exhibited c-kit epitope by immunofluorescence (Supporting Information Fig. S8).
After growing the c-kit+-derived clones in differentiation media (Fig. 5B), they all exhibited plasticity. In adipogenic medium, they accumulated lipid droplets that stained for Oil Red O, and exhibited upregulation of PPAR-γ and adiponectin (Fig. 5C). In osteogenic medium, Runx2, osteopontin, and AP were upregulated, and cultures stained for Alizarin Red S (Fig. 5D). In epithelial medium, embryoid body-like morphology was observed, and the genes involved in epithelial commitment were upregulated (Fig. 5E). Colocalization of pan-cytokeratin and E-cadherin was also observed in the epithelial spheres (Fig. 5E).
C-kit+ Cells Form Nephrospheres when Grown in Nonadherent Conditions
Sphere-forming assays have been used, both retrospectively and prospectively, to investigate stem cells and progenitors in many tissues during development and in the adult . A central tenet of sphere-forming assays is that each sphere is derived from a single cell and is therefore clonal.
Accordingly, nonclonal (nonsingle cell derived) and clonal (single cell derived) c-kit+ cells, previously grown in adherent conditions, were dissociated into single cells and grown at clonal density (1 × 103 cells per well), in six-well plates, in nonadherent conditions (Fig. 6A). Primary spheres were formed by proliferation instead of aggregation and were visible after 4 days (Fig. 6B) at a frequency of approximately 2.5% of the initially plated cells (Fig. 6C). The majority of the spheres were small, measuring 40–100 μm (Fig. 6C). The spheres were passaged a minimum of three times, demonstrating self-renewal capacity. Hence, c-kit+ cells clearly are clonogenic supporting their identity as a true stem cell. However, the higher proliferation rate was observed when plating cells in adherent conditions, likely reflecting the importance of cell–cell interaction and cell adhesion for c-kit+/Lin− cell growth.
C-kit-derived spheres exhibited markers from both neuroectoderm and mesoderm progeny (Fig. 6D), including nestin, β-3 tubulin, Acta2, isolectin, pan-cytokeratin, E-cadherin, and markers found in the kidney (NKCC2, NCCT, and AQP-1). Colocalization of c-kit receptor and NKCC2 was observed in the spheres (Fig. 6E). Upregulation of those genes was observed in primary and secondary spheres (Fig. 6F).
Regenerative Capacity of C-kit+/Lin- Cells After Acute IRI
As a final test of the regeneration capacity of kidney neonatal c-kit+ stem cells, we assessed their ability for in vivo tissue repair. To evaluate the potential of c-kit+/Lin− cells to improve renal function in vivo, we used the model of acute IRI . This model, which mainly affects proximal tubular function, also affects the glomeruli involving podocyte foot process effacement .
We injected c-kit+/Lin− cells (n = 8), MSCs (n = 6), or saline (n = 12) following IRI into the aorta immediately upstream of the renal arteries, while gently clamping the aorta below the kidneys. Animals were followed for 8 days. C-kit+/Lin− cells promoted renal functional recovery as demonstrated by improvement of creatinine and BUN at day 4. (Fig. 7A). MSCs improved renal function at day 2. C-kit+/Lin− cells and MSC-treated animals exhibited not only a higher proliferation of surviving epithelial tubular cells in comparison to the control (Fig. 7B) but also a less severe kidney injury score (Fig. 7C), as indicated by less tubular damage compared to the control group (Fig. 7D).
Immunofluorescence staining with an anti-GFP antibody and staining for E-cadherin indicated that c-kit+/Lin− cells were integrated into tubules in all eight animals studied (Fig. 7E). C-kit+/Lin− cells also engrafted into glomeruli and vessels in three of eight animals (Fig. 7E). Most of GFP-labeled c-kit+ cells engrafted into glomeruli were found in Bowman's capsule, while a few of them were also seen in podocytes, as demonstrated by the colocalization with WT-1 (Fig. 7E). On day 8 after IRI, the number of GFP-positive c-kit cells expressing E-cadherin was 11.5% ± 1.1% of all tubular cells counted in 20× fields with 2× zoom (∼1,000 epithelial tubular cells counted). For MSCs, that number was significantly lower: 7.7% ± 1.5% (∼800 epithelial tubular cells counted; p = .044) (Fig. 7F). No engraftment into vessels and glomeruli in MSC-treated animals was observed. There were also GFP+-cells observed within the lumen of the tubules, indicating that some cells may have been eliminated in the urine anti-GFP antibody in saline sections was used as a control (Fig. 7G).
Here, we demonstrate that c-kit+ cells found in neonatal rat kidneys exhibit the fundamental properties of stem cells, including clonogenicity, self-renewal, multipotential capacity for commitment to mesoderm and neuroectoderm progeny, and contribute to kidney repair.
We detected c-kit+ cells in the TAL of Henle's loop, a MM-derived structure. MM-derived cells also express epithelial, mesenchymal, endothelial, neuronal, and renal differentiated cell markers [32, 38–40], as did our neonatal c-kit+ cells. Notably, immature tubules can also express vimentin and epithelial markers, such as ZO-1 . More recently, human inducible pluripotent stem cell-derived OSR1+ cells differentiated into intermediate mesoderm, the precursor structure of the MM and ureteric bud, and exhibited not only tubular markers but also glomerular and vascular markers .
During neonatal rat kidney development, intense proliferation is seen in S-shaped bodies, immature tubules, and undifferentiated cells . Several genes are upregulated during that period, including the c-kit receptor in the MM and ureteric bud . Exogenous SCF expands c-kit+ populations from both renal interstitium and hemangioblasts and can accelerate kidney development . Additionally, studies on transgenic mice confirmed c-kit expression in hemangioblasts, MM, and also in the epithelial cells of the distal tubules, collecting ducts, ureter, and bladder . However, further studies on in vivo lineage tracing of endogenous c-kit+ population would bring additional information whether c-kit+ cells invade the condensing MM from outside, are a mesenchymal population of cells distinct from the MM cells that will became renal epithelia, or derive from some distinct MM progenitor cell [43, 45–47].
The TAL segments of the nephron reside in areas of low oxygen tension, suggesting that these segments may represent an hypoxic niche of stem cells, as described in other organs and tissues . Recently, the adult progenitor/stem cell marker Lgr5 was also identified in S-shaped body segments dedicated to generating the TAL . However, Lgr5 positive cells exhibited a different gene profile from the c-kit+ population and their expression was restricted to cell clusters within developing nephrons in the cortex until postnatal day 7, as opposed to c-kit+ cells that were detected in adult TAL segments.
Moreover, early stem cells and reprogramming genes Oct4, Sox2, Klf4, and c-myc are detected in developing kidneys according to the GUDMAP database . This is interesting in light of studies showing that inducible pluripotent stem cells were obtained from proximal tubular cells with only two transcription factors (Oct4 and Sox2)  and four transcription factors (Oct4, Sox2, Klf4, and c-myc)  from mesangial cells, suggesting that epigenetic memory might also exist in the kidney. Yet, Oct4 is dispensable for both self-renewal and maintenance of somatic stem cells in the adult mammal . Furthermore, Klf4 regulates kidney epithelial tubular differentiation , while c-myc promotes proliferation of renal progenitors .
Sphere-forming assays have been tested in different adult murine and human tissues, including anterior pituitary, prostate, dermis, pancreas, cornea, retina, breast and heart, and are a useful tool to test the potential of cells to exhibit stem cells traits, although not considered a read-out of in vivo stem cell activity . Here, we showed that c-kit-derived nephrospheres exhibited markers of neuroectoderm and mesoderm progeny. It is noteworthy that common sphere features include the presence of stem cells, progenitors, and differentiated cells, the expression of nestin, routinely used for detection of neural stem cells but also characteristic for progenitor epithelial cells, and the ability of the sphere-derived cells to differentiate into other cell types in addition to their own tissue-specific cell type. More recently, E-cadherin and KRT18 were described as early differentiation markers in embryonic stem cells , although contrasting data showed E-cadherin involvement in somatic cell reprogramming .
In this study, c-kit-mediated kidney regeneration involved multicompartment engraftment. Importantly, our study did not rule out a paracrine effect  or the intrinsic mechanism of repair due to the proliferating capacity of surviving tubular epithelial cells [15, 59]. Further lineage tracing studies could evaluate the involvement of c-kit+ cells in that mechanism. Engraftment of c-kit+ cells into Bowman's capsule and podocytes suggest that these cells may also play a role in repopulating kidney stem cell niches, as the one described in Bowman's capsule [7, 8]. Furthermore, c-kit+ cells from different organs, including the biliary , bronchiolar , and renal epithelia  exhibit stem cell characteristics and epithelial differentiation. Cardiac c-kit+ cells have also been therapeutically used after myocardial infarction [61, 62]. Moreover, c-kit+ cells can promote kidney regeneration by an autocrine mechanism, as demonstrated by the shift of these cells from the papilla and medullary rays to the corticomedullary area following acute IRI . However, c-kit+ cells described here are distinct from the kidney c-kit+ side population, because the latter exhibited variable differentiation potential and failed to integrate into tubules [64, 65].
Here, we identified a neonatal kidney-derived c-kit+ cell population that fulfills all of the criteria as a stem cell. These cells were found in the thick ascending limb of Henle's loop and exhibited clonogenicity, self-renewal, and multipotentiality with differentiation capacity into mesoderm and ectoderm progeny. Ex vivo expanded c-kit+ cells not only exhibited a paracrine effect but also integrated into several compartments of the kidney, including tubules, vessels, and glomeruli and contributed to functional and morphological improvement of the kidney following acute IRI in rats. Taken together, kidney-derived c-kit+ cells have important biological and therapeutic implications in regenerative medicine.
This work was supported by a postdoctoral research fellowship grant (1KD07-33958) from the James and Esther King Florida Biomedical Research Program to E.B.R., and National Institutes of Health RO1 Grants HL107110 and AG025017 to J.M.H. Special thanks to Carmen Perez for preparing the histologic sections, Irene Margitich for technical support, Wayne Balkan for carefully reading the paper, Shannon Opiela and Jay Enten for the assistance at FACS facility, and Qinghua Hu for providing the GFP-MSCs cells.
Disclosure of Potential Conflicts of Interest
The authors indicate no potential conflicts of interest.