ABSTRACT
Complete and robust human genome duplication requires loading MCM helicase complexes at many DNA replication origins, an essential process termed origin licensing. Licensing is restricted to G1 phase of the cell cycle, but G1 length varies widely among cell types. Using quantitative single cell analyses we found that pluripotent stem cells with naturally short G1 phases load MCM much faster than their isogenic differentiated counterparts with long G1 phases. During the earliest stages of differentiation towards all lineages, MCM loading slows concurrently with G1 lengthening, revealing developmental control of MCM loading. In contrast, ectopic Cyclin E overproduction uncouples short G1 from fast MCM loading. Rapid licensing in stem cells is caused by accumulation of the MCM loading protein, Cdt1. Prematurely slowing MCM loading in pluripotent cells not only lengthens G1 but also accelerates differentiation. Thus, rapid origin licensing is an intrinsic characteristic of stem cells that contributes to pluripotency maintenance.
INTRODUCTION
Metazoan DNA replication requires initiation at thousands of DNA replication origins during S phase of every cell cycle. Origins are genomic loci where DNA helicases first unwind DNA and DNA synthesis begins. Origins are made competent for replication during G1 phase of each cell cycle by the loading of Minichromosome Maintenance (MCM) complexes onto DNA. MCM is the core component of the replicative helicase, and the process of MCM loading is termed origin licensing. Total MCM levels remain constant throughout the cell cycle, but the levels of DNA-loaded MCM change as cells progress through the cell cycle. Cells can begin MCM loading as early as telophase, and loading continues throughout G1 until the G1/S transition, the point of maximum DNA-loaded MCM1,2. Throughout S phase, individual MCM complexes are activated for DNA unwinding as origins “fire”. MCM complexes travel with replication forks and are progressively unloaded as replication forks terminate (Fig. 1a)3–5.
The control of origin licensing is critical for genome stability. Origins must not be re-licensed after S phase begins because such re-licensing can cause a genotoxic phenomenon known as re-replication which may result in double strand breaks, gene amplification, aneuploidy, and general genome instability6,7. To avoid re-replication, MCM loading is tightly restricted to G1 phase by multiple overlapping mechanisms that destroy or inactivate MCM loading proteins to prevent any new origin licensing after S phase begins4,6,7. On the other hand, cells typically load 5-10-fold more MCM complexes in G1 than they strictly require under ideal circumstances, and the additional MCM loading ensures timely and complete genome duplication even if replication hurdles are encountered in S phase8–10. It is possible for cells to proliferate with less than optimal MCM loading, but such cells are hypersensitive to DNA damage and replication stress11,12.
MCM loading to license origins is restricted to G1, but G1 length varies widely among different cell types. For example, specialized developmental and immune cell cycles have minimal G1 lengths of mere minutes13–15. In cultured human embryonic stem cells, G1 is only 2-3 hours, and this short G1 is both a hallmark of and has been implicated in maintaining pluripotency16,17. G1 lengthens early in differentiation, and in cultured somatic cells is often greater than 12 hours18. Thus, different proliferating cells have drastically different amounts of time available to complete MCM loading before making the G1-to-S phase transition. In addition, pluripotent stem cells respond to differentiation stimuli specifically in G1 phase, suggesting that the balance among cell cycle phases influences differentiation potential19,20.
Given that MCM loading is restricted to G1 and the wide variation of G1 lengths, we postulated that the absolute amount of loaded MCM in S phase is a product of both the time spent in G1 and the rate of MCM loading. The combination of these two parameters has implications for genome stability because loading more or less MCM in G1 influences S phase length and how effectively S phase cells can accommodate both endogenous and exogenous sources of replication stress21,22. These implications are relevant both when cell cycle distributions change during development and during oncogenesis since many cancer cell lines also have short G1 phases. The actual rate of MCM loading in human cells has not yet been quantified however, and little is known about how the amount, rate or timing of MCM loading varies between cells with different G1 lengths. Here we utilized single cell flow cytometry to measure MCM loading rates in asynchronous populations of pluripotent and differentiated cells. We discovered that rapid MCM loading is intrinsic to pluripotency, slows universally during differentiation, and rapid replication licensing suppresses differentiation. These findings demonstrate that the rate of MCM loading is subject to developmental regulation, and we suggest that rapid origin licensing is a new hallmark of pluripotency.
RESULTS
Pluripotent cells load MCM significantly faster than differentiated cells
We considered two possibilities for how cells with varying G1 lengths load MCM onto DNA. One possibility is that cells with short G1 phases load MCM at the same rate as cells with long G1 phases resulting in less total loaded MCM. Alternatively, cells with short G1 phases could load MCM faster than cells with long G1 phases and reach similar levels of loaded MCM. To distinguish between these scenarios, we developed an assay to measure DNA-loaded MCM in individual cells of asynchronously proliferating populations by adapting a previously-reported flow cytometry method23,24. We extracted EdU-labelled immortalized epithelial cells (RPE1-hTERT) with nonionic detergent to remove soluble MCM. We then fixed the remaining chromatin-bound proteins for immunofluorescence with anti-MCM2 antibody as a marker of the MCM2-7 complex (Fig. 1b, left) or without primary antibody as a control (relevant flow cytometry gating schemes are shown in Supplementary Fig. 1). Interestingly, individual G1 cells have a very broad range of DNA-loaded MCM levels with a more than 100-fold difference between minimum and maximum (Fig. 1b, left). MCMs are unloaded during S phase, ending in G2/M with undetectable MCM on DNA (Fig. 1b, left). As expected, total MCM protein levels do not substantially change in the cell cycle (Fig. 1b, right). For comparison to commonly used cell fractionation methods to assess MCM dynamics, we also probed immunoblots of chromatin-enriched fractions, and noted a similar MCM expression, G1 loading, and S phase unloading pattern (Fig. 1c)25,26.
Loaded MCM complexes are extremely stable on DNA, both in vivo and in vitro27–30. In human cells, MCMs can persist on DNA for more than 24 hours during a cell cycle arrest and are typically only unloaded during S phase31. These properties result in MCM loading that occurs unidirectionally throughout G1 phase32. The unidirectional nature of MCM loading means that G1 cells with low MCM levels are in early G1, and G1 cells with high MCM levels are in late G1. Since we observed a broad distribution of MCM loading throughout G1 including many cells with low levels of loaded MCM, we conclude that RPE1-hTERT cells load MCM relatively slowly during their ∼9 hr G1.
We then used this method to analyze MCM loading in asynchronous cells with different G1 lengths. H9 human embryonic stem cells (hESCs) have a short G1 phase and spend most of the cell cycle in S phase. In contrast to the differentiated epithelial cells, the majority (∼80%) of G1 hESCs had high levels of loaded MCM; very few G1 cells had low levels of loaded MCM (blue cells, Fig. 1d). This difference suggests that hESCs load MCM rapidly to achieve abundant DNA-loaded MCM in a short time. To test if MCM loading varies in differentiated cells, we differentiated hESCs into neural progenitor cells (NPCs) to generate an isogenic pair of pluripotent and differentiated cells. In contrast to hESCs, differentiated NPCs had a wide distribution of DNA-loaded MCM in G1 (blue cells, Fig. 1d); they also spend approximately five time longer in G1 (Fig. 1e). Since the NPCs had many cells with low levels of DNA-loaded MCM, we conclude that these differentiated cells load MCM more slowly than hESCs.
To generate another isogenic pair of pluripotent and differentiated cells we reprogramed ARPE-19 primary retinal pigmented epithelial cells (RPE) into induced pluripotent stem cells (iPSCs). The iPSCs had hallmark features of pluripotency as measured by microscopy, bisulfite sequencing, gene expression, and teratoma formation (Supplementary Fig. 2a-e), and their G1 phases were typically seven times shorter than their differentiated parents (Fig. 1e). Like hESCs, the pluripotent iPSCs had predominantly high levels of DNA-loaded MCM in G1 (Fig. 1d). Importantly, both of the pluripotent cell lines reached approximately equal levels of DNA-loaded MCM at the start of S phase as their differentiated counterparts did, but in in less time (the absolute MCM loading intensities are comparable when samples are processed and analyzed with identical instrument settings). Taken together, these data demonstrate that pluripotent cells load MCMs rapidly in G1, but differentiated cells load MCMs slowly.
We then quantified the relative MCM loading rates in pluripotent and differentiated cells using ergodic rate analysis, a mathematical method that can derive rates from fixed, steady state populations33. Ergodic analysis can measure any unidirectional rate parameter from a steady state distribution, and is not limited to the cell cycle (e.g. car traffic jams)34. The ergodic analysis as applied to the cell cycle means that within a steady state population with a constant doubling time and cell cycle distribution, the number of cells at any point in the cell cycle is inversely related to the rate they move through that point. For any measured parameter, the density of cells indicates rate: low cell density on a flow cytometry plot indicates a fast rate passing through that cell cycle state whereas high cell density indicates a slow rate. This phenomenon is analogous to a high density of slow-moving cars observed at a given point on a road in a traffic jam compared to a low density of fast-moving cars on an open highway. We visualized MCM loading as histograms of the MCMDNA intensities in only the G1 cells for ergodic rate analysis (G1-MCMDNA, Fig. 2a, b and Supplementary Fig. 3).
To compute MCM loading rate per hour, we experimentally determined the cell cycle distributions and doubling times of each cell population (Supplementary Fig. 3). Pluripotent cells reached near equal levels of loaded MCM at the G1/S transition in less time than differentiated cells. To quantify the actual MCM loading rate difference, we subdivided the G1-MCMDNA population into 10 equally-sized bins, calculated the MCM loading rate for each bin, then the overall average MCM loading rate for each G1 population. These calculations revealed that pluripotent hESCs loaded MCM 6.5 times faster than differentiated NPCs and pluripotent iPSCs loaded MCM 3.9 times faster than differentiated RPEs (Fig. 2c). Thus, pluripotent cells with short G1 phases load MCMs significantly faster than differentiated cells with long G1 phases.
Differentiation, G1 length, and MCM loading rate are coupled
We hypothesized that MCM loading is fundamentally linked to pluripotency because MCM loading rate decreased during differentiation and increased during reprogramming. This idea predicts that slowed MCM loading is a phenomenon common to differentiation towards all germ layers. To test that hypothesis, we initiated differentiation in hESCs towards the three embryonic germ layers (neuroectoderm, mesoderm and endoderm) and extraembryonic trophectoderm, collecting cells at 24 hour and 48 hours after inducing differentiation (Supplementary Fig. 4b). We confirmed progress towards each lineage by the expected gene expression changes, particularly induction of lineage-specific markers and modest reduction of pluripotency markers – even at these very early time points (Fig. 3c). We assessed MCM loading rates during differentiation by flow cytometry as before. The MCM loading rate clearly decreased for all germ layers rapidly within the first 48 hours of initiating differentiation (Fig. 3a, compare the grey histograms for undifferentiated G1 cells to the green and blue histograms). The decrease in MCM loading rate also coincided with the increase in the proportion of G1 cells for each lineage. For example, within 24 hours of neuroectoderm differentiation, G1 had already lengthened and MCM loading had slowed, but during trophectoderm differentiation both G1 lengthening and slowed MCM loading took 48 hrs (Fig. 3a and b). The closely-coordinated changes that we universally observed during differentiation suggest that MCM loading rate is coupled to G1 length. Importantly, these results demonstrate that the rate of origin licensing by MCM loading is developmentally regulated.
We next asked if G1 length and MCM loading rate are obligatorily coupled, or if the link can be short-circuited by artificially advancing the G1/S transition. To distinguish between these possibilities, we constructed an RPE1-hTERT derivative with a CYCLIN E1 cDNA under doxycycline-inducible control. Cyclin E1 overproduction reproducibly shortened G1 length, consistent with previous studies (Fig. 4a, b)35,36. Strikingly, cells overproducing Cyclin E1 (designated as “↑Cyclin E1”) not only spent less time in G1 but also began S phase with much lower amounts of loaded MCM compared to the control; this new subpopulation appeared in the central triangular region of the plots that is typically clear of S phase cells (Fig. 4c, orange EdU+ MCMDNA+). We conclude that the MCM loading rate did not increase to accommodate the shorter G1 because the ↑Cyclin E1 cells had on average at least two-fold less DNA-loaded MCM in early S phase than control cells (Fig. 4d). Thus, shortening G1 length without increasing MCM loading rate causes G1 cells to enter S phase without the full complement of DNA-loaded MCM.
Fast loading hESCs have more Cdt1 in G1
Loading MCM complexes onto DNA requires the six subunit Origin Recognition Complex (ORC), Cdc6, and Cdt1. We hypothesized that fast MCM loading in pluripotent stem cells is achieved by increased levels of the loading proteins. To test this idea, we probed protein lysates of asynchronous cells to compare the amount of MCM loading proteins between isogenic cell lines. Total Mcm2 and ORC protein levels remained constant (Fig. 5a). The other MCM loading factors normally change in their abundance during the cell cycle due to regulated proteolysis. Cdc6 protein levels are low in G1 and high in S phase (Fig. 5b). Conversely, Cdt1 protein levels are high in G1 and low in S phase (Fig. 5b)37,38. Since an asynchronous population of pluripotent cells spends significantly more time in S phase than differentiated cells do, we expected Cdc6 levels to be higher in asynchronous pluripotent cells compared to their isogenic counterparts. Cdc6 was indeed higher in pluripotent cells, as was Geminin, a protein regulated in a similar manner as Cdc6 (Fig. 5a)39. To our surprise, even though the majority of asynchronous pluripotent cells were in S phase, a time when Cdt1 is degraded, Cdt1 levels were higher in asynchronous pluripotent cells than in isogenic differentiated cells (Fig. 5a). A similar observation was reported for mouse embryonic stem cells40. These data imply that Cdt1 levels are higher in G1 phase of pluripotent cells than G1 of differentiated cells, providing a potential explanation for fast MCM loading in pluripotent cells.
To directly measure Cdt1 levels specifically in G1, we collected asynchronous hESCs and NPCs then fixed and stained them for Cdt1, EdU and DAPI. S phase Cdt1 degradation in hESCs is similar to differentiated cells with very low levels in S phase (purple, Fig. 5c). In contrast, the hESCs had a large population of cells with high Cdt1 levels in G1 (green cells) and significant amounts of Cdt1 in G2/M phase (grey cells), whereas the NPCs had a broad and overall lower Cdt1 distribution in G1 and very little Cdt1 in G2/M (Fig. 5d). The G1 hESCs consistently harbored significantly more Cdt1 than G1 NPCs did (2.9 fold higher median, 2.2 fold higher mean, 3 replicates (Fig. 5d)). We note that CDT1 mRNA is modestly but consistently higher in asynchronous hESCs compared to differentiated derivatives, and that Cdt1 protein levels decrease during early differentiation coincident with the slowdown in licensing rate, but before loss of Oct4 (Fig. 3c and Supplementary Figure 4a). We postulate that the higher amount of this essential MCM loading protein specifically in G1 contributes to the fast MCM loading rate in hESCs.
Rapid MCM loading protects hESC pluripotency
Cdt1 is essential for MCM loading, therefore reducing Cdt1 levels is expected to slow MCM loading. If MCM loading rate is linked to G1 length, then slowing MCM loading by reducing Cdt1 levels should lengthen G1. To test this prediction, we used siRNA to reduce Cdt1 in hESCs and measured changes in both MCM loading rate and G1 length (Fig. 6a, b). As expected, Cdt1 depletion reduced MCM loading rate in hESCs (Fig. 6c). Strikingly, G1 length increased coincidentally with the decrease in MCM loading rate (Fig. 6d). We obtained a complementary result by depleting the orthogonal MCM loading protein, Cdc6 (Supplementary Fig. 5a-d). A more modest Cdc6 knockdown correlated with a weaker, but detectable effect on MCM loading. Taken together, these data corroborate the close link between MCM loading rate and G1 length in hESCs.
Our demonstration that slower MCM loading occurs universally during early differentiation suggested a functional link between the rate of MCM loading and pluripotency maintenance. We considered that slowing MCM loading might promote differentiation. To explore this idea, we prematurely slowed MCM loading in hESCs by Cdt1 depletion prior to inducing their differentiation (Fig. 7e). After Cdt1 depletion, we stimulated differentiation towards trophectoderm with BMP441. After 48 hours, we quantified Oct4 and Cdx2 by immunostaining. (Fig. 7a). The pluripotency transcription factor Oct4 and the trophectoderm transcription factor Cdx2 reciprocally repress one another’s expression, creating a clear distinction between Oct4 positive-Cdx2 negative pluripotent cells and Oct4 negative-Cdx2 positive differentiating cells42. We quantified the mean fluorescence intensity of both Oct4 and Cdx2 in >18,000 cells per condition with a customized, automated CellProfiler pipeline, plotting the signal intensities for each cell in a density scatter plot (Fig. 7b, c). Stimulating control hESCs with 10 ng/mL of BMP4 slightly shifted the population towards differentiation, but most cells remained pluripotent with high Oct4 levels at this time point. Strikingly, hESCs pretreated with Cdt1 siRNA to prematurely slow MCM loading gained a substantial population of Oct4 negative-Cdx2 positive differentiating cells relative to controls that were treated similarly. To quantify the extent of differentiation, we divided the Cdx2 intensity of each cell by its Oct4 intensity, creating a single differentiation score (Fig. 7d). After 10 ng/ml BMP4 treatment, Cdt1-depleted hESCs had significantly higher scores, indicating that prematurely slowing MCM loading promoted differentiation (p<0.0001, two-tailed Mann-Whitney test). Both control cells and Cdt1-depleted cells differentiated more fully at a higher concentration of 50 ng/mL BMP4, but the Cdt1-depleted cells still differentiated further than the controls (p<0.0001, two-tailed Mann-Whitney test, Fig. 7b and data not shown). Other combinations of BMP4 concentrations or treatment times also resulted in a consistent, significant increase in differentiation in cells pretreated to slow MCM loading (p<0.0001, two-tailed Mann-Whitney test, data not shown). Importantly, the phenotype was conserved across multiple differentiation lineages, as prematurely slowing MCM loading prior to endoderm differentiation also increased the number of cells positive for the endoderm transcription factor Sox17 relative to controls at the same time point (Supplementary Fig. 6b). Additionally, slowing MCM loading by depleting a different MCM loading factor, Cdc6, also significantly promoted differentiation (p<0.0001, two-tailed Mann-Whitney test), and the more modest reduction in MCM loading (i.e. Supplementary Fig. 5b) resulted in a correspondingly weaker differentiation acceleration (Fig. S7a-e). Thus, we conclude that slow MCM loading generally promoted differentiation and by extension, that rapid MCM loading preserves pluripotency.
DISCUSSION
In this study we demonstrate that rapid MCM loading to license replication origins is an intrinsic property of pluripotent cells. Human embryonic stem cells have a remarkably fast MCM loading rate, and reprogramming to create induced pluripotent stem cells increases MCM loading rate. Moreover, MCM loading slows concurrently with the G1 lengthening and extensive cell cycle remodeling that accompany the early stages of differentiation (Fig. 7f). To our knowledge this is the first demonstration that the rate of MCM loading is developmentally regulated. The regulated decrease in MCM loading rate is critical during differentiation, as rapid MCM loading protects pluripotency, and prematurely slowing MCM loading promotes differentiation.
Pluripotent stem cells load MCM complexes rapidly to reach similar total amounts of loaded MCM at the G1/S transition in less time than their isogenic differentiated counterparts. Although we did not detect substantial MCM loading in telophase, as suggested previously43, it is clear that telophase loading is an option in some cells and a requirement in cells with no detectable G1 such as S. pombe and in the first nuclear divisions in D. melanogaster44,45. Stem cells achieve faster MCM loading, at least in part, by particularly high Cdt1 levels in G1. These high levels are achieved not only by a modest difference in CDT1 mRNA (Fig. 3c) but also post-transcriptionally by specific re-accumulation of Cdt1 protein in the preceding G2 phase (Fig. 5c). Cdt1 stability in G2 phase has been attributed to geminin-mediated protection from the SCFSkp2 E3 ubiquitin ligase in non-stem cells46–48. It is not clear however that geminin levels are particularly high in stem cells relative to differentiated cells (aside from differences expected from cell cycle distribution changes (Fig. 5a)40), so it seems unlikely that geminin drives higher Cdt1 levels in stem cells. Skp2 levels also do not change in the earliest stages of stem cell differentiation49. Cdt1 is protected in late S phase and G2 by cyclin A/Cdk1 activity50, and we thus consider it likely that the documented high CDK activity in stem cells contributes to Cdt1 stabilization in G251,52. The anticipatory Cdt1 accumulation to promote MCM loading in G1 was originally proposed from experiments in cancer-derived cell lines. Our observations in stem cells suggest this strategy is employed by non-transformed cells during developmental stages that require short G1.
Other factors besides Cdt1 accumulation may also accelerate MCM loading. The hESCs we assayed have 2-3 fold greater Cdt1 protein levels in G1 relative to NPCs yet load MCM 6.5 times faster per hour than NPCs. One Cdt1 molecule can (in vitro) load multiple MCM complexes since Cdt1 is released into the soluble phase immediately after completing a loading reaction53. Stem cells may experience less Cdc6 degradation in early G1 due to nearly constitutive Cyclin E/Cdk2 activity and/or attenuated APCCdh1 activity40,52,54. Additionally, stem cells are enriched for euchromatin, an environment that may be particularly permissive for rapid MCM loading55.
Rapid MCM loading may itself contribute to mechanisms that maintain short G1 phases in pluripotent cells. The origin licensing checkpoint links the amount of loaded MCM to G1 length by controlling Cdk2 activity. In that regard, overproducing Cyclin E “short-circuited” the licensing checkpoint in slow loading differentiated cells. This checkpoint has thus far only been demonstrated in p53-proficient differentiated mammalian cells56, but the G1 lengthening of hESCs after Cdt1 depletion suggests that pluripotent stem cells also have a functioning licensing checkpoint. Cells with fast MCM loading could satisfy this checkpoint quickly, activate Cyclin E/Cdk2, and thus spend less time in G1. Mechanisms that support short G1 length preserve pluripotency in hESCs (and promote reprogramming to iPSCs) since cells are most sensitive to differentiation cues in G1; in that regard, extending G1 phase in hESCs can increase differentiation propensity16,20,54,57. Recent work with quintuple knockout mice lacking all D and E type cyclins also reported that Cyclin E/Cdk2 further contributes to maintaining pluripotency by stabilizing the Oct4, Sox2, and Nanog transcription factors58. Fast MCM loading may have evolved as an intrinsic property of pluripotent cells to maintain high Cdk2 activity and keep G1 phase short.
Cyclin/Cdk activity is not the sole connection between the cell cycle and pluripotency. Non-CDK cell cycle-associated proteins regulate expression of key pluripotency genes including SOX2 and NANOG19,59,60. Pluripotency transcription factors themselves regulate expression of cell cycle genes including those encoding cyclins, CDK inhibitors, and E2F3a61–63. On the other hand, pluripotency and cell cycle functions can be genetically uncoupled in experiments where manipulating the cell cycle didn’t alter pluripotency and vice versa64,65. We observe that licensing inhibition can accelerate differentiation even without greatly lengthening G1 (Supplementary Figs. 5, 7) which may point to an additional direct and cell cycle-independent link between MCM loading rate and differentiation.
We note that early differentiation is not the only setting in which rapid MCM loading during a short G1 may be relevant. Like hESCs, activated T cells have very fast cell cycles with short G1 phases14. Oncogenic transformation is also frequently associated with G1 shortening. It may be that the pathways linking differentiation to MCM loading rate are also coopted in some cancers to induce rapid licensing. On the other hand, a subset of cancers may proliferate in a perpetually underlicensed state that contributes to the genome instability characteristic of transformed cells. Future investigations will elucidate the molecular relationships among developmental signaling pathways, MCM loading rate, and cell cycle remodeling.
MATERIALS AND METHODS
Cell Culture
T98G, HEK293T, and RPE1-hTERT were cultured in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 2 mM L-glutamine and 10% fetal bovine serum (FBS) and incubated in 5% CO2 at 37 °C. ARPE-19 (male) were cultured in 1:1 DMEM:F12 supplemented with 2 mM L-glutamine and 10% fetal bovine serum and incubated in 5% CO2 at 37 °C. T98G, HEK293T, RPE1-hTERT, and ARPE-19 cells were from the ATCC and were passaged with trypsin and not allowed to reach confluency. WA09 (H9 hESCs) were cultured in mTeSR1 (StemCell Technologies) with media changes every 24 hours on Matrigel (Corning) coated dishes and incubated in 5% CO2 at 37 °C. H9s had normal diploid karyotype at passage 32 and were used from passage 32-42. ARPE-iPSCs were cultured in Essential 8 (Life Technologies) with media changes every 24 hours on Matrigel (Corning) coated dishes and incubated in 5% CO2 at 37 °C. iPSCs were used from passage 20-25. Both hESCs and iPSCs were routinely passaged every 4 days as aggregates using ReLeSR, according to manufacturer’s instructions (StemCell Technologies). The hESCs and iPSCs were only passaged as single cells in 10 μM Y-27632 2HCl (Selleck Chemicals) for experiments, as described previously 66. NPCs were cultured in Neural Progenitor Medium (StemCell Technologies) with media changes every 24 hours on poly-L-ornithine/Laminin (Sigma Aldrich) coated dishes and incubated in 5% CO2 at 37 °C. NPCs were passaged with StemPro Accutase (Gibco) weekly.
Total lysate and chromatin fractionation
Cells were collected via trypsinization. For total protein lysates, cells were lysed on ice for 20 minutes in CSK buffer (300 mM sucrose, 100 mM NaCl, 3 mM MgCl2, 10 mM PIPES pH 7.0) with 0.5% triton x-100 and protease and phosphatase inhibitors (0.1 mM AEBSF, 1 μg/ mL pepstatin A, 1 μg/ mL leupeptin, 1 μg/ mL aprotinin, 10 μg/ ml phosvitin, 1 mM β-glycerol phosphate, 1 mM Na-orthovanadate). Cells were centrifuged at 13,000 xg at 4 °C for 5 minutes, then the supernatants were transferred to a new tube for a Bradford Assay (Biorad) using a BSA standard curve. Chromatin fractionation for immunoblotting was performed as described previously25,26, using CSK buffer with 1 mM ATP, 5 mM CaCl2, 0.5% triton x-100 and protease and phosphatase inhibitors to isolate insoluble proteins and S7 nuclease (Roche) to release DNA bound proteins. A Bradford Assay (Biorad) was performed for equal loading.
Immunoblotting
Samples were diluted with SDS loading buffer (final: 1% SDS, 2.5% 2-mercaptoethanol, 0.1% bromophenol blue, 50 mM Tris pH 6.8, 10% glycerol) and boiled. Samples were run on SDS-PAGE gels, then the proteins transferred onto polyvinylidene difluoride membranes (Thermo Fisher). Membranes were blocked at room temperature for 1 hour in either 5% milk or 5% BSA in Tris-Buffered-Saline-0.1%-tween-20 (TBST). After blocking, membranes were incubated in primary antibody overnight at 4 °C in either 1.25% milk or 5% BSA in TBST with 0.01% sodium azide. Blots were washed with TBST then incubated in HRP-conjugated secondary antibody in either 2.5% milk or 5% BSA in TBST for 1 hour, washed with TBST, and then membranes were incubated with ECL Prime (Amersham) and exposed to autoradiography film (Denville). Equal protein loading was verified by Ponceau S staining (Sigma Aldrich). Antibodies used for immunoblotting were: Mcm2, (BD Biosciences, Cat#610700), Mcm3, (Bethyl Laboratories, Cat#A300-192A), Cdt1, (Cell Signaling Technologies, Cat#8064S), Cdc6, (Santa Cruz Biotechnology, Cat#sc-9964), Oct4, (Abcam, Cat#ab19857), Cdx2, (Abcam, Cat#ab76541), Cyclin E1, (Santa Cruz Biotechnology, Cat#sc-198), Orc1, (Bethyl Laboratories, Cat#A301-892A), Orc6, (Santa Cruz Biotechnology, Cat#sc-32735), geminin, (Santa Cruz Biotechnology, Cat#sc-13015), Histone H3, (Cell Signaling Technologies, Cat#4499S), TRA-1-60, (Invitrogen, Cat#41-1000), nestin, (Abcam, Cat#ab22035).
Flow Cytometry
For EdU labeled samples, cells were incubated with 10 uM EdU (Santa Cruz Biotechnology) for 30 minutes prior to collection. For total protein flow cytometry, cells were collected with trypsin and resuspended as single cells, washed with PBS, and fixed with 4% paraformaldehyde (Electron Microscopy Sciences) in PBS for 15 minutes at room temperature, then 1% BSA-PBS was added, mixed and cells were centrifuged at 1000 xg for 7 minutes (and for all following centrifuge steps) then washed with 1% BSA-PBS and centrifuged. Fixed cells were permeabalized with 0.5% triton x-100 in 1% BSA-PBS at room temperature for 15 minutes, centrifuged, then washed once with 1% BSA, PBS and centrifuged again before labeling. For chromatin flow cytometry, cells were collected with trypsin and resuspended as single cells, washed with PBS, and then lysed on ice for 5 minutes in CSK buffer with 0.5% triton x-100 with protease and phosphatase inhibitors. Next, 1% BSA-PBS was added and mixed, then cells were centrifuged for 3 minutes at 1000 xg, then fixed in 4% paraformaldehyde in PBS for 15 minutes at room temperature. Next, 1% BSA-PBS was added, mixed and cells were centrifuged then washed again before labeling. The labeling methods for total protein samples and chromatin samples were identical. For DNA synthesis (EdU), samples were centrifuged and incubated in PBS with 1 mM CuSO4, 1 μM fluorophore-azide, and 100 mM ascorbic acid (fresh) for 30 minutes at room temperature in the dark. 1% BSA-PBS + 0.1% NP-40 was added, mixed and centrifuged. Samples were resuspended in primary antibody in 1% BSA-PBS + 0.1% NP-40 and incubated at 37 °C for 1 hour in the dark. Next, 1% BSA-PBS + 0.1% NP-40 was added, mixed and centrifuged. Samples were resuspended in secondary antibody in 1% BSA-PBS + 0.1% NP-40 and incubated at 37 °C for 1 hour in the dark. Next, 1% BSA-PBS + 0.1% NP-40 was added, mixed and centrifuged. Finally, cells were resuspended in 1% BSA-PBS + 0.1% NP-40 with 1 μg/mL DAPI (Life Technologies) and 100 μg/mL RNAse A (Sigma Aldrich) and incubated overnight at 4 °C in the dark. Samples were run on a CyAn ADP flow cytometer (Beckman Coulter) and analyzed with FCS Express 6 software (De Novo). The following antibody/fluorophore combinations were used: (1): Alexa 647-azide (Life Technologies), primary: Mcm2 (BD Biosciences, Cat#610700), secondary: Donkey anti-mouse-Alexa 488 (Jackson ImmunoResearch), DAPI. (2): Alexa 488-azide (Life Technologies), primary: Cdt1 (Abcam, Cat#610700), secondary: Donkey anti-rabbit-Alexa 647 (Jackson ImmunoResearch), DAPI. Cells were gated on FS-area vs SS-area. Singlets were gated on DAPI area vs DAPI height. The positive/negative gates for EdU and MCM were gated on a negative control sample, which was treated with neither EdU nor primary antibody, but incubated with 647-azide and the secondary antibody Donkey anti-mouse-Alex 488 and DAPI to account for background staining (Supplementary Figure 1).
Doubling Time
Doubling time was calculated by plating equal number of cells as described above and counting cell number over time using a Luna II automated cell counter (Logos Biosystems) at 24, 48, and 72 hours after plating. Three or four wells were counted as technical replicates at each timepoint. GraphPad Prism’s regression analysis was used to compute doubling time, and multiple biological replicates were averaged for a final mean doubling time. ARPE-19s were counted 4 times, hESCs and NPCs 3 times, and iPSCs 2 times.
Cell Synchronization
To synchronize cells in G1, T98G cells were grown to 100% confluency, washed with PBS, and incubated for 72 hr in 0.1% FBS, DMEM, L-glutamine. After serum-starvation, cells were re-stimulated by passaging 1:3 with trypsin to new dishes in 20% FBS, DMEM, L-glutamine, collecting cells 10 hr and 12 hr post-stimulation. To synchronize T98G cells in early S phase, cells were treated as for G1, except 1 mM Hydroxyurea (Alfa Aesar) was added to the media upon re-stimulating and cells were collected 18 hr post-stimulation. To synchronize cells in mid-late S, cells were treated as in early S, then at 18 hr post-stimulation cells were washed with PBS and released into 10% FBS, DMEM, L-Glutamine, collecting 6 hr, 8 hr post release.
Cloning
The pInducer20-Cyclin E plasmid was constructed using the Gateway cloning method (Invitrogen). The attB sites were added to Cyclin E1 cDNA by PCR using Rc/CMV cyclin E plasmid as a template and BP-cycE-F (5′ GGGGACAAGTTTGTACAAAAAAGCAGGCTACCATGAAGGAGGACGGCGGC) and BP-cycE-R primers (5′ GGGGACCACTTTGTACAAGAAAGCTGGGTTCACGCCATTTCCGGCCCGCT) 67. The PCR product was recombined with pDONR221 plasmid using BP clonase (Invitrogen) according to the manufacturer’s instructions and transformed into DH5α to create pENTR221-Cyclin E1. Then the LR reaction was performed between pInducer20 and pENTR221-Cyclin E1 using LR Clonase (Invitrogen) according to manufacturer’s instructions and transformed into DH5α to create pInducer20-Cyclin E1.
Inducible Cyclin E
To package lentivirus, pInducer20-Cyclin E1 was co-transfected with ΔNRF and VSVG plasmids into HEK293T using 50 μg/mL Polyethylenimine-Max (Aldrich Chemistry). Viral supernatant was transduced with 8 ug/mL Polybrene (Millipore) onto RPE1-hTERT cells overnight. Cells were selected with 500 ug/mL neomycin (Gibco) for 1 week. To overproduce Cyclin E1, cells were treated with 100 ng/mL doxycycline (CalBiochem) for 72 hr in 10% FBS, DMEM, L-glutamine. Control cells were the Inducer20-Cyclin E1 without doxycycline.
siRNA transfections
For siRNA treatment, Dharmafect 1 (Dharmacon) was mixed in mTeSR1 with the appropriate siRNA according to the manufacturer’s instructions, then diluted with mTeSR1 and added to cells after aspirating old media. The final siRNA concentrations were: 100 nM siControl (Luciferase), 25 or 100 nM siCdt1, or a mixture of two siCdc6 (2144 and 2534 at 50 nM each). The Cdt1 siRNA mix was incubated on cells for either 20 or 24 hours, then changed to new mTeSR1 without siRNA. The Cdc6 siRNA mix was incubated on cells for 24 hours, then changed to new mTeSR1 without siRNA for 8 hours (32 total hours). The siRNA were described previously 68,69.
Differentiation
Trophectoderm: hESCs were passaged as single cells at 7x103 /cm2 in mTeSR1 with 10 μM Y-27632 2HCl onto Matrigel-coated plates. 24 hours later, the media was changed to start differentiation with fresh mTeSR1 with 100 ng/mL BMP4 (R&D Systems), and 24 hours later the media was changed to fresh mTeSR1 with 100 ng/mL BMP4 for 48 total hours of differentiation.
Neuroectoderm: hESCs were differentiated using a monolayer-based protocol in Neural Induction Medium: hESCs were passaged as single cells at 5.2x104 /cm2 in STEMdiff™ Neural Induction Medium (StemCell Technologies) with 10 μM Y-27632 2HCl onto Matrigel coated plates, and plating started the differentiation. 24 hours later, the media was changed to fresh Neural Induction Medium for another 24 hours for 48 hours total differentiation. To derive NPCs, hESCs were differentiated in Neural Induction medium (StemCell Technologies) using the Embryoid Body Neural Induction protocol according to manufacturer’s instructions, similar to previous reports 70. Once generated, NPCs were maintained in Neural Progenitor Medium (StemCell Technologies).
Mesoderm: hESCs were passaged as single cells at 3x104/cm2 in mTeSR1 with 10 μM Y-27632 2HCl onto Matrigel coated plates. 24 hours later, the media was changed to start differentiation. Cells were washed with Advanced RPMI 1640 (Gibco), then incubated in Advanced RPMI 1640 with B27 minus insulin (Gibco), 2 mM L-glutamine, and 8 μM CHIR-99021 (Selleck Chemicals). At 24 hours after changing the media, cells were washed with Advanced RPMI 1640 then incubated in Advanced RPMI 1640 with B27 minus insulin (Gibco), 2 mM L-glutamine, without CHIR-99021 for 24 hours for a total of 48 hours of differentiation.
Endoderm: hESCs were passaged as single cells at 4x103/cm2 in mTeSR1 with 10 μM Y-27632 2HCl onto Matrigel-coated plates. The next day the media was changed to fresh mTeSR1 without Y-27632 2HCl. 24 hours later, the media was changed to start differentiation. The cells were washed with Advanced RPMI 1640 (Gibco), then incubated in Advanced RPMI 1640 with 0.2% FBS, 2 mM L-glutamine, 100 ng/mL Activin A (R&D Systems) and 2.5 μM CHIR-99021. At 24 hours after changing the media, cells were washed with Advanced RPMI 1640 then incubated in Advanced RPMI 1640 with 0.2% FBS, 2 mM L-glutamine, 100 ng/mL Activin A, without CHIR-99021 for 24 hours for a total of 48 hours of differentiation.
Phase contrast microscopy
Phase contrast images were acquired with an Axiovert 40 CFL inverted microscope, 20x objective.
Immunofluorescence Microscopy
For immunofluorescence microscopy, hESCs were plated as single cells in mTeSR1 with 10 μM Y-27632 2HCl in Matrigel-coated, 24 well, #1.5 glass bottom plates (Cellvis) at 7x103/cm2 for siCdt1, Trophectoderm, at 5x103 for siCdc6, Trophectoderm, and at 4x103/cm2 for siCdt1, Endoderm. Cells were incubated with siCdt1 for 20 hours (Trophectoderm), 24 hours (Endoderm) or siCdc6 for 32 hours (Trophectoderm) all in parallel with siControl as described above (siRNA transfections). After siRNA treatment, cells were differentiated as described above (Differentiation) with the following modifications: For trophectoderm, multiple BMP4 concentrations and treatment times were used as indicated (Fig. 7, Supplementary Fig. 6). For treatment less than 48 hours, cells were incubated in mTeSR1 after siRNA treatment until starting differentiation. (Example: 12 hours of mTeSR1 then 36 hours of BMP4, for a total of 48 hours). For endoderm, the first RPMI/Activin/CHIR-99021 was immediately after siRNA, without a day of incubation in mTeSR1. After differentiation, cells were fixed in 4% paraformaldehyde in PBS for 15 minutes at room temperature, washed with PBS, and permeabalized with 5% BSA, PBS, 0.3% triton x-100 at 4 °C overnight. Next, cells were incubated in primary antibody in 5% BSA, PBS, 0.3% triton x-100 at 4 °C overnight. Cells were washed with PBS at room temperature, then incubated in secondary antibody in 5% BSA, PBS, 0.3% triton x-100 at room temperature for 1 hour. Cells were washed with PBS, then incubated in 1 μg/mL DAPI in PBS for 10 minutes at room temperature, then washed with PBS. For trophectoderm the primary antibodies were Oct4 (Millipore, Cat#MABD76) and Cdx2 rabbit (Abcam, Cat#ab76541), the secondary antibodies were goat anti-mouse-Alexa 594, donkey anti-rabbit-Alexa 488. For endoderm the primary antibody was Sox17 (R&D Systems, Cat#AF1924), the secondary antibody was donkey anti-goat-Alexa 594 (Jackson ImmunoResearch). Cells were imaged in PBS on a Nikon Ti Eclipse inverted microscope with an Andor Zyla 4.2 sCMOS detector. Images were taken as 3x3 scan of 20x fields with a 0.75 NA objective, stitched with 15% overlap between fields using NIS-Elements Advanced Research Software (Nikon). Shading correction was applied within the NIS-Elements software before acquiring images. Raw images were quantified using a custom CellProfiler pipeline.
qPCR (Figure 3)
RNA lysates were prepared using Norgen Biotek’s Total RNA Purification Kit (Cat. 37500). Lysates were first treated with Promega RQ1 RNase-Free DNase (Promega), and then converted to cDNA using Applied Biosystem’s High-Capacity RNA-to-cDNA Kit (Cat. 4387406). Quantitative real-time PCR (qPCR) with SYBR Green (Bio-Rad; SsoAdvanced Universal SYBR Green Supermix, Cat. 1725271) was carried out to assess gene expression. All results were normalized to ACTB. Primers for qPCR were ordered from Eton Bioscience. Primers:
POU5F1-F: 5′-CCTGAAGCAGAAGAGGATCACC,
POU5F1-R 5′-AAAGCGGCAGATGGTCGTTTGG,
CDX2-F 5′-ACAGTCGCTACATCACCATCCG,
CDX2-R 5′-CCTCTCCTTTGCTCTGCGGTTC,
T-F 5′-CTTCAGCAAAGTCAAGCTCACC,
T-R 5′-TGAACTGGGTCTCAGGGAAGCA,
SOX17-F 5′-ACGCTTTCATGGTGTGGGCTAAG,
SOX17-R 5′-GTCAGCGCCTTCCACGACTTG,
CDT1-F 5′-GGAGGTCAGATTACCAGCTCAC,
CDT1-R, 5′-TTGACGTGCTCCACCAGCTTCT,
SOX2-F 5′-CTACAGCATGATGCAGGACCA,
SOX2 -R 5′-TCTGCGAGCTGGTCATGGAGT,
PAX6-F 5′-AATCAGAGAAGACAGGCCA,
PAX6-R 5′-GTGTAGGTATCATAACTC,
ACTB-F 5′-CACCATTGGCAATGAGCGGTTC,
ACTB-R 5′-AGGTCTTTGCGGATGTCCACGT.
Generating ARPE-19-iPS cells
ARPE-19-iPS cells were derived from the human retinal pigment epithelial cell line ARPE-19 by reprogramming with CytoTune-iPS 2.0 Sendai reprogramming kit (Invitrogen) following the manufacturer’s instructions.
Briefly, two days before Sendai virus transduction, 100,000 ARPE-19 cells were plated into one well of a 6-well plate with ATCC-formulated DMEM:F12 medium and were transduced with the CytoTune™ 2.0 Sendai reprogramming vectors at the MOI recommended by the manufacturer 48 hours later (d0). The medium was replaced with fresh medium every other day starting from one day after transduction (d1). At day 7, transduced cells were replated on Matrigel coated 6-well plates. Cells were fed with Essential 8 medium every day. Colonies started to form in 2-3 weeks and were ready for transfer after an additional week. Undifferentiated colonies were manually picked and transferred to Matrigel coated 6-well plates for expansion. After two rounds of subcloning and expansion (after passage 10), RT-PCR was used to verify whether iPS cells were vector-free with the primer sequences published in the manufacturer’s manual.
After iPS cells became virus-free, they were submitted to the University of Minnesota Cytogenomic Laboratory for karyotype analysis. This analysis indicated that the ARPE-19-iPS cells have normal karyotypes.
Immunofluorescence characterization of ARPE-19-iPS cells
To examine pluripotency markers, iPS cells were fixed with 4% paraformaldehyde for 20 minutes. If nuclear permeation was required, cells were treated with 0.2% triton-x-100 in phosphate-buffered saline (PBS) for 30 minutes, blocked in 3% bovine serum albumin in PBS for 2 hours, and incubated with the primary antibody overnight at 4°C. Antibodies targeting the following antigens were used: TRA1-60 (MAB4360, 1:400), TRA1-81 (MAB4381, 1:400), stage-specific embryonic antigen-4 (MAB4304, 1:200), and stage-specific embryonic antigen-3 (MAB-4303, 1:200), all from Millipore/Chemicon (Billerica, MA), OCT3/4 (AB27985, 1:200) from Abcam (Cambridge, MA), and NANOG (EB068601:100) from Everest (Upper Heyford, Oxfordshire, UK). Cells were incubated with secondary Alexa Fluor Series antibodies (all 1:500, Invitrogen) for 1 hour at room temperature and then with DAPI for 10 minutes. Images were examined using an Olympus FluoView 1000m IX81 inverted confocal microscope and analyzed with Adobe Photoshop CS6. Direct alkaline phosphatase (AP) activity was analyzed as per the manufacturer’s recommendations (Millipore).
Bisulfite sequencing and methylation analysis
Genomic DNA was isolated using the DNeasy Blood and Tissue kit (Qiagen) per manufacturer’s recommendations for isolation from mammalian cells. Bisulfite conversion was performed using the Epitect Bisulfite kit (Qiagen) according to the manufacturer’s protocol for low amounts of DNA. Single-step PCR amplification of the NANOG and OCT4 promoter regions were conducted using Accuprime Supermix II (Invitrogen). Amplification products were visualized by gel electrophoresis and bands were excised and purified using the QIAquick Gel Extraction kit (Qiagen). Purified PCR products were inserted into the PCR4-TOPO vector (Invitrogen) and individual clones were sequenced. Alignment and methylation analysis were performed using the online QUMA program (http://quma.cdb.riken.jp/). Sequenced clones with at least 90% non-CpG cytosine conversion and at least 90% sequence homology were retained for analysis.
Quantitative reverse transcriptase PCR (Supplementary Figure 2)
RNA was isolated using RNeasy Mini kit (Qiagen) and treated with TURBO DNA-free (Ambion, Austin, TX). First-strand cDNA was synthesized using a Superscript III First-Strand Synthesis SuperMix (Invitrogen). Reverse transcriptase-PCR was performed using TaqMan Gene Expression Assays and TaqMan Universal PCR Master Mix, No AmpErase UNG (Applied Biosystems, Carslbad, CA) as per the manufacturer’s protocol.
TaqMan gene expression assays used were OCT4 (Hs04260367-gH), SOX2 (Hs01053049-sl), NANOG (Hs04399610-g1), KLF4 (Hs00358836-m1), MYC (Hs00153408-m1), LIN28 (Hs00702808-s1), REXO1 (Hs00810654-m1), ABCG2 (Hs1053790-m1), DNMT3 (Hs00171876-m1), with GAPDH (Hs99999905-m1) used as an endogenous control. Expression levels were measured in duplicate. For genes with expression below the fluorescence threshold, the cycle threshold (Ct) was set at 40 to calculate the relative expression. Analysis was performed using an ABI PRISM 7500 sequence detection system (Applied Biosystems).
Teratoma analysis
ARPE-19-iPS cells contained in a mixture of DMEM/F12, Matrigel and collagen were implanted onto the hind flank of NSG mice (n=5) until a palpable mass formed. Teratoma tissue was excised for histological examination following embedding and staining by haematoxylin and eosin. Experiments were conducted with the approval of the Institutional Animal Care and Research Committee at the University of Minnesota.
Ergodic Rate Analysis
Ergodic Rate Analysis for cell cycle data was based on previously published work 33. First, the raw data from flow cytometry files were extracted using FCSExtract Utility (Earl F Glynn) to comma separated value (.csv) files. The data in the .csv files were then gated in FCS Express 6 (De Novo Software) and the data for only G1-MCMDNA were exported. MCM negative cells were excluded based on the negative control sample (see Flow Cytometry). The mean MCM loading rate was calculated in MATLAB (MathWorks). To calculate the mean MCM loading rate, the G1-MCMDNA were subdivided into 10 equal sized bins, with rate calculated for each bin, and all 10 rates were averaged together for a mean MCM loading rate. The rate calculation was based on the formula from Kafri et al: wn = MCM loading rate in bin n
α = ln(2) / doubling time (Supplementary Figure 2)
F = number of G1-MCMDNA cells / total number of cells in sample. F was calculated from FCS Express and entered into MATLAB manually (Supplementary Figure 2).
fn= number of cells in bin n / number of G1-MCMDNA cells
The bins were created in MATLAB (Figure S2). To control for small day to day differences in raw data from staining intensities, the histogram edges were defined with the first bin starting at the lowest MCM value and the last bin ending at the highest MCM value, divided into 10 equal sized bins between the lowest and highest MCM value. The 10 wn were then averaged for a final mean w per sample. Sample MATLAB code:
alpha_iPSC=log(2)/15.64; F_iPSC_1=0.0801; %calculate lowest and highest MCM values% maxMCM_iPSC_1=max(iPSC_1(:,1)); minMCM_iPSC_1=min(iPSC_1(:,1)); %create histogram with 10 bins and specified first and last bin limits% h10_iPSC_1=histogram(iPSC_1(:,1), ‘NumBins’, 10, ‘BinLimits’, [minMCM_iPSC_1, maxMCM_iPSC_1]); %calculate fn within each bin% totalf_iPSC_1=size(iPSC_1); fn10_iPSC_1=(h10_iPSC_1.Values)/totalf_iPSC_1(1,1); %calculate mean w% w10mean_iPSC_1=mean(alpha_iPSC .*(2-F_iPSC_1)./fn10_iPSC_1);The mean MCM loading rate was calculated for 3 biological replicates for each cell line, and the replicates were averaged using GraphPad Prism for further statistical analysis. We cannot use ergodic rate analysis on actively differentiating cells (e.g. Figure 3) because they are not at steady state.
Quantification and Statistical Analysis
Statistical analysis was performed with GraphPad Prism 7 using unpaired, two-tailed t test (displayed as mean ±SD) or two-tailed Mann-Whitney test as indicated in figure legends. Significance levels were set at *p≤ 0.05, **p≤ 0.01, ***p≤ 0.001, ****p≤ 0.0001. All experiments were performed a minimum of 2 times, and representative data are shown in figures.
AUTHOR CONTRIBUTIONS
J.P.M. designed and performed the experiments and analyzed the results. R.D. and P.C. performed experiments. R.M.B. W.C, K.T., B.R.W., J.T. and A.K.B. generated, validated the iPSCs and associated data in Supplementary Figure 2. J.E.P. and J.G.C. designed experiments and supervised the project. J.P.M and J.G.C. wrote the manuscript with input from the other authors.
ACKNOWLEDGMENTS
We are grateful to Ran Kafri for advice on ergodic rate analysis. We thank Jeffrey Jones for managerial assistance, Dr. Sam Wolff for microscopy assistance, the UNC Human Pluripotent Stem Cell Core stem cell culture assistance and A. Adams, S. Wong, M. Consuegra, S. Goraya, and S.Sisk for technical assistance. We also thank Dr. Robert Duronio, Dr. Michael Emanuele, and the Cook lab for helpful discussions. We thank Ron McElmurry and Megan Riddle at the University of Minnesota for conducting teratoma assays, and the Cytogenomics Shared Resource (P30 CA077598) at the University of Minnesota Masonic Cancer Center for karyotype analysis. The UNC Flow Cytometry Core Facility is supported in part by P30 CA016086. This work was supported by a fellowship from the NSF (DGE-1144081) to J.P.M and by grants from the NIH to J.G.C. (GM083024 and GM102413), R.M.B. (T32-CA009138) J.E.P. (DP2-HD091800) and A.K.B. (GM074917). Additional funding was provided by the W.M. Keck foundation to J.E.P. and J.G.C; J.T. is supported by the Tulloch Chair in Stem Cell Biology, Genetics and Genomics at the University of Minnesota.
COMPETING INTERESTS
We have no competing financial or non-financial interests.