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[ { "id": "18162134__text", "type": "full_text", "text": [ "BackgroundMuch of our current knowledge of the molecular expression profile of human embryonic stem cells (hESCs) is based on transcriptional approaches. These analyses are only partly predictive of protein expression however, and do not shed light on post-translational regulation, leaving a large gap in our knowledge of the biology of pluripotent stem cells.ResultsHere we describe the use of two large-scale western blot assays to identify over 600 proteins expressed in undifferentiated hESCs, and highlight over 40 examples of multiple gel mobility variants, which are suspected protein isoforms and/or post-translational modifications. Twenty-two phosphorylation events in cell signaling molecules, as well as potential new markers of undifferentiated hESCs were also identified. We confirmed the expression of a subset of the identified proteins by immunofluorescence and correlated the expression of transcript and protein for key molecules in active signaling pathways in hESCs. These analyses also indicated that hESCs exhibit several features of polarized epithelia, including expression of tight junction proteins.ConclusionOur approach complements proteomic and transcriptional analysis to provide unique information on human pluripotent stem cells, and is a framework for the continued analyses of self-renewal.\nHuman embryonic stem cells (hESCs) are pluripotent cells isolated from the inner cell mass of the blastocyst [1]. They can be maintained for prolonged periods in culture and differentiate to representatives of the three germ layers as well as trophoblasts and germ cells. This differentiation potential may be used to model certain aspects of human embryogenesis, including the development and differentiation of pluripotent and other stem cell types during the processes of gastrulation, neurogenesis and organogenesis. Thus, hESCs provide a unique and powerful system to study otherwise intractable aspects of human development. Furthermore, these approaches have the potential to provide differentiated cell types for cell replacement therapies of degenerative disorders such as Parkinson's disease and Type I diabetes [2,3]. Before these cell therapy applications are developed, an understanding of the molecular and cellular mechanisms that drive self-renewal and differentiation is required. Fundamental to this understanding is the elucidation of the transcriptome and proteome of hESCs, using approaches that lay a framework for functional analyses of the unique properties of these cells. Large-scale gene expression analyses such as microarray, massive parallel signature sequencing (MPSS), expressed sequenced tag (EST) enumeration, and serial analysis of gene expression (SAGE) have been used to compare multiple hESC lines [4-7]; hESCs to germ cell tumors [8]; or to differentiated derivatives in embryoid bodies [9-11] or neural populations [12]. These approaches have highlighted an expanded set of transcripts that mark the pluripotent state [4,13,14], cross-species commonalities in the molecular profile of ESCs [6,12,15], prominent receptors expressed by hESCs [8] and pathways that may play a role in the regulation of pluripotency [16,17]. Nevertheless, cataloguing the cellular transcriptome is only predictive of protein expression and typically does not shed light on post-transcriptional regulation. For example, while tens of thousands of transcripts can be followed simultaneously with SAGE, microarrays and MPSS, these methods do not routinely detect differences in transcript splice variants, or polyadenylation status. These differences may have profound effects on translation, as well as the isoform and function of the protein produced. Finally, numerous post-translational modifications are known to regulate protein function, including enzymatic cleavage, covalent coupling to other molecules, glycosylation, phosphorylation and ubiquitination. These issues all highlight potential shortfalls in our understanding of the hESC proteome. Several practical approaches for proteomic analyses are currently available, the most established of which is the 2-dimensional (2D) separation of proteins by polyacrylamide gel electrophoresis (PAGE). HPLC-tandem mass spectrometry (HPLC-MS/MS) based technology is rapidly evolving and has recently been used to detect protein expression in multiple cell types. An alternate approach is the recent large-scale adaptation of standard western blotting [18]. In this procedure, a large well is used to separate the sample by PAGE and lanes are created on the membrane containing immobilized protein with the use of a manifold. Compatible combinations of primary antibodies are predetermined, with the criterion of being able to identify proteins that do not co-migrate. Different combinations of primary antibodies are added to each well, with appropriate dilutions of each primary antibody so that expressed proteins are detected in a single condition. The scalability of the system depends on defining suitable combinations of primary antibodies, with up to 1000 antibodies in 200 lanes being used in the largest screens thus far. Detection software is used to identify proteins based on their expected and observed gel mobility. Unlike 2D PAGE and HPLC-MS/MS, large-scale western blotting only identifies proteins for which antibodies are already available. While this is not an appropriate screen for identifying uncharacterized proteins, it greatly simplifies the verification and functional analyses of proteins that are detected. In addition, this approach is highly flexible, and if desired can be focused to particular sets of proteins or protein function, such as cell signaling molecules. Importantly, the foundation of this approach is the large amount of data on individual antibodies, which are already available and characterized in the literature. More recently, two research groups have conducted proteomic analyses of hESCs using MS [19-22]. In the present study, we used two large-scale western blot systems to examine the expression of > 1000 proteins in hESCs and detected > 600 proteins that were grouped into 18 functional classes. In addition, we identified 42 examples of multiple bands for a single protein, likely to be protein isoforms and/or post-translational modifications, and 22 phosphorylation events in cell signaling molecules. We correlated the expression of members of key active pathways in our transcriptional and proteomic databases and confirmed the validity of this approach. Using these approaches we identified new markers for undifferentiated hESCs and highlighted unrecognized epithelial characteristics of hESCs. Our data confirm the importance of proteomic analyses in complementing transcriptional profiling and provide a framework for continued analyses of the molecular and cellular biology of pluirpotent hESCs.\nPowerBlot analysis of hESCsWe first employed a large-scale western blot screen, the PowerBlot system, to profile protein expression in undifferentiated hESCs. This system used 934 antibodies toward proteins representing 22 diverse classes of function, such as transcription factors, the MAP kinase (MAPK) pathway, and apoptosis, among others. To expand a large-scale culture of BG01 cells for this assay, a collagenase- and trypsin- based passaging method was used [23]. While these conditions have been associated with the accumulation of trisomies of chromosomes 12, 17 and X [24], the ease of use of these cultures and similarity in gene expression and differentiation potential to karyotypically normal BG01 hESCs [11,24,25] make them suitable for such large scale applications. For the PowerBlot screen, whole cell lysate from BG01 hESCs was separated on five 4–15% gradient gels. Each blot contained size markers and 39 lanes. Each lane was screened with 1–8 antibodies in combinations that had been predetermined to enable accurate identification of well-separated proteins (Fig. 1A–E). The gels and blots were performed in duplicate and expressed proteins were identified by their predicted size and verified by visual inspection.Figure 1PowerBlot analysis of undifferentiated BG01 hESCs. This large-scale western blot consisted of five gels run in duplicate and probed with 934 antibodies. (A-E) One set of blots is shown at a contrast that highlights most bands. (F) A representative lane (gel C, lane 24) aligned with protein markers used for band identification. (G) Scatter-plot of the normalized average intensity (i.u.) values for each protein indicating a linear relationship between duplicate blots. Datasets for this analysis are in Additional Tables 1 and 2.A total of 545 antibodies detected bands of appropriate size, which could be compressed to 529 proteins with unique SwissProt identification numbers (Fig. 1A–E and Additional File 1). An enlargement of a representative lane (lane 24 of Blot C) alongside protein markers is shown in Fig. 1F. Thirteen proteins including AKT, caveolin1 and ERK1 were detected in multiple lanes using the same or different antibodies. Information on the antibody catalogue number and dilution, band intensity for each repeat and the averaged value, description of protein function, and Entrez gene and SwissProt database identification numbers is shown in Additional File 1. Three hundred and eighty three antibodies did not detect bands in this screen, indicating lack of expression, or possibly technical issues with detection under standard conditions (Additional File 1).The size of the detected proteins ranged from 15 kD (GS15) to 280 kD (ABP-280). The average intensity of the detected proteins ranged from 195 to 117926 normalized intensity units (i.u.), with an average of 5367 i.u. The proteins with the highest band intensity were the B2 Bradykinin Receptor (117926 i.u.), Karyopherin α (80698 i.u.), and BiP (74922 i.u.), whilst the proteins with the lowest intensity that could be verified by visual inspection were Inhibitor 2 (247 i.u.), Caspase 8 (201 i.u.), and OXA1Hs (195 i.u.). Finally, the consistency of this assay was demonstrated by plotting the normalized average intensity values for each protein, which revealed a linear relationship between the duplicate samples (Fig. 1G). Kinexus analysis of hESCsA more focused screen was used to profile expression of protein kinases, phosphatases and phosphorylated sites in cell signaling molecules in hESCs. The Kinexus assays contained 140 antibodies to these related classes of proteins and phospho-sites. Karyotypically normal BG03 hESCs grown on a fibronectin matrix in MEF-CM [26] were used for this analysis, and whole cell lysate was separated on four 12.5% gels for western blotting. Eighty five immunoreactive bands were identified, representing 38 protein kinases and 16 phosphatases, their isoforms, and 22 phosphorylated sites in signaling molecules (Fig. 2A–D, Additional File 1). Sixty-four antibodies did not detect their corresponding antigen (Additional File 1).Figure 2Kinexus blots of undifferentiated BG03 cells. Four blots were used to probe BG03 lysate with (A, B) 76 antibodies for protein kinases, (C) 27 antibodies for phosphatases and (D) 37 antibodies for phosphoylated sites in cell signaling molecules. Identified bands are indicated (*). Datasets for this analysis are in Additional Tables 1 and 2. Functional classification of proteins expressed in hESCsThe PowerBlot and Kinexus assays identified a diverse range of proteins expressed in hESCs. To further annotate these data, the detected proteins were ordered into 18 subgroups based on protein function (Additional File 2). For example, 16 factors with known or implied roles in the regulation of self-renewal or pluripotency of mESCs or hESCs, such as Oct4 [27], STAT3 [28], members of the FGF [29], PI3 kinase [30], Src [31] or MAPK pathways [32], and phosphorylated isoforms of GSK3, STAT3 and p38 MAPK, were grouped under \"Pluripotency\" (Fig. 3A and Additional File 2). Another functional group (Cell surface) consisted of 20 transmembrane or cell surface proteins (Additional File 2). This included several receptors for peptides and growth factors, such as neurotensin receptor 3, the B2 bradykinin, endothelin 1, and thrombin receptors, and the glial derived neurotrophic factor receptor α (Fig. 3B). These molecules may be useful as targets for cell sorting experiments, and expression of these receptors could identify bioactive peptides or growth factors that may influence hESC self-renewal or differentiation.Figure 3Functional classification and mobility variants of proteins detected in hESCs. (A) Proteins with known or suggested roles in self-renewal are shown, including Oct4, STAT3, Smad2/3 and FGF2 (Additional Table 2, \"Pluripotency\"). Isoforms of FGF2, and phospho-GSK3 are indicated (*). (B) Cell surface proteins are shown, including Connexin 43, E-Cad and GDNFRα (Additional Table 2, \"Cell Surface\"). Other functional classes of proteins are indicated in Additional Table 2. (C) A total of 42 proteins, including FGF2, HSP70 and ERK1, were found to have multiple bands in either the PowerBlot or Kinexus blots. These bands migrated closely but were sufficiently separated from other detected proteins. Bands predicted to be isoforms of the indicated protein are highlighted in some panels (*).Other functional classification of the proteins detected by the PowerBlot screen included: transcription factors (71 proteins), nucleus and nuclear transport (144), cytoskeleton (75), cell adhesion (45), MAP kinase pathway (24), protein kinase A (13), protein kinase C (20), tyrosine kinases (15), adaptors and tyrosine kinase substrates (51), protein phosphatases (17), GTPases and regulators (42), calcium signaling (23), cell cycle (87), apoptosis (61), membrane research (62), and other functions (51) (Additional File 1). Some proteins were included in multiple functional categories due to overlapping properties, such as AIM-1, which was included in the cell cycle as well as in the nucleus/nuclear transport categories. The Kinexus expression data was organized separately into cell signaling-related functional groups (Additional File 1). In addition, 35 proteins were detected by both the PowerBlot and Kinexus systems (Table 1).Table 1Proteins detected by both PowerBlot and Kinexus systemsProtein nameSwiss NrProtein nameSwiss NrBMXP51813MEK2P36506CaM Kinase KinaseQ64572MKP2Q62767Casein Kinase I epsilonP49674p38 alpha/SAPK2aQ16539Casein Kinase II alpha/CK2aP19139PaxillinP49024Cdk1/Cdc2P06493PKA CP17612Cdk5Q00535PKC betaP05771Cdk7P50613PKC deltaQ05655DAP KinaseP53355PP2A Catalytic alphaP05323DAP3P51398PP5/PPTP53042ERK1Q63538PTP1BP18031ERK2P27703PTP1C/SHP1P29350FAKQ00944PTP1D/SHP2Q06124GSK-3 betaP18266RbP13405I kappa B alphaP25963RskQ15418IKK betaO14920Stat1A46159JAK1P23458Stat3P52631JNK1P45983VHRP51452MEK1Q02750 Detection of protein isoforms or post-translational variantsUnlike many cDNA-based gene expression assays, western blotting has the capacity to detect multiple protein isoforms due to translation of different mRNA splice variants, as well as post-translational modifications such as enzymatic cleavage, glycosylation, or phosphorylation. Examination of the blots described here identified 42 examples of multiple banding for a single target antigen (Fig. 3C). These candidates exhibited closely migrating multiple bands, which were close to their predicted size but were sufficiently separated from other proteins. For example, four closely migrating bands were observed for FGF2 (Fig. 3C, top panel), which may represent known glycosylation variants of this growth factor [33]. Other known examples of post-translational modifications included those of HSP70, IKKgamma and ERK1. Verification of protein expression by immunocytochemistryThe PowerBlot and Kinexus assays identified proteins based on their expected and observed molecular weight, using combinations of antibodies that had been predetermined to detect proteins of sufficiently different sizes. Proteins known to be expressed by hESCs and also identified by these assays, included Oct4, E-CAD, Connexin 43 and Hsp70. To verify expression using a complementary approach, we performed immunoflurorescent staining for 10 proteins not previously reported to be expressed in hESCs by immunocytochemistry, using karyotypically normal BG01 cultures (Fig. 4A–K). These included ABP-280, a homodimeric actin-binding protein often associated with membrane glycoproteins; CtBP1 and CtBP2, two C terminal binding proteins that are a class of transcription corepressors; GS-28, a golgi protein; HDJ-2, a member of the DnaJ-related Hsp40 (heat shock protein 40) subfamily; L-Caldesmon, a cytoplasmic actin-binding protein; Rabaptin, a GTP-binding protein; phosphorylated-p130 Cas, a docking protein with an amino-terminal SH3 domain that may function as a molecular switch that regulates CAS (Crk-associated substrate) tyrosine phosphorylation; Ras-GAP and phosphorylated Ras-GAP (p-Y460), a protein that down-regulates the signal transducer p21ras; and ShcC, a protein with an N-terminal phosphotyrosine-binding domain. These proteins were all expressed by hESCs, with the expected subcellular localization (Fig. 4A–K). Oct4 was used as a positive control (Fig. 4L). These results suggested that most of the bands in the PowerBlot and Kinexus assays were likely to be correctly identified.Figure 4Verification of protein expression using immunocytochemistry. (A-K) Ten proteins that were detected in undifferentiated hESCs by western blotting were also detected by immunofluorescence of BG01 cells grown in MEF-CM. Ras-GAP (pY460) is a phosphorylated form of Ras-GAP. The same antibodies were used in this analysis as in the PowerBlot assay, except phospho-p130 Cas (Tyr165). (L) Oct4 was used as a positive control. (M-R) Oct4, TNIK and p130 Cas as markers of undifferentiated hESCs. BG01 cultures were partially differentiated by exposure to 10% fetal bovine serum for 3 days. (M) Oct4 was expressed uniformly in undifferentiated cells, (P) but was downregulated in morphologically differentiated areas after 3 days in serum (arrowhead). (N) TNIK expression was localized to the cytoplasm, and (N, Q) expression appeared to be restricted to morphologically undifferentiated cells (arrowhead). (O) p130 Cas was detected in a membrane/peripheral-cytoplasmic pattern in undifferentiated cells, (R) but this distribution was substantially altered in differentiating cells with a flattened morphology, which exhibited a general cytoplasmic, or perinuclear profile. Scale bar for A-L: (A, L) 200 μm; (C, D, F, H, I, J, K) 100 μm; (B, E, G) 50 μm. Scale bar for M-R: (M, N, P, Q): 100 μm ; (O, R): 50 μm.Preliminary analyses also indicated that expression of some of these proteins was downregulated in differentiated cells, including p130 Cas and the Traf2- and Nck-interacting kinase (TNIK). TNIK is known to be involved in the inhibition of cell spreading via disruption of F-actin [34,35]. Immunofluorescence was used to examine the expression of TNIK and p130 Cas during early differentiation of hESCs. BG01 cultures were partially differentiated by growth in serum containing media for 3 days. This condition generated heterogeneous populations containing Oct4+ cells with characteristic hESC morphology and less tightly packed, and morphologically differentiated areas, lacking expression of Oct4 (Fig 4M, P). TNIK was expressed highly in undifferentiated hESCs, and in the undifferentiated areas at day 3, but was downregulated in areas undergoing morphological differentiation (Fig 4N, Q). This may indicate that TNIK is active in hESCs and degraded rapidly upon differentiation. p130 Cas was detected in a membrane/peripheral-cytoplasmic pattern in hESCs (Fig 4O). The distribution of p130Cas was substantially altered in differentiating cells with a flattened morphology, exhibiting a general cytoplasmic, or perinuclear profile (Fig 4R). This could indicate an alteration in the function of p130 Cas as pluripotent cells differentiate. These analyses suggested that the change in expression or distribution of these proteins could be used as markers for undifferentiated hESCs. Comparison of proteomic and transcriptional profiles of hESCsWe have previously employed the Illumina Bead Array system for the large-scale profiling of gene expression in hESCs using 24,000 transcript probes [11]. To compare proteomic and transcriptional analyses of hESCs, the levels of > 600 proteins detected using large scale blotting were correlated with the levels of transcripts detected with the Illumina platform (Additional File 3). In general, a close match between the expression level of transcript and protein was observed: transcripts for nearly all the detected proteins were also identified in the Illumina analysis, and most proteins expressed at high levels also exhibited high mRNA levels.We reasoned that a focused comparison of specific signaling pathways using a combination of proteomic and transcriptional data was likely to be much more informative than a global interrogation of hESCs. Several major signal pathways that have been suggested to be involved in self-renewal were examined to test this approach. These included the FGF, TGFβ, GSK3β/Wnt/β-catenin and Jak/Stat pathways [17,29,36-39], as well as the more recently suggested MAPK/ERK and Gap junction pathways [32,40]. Correlating transcriptional and proteomic data provided direct confirmation that these pathways were present and likely functional in hESCs (Table 2). For example, FGF2 protein was expressed highly in hESCs and expression of key members of the TGFβ, Wnt, Jak/Stat and Gap junction pathways, namely Stat1, SMADs, GSK3β, β-catenin and Connexin 43, were detected in both transcriptional and proteomic databases.Table 2Signal pathways that may be active in hESCsNameProteinmRNATGF βStat1++++++PAI-1/SERPINE1+++-Smad2/3++++Jun+++Smad4/DPC4+++Endoglin+-WntCtBP2+++++++PP2A Catalytic alpha/PPP2CA+++++++EBP50/SLC9A3R1++++++beta-Catenin/Ctnnb1++++Cyclin D3/CCND3++++GSK-3 beta++++Jun+++Casein Kinase II alpha/CSNK2A1++++Jak-StatStat1++++++Crk++++Stat3/2+++++Stat6+++++PTP1B+++++JAK1++-Glucocorticoid R/NR3C1++-Thrombin Receptor/PAR1/F2R+++SHPS-1/PTPNS1++++MCM5+++++Smad2/3++++Tyk2++++Jun+++Bcl-x/BCL2L1++++Smad4/DPC4+++Stat5A++GPCRB2 Bradykinin Receptor/BDKRB2++++-Neurotensin Receptor 3/SORT1+++-Endopeptidase 3.4.24.16/NLN++++IP3R-3++++SHC++++Gap JunctionCdk1/Cdc2++++++GRB2++++++MEK1/MAP2K1++++++PKA C++-PKA RI alpha++-PKC alpha++-C-Raf/RAF1++++ZO-1/TJP1+++++Connexin-43/GJA1++++IGFPKC iota++++++MEK1/MAP2K1++++++Rsk/RPS6KA1+++++GRB2++++++MEK2/MAP2K2++++++PI3Kinase/PIK3R1+++++pan ERK/MAPK1+++++Crk++++eIF-4E+++++ShcC+++-PAI-1/SERPINE1+++-C-Raf++++SHC+++++PKC beta/PRKCB1++++NCK++++PKB alpha/Akt+++GSK-3 beta++++Ercc-1++++Fatty Acid Synthase/FASN+++++Jun+++RAFT1/FRAP++++PTP1D/SHP2/PTPN11++++SCAMP1++++Bcl-x/BCL2L1++++p70s6k/RPS6KB1+-PI3-Kinase p170/PIK3C2A++PTP1B/PTPN1+++Dok1/p62dok+++PI3-Kinase p110 alpha/PIK3CA+-ERBBEphA4/Sek++++-ShcC/SHC3+++-c-erb-B2/ERBB2++++C-Raf/RAF1++++SHC/SHC1+++++GDNFI kappa B epsilon/NFKBIE++++++GRB2++++++MEK2++++++NCK+++++C-Raf++++Ras-GAP/RASA1++++SHC+++++GDNFR-alpha/Gfra1++-Jun+++IKK beta++++pan-JNK/SAPK1/MAPK10+++NBS1/ARTN++Dok1/p62dok+++Tight JunctionPTEN++++++PP2A Catalytic alpha+++++++PKC iota++++++Sec8/SEC8L1+++++beta-Catenin/CTNNB1++++CDC42+++++AF6/MLLT4+++++PKC alpha++-Yes++++Rho/ARHA+++++ZO-1/TJP1+++++CASK+++Symplekin/SYMPK++-Ras/NRAS++++Casein Kinase II alpha/CSNK2A1++++VAP33/VAPA+++alpha-Catenin/Ctnna1+++MAPKpan ERK++++++MEK1++++++Rsk++++++ERK2++++++MEK2/Map2k2++++++MST3/STK25++++++ERK1+++++CDC42+++++C-Raf++++p38 alpha/SAPK2a++-G3BP+++++TFII-I/GTF2IRD1++++MST1/STK4++++MKP2/Dusp4++++Ras++++Phospho-p38MAPK (T180/Y182)+++pan-JNK/SAPK1+++Inhibitor2/PPP1R2++++ABP-280++++++14-3-3 epsilon/YWHAE++++MAPKAPK-5+++TAO1+*PBK+++MKK3b/Map2k3++Protein expression level: > 10,000: ++++; 5,000–10,000: +++; 1,000–5,000:++; 100–1,000: + mRNA gene expression level: > 5,000:++++; 1,000–5,000: +++; 100–1,000: ++; 30–100: +*: not included in the gene expression arrayThis independent confirmation of known networks led us to examine other pathways that showed a similar correlation but have not been identified as key regulators of either self-renewal or differentiation, or suggest unappreciated characteristics of hESCs. Four signaling pathways (IGF, ERBB2, GPCR, and GDNF) and the tight junction complex were highlighted by this analysis (Table 2), and expression of key proteins in these pathways was confirmed. A detailed study demonstrating the importance of the IGF and ERBB2 pathways in hESC self-renewal has been performed and enabled the development of a defined medium for hESC maintenance (TCS and AJR, submitted). Tight junctions are apical cell-cell junctions found in epithelia that establish a barrier to the extracellular environment and a border for apical-basolateral polarity. While hESCs grow in colonies that are highly reminiscent of epithelia, and have been shown to be coupled by gap junctions [40], the formation of tight junction complexes has not been described. hESCs expressed the ZO1 and occludin tight junction proteins along cell borders as expected in polarized epithelia. The distribution of ZO1 expression changed dramatically as hESCs proliferated in culture. When tight junction complexes were disrupted by disaggreagation to single cells, only a subset of cells showed ZO1 staining 4 days after plating (Fig. 5). Continued proliferation to a confluent monolayer on day 7 was accompanied by widespread expression of ZO1, suggesting the formation of a general tight junction barrier. These cultures were undifferentiated and retained uniform expression of Oct4 protein (not shown). ERBB2 and 3 are members of the epidermal growth factor (EGF)-receptor family, which regulate epithelial proliferation via EGF-family ligands. ERBB2 and 3 transcripts are expressed by hESCs [8], are known to function as a heterodimer [41], and transmit a strong proliferative signal for hESCs by Heregulin 1β (an EGF-family ligand) (TCS and AJR, submitted). Immunofluorescence revealed general cell surface expression of ERBB2 on hESCs. Conversely, ERBB3 was highly localized to a concentrated area, and observed in cells that also expressed ZO1. Epithelial cells are known to localize ERBB receptors to the basolateral side of tight junctions, which serves to functionally separate receptors from ligands [42,43]. This is a basic epithelial wound healing mechanism, whereby disruption of the tight junction barrier by injury immediately exposes receptors to extracelluar ligands [43]. These staining patterns are also suggestive of basolateral sorting of ERBB3 in hESCs. The pathways and complexes identified by these analyses lay a framework for future functional analyses of signaling networks in hESCs.Figure 5Tight junction proteins and ERBB2/3 expression in hESCs. BG01 hESCs were disaggregated to single cells using accutase [52] and cultured in defined conditions. (A) ZO1 expression four and (B) seven days after plating, indicating progressive tight junction formation. (C) Occludin expression 5 days after plating. (D) General cell surface expression of ERBB2, in the same field of view as (A). (E) Localized expression of ERBB3, in the same field of view as (B). (F) Higher magnification of ERBB3 localization in ZO1 expressing BG01 cells, 5 days after plating. Nuclei were stained with DAPI.\nAttempts to harness the potential of hESCs for models of human embryogenesis and cell therapy applications will be greatly enhanced by a detailed understanding of their molecular characteristics. This includes definition of the transcripts, splice variants, and protein isoforms expressed by these cells. Post-translational modifications such as phosphorylation and glycosylation, and the receptors and signaling pathways active in the pluripotent state, or during early differentiation, also need to be determined. This should also be complemented by an understanding of epigenetic characteristics of pluripotency, including methylation, imprinting and chromatin conformation. Such a comprehensive definition of the molecular state of hESCs will enable more accurate prediction and testing of the conditions used for growth and differentiation of hESCs, by precise genetic modification or application of specific growth factor cocktails and reagents. For example, a scalable, fully defined and GMP-certified culture system will need to be developed for the eventual development of hESC-based cellular therapies. Progress has been made in defining growth factor conditions that support self-renewal [44-46], and hESC lines have been isolated in the absence of mouse embryonic fibroblasts and in animal protein free culture conditions [47,48]. A more refined understanding of the biology of hESCs has contributed the development of a defined medium utilizing ligands for IGF1R and ERBB2/3 receptors to promote in self-renewal (TCS and AJR, submitted). We and others have performed transcriptional analyses of hESCs, using cDNA and oligonucleotide microarrays, SAGE, MPSS and EST enumeration. These techniques have enabled the collation and comparison of transcriptional profiles from multiple hESC lines and their differentiated derivatives and have highlighted an expanded set of hESC specific markers and signaling pathways that may regulate self-renewal or differentiation. Using pathway analysis we were also able to identify key pathways that are active in ESCs (reviewed in [16]). While these efforts have been highly valuable in defining the transcriptional profile of undifferentiated hESCs, they are only predictive of translation and do not shed light on post-translational events in this unique cell type. These processes may also be highly regulated, which could contribute significantly to the overall conversion of genetic information to actual protein function. We report here a proteomic analysis of pluripotent hESCs by using two large-scale western blotting systems and highlight post-translational events in undifferentiated hESCs. The expression of 545 bands was detected, potentially representing 529 proteins, or their migratory isoforms. In addition, one hundred and forty phospho-specific antibodies were used to identify 85 different phosphorylated sites, on 76 proteins in these cells. The detected proteins were annotated into functional classes representing diverse cellular processes. For example, multiple proteins were detected that have been suggested to regulate the pluirpotent state in mouse ESCs or hESCs. Defining the interplay of these multiple signaling pathways will be critical in understanding the self-renewal versus differentiation decisions of hESCs. Therefore, our data provide a powerful framework for the functional analysis of specific proteins, protein classes, or molecular pathways. In particular, the availability of antibodies for candidate proteins is a major benefit of this approach compared to 2D-gel or HPLC-MS/MS based proteomics. Although these western blotting approaches are currently more limited in scope than most large-scale cDNA based assays, detecting up to 1000 proteins compared to tens of thousands of transcripts, they have the potential to highlight translational events and post-translational modifications. By comparison, SAGE and MPSS are limited to detecting short sequence \"tags\" adjacent to the poly-A tail of transcripts, and may not distinguish splice variants with the same 3' exon. We detected 42 proteins with multiple closely migrating bands (Fig. 3C), suggestive of closely related isoforms or post-translational modifications such as phosphorylation. These focused proteomic approaches are therefore likely to be highly complimentary to transcriptional analyses in investigating the functional expression of the genome in hESCs and during cellular differentiation. One potential issue with this approach is that multiple antibodies are included in each lane, which could possibly lead to misidentification of bands. To demonstrate that identified proteins were expressed in hESCs, the same antibodies used in the PowerBlot assay were used to confirm expression of 10 representative proteins by immunofluorescence (Fig. 4). Furthermore, 13 proteins were detected with multiple different antibodies, and 35 proteins (Table 1) were detected in both the PowerBlot and Kinexus assays. This provided internal, or independent, confirmation of expression of these proteins. Other studies have also demonstrated the expression of several of the proteins we detected in hESCs. These include Oct4, a key marker of the pluripotent state, Connexin 43 and GSK3β, confirming the reliability of large-scale western blotting. Finally, several proteins detected by our assays were also detected in hESCs by MS approaches including Karyopherin α [19]. Additionally, the PowerBlot assay was performed in duplicate, and was shown to be highly reproducible. This suggested that this approach should be informative when comparing hESCs to their differentiated derivatives. Two candidate proteins, TNIK and p130 Cas, were downregulated, or exhibited altered localization upon spontaneous differentiation of hESCs, respectively. This indicated that they were novel markers of undifferentiated cells and molecules that could be functionally involved with self-renewal. It is impossible in an initial manuscript to analyze and rigorously test all the predictions that could be made from comparing transcriptional and proteomic data sets. However, we did examine key features to illustrate the power of this methodology. Potential new markers for hESCs were identified, the expression and activation of proteins in key self-renewal pathways were confirmed, and a diverse range of proteins were detected and expression correlated with transcriptional analyses. In addition, we highlighted several candidate signaling pathways that may be relevant to self-renewal. Examination of tight junction protein expression indicated that undifferentiated hESCs could form polarized epithelia, which has also been recently suggested by ultrastructural analyses [49]. Discrete localization of ERBB3 may also suggest basolateral separation of this receptor from soluble ligand. These analyses highlight that predictions from a combination of transcriptional and proteomic approaches will serve to focus the investigation of hESCs in the future.\nIn summary, we generated a focused proteome of hESCs using large-scale western blotting and sorted the detected proteins according to function and signaling pathways. This characterization provides important basic information on expressed proteins, their isoforms and post-translational modifications, and tools for the continued investigation of the underlying molecular characteristics of hESCs. Importantly, we provide a list of tools, in the form of commercially available antibodies, which can be used to interrogate the function of these molecules in self-renewal or differentiation.\nCulture of human embryonic stem cellsFor the PowerBlot analysis, enzymatically passaged BG01 hESCs were grown as described previously [23]. These conditions were necessary to scale up the culture to generate the milligram amounts of protein lysate required for this analysis. These conditions maintain cell populations that express the appropriate markers of pluripotency and can differentiate to representatives of all three germ layers, but may lead to eventual accumulation of trisomies for chromosomes 12, 17 or X [26]. For the Kinexus assays, BG03 hESCs were maintained in MEF-conditioned medium (MEF-CM) without the accumulation of karyotypic abnormalities as described previously [14,26].hESCs were also maintained in a defined medium as indicated. These conditions are described in detail elsewhere (TCS, AJR, submitted). Briefly, the media consisted of DMEM/F12 (Invitrogen), 2% fatty acid-free Cohn's fraction V BSA (Serologicals), 1× nonessential amino acids, 50 U/ml penicillin/streptomycin, 50 μg/ml ascorbic acid, 10 μg/ml bovine transferrin, 0.1 mM β-mecaptoethanol (all from Invitrogen), 1× Trace Elements A, B & C (Mediatech), 10 ng/ml hergulin1β (Peprotech), 10 ng/ml activinA (R&D Systems), 200 ng/ml LR3-IGF1 (JRH Biosciences), and 8 ng/ml FGF2 (R&D Systems). Cultures were passaged using Collagenase IV and plated on growth factor depleted Matrigel (BD Biosciences) diluted 1:200. These cultures were karyotypically normal.To partially differentiate hESC cultures for immunostaining analysis, karyotypically normal BG01 cells were plated on matrigel and grown for three days in DMEM/F12 containing 10% fetal calf serum (HyClone), 1× nonessential amino acids, 20 mM L-glutamine, 50 U/ml penicillin/streptomycin, and 0.1 mM β-mecaptoethanol. PowerBlot assaysBG01 hESC lysate was prepared in 10 mM Tris-HCl pH 7.4, 1 mM sodium orthovanadate and 1% SDS, and the PowerBlot assays were performed by BD Biosciences (BD Biosciences). Briefly, 200 μg of protein lysate was loaded in a single, gel-wide well, on a SDS-4–15% gradient polyacrylamide gel. The full PowerBlot screen consisted of five gels, which were blotted and probed with 934 antibodies, and was performed in duplicate with the same cell lysate. The gel dimensions were 130 × 100 × 0.5 mm, and proteins were separated at 150 volts for 1.5 hours, and transferred to an Immobilon-P membrane (Millipore). The membranes were blocked and clamped in a manifold that created 40 lanes across each membrane. A mix of 1 to 8 mouse monoclonal primary antibodies was added to each lane, in dilutions and combinations that had been predetermined to enable accurate identification of well-separated proteins. The predicted sizes of detectable proteins in the blots ranged from 10–540 kD, and the dilutions of the primary antibodies ranged from 1:250 to 1:15,000.The blots were removed from the manifolds, washed and incubated with goat anti-mouse secondary antibody conjugated to the Alexa680 fluorophore (Molecular Probes). The membranes were scanned using the Odyssey Imaging System (LI-COR). Molecular weight standards were generated by adding a cocktail of antibodies to P190 (190 kD), Adaptin beta (106 kD), STAT-3 (92 kD), PTP1D (72 kD), Mek-2 (46 kD), RACK-1 (36 kD), GRB-2 (24 kD) and Rap2 (21 kD) to lane 40 of gels A-D. Molecular standards for gel E were generated by adding a cocktail of antibodies to Exportin-1/CRM1 (112 kD), MCM (83 kD), Nucleoporin p62 (62 kD), α-tubulin (55 kD), Actin (42 kD), KNP-1/HES1 (28 kD) and NTF2 (15 kD) to lane 16, and antibodies to p190 (190 kD), Hip1R (120 kD), Transportin (101 kD), Calreticulin (60 kD), Arp3 (50 kD), eIF-6 (27 kD) and Rap2 (21 kD) to lane 17.Bands were detected and raw signal intensity captured automatically using the PDQuest software (Bio-Rad). To normalize the signal intensities, the total raw quantity of each band was divided by the average intensity value of the molecular standards in that image and the normalized values for the duplicate samples were averaged and expressed as normalized intensity units (i.u.). These values represent the relative signal intensity observed for each identified protein band, rather than relative expression levels of different proteins, due to differences in the efficiencies of antibody binding and dilution of the primary antibodies used. Proteins were identified based on the similarity of expected and observed band migration profiles and bands that could not be identified were excluded from the analysis. All identified proteins were verified by visual inspection, and proteins exhibiting a low signal intensity, with an averaged signal of < 1000 i.u., were verified by visual inspection using contrast enhancement in Adobe Photoshop. Bands with > 800 i.u. could typically be observed without additional image enhancement. Microsoft Excel files were generated that contained information on: gel number, lane number, antibody catalogue number (BD Biosciences), protein name, expected size, observed size, repeat 1 i.u. value, repeat 2 i.u. value, averaged i.u. value, antibody dilution, outline of protein function, Entrez gene and SwissProt identification numbers. These tables were used to list expressed proteins (Additional File 1). Kinexus assaysPreparation of the BG03 cell lysate and western blotting was performed according to published protocols [50]. Briefly, cell lysate was prepared in 20 mM MOPS pH 7.0, 2 mM EGTA, 5 mM EDTA, 30 mM sodium fluoride, 40 mM β-glycerolphosphate pH 7.2, 20 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM PMSF, 3 mM benzamidine, 5 μM pepstatin, 10 μM leupeptin, 0.5% nonidet P-40, with the final pH adjusted to 7.2. The Kinexus assays for protein kinases (KPKS-1.2A and B [76 antibodies]), phosphatases (KPPS-1.2 [27 antibodies]) and phosporylated sites in cell signaling molecules (KPSS-3.1 [37 antibodies]) were performed by Kinexus. The Bio-Rad Mini-PROTEAN 3 electrophoresis system was used to separate proteins by SDS-PAGE. For each assay, 250 μg of cell lysate was loaded in a single well spanning the width of the stacking gel, then separated through a 12.5% SDS-Polyacrylamide gel and transferred to a PVDF membrane. A 20-lane manifold was placed over the membrane and a different mixture of up to 3 primary antibodies was added to each well. The combinations of primary antibodies had been predetermined to detect well-separated proteins, avoiding crossreaction to different proteins that co-migrate. The primary antibodies were rabbit and goat polyclonal, and mouse monoclonal antibodies, diluted 1:1000. After incubation with the primary antibodies, the membranes were removed from the manifolds, washed and incubated with a mix of the appropriate secondary antibodies. The secondary antibodies were donkey anti-rabbit (at 1:5000), sheep anti-mouse (at 1:10,000) and bovine anti-goat (at 1:10,000), all conjugated with horse radish peroxidase. The membranes were washed and immunoreactive bands detected by enhanced chemiluminescence (Amersham-Pharmacia) using a FluorS Max Multi-imager (Bio-Rad). Prestained size markers (201.5, 156.8, 106, 79.7, 48.4, 37.8, 23.3, and 18.2 kD) and predetermined human-specific protein migration profiles were used to accurately identify proteins using the Kinexus immuno-reactivity identification system (IRIS) software. Detected proteins were verified by visual inspection. ImmunocytochemistryImmunocytochemistry and staining procedures were as described previously [51]. Briefly, cells were fixed with 4% paraformaldehyde for half an hour, blocked in blocking buffer (5% goat serum, 1% BSA, 0.1% Triton X-100) for 1 hour followed by incubation with the primary antibody at 4°C overnight. Appropriately coupled secondary antibodies (Molecular Probes) were used for single and double labeling. All secondary antibodies were tested for cross reactivity and non-specific immunoreactivity. The following antibodies were used: ABP-280 (1:250, BD Biosciences 610798), CtBP1 (1:1000, BD Biosciences 612042), CtBP2 (1:1000, BD Biosciences 612044), GS-28 (1:2000, BD Biosciences 611184), HDJ-2 (1:100, BD Biosciences 611872), L-Caldesmon (1:2000, BD Biosciences 610660), Rabaptin-5 (1:500, BD Biosciences 611080), phospho-p130 Cas (Tyr165) (1:50, Cell Signaling Technology 4015), phospho-Ras-GAP (pY460) (1:250, BD Biosciences 612736), Ras-GAP (1:250, BD Biosciences 610043), Shc-C (1:1000, BD Biosciences 610642), Oct-4 (Santa Cruz biotechnology, 1:200 SC-8628), TNIK (1:100, BD Biosciences, 612250), p130 Cas (1:100, BD Biosciences, 610272), ERBB2 (1:100, Lab Vision, 9G6.10), ERBB3 (1:100, R&D Systems, MAB348), ZO1 (1:100, Invitrogen, 61–7300), or Occludin (1:100, Invitrogen, 71–1500). Hoechst (Invitrogen) or DAPI (Sigma) were used to identify nuclei, and Triton X-100 was omitted when staining for extracellular antigens (ZO1, occludin, ERBB2/3). Images were captured on an Olympus or Nikon fluorescence microscope. lllumina data and comparison to proteomic databaseExpression levels of proteins detected by the PowerBlot assay were compared to our previous published database of multiple hESC lines examined using the Illumina bead array platform (Liu et al., 2006). Averaged transcript expression signals from the BG01, BG02 and BG03 cell lines were converted to a +/- format, based on the following criteria: A mean transcript detection level of > 5,000 was designated as ++++; 1,000–5,000 as +++; 100–1,000 as ++; 30–100 as +; and signals < 30 was represented as -. In parallel, the protein expression levels were converted to a +/- format based on these criteria: i.u. > 10,000 as ++++; 5,000–10,000 as +++; 1,000–5,000 as ++; 100–1,000 as +. In addition, genes were categorized into the same functional/signaling pathways as per the western blot database.\n" ], "offsets": [ [ 0, 44733 ] ] } ]

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[ { "id": "18286199__text", "type": "full_text", "text": [ "BackgroundHuman embryonic stem cells (hESCs) offer a virtually unlimited source of neural cells for structural repair in neurological disorders, such as stroke. Neural cells can be derived from hESCs either by direct enrichment, or by isolating specific growth factor-responsive and expandable populations of human neural stem cells (hNSCs). Studies have indicated that the direct enrichment method generates a heterogeneous population of cells that may contain residual undifferentiated stem cells that could lead to tumor formation in vivo.Methods/Principal FindingsWe isolated an expandable and homogenous population of hNSCs (named SD56) from hESCs using a defined media supplemented with epidermal growth factor (EGF), basic fibroblast growth factor (bFGF) and leukemia inhibitory growth factor (LIF). These hNSCs grew as an adherent monolayer culture. They were fully neuralized and uniformly expressed molecular features of NSCs, including nestin, vimentin and radial glial markers. These hNSCs did not express the pluripotency markers Oct4 or Nanog, nor did they express markers for the mesoderm or endoderm lineages. The self-renewal property of the hNSCs was characterized by a predominant symmetrical mode of cell division. The SD56 hNSCs differentiated into neurons, astrocytes and oligodendrocytes throughout multiple passages in vitro, as well as after transplantation. Together, these criteria confirm the definitive NSC identity of the SD56 cell line. Importantly, they exhibited no chromosome abnormalities and did not form tumors after implantation into rat ischemic brains and into naïve nude rat brains and flanks. Furthermore, hNSCs isolated under these conditions migrated toward the ischemia-injured adult brain parenchyma and improved the independent use of the stroke-impaired forelimb two months post-transplantation.Conclusions/SignificanceThe SD56 human neural stem cells derived under the reported conditions are stable, do not form tumors in vivo and enable functional recovery after stroke. These properties indicate that this hNSC line may offer a renewable, homogenous source of neural cells that will be valuable for basic and translational research.\nTo date there have been no effective treatments for improving residual structural and functional deficits resulting from stroke. Current therapeutic approaches, such as the use of thrombolytics, benefit only 1 to 4% of patients [1]. Consequently, the majority of stroke patients experience progression of ischemia associated with debilitating neurological deficits. Recent evidence has suggested that the transplantation of cells derived from cord blood, bone marrow or brain tissue (fetal and adult) enhances sensorimotor function in experimental models of stroke [2], [3]. However, the normal human-derived somatic stem cells used in these studies have a limited capacity to differentiate into the diverse neural cell types optimal for structural and physiological tissue repair and are not amenable for large-scale cell production. Unlike other sources of stem cells, hESC lines possess a nearly unlimited self-renewal capacity and the developmental potential to differentiate into virtually any cell type of the organism. As such, they constitute an ideal source of cells for regenerative medicine. The successful derivation of hESC lines from the inner cell mass of preimplantation embryos and their long-term maintenance in vitro over multiple passages has been demonstrated [4] and standardized. Differentiation and enrichment processes that direct hESCs towards a neural lineage have also been achieved. To promote neuralization, ESCs were cultured in a defined media supplemented with morphogens or growth factors [5], [6], [7] or cultured under conditions that promote “rosettes”, structures morphologically similar to the developing neural tube [8], [9]. This neuralization process has proven invaluable in understanding the specification of hESC-derived neural tissue [10], [11], [12]. However, the enriched neural progeny derived displayed overgrowth and limited migration after grafting into normal newborn mice [13] and lesioned adult rat striatum [12], [14], [15], [16]. The inner cores of these grafts contained tumorigenic precursor cells (reviewed in [17]). These findings suggest that neural cells generated by acute exposure to growth factors and/or morphogens may still be heterogeneous and potentially tumorigenic. We report an alternative method for the isolation and the perpetuation of a multipotent hNSC line from the hESCs with a primitive mode of self-renewal. We also demonstrate their long-term expansion, non-tumorigenic properties and functional engraftability in an experimental model of stroke.\n1. In vitro isolation, growth and differentiation of hESC-derived hNSCsThe hESCs were maintained and expanded on mouse feeder layer in media supplemented with bFGF (Figure 1A). After cell dissociation, a portion of the hESCs was cultured in serum free medium containing EGF, bFGF and LIF. These factors are known to stimulate the proliferation of human fetal-derived NSCs [18], [19]. After 3 days in vitro (DIV), there was selective survival and growth of cells that aggregated in clusters or spheres (Figure 1B). These primary spheres were harvested and replated in fresh media. During the following week, the spheres attached to the flask and a fibroblast-like cell population began to migrate out (Figure 1C). Secondary spheres (2° spheres) were generated from these cultures and lifted off by the end of the week leaving a hollow in the middle of the attached cells (Figure 1D). The floating 2° spheres were collected and replated in fresh growth medium for 2 weeks. The cultures were then passaged by collagenase cell dissociation every 7 DIV for an additional 4 passages (Figure S1). At the 5th and 6th passages, spheres were dissociated into a single-cell suspension using trypsin-EDTA. At this stage there was a change in the hNSCs' adherent properties, and the cells began to grow as a monolayer with multiple foci of cells throughout the culture (Figure 1F). The adherent hNSC culture stained uniformly for nestin (Figure 1K), vimentin (Figure 1L) and with the radial glial marker 3CB2 (Figure 1M) indicating their homogeneity and NSC identity. Under these culture conditions, it is noteworthy that we did not observe the formation of rosettes which has been previously reported to occur under certain conditions during neuralization of hESCs [8], [20], [21]. RT-PCR analysis confirmed that these hNSCs did not express the pluripotency transcripts Oct-4 and Nanog (Figure 1I). Furthermore, the hNSCs did not express transcripts for brachyury and foxa2, marker genes for mesoderm and endoderm respectively (negative result, data not shown).10.1371/journal.pone.0001644.g001Figure 1Isolation and purification of hNSCs from the hESCs.The hESCs were grown on a mouse feeder layer (A). Primary neurospheres (B) were isolated and replated to eliminate other non-neural cells. The selectively harvested secondary neurospheres (arrow in C), left behind hollow cores in the surface area (star in D) where they attached earlier. They were perpetuated for an additional 5 passages (E). These 2° spheres were then passaged using a single cell dissociation protocol (F). Arrow in F shows an example of a focus of proliferating cells. (G, H) The hNSCs were passaged every 5–7 days, as described in the Methods section. Starting from an initial population of 1 million cells, the cumulative cell number was calculated at each passage as the fold of increase×the total cell number and plotted as logarithm with base 2 in function of time (G). The cell perpetuation (G) and population doubling (H) analysis demonstrated the continuous and stable growth of the hNSCs. (I) RT-PCR analysis showing the down regulation of the pluripotency transcripts Oct4 and Nanog in secondary neurospheres and in expanded hNSCs at passage 8 (P8). (J) Cytogenetic evaluation of the SD56 hNSCs line at passage 12 by standard G-banding was performed. Twenty metaphase cells were analyzed and showed a normal female chromosome complement (46,XX). Isolated and expanded hNSCs expressed the neural precursor cell markers nestin (K), Vimentin (L) and the radial glial cell marker 3CB2 (M) in virtually all the progeny. (N-P) Clonal self-renewal ability of the isolated hNSCs through symmetrical cell division. Single (N), two-cell stage (O) and four-cell stage (P) of a hNSC proliferating over a 3-day culture period. Note the symmetrical segregation of BrdU and nestin in the progeny. Bars: (A, B, C, D) 200 µm; (E, F) 100 µm; (K–M) 20 µm; (N–P) 10 µm.To ascertain self-renewal ability under clonal conditions, a single cell suspension was plated at clonal density (1–2 cell/10 µl). To determine if the hNSCs divide symmetrically, we pulsed cultures with the thymidine analog, bromodeoxyuridine (BrdU), after plating and looked for the expression of nestin in the progeny. Cultures were fixed after 1, 2 or 3 DIV (Figure 1N–P). After 2 days, plated single cells first underwent a symmetric cell division and gave rise to daughter cells that were both positive for BrdU and nestin. The clone of cells continued to grow over the 3 DIV and all the progeny expressed nestin. BrdU labeling demonstrated that it was rare for only one daughter cell to inherit the BrdU and thus had undergone asymmetric segregation of the chromatids (Figure S2). G-band karyotyping of these hNSCs confirmed the normal, non-transformed nature of cells after 12 passages (Figure 1J). We named the derived hNSCs SD56 (intermittently referred to as SD56 hNSCs or hNSCs).Under these defined growth conditions, the hNSCs showed stable growth with a 2.7±0.2 fold increase every 5 to 7 days (Figure 1G). The population doubling at each passage averaged at 1.4±0.1 (Figure 1H). The viability of hNSCs at each passage was consistent at the approximate value of 98%. The projected cumulative cell numbers demonstrated that trillions of cells could be generated over a period of 5 months (Figure 1G). We expanded the isolated hNSCs lines for over 20 passages with a stable phenotype. An initial cell bank of 75 vials containing 2 to 5 million cells each was generated and cryopreserved.Upon removal of the mitogenic factors and plating on a coverslip pre-coated with poly-L-ornithine (PLO) substrate, the hNSCs spontaneously differentiated into neurons, astrocytes and oligodendrocytes, a property that is consistent with normal multipotent hNSCs (Figure 2). After 2 DIV, hNSCs expressed transcripts for the neural-specific genes nestin, Notch1 and neural cell adhesion molecule (N-CAM) (Figure 2A) and for the lineage specific markers β-tubulin class III, medium-size neurofilament (NF-M) and microtubule-associated protein 2 (MAP-2) for neurons, GFAP for astrocytes and myelin basic protein (MBP) for oligodendrocytes (Figure 2A). Transcripts for the GABAergic cell marker glutamic acid decarboxylase (GAD) were expressed, but transcripts for the tyrosine hydroxylase (TH), a marker for dopaminergic neurons, were undetectable. Immunocytochemical analysis (Figure 2B–F) of 10 day-old cultures demonstrated that the proportion of nestin+ cells was 36.6±2.7%, 62.5±2.8% expressed the neuronal marker TuJ1, 1.9±0.3% expressed the astrocytic marker GFAP and 7.1±0.4% were oligodendrocytes and expressed galactocerebrocide (GC) (Figure 2F). A subset (9.8±1.6%) of the GFAP+ astrocytes co-expressed nestin.10.1371/journal.pone.0001644.g002Figure 2hESC-derived hNSCs spontaneously differentiated into the 3 principal central nervous system cell types.Dissociated hNSCs were washed free of growth factors and plated on poly-L-onithine coated glass coverslips. Differentiated cultures were either harvested after 2 DIV for total RNA extraction and RT-PCR analysis or fixed after 10 DIV and processed for indirect immunocytochemistry. (A) Differentiated hNSCs expressed the neural-specific transcripts nestin and Notch1 as well as transcripts: for neurons (β-tubulin class III, MAP-2, NCAM and medium-size neurofilament, NF-M), for astrocytes (GFAP) and for oligodendrocytes (MBP). The hNSCs expressed transcripts for GAD, but not for TH. B, C & D, morphology of NSC-derived progeny differentiated into GFAP+ astrocytes (B), GC-expressing oligodendrocytes (C) and TuJ1+ neurons (D), DAPI (blue) show life cell nuclei. (E) Photo showing cultures double-immunostained for TuJ1 (green) and nestin (red) (DAPI, blue). (F) Quantitative analysis of immunostained 10 day-old cultures for the 3 neural cell types. Results are mean±s.e.m. of three independent experiments, each performed in duplicate. Bars: (c) 20 µm; (d, e) 10 µm. 2. The isolated hNSCs are normal and do not form tumors in normal nude ratsThe self-renewal and pluripotent abilities of the hESCs are also associated with tumorigenic properties. Therefore, the first critical step toward developing therapeutic hNSCs is to verify that they are non-tumorigenic. The monolayer culture of SD56 hNSCs clearly demonstrated contact inhibition of growth, a normal karyotype and did not express the pluripotency transcripts Oct-4 and Nanog. Removal of mitogenic factors and continued culture on plastic resulted in cell senescence that is characteristic of non-transformed cells. To determine whether SD56 hNSCs form tumors in vivo, we transplanted them at high density (see Methods) into the forebrain and subcutaneously into the flank of nude rats. The animals were kept for a two-month post-transplant survival period. To label mitotically active cells in vivo during S-phase, the rats were injected IP with BrdU (50 mg/kg) every 8 hours during the last 24 hours before euthanasia. The transit amplifying endogenous precursors located in the subventricular zone (SVZ) were labeled (Figure S3); however, we were unable to detect grafted SD56 hNSCs co-localizing the human-specific nuclear marker hNuc and BrdU (Figure S3). No surviving SD56 hNSCs were detected in the flank of the transplanted animals suggesting that the grafted cells are not tumorigenic. 3. Transplanted cells survived, migrated toward and engrafted into the stroke-damaged host tissueTo investigate the survival and functional engraftment in an injury environment, hNSCs (4×105) were transplanted into the ischemic boundary zone in the rat striatum one week after the middle cerebral artery occlusion (MCAO) was performed. Animals were euthanized two months later and the brains processed for histo-pathology and immunocytochemistry. Grafted SD56 hNSCs, identified with hNuc, demonstrated a 37.0±15.8% survival rate and a remarkable dispersion toward the stroke-damaged tissue with no sign of overgrowth or tumorigenesis. The majority of grafted cells (61.2±4.7%) migrated at least 200 µm away from the injection site and penetrated an average distance of 806.4±49.3 µm into the stroke-damaged tissue (Figure 3A–C). Immunostaining with the blood vessel marker, GluT1, revealed dilated vessels in the infarcted striatum and a close association between vessels and the grafted hNSCs (Figure 3B, 3C). The grafted cells rarely expressed the proliferation marker Ki67 (Figure 3D), 29.8±4.4% expressed nestin (Figure 3E), 6.5±0.9% expressed doublecortin (DCX) and 60.8±8.1% were TuJ1+ (Figure 3F, G). Grafted cells rarely co-expressed the astroglial marker GFAP (Figure 3H) or differentiated into CNPase-expressing oligodendrocytes (Figure 3I). Immunostaining for GAD demonstrated that 25.1±2.3% of grafted hNSCs differentiated into GABAergic neurons while less than 2% were positive for glutamate (Figure 3J, K).10.1371/journal.pone.0001644.g003Figure 3Dispersion, engraftment and differentiation of the hNSCs in stroke-lesioned animals.(A) Schematic drawing of a frontal section through the striatum illustrating the dispersion of grafted hNSCs in the focal ischemia-lesioned parenchyma (shaded area). (B, C) Photos show frontal sections through the graft in the striatum immunostained with the human specific antibodies: anti-hNuc (green in B & C) and anti-GluT1 (red, B & C) showing blood vessels and dispersed hNSCs in the graft zone. C: higher magnification of the inset in B. (D–I) Photos taken from frontal sections through the graft in the striatum double immunoprocessed for cell proliferation and neural lineage markers. (D) Note the endogenous Ki67+ cells (red cells, arrow) in the stroke damaged area and the hNuc+ (green)/Ki67- grafted hNSCs (arrowheads). (E) Examples of grafted SD56 hNSCs showing co-expression of hNuc (green) and nestin (red). (F) Confocal 3D reconstructed orthogonal images of the hNuc+(green)/DCX+(red) NSCs (arrowheads) viewed in the x-z plan on the top and y-z plan on the right. (G) Examples show the majority of grafted NSC progeny co-expressing hNuc (red) and the neuronal marker TuJ1 (green). Grafted NSCs rarely differentiate into GFAP+ astrocytes (H). In I, rare example of grafted NSC progeny becoming an oligodendrocyte identified by the expression of CNPase (green). Grafted NSCs expressed the GABAergic marker GAD65/67 (J) and rarely expressed glutamate (K). (Abbreviations: Cx: cortex, Str: striatum). Bars: (B, C) 100 µm; (D, F) 20 µm; (E, G–K) 10 µm. 4. Transplanted cells improve sensorimotor function of the stroke-disabled forelimbWe asked whether transplanted SD56 hNSCs could enhance the recovery of sensorimotor function that is compromised in the stroke-injured rats. We used the cylinder test to measure the sensorimotor asymmetry in forelimb use during spontaneous exploration [22]. To establish the baseline of the stroke-induced sensorimotor deficit, spontaneous behavior of rats in a transparent cylinder was videotaped one week after stroke (pre-transplant, Figure 4). Tests were then conducted 4 and 8 weeks after vehicle and SD56 hNSCs transplantation. Stable asymmetry in forelimb use was observed 7 days post-stroke (pre-transplant, Figure 4). Ischemic rats used their impaired forelimbs (contralateral to lesion) during lateral exploration less than they did before stroke. Transplantation of SD56 hNSCs significantly enhanced the independent use of the impaired contralateral forelimb 4 weeks post transplantation (P<0.05 vs pre-transplant). Eight weeks after transplantation the improvement in the use of the impaired forelimb was stable and significant when compared to the pre-transplant group and significantly improved in comparison to vehicle treated group at 8 weeks (Figure 4). In the vehicle treated group, the independent use of the contralateral forelimb remained impaired 4 and 8 weeks post-injection. In an independent study and using the same MCAO rat animal model, we found that transplantation of dermal fibroblasts did not improve the stroke-induced motor deficits (unpublished data).10.1371/journal.pone.0001644.g004Figure 4Transplantation of NSCs improves sensorimotor function of the stroke-disabled forelimb.Forelimb use during spontaneous lateral exploration was measured in the cylinder test (see Method and Results sections for details). Groups of vehicle injected (n = 7) and hNSCs (n = 10) transplanted are represented. The animals were tested as described in Method section. Note the significant increase in the independent use of the impaired contralateral forelimb at 4 and 8 weeks post transplantation (P<0.05 vs pre-transplant group). The contralateral forelimb remained impaired in the vehicle treated group at 4 and 8 weeks post-injection. Bars represent percentages±s.e.m. of steps taken by the ipsilateral, contralateral and both forelimbs simultaneously. *P<0.05 vs pre-transplant group; #P<0.05 vs vehicle groups.\nOur results indicate that a self-renewable and homogenous population of hNSCs, SD56, was derived from hESCs using defined media supplemented with a specific combination of growth factors. The SD56 hNSCs grew as an adherent monolayer culture, uniformly expressed molecular features of hNSCs including nestin, vimentin and the radial glial marker 3CB2, and did not express the pluripotency markers Oct4 or Nanog. The self-renewal property of the hNSCs was characterized by a predominant symmetrical mode of cell division. They exhibited no chromosomal abnormalities and demonstrated non-tumorigenic properties after implantation into ischemic brains and into naïve nude rat brains and flanks. Furthermore, the transplanted SD56 hNSCs migrated toward the stroke-damaged adult brain parenchyma, engrafted and improved the independent use of the stroke-impaired forelimb. Maintenance of stem cells requires symmetrical and asymmetrical cell divisions to both expand and to give rise to specialized progeny of a specific tissue (reviewed in [23]). In vivo, a complex cellular micro-environment or niche ensures the self-maintenance property of NSCs [24], [25], [26], [27]. In vitro, defined growth factors and extracellular matrices support stem cell self-renewal [28], [29]. The embryonic stem cells can propagate in a predominantly proliferative symmetrical mode, leading to homogeneous cell cultures growing relatively quickly with minimal cell differentiation [30], [31], [32], [33], [34]. Tissue specific stem cells, however, self-renew in a predominant asymmetric mode to maintain them selves and compensate for the loss of differentiated cells due to disease or injury. Thus, NSCs isolated from developing or adult brain grow as a mixture of undifferentiated and differentiated cells due the predominant asymmetrical mode of cell division [35], [36], [37], [38], [39], [40], [41]. A recent study has reported that a murine ESC-derived NSC line (LC1) is propagated as an adherent homogenous culture with a dominant mode of symmetrical self-renewal [21]. A combination of EGF and FGF2 was sufficient to propagate these NSCs as an adherent monolayer. However, the SD56 hNSC line described here required the combination of EGF, bFGF and LIF for self-maintenance. Although there are morphological and molecular similarities between our hNSCs and the NSCs previously described [21], the methods of isolation and growth are different. In addition to the different combination of growth factors used, the hNSC line we have isolated did not go through the rosette-structure stage. The in vitro analysis of BrdU incorporation and nestin expression indicated that our hNSCs divide predominantly symmetrically. This type of growth pattern is characteristic of primitive normal stem cells undergoing mostly symmetric cell division to increase the stem cell pool at the early stage of development or during tissue regeneration after injury [23]. RT-PCR and immunocytochemistry analysis demonstrated that these undifferentiated SD56 cells did not express any pluripotency, endodermal or mesodermal markers. Furthermore, the SD56 hNSCs described here exhibited the multipotential characteristic to differentiate into neurons, astrocytes and oligodendrocytes both in vitro and upon transplantation. Together these findings suggest that the hNSC line we isolated are appropriately programmed and share similar characteristics with the definitive NSCs of the developing brain. The SD56 hNSCs demonstrated a remarkable ability to migrate toward the stroke-damaged parenchyma of the adult rat brain. This directed migration by the majority of the grafted cells could be due to their uniform cellular composition, which results in an equal response to the host microenvironment. In previous studies, subpopulations of transplanted hESCs that were enriched in neural cells migrated in host microenvironments conducive to cell migration, such as the developing brain or in structures such as the rostral migratory stream [13], [20]. In the adult lesioned brain, the grafted hESC-derived neural cells proliferated and formed rosettes [14], teratomas [12], [15] or a cellular mass that induced a gliotic host response whereby local astrocytes demarcated the grafts [16]. Enriched neural cultures derived from mouse [42] and monkey ESCs [43] have produced behavioral improvements when transplanted into animal models of stroke and brain injury. However, in these cases, the transplanted non-human ESCs also formed a mass with signs of overgrowth in the core, as well as deformations [44], [45], [46]. ESCs plated at low density acquire neural identity within few hours after plating [47]. Interestingly, nearly all viable cells expressed nestin, the early neural fate marker Sox1, and the pluripotency marker Oct4. Together, these studies are seminal and suggest that complete neuralization may not be achieved through certain enrichment processes, consequently the neural cells could revert to a pluripotent stage [17]. The dispersion of the grafted hNSCs within host parenchyma may allow for more graft-host interactions that could stabilize differentiation, inhibit growth and prevent gliotic host response. In the present study, SD56 hNSC-transplanted animals demonstrated a stable improvement in the sensorimotor function when evaluated for spontaneous exploratory activity in the cylinder test that detects long-lasting deficits in forelimb use in the experimental models of stroke [22]. The transplantation of hNSCs significantly enhanced the independent use of the impaired contralateral forelimb 8 weeks post transplantation. This is the first report demonstrating that the transplantation of hNSCs derived from hESCs can improve neurologic behavior after experimental stroke. Together, these findings are encouraging and suggest that these cells are promising for future development. In addition to their therapeutic application, the hNSCs isolated under the reported conditions offer a means to interrogate host environments and unravel mechanistic features of self-renewal, non-tumorigenicity and functional engraftability in animal models of neurological disorders.\nhESC and NSC CulturesThe hESC line H9 (WiCell Research Institute) was propagated every 5 to 7 days on irradiated mouse embryonic fibroblasts. The cell culture media consisted of a 1∶1 mixture of Dulbecco's modified Eagle's medium (DMEM) and F12 nutrient, 20% serum replacement (Invitrogen), 0.1 mM β-mercaptoethanol, 2 µg/ml heparin and 4 ng/ml bFGF (R&D Systems). To generate the NSCs, dissociated hESCs were cultured in a chemically defined medium composed of DMEM/F12 (1∶1) including glucose (0.6%), glutamine (2 mM), sodium bicarbonate (3 mM), and HEPES buffer (5 mM) [all from Sigma except glutamine (Invitrogen)]. A defined hormone mix and salt mixture (Sigma), including insulin (25 mg/ml), transferrin (100 mg/ml), progesterone (20 nM), putrescine (60 mM), and selenium chloride (30 nM) was used in place of serum. The medium was supplemented with EGF (20 ng/ml), bFGF (10 ng/ml) and LIF (10 ng/ml). Dissociated hNSCs were plated at a density of 100,000 cell/ml in Corning T75 (Invitrogen) culture flasks in the defined media together with the growth factors. After 5–7 DIV, the adherent culture was incubated in 0.025%trypsin/0.01% EDTA (w/v) for 1 min followed by the addition of trypsin inhibitor (Invitrogen) then gently triturated to achieve single cell suspension. The cells were then washed twice with fresh medium and reseeded in fresh growth factor-containing media at 100,000 cells/ml. This procedure was performed for 21 passages and the fold of increase and population doubling were calculated at each passage. For clonal analysis, single spheres or confluent hNSC cultures were single cell dissociated and serially diluted to yield 1–2 cell/10 µl. A 10-µl-cell suspension was then added to each of 96 or 24 well plates containing 200–300 µl of growth media. Wells containing one viable cell were marked the next day and re-scored 5 to 7 days later for cell proliferation. The differentiation of the hNSCs was performed as previously described [48]. Dissociated hNSCs were plated at a density of 106 cells/ml in control (media/hormone mix) medium devoid of any growth factors and supplemented with 1% fetal bovine serum (FBS) on poly-L-ornithine-coated (15 mg/ml; Sigma) glass coverslips in 24-well Nunclon culture dishes with 0.5 ml/well. After 2, 7–15 DIV cultures were fixed and processed for immunocytochemistry or used for RT-PCR analysis. Karyotype analysisLong-term cultures of hNSCs were incubated at 37°C and harvested for metaphase chromosomes when the cultures were 75% confluent. Metaphase chromosomes were obtained by standard chromosome harvest methods by exposure to Colcemid at 0.1 µg/ml for 1 hour at 37°C, a 2-minute exposure to trypsin/EDTA, hypotonized with 0.057 M KCl and fixed with 3∶1 methanol:acetic acid. Slide preparations were made by dropping the fixed cell pellet onto cold, wet slides and air-dried. After incubating the slides at 90°C for 30 minutes, chromosomes were trypsin banded and then Wright/Giemsa stained for G-banding analysis. Twenty metaphase cells were completely analyzed and a normal female chromosome complement was found (46,XX). Tumorigenicity in nude ratsAll animal experiments were conducted according to the National Institute of Health (NIH) guidelines and approved by the IACUC. Normal adult NIH-Nude rats (n = 5, 8 week-old, Taconic, Germantown, New York, United States) were used to test the tumorigenic potential of the SD56 hNSCs. Undifferentiated hNSCs from passage 9 were single cell dissociated using trypsin-EDTA and suspended at the concentration of 125,000 cell/µl in preparation for cell transplantation. Two µl of the cell suspension were stereotaxically transplanted into 4 sites within the striatum at the following coordinates: AP: +1.0 mm, ML: +3.2 mm, DV: −5.0; AP: +0.5 mm, ML: +3.0 mm, DV: −5.0; AP: −0.5 mm, ML: +3.0 mm, DV: −5.0; AP: −1.0 mm, ML: +3.5 mm, DV: −5.0 mm with the incisor bar set at 3.4 mm. The injection rate was 1 µl/min, and the cannula was left in place for an additional 5 min before retraction. For the flank tumor assay, 2×106 cells (125,000 cell/µl) were injected subcutaneously to the side of the adult nude rats. To label mitotically active cells in vivo during S-phase, the rats were injected IP with the BrdU (50 mg/kg, Sigma) every 8 hours during the last 24 hours before euthanasia. After 2-month survival time, rats were euthanized and a postmortem examination for tumor formation was performed. Induction of Focal Ischemia and Cell TransplantationAll animal experimentations were conducted according to the National Institute of Health (NIH) guidelines and approved by the IACUC. Sprague Dawley adult male rats (n = 17, 275g–310g, Charles River Laboratories, Wilmington, Massachusetts, United States) were subjected to one and a half hour suture occlusion of the middle cerebral artery (MCAO), as previously described [49] and immunosuppressed 2 days before cell transplantation and daily thereafter for one week with i.p. injections of cyclosporine A (20 mg/ml, Sandimmune, Novartis Pharmaceuticals). Thereafter oral cyclosporine was used at 210 µg/ml in drinking water until euthanasia. Undifferentiated SD56 hNSCs from passages between P9 and P13 were single cell dissociated using trypsin-EDTA in preparation for cell transplantation. One week after the stroke lesion, 2 µl of the hNSCs, at a concentration of 50,000 cell/µl, were stereotaxically transplanted into 4 sites within the lesioned striatum (n = 10) at the following coordinates: AP: +1.0 mm, ML: +3.2 mm, DV: −5.0; AP: +0.5 mm, ML: +3.0 mm, DV: −5.0; AP: −0.5 mm, ML: +3.0 mm, DV: −5.0; AP: −1.0 mm, ML: +3.5 mm, DV: −5.0 mm with the incisor bar set at 3.4 mm. The injection rate was 1 µl/min, and the cannula was left in place for an additional 5 min before retraction. As a control group, we used rats subjected to ischemia and injected with the vehicle (n = 7). All animals underwent baseline motor behavioral assessment before and after the ischemic lesion, and 4 & 8 weeks after cell transplantation. The animals were killed after 2-month survival time by transcardial perfusion with phosphate buffered saline (PBS) followed by 4% paraformaldehyde. The brains were cryoprotected in an increasing gradient of 10, 20 and 30% sucrose solution and cryostat sectioned at 40 µm and processed for immunocytochemistry. ImmunocytochemistryCultures were fixed with 4% paraformaldehyde for 15 min. Both cells and brain sections were rinsed in PBS for 3×5 min then incubated for 2 hrs (cultures) or overnight (brain sections) with the appropriate primary antibodies for multiple labeling. Secondary antibodies raised in the appropriate hosts and conjugated to FITC, RITC, AMCA, CY3 or CY5 chromogenes (Jackson ImmunoResearch) were used. Cells and sections were counterstained with the nuclear marker 4′,6-diamidine-2′-phenylindole dihydrochloride (DAPI). Positive and negative controls were included in each run. Immunostained sections were coverslipped using fluorsave (Calbiochem) as the mounting medium. The following antibodies were used: Anti-human Nuclei (hNuc, monoclonal 1∶100, Chemicon), Anti-TuJ1 (monoclonal 1∶100, Covance; Polyclonal 1∶200, Aves Labs); anti-GAD65/67 (polyclonal 1∶1000, Chemicon); Anti-glial fibrillary acidic protein (GFAP, monoclonal 1∶1000, Chemicon; polyclonal 1∶200, Aves Labs); Anti-galactocerebrocide (GC, monoclonal 1∶250, Chemicon); Anti-CNPase (polyclonal 1∶200, Aves Labs); Anti-Glucose Transporter type 1 (Glut-1 polyclonal, 1∶500, Chemicon); Anti-Nestin (polyclonal 1∶1000, Chemicon); Anti-vimentin (monoclonal 1∶500, Calbiochem); Anti-3CB2 (monoclonal 1∶500, Developmental Studies Hybridoma Bank); Anti-doublecortin (DCX, polyclonal 1∶250, SantaCruz Biotechnology); Anti-Ki67 (polyclonal 1∶250, SantaCruz Biotechnology). Fluorescence was detected, analyzed and photographed with a Zeiss LSM550 laser scanning confocal photomicroscope. For each animal, quantitative estimates of the total number of grafted cells were stereologically determined using the optical fractionator procedure [50]. A computer-assisted image analysis system was performed using Stereo Investigator software (MicroBrightField, Inc.). The rostral and caudal limits of the reference volume were determined by first and last frontal sections containing grafted cells. The striatum and cortex were accurately outlined at low magnification (2.5× objective). The optical fractionator probe was selected to perform systematic sampling of the immunoreactive cell population distributed within the serial sections to estimate the population number in the volume of tissue. The counting frame of the optical fractionator was defined at 50×50 µm squares and the systematic sampling was performed by translating a grid with 200×200 µm squares onto the sections of interest using the Stereo Investigator software. The sample sites were systematically and automatically generated by the computer and examined using a 60× objective of a Nikon Eclipse TE 300 microscope. The counting frame displayed inclusion and exclusion lines and only immunoreactive cell bodies falling within the counting frame with no contact with the exclusion lines were counted. The cell dispersion was measured by counting the number of cells within 200 µm distance from the graft site. The number and distance in µm of cells dispersed beyond 200 µm was also measured. An average of 2,000 cells was counted per animal. Double labeling was determined using the confocal laser scanning microscope by random sampling of 100 or more cells per marker for each animal, scoring first for hNuc+, followed by DAPI+ nuclei and then the marker of choice. The double labeling was always confirmed in x-z and y-z cross-sections produced by the orthogonal projections of z-series. Reverse Transcription-Polymerase Chain Reaction (RT-PCR) analysisTotal RNA was extracted from cultured cells using RNAeasy kit (Quiagen). Aliquots (1 µg) of total RNA from the cells were reverse transcribed in the presence of 50 mM Tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2, 10 mM DTT, 0.5 µM dNTPs, and 0.5 µg oligo-dT(12–18) with 200 U Superscript RNase H-Reverse Transcriptase (Invitrogen). PCR amplification was performed using standard procedure with Taq Polymerase. Aliquots of cDNA equivalent to 50 ng of total RNA were amplified in 25 µl reactions containing 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 1.5 mM MgCl2 , 50 pmol of each primer, 400 µM dNTPs, and 0.5 U AmpliTaq DNA polymerase (Perkin-Elmer). PCR was performed using the following thermal profile: 4 min at 94°C; 1 min at 94°C, 1 min at 60°C, 1.5 min at 72°C, for 30–40 cycles; 7 min at 72°C, and finally a soak at 4°C overnight. The following day, 10 µl aliquots of the amplified products were run on a 2% agarose Tris–acetate gel containing 0.5 mg/ml ethidium bromide. The products were visualized through a UV transilluminator, captured in a digital format using Quantify One Gel Analysis software (Bio-Rad Laboratories) on a Macintosh G4 computer.The PCR primers specific to each transcript were as follows: GFAP, forward (F), 5′-TCATCGCTCAGGAGGTCCTT–3′ Reverse (R), 5′-CTG TTGCCAGAGATGGAGGTT–3′; MAP2 (F) 5′-GAAGACTCGCATCCGAATGG–3′, (R) 5′-CGCAGGATAGGAGGAAGAGACT–3′; MBP (F) 5′-TTAGCTGAATTC GCGTGTGG–3′, (R) 5′-GAGGAAGTGAATGAGCCGGTTA-3′ were deigned using the Primer Designer software, Version 2.0 (Scientific and Educational Software) [48]. 18S, β-tubulin class III, N-CAM, Nestin, NF-M, Notch-1 primers [51]. Oct4, Nanog primers [11]. FOXa2 (HNF3B), Brachyury primers [52]. Behavioral testsThe cylinder test was used to assess the spontaneous forelimb use during lateral exploration movement [22]. Rats were placed in a transparent acrylic cylinder (20 cm diameter) for 5 minutes. The cylinder encourages use of the forelimbs for vertical exploration. A mirror was placed behind the cylinder so that the forelimbs could be viewed at all times. Testing sessions were videotaped and forelimb use was scored by a blinded operator. Movements scored were the independent use of the left or right forelimb or simultaneous use of both the left and right forelimb to contact the wall of the cylinder during a full rear, to initiate a weight-shifting movement, or to regain center of gravity while moving laterally in a vertical posture along the wall. Animals were tested for their baselines after stroke and 4 and 8 weeks after cell transplantation. Statistical analysisOutcome measurement for each experiment was reported as mean±SEM. All data were analyzed using SPSS 11 for Mac OS X (SPSS Inc.). Significance of inter-group differences was performed by applying Student's t-test where appropriate. The One-Way ANOVA analysis was used to compare group differences for the forelimb use as the dependant variable and groups as the single independent factor variable. Differences between the groups were determined using Bonferroni's post hoc test. A P-value of less than 0.05 was considered to be statistically significant.\n" ], "offsets": [ [ 0, 38643 ] ] } ]

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[ { "id": "16316465__text", "type": "full_text", "text": [ "BackgroundUsing antibodies to specific protein antigens is the method of choice to assign and identify cell lineage through simultaneous analysis of surface molecules and intracellular markers. Embryonic stem cell research can be benefited from using antibodies specific to transcriptional factors/markers that contribute to the \"stemness\" phenotype or critical for cell lineage.ResultsIn this report, we have developed and validated antibodies (either monoclonal or polyclonal) specific to human embryonic stem cell antigens and early differentiation transcriptional factors/markers that are critical for cell differentiation into definite lineage.ConclusionThese antibodies enable stem cell biologists to conveniently identify stem cell characteristics and to quantitatively assess differentiation.\nAlthough the stem cell concept was introduced decades ago, to date, stem cells can only be defined functionally, not morphologically or phenotypically. Two functions define stem cells. Firstly, they are self-renewing, thus able to propagate to generate additional stem cells. Secondly they can differentiate into various progenitor cells, which commit to further maturation along a specific lineage. While stem cells can be best defined functionally, a good number of molecular markers have been used to prospectively identify various stem cell populations. Although the functional importance of many of these antigens remains unknown, their unique expression pattern and timing of expression provide a useful tool for scientists to identify as well as isolate stem cells. Embryonic stem cells (ESC), derived from the inner cell mass of pre-implantation embryos, have been recognized as the earliest stem cell population [1,2]. This pluripotent population can differentiate into all somatic tissue including germ cells. In the case of human ESC, they can differentiate into some extra-embryonic derivatives as well. Like mouse ESC, human ES cells can be maintained and propagated on mouse fibroblast feeders for extended periods in media containing basic fibroblast growth factor (bFGF) [3]. Gene expression of undifferentiated human ES cells has been investigated among several ES cell lines by a variety of techniques. They include comparison with databases, reverse transcriptase-polymerase chain reaction, focused cDNA microarrays, and immunocytochemistry. A list of molecules comprised of known ES-specific or -highly expressed genes and candidates that can serve as markers for human ESCs and may also contribute to the \"stemness\" phenotype has been established [3-11]. For example, pluripotent ESC can be characterized by high level expression of Oct3/4 (POU domain, class 5, transcription factor 1, Pou5f1) and Nanog, which are a member of POU domain and homeobox transcription factors respectively. A critical amount of Oct3/4 and Nanog expression is required to sustain stem-cell pluripotency and both of these markers are downregulated as cells differentiate in vitro and in vivo [4-9]. Antibodies to Oct3/4 which cross react with human Oct 3/4 have been widely used to monitor the presence of undifferentiated ESC. No single marker however is sufficient or unique for identifying ESCs. Oct3/4 for example is expressed by germ cells and may be expressed by specific populations later in development. Likewise, Nanog has been shown to express in other tissues. We and other have noted however, that while no single marker is sufficient a constellation of positive and negative markers can in concert unambiguously allow one to define the state of ESC cultures and that surface markers in combination can be used to prospectively sort for ESC. Based on published data at the level of gene expression, we have cloned a number of candidate marker genes. We have also expressed the recombinant protein and generated a panel of monoclonal or polyclonal antibodies to these proteins. Using these antibodies we have confirmed the specificity and selectivity of these antibodies on several ESC lines and established their utility as stem cells markers. Our results confirm the expression pattern and timing of these cell markers at the protein level, whereas previous data reported at the level of gene expression.\nCharacterization of undifferentiated human ES cells and differentiated EBs by antibodiesAll monoclonal antibodies were initially selected for their abilities to recognize recombinant proteins in direct ELISAs. A subset were also tested by Western Blot analysis using recombinant proteins and cell lysate to confirm binding to a single epitope. The best clone was later screened for its applications for immunocytochemistry and flow cytometry using various cell lines. Human peripheral blood platelets were used for screening mouse anti-human CD9 antibody. MCF-7 cells were used for screening mouse anti-human E-Cadherin and PODXL (podocalyxin-like) antibodies. MG-63 cells were used for screening mouse anti-human GATA1 (GATA binding protein 1) antibody. Beta-TC6 cells were used for screening for mouse anti-human/mouse PDX-1 (pancreatic duodenal homeobox-1) antibody. NTERA-2 cells were used for screening mouse anti-human Oct3/4 and SOX2 (sex-determining region Y-box 2) antibodies. All polyclonal antibodies were affinity-purified using recombinant proteins and validated by direct ELISAs and Western. Caco-2 cells were used for validation of goat anti-human GATA6 antibody and NTERA-2 cells were used for validation of goat anti-human Nanog and anti-human Oct3/4 antibodies (Summarized in Table 1).Table 1Summary list of antibody verification by western blot.AntibodySample used for analysisMol. Wt. (KD)Gt × hBrachyurymouse ES-derived EB lysate48Ms × hDPPA5N/AN/AGt × hGATA6Caco2 cell lysate65Gt × hNanogNTERA-2 cell lysate33Gt × hOct 3/4NTERA-2 cell lysate39Gt × hPDX1beta-TC 6 cell lysate32Gt × hSOX17mouse ES-derived EB lysate45Ms × hCD9PBMC25Rt × hGATA-1N/AN/AMs × hE-CadherinMCF-7 cell lysate97Ms × hPODXLMCF-7 cell lysate57Ms × hSOX2NTERA-2 cell lysate36N/A: 1. DPPA5 is still being subcloned. Only Elisa verification is available.2. The clone for GATA-1 (MAB1779) does not work for Western blot application but is useful for IHC, The clone picked for Western blot analysis does not work for IHC (MAB17791, see data in ).After antibodies were validated in direct ELISAs, Western blot or cell lines (Fig. 1 and data not shown), they were used to examine the expression of individual molecules in undifferentiated human ES cells and differentiated EBs. When examined by immunohistochemistry, high level of expressions of Oct3/4, SOX2, E-Cadherin, PODXL and Nanog were observed in undifferentiated human ES cells (Fig. 2A, 2B and 2C). DPPA5 (developmental pluripotency associated 5) expression was also observed in undifferentiated human ES cells (data not shown). We noted that a subset of the proteins used were membrane bound proteins. To test if any of the antibodies generated could recognize an extracellular epitope and thus be used for live cell sorting, we repeated staining of live cells as previously described. The CD9, E-Cadherin and PODXL antibodies recognized an extracellular epitope and their ability to select cells by FACS was confirmed (Fig. 3). Minimal or no expressions of Oct3/4, E-Cadherin, PODXL and Nanog were detected in the differentiated EBs (Fig. 2D, 2E and 2F). However, SOX2 expression, which is observed in neural progenitor cells, is persistent in subsets of EBs.Figure 1Western blot analysis for Gt × hOct3/4 (A), Gt × hNanog (B) and Ms × hSOX2 (C) in NTERA-2 cell lysate, Ms × hE-Cadherin (D) in MCF-7 cell lysate, Ms × hCD9 (E) in PBMC lysate and Ms × hPDX-1(F) in β-TC-6 cell lysate. Numbers indicate the positions of molecular weight markers.Figure 2Undifferentiated human ES cells (A, B, and C) and differentiated EBs (D, E and F) were analyzed using antibodies to indicated molecular markers. Immunostaining with goat anti-human Oct3/4 (Red in A and D), mouse anti-human SOX2 (Green in A and D), goat anti-human E-Cadherin (Red in B and E), mouse anti-human PODXL (Green in B and E), and goat anti-human Nanog (Red in C and F), are contrasted with DAPI nuclear staining (Blue in C-F). Note the dramatic downregulation of ESC specific markers (Oct3/4, E-Cadherin, PODXL, and Nanog) in EBs. However, SOX2 expression is persistent in subsets of EB cells. Scale bars = 100 μm.Figure 3Human embryonic stem cells stained with anti-CD9 (A), anti-E-Cadherin (B), and anti-PODXL (C) and antigen expression detected by a flow cytometer. The specific staining is indicated by green histogram and corresponding isotype control is indicated by black histogram.Suspension culture with FGF withdrawal is known to induce differentiation of ES cells to all three germ layer precursors [12]. The differentiation status of the EB used here was detected to contain all germ cell markers by RT-PCR (Fig. 4). In order to examine how more antibodies can be used for characterization of early differentiation events from human ES cells, we examined the expressions of endodermal markers, SOX17, GATA6 and PDX-1, and mesodermal markers, Brachyury and GATA1, in the undifferentiated human ES cells and differentiated EBs. Expressions of SOX17, GATA6, PDX-1, Brachyury and GATA1 were not detected in undifferentiated human ES cells (data not shown). In contrast to the undifferentiated ES cells, subpopulations of SOX17-, GATA6-, Brachyury- and GATA1-positive cells were observed (Fig 4). These results suggest that both endodermal and mesodermal precursors exist in EBs with FGF withdrawal for 8 days. However, no PDX-1-positive cells were seen in EBs differentiated with the same treatment (data not shown).Figure 4Differentiated EBs were analyzed by either immunocytochemistry or RT-PCR to the indicated molecular markers. (A) Immunostaining with goat anti-human SOX17 (Red), is contrasted with Fluoro Nissl nuclear staining (Green). (B) Immunostaining with goat anti-human GATA6 (Red), is contrasted with DAPI nuclear staining (Blue). (C) Immunostaining with goat anti-human brachyury (Red), is contrasted with DAPI nuclear staining (Blue). (D) Immunostaining with mouse anti-human GATA1 (Red). Note that each antibody recognizes subsets of EB cells. Scale bars = 100 μm. (E) The differentiation status of EB is detected by RT-PCR using different germ layer cell markers. Selected endoderm markers AFP, FoxA2; mesoderm markers Hand1, MSX1 and ectoderm marker Msl1 were all highly expressed in the EB samples while their expression was either undetectable or at low level in the ES samples. G3PDH was a positive control showing similar amount of RNA samples were used for analysis. Examination of cross-reactivity of antibodies on mouse ES and differentiated cellsWe have also examined the cross-reactivities of these antibodies to mouse ES cells using mouse D3 ES cell line and mouse fetal endodermal tissue. Cross-reactivity to mouse of goat anti-Oct3/4, goat anti-PDX-1, goat anti-SOX17 and mouse anti-SOX2 was detected. Minimal cross-reactivity to mouse, measured by 10% intensity to human by higher than control cells, was observed in mouse anti-CD9 and mouse anti-E-cadherin antibodies. Goat anti-Nanog and mouse anti-PODXL antibodies appear to be human-specific as well (data not shown). The subtypes of monoclonal antibodies were also identified in the best clones. These results are summarized in Table 2.Table 2Summary of antibodies detection in ES and EB samples.AntibodyESEBReactivity to mouseIsotype of monoclonal antibody (Clone No.)Gt × hBrachyuryNoYesNT*Ms × hDPPA5YesNT*NT*ND*Gt × hGATA6NoYesNT*Gt × hNanogYesDownNoGt × hOct 3/4YesDownYesGt × hPDX-1NoNoYesGt × hSOX17NoYesYesMs × hCD9YesNoMinimalMouse IgG2B (clone 209306)Ms × hE-cadherinYesNoMinimalMouse IgG2B (clone 180224)Ms × hGATA1NoYesNT*Rat IgG2B (clone 234732)Ms × hPODXLYesNoNoMouse IgG2A (clone 222328)Ms × hSOX2YesYesYesMouse IgG2A (clone 245610)*NT, Not tested; ND, Not determined.\nThe expression patterns detected using antibodies developed in our facility are consistent with data reported using reverse transcriptase-polymerase chain reaction or cDNA microarrays. Moreover several of the monoclonal antibodies have differing heavy chain subunits allowing double labeling using subtype specific markers to be performed. In summary, we have developed a useful collection of antibodies that would be useful for identification of stem cell characteristics and assessment of differentiation. Several additional antibodies to the molecules that have been identified as potential cell lineage markers [13] are currently under development using the same approach.\nCloning and expression of Brachyury, DPPA5, CD9, E-Cadherin, GATA1, GATA6, Nanog, Oct3/4, PDX-1, PODXL, SOX2 and SOX17Brachyury (aa. 1–202), DPPA5 (a.a. 1–116), GATA1 (a.a. 1–413), GATA6 (aa. 1–449), Nanog (aa. 153–305), Oct3/4 (aa. 1–265), PDX-1 (aa. 1–283), SOX2 (aa. 135–317) and SOX17 (aa. 177–414) were expressed in E. Coli and extracellular domains of CD9, E-Cadherin, PODXL were expressed in mouse NSO cells. All proteins were purified and sequenced before they were used as antigens for immunizations and as substrate for antibody screening and subcloning. Production and purification of antibodiesAll monoclonal antibodies were derived from fusions of mouse myeloma with B cells obtained from BALB/c mice which had been immunized with purified antigen. The IgG fraction of the culture supernatant was purified by Protein G affinity chromatography (Sigma). Each panel of antibodies was screened and selected for their abilities to detect purified recombinant antigen in direct ELISA and Western blot. All polyclonal antibodies were derived from sera of goats which had been immunized and boost it with purified antigen. Antibody was purified from the sera by an antigen-affinity chromatography. Cells and cell cultureHuman Caco-2, MG-63, MCF-7, NTERA-2 and mouse D3 cells were purchased from American Type Culture Collection (ATCC). Cells were cultured according to the ATCC instructions. Information regarding human ES cell line HSF-6 (NIH code UC06) can be obtained at the website [14]. Undifferentiated human ES cells were cultured according to the protocol provided by the University of California, San Francisco in human ES culture medium [DMEM supplemented with 20% KnockOut Serum Replacement (Invitrogen) and 5 ng/mL of bFGF (R&D Systems)]. To induce formation of embryoid bodies (EBs), ES colonies were harvested, separated from the MEF feeder cells by gravity, gently resuspended in ES culture medium and transferred to non-adherent suspension culture dishes (Corning). Unless otherwise noted, EBs derived from human ES cell aggregates were cultured for 8 days in ES culture medium deprived of bFGF and used for analysis by immunohistochemistry as described. Western blotCells are solubilized in hot 2× SDS gel sample buffer (20 mM dithiothreitol, 6% SDS, 0.25 M Tris, pH 6.8, 10% glycerol, 10 mM NaF and bromophenyl blue) at 2 × 106 per mL. The extracts are heated in a boiling water bath for 5 minutes and sonicated with a probe sonicator with 3–4 bursts of 5–10 seconds each. Samples are diluted with 1× SDS sample buffer to the desired loading of 1–5 × 103 per lane. Lysates were resolved by SDS-PAGE, transferred to Immobilon-P membrane, and immunoblotted with 0.5 μg/mL primary Abs as described in R&D Systems Website [15]. ImmunohistochemistryAntibodies were used with the appropriate secondary reagents at a concentration of 5 to 10 μg/ml. Cells or sections of EBs were fixed with 4% paraformaldehyde in PBS at room temperature for 20 min, then blocked and permeabilized with 0.1% Triton X-100, 1% BSA, 10% normal donkey serum in PBS at room temperature for 45 min. After blocking, cells were incubated with diluted primary antibody overnight at 4°C followed by coupled anti-mouse or anti-goat IgG (Molecular Probes) at room temperature in the dark for an hour. Between each step cells were washed with PBS with 0.1% BSA. RT-PCRTotal RNA was extracted from EBs using Trizol LS (Invitrogen). cDNA was synthesized by using Superscript II reverse transcriptase (Invitrogen) according to the manufacturer's recommendations. The PCR primers are available upon request. Flow cytometryAntibodies were prepared at the concentration of 0.1 mg/mL. 10 μL of the stock solution was added to 1 – 2.5 × 105 cells in a total reaction volume not exceeding 200 μL. The sample was then incubated for 20 min at 2–8 °C. Following incubation, excess antibody was removed by washing cells twice with FACS buffer (2% FCS and 0.1% sodium azide in Hank's buffer). After wash, cells were resuspend in 200 μL of FACS buffer and the binding of unlabeled monoclonal antibodies was visualized by adding 10 μL of a 25 μg/mL stock solution of a secondary developing reagent such as goat anti-mouse IgG conjugated to a fluorochrome for 20 min at 2–8°C. Following incubation, cells were washed once with FACS buffer, once with PBS. After wash, cells were resuspend in 400 μL of PBS and analyzed on a FACScant flow cytometer (Becton-Dickinson, Mountain View, CA). Five thousand events were collected and analyzed using CELL Quest software.\n" ], "offsets": [ [ 0, 17113 ] ] } ]

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[ { "id": "17389645__text", "type": "full_text", "text": [ "Mapping sites within the genome that are hypersensitive to digestion with DNaseI is an important method for identifying DNA elements that regulate transcription. The standard approach to locating these DNaseI-hypersensitive sites (DHSs) has been to use Southern blotting techniques, although we, and others, have recently published alternative methods using a range of technologies including high-throughput sequencing and genomic array tiling paths. In this article, we describe a novel protocol to use real-time PCR to map DHS. Advantages of the technique reported here include the small cell numbers required for each analysis, rapid, relatively low-cost experiments with minimal need for specialist equipment. Presented examples include comparative DHS mapping of known TAL1/SCL regulatory elements between human embryonic stem cells and K562 cells.\nMapping the location of DNaseI-hypersensitive sites (DHSs) remains central to developing our understanding of transcriptional regulation. Elements with a range of transcriptional regulatory functions have been identified initially as DHSs. These include transcriptional enhancers (1,2) and repressors (3,4) as well as chromatin insulators and barrier elements (5,6). A number of techniques have been published recently that permit the mapping of DHSs without the need for Southern blotting (7–13). These include high-throughput sequencing of cloned DNA libraries derived from DNaseI-digested chromatin (8,9), and a number of different approaches that use genomic tiling path arrays to map DHSs (7,11,13). While these approaches have the advantage of covering large genomic regions with a limited number of experiments, they are inherently costly and less applicable to the rapid DHS mapping of specific genomic sites in a range of cell types. An alternative approach that does permit the targeted semi-quantitative DHS mapping of specific loci has used quantitative real-time PCR to map sites from digested DNA (10). This technique depends on the quantification of relative loss of PCR signal observed when PCR primers amplify across regions of digested DNA compared with amplification of undigested DNA. This approach has been reported as showing good sensitivity, but a limitation is the large number of PCR reactions required to quantify the calculated loss of signal with significant certainty. In this article, we present an alternative method to identify DHS using real-time PCR by adapting the protocol we have used to map DHS using genomic tiling path arrays (11). We have previously demonstrated the high sensitivity and specificity of the basic protocol with regard to identifying DHS across large genomic regions (11). Here, we detail the laboratory protocol that permits the rapid comparative mapping of known and candidate DHS between different cell types using real-time PCR. As examples, we include comparative DHS mapping of regulatory elements located across the extended TAL1 (T-cell acute lymphocytic leukaemia-1, also known as SCL (stem cell leukaemia)) locus in human embryonic stem (hES) cells and the leukaemia cell line K562.\nGenerating a library of DHSsThe basic protocol is outlined in Figure 1. Nuclei are extracted from living cells, then digested on ice for 1 h with a range of DNaseI concentrations, as detailed in the ‘Materials and methods’ sections. Following RNaseA and proteinaseK treatment, the DNA is extracted and run on a 1% agarose gel to check for the size of digested DNA. The gel in Figure 1A shows the DNA from mouse thymocyte nuclei digested with 0, 40 and 120 units of DNaseI. Maximal enrichment at DHS is usually observed with samples that are not over-digested. In the experiment shown, maximal enrichment at a known control DHS was seen with 40 units DNaseI (real-time PCR quantification shown in Figure 2B). The DNA is then blunt-ended using T4 polymerase (Figure 1B) and ligated with an asymmetric double-stranded linker. After extraction, DNA is amplified using a biotinylated linker-specific primer and 35 thermal polymerase cycles, as detailed in ‘Materials and methods’ section. As the linker will ligate to both ends of digested DNA, the amplification will represent a mix of primer extension and PCR, depending on the length of DNA amplified. As has been previously reasoned (13), one double strand of DNA in a region of DNaseI hypersensitivity is more likely to be digested twice within a short distance than non-hypersensitive DNA. This will lead to the preferential amplification of DNA from regions of DHS. The amplified DNA is then extracted using para-magnetic streptavidin beads, which provides a DNA library representative of whole-genome DHS. Agarose gel electrophoresis of the DNA recovered from the beads confirms that the vast majority of these products are between 300 and 500 base pairs in length (Figure 1B). Real-time PCR quantification of DNA within the DHS libraryFigure 2 documents the quantification of specific DNA sites from different mouse-cell-derived DNA libraries. Figure 2A shows the quantification of material from two primer sets, which are within (primer pair A) and 3′ (primer pair B) to the mouse Stil promoter. The left-hand panels of Figure 2A show the Sybr-green real-time fluorescence profiles using serial 5-fold dilutions of quantified mouse genomic DNA standards with primers A and B. A calculated standard curve then permits the quantification of DNA from this sequence in library samples. The right-hand panels of Figure 2A show the quantification of samples from 0 and 40 units DNaseI-treated material using primers A and B. While there is no difference in amplification between the samples using primer pair B, with primer pair A, the 40 units DNaseI sample amplifies ∼5.5 PCR cycles before the 0 units sample. This equates to greater than 40-fold enrichment at the Stil promoter compared with no observed enrichment downstream of the promoter. Figure 2B shows the quantifiable differences in enrichment at the porphobilinogen deaminase (Hmbs) promoter between primary mouse thymocytes and the mouse T-cell line BW5147. This representative primary mouse cell experiment was performed with the DNA shown in Figure 1A, which confirms that with these cells, maximal enrichment is observed from DNA that is not over-digested. The comparison of the thymocytes with the BW5147 cells illustrates another common finding that, in our experience of primary cell experiments, maximal enrichment is often obtained using lower amounts of DNaseI compared with cell lines. Comparison of real-time PCR DHS with Southern blottingAs the protocol generates template by primer extending away from a DNaseI-digestion site, as well as PCR amplification between DNaseI-digestion sites, there is the potential for the genomic size of the real-time DHS to be larger than sites revealed by Southern blotting. There is also potential for signal to be lost at the most ‘open’ stretch of DNA, due to over-digestion with DNaseI. These issues are addressed in Figure 3. The upper panel shows a Southern blot of the human STIL (SCL/TAL1 interrupting locus) promoter in K562 cells. The BglII restricted fragment is probed from the 3′ end, as shown in the upper panel. This reveals a central DHS ∼500 bp wide, with a suggestion of weaker hypersensitivity for a few hundred base pairs 5′ and 3′ to the central region. The lower panel shows real-time PCR data from K562 material using 10 primer sets, each generating an amplicon ∼120 bp long. This permits the quantification of enrichments over 1200 bp, centred around the STIL transcription start site. The lower panel represents the mean ± SD enrichment from three independent biological replicates from K562 cells. The black bar denotes the location of the ‘core’ hypersensitive site as defined by Southern blotting. There is good correlation between the location of the DHS between the two techniques. The 5′ extension of enrichment seen in the lower panel appears to reflect the weaker 5′ extension seen in the Southern blot in the upper panel. A dip in enrichment is observed in the lower panel over the previously mapped transcription-factor-binding sites (14), which may represent over-digestion of the most accessible core region of the DHS. Quantification of DHS across the TAL1 locus in hES and K562 cellsAs examples of relative enrichments at different regulatory elements, we present a comparison of enrichments across the extended TAL1 locus from human embryonic stem (hES) cells and K562 cells in Figure 4B. Relative quantification of mRNA expression using real-time PCR shows that both of these cell types express STIL, but only K562 cells express TAL1 (Figure 4A). TAL1 expression is critical to the establishment of haematopoiesis in a developing embryo (15). Post embryogenesis, TAL1 expression is maintained in the non-lymphoid haematopoietic system, although in addition, expression is observed in a range of non-haematopoietic tissues, including endothelium and brain (16–18). Over the past 10 years, we and others have dissected the regulatory elements that direct the expression of TAL1 to different tissues (11,19–31). These include the 3′ haematopoietic stem cell enhancer (+19) (24,30), the 5′ endothelial-haematopoietic enhancer (−4) (31), the 3′ erythroid enhancer (+40 in mouse, +50 in human) (28), and a number of neuronal elements (25,26). The location of the STIL promoter has been previously mapped (14), although no other STIL regulatory elements have yet been identified. Figure 4B shows the mean ± SD ratio (DNaseI-treated enrichment/DNaseI-untreated enrichment) of quantitative enrichments from two independent hES cell biological replicates and two independent K562 biological replicates, using primer sets from different locations as indicated in the figure. Primers indicated in red correspond to known regulatory elements. The hES cell cultures were maintained as detailed in methods. Both cell types show significant enrichment at the STIL promoter. Four control regions were selected: −16 kb, upstream of TAL1; +5 and +22 kb, within the TAL1 locus; +70 kb, downstream of TAL1. These sequences were chosen as control regions as they show limited homology between a number of species (26) and were considered highly unlikely to represent regulatory elements. All four regions showed no enrichments between DNaseI-treated and -untreated samples in either the hES cells or K562 cells. The most striking differences between hES cells and K562 cells were found at the TAL1 regulatory elements. With K562 cells, there is prominent enrichment at the TAL1 promoter 1a, and the −4, +19 and +50 enhancers. These cells express high levels of TAL1 transcripts, and were originally derived from a patient with blastic transformation of chronic myeloid leukaemia. K562 cells are relatively undifferentiated, although they have a partial erythroid phenotype. This potentially explains the marked enrichment seen at the +19 stem cell enhancer and the +50 erythroid enhancer. This is similar to data obtained from primary mouse erythroid cells (day 14.5 foetal liver), which show the highest enrichments at the mouse +40 enhancer (homologous to the human +50) (data not shown). In contrast, the hES cells show markedly reduced enrichment at the TAL1 regulatory elements when compared with K562 cells. Although there is enrichment at the +50 enhancer, there is minimal enrichment at the −4, promoter 1A and +19 elements. Reduced/absent accessibility of regulatory elements to DNaseI digestion is consistent with the lack of expression of TAL1 in hES cells (Figure 4A). As ES cells differentiate to form embryoid bodies, expression of TAL1 is rapidly switched on, appearing from day 3 of mouse ES cell differentiation (32). The relative accessibility of the +50 enhancer may reflect a poised chromatin state in hES cells, which permits a rapid response to the changing transcription factor environment that accompanies differentiation.\nThe development of techniques that permit the rapid comparative mapping of DHS between different cell types will greatly facilitate the study of transcriptional regulation in both normal and diseased cells. Recently published high-throughput techniques that map DHS sites using high-throughput sequencing (8,9) and genomic array tiling paths (7,11,13) have clear advantages of scale over more targeted approaches. Major disadvantages of these approaches include cost and lack of focus. This makes them less suitable for many laboratories that want to assess the chromatin accessibility of a number of defined or presumed regulatory elements in a range of cell types. A real-time-PCR-based approach to DHS mapping has, therefore, a number of potential advantages for researchers interested in specific regulatory questions at defined loci. Real-time PCR is relatively inexpensive compared with the large-scale techniques, and permits a rapid, focused DHS analysis of defined regions of DNA from multiple cell types. It also provides flexibility, as any genomic region can be analysed from the DNA library derived from the DNaseI-treated samples by designing further real-time PCR primer sets. We have previously shown that our basic technique for amplifying a library of DHS generates a template representative of known DHS with excellent sensitivity and specificity (11). Although the experiments presented in this paper were each performed using 5 million cells/digestion condition, we have obtained reproducible data using 5-fold fewer cells as starting material. We feel our technique can deliver acceptable specificity and sensitivity for DHS mapping with small numbers of cells, and will therefore be of use to those researchers working with limited numbers of primary cells. An alternative approach using real-time PCR to define DHS has been previously published (10). The two approaches differ in that Dorscher et al. quantify the DHS through the loss of PCR signal obtained from DHS when DNA is digested, whereas our approach uses real-time PCR to quantify a gain of signal observed from DHS. Dorscher et al. report excellent sensitivity using their approach to map DHS. However, the technique depends on large numbers of comparative quantitative real-time PCR reactions across a region in both digested and undigested material, in order to quantify the loss of enrichment. One advantage of our technique published here is that data can be obtained using far fewer quantitative PCR reactions. The technique is highly reproducible, with relatively little variation in quantifiable enrichments observed between different biological replicates. Moreover, we demonstrate tissue specificity, with variable enrichment at known regulatory elements between different cell types. The technique published here permits the rapid comparative analysis of DHS between different cell types from relatively small numbers of cells. It will have potential use for researchers across a broad spectrum of biology for the study of transcriptional regulation in both healthy and diseased tissues.\nCell cultureCell lines were maintained in culture as previously described (11). Care was taken to ensure maximum viability of cells when taken for experiments. The primary thymocytes used for Figure 2 were prepared by gentle physical disassociation of a whole thymus gland into PBS supplemented with 2% FCS. Cells were filtered to ensure that a single-cell suspension was taken forward for nuclei isolation. Human embryonic stem cells (hES) were grown in chemically defined media in the presence of Activin and FGF2, as detailed previously (33). In these conditions, hES cells remain homogenously undifferentiated. Nuclei preparationUp to 3 × 107 cells were washed in ice-cold PBS and resuspended in 2 ml of ice-cold cell lysis buffer [300 mM sucrose, 10 mM Tris pH 7.4, 15 mM NaCl, 5 mM MgCl, 0.1 mM EGTA, 60 mM KCl, 0.2% NP-40, 0.5 mM DTT, 0.5 μM spermidine, 1× protease inhibitor (complete, Roche)]. After 5 min, the lysed cells were spun at 500 g for 5 min at 4°C with brakes off. After careful removal of supernatant, the nuclei were gently resuspended in 200 μl of ice-cold reaction buffer (20 μl 10× DNaseI buffer, 4 μl glycerol, 176 μl water) using pipette tips with cut off ends. The nuclei were spun again at 500 g for 5 min at 4°C and, following supernatant removal, were resuspended in 30 μl of reaction buffer per DNaseI condition. For example, if 2 × 107 cells were taken to look at four different conditions (e.g. 0, 20, 60, 120 units DNaseI), the nuclei were resuspended in 120 μl of reaction buffer. Separate 30 μl aliquots were then taken and gently mixed with 70 μl of DNaseI mix (see Table 1) on ice. This made a final digestion volume of 100 μl for each sample, which was left to incubate for 1 h on ice in the cold room. After 1 h, 700 μl of nuclei lysis buffer (100 mM tris HCL pH 8, 5 mM EDTA pH 8, 200 mM NaCl, 0.2% SDS) was added to each sample with 50 μg proteinase K. Following gentle mixing with inversion, the lysed samples were incubated at 55°C for 1 h. RNaseA (10 μg (Ambion)) was then added to each sample and further incubated at 37°C for 30 min. DNA was then extracted using standard phenol–chloroform techniques. Care was taken to use cut-off tips and very gentle pipetting to reduce non-specific DNA sheering. Following precipitation, DNA was resuspended in 200 μl of 0.1 TE and quantified using spectophotometry. Samples (1 μg) were analysed using gel-electrophoresis, as shown in Figure 1A. DNA library preparationFollowing quantification, 7.5 μg of DNA was taken for each sample condition. This was blunt-ended using T4 polymerase (10 μl 10× buffer, 0.5 μl 25 mM dNTP, 3 μl BSA, 1 μl T4 polymerase (3 U/μl NE Biolabs), 85.5 μl of water/DNA in 0.1 TE). The samples were mixed gently on ice, then put at 12°C for 16 min. The reaction was stopped with excess EDTA (4 μl 0.5 M) followed by 75°C for 10 min. The DNA was further extracted using phenol–chloroform and precipitated with ethanol. Glycogen carrier (5 μg) was used at this stage, and the pellet resuspended in 80 μl water.The blunt-ended DNA was then ligated to the LP21–25 linker. A stock of linker was prepared by mixing 80 μl LP21 (100 μM: GAATTCAGATCTCCCGGGTCA) with 80 μl LP25 (100 μM: GCGGTGACCCGGGAGATCTG AATTC) and 240 μl water. The mix was then placed on a 95°C hot block and the power supply turned off. When the block had cooled to room temperature, the LP21–25 linker was aliquoted and frozen. The 80 μl DNA sample was split into two samples for ligation (5 μl 10× buffer, 3 μl LP21–25 linker, 3 μl ligase (NE biolabs 400 U/μl and 39 μl DNA). This was mixed well with a pipette and left at 16°C overnight.Following ligation, the DNA was precipitated with ethanol, and resuspended in 42 μl water. The sample was amplified using Vent exo-polymerase and a biotinylated LP25 primer. The mix (5 μl ThermoPol 10× buffer, 0.5 μl B-LP25 (200 μM), 2 μl Vent exo-(NE Biolabs 2 U/μl), 1 μl dNTP (25 mM) and 41.5 μl DNA) was amplified with the following thermal cycle: (95°C 3 min, 95°C 30 s, 61°C 30 s, 72°C 30 s) × 35 cycles.Following amplification, the biotinylated products were extracted using Dynal streptavidin beads. For each sample, 30 μl of beads (Dyabeads M-270; Dynal biotech) were washed twice in 2× binding buffer (10 mM Tris HCL pH7.5, 1 mM EDTA, 2 M NaCl) using a magnet, and then resuspended in 50 μl of 2× binding buffer/sample. Each post amplification sample (50 μl) was then mixed with 50 μl of resuspended beads and incubated at room temperature on a rotator for 1 h. After binding, the samples were washed twice with 2× binding buffer, then once with 1× TE. After the last wash, each sample was resuspended in 30 μl 0.1 TE. The paired samples that were split before ligation were then pooled, making a 60 μl final aliquot. This was heated at 95°C for 10 min to free the DNA from the beads, and then stored at 4°C. Real-time PCR quantificationWe have used Stratagene Brilliant SYBR Green QPCR Master Mix as our standard kit for quantification of genomic controls and samples. We have used a range of primer sets. The HMBS primers are: human—Forward; ATGCTGCCTATTTCAAGGTTGT, Reverse; GAATT GGAACATTGCGACAGT, and mouse—Forward; CGGAGTCATGTCCGGTAAC, Reverse; CGACCAA TAGACGACGAGAA. Primers from the TAL1 locus (human and mouse) are available on request.Amplification conditions and PCR mixes were as recommended by Stratagene. Genomic standard curves were calculated for each primer using serial 5-fold dilutions from 50 ng genomic DNA/PCR reaction. For each sample PCR, a 45 μl stock was made using 1.3 μl of sample DNA + 43.7 μl water. This stock was used as 5 μl per real-time PCR reaction, and following amplification, quantified relative to the genomic DNA standard curve, for each primer set. The quantifications shown in Figure 3 represent the absolute quantification of DNA at each primer set for the 5 μl PCR sample.The mRNA expression levels of TAL1 and STIL in Figure 4A were normalised relative to β-actin and plotted relative to mRNA expression from normal human donor CD34 + cells, as described previously (11).\n" ], "offsets": [ [ 0, 21052 ] ] } ]

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[ { "id": "17381551__text", "type": "full_text", "text": [ "This work uncovers novel mechanisms of aging within stem cell niches that are evolutionarily conserved between mice and humans and affect both embryonic and adult stem cells. Specifically, we have examined the effects of aged muscle and systemic niches on key molecular identifiers of regenerative potential of human embryonic stem cells (hESCs) and post-natal muscle stem cells (satellite cells). Our results reveal that aged differentiated niches dominantly inhibit the expression of Oct4 in hESCs and Myf-5 in activated satellite cells, and reduce proliferation and myogenic differentiation of both embryonic and tissue-specific adult stem cells (ASCs). Therefore, despite their general neoorganogenesis potential, the ability of hESCs, and the more differentiated myogenic ASCs to contribute to tissue repair in the old will be greatly restricted due to the conserved inhibitory influence of aged differentiated niches. Significantly, this work establishes that hESC-derived factors enhance the regenerative potential of both young and, importantly, aged muscle stem cells in vitro and in vivo; thus, suggesting that the regenerative outcome of stem cell-based replacement therapies will be determined by a balance between negative influences of aged tissues on transplanted cells and positive effects of embryonic cells on the endogenous regenerative capacity. Comprehensively, this work points toward novel venues for in situ restoration of tissue repair in the old and identifies critical determinants of successful cell-replacement therapies for aged degenerating organs.\nEmbryonic stem cells (ESCs) are distinguished by their ability to self-renew and to differentiate into any other cell type via asymmetric cell divisions, in which one daughter cell maintains ‘stemness’ while the other daughter cell differentiates into a particular tissue type. ESCs, including those of human origin (hESCs), are derived from the blastocyst and can be propagated in vitro (Evans & Kaufman, 1981; Thomson et al., 1998; Wobus & Boheler, 2005). Their tremendous potential for organogenesis has created a great interest in using hESCs for replacing tissues and organs lost to disease, or old age (reviewed in Wobus & Boheler, 2005). As such, the use of hESCs is particularly important, due to the fact that adult organ stem cells are often limited in number, cell-fate plasticity, expansion capacity, telomere length, and lifespan (Mayhall et al., 2004). The general goal behind most cell-replacement approaches is to expand and then differentiate hESCs in vitro, thus producing a cell type of interest, such as neuronal, blood, endothelial, pancreatic, bone, and others. These differentiated cells are expected to replace their dysfunctional counterparts in vivo. The scope of disorders that can be potentially treated with a neoorganogenesis approach is large and includes many that are currently incurable, such as muscle atrophy, diabetes, Alzheimer's disease, Parkinson's disease, and other degenerative diseases that often accompany human aging (McDonald et al., 1999; Liu et al., 2000; Hori et al., 2002; Kim et al., 2002; Blyszczuk et al., 2003). While many studies have focused on the derivation, propagation, and in vitro differentiation of hESCs (reviewed in Hoffman & Carpenter, 2005; Wobus & Boheler, 2005), relatively few have examined the properties of these cells and their more differentiated progeny in the aged, as opposed to the young, systemic and local organ environments. Recently published data suggest that these extrinsic cues become altered with age in ways that preclude activation of organ stem cells (such as satellite cells), inhibit repair-specific molecular signaling (such as delta-Notch), and interfere with productive tissue repair (Conboy et al., 2003, 2005; Janzen et al., 2006; Krishnamurthy et al., 2006; Molofsky et al., 2006). Furthermore, at least two lines of evidence suggest that stem cell-based tissue-replacement therapies might be hindered in the elderly, because all cells along the developmental lineage (e.g., stem cells, more differentiated progenitor cells or even tissues containing a pool of precursors) might rapidly ‘age’ and fail to contribute to organ repair when introduced into the old organism in vivo. First, in heterochronic tissue-transplantation studies, the age of the host environment determined the regenerative outcome, as both young and old skeletal muscle explants containing differentiated and precursor cells effectively regenerated in young, but not in old animals (Zacks & Sheff, 1982; Carlson & Faulkner, 1989). Second, using parabiotically paired young and old mice, the regenerative potential of muscle and liver was shown to be influenced by the age of the systemic environment (Conboy et al., 2005). Thus, we sought to determine whether key molecular identifiers of stem cell properties, the rate of cell proliferation, and the myogenic capacity would be influenced by the age of extrinsic milieu, regardless of whether stem cells are embryonic or the more differentiated, muscle-specific satellite cells. Satellite cells are muscle stem cells situated in direct contact with myofibers, the differentiated muscle cells. When myofibers are damaged, quiescent satellite cells are activated to proliferate and then differentiate into fusion-competent myoblasts that continue to proliferate and can form primary cultures, but are also capable of producing new, multinucleated myofibers or myotubes in vitro and in vivo (Morgan et al., 2002; Collins et al., 2005; Wagers & Conboy, 2005). Activated satellite cells express myogenic markers, such as Myf5, M-cadherin, and Paired box gene 7 (Pax7); fusion-competent myoblasts express high levels of desmin, and de novo generated myofibers or myotubes express embryonic myosin heavy chain (eMyHC) and continue to express desmin (Schultz & McCormick, 1994; Wagers & Conboy, 2005). While desmin can be also present in smooth and cardiac muscle cells, the isolation of hind limb skeletal muscle with subsequent purification of myofibers away from all interstitial cells, as well as purification of associated muscle stem cells results in primary cultures that are uniformly of skeletal muscle lineage. Every desmin+ cell in such cultures is a fusion-competent myoblast, and is able to produce multinucleated myotubes after 48 h of culture in differentiation-promoting medium [Dulbecco's modified Eagle's medium (DMEM) with 2% horse serum]. Some of these myogenic cells fuse into myotubes, even in the mitogen-rich medium [(Opti-MEM (Invitrogen, Carlsbad, CA, USA) with 5–10% mouse serum or DMEM with 10% fetal bovine serum, FBS] (Conboy & Rando, 2002; Conboy et al., 2003; and see below). An experimental system was developed that (i) provided the ability to study the regenerative response of hESCs and of muscle stem cells in various heterochronic environments in vitro; and (ii) allowed examination of the effects of hESCs on muscle repair, in vivo, after transplantation into young vs. old hosts. This model allowed us to address both the negative effects of the aged niche on key stem cell properties and the positive effects of hESCs on the aged muscle-specific organ progenitor cells in vitro, and on the regenerative capacity of old muscle in vivo. The resulting data demonstrate that the composition of conserved extrinsic cues, regulating stem cell responses, becomes altered with age in ways that inhibit both hESCs and adult stem cell regenerative potential. Specifically, molecular markers of stem cell functionality, e.g. Oct4 (in hESCs) and Myf5 (in muscle stem cells), the rate of cell proliferation, and the capacity for myogenic differentiation are all dominantly inhibited by the aged systemic milieu, and by the old differentiated muscle tissue. However, while satellite cells are unable to deter the inhibitory affects of aged systemic and local niches, hESCs are capable of antagonizing the aged environments, thereby enhancing the regenerative potential of both young and old muscle stem cells in vitro and in vivo. Thus, a complex interplay between negative regulation of hESCs and adult muscle stem cells by the aged niche, and positive regulation of the host's regenerative responses by hESCs will likely determine the success of hESC-based cell-replacement therapies in the old.\nRegenerative responses of adult muscle stem cells and hESCs are dominantly inhibited by the aged systemic milieuPrevious work established that the upregulation of repair-specific molecular signaling mechanisms, such as Notch, and successful engagement of resident muscle stem cells in tissue repair are largely determined by the age of the systemic milieu, rather than by the cell-autonomous age of muscle cells, or by the differences in their numbers (Conboy & Rando, 2005; Conboy et al., 2005). Intriguingly, these experiments also hinted at a small but persistent inhibitory effect of the aged systemic milieu on the performance of young stem cells. Exploring this further, we found that young serum permits satellite cells to be myogenic, while old serum inhibits the satellite cell regenerative potential not only alone, but also when mixed with young serum, suggesting a dominant over-riding of ‘young’ serum factors (Fig. 1). Myofiber cultures, in which satellite cells have been activated by injury in vivo, were established from young (2–3 months) and old (22–24 months) C57-BL/6 male mice, as previously described (Conboy & Rando, 2002; Conboy et al., 2005). As previously shown, this method is well suited for the assessment of satellite cell regenerative myogenic capacity (Conboy & Rando, 2002; Wagers & Conboy, 2005). Isolated myofiber explants with associated satellite cells were cultured overnight in the presence of young or old serum (alone at 5% and 10%, and mixed at 5% young + 5% old); bromodeoxyuridine (BrdU) was added for the last 2 h of culture to measure the rate of cell proliferation. The effects of heterochronic systemic milieu on myogenic potential were examined as generation of proliferating myoblasts that express desmin and Myf5, and that spontaneously form multinucleated nascent myotubes. As shown in Fig. 1A and quantified in Fig. 1B, the age of sera clearly determined satellite cell regenerative potential and old serum strongly inhibited the myogenic potential of young satellite cells either when present alone, or when mixed with young sera. Similar data was obtained by using another myogenic marker, Pax7 (Supplementary Fig. S1). Additionally, there were two to three times fewer total cells generated in the presence of aged serum (not shown).Fig. 1The age of sera determined the regenerative potential of satellite cells. (A) Young satellite cells were cultured either in 5% or 10% young (Young), 10% old (Old), or in a 5%+ 5% mouse sera combination (young + old). Cells were analyzed by immunofluorescence microscopy, using anti-BrdU (red), antidesmin (green) or anti-Myf5 antibodies (green, small panels). Similar results are shown for Pax7 immunodetection (Supplementary Fig. S1). Hoechst (blue) labeled nuclei. (B) Three independent experiments were quantified [300 young myofibers per experiment] as percentage of desmin+/Myf5+/BrdU+ de novo generated cells for each age and culture condition. On average, two to three fewer cells were generated when cultured in the presence of old. Shown are identical microscope fields at ×40 magnification. At least three independent experiments produced similar results. (*) indicates P≤ 0.001 as compared to young sera.Importantly, it was not simply the dilution of young serum factors that resulted in diminished myogenic capacity when young and old sera were mixed, because young sera promoted robust myogenesis both at 10% and 5%. Thus, old serum factors dominantly inhibited the myogenic capacity of young satellite cells even in the presence of young serum. This observation suggests that satellite cells of young mice engage in efficient myogenic responses, in part, because the inhibitory influence of old circulatory milieu is absent.These data reveal that the regenerative potential of young muscle stem cells is determined by the age of the systemic milieu, prompting us to investigate whether hESCs would similarly succumb to inhibitory factors present in the aged circulation.To determine the effects of aged serum on stem cell self-renewal/pluripotency, we analyzed hESC expression of Oct4 and studied the rate of hESC proliferation, by assessing BrdU incorporation (Fig. 2) and Ki67 expression (Supplementary Fig. S2). Specifically, these determinants of hESC regenerative potential were examined in the presence of heterochronic (young vs. old) mouse sera added to typical hESC medium, e.g., MEF-conditioned medium (MCM). Oct4 is expressed by self-renewing, pluripotent ESCs in culture, by the totipotent inner cell mass of the blastocyst and by the germ cells (Nichols et al., 1998; Pesce et al., 1999). Most cells in control cultures or young conditions expressed high levels of this marker of ‘stemness’, and maintained their normal phenotype and morphology throughout the various co-culture experiments performed in this study (see below).Fig. 2The regenerative potential of embryonic stem cells was negatively affected by aged mouse sera. (A) hESCs were cultured in MCM with 10% young (young) or old (old) mouse serum, or in three control media: MCM without mouse sera; GM (myoblast medium of Ham's F10 with 20% FBS) and DMEM/FBS (hESC differentiation medium of DMEM with 10% FBS). BrdU was added for the last 2 h of culture to measure the rate of cell proliferation. Immunodetection assays were performed for BrdU (red), Oct4 (red), and Ki67 (Supplementary Fig. S2). Hoechst (blue) labels nuclei. A high rate of hESC proliferation and Oct4 expression is displayed in all control media and in the presence of young mouse serum. In contrast, hESC proliferation and Oct4 expression are inhibited in the presence of old mouse serum, either alone or when mixed with young serum. MCM with mouse sera at 5% gave results similar to those observed with 10% young mouse sera or in control media (Supplementary Fig. S3). (B) Three independent experiments yielded similar results and were quantified as percentage of BrdU+ and Oct4+ cells for each culture condition. * indicates P < 0.001 as compared to young serum.Importantly, at 10% aged serum dramatically inhibited the self-renewal and proliferative potential of hESCs, as judged by highly diminished Oct4 expression and a lack of BrdU incorporation. Again, the inhibitory factors in the aged milieu were dominant over the young, as evidenced by a decline in Oct4 expression, the low rate of BrdU incorporation, and Ki67 expression in young and old mixed environments (5% young + 5% old sera in MCM). Similar to the data shown for adult stem cells (ASCs) (Fig. 1), it was not simply a dilution of young serum factors as hESCs robustly proliferated and expressed high levels of Oct4 when cultured with 5% young sera in MCM (Supplementary Fig. S3). Quantification of multiple independent experiments has demonstrated that hESC expression of Oct4 and BrdU incorporation have been reduced by two- to threefold in the aged milieu (Fig. 2B).As expected, hESCs cultured in control media, including MCM alone that does not contain either young or old serum, also displayed a high rate of proliferation and Oct4 expression (Fig. 2, control medium). Additionally, in this experimental set-up there was no general inhibitory effect of sera per se on hESC proliferation and Oct4 expression, as 10% young mouse sera (young) and 10–20% of FBS (growth medium and DMEM/FBS) allowed for a high rate of cell proliferation and for uniformly high Oct4 levels (Fig. 2).When instead of immediate exposure to aged mouse serum, hESCs were first cultured overnight in MCM, these cells were no longer susceptible to the negative effects of old systemic milieu (Fig. 3), suggesting that hESC-produced factors established an embryonic microniche that may provide temporary protection from the aged environment. It appears that satellite cells do not have such anti-aging ability, because despite an initial activation in entirely young environments, e.g., after muscle injury to young muscle, isolated satellite cells remain susceptible to inhibition by the old mouse serum (Figs 1 and 4C). Similarly, culturing satellite cells isolated from noninjured muscle in growth-promoting medium for 1–2 days does not protect against the inhibitory affects of aged systemic milieu (not shown).Fig. 3Embryonic stem cells produce youthful microniche in culture. (A) As opposed to immediate exposure to old mouse serum after passaging (10% old), preculturing of hESCs for 24 h in feeder-free conditions, e.g., Matrigel™ + MCM, prior to replacing MCM with MCM + 10% old mouse sera, resulted in continuously high BrdU incorporation and Oct4 expression (embryonic microniche + 10% old). BrdU was added for the last 2 h of culture to measure the rate of cell proliferation. Immunodetection of BrdU and Oct4 (both in red) was performed as described in Experimental procedures. Hoechst (blue) labels nuclei. (B) Three independent experiments yielded similar results and were quantified as percentage of BrdU+/Oct4+ for each condition. * indicates P < 0.001 as compared to ‘old + MCM’.Fig. 4Aged muscle niche inhibits the regenerative potential of hESCs and satellite cells. (A) Immunodetection of a mouse-specific M-cadherin (green) or desmin (red; both human and mouse proteins are detected) revealed that hESCs underwent muscle lineage differentiation when co-cultured with young, but not old myofibers. The myogenic progeny of hESCs appears M-cadherin−/desmin+ (white arrow in young), as opposed to M-cadherin−/desmin− hESCs that lack myogenic commitment (white arrow in old). M-cadherin+/desmin+ cells are the myogenic progeny of mouse satellite cells (yellow arrows). To assess the effects of secreted factors produced by young vs. old myofibers on the rate of hESC proliferation, transient, 2 h BrdU incorporation was examined in hESCs cultured for 48 h with supernatants produced by heterochronic myofiber explants (See Experimental procedures for details). As compared to young myofiber-derived supernatants (young myofiber supernant), exposure to old myofiber-derived supernatants (old myofiber supernant) inhibited hESCs proliferation, as judged by BrdU immunodetection (red). As expected, the rate of hESCs proliferation was high in control media (shown in Fig. 2). Hoechst (blue) labels nuclei in all experiments. Quantification of desmin+/BrdU+ hESCs in direct myofiber cocultures, or with muscle supernatants, is shown in (B). * indicates P≤ 0.001 as compared to young. (C) Transwell co-cultures between purified young satellite cells and myofibers isolated from uninjured young (young myofiber) and old (old myofiber) muscle demonstrated that satellite cell regenerative myogenic capacity was inhibited by the aged differentiated muscle. Myogenic potential was determined by the ability of satellite cells to generate proliferating desmin+ myoblasts (immunodetection shown in green) and by rate of proliferation (2 h BrdU incorporation; immunodetection shown in red). (D) Satellite cell regenerative potential was quantified as percentage of desmin+/BrdU+ cells for transwell co-cultures with young or old uninjured myofibers (i.e., RM, resting muscle). n = 3; * indicates P≤ 0.05 as compared to young.Comprehensively, these data establish that the inhibition of stem cell regenerative potential by the aged systemic milieu is conserved between species (mouse vs. human) and cell types (adult vs. embryonic stem cells). As summarized in Table 1, aged mouse sera similarly affected the expression of key molecular identifiers of both embryonic and adult stem cells, e.g., Oct4 in hESCs and Myf5 in mouse ASCs. As expected, adult mouse stem cells did not express Oct4, and hESCs did not express Myf5 in these experimental conditions (not shown). Moreover, aged systemic milieu had similar inhibitory effects on proliferation of hESCs and ASCs, suggesting that not only the regenerative capacity, but also the presence and expansion of stem cells will be significantly restricted in aged organs. Intriguingly, prolonged culturing of hESCs in their preferred in vitro conditions enables generation of an embryonic microniche that antagonizes the inhibitory influences of aged circulatory factors.Table 1Conservation of stem cell aging in the systemic environmentRate of proliferation ESC/ASC (percentage of BrdU)Call-fate identifier ESC (percentage of Oct4)/ ASC (percentage of Myf5)10% young59.5 ± 0.8, 59.3 ± 4.099.0 ± 0.1, 50.7 ± 9.510% old32.7 ± 2.1, 27.3 ± 3.517.6 ± 3.2, 18.1 ± 5.95% young + 5% old31.0 ± 2.6, 38.0 ± 2.020.6 ± 3.5, 17.1 ± 4.2Quantified results from Figs 1, 2 are summarized and presented as mean percentages from experimental replicates ± SE. Rate of proliferation (BrdU) and cell-fate identifier (Oct4 or Myf5) are shown for both ESCs and ASCs cultured in heterochronic systemic conditions of 10% young (young), 10% old (old) or in 5%+ 5% mouse sera combination (young + old). Results for 5% young mouse sera are very similar to those for 10% young mouse sera and are shown in Fig. 1 (ASCs) and Supplementary Fig. S3 (hESCs). The regenerative potential of hESCs and ASCs is inhibited by aged differentiated muscleAfter establishing that the aged systemic niche negatively affects the regenerative capacity of hESCs and of ASCs, we then assessed whether myogenic potential and the rate of cell proliferation would be inhibited in hESCs and ASCs by the aged local muscle niche. Myofibers with associated satellite cells were isolated from young and old injured muscle, and were directly co-cultured with hESCs in typical hESC differentiation medium (DMEM/FBS). Similar to Fig. 1, the myogenic potential in these co-cultures was assayed by the expression of desmin, which is present in both fusion-competent myoblasts and newly formed myotubes. To analyze whether hESCs, mouse myogenic progenitor cells or both could express desmin in direct co-cultures, we costained these cells with a mouse-specific antibody to a myogenic marker, M-cadherin, which does not react with human protein, and a desmin-specific antibody that recognizes both mouse and human proteins. As shown in Fig. 4A, hESCs underwent myogenic differentiation in co-cultures with young myofibers (M-cadherin−/desmin+ mononucleated cells, white arrow in young). These myogenic progeny of hESCs in co-cultures with young myofibers could be of skeletal, smooth or cardiac muscle lineages (Debus et al., 1983; Fischman & Danto, 1985; Schultz & McCormick, 1994). As expected, the young mouse muscle progenitor cells (M-cadherin+/desmin+) were more advanced in their degree of myogenic differentiation, which was of skeletal muscle lineage, as judged by the formation of large, multinucleated de novo myotubes (yellow arrow in young). In addition to the myogenically differentiated human cells, co-cultures with young myofiber explants also contained some small undifferentiated hESC colonies, as determined by immunoreactivity to a human-specific antibody to the nuclear mitotic apparatus protein, NuMA and Oct4 expression (Supplementary Fig. S4).In contrast, when co-cultured with the aged mouse myofibers, only mouse cells appeared desmin+ (Fig. 4A, yellow arrow in old). These aged myogenic cells were of skeletal muscle lineage, based on spontaneous generation of multinucleated myotubes (see Fig. 5B) and based on induced differentiation into myotubes in DMEM + 2% horse serum (not shown). Importantly, the myogenic differentiation of hESCs failed in the aged co-cultures (Fig. 4A, white arrow in old). Furthermore, colonies of hESCs in co-cultures with aged myofibers typically differentiated into cells with fibroblast morphology, which lacked Oct4 expression (not shown). Spontaneous production of desmin+ myogenic cells in control hESC cultures without myofibers, or with young/old mouse sera was less than 0.1% (not shown).Fig. 5In vitro co-culture with hESCs enhanced myogenesis of mouse cells. (A) 1 × 105 hESCs or control hMSCs were co-cultured with 5 × 106 primary mouse myoblasts. hESCs expressing Oct4 (immunodetection shown in red) dramatically enhanced myotube formation of co-cultured mouse myoblasts (immunodetection of eMyHC is shown in green), as compared to co-cultures between mouse myoblasts and human mesenchymal stem cells (Mb + hMSCs) or myoblasts alone (Mb alone). Experiments were carried out in myoblast differentiation medium. Hoechst (blue) labels nuclei throughout this figure. (B) 1 × 105 hESCs or control hMSCs were co-cultured with young or old myofiber-associated satellite cells, as described in Experimental procedures. Co-culture with hESCs (myofiber + hESC), but not hMSCs (myofiber + hMSC) or control medium (DMEM/FBS), greatly enhanced the myogenic potential of both young and old myofiber-associated satellite cells, based on immunodetection of percentage of desmin+ de novo generated myoblasts and multinucleated myotubes. These experiments were carried out in GM. Shown are myogenic responses of mouse cells only, judged by lack of immunoreactivity to human-specific/hESC-specific antigens, such as NuMA and Oct4; and presence of mouse-specific immunoreactivity, e.g., M-cadherin (not shown). Both young and old myofiber associated satellite cells exhibited considerable myogenic improvement over control conditions. n = 3.In concert with the conservation of inhibitory affects of aged systemic niche, the negative influence of local muscle niche was also found to be conserved in its inhibition of hESC and ASC regenerative responses. Specifically, the myogenic capacity (generation of desmin+ myoblasts) was inhibited in young satellite cells co-cultured in a transwell system with aged myofibers (Fig. 4B). In addition, hESC and ASC proliferation (BrdU incorporation) was also inhibited by aged differentiated muscle (Fig. 4A,C). These data suggest that not only systemic but also local organ niches would inhibit key stem cell properties, e.g., myogenic capacity and the rate of proliferation in the aged organism. The conserved inhibitory influences of the differentiated muscle niche on hESC and ASC regenerative responses are summarized in Table 2.Table 2Conservation of stem cell aging in the local organ nicheRate of proliferation ESC/ASC (percentage of BrdU)Myogenic differentiation ESC/ASC (percentage of desmin)Young myofiber60.2 ± 2.5, 40.5 ± 2.67.4 ± 0.9, 47.6 ± 5.0Old myofiber30.1 ± 4.3, 21.5 ± 4.11.3 ± 0.7, 19.7 ± 4.7Quantified results from Fig. 4 are summarized and presented as mean percentages from experimental replicates ± SE. Rate of proliferation (BrdU) and myogenic differentiation (desmin) are shown for both ESCs and ASCs, in the presence of young vs. old differentiated muscle environments (young myofiber or old myofiber). hESCs indirectly enhance and rejuvenate the regeneration of skeletal muscleWhile hESC properties were inhibited by aged differentiated muscle, the myogenic potential of aged satellite cells seemed to be enhanced by co-cultures with hESCs (Fig. 4A). Therefore, we further explored the enhancing and rejuvenating effects of hESCs on myogenic potential in vitro and in vivo, using human mesenchymal stem cells (hMSCs) as a negative control. First, we examined the effects of hESCs on myotube generation by co-culture with primary myoblasts freshly derived from activated-by-injury satellite cells (Conboy et al., 2003). As shown in Fig. 5A (Mb + hESC), primary myoblasts underwent very rapid and robust nascent myotube formation, when co-cultured with hESCs for 48 h in myoblast differentiation medium. Namely, remarkably large fused myotubes containing approximately 50–70 nuclei formed around hESCs colonies (Fig. 5A). In contrast, when co-cultured with hMSCs, myotube formation was no greater than in myoblast cultures alone (Fig. 5A, Mb + hMSC and Mb alone). Encouraged by these data, we analyzed the myogenic potential of young and old satellite cells co-cultured with hESCs for 48 h. As shown in Fig. 5B, hESCs conferred a much-enhanced myogenic capacity on both young and, importantly, old myofiber-associated satellite cells (rapid formation of desmin+ myogenic cells, many of which formed de novo multinucleated myotubes). Control co-cultures of these satellite cells with hMSCs displayed no enhanced myogenicity. In summary, while the myogenic potential (production of desmin+ fusion-competent cells) was more pronounced in young vs. old myofiber-associated satellite cells under all experimental conditions, a finding that is consistent with previous data (Conboy et al., 2003), a clear increase in myogenic potential of old satellite cells was noted in co-cultures with hESCs, as compared to control cultures devoid of hESCs (Fig. 4A,B).Interestingly, in addition to the rejuvenating effects of direct co-cultures shown in Fig. 5, soluble factors present in hESC-conditioned culture supernatants were also able to enhance myogenesis of aged satellite cells (Supplementary Fig. S5). Thus, in agreement with the notion that an established embryonic microniche antagonizes the inhibitory effects of the aged environment on stem cell responses (Fig. 3), the hESC-produced factors enhanced myogenic capacity of even old mouse satellite cells.Establishing that hESC-produced factors enhance adult myogenesis and rejuvenate the regenerative capacity of even aged satellite cells in vitro prompted us to examine whether the regeneration of old injured muscle will be improved by hESC transplantation in vivo. Additionally, based on the data shown above, we speculated that even if the host's repair capacity is improved, hESCs themselves will not be efficiently maintained or expanded in the context of old systemic and local organ environments, and will not directly contribute to the repair of aged skeletal muscle. To test these hypotheses, we injected 5 × 105 hESCs or control hMSCs into the tibialis anterior (TA) and gastrocnemius muscles of young and old mice at 24 h after cardiotoxin-induced injury, when activation/proliferation of endogenous satellite cells normally begins (Conboy et al., 2003, 2005; Wagers & Conboy, 2005). To avoid immune response against hESC antigens, mice were immunosuppressed using FK506 (Ito & Tanaka, 1997; Dumont, 2000). Muscle was isolated 5 days post-injury, when nascent differentiated myofibers normally replace the damaged tissue (Conboy et al., 2003), and 10 µm cryosections were analyzed for the success in tissue repair using hematoxylin and eosin (H&E) histochemistry and eMyHC immunodetection. H&E analysis reveals newly formed myofibers, based on their smaller size and centrally located nuclei. Additionally, de novo myofibers in the damaged area appear positive for eMyHC, while undamaged myofibers remain negative. As shown in Fig. 6A and quantified in 6B, injection of hESCs significantly enhanced regeneration of skeletal muscle. Remarkably, this positive embryonic effect was especially pronounced in old tissue.Fig. 6Skeletal muscle regeneration following hESC transplantation is a balance between the inhibitory influence of aged niches and the rejuvenating effects of hESCs. Young and old tibialis anterior and gastrocnemius muscles were injured by cardiotoxin injection. hESCs or hMSCs were transplanted at the site of injury and were analyzed by cryosectioning at Day 5 after injury (as described in Experimental procedures). (A) Newly regenerated myofibers were detected using eMyHC-specific antibody (green) and staining with H&E. In H&E staining, newly regenerated areas contain smaller, immature myofibers with centrally located nuclei. Uninjured myofibers are much larger, by comparison, with peripherally restricted nuclei. Poorly regenerated areas lack new myofibers and contain areas of fibrosis and inflammation. eMyHC immunodetection is specific for regenerating areas of muscle only. Both assays showed dramatic enhancement of muscle regeneration in ‘old + hESC’ vs. ‘old + hMSC’. Regeneration improvement was also seen in young + hESC, as compared to young + hMSC. (B) Quantification of muscle regeneration was performed by analyzing the density of newly formed myofibers per mm2 of injury site, which is the volume that typically covers the whole injured area. Multiple, 10 µm H&E sections were examined through the entire volume of injury in multiple, independently injured muscles. n = 20; * indicates P < 0.001 (‘old + hMSC’ compared to young + hMSC and ‘old + hMSC’ compared to ‘old + hESC’. (C) H&E and immunofluoresence staining for Oct4, and a human-specific antibody to NuMA, revealed the failure of hESCs to expand or persist in old, but the presence of hESCs in young muscle at 5 days post-transplantation. Hoechst (blue) labels nuclei.Importantly, such enhanced and rejuvenated muscle repair stems from an indirect induction, as hESCs themselves (or control hMSCs) did not physically contribute to the mouse myofibers, as judged by near absence (less than 0.1%) of human-specific NuMA+ nuclei in de novo desmin+ myofibers, analyzed through multiple injury sites. An example of one regenerated myofiber from young muscle injected with hESCs, with NuMA+ nucleus in a field of NuMA−/desmin+ mouse myofibers, is shown in Supplementary Fig. S6. No such NuMA+/desmin+ myofibers were detected in aged regenerated muscle (not shown).In agreement with the in vitro data, establishing that aged systemic and local niches inhibit hESC proliferation and Oct4 expression (Figs 2 and 4 and Supplementary Fig. S2), hESCs failed to expand or even persist in old muscle, as judged by the absence of NuMA+/Oct4+ hESC-derived cells in the aged tissue. In contrast, colonies of Numa+/Oct4+ hESC-derived cells that did not undergo myogenic differentiation were easily detected in young regenerating muscle (Fig. 6C). This finding validates several technical aspects of these experiments, and confirms the contrasting effects of young and old systemic and local organ niches on hESC self-renewal.These data further confirm and extrapolate our findings and demonstrate that when exposed to both aged systemic and local organ niches, hESCs fail to persist and do not contribute to tissue repair directly. At the same time, these embryonic cells indirectly but significantly improve the repair of aged injured muscle in vivo.\nThe data presented here establish for the first time that both the local environment of old differentiated organ, e.g., skeletal muscle and the systemic milieu dramatically affect the regenerative potential of both hESCs and mouse post-natal myogenic progenitor cells. Not only are the factors promoting myogenic differentiation and proliferation of hESCs likely to become depleted with age, but the aged systemic and local organ niches are likely to contain dominant inhibitors of ASC and hESC regenerative potential (Figs 1, 2, and 4, summarized in Tables 1 and 2). Importantly, the similar inhibitory effects of old mouse serum and old myofibers on satellite cell (Figs 1 and 4C) and hESC (Figs 2 and 4A) proliferation and regenerative capacity suggest the conservation of elements in age-specific extrinsic regulatory mechanisms between evolutionarily distinct species and stem cell types. Additionally, a similarity in the inhibitory properties between systemic and local organ niches is also of interest and may indicate that molecules produced by old tissues have circulatory/endocrine activity; and/or that age-specific systemic inhibitory components become deposited in the old tissues. Humans display broad genetic polymorphisms and behavioral variations, which makes the identification of age-specific molecular changes complicated. In contrast, laboratory mice are genetically and environmentally controlled. Establishing that age-specific signals, regulating stem cell responses, are evolutionarily conserved and soluble enables the formation of rational approaches for the identification and characterization of the inhibitors involved, and for revealing the precise timing of their first appearance in serum and differentiated tissues with advancing age. Significantly, these experiments have also revealed that not only are hESCs able to protect themselves against the negative influences of aged mouse sera (Fig. 3), but these cells also produce factors that dramatically enhance the myogenic capacity of primary myoblasts and young and old satellite cells (Fig. 5), and also significantly improve repair of young and old injured muscle in vivo (Fig. 6). Identification of these embryonic factors would allow us to potentially enrich the arsenal of therapeutic tools for combating age-specific degenerative disorders. The interactions between hESCs and heterochronic differentiated niches, initially identified in vitro, have been confirmed by in vivo experiments. Namely, while the regenerative capacity, or presence, of hESCs is greatly restricted in aged, as compared to young skeletal muscle (where transplanted cells experience both old systemic and local environments), embryonic cells indirectly enhance and rejuvenate muscle repair when introduced at the time of muscle stem cell activation in the host, e.g., at Day 1 after the injury (Fig. 6). It remains to be determined whether the percentage of hESCs direct contribution to desmin+ myofibers in young muscle will be increased by transplanting these cells at a different time-point after muscle injury, e.g., at Days 3–5 (as in co-cultures with myofibers pre-injured for 3 days, Fig. 4A). In any case, the virtual lack of hESC and hMSC direct contribution to the newly regenerated skeletal muscle, when small numbers of these cells were injected into injured tissue, is completely consistent with the body of previous data demonstrating that myofiber-associated satellite cells conduct rapid and robust muscle repair and greatly outnumber injected human cells (Collins et al., 2005; Wagers & Conboy, 2005); that compared to muscle-specific satellite cells, the myogenic differentiation of hESCs in vitro remains very small (Fig. 5, Table 2), and that control hMSCs are not normally myogenic unless these cells overexpress exogenous constitutively active domain of Notch (Dezawa et al., 2005). Intriguingly, the failure of hESCs to strive in old skeletal muscle might represent a therapeutically desirable outcome. For example, while in young tissue hESC derivatives putatively would go on to produce teratomas, it is unlikely that teratoma formation would occur after hESC transplantation into aged skeletal muscle. Thus, the indirect beneficial effects of hESCs on tissue repair are unlikely to be compromised by the oncogenic properties of these embryonic cells in the context of old skeletal muscle. Comprehensively, the results of this work increase our understanding of aging as a process, reveal evolutionary conserved age-specific interactions between stem cells and their differentiated niches, and suggest novel therapeutic approaches for improving the regenerative responses of endogenous or transplanted stem cells in old individuals.\nAnimal strainsYoung (2–3 months), C57-BL/6 male mice were obtained from pathogen-free breeding colonies at Jackson Laboratories (Bar Harbor, ME, USA). Aged 22–24 months C57-BL/6 male mice were obtained from the National Institute on Aging (NIH). Animals were maintained in the North-West Animal Facility of the University of California, Berkeley, CA, USA, and handled in accordance with the Administrative Panel on Laboratory Animal Care at UC Berkeley. Muscle injury and isolationMyofiber cultures, in which satellite cells were activated by in vivo injury, were set up as previously described (Conboy & Rando, 2002; Conboy et al., 2005). Briefly, mice were injured by direct injection with 5 ng cardiotoxin (CTX-1) (Sigma, St Louis, MO, USA) into the tibialis anterior and gastrocnemius muscles using a 28-gauge needle. After 1–5 days post-injection, injured or uninjured muscle tissue was dissected out. Once isolated, whole muscle was prepared for cryosectioning (see below) or myofiber fragments were obtained from hind limb muscles by enzymatic digestion (see below), trituration, and multiple sedimentation and washing procedures. Additionally, blood was collected from mice for the isolation of sera. Briefly, blood cells were coagulated at 37 °C for 15’ and then were centrifuged repeatedly at 5900 g, 4 °C in a microfuge for 3’ to isolate sera. Mixtures of young and old sera were made 1 : 1. For example, in 5%+ 5% conditions, 50 µL of young and 50 µL old serum were added to 900 µL of culture medium (Opti-MEM or MCM, see co-culture procedures below). Myofiber explant culturesExplant and primary cell cultures were generated from C57-BL/6 mice, as previously described (Conboy & Rando, 2002; Conboy et al., 2003). Dissected gastrocnemius and tibialis anterior muscles underwent enzymatic digestion at 37 °C in DMEM (Invitrogen)/Pen-Strep (Invitrogen)/0.2% Collagenase Type IIA (Sigma) solution. Isolated fibers were resuspended in GM (Ham's F10 nutrient mixture (Mediatech, Inc., Herndon, VA, USA), 20% FBS (Mediatech), 5 ng mL−1 bFGF (Chemicon, Temecula, CA, USA) and 1% Pen-Strep, and cultured on ECM-coated (BD Biosciences, San Jose, CA, USA) plates (diluted 1 : 500 in PBS). Cultures of primary myoblasts were derived from isolated fibers, through repeated passaging, and were maintained in GM. Myoblast differentiation medium [DMEM, supplemented with 2% horse serum (Mediatech)] was used to promote rapid formation of myotubes from cultured myoblasts (Morgan & Partridge, 2003). Human embryonic and mesenchymal stem cell cultureThe federally approved hESC line, H7 (NIH no. WA07, obtained from WiCell Research Institue, Madison, WI, USA), was used in accordance with the UC Berkeley and UC San Francisco Committee on Human Research guidelines, and in accordance with NIH guidelines. To propagate hESCs, routine culturing and maintenance was performed using standard in vitro conditions for both feeder-dependent and feeder-free cultures (Geron Corporation, 2002). Briefly, hESCs grown on MEFs were cultured in standard hESC medium [Knockout™ DMEM, 20% KSR, 1% NEAA, 1 mm l-glutamine (Invitrogen), 0.1 mmβ-mercaptoethanol (Sigma)] and were supplemented with 4 ng mL−1 hbFGF (Invitrogen). Feeder-free hESC cultures were maintained in MEF-conditioned hESC medium (MCM), 4 ng mL−1 hbFGF. Differentiation medium for hESCs (DMEM/FBS) was made by replacing KSR with 20% FBS (Hyclone, Logan, UH, USA). hMSCs were maintained in mesenchymal stem cell GM, MSC-GM™ and were cultured according to supplier recommendations (Cambrex Walkersville, MD, USA). hESCs and hMSCs were typically seeded onto chambered slides coated with a 3% GFR Matrigel™ (BD Biosciences) substrate in PBS. Cells were typically incubated for 48 h at 37 °C, 5% CO2, under the various experimental conditions employed, then were fixed with 70% EtOH/PBS at 4 °C. hESCs and hMSCs were analyzed 24–48 h after experimental treatments, during which no apoptosis-related differences in cell numbers were observed. Heterochronic co-culture systemsHeterochronic systemic cultures were established by culturing myofiber explants (in GM) or hESCs (in MCM) in the presence of young, old or young + old sera for 48 h (Figs 1 and 2 and Supplementary Figs S1–3). In such cultures, hESCs were passaged immediately prior to sera exposure. In contrast, preculturing of hESCs for 24 h in MCM, prior to replacing MCM with MCM + 10% old mouse sera was done for embryonic microniche experiments (Fig. 3). For heterochronic local organ niche cultures, hESCs were co-cultured directly with myofiber explants for 48 h in GM, or were cultured in the presence of supernatants derived from cultured myofiber explants for 48 h (Figs 4A and 5). Specifically, 1 × 105 hESCs or control hMSCs were co-cultured with identical volume, e.g., 100 µL, of young or old myofiber fragments with their associated satellite cells (Fig. 5). In experiments shown in Supplementary Fig. S5, culture-conditioned supernatant produced by hESCs grown in MCM was used as a medium in which 1 × 105 of myofiber-associated young or old satellite cells were cultured for 48 h. In direct co-cultures, mouse vs. human cells were distinguished by immunodetection with human-specific/hESC-specific and mouse-specific antibodies (Supplementary Fig. S4 and see below). To prepare muscle supernatants, explants were cultured for 24 h in GM and cellular debris was removed from conditioned media by multiple rounds of centrifugation. The absence of cells was confirmed by microscopic examination. To mimic the local organ niche for satellite cell assays (Fig. 4B), 1.0 µm transwell (Corning, NY, USA) co-cultures of uninjured explants with activated satellite cells were established. Activated-by-injury (24 h post-injury) satellite cells were seeded onto ECM-coated 12-well plates in Opti-MEM (Invitrogen) and 5% FBS. Transwells were placed over satellite cells and contained isolated myofiber explants from uninjured young or old muscle (i.e., resting muscle). Satellite cells were cultured for 72–96 h in the presence of myofiber explants and were fixed for immunodetection, as described above. Cell transplantationhESCs were grown on MEFs and expanded in 6-well plates. Cells were treated with 1 mg mL−1 Collagenase Type IV (Invitrogen) for 5–10 min, were washed and then incubated with 0.5 mg mL−1 Dispase (Invitrogen) to lift only human cell colonies. Isolated hESCs were washed several times and resuspended in 100 µL hESC medium. Similarly, hMSCs were expanded in 6-well plates, lifted with Trypsin/EDTA (Invitrogen), washed and resuspended in 100 µL hESC medium. Approximately 5 × 105 hESCs or hMSCs were injected into 24 h post-injured gastrocnemius and tibialis anterior muscles of young and old mice, using a 21-gauge needle. Immunosuppression of animals was achieved by intraperitoneal injection of 1 mg kg−1 FK506 (Sigma) at 48 h prior to cell transplantation, and on each day following transplantation. Immunodetection and histological analysisTo assay the affects of heterochronic local and systemic environments on stem cell regenerative potential, hESC, hMSC, and myofiber-derived precursor cell cultures were fixed with 70% EtOH/PBS at 4 °C, and were analyzed by indirect immunofluorescence. Combinations of antibodies were used to co-stain cultures and histosections, in order to determine the percentages of cells that proliferated or differentiated and to distinguish hESCs from mouse cells. Antibodies to the myogenic transcription factors, Myf5/Pax7, the intermediate filament protein, desmin, and the marker of newly formed myotubes, eMyHC, were used to reveal commitment to myogenic differentiation. Cell commitment to this differentiation program was assessed by the efficiency of myotube formation, estimated by the number of nuclei per myotube. Ki67, a cell cycle related nuclear protein consistently absent in quiescent cells, was used as a marker for proliferation. Whereas Ki67 appears in all active phases of the cell cycle, BrdU staining allowed exclusive detection of cells in S-phase, thereby enabling accurate quantification of DNA synthesis. In select cultures, 10 µm BrdU was added for 2 h prior to fixation. BrdU-specific immunostaining required nuclear permeabilization with treatment of 4N HCl. hESCs were distinguished from mouse cells by using a species-specific antibody to the cell-surface marker M-cadherin for murine and the nuclear marker NuMA for human cells. Antibodies to Oct4 were used as a marker of hESC self-renewal/pluripotency. Following permeabilization in PBS, +1% FBS, +0.25% Triton X-100, cells were incubated with primary antibodies (concentration determined as per manufacturer's recommendations) for 1 h at room temperature in PBS, +1% FBS, washed several times, and then incubated with fluorophore-conjugated, species-specific secondary antibodies (diluted 1 : 500 in PBS + 1% FBS) for 1 h at room temperature. For histological analysis, dissected muscle was treated in a 25% sucrose/PBS solution, frozen in OCT compound (Tissue Tek) and cryosectioned at 10 µm. Immunostaining was performed in the manner described above, or H&E staining of cryosections was performed. Nuclei were visualized by Hoechst staining for all immunostains. Samples were analyzed at room temperature by using a Zeiss Axioscope 40 fluorescent microscope, and imaged with an Axiocan MRc camera and AxioVision software. All images depict identical microscope fields at ×20 magnification, unless otherwise noted. ReagentsAntibodies to Oct4 (ab18976), BrdU (BU1/75 (ICR1), and Ki67 (ab15580) were purchased from Abcam (Cambridge, MA, USA). Antibody to M-cadherin (clone 12G4) was acquired from Upstate Biotechnology (Lake Placid, NY, USA), and NuMA antibody (Catalog number NA09L) from EMD Biosciences (San Diego, CA, USA). Antibody to developmental eMyHC (clone RNMy2/9D2) was acquired from Vector Laboratories (Burlingame, CA, USA). Myf5 (GTX77876) and Pax7 (GTX77888) antibodies were obtained from GeneTex (San Antonio, TX, USA). Desmin antibodies (clone DE-U-10 and Catalog number D8281), BrdU labeling reagent and FK506 (Catalog number F4679) were obtained from Sigma. Fluorophore-conjugated secondary antibodies (Alexa Fluor) were obtained from Molecular Probes (Eugene, OR, USA). Statistical analysesA minimum of three replicates were undertaken for each experimental condition. Quantified data are presented as means ± SE. Significance testing was performed using one-way analysis of variance (anova) to compare data from different experimental groups. P values of < 0.05 were considered as statistically significant.\n" ], "offsets": [ [ 0, 49710 ] ] } ]

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