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Cellular aging is linked to deficiencies in efficient repair of DNA double strand breaks and authentic genome maintenance at the chromatin level. Aging poses a significant threat to adult stem cell function by triggering persistent DNA damage and ultimately cellular senescence. Senescence is often considered to be an irreversible process. Moreover, critical genomic regions engaged in persistent DNA damage accumulation are unknown. Here we report that 65% of naturally occurring repairable DNA damage in self-renewing adult stem cells occurs within transposable elements. Upregulation of Alu retrotransposon transcription upon ex vivo aging causes nuclear cytotoxicity associated with the formation of persistent DNA damage foci and loss of efficient DNA repair in pericentric chromatin. This occurs due to a failure to recruit of condensin I and cohesin complexes. Our results demonstrate that the cytotoxicity of induced Alu repeats is functionally relevant for the human adult stem cell aging. Stable suppression of Alu transcription can reverse the senescent phenotype, reinstating the cells' self-renewing properties and increasing their plasticity by altering so-called “master” pluripotency regulators.
The stem cell hypothesis of aging1,2 postulates that the gradual and coordinated age-related loss of DNA damage repair capacity results in DNA damage accumulation over time; by triggering the persistent DNA damage response (DDR), cellular senescence and ultimately, stem cell aging. Although senescence was previously reported to be an irreversible process,3 several line of evidence indicate that senescence can be suppressed pharmacologically.4,5 Recent investigations of the distribution of the DNA damagespecific chromatin modification (γH2AX) in the genome of cycling Saccharomyces cerevisiae cells revealed “fragile” genomic locations.6 This suggests that mapping sites of γH2AX enrichment could be fruitful to pinpoint at-risk genomic elements in other genomes, perhaps indicating that specific genomic regions or elements exist where DNA damage is less efficiently repaired in aged cells. These functional elements have not been previously identified in the context of human adult stem cell aging. Here, we report that Alu retrortansposal RNA drives persistent DDR through the alteration of chromatin structure and that this event is functionally important as an integral component to human adult stem cell senescence ex vivo.
Here, we isolated adult adipose derived mesenchymal stem cells (hADSCs) (Fig. S1A), and investigated the mechanisms leading to their aging upon ex vivo expansion. As previously reported, upon isolation, hADSCs exhibit consistent self-renewing (SR) capacity until population doubling (PD)17, after which they displayed characteristic aging phenotypes7–9 (Fig. S1B). By PD37 (SR), hADSC cultures manifested a dramatic downregulation of the genes encoding cell cycle progression functions (Table 1) and accumulated non-dividing giant cells expressing the enzyme lysosomal pH 6 senescence-associated β-galactosidase (SA-β-Gal)10 (Fig. 1A). Cells self-renewed poorly, as determined by incorporation of 3[H] thymidine and bromodeoxyuridine (BrdU) into DNA (Fig. S1C and D). As hADSCs approached senescence, both mediators of DDR, phosphorylated form of histone variant H2AX (γH2AX),11 and p53 binding protein-1 (53BP1),12 form characteristic persistent DNA damage foci (Fig. 1B and C).11,13,14 The presence of these foci drastically increased from very rare in self-renewing ADSCs, to almost 90% in hADSCs approaching senescence (SEN hADSCs) (Fig. 1C).
In concert with the suggested association between genotoxic stress-induced DDR and retrotransposon activation,15,16 we observed a dramatic increase in Alu transcriptional activity in senescent hADSC (Fig. 1D). We did not observe a generalized upregulation of other major Pol III dependent genes, such as 7SL (Fig. 1D) or rRNA genes (Fig. 1E). Thus, this robust Pol III-dependent transcriptional activation upon senescence of hADSCs appears to specific to Alu retrotransposons.
Persistent DNA damage 53BP-1/γH2AX foci apparently were transcriptionally active in SEN hADSC (Fig. 1F) as indicated by in vivo nascent transcript labeling with FUr17,18 as described in Materials and Methods, and their co-localization with nuclear PML bodies implicated many cellular processes in DNA repair and transcription19–21 (Fig. 1G). These observations are consistent with previous reports indicating a possible involvement of RNA component(s) in 53BP1 foci formation after IR-induced damage in NIH3T3 and HeLa cells.22 The transcriptional activity within persistent DNA damage foci seems to be PolIII dependent, because treatment with the inhibitor of Pol-III, tagetin, abrogates FUr-incorporation in the transcripts and impairs 53BP1 focus-forming capacity in SEN hADSCs (Fig. 2). These results suggest a possible connection between transcriptional activity of Alu retrotransposons and genomic locations engaged in persistent DNA damage foci formation.
To identify genomic loci directly engaged in DDR in SEN hADSCs, we performed genome-wide profiling of hADSC chromatin using chromatin immunoprecipitation with an antibody against the γH2AX modified histone, followed by next generation sequencing (ChIP-seq) on the ABI SOLiD platform (Fig. 3A and Sup. Material). Asynchronous SR samples of hADSCs were used in similar ChIP-seq experiments to establish an overall representation of γH2AX modified chromatin and to define the locations of repairable DNA damage that does not affect the SR properties of the cells. Four replicate ChIPseq experiments each were performed for SR and SEN cells, and the resulting sequence tags were mapped to the human genome reference sequence (Table 2 and Sup. Material). Genomic maps of γH2AX sequence tags were subject to outlier removal, noise reduction, data merging from replicate experiments and clustering into contiguous mono-nucleosomal, midsize and large clusters (Sup. Material). The resulting genomic distributions of γH2AX modified nucleosomes and clusters were compared for SR and SEN cells (Fig. 3A) in order to assess the positional and quantitative changes in γH2AX-modified chromatin associated with these phenotypes.
Our mapping analysis revealed that γH2AX-modified chromatin was distributed non-randomly, with the majority of γH2AX modifications (SR: 65.4%, SEN: 65.2%) mapping to transposable elements (TEs), most of which were Alu (SINE), L1 (LINE) or LTR retrotransposons (Table S1). Alu elements were the most enriched for γH2AX-modified nucleosomes in SR cells compared with SEN cells, whereas L1s had the greatest relative increase of γH2AX-modified nucleosomes in SEN cells (Fig. 3B). The majority of the mono-nucleosomal γH2AX sites (67.3% in SR and 65.8% in SEN) were associated with the retrotransposal portion of the genome, with SINEs contributing 34.9% and 30.0% and LINEs accounting for 32.4% and 35.8% of these sites in SR and SEN hADSCs, respectively (Table S1). Because the retrotransposon portion of the genome is known to impede the progression of replication machinery,23–25 a portion of γH2AX detected in asynchronously dividing cells (the SR sample) might be caused by replication-fork pausing or collapse as described previously in yeast.23,26–28 Our data also indicates that the majority of damage in SEN cells is not related to unrepaired damage accumulation caused by the collision of replication forks and transcriptional complexes in SR cells, since gene density and γH2AX tag density were significantly positively correlated in SR samples and negatively correlated in SEN samples (Table S2).
We also observed substantial differences between the SR and SEN samples in the amount of (as well as the genomic features occupied by) large contiguous γH2AX clusters. The total amount of large γH2AX clusters dropped more than 3-fold comparing SR (37,467,892 bp) and SEN (11,243,429 bp) samples (Table S1). In SEN non-replicating samples, large γH2AX clusters were shifted to intergenic regions, with a decrease in intragenic sequences from 47.9% in SEN samples to 35.2% in SR (Table S1). Together, these data suggested a quantitative and qualitative drift in DNA damage susceptibility occurring upon human adult stem cell ex vivo aging.
Bioinformatic analysis allowed us to pinpoint genomic locations responsible for the formation of persistent DNA damage foci. Further analysis of ChIP-seq tags that bore a specific telomeric repeat sequence motif (TTA GGG/AAT CCC) among all sequencing tags (mapped uniquely combined with unmapped due to their repetitive nature) revealed that SR and SEN cell samples had similar numbers of γH2AX telomeric tags (1.66% vs. 1.63%). Similarly, peri-telomeric regions showed 1.5-fold (SR) and 2.1-fold (SEN) depletion in γH2AX-modified nucleosomes (Figs. 3C, S2 and S3), suggesting that in contrast to senescence of somatic cells driven by telomeric erosion-associated DNA damage accumulation,29–31 global telomeric and peri-telomeric DNA damage was an unlikely causal factor for hADSCs senescence.
Our data revealed that the distribution of γH2AX modified chromatin at pericentric regions was markedly different from those seen for peri-telomeres. There was a substantial over-representation of γH2AX modified chromatin in pericentromeres (Figs. 3D and S2), and a number of individual chromosomes exhibited highly enriched levels of pericentromeric γH2AX accumulation (Fig. 3E). Overall, pericentromeres exhibited 2.0- and 2.1-fold enrichments in γH2AX accumulation for SR and SEN cells, respectively (Fig. S3), thus indicating intrinsic susceptibility to DNA damage (i.e., “fragility”) of these chromosomal regions. Pericentromeres were not only enriched for γH2AX, but they were also one of the few genomic features that showed relatively greater γH2AX presence in SEN cells (Fig. 3B and D). Chromosomes 6, 10, 2 and 19 demonstrated the most significant enrichment of large γH2AX clusters in SEN cells (Fig. 3E). An increase in pericentric γH2AX accumulation on a subset of human chromosomes in SEN hADSCs could be indicative of: (1) defects in the assembly of kinetochores that provoke spindle-generated DNA damage, resulting in a compromised spindle-assembly checkpoint (SAC),32 and/or, (2) unresolved DSB, possibly due to changes in the characteristics of their pericentric heterochromatin and the molecular machinery responsible for its repair. Both of these events, acting separately or synergistically, could potentially trigger senescence of human adult stem cells.
We investigated both of these possibilities. We tested whether or not centromeric histone H3 variant (CENP-A), which is required to recruit many other centromere and kinetochore proteins,33 colocalizes with persistent DNA damage foci upon senescence. By using confocal fluorescent microscopy, we observed that CENP-A co-localized with persistent γH2AX/53BP1 foci in nearly 75% of the cases upon senescence of hADSCs (Fig. 4B and C), indicating that centromeric regions are embedded within persistent γH2AX/53BP1 DNA damage foci. Similar overlapping immunostaining patterns were observed when human anti-kinetochore serum (α-CREST) was prepped with γH2AX (Fig. S4A) or anti-53BP1 (Fig. S4B) antibodies. No detectable co-localization of centromeric regions with either 53BP1 (Fig. 4A) or γH2AX (data not shown) was observed in interphase of SR hADSCs, consistent with the fact that no prolonged cell cycle arrest was triggered in these cells (Fig. S1B–D), nor were γH2AX/53BP1 foci formed (Fig. 1B and C). These data further support the notion that the pericentric chromatin, rather then centromeres, per se, might drive DNA damage susceptibility, since no defects in the incorporation of CENP-A in centromeric chromatin or inner kinetochore assembly were observed.
The pericentric regions of the chromosomes shown to accumulate large clusters of γH2AX upon senescence (Figs. 3D, E, S3 and Table S1) are known to be enriched for Alu retrotransposal repeats.34 In multiple organisms, it has been shown that transcription from within pericentric heterochromatin is under tight control during the cell cycle and is required for the formation and maintenance of heterochromatin and sister chromatid cohesion.35,36 Because the persistent DNA damage foci associated with pericentric regions (Figs. 4B, C and S4) are actively transcribed (Fig. 1F and G) and sensitive to Pol-III inhibition (Fig. 2), and because we observed significant upregulation of transcriptional activity of Alu repeats upon hADSCs ex vivo aging (Fig. 1D), we next investigated if increased transcriptional activity of Alu retrotransposons could be recorded from the pericentric regions of the chromosomes susceptible for DNA damage accumulation in SEN hADSCs (Fig. 3E).
Chromosome 10 showed an increase of pericentric γH2AX tag accumulation in the SEN sample, including an excess of large clusters, whereas chromosome 21 had more pericentric γH2AX in the SR sample (Fig. 3E). RT-PCR was used to evaluate the transcriptional activity of individual Alu and MIR SINE sequences co-located with large γH2AX clusters in the pericentromeric regions of chromosome 10 (Fig. 5A) and chromosome 21 (Fig. S5). No transcriptional activity from MIR on ch10 was observed in either SR or SEN samples, whereas the transcription of Alus on both chromosomes were recorded in both cases. The chromosome 10 Alu sequence associated with a large cluster of γH2AX in the SEN sample was significantly upregulated in SEN vs. SR cells (Fig. 5A). On the other hand, AluJb and AluSx on chromosome 21 did not demonstrate significant differences in their transcriptional activity between SEN vs. SR cells (Fig. S5). Thus, the upregulation of Alu transcriptionl activity correlates with the presence of persistent DNA damage in SEN samples. Together, these data along with multiple reports from yeast and mammals,37–41 suggest a general role for the Pol-III transcription in genome organization, and prompted us to consider the hypothesis that, similar to centromeric tRNA genes in yeast,42 pericentric Alu retrotransposons in human adult stem cells might be integral to centromere function in self-renewing hADSCs, and an increase in the rate/level of their transcriptional activity upon ex vivo aging of hADSCs might be toxic to the chromatin environment and cause senescence.
Transcriptional activity of SINE/Alu retrotransposons is driven by the Pol-III transcriptional complex,43 and its general transcription factor TFIIIC, reportedly involved in the maintenance of high-order chromatin structure37,44–45 through structural organization of chromatin cohesion and condensation in yeast.42 We observed stable recruitment of TFIIIC to the analyzed retrotransposal repeats on chromosome 10 in both SR and SEN hADSC. Surprisingly, we observed a senescence-associated increase in the recruitment of TFIIIC to the MIR retrotransposal element located within the large γH2AX cluster even in the absence recorded MIR transcription (Fig. 5C). The specific association of cohesin with Alu elements has been reported in human cells,46 and multiple lines of evidence point to the critical role of the cohesin complex in DNA repair by homologous recombination47 as well as in spindle assembly and chromosome segregation during mitosis.34,47,48 Both cohesin and condensin complexes are indispensable for sister chromatid cohesion49 and DNA repair by homologous recombination (HR),50,51 while their deficiency has been shown to be responsible for triggering G1 and G2-M DNA damage checkpoints.52 Also, spindle assembly checkpoint (SAC) acts semi-redundantly with the DNA damage checkpoint to enact long-term cell cycle arrest in the presence of unresolved pericentric DSB, thereby preventing the segregation of the damaged chromatin.53–56
Therefore, we elected to investigate whether the reported toxicity of Alu RNA is related to an inability to repair pericentric DNA damage due to deficiencies in recruitment of the key complexes to the chromatin within or in direct proximity to DNA damage clusters. We investigated the recruitment of components of the cohesin complex Scc1/Rad21/Mcd1, the condensin I complex (Cap-H), and the histone acetylase Eco1, a regulator of Scc1/Rad21/Mcd1 cohesive state57–59 (Fig. 5B), to the Alu repeats in the vicinity of a persistent pericentric γH2AX cluster on chromosome 10. Data obtained by conventional ChIP analysis demonstrated a statistically significant loss in the recruitment of Scc1/Rad21/Mcd1 and Cap-H to the MIR and Alu transcription units during senescence of hADSC (SEN red bars in Fig. 5C) when compared with the status of the same genomic locations in SR cells (SR blue bar in Fig. 5C). Surprisingly, the recruitment of the histone acetylase Eco1 was largely correlated with the formation of persistent γH2AX clusters in senescence, and appeared to be Scc1/Rad21/Mcd1-independent. This suggests another, not yet identified, role for the Eco1 protein in DDR, different from its previously reported participation in the modulation of cohesiveness of Scc1/Rad21/Mcd1.46,57–59 Together these data further support the correlation between an increase in Alu transcription, persistent DNA damage and defects in pericentric recruitments of condensin I (Cap-H) and cohesin (Scc1/Rad21/Mcd1), and suggest an experimental approach to evaluate the functional significance of Alu toxicity in establishing the irreversible3 senescence of hADSCs.
In order to evaluate the functional significance of Alu transcription in establishing and/or mediating cellular senescence upon ex vivo aging, we took advantage of the common sequence feature of Alu retrotransposons to generate a number of lenti-virus-delivered shRNA constructs carrying GFP for assessment of transduction efficiency (Fig. 6A and Materials and Methods). hADSCs transduced with lentiGFP expressing sh-132Alu RNA (Fig. 6B) demonstrated a near complete knockdown of the generic Alu transcript (Fig. 6C). hADSCs transduced with lentiGFP (control) or lentiGFP sh-193Alu exhibited little or no change at the Alu transcription level. Surprisingly, SEN hADSCs lines stably expressing sh-132Alu RNA exhibited a dramatically altered morphology and exhibited an increase in proliferation as detected in 3[H] thymidine uptake DNA synthesis experiments (Fig. 6D). In contrast, SEN hADSCs transduced with lentiGFP (control), or treated with polybrene alone (PB), did not show any significant changes in their proliferation rate when compared with wild type (wt) SEN hADSCs (Fig. 6D). These results suggest that forced suppression of Alu transcription was sufficient to overcome persistent DDR in senescent hADSCs. sh-132Alu expressing cells were uniformly positive for proliferation specific antigen Ki67, expression of which is strictly associated with cell proliferation (Fig. 6E). Loss of senescence-associated β-galactosidase activity and disassembly of persistent DNA damage foci (γH2AX) was recorded in sh-132Alu expressing cells. Infected and uninfected cell populations are shown in the same field (Fig. 6F). Faithful re-establishment of DNA synthesis upon knockdown of the Alu transcript was further recorded 96 h after transduction and is sustainable as sh-132Alu expressing cells demonstrate proliferation upon culturing for 40 d or longer (Fig. S6 and data not shown). Unexpectedly, when compared with wt SEN hADSCs, prior SEN hADSCs lines stably expressing sh-132Alu RNA demonstrated a statistically significant upregulation of pluripotency factors Nanogand Oct4 as detected by qPCR analysis (Fig. 6G). The levels of Nanog and Oct4 in wt SEN hADSCs were below the threshold of the qPCR method.
In conclusion, our data support the observation that a retrotransposal portion of the genome is particularly sensitive to DSBs, a type of DNA breakage recently linked directly to genomic rearrangements in cancer.15,60–63 Unexpectedly, our results imply the importance of a fine-turned balance between beneficial (allowed) rate of Alu retrotransposons transcription, critical for mediating pericentromeric integrity during cell cycle progression and the toxicity of elevated transcriptional activity from the Alu retrotransposons, ultimately leading to human adult stem cell aging ex vivo. Our data indicate that such toxicity of Alu transcription is associated with loss of the recruitment cohesin and condensin I complexes to pericentromeric regions, thus blocking efficient repair of pericentric regions, leading to formation of persistent DNA damage foci (Fig. 6H). This impairment of condensin I loading is likely due to an absence in the recruitment of condensin complex, rather than a deficiency in Eco1, consistent with previous reports demonstrating that condensin is indispensible for cohesin loading onto the chromosome64 and that depletion of the Cap-H subunit of the condensin I complex results in defects associated with alterations in the structural integrity of centromere-proximal heterochromatin in Drosophila.65 Our data further linking together physiological impairment in recruitment of cohesin/condensin I complexes and Alu toxicity to persistent DDR upon adult stem cell ex vivo aging, suggests that such regulation might be an integral part of cellular aging or age-associated disease.66 Our results, in concert with previously published data,4,5 also challenge the notion that cellular senescence and ultimately cellular aging, is always an irreversible process.
Primary antibodies used were: 53BP1 (Bethyl Laboratories #A300-273A), BrdU (BD, 7580), CD31 (Invitrogen #HMCD3101), CD44 (Invitrogen #HMCD4401), CD45 (Invitrogen #HMCD4501), CD105 (Invitrogen #HMCD10520), Phospho-Chk1 (Ser345) (CST #2341), Phospho-Chk2 (Thr68) (CST #2661), CENP-A (Abcam #ab13939), human anti-centromeric autoantibody, α-CREST (Antibodies Inc., #15-235), γH2AX (Millipore #05-636). Secondary antibodies were AlexaFlour® conjugated donkey antibodies (Invitrogen).
Human adipose derived stem cells were isolated from human subcutaneous white adipose tissue collected during liposuction procedures. The lipoaspirate was suspended in Hank's Buffered Salt Solution (HBSS), 3.5% Bovine Serum Albumin (BSA), 1% Collagenase, type II (Sigma) in 1:3 w/v ratio and shaken at 37°C for 50 min. The cells were filtered through a 70 µm mesh cell strainer (BD Falcon #352350), treated with Red Blood Cell Lysis buffer (150 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, pH 7.3), and expanded ex vivo in DMEM/F12 complete medium (DMEM/F12, 10% FBS, 100 U/ml penicillin, 100 µg/ml streptomycin, 2.5 µg/ml amphotericin B; Invitrogen) in 10% CO2 at 37°C and passaged at 80% confluency, changing medium every 72–96 h. Cumulative population doublings were calculated by summing the population doublings (PD = log(N/N0) × 3.33, where N0 is the number of cells plated in the flask and N is the number of cells harvested at this passage) across multiple passages as a function of the number of days it was grown in culture.
Five × 105 cells each were incubated for 30 min on ice in the dark with fluorochrome-conjugated antibodies (CD31, CD44, CD45 and CD105; Invitrogen) in PBS with 1% BSA (Sigma), washed and analyzed in a Guava EasyCyte Mini System (Guava Technologies, Millipore). Data analysis was done with FlowJo software (Tree Star, Ashland, OR).
Senescence-associated β-galactosidase activity assay was done as described in manufacturer's kit (BioVision).
1–3 × 104 cells/well in 4-well slides were fixed with 4% paraformaldehyde and permeabilized with PBS, 0.5% Triton X-100. The blocking and antibody incubations were performed in 4% normal donkey serum (NDS; Jackson Immunochemicals) in PBS. The nuclei were counter-stained with 100 ng/ml 4′, 6-diamidino-2-phenylindole (DAPI; Sigma), and the slides were mounted in ProLong® Gold antifade aqueous mounting medium (Invitrogen). Analysis was performed as described in details in Supplemental Materials and Methods.
Human Cell Cycle qRCR Array (SABiosciences #PAHS-020C-2) was used for the profiling of self-renewing (SR) and senescent (SEN) hADSCs. Details of the experimental protocol and data analysis are given in Supplemental Materials and Methods online.
Nucleosomal ChIP was performed as described in reference 31, and in Supplemental Materials and Methods online. Samples were sequenced on SOLiD™ System 2.0 (Applied Biosystems) according to manufacturer's protocol.
Senescent hADSCs, 1 × 104 cells/well in 4-well slides, were treated with 2 mM 5-Fluorouridine (FUr; Sigma) for 10 min. The nuclei were exposed with ice-cold CSK buffer (100 mM KCl, 300 mM sucrose, 10 mM Pipes pH 6.8, 3 mM MgCl2, 1 mM EGTA, 1.2 mM phenylmethylsulfonyl fluoride) with 0.5% Triton X-100 and fixed with 4% paraformaldehyde. Immunostaining was performed as described above for immunoflourescence.
Genomic coordinates of the Alu elements tested in RT-PCR were from the March 2006 Assembly (NCBI36/hg18) of the Human Genome Browser at UCSC (http://genome.ucsc.edu); MIR: chr10: 41922815–41922906, Alu: chr10:41928992–41929118, AluJb: chr21:10141132–10141429 and AluSx: chr21:10145344–10145644. 100 ng of total RNA was used with the RT2 First Strand Kit (SABiosciences) per reaction. The primers for first strand synthesis are at locations outside of the Alu element sequences (external or reverse primers, Table S3) and forward primers within the Alu element sequence (internal forward primers, Table S3). RPL13A was used as a positive control.
Lentiviral shRNA constructs to knockdown genetic Alu transcript were designed as follows: oligonucleotides Lenti sh-132 Alu RNA 5′-GAT CCC CCC ACC ACG CCC GGC TAA TTT TCA AGA GAA ATT AGC CGG GCG TGG TGG TTT TTG GAA A-3′ and Lenti sh-193 Alu RNA 5′-GAT CCC CCC CGG GTT CAA GCG ATT CTT TCA AGA GAA GAA TCG CTT GAA CCC GGG TTT TTG GAA A-3′, were annealed with equal amounts of their complementary strands, creating restriction site specific overhangs for cloning, and ligated into HindIII and BglII digested, gel purified pENTR/pTER+ vector.67 The constructs were confirmed by sequencing (sense strand sequence is shown above). Equal amounts of each constructs was mixed with pLenti-CMV-GFP DEST vector67 in LR Clonase reaction to recombine cloned shRNA production elements into a destination vector according to manufacturer's instructions (Invitrogen). The produced lentiviral plasmid was transformed into E. coli Stbl3 cells (Invitrogen) for amplification.
293T cells were grown in DMEM complete medium (DMEM, high glucose, 10% FBS, 0.1 M nonessential amino acid, 6 mM L-glutamine, 1 mM pyruvate, 100 U/ml penicillin, 100 µg/ml streptomycin) for lentiviral production and transfected for 12 h with pLenti sh-132Alu or pLenti sh-193Alu and pCgpV, pRSV-Rev and pCMVVSV-G helper plasmids (Allele Biotechnology) in 2:1:1:1 molar ratio using Lipofectamine 2000 (Invitrogen) according to standard protocol.67 Medium was collected at 48 and 72 h and filtered (0.45 µm). Virus was precipitated with PEG and frozen in aliquots (−80°C). Lentiviral transductions were done in complete medium with 5 µg/ml Polybrene (Santa Cruz Biotechnology) for 12 h. Viral titers were determined by comparing GFP positive cells counts to total population.
For each condition in 3[H]-thymidine uptake assay, 10,000 cells were treated with 1 µCi 3[H]-thymidine (Perkin-Elmer, Boston, MA) in DMEM/F12 complete for 24 h. DNA was isolated from harvested cells and quantified with NanoDrop (ND-1000; NanoDrop Technologies Inc.). 3[H]-thymidine uptake into cellular DNA was measured with liquid scintillation counter (LS 6500; Beckman Instruments). Lentiviral transduction was done as above, and untreated and polybrene treated controls were performed in parallel.
RNA was isolated from SR and SEN hADSCs (with or without lentiviral transduction) with mir-VANA (Ambion, Invitrogen) kit and 2 mg of total RNA/lane was run on a 7 M Urea, 6% polyacrylamide, TBE. Gel was stained with ethidium bromide, photographed and electroblotted onto Hybond™-N+ (Amersham, GE Healthcare). Hybridizations were performed in 6x SSC, 4x Denhardts', 0.1% SDS at 37°C. Oligonucleotide probes were labeled with Biotin-16-dUTP (Roche) by terminal transferase (NEB). Northern was visualized with streptavidin-HRP (Invitrogen) using ECL Plus western Blotting Detection Reagent (Amersham, GE Healthcare) and Amersham Hyperfilm (GE Healthcare). Oligonucleotide used as probes were:
Alu 132: 5′-CCA CCA CGC CCG GCT AAT TT-3′ and Alu 90: 5′-CGC GCG CCA CCA CGC CCG GCT AAT TTT TGT ATT TTT AGT AGA GAC GGG GTT TCA CCA TGT TGG CC-3′.
No potential conflicts of interest were disclosed.
We thank Jeri Jenkins for help with manuscript preparation, Max Sarrazin for help with figures, and ABI Life Technologies for reagents and technical assistance for SOLiD instrumentation. For critical reading of the manuscript we are grateful to Drs. B. Kennedy, G. Lithgow and M. Classon. We thank Dr. Denise Muñoz and Justin Winstead for assistance with library preparation and sequencing runs. This work was supported by National Institutes of Health pilot projects on UL1 DE019608, Buck Institute Trust Fund to V.V.L. and by the Alfred P. Sloan Foundation (BR-4839) to I.K.J. V.V.L. designed the experiments; J.W. and I.K.J. developed and applied bioinformatics method for data analysis; G.G., S.L.H., M.A., B.B. and E.L.L. performed the experiments; P.J.C. and M.D. provided the primary cells; B.P. and E.H. mapped the SOLiD data, G.S. provided technical assistance for SOLiD instrumentation; I.K.J., M.G.R. and V.V.L. discussed the results and wrote the paper.