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Ectopically hTERT expressing adult human mesenchymal stem cells are less radiosensitive than their telomerase negative counterpart
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w w w. e l s e v i e r. c o m / l o c a t e / y e x c r
Research Article
Ectopically hTERT expressing adult human mesenchymal stem cells are less radiosensitive than their telomerase negative counterpart Nedime Serakinci a,b,⁎,1 , Rikke Christensen a,1 , Jesper Graakjaer f , Claire J. Cairney c , W. Nicol Keith c , Jan Alsner d , Gabriele Saretzki e , Steen Kolvraa f a
Department of Human Genetics, University of Aarhus, Aarhus, Denmark Institute of Medical Biology, Department of Anatomy and Neurobiology, Southern Denmark University, Odense, Denmark c Centre for Oncology and Applied Pharmacology, University of Glasgow, Cancer Research UK, Beatson Laboratories, Garscube Estate, Glasgow, UK d Department of Experimental Clinical Oncology, Aarhus University Hospital, Aarhus, Denmark e Henry Wellcome Laboratory for Biogerontology, Newcastle General Hospital, University of Newcastle upon Tyne, Newcastle, UK f Department of Clinical Genetics, Vejle County Hospital, Vejle, Denmark b
ARTICLE INFORMATION
ABS T R AC T
Article Chronology:
During the past several years increasing evidence indicating that the proliferation capacity
Received 30 June 2006
of mammalian cells is highly radiosensitive, regardless of the species and the tissue of origin
Revised version received
of the cells, has accumulated. It has also been shown that normal bone marrow cells of mice
21 December 2006
have a similar radiosensitivity to other mammalian cells so far tested. In this study, we
Accepted 3 January 2007
investigated the genetic effects of ionizing radiation (2.5–15 Gy) on normal human
Available online 8 January 2007
mesenchymal stem cells and their telomerised counterpart hMSC-telo1. We evaluated overall genomic integrity, DNA damage/repair by applying a fluorescence-detected alkaline
Keywords:
DNA unwinding assay together with Western blot analyses for phosphorylated H2AX and
Telomere dysfunction
Q-FISH was applied for investigation of telomeric damage. Our results indicate that hMSC
Telomerase
and TERT-immortalized hMSCs can cope with relatively high doses of γ-rays and that
Mesenchymal stem cells
overall DNA repair is similar in the two cell lines. The telomeres were extensively destroyed
Ionizing radiation
after irradiation in both cell types suggesting that telomere caps are especially sensitive to
p16INK4a
radiation. The TERT-immortalized hMSCs showed higher stability at telomeric regions than
Telomere capping
primary hMSCs indicating that cells with long telomeres and high telomerase activity have
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the advantage of re-establishing the telomeric caps. © 2007 Elsevier Inc. All rights reserved.
Introduction Lack of telomere maintenance is a genetic feature that appears to be associated with multiple genetic defects, all
conferring radiosensitivity. Human radiosensitive cell lines have thus been shown either to have short or dysfunctional telomeres and in a mouse model it has been shown directly that mice with critically short telomeres are more
⁎ Corresponding author. Institute of Medical Biology, Department of Anatomy and Neurobiology, Southern Denmark University, Winsløvparken 21, 5000 Odense C, Denmark. Fax: +45 6590 6321. E-mail addresses: [email protected], [email protected] (N. Serakinci). 1 Authors equally contributed to the 1st authorship of the work. 0014-4827/$ – see front matter © 2007 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2007.01.002
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radiosensitive to ionizing radiation (gamma rays) and show an increased chromosomal instability [1]. Telomeres are the physical ends of chromosomes and are responsible for the maintenance of chromosome stability and therefore dysfunctional telomeres affect genome stability and cell proliferative capacity. The most terminal region of telomeres exists as a single-stranded 3′ overhang of the G-rich strand that is believed to loop back and pair with the C-rich strand at an internal double-stranded location forming a stable loop structure—a “telomere cap” [2]. It has been shown that constitutive expression of the catalytic subunit of telomerase, hTERT, prevents the onset of replicative senescence and crisis by maintaining telomere length and function in several cell types including mesenchymal stem cells, fibroblasts and epithelial cells [3–7]. The telomere cap is therefore believed to be an important factor for cellular senescence [8]. The telomere is a dynamic nucleoprotein complex that can switch between capped and uncapped states. When the telomeres are in a capped “protected” state, the cells are allowed to proliferate whereas uncapped telomeres signal cell cycle arrest. Thus, the telomere capping status and not only the telomere length is believed to be important for telomere function. Ionizing radiation (IR) is a potent inducer of DNA doublestrand breaks (DSB) and has been shown to induce cellular growth arrest [9–11]. This stress-induced growth arrest has been suggested to be a premature form of senescence that closely resembles replicative senescence and has been termed stress-induced premature senescence (SIPS) [12]. It has been suggested that SIPS is induced by a p53-dependent cell cycle arrest via generation of non-specific as well as telomerespecific DNA damage [12]. Several studies have, however, shown that ionizing radiation does not lead to telomere shortening [1,13,14] and overexpression of hTERT is not able to protect the cells from SIPS [15]. In order to further study the role of telomere integrity and telomerase expression in relation to radiosensitivity, the short-term effect of ionizing radiation on human mesenchymal stem cells (MSC) derived from bone marrow stromal cells, stably transduced with the hTERT gene and their nontransduced counterpart has been investigated. Stem cells are unique in their ability to both self renew and to give rise to differentiated tissues. Presumably, stem cells have a higher degree of repair ability [16]. The human mesenchymal stem cells (MSCs) are multipotent precursors present in adult bone marrow and can differentiate into osteoblasts, adipocytes and myoblasts and play important roles in hematopoiesis. Many of these tissues of mesenchymal origin can give rise to cancer. Thus, these cells present a good model system for studying radiosensitivity and the consequences of this. In the present study, telomerase positive hMSCs with long telomeres and telomerase negative primary hMSCs with shorter telomeres were treated with varying doses from 2.5 to 15 Gy of γ-rays. The overall repair capabilities were similar in the primary and immortalized hMSCs. Upon IR, both cell types lost their telomere profile although the mean telomere length was unaltered. We interpret the loss of telomere profile as an indication of extensive telomeric damage, most likely indicating the occurrence of many telomere-near double-stranded breaks. We furthermore believe that cells with long telomeres
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and high telomerase activity in spite of this telomeric damage have an increased ability to rebuild the telomere cap as they showed lower frequency of anaphase bridges and lost telomeric signals than telomerase negative hMSCs.
Materials and methods Cell culture Primary hMSCs were aspirated from the bone marrow and isolated by centrifugation (700×g for 15 min at 4 °C) over a Ficoll Hypaque gradient (Sigma, St. Louis, MO, USA) as previously described [17]. The resulting cells were cultured as described below. Human mesenchymal stem cells immortalized with the hTERT gene at population doubling level 4 (PDL4) (hMSC-telo1) were generated by using a standard retroviral transduction protocol. In brief, the cDNA of the hTERT gene was inserted into the GCsam retroviral vector in which the transgene expression was driven by a Moloney murine leukemia virus long-terminal repeat. Thereafter, the construct was packaged in PG13 cells (American Type Culture Collection, Manassas, VA, USA) by a two-step procedure as previously described [18]. Normal and immortal hMSCs were cultivated in high glucose (4.5 g/l) Dulbecco's modified Eagle medium (D-MEM, GIBCO, Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal calf serum (GIBCO, Invitrogen), 100 U/ml of penicillin and streptomycin (GIBCO, Invitrogen) and 2 mM of L-glutamine.
Radiation of cells Cells were plated with a confluency of 25% and 48 h later irradiated by using a 137Cs radiator (Gammacelle 2000 RH, AEK Risø, Denmark) with varying doses from 2.5 to 15 Gy of γ rays delivered at a rate of 2.5 Gy/min. Cells were analyzed at different time points after irradiation. The culture medium was changed immediately after irradiation.
RNA extraction and RT-PCR analysis (hTERT, osteocalcin, alkaline phosphatase, collagen, β-actin) RNA was isolated from cells using the High Pure RNA Tissue Kit (Roche, Hvidovre, Denmark). cDNA was synthesized from RNA using the 1st Strand cDNA Synthesis Kit for RT-PCR (AMV) (Roche) according to manufacturer's instructions. Ectopic expression of hTERT was analyzed using ectoTERT primers (sense: 5′-GGACCATCTCTAGACTGACG-3′, antisense: 5′GGAGCGCACGGCTCGGCAGC-3′) and endogenous expression of hTERT was analyzed using endoTERT primers (sense: 5′CTGCTGCGCACTGGGAAGC-3′, antisense: 5′-GGACACCTGGCGGAAGGAG-3′). Expression of osteoblastic markers was analyzed using osteocalcin primers (sense: 5′-CATGAGAGCCCTCACA-3′, antisense: 5′-AGAGCGACACCCTAGAC-3′), alkaline phosphatase primers (sense: 5′-ACGTGGCTAAGAATGTCATC3′, antisense: 5′-CTGGTAGGCGATGTCCTTA-3′) and Collagen type 1 primers (sense: 5′-TGACGAGACCAAGAACTG-3′, antisense: 5′-CCATCCAAACCACTGAAACC-3′). β-Actin primers (sense: 5′-CTGTGCTGTCCCTGTATGCC-3′, antisense: 5′-GTGGTGGTGAAGCTGTAGCC-3′) were used in parallel reactions as a control of the cDNA preparation. Furthermore, β-actin primers
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were used in a PCR reaction performed on RNA to verify that no contaminating DNA was present in the RNA preparation. Amplification parameters for β-actin, ectoTERT, alkaline phosphatase and collagen type I were as follows: After an initial denaturation step at 94 °C for 3 min, amplification was performed for 30 cycles at 94 °C for 30 s, 59 °C for 30 s and 72 °C for 1 min, followed by a final extension step at 72 °C for 10 min. Osteocalcin primers used same conditions but an annealing temperature of 60 °C. Amplification parameters for endoTERT were as follows: After an initial denaturation step at 95 °C for 15 min, amplification was performed for 40 cycles at 94 °C for 30 s, 60 °C for 30 s and 72 °C for 1 min, followed by a final extension step at 72 °C for 10 min.
RNA extraction and RT-PCR analysis (CDKN1A, CDKN2A, β-actin) RNA was isolated from cells using the RNeasy Mini Kit (QIAGEN Nordic, Crawley, UK) and cDNA was synthesized using the High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA, USA) according to manufacturer's instructions. CDKN1A and CDKN2A expression were analyzed using pre-developed TaqMan® Gene Expression Assays (Hs00355782 _ m1 and Hs00233365 _ m1) and β-actin by the ACTB TaqMan® Endogenous Controls Assay Kit on an ABI 7000 (Applied Biosystems, Foster City, CA, USA).
TP53 mutation analysis
61 °C) for 30 s and 72 °C for 30 s). Primers used were as follows: CDKN2A Exon1Ase: GAAGAAAGAGGAGGGGCTG; Exon1Aas: GCGCTACCTGATTCCAATTC; Exon1Bse: TCCCAGTCTGCAGTTAAGG; Exon1Bas: GTCTAAGTCGTTGTAACCCG; Exon2se: GGAAATTGGAAACTGGAAGC; Exon2as: TCTGAGCTTTGGAAGCTCT; Exon3se: GACCTGGAGCGCTTGAGCGGT; Exon3as: CTGTAGGACCTTCGGTGACTGATGA; ACTB Exon4se: ACCCCAGCACACTTAGCCG; Exon4as: CACGATGCCAGTGGTACGG. Amplicons were analyzed on a 3% MetaPhor Agarose gel (Cambrex, East Rutherford, NJ, USA).
Osteoblast differentiation Three days after irradiation part of the cells were incubated with a new medium containing 5 × 10− 9 M calcitriol, (1,25-(OH)2 vitamin D3, Leo Pharma, Ballerup, Denmark) for 3 days. Afterwards, RNA was isolated from the cells for expression analysis as described above.
Proliferation, senescence and apoptosis assay Cell proliferation studies Cells were expanded by consecutive subcultivations in D-MEM with 10% FCS. Long-term cell growth in vitro was determined by calculating population doubling levels (PDL). The initial seeding number (Nstart) and the 80% confluence harvested cell number (Nfinish) were used to calculate the population doubling level PD = ln[(Nfinish) / (Nstart)] / ln2. Thus, the cumulative population doubling level is the sum of PDs.
Coding exons (2–11) of TP53 were amplified for 40 cycles (95 °C for 30 s, 62 °C for 30 s and 72 °C for 30 s). Primers used were: Exon02se: GACCCAGGGTTGGAAGCGTCTC; Exon02as: GGGGTCGGGGTGGTGGCC; Exon03se: CATGGGACTGACTTTCTGCTCTTG; Exon03as: CGGGGACAGCATCAAATCATC; Exon04Nse: GCTGGGGGGCTGAGGACC; Exon04Nas: GCCGCCGGTGAAAATAGGAGCTG; Exon04Cse: AGCTCCTACACCGGCGG; Exon04Cas: GCCAGGCATTGAAGTCTCATGG; Exon05Nse: TTCAACTCTGTCTCCTTCCTCTTCC; Exon05Nas: CAGCGCCTCACAACCTCCG; Exon05Cse: CGCGCCATGGCCATCTAC; Exon05Cas: CAGCGCCTCACAACCTCCG; Exon06se: GTCCCCAGGCCTCTGATTCC; Exon06as: CGGAGGGCCACTGACAACC; Exon07se: CAGGTCTCCCCAAGGCGCAC; Exon07as: GCAAGCAGAGGCTGGGGCAC; Exon08se: ACTGCCTCTTGCTTCTCTTTTCC; Exon08as: AATCTGAGGCATAACTGCACCC; Exon09se: GGGTGCAGTTATGCCTCAGATT; Exon09as: CGGCATTTTGAGTGTTAGACTGG; Exon10se: TGTATATACTTACTTCTCCCCCTCC; Exon10as: AGGGGAGTAGGGCCAGTAAGG; Exon11se: TGGTCAGGGAAAAGGGGCAC; Exon11as: GAGAGATGGGGGAGGGAGGC. All primers were extended at the 5' end with universal primer sequences; sense: CTCCTGTTCCGACCCTGCC and antisense: CGGAACAGGAGAGCGCACG. PCR products were purified using the QIAquick Gel Extraction Kit (Qiagen, West Sussex, UK), sequenced using the universal sequence primers and the BigDye DyeTerminator Cycle Sequencing Kit v. 1.1 and analyzed on an ABI 310 (Applied Biosystems).
Cells were first exposed to varying doses of radiation then washed in PBS, trypsinized and plated in 100 mm dishes in triplicate. Primary hMSCs were plated at about 10 cells per cm2 whereas hMSC-telo1 cells were plated at about 7 cells per cm2. After incubation for 14 days at 37 °C in 5% humidified CO2, the cells were washed twice with PBS, stained with 0.5% Crystal Violet in methanol for 5–10 min at room temperature and washed twice with PBS. Visible colonies were counted and the colony forming efficiency calculated in percent of plated cells.
CDKN2A genomic integrity analysis
Apoptosis assay (TUNEL)
All exons of CDKN2A and exon 4 of ACTB (β-actin) were amplified for 38 cycles (95 °C for 30 s, 58 °C (CDKN2A Exon1A:
Cells were seeded in 75 cm2 culture flasks and irradiated with different doses of γ rays. At different time points (4, 24 and 96 h after), cells were trypsinized, fixed in 4% paraformaldehyde
β-Galactosidase staining 2500 cells were seeded in slide flasks and irradiated 2 days later with different doses of γ rays. At different time points after irradiation (4, 7 and 10 days after), the cells were washed in PBS and fixed for 5 min at room temperature in 2% formaldehyde/0.2% glutaraldehyde. After washing, the cells were incubated at 37 °C (no CO2) over night with fresh β-gal solution containing 1 mg of 5-bromo-4-chloro-3-indolyl β-Dgalactoside (X-Gal, Sigma) per ml, 40 mM citric acid (pH 6.0), 5 mM potassium ferrocyanide, 5 mM potassium ferricyanide, 150 mM NaCl, 2 mM MgCl2. From each slide flask, minimum of 500 cells were counted under the microscope. The experiment was repeated 3 times and the mean percent of β-galactosidase-positive cells was then calculated.
CFU assay
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solution for 60 min on a shaker at room temperature and permeabilised in 0.1% Triton X-100/0.1% sodium citrate for 5 min at room temperature. Cells were stained using the In Situ Cell Death Detection Kit, Fluorescein (Roche) according to manufacturer's instructions. Green (FITC) fluorescence after 488 nm laser excitation was measured on the flow cytometer (BS FACSAria™ Cell Sorter, BD Biosciences, Erembodegem, Belgium).
DNA damage and repair assay (Fluorescence-Detected Alkaline DNA unwinding) DNA damage was measured as previously described [16]. The method detects single and double-strand breaks by their effect on the rate of DNA denaturation in alkali, monitored by the fluorescence intensity of an intercalating dye. The fluorescence intensity is inversely correlated to the number of DNA strand breaks present at the time of lysis. DNA damage percentage was calculated as 100 × (P0 − Px) / P0, with P0: fluorescence intensity of an unirradiated sample and Px: fluorescence intensity of the irradiated cell sample. Repair capacity was assayed by leaving the cells for the indicated time points (10′ till 4h), lysing them thereafter and compare them to the values immediately after irradiation.
H2AX Western analysis Cells were lysed in SDS lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris–HCl, pH 8.1) 1 h, 4 h and 1 day after irradiation and subjected to 2 rounds of freeze thawing on ethanol/dry ice. Following cell lysis, 40 μg protein was electrophoresed on NuPAGE 4–12% Bis-Tris polyacrylamide gels (Invitrogen) and transferred to nitrocellulose membranes. Membranes were incubated with mouse Anti-phospho-Histone H2A.X antibody (Upstate Biotechnology, Waltham, MA, USA) followed by incubation with goat anti-mouse HRP-conjugated secondary antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Immunoreactive bands were detected using the ECL detection system (Amersham Bioscience/GE Healthcare Bio-Sciences, Little Chalfont, UK). To assess protein loading, stripped membranes were re-probed using rabbit anti-Erk antibody (Santa Cruz Biotechnology).
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Normalization of telomere length values and Statistical analysis Telomere length values were normalized for each metaphase using the following equation: Z ¼ X=l where Z = normalized telomere length value; X = raw telomere length value; μ = mean telomere length of all telomeres in the metaphase. The rationale and normalization procedures were done as previously described in [19,20].
Telomere restriction fragment assay (TRF) Telomere lengths were determined using the Telo TAGGG Telomere Length Assay (Roche). Briefly, genomic DNA was embedded in 1% agarose plugs and a plug containing 3 μg of genomic DNA was digested with the restriction enzymes HinfI and RsaI. Plugs were analyzed in a 1% agarose gel at 14 °C by pulsed field gel electrophoresis (CHEF-DRII-SYSTEM, Biorad, Herlev, Denmark) at 3.5 V/cm for 17 h at a ramp pulse of 2–10 s. The DNA was Southern-blotted to charged nylon membranes (Hybond-N+, Amersham Bioscience) and hybridized with a digoxenin (DIG)-labelled TTAGGG probe. The average TRF lengths were determined by comparing the signals relative to a molecular weight standard.
Chromosome analysis The cells were cultured in slide flasks (Nunc, Rochester, NY, USA). Before harvest, 10 mg/ml Colcemid (GIBCO, Invitrogen) was added to a final concentration of 0.2 μg/ml and culturing continued for another 35 min. The cells were lysed by adding 60 mM KCl and thereafter standard harvesting took place including the Carnoy's fixative. Giemsa bands were produced with Wright's stain. From each cell type and dose 45 metaphases were analyzed.
Results Experimental system
Telomere profiles The telomere profile was measured on chromosomes prepared as described below. A telomere FISH kit (DakoCytomation, Glostrup, Denmark) was used for telomere staining according to manufactures instructions. For subsequent karyotyping, the DNA was stained with a DAPI solution (0.5 μg/ml) also containing antifade (0.11% phenylenediamine dihydrochloride).
Image acquisition and analysis Fluorescent-specific signals were visualized by fluorescence microscopy and captured at 100× magnification with a cooled CCD camera using IpLab software (Scanalytics, Fairfax, VA, USA). For each sample, twenty metaphases were analyzed. Image acquisition and analysis were done as previously described in [19,20].
To explore the roles of telomeres and telomerase activity in relation to radiosensitivity, we investigated the effect of ionizing radiation on normal primary human mesenchymal stem cells (hMSC) and telomerase-immortalized hMSCs (hMSCtelo1) established by transducing normal primary hMSCs with a retrovirus carrying the hTERT gene. Analysis of the endogenous and ectopic expression of hTERT showed that neither primary hMSC nor the hMSC-telo1 line expressed endogenous hTERT; however, the immortalized hMSC-telo1 cells expressed high levels of ectopic hTERT that resulted in an increased mean telomere length of 21 kb already at PDL 43 while the mean telomere length of the primary cells was 8 kb (Fig. 1A).
Radiation of hMSC and hMSC-telo1 cells Normal primary (hMSC) and immortal hMSCs (hMSC-telo1) at PDL 114 were treated with varying doses from 2.5 to 15 Gy of γ
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Fig. 1 – (A) TRF analysis of primary hMSC and telomerase-immortalized hMSC (hMSC-telo1 cells) at the indicated PD levels. TRF lengths were determined as described in Materials and methods. Molecular size markers (kb) are indicated to the left of the blot. (B, C) Standard RT-PCR analysis of hMSC and hMSC-telo1 cells. (B) The figure illustrates non-irradiated cells and cells irradiated with 2.5 and 5 Gy of γ-rays. RNA was isolated from the cells and subjected to RT-PCR using primers amplifying the ectopic hTERT gene and the β-actin gene. A sample without template was included as a negative control. (C) The figure shows cells irradiated with a 5-Gy dose (+) or non-irradiated (−). Three days after IR, some of the cells were treated with VitD3 (+). RNA was isolated from the cells and subjected to RT-PCR using primers amplifying a region in the osteocalcin gene (Ost), the alkaline phosphatase gene (AP), the collagen type I gene (Col-1) and in the β-actin gene. A sample without template was included as a negative control. PCR was performed on RNA to verify that no DNA was present in the RNA preparation (data not shown). (D) Induction of p21CIP1 (CDKN1A) expression following irradiation. RNA was isolated from the cells and subjected to real-time RT-PCR. Expression was measured relative to β-actin before (Control) and 4 h after irradiation in hMSC (light gray) and hMSC-telo1 cells (dark gray) with doses from 2.5 to 15 Gy. Data shown are mean ± SEM for three independent experiments. (E) PCR analysis on genomic DNA for CDKN2A copy number analysis. Figure shows pooled PCR products from each of the different cell lines and passages.
rays. Irradiation of normal hMSC and hMSC-telo1 cells did not lead to immediate cell death, not even at the highest dose.
either hMSC or hMSC-telo1 cells (data not shown) or modulated the expression of ectopic hTERT in hMSC-telo1 cells (Fig. 1B).
Endogenous and ectopic hTERT expression
Expression of differentiation markers
The endogenous and ectopic expression of hTERT was analyzed after irradiation of the cells by standard RT-PCR analysis. Irradiation did not induce expression of endogenous TERT in
It has previously been shown that introduction of hTERT into primary hMSCs does not change the ability of the cells to differentiate into various cell types [3,21]. After irradiation, the
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cells were incubated in medium containing 1,25-(OH)2-vitamin D3 (vitamin D3) that induces differentiation along the osteogenic pathway. The expression of different osteoblastic markers (osteocalcin (Ost), alkaline phosphatase (AP) and collagen type I (Col-1)) was analyzed by standard RT-PCR analysis. As seen in Fig. 1C, presence of vitamin D3 in the culture medium leads to increased expression of osteocalcin in both primary hMSC and hMSC-telo1 cells before irradiation. The expression levels of AP and Col-1 were at the same level with or without vitamin D3. Irradiation of the cells did not change the ability of the cells to express these markers (Fig. 1C).
Wild-type p53 but inactivation of p16Ink4A and p14ARF DNA sequence analysis of the coding regions of the p53 gene, TP53, was wild type in both hMSC and hMSC-telo1 (data not shown). Four hours after irradiation, CDKN1A (p21CIP1) expression was induced in hMSC and hMSC-telo1, indicating that the p53 protein was functioning normally in both cell lines (Fig. 1D). CDKN2A (p16Ink4A) expression could not be detected in hMSC-telo1 (data not shown) and DNA copy number analysis showed deletion of both copies of CDKN2A (Fig. 1E). Thus, p16Ink4A and p14ARF are fully inactivated in hMSC-telo1, whereas active in hMSC.
Proliferation and survival Irradiation of primary hMSC and hMSC-telo1 cells resulted in a dose-dependent growth arrest (Figs. 2A and B). At the high doses (10 and 15 Gy), many cells, both hMSC and hMSC-telo1, acquired an enlarged flat morphology typical of senescent cells after 3–5 days. Analysis of the colony forming ability (CFU assay) after irradiation demonstrated that both primary hMSCs and hMSC-telo1 cells showed a decrease in the CFUs already at 2.5 Gy and at the high dose (15 Gy) no colonies were formed within 14 days in either of the cell types (Fig. 2C) meaning that both primary hMSCs and hMSC-telo1 cells already showed considerable cell cycle arrest at the low dose 2.5 Gy. hMSCs irradiated with the higher doses (10–15 Gy) did not start to divide within 3 months following irradiation whereas few dividing cells were observed 1–2 weeks after irradiation with the 2.5-Gy dose as demonstrated by the CFU assay after 2 weeks. hMSC-telo1 cells that received the high doses (10–15 Gy) started to divide 2–3 weeks after irradiation. Thus, hMSC-telo1 cells required shorter time (days in culture) to return to cell cycle than hMSCs.
DNA-damage and phosphorylation of H2AX Irradiation-induced DNA damage/repair was analyzed by Fluorescence-detected alkaline DNA unwinding and it was found that irradiation induced the same extent of DNA damage in both cell types (Figs. 3A and B) and the amount of damage increased with the dose. After 4 h, the DNA damage percentage in hMSC and hMSC-telo1 cells receiving both the low (2.5 Gy) and the high (15 Gy) doses had decreased to a level of 20% damage indicating that the two cell types did not differ in their overall genomic repair capabilities. DNA damage was furthermore investigated by measuring the kinetics of H2AX phosphorylation after subjecting the cells
Fig. 2 – Growth curve of primary hMSC (A) and the telomerase-immortalized hMSC-telo1 cells (B) after irradiation. Cells were plated and treated with different doses of γ-rays: 2.5 Gy (■), 5 Gy (▲), 10 Gy (×) and 15 Gy (●) or non-treated cells (O). The cells were irradiated at the indicated time point and the cumulative PD levels followed. (C) CFU assay of primary hMSC (●) and hMSC-telo1 cells (■). Cells were irradiated with 2.5 or 15 Gy of γ-rays, plated in 100 mm dishes and scored for colony formation after 14 days. Data shown are mean ± SD.
to IR. For both hMSC and hMSC-telo1 cells the phosphorylation of H2AX increased in a dose-dependent way upon irradiation with doses from 2.5 to 15 Gy of γ-rays (Fig. 3C). In hMSC-telo1 cells, the level of H2AX phosphorylation was decreased
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Fig. 4 – Beta-galactosidase-associated senescence in irradiated primary hMSC (grey lines) and telomerase-immortalized hMSC-telo1 cells (black lines). Cells were irradiated with 2.5 (dotted line) or 15 Gy (broken line) of γ-rays and stained with X-gal substrate 4, 7 and 10 days after irradiation. Control cells were not irradiated (full line). The percentage of blue cells is given in the figure. The data shown are mean percent ± SD estimated from a binomial distribution for three experiments.
considerable 4 h after irradiation, whereas in hMSCs this was not the case. One day after, no phosphorylation could be detected in either of the cell types (data not shown). This finding was in accordance with the DNA damage/repair analysis performed by fluorescence activated DNA unwinding (FADU).
Senescence-associated β-galactosidase and TUNEL assays
Fig. 3 – DNA damage in primary hMSC (A) and telomerase-immortalized hMSC-telo1 cells (B) after irradiation. DNA damage and repair was measured by a fluorescence-detected alkaline DNA unwinding assay in 3 independent experiments. The cells were trypsinized and irradiated with doses from 2.5 to 15 Gy of γ-rays. At different time points, the initial and the remaining damage (remdam) in the cells were analyzed. Error bars are standard errors. (C) Western blot analysis of induction of H2AX phosphorylation in irradiated hMSC and hMSC-telo1 cells. Protein extracts were prepared from non-irradiated cells or cells irradiated with doses from 2.5 to 15 Gy of γ-rays. Size markers are indicated to the left of the blot. ERK staining of the same samples was used as loading control.
The proportion of senescent cells after irradiation was analyzed by staining the cells for senescence-associated β-galactosidase (SA β-gal). Before irradiation, 12%± 0.8% of normal hMSCs and 2% ± 0.4% of hMSC-telo1 cells showed positive staining for SA βgal (Fig. 4). Irradiation increased the proportion of SA β-gal positive cells in a dose-dependent way in both hMSC and hMSCtelo1 cells. Ten days after irradiation, a proportion of 91% ± 0.6% and 97% ± 0.4% of the hMSCs stained positive for SA β-gal at the low (2.5 Gy) and high (15 Gy) doses, respectively. However, in hMSC-telo1 cells only 30% ± 1% and 76%± 1% of the cells showed SA β-gal positive staining 10 days after irradiation with 2.5 Gy and 15 Gy of γ-rays, respectively (Fig. 4). The TUNEL assay was used to measure the proportion of apoptotic cells in the cell cultures. Before irradiation, 0.25% ± 0.14% (n = 3) and 0.10% ± 0.03% (n = 3) apoptotic cells were observed in hMSC and hMSC-telo1 cell cultures, respectively. Irradiation of hMSC and hMSC-telo1 cells with 2.5 and 15 Gy of γrays did not lead to increased apoptotic cell death when measured 4, 24 and 96 h after irradiation by TUNEL assay (data not shown).
Telomere length and profile To investigate the effect of irradiation on the telomere length, we performed a TRF analysis. Neither hMSC nor hMSC-telo1 cells irradiated with doses from 2.5 to 15 Gy exhibited detectable telomere shortening when measured up to 10 days after irradiation (Fig. 5A).
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Fig. 5 – Telomere length and profile analysis. (A) The TRF length was determined in primary hMSC (lanes 2–14) and telomerase-immortalized hMSC-telo1 cells (lanes 15–27) 1, 4 and 10 days after irradiation of the cells with doses from 2.5 to 15 Gy of γ-rays. TRF lengths were determined as described in Materials and methods. Molecular size markers (kb) are indicated to the left of the blot. (B) Comparison of the telomere profiles in two different unirradiated hMSC-telo1 samples at PDL 128. (C) Comparison of the telomere profiles in two different hMSC-telo1 samples irradiated with 15 Gy. (D, E) The telomere profiles of hMSC-telo1 cells irradiated with 2.5 Gy (D) and 15 Gy (E) of γ-rays were compared to the telomere profile of non-irradiated hMSC-telo1 cells. The mean values of normalized telomere lengths from 20 metaphases are assigned to the y-axis. Error bars describe the 95% confidence limits for the mean.
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Fig. 5 (continued ).
The effect of irradiation on the cells telomere profile was measured by using a PNA-FISH method and computer-assisted telomere quantifier software. Analysis of hMSCs after irradiation was hampered by the difficulties of obtaining metaphases due to low cell cycling. As previously shown by our group, the hMSC and hMSC-telo1 cells have a telomere profile as first described in lymphocytes (Graakjær et al, submitted work). A high and significant correlation is obtained when the telomere profiles of two hMSC-telo1 samples from the same passage (PDL 128: correlation = 0.71, p < 0.001, Fig. 5B) or from different passages (PDL 43 compared to PDL 167: correlation = 0.62, p < 0.001) are compared indicating that the telomere profiles of un-irradiated samples are stable even after long-term cell growth. However, no significant correlation is found when comparing the telomere profiles of two samples irradiated with the same dose (hMSC-telo1 irradiated with 15 Gy: correlation = −0.06, p = 0.63, Fig. 5C) indicating that the telomere length pattern is not reproducible after irradiation. Comparison of the telomere profiles before and after irradiation show that the telomere profile is lost in hMSC-telo1 cells at both low (2.5 Gy) and high doses (15 Gy) (Figs. 5D and E). The difference in profile changes between 2.5 and 15 Gy can be explained by the fact that irradiation destroys the telomere length pattern in a random manner and the cells that establish the cell culture after irradiation have different telomere profiles.
Chromosome analyses After irradiation, random chromosomal aberrations have been observed but no clonal aberration except in 15 Gy
irradiated hMSC-telo1 cells. hMSCs could only be analyzed at the low dose 2.5 Gy because cells receiving higher doses did not proliferate after irradiation. After irradiation with a 2.5Gy dose, hMSCs showed a 3-to 4-fold increase in anaphase bridge/micronuclei formation, while hMSC-telo1 cells only showed a 0- to 1-fold increase. In addition, it was observed that the frequency of lost telomeric signals after irradiation with a 2.5-Gy dose had increased 2- to 3-fold in hMSCs but only 0- to 1-fold in hMSC-telo1 cells. Overall, these results indicate that cells with long telomeres due to high telomerase expression show higher stability at telomeric regions than cells with shorter telomeres and no telomerase expression.
Discussion In this study, we have investigated the radiosensitivity of telomerase-immortalized hMSCs versus normal primary hMSCs. The hMSC-telo1 cells have high telomerase activity and long telomeres compared to their normal counterpart hMSCs that have shorter telomeres and no telomerase activity. Irradiation of both cell types resulted in a dosedependent growth arrest, indicating that both cell types have at least one intact DNA damage control pathway. This assumption is further supported by the finding that both cell types had functional p53, although telomerase-transduced cells as expected contained inactivating mutations in the CDKN2A gene, resulting in lack of p16 and p14 protein. It seems likely that compromising the p16-checkpoint increased the
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ability to extend cellular life span while maintaining the diploid status of telomerase-immortalized cells. hMSC-telo1 cells returned to cell cycle within a few weeks after irradiation even after subjected to the high dose (15 Gy). In contrast, primary hMSCs receiving the same high dose did not return to cell cycle within 3 months of radiation. The faster recovery after irradiation of telomerase-immortalized cells might be explained by the fact that these cells do not express p16 since this may result in these cells returning to cell cycle in spite of remaining DNA damage. A number of other observations point, however, towards telomeres as a central player in the induction and subsequent release from cell cycle arrest. It is well established that radiation using γ rays induces singlestrand and double-strand breaks (DSB) in the DNA and that the cell cycle machinery arrests the cells until the damage has been repaired [9–11]. Our data show that the level of overall DNA damage/repair did not differ between hMSC and hMSCtelo1 cells after irradiation with the same doses and the amount of damage remaining after 4 h was the same in the two cell types indicating that the cells had similar repair capabilities. This result was confirmed by analysis of phosphorylation of histone H2AX. Upon irradiation, H2AX imme-
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diately becomes phosphorylated on Ser-139 and is believed to recruit DNA repair factors to the site of DNA double-strand breaks [22]. Hence, the return of H2AX to its non-phosphorylated state within a day also suggests that overall DNA repair in both cell systems are finalized within a day. Thus, DNA repair is not dependent on telomere length and/or telomerase expression in our cell system. Likewise, it has been shown that double-strand break repair induced by γ-radiation is identical in normal and telomerase-immortalized human foreskin fibroblasts [23] and furthermore that repair of pyrimidine dimers and dimethylsulfate damage induced by ultraviolet radiation or dimethylsulfate is identical in telomerase-immortalized human skin fibroblasts and normal skin fibroblasts [24]. In spite that overall DNA repair was finalized within a day even with the high doses of γ-rays both cell types remained in cell cycle arrest for weeks to months. Since all the hallmarks of stress-induced senescence are present in the cells, this finding very strongly suggests that part of the DNA was not repaired within these first hours and that this remaining damage maintains the cells in cell cycle arrest. We have in this connection obtained results indicating that radiation leads to
Fig. 6 – Illustration of our current hypothesis on p53 and p16 pathways in our cell system.
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extensive damage of the telomeres. Since in general telomeric DNA is less efficiently repaired than other parts of the genome [25], it can therefore be imagined that the remaining DNA damage is associated with the telomeres. The loss of telomere profile indicates telomere damage, namely suggesting primary occurrence of double-strand breaks in the telomeric regions. At first glance it may seem puzzling that the relatively few damaged/dysfunctional telomeres per cell can cause such an extensive loss of profile. We hypothesize that the extensive profile loss is a consequence of not only the initial telomere damage, but also of subsequent fusion–break–fusion cycles, occurring after the cells have re-entered the cell cycle, resulting in substantial re-shuffling of telomeric regions, until finally all chromosome ends have achieved stable telomere coverage, at which point fully normal cell proliferation is established, but with a different profile. It should be pointed out, as also previously stated [26], that full development of this cascade of events require that cells have not totally left the cell cycle, since fusion–break–fusion cycles require at least some degree of cell division. This condition is, however, fulfilled for the cells where we analyzed telomere profiles, simply because these cells go into mitosis, which means that they are cycling. Surprisingly, the overall telomere length did not decrease after irradiation. This might, however, be explained by the appearance of fusion breakage fusion cycles that would lead to rearrangements, but not necessarily result in telomeric loss. It may thus be possible that irradiation causes destabilization of the telomeric loop and uncapping of telomeres without initial double-strand breaks. This then leads to fusion breakage fusion cycles without net loss of telomere sequences. In agreement with this, it has previously been shown that irradiation can induce senescence without reducing telomere length [13–15] also suggesting that one or more aspects of telomeres other than length per se contribute to the response. Furthermore, Ku86−/− mouse embryonic fibroblasts with long but dysfunctional telomeres resulting from loss of their endcapping function showed a senescence-like arrest [27]. Our results thus support the hypothesis that telomere dysfunction can induce stress-induced premature senescence by a telomere-length-independent mechanism most likely involving loss of the telomere capping function. It has previously been shown that dysfunctional, uncapped telomeres associate with γ-H2AX [28]. In our study, when the irradiated cells were stained with antibodies against H2AX and TRF1 we also observed that there was a tendency of colocalization at the telomere sides. Furthermore, a small proportion of the H2AX foci were remaining after a day. The appearance of anaphase bridges can be used as a measure of telomere dysfunction [29]. Based on the fact that the hMSC-telo1 cells have fewer cells positive for SA β-gal stain and a lower frequency of anaphase bridges and lost telomeric signals, it can be suggested that cells with long telomeres and high telomerase activity re-establish telomeric function more efficiently than cells with shorter telomeres and no telomerase expression and that this ability is important for returning to cell cycle. This is in agreement with previous reports, where it has been shown that telomerase has a protective effect on very short telomeres that, in its absence, would have caused cells to stop dividing. Thus, active
telomerase helps even very short telomeres to be functionally capped [8,30,31]. Furthermore, it has been indicated that longer telomeres are more likely to rebuild the cap than are shorter telomeres [32,33]. It has recently been shown that fibroblasts expressing hTERT showed high level of stability even after exposure to ionizing radiation [34]. In agreement with this, our cell systems showed low sensitivity to radiation. Clearly, hMSCtelo1 cells showed a higher survival effect than hMSCs, which might be explained by a telomerase survival effect. Overall, our data suggest that uncapping of telomeres induced by irradiation plays a role in cell cycle depression. Furthermore, our results are compatible with the hypothesis that the signaling pathway goes through p53 and not p16. Fig. 6 illustrates our current hypothesis with regards to signaling pathways in our cell system. It is here illustrated, that we, like others [35–38] believe that p16 and p53 are located in two different pathways, and that telomere damage is signaled through the pathway involving p53. Indirect support for this latter notion comes from the fact that telomerase-immortalized cells often have inactivation of p16 suggesting that telomere elongation and loss of p16 are two independent events in the immortalization process [36–38]. In addition, we find that irradiation does not change the differentiation ability of mesenchymal stem cells. Overall, our observations point out some of the key properties of these cell populations that allow them to survive under high doses of γ-rays.
Acknowledgments The authors wish to thank Anette Thomsen, Christian Knudsen, Bente Kierkegaard, Inger Marie Thuesen, Alan Leake and Sharon Burns for their excellent technical assistance. The work was supported by Danish Cancer Society, Cancer Research UK, Glasgow University and European Union contracts EC-FP5-FI6R-CT-2002-0021 (TELOSENS) and LSHCCT-2004-502943 (MOL Cancer Med).
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a v a i l a b l e a t w w w. s c i e n c e d i r e c t . c o m
w w w. e l s e v i e r. c o m / l o c a t e / y e x c r
Research Article
Ectopically hTERT expressing adult human mesenchymal stem cells are less radiosensitive than their telomerase negative counterpart Nedime Serakinci a,b,⁎,1 , Rikke Christensen a,1 , Jesper Graakjaer f , Claire J. Cairney c , W. Nicol Keith c , Jan Alsner d , Gabriele Saretzki e , Steen Kolvraa f a
Department of Human Genetics, University of Aarhus, Aarhus, Denmark Institute of Medical Biology, Department of Anatomy and Neurobiology, Southern Denmark University, Odense, Denmark c Centre for Oncology and Applied Pharmacology, University of Glasgow, Cancer Research UK, Beatson Laboratories, Garscube Estate, Glasgow, UK d Department of Experimental Clinical Oncology, Aarhus University Hospital, Aarhus, Denmark e Henry Wellcome Laboratory for Biogerontology, Newcastle General Hospital, University of Newcastle upon Tyne, Newcastle, UK f Department of Clinical Genetics, Vejle County Hospital, Vejle, Denmark b
ARTICLE INFORMATION
ABS T R AC T
Article Chronology:
During the past several years increasing evidence indicating that the proliferation capacity
Received 30 June 2006
of mammalian cells is highly radiosensitive, regardless of the species and the tissue of origin
Revised version received
of the cells, has accumulated. It has also been shown that normal bone marrow cells of mice
21 December 2006
have a similar radiosensitivity to other mammalian cells so far tested. In this study, we
Accepted 3 January 2007
investigated the genetic effects of ionizing radiation (2.5–15 Gy) on normal human
Available online 8 January 2007
mesenchymal stem cells and their telomerised counterpart hMSC-telo1. We evaluated overall genomic integrity, DNA damage/repair by applying a fluorescence-detected alkaline
Keywords:
DNA unwinding assay together with Western blot analyses for phosphorylated H2AX and
Telomere dysfunction
Q-FISH was applied for investigation of telomeric damage. Our results indicate that hMSC
Telomerase
and TERT-immortalized hMSCs can cope with relatively high doses of γ-rays and that
Mesenchymal stem cells
overall DNA repair is similar in the two cell lines. The telomeres were extensively destroyed
Ionizing radiation
after irradiation in both cell types suggesting that telomere caps are especially sensitive to
p16INK4a
radiation. The TERT-immortalized hMSCs showed higher stability at telomeric regions than
Telomere capping
primary hMSCs indicating that cells with long telomeres and high telomerase activity have
p21
the advantage of re-establishing the telomeric caps. © 2007 Elsevier Inc. All rights reserved.
Introduction Lack of telomere maintenance is a genetic feature that appears to be associated with multiple genetic defects, all
conferring radiosensitivity. Human radiosensitive cell lines have thus been shown either to have short or dysfunctional telomeres and in a mouse model it has been shown directly that mice with critically short telomeres are more
⁎ Corresponding author. Institute of Medical Biology, Department of Anatomy and Neurobiology, Southern Denmark University, Winsløvparken 21, 5000 Odense C, Denmark. Fax: +45 6590 6321. E-mail addresses: [email protected], [email protected] (N. Serakinci). 1 Authors equally contributed to the 1st authorship of the work. 0014-4827/$ – see front matter © 2007 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2007.01.002
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radiosensitive to ionizing radiation (gamma rays) and show an increased chromosomal instability [1]. Telomeres are the physical ends of chromosomes and are responsible for the maintenance of chromosome stability and therefore dysfunctional telomeres affect genome stability and cell proliferative capacity. The most terminal region of telomeres exists as a single-stranded 3′ overhang of the G-rich strand that is believed to loop back and pair with the C-rich strand at an internal double-stranded location forming a stable loop structure—a “telomere cap” [2]. It has been shown that constitutive expression of the catalytic subunit of telomerase, hTERT, prevents the onset of replicative senescence and crisis by maintaining telomere length and function in several cell types including mesenchymal stem cells, fibroblasts and epithelial cells [3–7]. The telomere cap is therefore believed to be an important factor for cellular senescence [8]. The telomere is a dynamic nucleoprotein complex that can switch between capped and uncapped states. When the telomeres are in a capped “protected” state, the cells are allowed to proliferate whereas uncapped telomeres signal cell cycle arrest. Thus, the telomere capping status and not only the telomere length is believed to be important for telomere function. Ionizing radiation (IR) is a potent inducer of DNA doublestrand breaks (DSB) and has been shown to induce cellular growth arrest [9–11]. This stress-induced growth arrest has been suggested to be a premature form of senescence that closely resembles replicative senescence and has been termed stress-induced premature senescence (SIPS) [12]. It has been suggested that SIPS is induced by a p53-dependent cell cycle arrest via generation of non-specific as well as telomerespecific DNA damage [12]. Several studies have, however, shown that ionizing radiation does not lead to telomere shortening [1,13,14] and overexpression of hTERT is not able to protect the cells from SIPS [15]. In order to further study the role of telomere integrity and telomerase expression in relation to radiosensitivity, the short-term effect of ionizing radiation on human mesenchymal stem cells (MSC) derived from bone marrow stromal cells, stably transduced with the hTERT gene and their nontransduced counterpart has been investigated. Stem cells are unique in their ability to both self renew and to give rise to differentiated tissues. Presumably, stem cells have a higher degree of repair ability [16]. The human mesenchymal stem cells (MSCs) are multipotent precursors present in adult bone marrow and can differentiate into osteoblasts, adipocytes and myoblasts and play important roles in hematopoiesis. Many of these tissues of mesenchymal origin can give rise to cancer. Thus, these cells present a good model system for studying radiosensitivity and the consequences of this. In the present study, telomerase positive hMSCs with long telomeres and telomerase negative primary hMSCs with shorter telomeres were treated with varying doses from 2.5 to 15 Gy of γ-rays. The overall repair capabilities were similar in the primary and immortalized hMSCs. Upon IR, both cell types lost their telomere profile although the mean telomere length was unaltered. We interpret the loss of telomere profile as an indication of extensive telomeric damage, most likely indicating the occurrence of many telomere-near double-stranded breaks. We furthermore believe that cells with long telomeres
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and high telomerase activity in spite of this telomeric damage have an increased ability to rebuild the telomere cap as they showed lower frequency of anaphase bridges and lost telomeric signals than telomerase negative hMSCs.
Materials and methods Cell culture Primary hMSCs were aspirated from the bone marrow and isolated by centrifugation (700×g for 15 min at 4 °C) over a Ficoll Hypaque gradient (Sigma, St. Louis, MO, USA) as previously described [17]. The resulting cells were cultured as described below. Human mesenchymal stem cells immortalized with the hTERT gene at population doubling level 4 (PDL4) (hMSC-telo1) were generated by using a standard retroviral transduction protocol. In brief, the cDNA of the hTERT gene was inserted into the GCsam retroviral vector in which the transgene expression was driven by a Moloney murine leukemia virus long-terminal repeat. Thereafter, the construct was packaged in PG13 cells (American Type Culture Collection, Manassas, VA, USA) by a two-step procedure as previously described [18]. Normal and immortal hMSCs were cultivated in high glucose (4.5 g/l) Dulbecco's modified Eagle medium (D-MEM, GIBCO, Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal calf serum (GIBCO, Invitrogen), 100 U/ml of penicillin and streptomycin (GIBCO, Invitrogen) and 2 mM of L-glutamine.
Radiation of cells Cells were plated with a confluency of 25% and 48 h later irradiated by using a 137Cs radiator (Gammacelle 2000 RH, AEK Risø, Denmark) with varying doses from 2.5 to 15 Gy of γ rays delivered at a rate of 2.5 Gy/min. Cells were analyzed at different time points after irradiation. The culture medium was changed immediately after irradiation.
RNA extraction and RT-PCR analysis (hTERT, osteocalcin, alkaline phosphatase, collagen, β-actin) RNA was isolated from cells using the High Pure RNA Tissue Kit (Roche, Hvidovre, Denmark). cDNA was synthesized from RNA using the 1st Strand cDNA Synthesis Kit for RT-PCR (AMV) (Roche) according to manufacturer's instructions. Ectopic expression of hTERT was analyzed using ectoTERT primers (sense: 5′-GGACCATCTCTAGACTGACG-3′, antisense: 5′GGAGCGCACGGCTCGGCAGC-3′) and endogenous expression of hTERT was analyzed using endoTERT primers (sense: 5′CTGCTGCGCACTGGGAAGC-3′, antisense: 5′-GGACACCTGGCGGAAGGAG-3′). Expression of osteoblastic markers was analyzed using osteocalcin primers (sense: 5′-CATGAGAGCCCTCACA-3′, antisense: 5′-AGAGCGACACCCTAGAC-3′), alkaline phosphatase primers (sense: 5′-ACGTGGCTAAGAATGTCATC3′, antisense: 5′-CTGGTAGGCGATGTCCTTA-3′) and Collagen type 1 primers (sense: 5′-TGACGAGACCAAGAACTG-3′, antisense: 5′-CCATCCAAACCACTGAAACC-3′). β-Actin primers (sense: 5′-CTGTGCTGTCCCTGTATGCC-3′, antisense: 5′-GTGGTGGTGAAGCTGTAGCC-3′) were used in parallel reactions as a control of the cDNA preparation. Furthermore, β-actin primers
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were used in a PCR reaction performed on RNA to verify that no contaminating DNA was present in the RNA preparation. Amplification parameters for β-actin, ectoTERT, alkaline phosphatase and collagen type I were as follows: After an initial denaturation step at 94 °C for 3 min, amplification was performed for 30 cycles at 94 °C for 30 s, 59 °C for 30 s and 72 °C for 1 min, followed by a final extension step at 72 °C for 10 min. Osteocalcin primers used same conditions but an annealing temperature of 60 °C. Amplification parameters for endoTERT were as follows: After an initial denaturation step at 95 °C for 15 min, amplification was performed for 40 cycles at 94 °C for 30 s, 60 °C for 30 s and 72 °C for 1 min, followed by a final extension step at 72 °C for 10 min.
RNA extraction and RT-PCR analysis (CDKN1A, CDKN2A, β-actin) RNA was isolated from cells using the RNeasy Mini Kit (QIAGEN Nordic, Crawley, UK) and cDNA was synthesized using the High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA, USA) according to manufacturer's instructions. CDKN1A and CDKN2A expression were analyzed using pre-developed TaqMan® Gene Expression Assays (Hs00355782 _ m1 and Hs00233365 _ m1) and β-actin by the ACTB TaqMan® Endogenous Controls Assay Kit on an ABI 7000 (Applied Biosystems, Foster City, CA, USA).
TP53 mutation analysis
61 °C) for 30 s and 72 °C for 30 s). Primers used were as follows: CDKN2A Exon1Ase: GAAGAAAGAGGAGGGGCTG; Exon1Aas: GCGCTACCTGATTCCAATTC; Exon1Bse: TCCCAGTCTGCAGTTAAGG; Exon1Bas: GTCTAAGTCGTTGTAACCCG; Exon2se: GGAAATTGGAAACTGGAAGC; Exon2as: TCTGAGCTTTGGAAGCTCT; Exon3se: GACCTGGAGCGCTTGAGCGGT; Exon3as: CTGTAGGACCTTCGGTGACTGATGA; ACTB Exon4se: ACCCCAGCACACTTAGCCG; Exon4as: CACGATGCCAGTGGTACGG. Amplicons were analyzed on a 3% MetaPhor Agarose gel (Cambrex, East Rutherford, NJ, USA).
Osteoblast differentiation Three days after irradiation part of the cells were incubated with a new medium containing 5 × 10− 9 M calcitriol, (1,25-(OH)2 vitamin D3, Leo Pharma, Ballerup, Denmark) for 3 days. Afterwards, RNA was isolated from the cells for expression analysis as described above.
Proliferation, senescence and apoptosis assay Cell proliferation studies Cells were expanded by consecutive subcultivations in D-MEM with 10% FCS. Long-term cell growth in vitro was determined by calculating population doubling levels (PDL). The initial seeding number (Nstart) and the 80% confluence harvested cell number (Nfinish) were used to calculate the population doubling level PD = ln[(Nfinish) / (Nstart)] / ln2. Thus, the cumulative population doubling level is the sum of PDs.
Coding exons (2–11) of TP53 were amplified for 40 cycles (95 °C for 30 s, 62 °C for 30 s and 72 °C for 30 s). Primers used were: Exon02se: GACCCAGGGTTGGAAGCGTCTC; Exon02as: GGGGTCGGGGTGGTGGCC; Exon03se: CATGGGACTGACTTTCTGCTCTTG; Exon03as: CGGGGACAGCATCAAATCATC; Exon04Nse: GCTGGGGGGCTGAGGACC; Exon04Nas: GCCGCCGGTGAAAATAGGAGCTG; Exon04Cse: AGCTCCTACACCGGCGG; Exon04Cas: GCCAGGCATTGAAGTCTCATGG; Exon05Nse: TTCAACTCTGTCTCCTTCCTCTTCC; Exon05Nas: CAGCGCCTCACAACCTCCG; Exon05Cse: CGCGCCATGGCCATCTAC; Exon05Cas: CAGCGCCTCACAACCTCCG; Exon06se: GTCCCCAGGCCTCTGATTCC; Exon06as: CGGAGGGCCACTGACAACC; Exon07se: CAGGTCTCCCCAAGGCGCAC; Exon07as: GCAAGCAGAGGCTGGGGCAC; Exon08se: ACTGCCTCTTGCTTCTCTTTTCC; Exon08as: AATCTGAGGCATAACTGCACCC; Exon09se: GGGTGCAGTTATGCCTCAGATT; Exon09as: CGGCATTTTGAGTGTTAGACTGG; Exon10se: TGTATATACTTACTTCTCCCCCTCC; Exon10as: AGGGGAGTAGGGCCAGTAAGG; Exon11se: TGGTCAGGGAAAAGGGGCAC; Exon11as: GAGAGATGGGGGAGGGAGGC. All primers were extended at the 5' end with universal primer sequences; sense: CTCCTGTTCCGACCCTGCC and antisense: CGGAACAGGAGAGCGCACG. PCR products were purified using the QIAquick Gel Extraction Kit (Qiagen, West Sussex, UK), sequenced using the universal sequence primers and the BigDye DyeTerminator Cycle Sequencing Kit v. 1.1 and analyzed on an ABI 310 (Applied Biosystems).
Cells were first exposed to varying doses of radiation then washed in PBS, trypsinized and plated in 100 mm dishes in triplicate. Primary hMSCs were plated at about 10 cells per cm2 whereas hMSC-telo1 cells were plated at about 7 cells per cm2. After incubation for 14 days at 37 °C in 5% humidified CO2, the cells were washed twice with PBS, stained with 0.5% Crystal Violet in methanol for 5–10 min at room temperature and washed twice with PBS. Visible colonies were counted and the colony forming efficiency calculated in percent of plated cells.
CDKN2A genomic integrity analysis
Apoptosis assay (TUNEL)
All exons of CDKN2A and exon 4 of ACTB (β-actin) were amplified for 38 cycles (95 °C for 30 s, 58 °C (CDKN2A Exon1A:
Cells were seeded in 75 cm2 culture flasks and irradiated with different doses of γ rays. At different time points (4, 24 and 96 h after), cells were trypsinized, fixed in 4% paraformaldehyde
β-Galactosidase staining 2500 cells were seeded in slide flasks and irradiated 2 days later with different doses of γ rays. At different time points after irradiation (4, 7 and 10 days after), the cells were washed in PBS and fixed for 5 min at room temperature in 2% formaldehyde/0.2% glutaraldehyde. After washing, the cells were incubated at 37 °C (no CO2) over night with fresh β-gal solution containing 1 mg of 5-bromo-4-chloro-3-indolyl β-Dgalactoside (X-Gal, Sigma) per ml, 40 mM citric acid (pH 6.0), 5 mM potassium ferrocyanide, 5 mM potassium ferricyanide, 150 mM NaCl, 2 mM MgCl2. From each slide flask, minimum of 500 cells were counted under the microscope. The experiment was repeated 3 times and the mean percent of β-galactosidase-positive cells was then calculated.
CFU assay
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solution for 60 min on a shaker at room temperature and permeabilised in 0.1% Triton X-100/0.1% sodium citrate for 5 min at room temperature. Cells were stained using the In Situ Cell Death Detection Kit, Fluorescein (Roche) according to manufacturer's instructions. Green (FITC) fluorescence after 488 nm laser excitation was measured on the flow cytometer (BS FACSAria™ Cell Sorter, BD Biosciences, Erembodegem, Belgium).
DNA damage and repair assay (Fluorescence-Detected Alkaline DNA unwinding) DNA damage was measured as previously described [16]. The method detects single and double-strand breaks by their effect on the rate of DNA denaturation in alkali, monitored by the fluorescence intensity of an intercalating dye. The fluorescence intensity is inversely correlated to the number of DNA strand breaks present at the time of lysis. DNA damage percentage was calculated as 100 × (P0 − Px) / P0, with P0: fluorescence intensity of an unirradiated sample and Px: fluorescence intensity of the irradiated cell sample. Repair capacity was assayed by leaving the cells for the indicated time points (10′ till 4h), lysing them thereafter and compare them to the values immediately after irradiation.
H2AX Western analysis Cells were lysed in SDS lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris–HCl, pH 8.1) 1 h, 4 h and 1 day after irradiation and subjected to 2 rounds of freeze thawing on ethanol/dry ice. Following cell lysis, 40 μg protein was electrophoresed on NuPAGE 4–12% Bis-Tris polyacrylamide gels (Invitrogen) and transferred to nitrocellulose membranes. Membranes were incubated with mouse Anti-phospho-Histone H2A.X antibody (Upstate Biotechnology, Waltham, MA, USA) followed by incubation with goat anti-mouse HRP-conjugated secondary antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Immunoreactive bands were detected using the ECL detection system (Amersham Bioscience/GE Healthcare Bio-Sciences, Little Chalfont, UK). To assess protein loading, stripped membranes were re-probed using rabbit anti-Erk antibody (Santa Cruz Biotechnology).
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Normalization of telomere length values and Statistical analysis Telomere length values were normalized for each metaphase using the following equation: Z ¼ X=l where Z = normalized telomere length value; X = raw telomere length value; μ = mean telomere length of all telomeres in the metaphase. The rationale and normalization procedures were done as previously described in [19,20].
Telomere restriction fragment assay (TRF) Telomere lengths were determined using the Telo TAGGG Telomere Length Assay (Roche). Briefly, genomic DNA was embedded in 1% agarose plugs and a plug containing 3 μg of genomic DNA was digested with the restriction enzymes HinfI and RsaI. Plugs were analyzed in a 1% agarose gel at 14 °C by pulsed field gel electrophoresis (CHEF-DRII-SYSTEM, Biorad, Herlev, Denmark) at 3.5 V/cm for 17 h at a ramp pulse of 2–10 s. The DNA was Southern-blotted to charged nylon membranes (Hybond-N+, Amersham Bioscience) and hybridized with a digoxenin (DIG)-labelled TTAGGG probe. The average TRF lengths were determined by comparing the signals relative to a molecular weight standard.
Chromosome analysis The cells were cultured in slide flasks (Nunc, Rochester, NY, USA). Before harvest, 10 mg/ml Colcemid (GIBCO, Invitrogen) was added to a final concentration of 0.2 μg/ml and culturing continued for another 35 min. The cells were lysed by adding 60 mM KCl and thereafter standard harvesting took place including the Carnoy's fixative. Giemsa bands were produced with Wright's stain. From each cell type and dose 45 metaphases were analyzed.
Results Experimental system
Telomere profiles The telomere profile was measured on chromosomes prepared as described below. A telomere FISH kit (DakoCytomation, Glostrup, Denmark) was used for telomere staining according to manufactures instructions. For subsequent karyotyping, the DNA was stained with a DAPI solution (0.5 μg/ml) also containing antifade (0.11% phenylenediamine dihydrochloride).
Image acquisition and analysis Fluorescent-specific signals were visualized by fluorescence microscopy and captured at 100× magnification with a cooled CCD camera using IpLab software (Scanalytics, Fairfax, VA, USA). For each sample, twenty metaphases were analyzed. Image acquisition and analysis were done as previously described in [19,20].
To explore the roles of telomeres and telomerase activity in relation to radiosensitivity, we investigated the effect of ionizing radiation on normal primary human mesenchymal stem cells (hMSC) and telomerase-immortalized hMSCs (hMSCtelo1) established by transducing normal primary hMSCs with a retrovirus carrying the hTERT gene. Analysis of the endogenous and ectopic expression of hTERT showed that neither primary hMSC nor the hMSC-telo1 line expressed endogenous hTERT; however, the immortalized hMSC-telo1 cells expressed high levels of ectopic hTERT that resulted in an increased mean telomere length of 21 kb already at PDL 43 while the mean telomere length of the primary cells was 8 kb (Fig. 1A).
Radiation of hMSC and hMSC-telo1 cells Normal primary (hMSC) and immortal hMSCs (hMSC-telo1) at PDL 114 were treated with varying doses from 2.5 to 15 Gy of γ
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Fig. 1 – (A) TRF analysis of primary hMSC and telomerase-immortalized hMSC (hMSC-telo1 cells) at the indicated PD levels. TRF lengths were determined as described in Materials and methods. Molecular size markers (kb) are indicated to the left of the blot. (B, C) Standard RT-PCR analysis of hMSC and hMSC-telo1 cells. (B) The figure illustrates non-irradiated cells and cells irradiated with 2.5 and 5 Gy of γ-rays. RNA was isolated from the cells and subjected to RT-PCR using primers amplifying the ectopic hTERT gene and the β-actin gene. A sample without template was included as a negative control. (C) The figure shows cells irradiated with a 5-Gy dose (+) or non-irradiated (−). Three days after IR, some of the cells were treated with VitD3 (+). RNA was isolated from the cells and subjected to RT-PCR using primers amplifying a region in the osteocalcin gene (Ost), the alkaline phosphatase gene (AP), the collagen type I gene (Col-1) and in the β-actin gene. A sample without template was included as a negative control. PCR was performed on RNA to verify that no DNA was present in the RNA preparation (data not shown). (D) Induction of p21CIP1 (CDKN1A) expression following irradiation. RNA was isolated from the cells and subjected to real-time RT-PCR. Expression was measured relative to β-actin before (Control) and 4 h after irradiation in hMSC (light gray) and hMSC-telo1 cells (dark gray) with doses from 2.5 to 15 Gy. Data shown are mean ± SEM for three independent experiments. (E) PCR analysis on genomic DNA for CDKN2A copy number analysis. Figure shows pooled PCR products from each of the different cell lines and passages.
rays. Irradiation of normal hMSC and hMSC-telo1 cells did not lead to immediate cell death, not even at the highest dose.
either hMSC or hMSC-telo1 cells (data not shown) or modulated the expression of ectopic hTERT in hMSC-telo1 cells (Fig. 1B).
Endogenous and ectopic hTERT expression
Expression of differentiation markers
The endogenous and ectopic expression of hTERT was analyzed after irradiation of the cells by standard RT-PCR analysis. Irradiation did not induce expression of endogenous TERT in
It has previously been shown that introduction of hTERT into primary hMSCs does not change the ability of the cells to differentiate into various cell types [3,21]. After irradiation, the
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cells were incubated in medium containing 1,25-(OH)2-vitamin D3 (vitamin D3) that induces differentiation along the osteogenic pathway. The expression of different osteoblastic markers (osteocalcin (Ost), alkaline phosphatase (AP) and collagen type I (Col-1)) was analyzed by standard RT-PCR analysis. As seen in Fig. 1C, presence of vitamin D3 in the culture medium leads to increased expression of osteocalcin in both primary hMSC and hMSC-telo1 cells before irradiation. The expression levels of AP and Col-1 were at the same level with or without vitamin D3. Irradiation of the cells did not change the ability of the cells to express these markers (Fig. 1C).
Wild-type p53 but inactivation of p16Ink4A and p14ARF DNA sequence analysis of the coding regions of the p53 gene, TP53, was wild type in both hMSC and hMSC-telo1 (data not shown). Four hours after irradiation, CDKN1A (p21CIP1) expression was induced in hMSC and hMSC-telo1, indicating that the p53 protein was functioning normally in both cell lines (Fig. 1D). CDKN2A (p16Ink4A) expression could not be detected in hMSC-telo1 (data not shown) and DNA copy number analysis showed deletion of both copies of CDKN2A (Fig. 1E). Thus, p16Ink4A and p14ARF are fully inactivated in hMSC-telo1, whereas active in hMSC.
Proliferation and survival Irradiation of primary hMSC and hMSC-telo1 cells resulted in a dose-dependent growth arrest (Figs. 2A and B). At the high doses (10 and 15 Gy), many cells, both hMSC and hMSC-telo1, acquired an enlarged flat morphology typical of senescent cells after 3–5 days. Analysis of the colony forming ability (CFU assay) after irradiation demonstrated that both primary hMSCs and hMSC-telo1 cells showed a decrease in the CFUs already at 2.5 Gy and at the high dose (15 Gy) no colonies were formed within 14 days in either of the cell types (Fig. 2C) meaning that both primary hMSCs and hMSC-telo1 cells already showed considerable cell cycle arrest at the low dose 2.5 Gy. hMSCs irradiated with the higher doses (10–15 Gy) did not start to divide within 3 months following irradiation whereas few dividing cells were observed 1–2 weeks after irradiation with the 2.5-Gy dose as demonstrated by the CFU assay after 2 weeks. hMSC-telo1 cells that received the high doses (10–15 Gy) started to divide 2–3 weeks after irradiation. Thus, hMSC-telo1 cells required shorter time (days in culture) to return to cell cycle than hMSCs.
DNA-damage and phosphorylation of H2AX Irradiation-induced DNA damage/repair was analyzed by Fluorescence-detected alkaline DNA unwinding and it was found that irradiation induced the same extent of DNA damage in both cell types (Figs. 3A and B) and the amount of damage increased with the dose. After 4 h, the DNA damage percentage in hMSC and hMSC-telo1 cells receiving both the low (2.5 Gy) and the high (15 Gy) doses had decreased to a level of 20% damage indicating that the two cell types did not differ in their overall genomic repair capabilities. DNA damage was furthermore investigated by measuring the kinetics of H2AX phosphorylation after subjecting the cells
Fig. 2 – Growth curve of primary hMSC (A) and the telomerase-immortalized hMSC-telo1 cells (B) after irradiation. Cells were plated and treated with different doses of γ-rays: 2.5 Gy (■), 5 Gy (▲), 10 Gy (×) and 15 Gy (●) or non-treated cells (O). The cells were irradiated at the indicated time point and the cumulative PD levels followed. (C) CFU assay of primary hMSC (●) and hMSC-telo1 cells (■). Cells were irradiated with 2.5 or 15 Gy of γ-rays, plated in 100 mm dishes and scored for colony formation after 14 days. Data shown are mean ± SD.
to IR. For both hMSC and hMSC-telo1 cells the phosphorylation of H2AX increased in a dose-dependent way upon irradiation with doses from 2.5 to 15 Gy of γ-rays (Fig. 3C). In hMSC-telo1 cells, the level of H2AX phosphorylation was decreased
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Fig. 4 – Beta-galactosidase-associated senescence in irradiated primary hMSC (grey lines) and telomerase-immortalized hMSC-telo1 cells (black lines). Cells were irradiated with 2.5 (dotted line) or 15 Gy (broken line) of γ-rays and stained with X-gal substrate 4, 7 and 10 days after irradiation. Control cells were not irradiated (full line). The percentage of blue cells is given in the figure. The data shown are mean percent ± SD estimated from a binomial distribution for three experiments.
considerable 4 h after irradiation, whereas in hMSCs this was not the case. One day after, no phosphorylation could be detected in either of the cell types (data not shown). This finding was in accordance with the DNA damage/repair analysis performed by fluorescence activated DNA unwinding (FADU).
Senescence-associated β-galactosidase and TUNEL assays
Fig. 3 – DNA damage in primary hMSC (A) and telomerase-immortalized hMSC-telo1 cells (B) after irradiation. DNA damage and repair was measured by a fluorescence-detected alkaline DNA unwinding assay in 3 independent experiments. The cells were trypsinized and irradiated with doses from 2.5 to 15 Gy of γ-rays. At different time points, the initial and the remaining damage (remdam) in the cells were analyzed. Error bars are standard errors. (C) Western blot analysis of induction of H2AX phosphorylation in irradiated hMSC and hMSC-telo1 cells. Protein extracts were prepared from non-irradiated cells or cells irradiated with doses from 2.5 to 15 Gy of γ-rays. Size markers are indicated to the left of the blot. ERK staining of the same samples was used as loading control.
The proportion of senescent cells after irradiation was analyzed by staining the cells for senescence-associated β-galactosidase (SA β-gal). Before irradiation, 12%± 0.8% of normal hMSCs and 2% ± 0.4% of hMSC-telo1 cells showed positive staining for SA βgal (Fig. 4). Irradiation increased the proportion of SA β-gal positive cells in a dose-dependent way in both hMSC and hMSCtelo1 cells. Ten days after irradiation, a proportion of 91% ± 0.6% and 97% ± 0.4% of the hMSCs stained positive for SA β-gal at the low (2.5 Gy) and high (15 Gy) doses, respectively. However, in hMSC-telo1 cells only 30% ± 1% and 76%± 1% of the cells showed SA β-gal positive staining 10 days after irradiation with 2.5 Gy and 15 Gy of γ-rays, respectively (Fig. 4). The TUNEL assay was used to measure the proportion of apoptotic cells in the cell cultures. Before irradiation, 0.25% ± 0.14% (n = 3) and 0.10% ± 0.03% (n = 3) apoptotic cells were observed in hMSC and hMSC-telo1 cell cultures, respectively. Irradiation of hMSC and hMSC-telo1 cells with 2.5 and 15 Gy of γrays did not lead to increased apoptotic cell death when measured 4, 24 and 96 h after irradiation by TUNEL assay (data not shown).
Telomere length and profile To investigate the effect of irradiation on the telomere length, we performed a TRF analysis. Neither hMSC nor hMSC-telo1 cells irradiated with doses from 2.5 to 15 Gy exhibited detectable telomere shortening when measured up to 10 days after irradiation (Fig. 5A).
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Fig. 5 – Telomere length and profile analysis. (A) The TRF length was determined in primary hMSC (lanes 2–14) and telomerase-immortalized hMSC-telo1 cells (lanes 15–27) 1, 4 and 10 days after irradiation of the cells with doses from 2.5 to 15 Gy of γ-rays. TRF lengths were determined as described in Materials and methods. Molecular size markers (kb) are indicated to the left of the blot. (B) Comparison of the telomere profiles in two different unirradiated hMSC-telo1 samples at PDL 128. (C) Comparison of the telomere profiles in two different hMSC-telo1 samples irradiated with 15 Gy. (D, E) The telomere profiles of hMSC-telo1 cells irradiated with 2.5 Gy (D) and 15 Gy (E) of γ-rays were compared to the telomere profile of non-irradiated hMSC-telo1 cells. The mean values of normalized telomere lengths from 20 metaphases are assigned to the y-axis. Error bars describe the 95% confidence limits for the mean.
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Fig. 5 (continued ).
The effect of irradiation on the cells telomere profile was measured by using a PNA-FISH method and computer-assisted telomere quantifier software. Analysis of hMSCs after irradiation was hampered by the difficulties of obtaining metaphases due to low cell cycling. As previously shown by our group, the hMSC and hMSC-telo1 cells have a telomere profile as first described in lymphocytes (Graakjær et al, submitted work). A high and significant correlation is obtained when the telomere profiles of two hMSC-telo1 samples from the same passage (PDL 128: correlation = 0.71, p < 0.001, Fig. 5B) or from different passages (PDL 43 compared to PDL 167: correlation = 0.62, p < 0.001) are compared indicating that the telomere profiles of un-irradiated samples are stable even after long-term cell growth. However, no significant correlation is found when comparing the telomere profiles of two samples irradiated with the same dose (hMSC-telo1 irradiated with 15 Gy: correlation = −0.06, p = 0.63, Fig. 5C) indicating that the telomere length pattern is not reproducible after irradiation. Comparison of the telomere profiles before and after irradiation show that the telomere profile is lost in hMSC-telo1 cells at both low (2.5 Gy) and high doses (15 Gy) (Figs. 5D and E). The difference in profile changes between 2.5 and 15 Gy can be explained by the fact that irradiation destroys the telomere length pattern in a random manner and the cells that establish the cell culture after irradiation have different telomere profiles.
Chromosome analyses After irradiation, random chromosomal aberrations have been observed but no clonal aberration except in 15 Gy
irradiated hMSC-telo1 cells. hMSCs could only be analyzed at the low dose 2.5 Gy because cells receiving higher doses did not proliferate after irradiation. After irradiation with a 2.5Gy dose, hMSCs showed a 3-to 4-fold increase in anaphase bridge/micronuclei formation, while hMSC-telo1 cells only showed a 0- to 1-fold increase. In addition, it was observed that the frequency of lost telomeric signals after irradiation with a 2.5-Gy dose had increased 2- to 3-fold in hMSCs but only 0- to 1-fold in hMSC-telo1 cells. Overall, these results indicate that cells with long telomeres due to high telomerase expression show higher stability at telomeric regions than cells with shorter telomeres and no telomerase expression.
Discussion In this study, we have investigated the radiosensitivity of telomerase-immortalized hMSCs versus normal primary hMSCs. The hMSC-telo1 cells have high telomerase activity and long telomeres compared to their normal counterpart hMSCs that have shorter telomeres and no telomerase activity. Irradiation of both cell types resulted in a dosedependent growth arrest, indicating that both cell types have at least one intact DNA damage control pathway. This assumption is further supported by the finding that both cell types had functional p53, although telomerase-transduced cells as expected contained inactivating mutations in the CDKN2A gene, resulting in lack of p16 and p14 protein. It seems likely that compromising the p16-checkpoint increased the
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ability to extend cellular life span while maintaining the diploid status of telomerase-immortalized cells. hMSC-telo1 cells returned to cell cycle within a few weeks after irradiation even after subjected to the high dose (15 Gy). In contrast, primary hMSCs receiving the same high dose did not return to cell cycle within 3 months of radiation. The faster recovery after irradiation of telomerase-immortalized cells might be explained by the fact that these cells do not express p16 since this may result in these cells returning to cell cycle in spite of remaining DNA damage. A number of other observations point, however, towards telomeres as a central player in the induction and subsequent release from cell cycle arrest. It is well established that radiation using γ rays induces singlestrand and double-strand breaks (DSB) in the DNA and that the cell cycle machinery arrests the cells until the damage has been repaired [9–11]. Our data show that the level of overall DNA damage/repair did not differ between hMSC and hMSCtelo1 cells after irradiation with the same doses and the amount of damage remaining after 4 h was the same in the two cell types indicating that the cells had similar repair capabilities. This result was confirmed by analysis of phosphorylation of histone H2AX. Upon irradiation, H2AX imme-
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diately becomes phosphorylated on Ser-139 and is believed to recruit DNA repair factors to the site of DNA double-strand breaks [22]. Hence, the return of H2AX to its non-phosphorylated state within a day also suggests that overall DNA repair in both cell systems are finalized within a day. Thus, DNA repair is not dependent on telomere length and/or telomerase expression in our cell system. Likewise, it has been shown that double-strand break repair induced by γ-radiation is identical in normal and telomerase-immortalized human foreskin fibroblasts [23] and furthermore that repair of pyrimidine dimers and dimethylsulfate damage induced by ultraviolet radiation or dimethylsulfate is identical in telomerase-immortalized human skin fibroblasts and normal skin fibroblasts [24]. In spite that overall DNA repair was finalized within a day even with the high doses of γ-rays both cell types remained in cell cycle arrest for weeks to months. Since all the hallmarks of stress-induced senescence are present in the cells, this finding very strongly suggests that part of the DNA was not repaired within these first hours and that this remaining damage maintains the cells in cell cycle arrest. We have in this connection obtained results indicating that radiation leads to
Fig. 6 – Illustration of our current hypothesis on p53 and p16 pathways in our cell system.
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extensive damage of the telomeres. Since in general telomeric DNA is less efficiently repaired than other parts of the genome [25], it can therefore be imagined that the remaining DNA damage is associated with the telomeres. The loss of telomere profile indicates telomere damage, namely suggesting primary occurrence of double-strand breaks in the telomeric regions. At first glance it may seem puzzling that the relatively few damaged/dysfunctional telomeres per cell can cause such an extensive loss of profile. We hypothesize that the extensive profile loss is a consequence of not only the initial telomere damage, but also of subsequent fusion–break–fusion cycles, occurring after the cells have re-entered the cell cycle, resulting in substantial re-shuffling of telomeric regions, until finally all chromosome ends have achieved stable telomere coverage, at which point fully normal cell proliferation is established, but with a different profile. It should be pointed out, as also previously stated [26], that full development of this cascade of events require that cells have not totally left the cell cycle, since fusion–break–fusion cycles require at least some degree of cell division. This condition is, however, fulfilled for the cells where we analyzed telomere profiles, simply because these cells go into mitosis, which means that they are cycling. Surprisingly, the overall telomere length did not decrease after irradiation. This might, however, be explained by the appearance of fusion breakage fusion cycles that would lead to rearrangements, but not necessarily result in telomeric loss. It may thus be possible that irradiation causes destabilization of the telomeric loop and uncapping of telomeres without initial double-strand breaks. This then leads to fusion breakage fusion cycles without net loss of telomere sequences. In agreement with this, it has previously been shown that irradiation can induce senescence without reducing telomere length [13–15] also suggesting that one or more aspects of telomeres other than length per se contribute to the response. Furthermore, Ku86−/− mouse embryonic fibroblasts with long but dysfunctional telomeres resulting from loss of their endcapping function showed a senescence-like arrest [27]. Our results thus support the hypothesis that telomere dysfunction can induce stress-induced premature senescence by a telomere-length-independent mechanism most likely involving loss of the telomere capping function. It has previously been shown that dysfunctional, uncapped telomeres associate with γ-H2AX [28]. In our study, when the irradiated cells were stained with antibodies against H2AX and TRF1 we also observed that there was a tendency of colocalization at the telomere sides. Furthermore, a small proportion of the H2AX foci were remaining after a day. The appearance of anaphase bridges can be used as a measure of telomere dysfunction [29]. Based on the fact that the hMSC-telo1 cells have fewer cells positive for SA β-gal stain and a lower frequency of anaphase bridges and lost telomeric signals, it can be suggested that cells with long telomeres and high telomerase activity re-establish telomeric function more efficiently than cells with shorter telomeres and no telomerase expression and that this ability is important for returning to cell cycle. This is in agreement with previous reports, where it has been shown that telomerase has a protective effect on very short telomeres that, in its absence, would have caused cells to stop dividing. Thus, active
telomerase helps even very short telomeres to be functionally capped [8,30,31]. Furthermore, it has been indicated that longer telomeres are more likely to rebuild the cap than are shorter telomeres [32,33]. It has recently been shown that fibroblasts expressing hTERT showed high level of stability even after exposure to ionizing radiation [34]. In agreement with this, our cell systems showed low sensitivity to radiation. Clearly, hMSCtelo1 cells showed a higher survival effect than hMSCs, which might be explained by a telomerase survival effect. Overall, our data suggest that uncapping of telomeres induced by irradiation plays a role in cell cycle depression. Furthermore, our results are compatible with the hypothesis that the signaling pathway goes through p53 and not p16. Fig. 6 illustrates our current hypothesis with regards to signaling pathways in our cell system. It is here illustrated, that we, like others [35–38] believe that p16 and p53 are located in two different pathways, and that telomere damage is signaled through the pathway involving p53. Indirect support for this latter notion comes from the fact that telomerase-immortalized cells often have inactivation of p16 suggesting that telomere elongation and loss of p16 are two independent events in the immortalization process [36–38]. In addition, we find that irradiation does not change the differentiation ability of mesenchymal stem cells. Overall, our observations point out some of the key properties of these cell populations that allow them to survive under high doses of γ-rays.
Acknowledgments The authors wish to thank Anette Thomsen, Christian Knudsen, Bente Kierkegaard, Inger Marie Thuesen, Alan Leake and Sharon Burns for their excellent technical assistance. The work was supported by Danish Cancer Society, Cancer Research UK, Glasgow University and European Union contracts EC-FP5-FI6R-CT-2002-0021 (TELOSENS) and LSHCCT-2004-502943 (MOL Cancer Med).
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