TRF2 dysfunction elicits DNA damage ... - Wiley Online Library

Journal of Neurochemistry, 2006, 97, 567–581

doi:10.1111/j.1471-4159.2006.03779.x

TRF2 dysfunction elicits DNA damage responses associated with senescence in proliferating neural cells and differentiation of neurons Peisu Zhang,* Katsutoshi Furukawa,* Patricia L. Opresko, Xiangru Xu,* Vilhelm A. Bohr and Mark P. Mattson*,à *Laboratory of Neurosciences and Laboratory of Molecular Gerontology, National institute on Aging Intramural Research Program, 5600 Nathan Shock Drive, Baltimore, Maryland, USA àDepartment of Neuroscience, Johns Hopkins University School of Medicine, Baltimore, Maryland, USA

Abstract Telomeres are specialized structures at the ends of chromosomes that consist of tandem repeats of the DNA sequence TTAGGG and several proteins that protect the DNA and regulate the plasticity of the telomeres. The telomere-associated protein TRF2 (telomeric repeat binding factor 2) is critical for the control of telomere structure and function; TRF2 dysfunction results in the exposure of the telomere ends and activation of ATM (ataxia telangiectasin mutated)-mediated DNA damage response. Recent findings suggest that telomere attrition can cause senescence or apoptosis of mitotic cells, but the function of telomeres in differentiated neurons is unknown. Here, we examined the impact of telomere dysfunction via TRF2 inhibition in neurons (primary embryonic hippocampal neurons) and mitotic neural cells (astrocytes and neuroblastoma cells). We demonstrate that telomere dysfunction induced by adenovirus-mediated expression of dominant-negative TRF2 (DN-TRF2) triggers a DNA damage

response involving the formation of nuclear foci containing phosphorylated histone H2AX and activated ATM in each cell type. In mitotic neural cells DN-TRF2 induced activation of both p53 and p21 and senescence (as indicated by an up-regulation of b-galactosidase). In contrast, in neurons DN-TRF2 increased p21, but neither p53 nor b-galactosidase was induced. In addition, TRF2 inhibition enhanced the morphological, molecular and biophysical differentiation of hippocampal neurons. These findings demonstrate divergent molecular and physiological responses to telomere dysfunction in mitotic neural cells and neurons, indicate a role for TRF2 in regulating neuronal differentiation, and suggest a potential therapeutic application of inhibition of TRF2 function in the treatment of neural tumors. Keywords: astrocytes, ataxia telangiectasia mutated, hippcampal neurons, senescence, telomeric DNA damage, tumor cells. J. Neurochem. (2006) 97, 567–581.

The ends of chromosomes in mammalian cells are stabilized by telomeres, which consist of a repeating DNA sequence (TTAGGG) that forms a duplex loop structure with telomere-associated proteins. This nucleoprotein complex forms a ‘cap’ that prevents the chromosome ends from being recognized as damaged DNA, and also prevents end-to-end fusion of chromosomes (Chan and Blackburn 2004). Two proteins called telomeric repeat binding factors, TRF1 and TRF2, form homodimers that directly bind to the telomere duplex region and facilitate formation of the telomere t-loop (Broccoli et al. 1997; Griffith et al. 1999; Stansel et al. 2001). Telomere dysfunction can occur as the result of telomere shortening in late generation somatic cells (Karlseder et al. 2002; Herbig

et al. 2004), mutated telomeric DNA (Guiducci et al. 2001; Lin et al. 2004) and through inhibition of TRF2 function by overexpression of a dominant negative form of

Resubmitted manuscript received November 30, 2005; accepted January 8, 2006. Address correspondence and reprint requests to Mark P. Mattson, Laboratory of Neurosciences, National institute on Aging Intramural Research Program, 5600 Nathan Shock Drive, Baltimore, MD 21224, USA. E-mail: [email protected] Abbreviations used: ATM, ataxia telangiectasia mutated; DN-TRF2, dominant negative telomeric repeat binding factor 2; MAP1b, microtubule-associated protein 1b; MAP2, microtubule-associated protein 2; NBS1, Nijmegen breakpoint syndrome 1; NF-M, neurofilament medium; TRF, telomeric repeat binding factor.

2006 The Authors Journal Compilation 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 567–581

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TRF2 (DN-TRF2) which lacks the NH2-terminal basic domain and the COOH-terminal Myb domain (Karlseder et al. 1999). Overexpression of DN-TRF2 achieves rapid and extensive telomere decapping by binding with endogenous TRF2 (Broccoli et al. 1997; Van Steensel et al. 1998; d’Adda di Fagagna et al. 2003; Takai et al. 2003), thereby displacing it from the telomere. This restricted telomere damage can activate ATM-dependent signaling pathways in association with nuclear foci of phosphorylated DNA damage response proteins including cH2AX, 53BP1, MDC1 and NBS1 (d’Adda di Fagagna et al. 2003; Takai et al. 2003). Thus, in mitotic cells TRF2 is a pivotal protein in telomere maintenance and recruitment of nucleases and other DNA repair factors to the telomere. Mutations in the telomere-associated proteins ATM (ataxia telangiectasia mutated), Werner and NBS1 (Nijmegen breakpoint syndrome 1) are responsible for human disorders characterized by developmental abnormalities and increased cancers (Digweed et al. 1999; Opresko et al. 2003; McKinnon 2004). Each of these proteins has been shown to be involved in DNA repair processes and their mutations therefore result in the accumulation of damaged DNA in mitotic cells. TRF2 has been shown to bind to ATM, NBS1 and Werner proteins (Karlseder et al. 2004; Zhu et al. 2000; Opresko et al. 2004), suggesting possible functional relationships between these proteins. In addition to abnormalities in proliferative tissues, patients with ataxia telangiectasia, NBS and Werner syndrome also manifest prominent neurological symptoms and analyses of the patients suggest abnormalities in development of neuronal circuits (De Leon et al. 1976; De Stefano et al. 2003; Lammens et al. 2003). During development of the nervous system, neural precursor cells proliferate and then differentiate into either neurons or mitotic glial cells. Recent studies of mice deficient in telomerase have provided evidence that telomere erosion adversely affects the survival of neural progenitor cells, causing a depletion of this cell population (Wong et al. 2003; Ferron et al. 2004). Studies of cultured embryonic hippocampal neurons suggest that telomerase can promote the survival of neural tumor cells and neurons during brain development (Fu et al. 1999; Du et al. 2004). Therefore, increasing evidence suggests important roles for telomeres in the processes of neurogenesis and survival of neural progenitor cells and newly generated neurons. In the present study we examined telomeredirected DNA damage responses in neural tumor cells, astrocytes and neurons. We demonstrate that TRF2 inhibition is sufficient to activate the ATM-mediated DNA damage response pathway in all three cell types, but that the molecular and cellular responses to the telomere dysfunction are different in neurons and mitotic neural cells.

Experimental procedures Cell cultures SH-SY5Y human neuroblastoma cells were obtained from American type culture collection (CRL-2266; Vienna, VA, USA). Cell cultures were maintained at 37C in a humidified 5% CO2/95% air atmosphere. The culture medium consisted of Advanced-Dulbecco’s modified Eagle’s medium (Gibco Life Technologies, Gaithersburg, MD, USA), 5% fetal CloneIII bovine serum (HyClone, Logan, UT, USA), 2 mM L-glutamine, 100 U/mL penicillin G and 100 lg/mL streptomycin. Primary neuronal cultures were prepared from embryonic day 18 rat hippocampus as described previously (Mattson et al. 1995). Neurons were plated at a density of 70–100 neurons/mm2 of culture surface in 60-mm or 35-mm culture dishes that had been coated with polyethyleneimine. The culture maintenance medium consisted of Neurobasal medium with B27 supplements and 0.01% gentamycin sulfate. Primary cortical astrocytes were cultured from postnatal day 2 Sprague-Dawley rats as described previously (Blanc et al. 1998) and were subcultured no more then four times prior to experiments Gene cloning and cell transfection The rat TRF2 gene was cloned using primers derived from putative rTRF2 (GenBank accession number XM341683): 5¢-TCAGACAGATCCGGGACATC-3¢ and 5¢-TTTCATGGTCCGCCAGCGATC3¢. RT–PCR was performed using total RNA and cDNA from cultured embryonic rat cortical neurons using methods described previously (Zhang et al. 2003). Full length human TRF2 cDNA was subcloned into pcDNA3.1 plasmid. Dominant negative TRF2 (DN-TRF2) cDNA (Karlseder et al. 1999; Zhang et al. 2003) was generated by PCR using full length human TRF2 cDNA as template with forward primer 5¢-CACCAGATCTACCCTGGAGGCACGGCTGGAAG AGGCAGTCAAT-3¢ and reverse primer 5¢-GAATTCTTACTTCTGCTTTTTTGTTATATTGGTTGT-3¢, and was then ligated with pENTER directional TOPO cloning vector (Invitrogen, Carlsbad, CA, USA). The DN-TRF2 cDNA was then subcloned either into pIRES-eGFP vector (Clontech Mountain View, CA, USA) with restriction enzymes BglII and EcoRI, or into two adenoviral Gateway Destiny vectors (pAD/CMV-V5-DEST or pAD/PL-DEST) using the manufacturer’s protocols modified for co-expression of eGFP reporter genes under CMV promoter. Lipofectamine 2000 (Invitrogen) was used for the transfection of SH-SY5Y cells, astrocytes and primary hippocampal neurons. Stable transfection of SH-SY5Y cells was performed in the presence of 800 lg/mL G418 selection for 2 days and 400 lg/mL G418 thereafter. For electrophysiology studies, primary hippocampal neurons (culture day 3) were transfected with either GFP only vector or DN-TRF2 together with IRES-GFP using a DNA/calciumphosphate co-precipitation method described previously (Kohrmann et al. 1999). For transduction of astrocytes and neurons, adenoviral vectors containing the genes for beta-galactosidase, eGFP and DN-TRF2 (alone or in combinaton with an IRES-eGFP reporter gene) were packaged into 293A cells using the ViraPower Adenoviral expression system using the manufacturer’s protocol (Invitrogen). Adenoviral particles were purified by ultracentrifugation in a gradient of cesium chloride (1.50, 1.35 and 1.25 mg/mL) for 1 h at 15 000 g first, followed by centrifugation for 18 h at 15 000 g in a solution of 1.35 mg/mL cesium chloride. The band


TRF2 function and telomere damage in neural cells 569

containing mature viral particles was removed and desalted against phosphate-buffered saline in a Vivaspin 20 mL column using the manufacturer’s protocol (Vivascience AG, Hanover, Germany). Infection units used for transfections were10 pfu/cell for SH-SY5Y cells, and 50–200 pfu/cell for rat hippocampal neurons and astrocytes (Filippova et al. 2001). Real time reverse transcription–polymerase chain reaction Total RNA was purified using an Absolutely RNA miniprep kit (Stratagene, La Jolla, CA, USA). First strand cDNA was synthesized with 1 lg of RNA using an iScript cDNA synthesis kit (Bio-Rad, Hercules, CA, USA). Real time PCR was performed in a 20 lL reaction with 25 ng of first strand cDNA, 0.5 lM of each primer and iQSYBR green supermix (Bio-Rad) in a DNA Engine Opticon real time PCR system (Bio-Rad Laboratories, Waltham, MA, USA). All primers were designed based on the Rattus norvegicus gene and generation of 160–200 bp PCR products. The primer sequences were: rTRF2 (accession XM341683) 5¢GCACACAGAGCCAGTGGAGAA3¢ and 5¢TGGGGATGCTAGGTTAGGAAG3¢; NF-M (neurofilament medium; accession NM017029) 5¢CCAGCTGCAGTCCAAGAGCA-3¢ and 5¢TCCCCGAAGCTCATTTTCCA3¢; MAP1b (microtubule-associated protein 1b; accession X60370) 5¢GAAGGCTCAGTGGGGAAGCA3¢ and 5¢GCATGCAGGGAAGGACTCGT3¢; MAP2 (microtubule-associated protein 2; accession X53455) 5¢ACGGGATCAACGGAGAGCTG3¢ and AGCAGAGCTGCAGGCTGAT; nestin (accession NM012987) 5¢CCTCCAGGAGCGCAGAGAAG3¢ and 5¢AGTTGCTGCCCACCTTCCAG3¢; b-actin (accession BC063166) 5¢-CCATCATGAAGTGTGACGTTG-3¢ and 5¢-GGAGGAGCAATGATCTTGATC-3¢. The thermal cycle conditions were 95C for 10 min, 40 cycles at 94C for 30 s, 60C for 30 s and 72C for 45 s. For each primer set, a standard curve was generated with five-fold serial dilution of four to five dilutions. All PCR reactions were performed with amplification efficiency greater than 97%. Data were collected using DNA Engine Opticon system (MJ Research). Data were analyzed by the relative comparative Ct method (DDCt) using gene expression macro software. Samples for each treatment were collected from four separate cultures. Each sample was tested in triplicate in each PCR run. Analysis of cell proliferation and senescence Bromodeoxyuridine (BrdU) incorporation was quantified as described previously (Cheng et al. 2003) using a biotinylated antimouse IgG secondary antibody and a Vectastain Elite ABC kit (Vector Laboratories, Burlingame, CA, USA). Cell growth curves were plotted by counting cell numbers in eight fixed areas (1 mm2) in each culture daily for 7 days. Senescence was assessed by staining for senescence-associated protein b-galactosidase as described previously (Dimri et al. 1995). Immunocytochemistry and confocal microscopy Cells were washed once with phosphate-buffered saline and fixed in 4% paraformaldehyde in phosphate-buffered saline for 30 min at room temperature. After pre-incubation with blocking buffer (2% non-fat powdered milk, 2% normal serum and 0.2% Triton X-100 in phosphate-buffered saline) for 1 h, cells were incubated with a primary antibody in blocking buffer overnight at 4C. The primary antibodies used were: anti-TRF2 monoclonal antibody (Imgenex,

San Diego, CA, USA); anti-cH2AX monoclonal and polyclonal antibodies (Upstate Biotechnology, Lake Placid, NY, USA); ATM 1918 monoclonal antibody (Upstate Biotechnology) and ATM S/TQ substrate polyclonal antibody (Cell Signaling Technology, Beverly, MA, USA). After thorough washing, cells were incubated with Alexa 568 or Alexa 633 conjugated secondary antibodies appropriate for the specific primary antibodies. In most experiments, cells were further incubated with DAPI in phosphate-buffered saline or propidium iodide in phosphate-buffered saline containing 1% RNase and 0.2% Triton X-100 for 10 min, and were then mounted in PermaFluor aqueous mounting medium (Immunon, Pittsburg, PA, USA). The cells were examined under a Zeiss LSM510 confocal laser-scanning microscope with 63 · oil immersion or 40 · water immersion objective. Immunoblot analysis Proteins (50 lg) in cell homogenates were separated by electrophoresis in 4–20% or 4–12% pre-cast gradient sodium dodecyl sulfate–polyacrylamide gel electrophoresis gels (Cambrex, East Rutherford, NJ, USA). Proteins were transferred electrophoretically to a polyvinylidene difluoride membrane sheet (Bio-Rad), which then was incubated in blocking solution [5% milk powder in Tris Tween Buffered Saline (TTBS)] for 1 h and incubated overnight at 4C in the presence of antibodies against TRF2 (Imgenex), p21 (Santa Cruz Biotechnology, Santa Cruz, CA, USA), p53 (Santa Cruz Biotechnology), phosphorylated p53 (Cell Signaling Technology), phosphorylated Chk2 T68 and Chk2 (Cell Signaling Technology). After washes in TTBS, the membrane was incubated for 1 h in the presence of the species-appropriate peroxidase-conjugated secondary antibody (Jackson Immunoresearch, West Grove, PA, USA) and washed in TTBS. Immunolabeled proteins were visualized using an enhanced chemiluminescence kit (Amersham). Subsequently, the membrane was washed with TTBS, incubated for 30 min in strip buffer (100 nM glycine, 0.1% sodium dodecyl sulfate, pH 2.0) and washed with TTBS. The stripped membrane was then reprobed with an antibody recognizing b-actin or tubulin (Sigma). Electrophysiology Electrophysiological recordings were performed on primary embryonic hippocampal neurons 3 days after transfection using the nystatin perforated patch technique as described previously (Furukawa et al. 1996, 2003; Cai et al. 2004). The resistance between the electrode filled with internal solution and the reference electrode was 4–10 MW. All external and internal solutions were adjusted to pH 7.4 and 7.2 with Tris base. The current and voltage were measured with a patch clamp amplifier Axopatch 200B (Axon Instruments, Foster City, CA, USA) linked through a Digidata 1320A interface and analyzed using pCLAMP software (Axon Instruments). Series resistance was compensated more than 70%, and linear leak and capacitance currents were subtracted using scaled currents for a 5 mV hyperpolarization from a holding potential in voltage-activated channel recordings. Cell membrane capacitance was calculated by integrating the charge under the current transient resulting from the 10 mV step. For comparison of current densities, voltage steps which elicited maximum current amplitudes were used for Na+, Ca2+ and K+ currents. All experiments were performed at 22–24C.


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Results

Dominant-negative TRF2 displaces TRF2 from telomeres in neural cells Previous studies have shown that acute telomere dysfunction (telomere ‘decapping’ and activation of a DNA damage response) can be induced in cultured tumor cells and fibroblasts (Takai et al. 2003; d’Adda di Fagagna et al. 2003). We therefore employed this molecular approach to determine the effects of telomere dysfunction on neural cells. A DN-TRF2 cDNA construct was made as described previously (Karlseder et al. 1999) and was expressed in the human neuroblastoma cell line SH-SY5Y (Figs 1b and c). To investigate the presence and subcellular localization of endogenous and vector-expressed TRF2 we employed a TRF2 antibody previously shown to specifically recognize human and mouse TRF2 (Opresko et al. 2002; Fotiadou et al. 2004). Immunoblot analysis using this TRF2 antibody demonstrated immunoreactive bands corresponding to the molecular weights of wild-type TRF2 (66 kDa) and DN-TRF2 (50 kDa) (Fig. 1b). Endogenous TRF2 protein was also present in vector transfected SH-SY5Y cells (Fig. 1b). Human DN-TRF2 can disrupt telomere function in both human and mouse non-neural cells (Smogorzewska and de Lange 2002). However, whether rat TRF2 can be inhibited by overexpression of DN-TRF2 human cDNA is not known. We therefore cloned a portion of rat TRF2 that included its dimerization domain and compared its predicted amino acid sequence with the sequences of human and mouse TRF2. Overall, TRF2s are conserved in human, mouse and rat with more than 88% similarity (rat : human ¼ 88%; rat : mouse ¼ 94%), and the sequences within the TRF2 dimerization domain are highly homologous between rat and human with more than 96% similarity (Fig. 1a), suggesting that endogenous rat TRF2 would likely interact with human DN-TRF2. We next determined the effect of overexpression of DN-TRF2 on the subcellular localization of wild-type TRF2 (Fig. 1c). Because DN-TRF2 inhibits binding of wild-type TRF2 to telomeres without affecting TRF1 binding, YFP-TRF1 served as a telomere marker to evaluate the association of TRF2 with telomeres (Opresko et al. 2002, 2004). The co-localization of YFP-TRF1 with either wildtype TRF2 or DN-TRF2 was examined in SH-SY5Y cells transfected with the following combinations of expression plasmids: YFP-TRF1 and wild-type TRF2; YFP-TRF1 and DN-TRF2; or YFP-TRF1 and wild-type TRF2/DN-TRF2. Nuclear foci of TRF2 immunoreactivity co-localized with YPF-TRF1 in cells co-expressing YFP-TRF1 and wild-type TRF2 (Fig. 1c). In contrast, TRF2 immunoreactivity exhibited a diffuse nuclear staining pattern and did not co-localize with TRF1 in cells co-expressing DN-TRF2 and YFP-TRF1 (Fig. 1c). Furthermore, when we co-expressed YFP-TRF1 and wild-type TRF2, DN-TRF2 expression displaced TRF2

from telomeres, as demonstrated by the lack of TRF2 co-localization with YFP-TRF1 (Fig. 1c). These results indicate that, in addition to lacking the ability to bind to the telomere, DN-TRF2 inhibits wild-type TRF2 telomere association in neural cells. To establish the relative levels of expression of TRF2 in rat brain and other tissues, we measured TRF2 mRNA levels by RT–PCR analysis in samples from different brain regions and peripheral tissues. The results showed that rat TRF2 mRNA is widely expressed (Fig. 1d), a result consistent with previously published data (Klapper et al. 2001). We further examined the cellular expression of TRF2 in rat brain cells in vivo and in cell culture. Immunoblot analysis showed that TRF2 was present in each region of the adult rat brain examined (cerebral cortex, hippocampus, midbrain and cerebellum) and in neuron-enriched embryonic rat brain primary cortical (Fig. 1e) and hippocampal (data not shown) cultures. TRF2 immunoreactivity was present in the nuclei of cultured rat hippocampal neurons, where it is located in discrete foci consistent with a telomeric localization (Fig. 1f). Displacement of TRF2 from telomeres results in ATMmediated H2AX phosphorylation in neural tumor cells, astrocytes and neurons Phosphorylation of the histone H2AX at serine 139 (cH2AX) occurs in response to DNA damage, and H2AX plays important roles in cell cycle checkpoint and DNA repair processes (Fernandez-Capetillo et al. 2004). Telomere dysfunction results in the accumulation of cH2AX at telomeres in human fibroblasts (d’Adda di Fagagna et al. 2003; Takai et al. 2003). We therefore determined if DN-TRF2 affected H2AX phosphorylation in astrocytes (a mitotic type of glial cell) and primary hippocampal neurons. Astrocytes and neurons were infected with 50 pfu/cell of adenoviral expression vectors containing either eGFP or DN-TRF2 in conjunction with IRES-eGFP. In uninfected cells (data not shown) or eGFP-expressing neurons and astrocytes (Figs 2a and b), only very weak diffuse nuclear cH2AX immunoreactivity was observed. DN-TRF2 expression triggered the appearance of multiple cH2AX foci in the nuclei of both astrocytes and neurons. The number of cells with at least three cH2AX-positive foci and the number of foci per cell was greater in the astrocytes than in the hippocampal neurons (Fig. 2c). In each cell type, cH2AX positive foci were detected within 3 days of infection with DN-TRF2 adenovirus and their numbers increased greatly through 8 days post-infection (Fig. 2c). Similar results were obtained in SH-SY5Y neuroblastoma cells using liposome-mediated transfection (data not shown). To further evaluate the effects of TRF2 inhibition on the DNA damage response in mitotic neural cells and neurons, we performed immunoblot analyses of TRF2 and phosphorylated H2AX levels in cell lysates from control and



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Fig. 1 Expression of dominant negative telomeric repeat binding factor 2 (DN-TRF2) displaces telomeric repeat binding factor 2 (TRF2) from telomeres in neural cells. (a) Structure of full-length and DN-TRF2 constructs. The cDNA sequence of the cloned rat TRF2 (rTRF2) in the dimerization domain region was confirmed by sequencing and is highly homologous with human TRF2 (hTRF2). (b) A representative immunoblot illustrating the overexpression of wildtype TRF2 (WT-TRF2) and DN-TRF2 in SH-SY5Y cells 24 h after transfection. The blot was probed with a TRF-2 antibody and shows relative levels of WT-TRF2 (66 kDa, single arrow) and DN-TRF2 (50 kDa, double arrow). (c) DN-TRF2 displaces WT-TRF2 from telomeres. SH-SY5Y human neuroblastoma cells were transfected with YFP-TRF1 to label telomeres, in combination with WT-TRF2 or DNTRF2. Cells were then immunostained 24 h after transfection with a TRF2 antibody. Confocal images show YFP-TRF1 (green) and TRF2 (red) associated fluorescence; these images are representative of

20–25 cells analyzed per group. YFP-TRF1 and TRF2 were co-localized in nuclear foci in cells expressing WT-TRF2 (left). In contrast, TRF2 immunoreactivity in cells expressing DN-TRF2 was diffusely distributed in the nucleus with no co-localization of TRF2 and YFP-TRF1 foci (middle and right), suggesting that TRF2 was displaced from telomeres. Similar results were obtained in two separate experiments. (d) Relative levels of TRF2 mRNA in samples of the indicated tissues from adult rats. Values are expressed as fold difference from the TRF2 mRNA level in stomach tissue and represent the mean and SEM of determinations made in samples from four rats. (e) Immunoblot analysis showing that TRF2 is present in cultured embryonic cortical neurons and in different brain regions of the adult rat brain. (f) TRF2 immunoreactivity (green) and propidium iodide fluorescence (red) in nuclei of cultured embryonic rat hippocampal neurons (5 days in culture). Note that TRF2 immunoreactivity is concentrated in discrete foci within the nucleus.

DN-TRF2 expressing SH-SY5Y neuroblastoma cells and primary hippocampal neurons. With similar levels of DN-TRF2 expression, phosphorylated H2AX was observed

in both SH-SY5Y cells and neurons (Fig. 2d). However, the cH2AX protein level was increased more slowly and to a lesser magnitude in neurons compared to neuroblastoma cells


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Fig. 2 Evidence that inhibition of telomeric repeat binding factor 2 (TRF2) function induces a DNA damage response in hippocampal neurons and proliferating neural cells. (a and b) cH2AX foci formation, an early DNA damage response indicator, was increased after dominant negative TRF2 (DN-TRF2) transduction. Hippocampal neurons (a) and astrocytes (b) were infected with adenoviral GFP alone or adenoviral DN-TRF2 in combination with IRES-GFP (green) with the same dose of virus (50 pfu/cell) for 8 days. The cells were then immunostained with an antibody against cH2AX (red). Compared with the GFP vector control (upper), the cells expressing DN-TRF2 exhibited large numbers of cH2AX-positive foci in their nuclei (lower). Greater numbers of cH2AX-positive foci were observed in astrocytes (b, lower) than in hippocampal neurons (a, lower) when expressing DN-TRF2. (c) The cH2AX index (percentage of cells with three or

more cH2AX foci were scored as positive) was determined in cultures of astrocytes and neurons that had been expressing GFP or DN-TRF2 for 3 or 8 days. Values are the mean and SEM of determinations made in at least four cultures. *p < 0.01, **p < 0.001 compared to the corresponding value for GFP-expressing control cells. (d) Immunoblots of samples from cultured hippocampal neurons and SH-SY5Y cells that expressed either DN-TRF2 or b-galactosidase by adenoviral infection for the indicated number of days. Samples (50 lg/lane) were subjected to immunoblotting on the same membrane. Levels of endogenous TRF2 and DN-TRF2 are shown in the upper blot. With similar levels of DN-TRF2 expression in SH-SY5Y cells (5 days of Ad.DN-TRF2 infection) and hippocampal neurons (10 days of ad.DN-TRF2 infection), the level of cH2AX (middle blot) was greater in the SH-SY5Y cells compared to neurons.

(Fig. 2d). These findings suggest that inhibition of TRF2 function triggers DNA damage in both dividing neural cells and neurons, although to a greater extent in mitotic cells. ATM is an important sensor and transducer of DNA damage responses (Sancar et al. 2004). After sensing DNA double-strand breaks, ATM undergoes autophosphorylation on serine 1981, leading to dimer dissociation, enzyme activation and phosphorylation of downstream substrates including the checkpoint proteins p53 and Chk2, and the DNA repair factors H2AX and NBS1 (Andegeko et al. 2001; Burma et al. 2001). It was recently reported that TRF2 can

bind to ATM and thereby prevent its activation (Karlseder et al. 2004). We therefore examined ATM activation and ATM substrate phosphorylation induced by telomere dysfunction in mitotic neural cells and neurons. Hippocampal neurons that had been transfected with DN-TRF2 in conjunction with IRES-GFP or GFP vector were doublelabeled by immunostaining with an antibody against cH2AX and an antibody against autophosphorylated ATM1981. In vector-transfected hippocampal neurons, numerous ATM1981 immunoreactive foci were present within the nucleus, but they did not co-localize with cH2AX, which



Fig. 3 Dysfunctional telomeric repeat binding factor 2 (TRF2) induces divergent ATM-mediated DNA damage responses in hippocampal neurons and mitotic neural cells. (a and b) Telomere-directed ATM and cH2AX activations were triggered by the expression of dominant negative TRF2 (DN-TRF2) in hippocampal neurons and SH-SY5Y cells. (a) Hippocampal neurons were transfected either with eGFP vector or with DN-TRF2 in combination with the IRES-GFP reporter gene, and were then immunostained with anti-cH2AX polyclonal (green) and anti-ATM1981 (red) monoclonal antibodies. Note that in cells expressing DN-TRF2, many cH2AX foci co-localize with phosphorylated ATM1981 (single arrows in inset point to dual stained foci). (b) SH-SY5Y cells were co-transfected with YFP-TRF1 and wild-type TRF2 (WT-TRF2) or with YFP-TRF1 and DN-TRF2. The cells were then immunostained with anti-cH2AX polyclonal (green) and antiATM1981 (red) monoclonal antibodies. Note that in cells expressing

DN-TRF2, many cH2AX- and ATM1981-positive foci (double arrows in inset point to triple stained foci) are co-localized with the telomere marker TRF1 (blue). (c and d) Divergent ATM downstream substrate activation in primary neurons (c) and astrocytes (d) expressing DN-TRF2 or b-galactosidase. Both cell types were immunostained with cH2AX antibody (green) and an antibody against the ATM phosphorylated substrate (pS/TQ; red). pS/TQ foci were absent in nuclei and did not co-localize with cH2AX in hippocampal neurons expressing DN-TRF2. In contrast, numerous pS/TQ and cH2AX foci were co-localized in astrocytes infected with adenoviral DN-TRF2. The results shown in panels (a)–(d) are representative of findings obtained in at least three separate experiments (25–35 cells evaluated in each experiment). (e) Immunoblots show that levels of activated Chk2 (Chk2T68) were increased in neurons expressing DN-TRF2, but were not changed in etoposide (ET)-treated neurons.

exhibited diffuse immunoreactivity (Fig. 3a). In contrast, the nuclei of neurons expressing DN-TRF2 exhibited large foci of cH2AX immunoreactivity which often co-localized with ATM1981 immunoreactive foci. ATM activation also occurred in response to expression of DN-TRF2 in neuroblastoma cells (Fig. 3b) and astrocytes (data not shown) compared to cells overexpressing wild-type TRF2 or to vector-transfected cells. As was the case in neurons, ATM1918 immunoreactivity in neuroblastoma cells and astrocytes formed distinct foci in the nucleus, many of which were co-localized with cH2AX. YFP-TRF1 (a telomere marker) was co-transfected with either wild-type TRF2

or DN-TRF2 into SH-SY5Y cells. In cells expressing DN-TRF2, some of the cH2AX and ATM double-labeled foci co-localized with YFP-TRF1 (Fig. 3b), indicating that an ATM-mediated DNA damage response occurred in response to telomere dysfunction. In order to further evaluate ATM phosphorylation of downstream checkpoint proteins, an antibody that recognizes ATM substrates with a phosphorylated SQ/T motif was used for immunostaining in combination with the cH2AX antibody. Neurons expressing DN-TRF2 exhibited pSQ/T immunoreactivity which was present diffusely in the cytosol but not in the nucleus; no co-localization of pSQ/T with


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cH2AX nuclear foci was detected (Fig. 3c). In contrast, astrocytes infected with adenoviral DN-TRF2 displayed numerous pST/Q foci that co-localized with cH2AX foci (Fig. 3d). These results agree with the observation that p53 phosphorylation at serine 15 was only present in SH-SY5Y cells but not in neurons after transduction with adenoviral DN-TRF2 (Fig. 3d); p53 is an ATM substrate with a ST/Q phosphorylation motif. The activation of ATM in response to TRF2 inhibition was confirmed by immunoblot analysis which revealed large increases in the amount of phosphorylated Chk2 (an ATM substrate) in neurons expressing DN-TRF2 (Fig. 3e). TRF2 inhibition induces p53 expression and senescence in neuroblastoma cells and astrocytes but not in postmitotic neurons In mitotic cells, telomere dysfunction can activate DNA double-strand break repair processes that lead to the up-regulation of p53, retinoblastoma (Rb) G1 checkpoint and cell cycle arrest, and increased expression of b-galactosidase, a marker of senescence (Elmore et al. 2002; Karlseder et al. 2002; Herbig et al. 2004). To determine the consequences of TRF2 inhibition in mitotic neural cells, the effects of DN-TRF2 in proliferating human neuroblastoma cells and primary rat cortical astrocytes were evaluated. DN-TRF2 expression in neuroblastoma cells resulted in a significant reduction in their proliferation as indicated by a decrease of BrdU incorporation, limited growth rate and induction of expression of b-galactosidase (Fig. 4a). The levels of b-galactosidase were also increased in astrocytes in response to expression of DN-TRF2, although cell proliferation was not affected significantly during the 7 day post-transfection period (Fig. 4b). To determine the effects of TRF2 inhibition in post-mitotic neurons, we expressed DN-TRF2 in conjunction with IRES-eGFP as a reporter gene in primary rat hippocampal neurons in culture. Neurons expressing DN-TRF2 that was delivered either by liposome-mediated transfection (Fig. 4c) or adenovirus (data not shown) did not exhibit increased levels of b-galactosidase compared with vector-transfected control neurons. In contrast, proliferating neural progenitor cells in neurosphere cultures exhibited robust increases in b-galactosidase activity upon overexpression of DN-TRF2 (Fig. 4c). DN-TRF2 expression did not result in increased apoptosis of SH-SY5Y cells, astrocytes or neurons during 7–15-day periods after transfection (data not shown). These results suggest that impairment of TRF2 function at telomeres can induce senescence in proliferating neural cells, but not in hippocampal neurons. To evaluate the molecular mechanism responsible for senescence induced by TRF2 inhibition in mitotic neural cells, we measured levels of p53 and p21, two G1 checkpoint proteins linked to cell cycle arrest (Gartel and Tyner 2002; Vousden 2002). Levels of p21 and p53 were increased by more than 10-fold in neuroblastoma cells expressing

DN-TRF2 (Fig. 4d). Increased phosphorylation of p53 at serine 15 was detected within 5 days of transfection with adenoviral DN-TRF2. In hippocampal neurons, levels of p21 were greatly increased within 5 days of expression of DN-TRF2, whereas levels of p53 and p53 activity were unchanged (Fig. 4d). Neurons exhibited a large increase in p53 levels and activity following exposure to the topoisomerase inhibitor etoposide, demonstrating the ability of these cells to induce p53 in response to DNA damage (Fig. 4d). Collectively, these findings suggested that telomere dysfunction has a differential influence on dividing cells and neurons. Inhibition of TRF2 enhances the differentiation of embryonic hippocampal neurons We observed that, in addition to suppressing their proliferation (Fig. 4a), inhibition of TRF-2 function also stimulated neurite outgrowth in SH-SY5Y cells (Fig. 5a). To further evaluate the role of TRF-2 in neuronal differentiation we measured the length of neurites in SH-SY5Y cells that had been stably transfected with either empty vector or DN-TRF2 in the absence or presence of 10 lM of retinoic acid, an agent known to induce neuronal differentiation of these cells (Pahlman et al. 1984). Neurites were significantly longer in cells expressing DN-TRF2 compared to the control cells, and this was the case in either the absence or presence of retinoic acid (Fig. 5a). To determine if inhibition of TRF-2 function affects the differentiation of primary neurons, we transfected embryonic rat hippocampal neurons on culture day 3 with control and DN-TRF2 adenoviral vectors, and measured the length of neurites of randomly chosen pyramidal neurons on culture day 7. The neurites of hippocampal neurons expressing DN-TRF2 were significantly longer than the neurites of the vector-transfected control neurons (Fig. 5b). The DN-TRF2 transfected neurons exhibited increased numbers of cH2AX immunoreactive nuclear foci compared to vector-transfected cells (Fig. 5b, inset), demonstrating an association of a molecular responses to telomere alterations resulting from TRF2 inhibition and neurite outgrowth. Neuronal differentiation is accompanied by the expression of neuron-specific intermediate filament proteins (NF-L, NF-M and NF-H) and microtubule-associated proteins (MAP1b and MAP2) (Riederer 1990). In order to provide further evidence that expression of DN-TRF2 induces neuronal differentiation, we used real-time RT–PCR to determine relative levels of expression of molecular markers of differentiated neurons in embryonic neurons transfected with adenovirus containing b-galactosidase or DN-TRF2 genes. Levels of NF-M, MAP1b and MAP2 mRNAs were significantly increased within 3–5 days post-infection with DN-TRF2 (Fig. 5c). In contrast, levels of nestin mRNA (a protein expressed in neural progenitor cells and immature neurons) was decreased in neurons overexpressing DN-TRF2 compared to cells overexpressing b-galactosidase (Fig. 5c).



Fig. 4 Inhibition of telomeric repeat binding factor 2 (TRF2) function induces senescence and p53 expression in proliferating neural cells, but not in neurons. (a) Dominant negative TRF2 (DN-TRF2) induced senescence in SH-SY5Y cells. The cells were stably transfected with either vector or DN-TRF2 for 15 days. The growth rate of the cells was then quantified by counting cells in defined microscope fields during the next 7 days. The cells were then treated with 1 lM BrdU for 1 h, and then fixed and stained with an anti-BrdU antibody or SA-bgalactosidase (SA-b-Gal) reagent. Compared with vector control (i), expression of DN-TRF2 (ii) resulted in a significant increase of b-galactosidase levels (v; **p < 0.001), a significant decrease in the number of BrdU-labeled cells (v; **p < 0.001) and a significant decrease in the cell growth rate (vi; p < 0.001). Values are the mean and SEM of determinations made in at least six different cultures. (b) DN-TRF2 induced senescence in astrocytes. Astrocytes were infected with adenoviral vectors containing either GFP or DN-TRF2 (50 pfu/ cell). Compared with controls (i), expression of DN-TRF2 (ii) resulted in a significant increase in the number of b-galactosidase positive

astrocytes (iii; **p < 0.001), but did not significantly affect the growth rate of the astrocytes (iv). (c) Expression of DN-TRF2 (200 pfu/cell) did not induce b- galactosidase expression in rat hippocampal neurons. Images showing lack of b-galactosidase staining in neurons 4 days after transfection with vectors containing GFP (i) or DN-TRF2 (ii) in conjunction with IRES-GFP reporter gene (arrows indicates GFP positive neurons). In contrast, the expression of DN-TRF2 induced b-galactosidase production in proliferating neural progenitor cells in neurosphere cultures (iii and iv). (d) DN-TRF2 induced p53 expression in neuroblastoma cells, but not in hippocampal neurons. Immunoblots of cell lysates from neuroblastoma cells (left) or primary hippocampal neurons (right) that had been infected for the indicated number of days with adenoviral vectors containing GFP or DN-TRF2 constructs. Each lane was loaded with 50 lg of protein. Blots were probed with antibodies against the indicated proteins. DN-TRF2 expression induced increases in the levels of both p21 and p53 in SH-SY5Y cells, whereas only p21 was up-regulated by DN-TRF2 in hippocampal neurons.

As neurons differentiate they become electrically excitable, a process that involves up-regulation of voltagedependent Na+, Ca2+ and K+ channels (Dietzel 1995). To

determine if TRF2 plays a role in regulating the development of neuronal excitability, we performed whole-cell patch clamp analyses of Na+, Ca2+ and K+ currents in embryonic


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Fig. 5 Telomere dysfunction enhances neurite outgrowth in SH-SY5Y cells and hippocampal neurons. (a) SH-SY5Y cells stably expressing GFP alone or dominant negative telomeric repeat binding factor 2 (DN-TRF2) in conjunction with IRES-GFP were treated with either retinoic acid or vehicle on post-transfection day 15. Seven days later neurite length was quantified. The fluorescence micrographs show representative microscope fields and the graph shows the results of neurite length measurements (mean and SEM of measurements made in two cultures (total of 88–100 cells analyzed). *p < 0.05 compared to the corresponding vector value. (b) Cultured hippocampal neurons were transfected with either GFP vector or DN-TRF2 in conjunction with IRES-GFP on culture day 3. Four days later neurite length was quantified. The fluorescence micrographs show representative microscope fields and the graph shows the results of neurite

length measurements (mean and SEM of measurements made in three cultures, total of 45–50 neurons analyzed in each culture). *p < 0.05 compared to the vector value. Neurons transfected with DN-TRF2 exhibited increased numbers of cH2AX positive nuclear foci compared to control neurons. The upper inserts show cH2AX immunoreactivity (red) and the lower inserts show phase-contrast images of the neuronal cell body. (c) Levels of the indicated mRNAs were quantified in cultured embryonic hippocampal neurons that had been infected with adenoviral vectors to express either DN-TRF2 or b-galactosidase (b-gal). Expression of DN-TRF2 resulted in an increase in levels of mRNAs for the neuronal markers NF-M (neurofilament medium), MAP1b (microtubule-associated protein 1b) and MAP2 (microtubule-associated protein 2), whereas levels of nestin mRNA (a marker for undifferentiated neural cells) decreased.



Fig. 6 Inhibition of telomeric repeat binding factor 2 (TRF2) increases voltage-gated ion currents in embryonic rat hippocampal neurons. (a) Comparison of membrane capacitance in control (GFP-expressing) and dominant negative TRF-2 (DN-TRF2) expressing neurons (n ¼ 23 and 22 for control and DN-TRF2 transfected neurons, respectively). Embryonic day 18 hippocampal neurons were transfected with GFP vector or DN-TRF2 in conjunction with GFP at 3 DIV and recordings were made 3 days after transfection. Neurons were infected with

adenovirus-DN-TRF2 (200 pfu/cell). (b, c and d) Representative recording traces of voltage-activated Na+ (b), Ca2+ (c) and K+ (d) currents in control and in DN-TRF2 transfected neurons (upper panels). The middle panels show I/V relationships and the lower panels show quantitative comparisons of current densities (values are the mean and SEM; n ¼ 20–23 neurons). *p < 0.05 compared to the value for GFP vector-transfected control neurons.

hippocampal neurons 3 days after transfection with GFP or DN-TRF2-GFP. Recordings were made from DN-TRF2 expressing (GFP-positive) neurons. Resting membrane potentials ranged from )40.3 to )78.3 mV and averaged )65.5 mV in control neurons, and ranged from )43.2 to )75.3 mV and averaged )66.7 mV in DN-TRF2 expressing neurons. Twelve of 23 control (52.2%) and 19 of 22 (86.4%) DN-TRF2 neurons exhibited action potentials during injection of small depolarizing currents under current-clamp conditions. The mean action potential frequencies were 0.35 and 0.56 Hz for the control and the DN-TRF2 neurons, respectively. Neurons expressing DN-TRF2 showed a slight increase in membrane capacitance, but the difference was not statistically significant (Fig. 6a). Voltage-activated Na+ currents were recorded by applying rectangular positive voltage step pulses. Twenty of 23 control neurons (87.0%) and 21 of 22 DN-TRF2 neurons (95.5%) exhibited Na+ currents that were completely blocked by 0.3 lM tetrodotoxin. DN-TRF2 expression induced a significant increase in the Na+ current density without affecting the current–voltage (I/V) relation-

ships (Fig. 6b). Similarly, the densities of both Ca2+ and K+ currents were significantly increased, and the I/V relationships for these currents were unaffected by DN-TRF2 expression (Figs 6c and d). These results show that inhibition of TRF2 increases Na+, Ca2+ and K+ currents, suggesting a role for TRF2 in regulating the acquisition of neuronal excitability during development of the hippocampus. Discussion

Expression of DN-TRF2 resulted in reduced cell proliferation and senescence (as indicated by the accumulation of b-galactosidase) in neuroblastoma cells, astrocytes and neural progenitor cells, effects that are similar to those documented in previous studies of various types of nonneural tumor cells and mitotic somatic cells (Van Steensel et al. 1998; Karlseder et al. 2002). In contrast, DN-TRF2 expression did not induce a senescence phenotype in hippocampal neurons. This differential effect of DN-TRF2 in neurons and mitotic neural cells was apparently not the result of a differential effect on telomeres because DN-TRF2


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expression resulted in displacement of wild-type TRF2 from telomeres, ATM activation and the formation of cH2AXpositive nuclear foci in neuroblastoma cells, astrocytes and hippocampal neurons. ATM is recruited to sites of DNA double-strand breaks and H2AX is a substrate phosphorylated by ATM (Ha et al. 2004). We found that both activated ATM and cH2AX were co-localized at nuclear foci induced by DN-TRF2, suggesting that TRF2 dysfunction is sensed by ATM and/or associated proteins resulting in the recruitment and activation of DNA damage response signaling pathways. Interestingly, despite activation of ATM in each type of cell, telomere dysfunction caused by TRF2 displacement resulted in p53 accumulation in neuroblastoma cells and astrocytes, but not in neurons. Previous studies have provided evidence that p53 plays a pivotal role in the cell cycle arrest and senescence induced by natural telomere shortening (Vaziri and Benchimol 1996) and experimentally induced telomere dysfunction in fibroblasts (Karlseder et al. 1999, 2002). It is therefore likely that p53 also plays an important role in the induction of a senescence-like phenotype in neuroblastoma cells and astrocytes. We found that Chk2 was activated in hippocampal neurons in response to inhibition of TRF2 function, but not in response to treatment with the topoisomerase inhibitor etoposide. Inhibition of TRF2 function did not induce p53 in neurons, whereas etoposide did induce p53. Others have shown that Chk2 activation is not required for topoisomerase inhibitor-induced p53 activation in cultured neurons (Keramaris et al. 2003). The pathways triggered by TRF-2 inhibition and topoisomerase inhibitors in neurons therefore diverge downstream of ATM activation. Although DN-TRF2 elicited a telomere-associated DNA damage response in neurons, p53 was not induced and no apparent adverse consequences were observed. However, previous studies have provided evidence that TERT, the catalytic subunit of telomerase, serves a cell survivalpromoting function in embryonic hippocampal neurons and that genetic depletion of TERT sensitizes neurons to apoptosis under conditions of trophic factor deprivation or exposure to oxidative and metabolic insults (Fu et al. 2000, 2002). TERT also protected cultured hippocampal neurons against death induced by DNA damaging agents that induce double-strand breaks (Lu et al. 2001). Moreover, transgenic mice overexpressing TERT exhibit increased resistance of neurons to ischemic brain injury in a model of stroke (Kang et al. 2004). In light of the accumulating evidence that proteins involved in telomere function can influence DNA damage-related responses in neurons, it will be of considerable interest to elucidate the roles of TRF2 and other telomere-associated proteins in neuronal survival and plasticity in the contexts of aging and neurodegenerative conditions. Expression of DN-TRF2 enhanced the differentiation of cultured hippocampal neurons as demonstrated by enhanced neurite outgrowth, increased expression of MAP2 and neurofilament-M (markers of mature neurons), and decreased

expression of nestin (a marker of neural stem cells and immature neurons). Moreover, our whole cell patch clamp data showed that neurons expressing DN-TRF2 exhibit larger Na+ and Ca2+ currents, suggesting enhanced membrane excitability, a property of mature neurons. p21 may be involved in both early and later stages of neuronal differentiation induced by TRF2 inhibition. Increasing evidence suggests that p21 plays a key role in cell cycle exit and neuronal differentiation of neural progenitor cells, and in the acquisition of mature neuronal phenotypes. A role for p21 in the differentiation of stem cells into neurons in vivo is suggested from studies of hippocampal neurogenesis in rats (Takahashi et al. 1999) and olfactory neurogenesis in mice (Legrier et al. 2001). Similar to neuroblastoma cells in which TRF2 function is inhibited, p21 is up-regulated during differentiation of neuroblastoma cells in culture (Yano et al. 2005) and mediates nerve growth factor-induced differentiation of PC12 cells (Erhardt and Pittman 1998). In addition, expression of p21 induces neurite outgrowth and branching in cultured hippocampal neurons (Tanaka et al. 2002), an effect similar to that observed in hippocampal neurons expressing DN-TRF2. Although the present and previous findings suggest a key regulatory role for p21 in neurogenesis and neuronal maturation, the specific molecular cascades downstream of p21 remain to be determined. The proteins encoded by the genes responsive to TRF2 inhibition are therefore among the candidates for such downstream effectors of p21-mediated neuronal differentiation. Expression of DN-TRF2 resulted in the formation of nuclear foci of ATM and phosphorylated H2AX that colocalized with TRF1, consistent with recruitment of DNA damage response proteins to telomeres. Roles for ATM and/ or p21 in the enhancement of neuronal differentiation induced by DN-TRF2 are suggested by data showing that neurite outgrowth is impaired in Purkinje neurons from ATM-deficient mice (Cheng et al. 2003) and that p21 enhances neurite outgrowth in cultured hippocampal neurons (Tanaka et al. 2002). A role for p21 in neurite outgrowth is suggested by studies showing that p21 expression is increased during the regenerative outgrowth of axons (Bonilla et al. 2002). During development of the nervous system, telomerase is down-regulated as neural precursor cells differentiate into neurons and as the neurons mature, while TRF2 expression is maintained during this period of development (Klapper et al. 2001). Our data therefore suggest a role for TRF2 in maintenance of telomere function during and after the transition of mitotic cells with telomerase activity to post-mitotic neurons that lack telomerase activity. The present study is the first to examine the consequences of telomere dysfunction in astrocytes, the most abundant cell type in the brain. Astrocytes serve many important functions, including providing trophic and metabolic support to neurons, modulating neurotransmitter signaling and regulating responses of the brain to injury and inflammation (Ridet



et al. 1997; Schousboe et al. 2004). During normal aging, astrocyte proliferation is decreased and these cells acquire phenotypes consistent with a senescence-like state; they hypertrophy, accumulate intermediate filaments and express b-galactosidase (Fanton et al. 2001; Cotrina and Nedergaard 2002). Similar, but more pronounced changes in astrocytes occur in several different neurodegenerative disorders including Alzheimer’s and Parkinson’s diseases (Schipper 1996). We found that telomere dysfunction induced p53 and senescence in astrocytes. Astrocytes lacking p53 exhibit resistance to senescence (Evans et al. 2003), which is consistent with a role for up-regulation of p53 in astrocyte senescence induced by TRF2 dysfunction. A role for telomere damage in astrocyte senescence in vivo is consistent with studies showing that DNA damage, including both single and double-strand breaks, are increased in association with senescence-like changes in astrocytes in the brain during aging and in patients with dementia (Bhaskar and Rao 1994; Martin et al. 2001). In addition to an adverse effect on astrocytes themselves, telomere dysfunction in astrocytes might adversely affect neighboring neurons. Indeed, studies of fibroblasts have suggested that senescent cells may alter the function of neighboring cells by secreting aging related factor such as inflammatory cytokines (Campisi 2005). Because the inhibition of endogenous TRF2 functions by DN-TRF2 results from displacement of endogenous TRF2 from telomeres, the biological effect of DN-TRF2 is expected to be maximal with an amount of DN-TRF2 equal to the level of endogenous TRF2. Our immunoblot analyses showed that levels of DN-TRF2 in cells transfected with adenovirus-DN-TRF2 were at least two-fold greater than the level of endogenous TRF2 in all three cell types, although SH-SY5Y cells were more efficiently infected than were astrocytes and primary neurons. The amount of adenovirus entering cells depends on the amount of CAR receptor on the cell surface which is cell type-dependent. We chose doses of adenovirus that resulted in relatively high levels of expression, but with negligible cytotoxicity. In order to allow a valid comparison between astrocytes and neurons, or SH-SY5Y cells and neurons, we used a five times greater dose of adenovirus in astrocytes and neurons (50 pfu/cell) than in SH-SY5Y cells (10 pfu/cell). This resulted in similar levels of DN-TRF2 protein in SH-SY5Y cells and neurons (Fig. 2d) and astrocytes (data not shown). The results showed that when DN-TRF2 levels are similar, the magnitude of the DNA damage response (phosphorylated H2AX) in mature neurons is much less than in SH-SY5Y cells. Even when hippocampal neurons were transfected with a higher dose of adenovirus (200 pfu/cell) (Figs 5b and c), the resulting very high levels of expression of DN-TRF2 enhanced c-H2AX levels and promoted neuronal differentiation, but did not adversely affect cell survival. Our findings have implications, not only for fundamental mechanisms of neurogenesis and neuronal differentiation,

but also for the pathogenesis and treatment of neural tumors. Cancers form when damaged cells escape apoptosis and senescence, failsafe mechanisms that counteract excessive mitogenic and cell survival signalling from activated oncogenes (Wechsler-Reya and Scott 2001; Campisi 2005). Cell senescence involves a permanent form of cell cycle arrest that can be driven by telomere attrition (Herbig et al. 2004). Several recent studies have shown that cellular senescence can also occur in vivo (Chen et al. 2005; Michaloglou et al. 2005). Although the molecular mechanisms of senescence are not fully understood, a telomere damage response (Karlseder et al. 1999), oncogenes (Chen et al. 2005; Michaloglou et al. 2005) and accumulation and activation of p53 play important roles. There is therefore considerable overlap in mechanisms of apoptosis and senescence. We found that overexpression of DN-TRF2 using an adenoviral vector induced senescence of neuroblastoma cells with no apparent adverse effect on neurons. This suggests that it may be possible to use telomere-directed therapies to halt the growth of brain tumors without damaging neurons. Acknowledgements This research was supported the National Institute on Aging Intramural Research Program.

References d’Adda di Fagagna F., Reaper P. M., Clay-Farrace L., Fiegler H., Carr P., Von Zglinicki T., Saretzki G., Carter N. P. and Jackson S. P. (2003) A DNA damage checkpoint response in telomere-initiated senescence. Nature 426, 194–198. Andegeko Y., Moyal L., Mittelman L., Tsarfaty I., Shiloh Y. and Rotman G. (2001) Nuclear retention of ATM at sites of DNA double strand breaks. J. Biol. Chem. 276, 38 224–38 230. Bhaskar M. S. and Rao K. S. (1994) Altered conformation and increased strand breaks in neuronal and astroglial DNA of aging rat brain. Biochem. Mol. Biol. Int. 33, 377–384. Blanc E. M., Keller J. N., Fernandez S. and Mattson M. P. (1998) 4-Hydroxynonenal, a lipid peroxidation product, inhibits glutamate transport in astrocytes. Glia 22, 149–160. Bonilla I. E., Tanabe K. and Strittmatter S. M. (2002) Small proline-rich repeat protein 1A is expressed by axotomized neurons and promotes axonal outgrowth. J. Neurosci. 22, 1303– 1315. Broccoli D., Smogorzewska A., Chong L. and de Lange T. (1997) Human telomeres contain two distinct Myb-related proteins, TRF1 and TRF2. Nat. Genet. 17, 231–235. Burma S., Chen B. P., Murphy M., Kurimasa A. and Chen D. J. (2001) ATM phosphorylates histone H2AX in response to DNA doublestrand breaks. J. Biol. Chem. 276, 42 462–42 467. Cai J., Cheng A., Luo Y., Lu C., Mattson M. P., Rao M. S. and Furukawa K. (2004) Membrane properties of rat embryonic multipotent neural stem cells. J. Neurochem. 88, 212–226. Campisi J. (2005) Senescent cells, tumor suppression, and organismal aging: good citizens, bad neighbors. Cell 120, 513–522. Chan S. R. and Blackburn E. H. (2004) Telomeres and telomerase. Philos. Trans. R. Soc. Lond. B Biol. Sci. 359, 109–121.


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Chen Z., Trotman L. C., Shaffer D. et al. (2005) Crucial role of p53dependent cellular senescence in suppression of Pten-deficient tumorigenesis. Nature 436, 725–730. Cheng A., Wang S., Rao M. S. and Mattson M. P. (2003) Nitric oxide acts in a positive feedback loop with BDNF to regulate neural progenitor cell proliferation and differentiation in the mammalian brain. Dev. Biol. 258, 319–333. Cotrina M. L. and Nedergaard M. (2002) Astrocytes in the aging brain. J. Neurosci. Res. 67, 1–10. De Leon G. A., Grover W. D. and Huff D. S. (1976) Neuropathologic changes in ataxia-telangiectasia. Neurology 26, 947–951. De Stefano N., Dotti M. T. and Battisti C. (2003) MR evidence of structural and metabolic changes in brains of patients with Werner’s syndrome. J. Neurol. 250, 1169–1173. Dietzel I. D. (1995) Voltage-gated ion currents in embryogenesis. Perspect. Dev. Neurobiol. 2, 293–308. Digweed M., Reis A. and Sperling K. (1999) Nijmegen breakage syndrome: consequences of defective DNA double strand break repair. Bioessays 21, 649–656. Dimri G. P., Lee X., Basile G., Acosta M., Scott G., Roskelley C., Medrano E. E., Linskens M., Rubelij I. and Pereira-Smith O. (1995) A biomarker that identifies senescent human cells in culture and in aging skin in vivo. Proc. Natl Acad. Sci. USA 92, 9363– 9367. Du B., Ohmichi M., Takahashi K. et al. (2004) Both estrogen and raloxifene protect against b-amyloid-induced neurotoxicity in estrogen receptor-transfected PC12 cells by activation of telomerase activity via Akt cascade. J. Endocrinol. 183, 605–615. Elmore L. W., Di Rehder C. W. X., McChesney P. A., Jackson-Cook C. K., Gewirtz D. A. and Holt S. E. (2002) Adriamycin-induced senescence in breast tumor cells involves functional p53 and telomere dysfunction. J. Biol. Chem. 277, 35 509–35 515. Erhardt J. A. and Pittman R. N. (1998) p21WAF1 induces permanent growth arrest and enhances differentiation, but does not alter apoptosis in PC12 cells. Oncogene 16, 443–451. Evans R. J., Wyllie F. S., Wynford-Thomas D., Kipling D. and Jones C. J. (2003) A P53-dependent, telomere-independent proliferative life span barrier in human astrocytes consistent with the molecular genetics of glioma development. Cancer Res. 63, 4854– 4861. Fanton C. P., McMahon M. and Pieper R. O. (2001) Dual growth arrest pathways in astrocytes and astrocytic tumors in response to Raf-1 activation. J. Biol. Chem. 276, 18871–18877. Fernandez-Capetillo O., Lee A., Nussenzweig M. and Nussenzweig A. (2004) H2AX: the histone guardian of the genome. DNA Repair 3, 959–967. Ferron S., Mira H., Franco S., Cano-Jaimez M., Bellmunt E., Ramirez C., Farinas I. and Blasco M. A. (2004) Telomere shortening and chromosomal instability abrogates proliferation of adult but not embryonic neural stem cells. Development 131, 4059–4070. Filippova N., Sedelnikova A., Tyler W. J., Whitworth T. L., Fortinberry H. and Weiss D. S. (2001) Recombinant GABA (C) receptors expressed in rat hippocampal neurons after infection with an adenovirus containing the human rho1 subunit. J. Physiol. 535, 145–153. Fotiadou P., Henegariu O. and Sweasy J. B. (2004) DNA polymerase beta interacts with TRF2 and induces telomere dysfunction in a murine mammary cell line. Cancer Res. 64, 3830–3837. Fu W., Begley J. G., Killen M. W. and Mattson M. P. (1999) Antiapoptotic role of telomerase in pheochromocytoma cells. J. Biol. Chem. 274, 7264–7271. Fu W., Killen M., Culmsee C., Dhar S., Pandita T. K. and Mattson M. P. (2000) The catalytic subunit of telomerase is expressed in devel-

oping brain neurons and serves a cell survival-promoting function. J. Mol. Neurosci. 14, 3–15. Fu W., Lu C. and Mattson M. P. (2002) Telomerase mediates the cell survival-promoting actions of brain-derived neurotrophic factor and secreted amyloid precursor protein in developing hippocampal neurons. J. Neurosci. 22, 10710–10719. Furukawa K., Barger S. W., Blalock E. M. and Mattson M. P. (1996) Activation of K+ channels and suppression of neuronal activity by secreted beta-amyloid-precursor protein. Nature 379, 74–78. Furukawa K., Wang Y., Yao P. J., Fu W., Mattson M. P., Itoyama Y., Onodera H., D’Souza I., Poorkaj P. H., Bird T. D. and Schellenberg G. D. (2003) Alteration in calcium channel properties is responsible for the neurotoxic action of a familial frontotemporal dementia tau mutation. J. Neurochem. 87, 427–436. Gartel A. L. and Tyner A. L. (2002) The role of the cyclin-dependent kinase inhibitor p21 in apoptosis. Mol. Cancer Ther. 1, 639–649. Griffith J. D., Comeau L., Rosenfield S., Stansel R. M., Bianchi A., Moss H. and de Lange T. (1999) Mammalian telomeres end in a large duplex loop. Cell 97, 503–514. Guiducci C., Cerone M. A. and Bacchetti S. (2001) Expression of mutant telomerase in immortal telomerase-negative human cells results in cell cycle deregulation, nuclear and chromosomal abnormalities and rapid loss of viability. Oncogene 20, 714–725. Ha L., Ceryak S. and Patiemo S. R. (2004) Generation of S phasedependent DNA double-strand breaks by Cr(VI) exposure: involvement of ATM in Cr(VI) induction of gamma-M2AX. Carcinogenesis 25, 2265–2274. Herbig U., Jobling W. A., Chen B. P., Chen D. J. and Sedivy J. M. (2004) Telomere shortening triggers senescence of human cells through a pathway involving ATM, p53, and p21 (CIP1), but not p16 (INK4a). Mol. Cell 14, 501–513. Kang H. J., Choi Y. S., Hong S. B. et al. (2004) Ectopic expression of the catalytic subunit of telomerase protects against brain injury resulting from ischemia and NMDA-induced neurotoxicity. J. Neurosci. 24, 1280–1287. Karlseder J., Broccoli D., Dai Y., Hardy S. and de Lange T. (1999) p53and ATM-dependent apoptosis induced by telomeres lacking TRF2. Science 283, 1321–1325. Karlseder J., Smogorzewska A. and de Lange T. (2002) Senescence induced by altered telomere state, not telomere loss. Science 295, 2446–2449. Karlseder J., Hoke K., Mirzoeva O. K., Bakkenist C., Kastan M. B., Petrini J. H. and de Lange T. (2004) The telomeric protein TRF2 binds the ATM kinase and can inhibit the ATM-dependent DNA damage response. PLoS Biol. 2, E240. Keramaris E., Hirao A., Slack R. S., Mak T. W. and Park D. S. (2003) Ataxia telangicctasia-mutated protein can regulate p53 and neuronal death independent of Chk2 in response to DNA damage. J. Biol. Chem. 278, 37 782–37 789. Klapper W., Shin T. and Mattson M. P. (2001) Differential regulation of telomerase activity and TERT expression during brain development in mice. J. Neurosci. Res. 64, 252–260. Kohrmann M., Haubensak W., Hemraj I., Kaether C., Lessmann V. J. and Kiebler M. A. (1999) Fast, convenient, and effective method to transiently transfect primary hippocampal neurons. J. Neurosci. Res. 58, 831–835. Lammens M., Hiel J. A., Gabreels F. J., ban Engelen B. G., van den Heuvel L. P. and Weemaes C. M. (2003) Nijmegen breakage syndrome: a neuropathological study. Neuropediatrics 34, 189– 193. Legrier M. E., Ducray A., Propper A., Chao M. and Kastner A. (2001) Cell cycle regulation during mouse olfactory neurogenesis. Cell Growth Differ. 12, 591–601.



Lin J., Smith D. L. and Blackburn E. H. (2004) Mutant telomere sequences lead to impaired chromosome separation and unique checkpoint response. Mol Biol. Cell 15, 1623–1634. Lu C., Fu W. and Mattson M. P. (2001) Telomerase protects developing neurons against DNA damage-induced cell death. Dev. Brain Res. 131, 167–171. Martin J. A., Craft D. K., Su J. H., Kim R. C. and Cotman C. W. (2001) Astrocytes degenerate in frontotemporal dementia: possible relation to hypoperfusion. Neurobiol. Aging 22, 95–207. Mattson M. P., Barger S. W., Begley J. G. and Mark R. J. (1995) Calcium, free radicals, and excitotoxic neuronal death in primary cell culture. Meth. Cell Biol. 46, 187–216. McKinnon P. J. (2004) ATM and ataxia telangiectasia. EMBO Rep. 5, 772–776. Michaloglou C., Vredeveld L. C., Soengas M. S., Denoyelle C., Kuilman T., van der Horst C. M., Majoor D. M., Shay J. W., Mooi W. J. and Peeper D. S. (2005) BRAFE600-associated senescence-like cell cycle arrest of human naevi. Nature 43, 720–724. Opresko P. L., von Kobbe C., Laine J. P., Harrigan J., Hickson I. D. and Bohr V. A. (2002) Telomere-binding protein TRF2 binds to and stimulates the Werner and Bloom syndrome helicases. J. Biol. Chem. 277, 41 110–41 119. Opresko P. L., Cheng W. H., von Kobe C., Harrigan J. A. and Bohr V. A. (2003) Werner syndrome and the function of the Werner protein; what they can teach us about the molecular aging process. Carcinogenesis 24, 791–802. Opresko P. L., Otterlei M., Graakjaer J., Bruheim P., Dawut L., Kolvraa S., May A., Seidman M. M. and Bohr V. A. (2004) The Werner syndrome helicase and exonuclease cooperate to resolve telomeric D loops in a manner regulated by TRF1 and TRF2. Mol. Cell 14, 763–774. Pahlman S., Ruusala A. I., Abrahamsson L., Mattsson M. E. and Esscher T. (1984) Retinoic acid-induced differentiation of cultured human neuroblastoma cells: a comparison with phorbolester-induced differentiation. Cell Differ. 14, 135–144. Ridet J. L., Malhotra S. K., Privat A. and Gage F. H. (1997) Reactive astrocytes: cellular and molecular cues to biological function. Trends Neurosci. 20, 570–577. Riederer B. M. (1990) Some aspects of the neuronal cytoskeleton in development. Eur. J. Morphol. 28, 347–378. Sancar A., Lindsey-Boltz L. A., Unsal-Kaccmaz K. and Linn S. (2004) Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annu. Rev. Biochem. 73, 39–85. Schipper H. M. (1996) Astrocytes, brain aging and neurodegeneration. Neurobiol. Aging 17, 467–480.

Schousboe A., Sarup A., Bak L. K., Waagepetersen H. S. and Larsson O. M. (2004) Role of astrocytic transport processes in glutamatergic and GABAergic neurotransmission. Neurochem. Int. 45, 521–527. Smogorzewska A. and de Lange T. (2002) Different telomere damage signaling pathways in human and mouse cells. EMBO J. 21, 4338– 4348. Stansel R. M., de Lange T. and Griffith J. D. (2001) T-loop assembly in vitro involves binding of TRF2 near the 3¢ telomeric overhang. EMBO J. 20, 5532–5540. Takahashi J., Palmer T. D. and Gage F. H. (1999) Retinoic acid and neurotrophins collaborate to regulate neurogenesis in adult-derived neural stem cell cultures. J. Neurobiol. 38, 65–81. Takai H., Smogorzewska A. and de Lange T. (2003) DNA damage foci at dysfunctional telomeres. Curr. Biol. 13, 1549–1556. Tanaka H., Yamashita T., Asada M., Mizutani S., Yoshikawa H. and Tohyama M. (2002) Cytoplasmic p21 (Cip1/WAF1) regulates neurite remodeling by inhibiting Rho-kinase activity. J. Cell Biol. 158, 321–329. Van Steensel B., Smogorzewska A. and de Lange T. (1998) TRF2 protects human telomeres from end-to-end fusions. Cell 92, 401–413. Vaziri H. and Benchimol S. (1996) From telomere loss to p53 induction and activation of a DNA-damage pathway at senescence: the telomere loss/DNA damage model of cell aging. Exp. Gerontol. 31, 295–301. Vousden K. H. (2002) Switching from life to death: the Miz-ing link between Myc and p53. Cancer Cell 2, 351–352. Wechsler-Reya R. and Scott M. P. (2001) The developmental biology of brain tumors. Annu. Rev. Neurosci. 24, 385–428. Wong K. K., Maser R. S., Bachoo R. M., Menon J., Carrasco D. R., Gu Y., Alt F. W. and DePinho R. A. (2003) Telomere dysfunction and Atm deficiency compromises organ homeostasis and accelerates ageing. Nature 421, 643–648. Yano M., Okano H. J. and Okano H. (2005) Involvement of Hu and heterogeneous nuclear ribonucleoprotein K in neuronal differentiation through p21 mRNA post-transcriptional regulation. J. Biol. Chem. 280, 12 690–12 699. Zhang P., Chan S. L., Fu W., Mendoza M. and Mattson M. P. (2003) TERT suppresses apoptotis at a premitochondrial step by a mechanism requiring reverse transcriptase activity and 14-3-3 protein-binding ability. FASEB J. 17, 767–769. Zhu X. D., Kuster B., Mann M., Petrini J. H. and de Lange T. (2000) Cell-cycle-regulated association of RAD50/MRE11/NBS1 with TRF2 and human telomeres. Nat. Genet. 25, 347–352.
