Accumulation of annexin A5 at the nuclear envelope is a biomarker of cellular aging

Mechanisms of Ageing and Development 133 (2012) 508–522

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Accumulation of annexin A5 at the nuclear envelope is a biomarker of cellular aging Karolin Klement a, Christian Melle b, Ulrike Murzik a, Stephan Diekmann a, Johannes Norgauer c, Peter Hemmerich a,* a b c

Leibniz-Institute for Age Research – Fritz Lipmann Institute, Jena, Germany Biomolecular Photonics Group, University Hospital Jena, Germany Department of Dermatology and Allergology, University of Jena, Germany

A R T I C L E I N F O

A B S T R A C T

Article history: Received 22 December 2011 Received in revised form 20 April 2012 Accepted 13 June 2012 Available online 20 June 2012

Cellular senescence is a permanent cell cycle arrest induced by short telomeres or oncogenic stress in vitro and in vivo. Because no single of the established biomarkers can reliably identify senescent cells, the application of new ones may aid the diagnosis of aged cells. Here we show that annexin A5 accumulates at the nuclear envelope during replicative and drug-induced cellular senescence in primary human fibroblasts. This new cellular aging phenotype that we have termed SA-ANX5 (senescence-associated accumulation at the nuclear envelope of annexin A5) is as efficient and quantitative as the wellestablished senescence-associated b-galactosidase activity assay and p21 immunoreactivity. SA-ANX5 is also observed in aged human skin where is exclusively detected in DNA damage foci-positive/Ki-67negative cells. We also observed that depletion of annexin A5 by siRNA in human fibroblasts accelerates premature senescence through the p38MAP kinase pathway. These observations establish SA-ANX5 as a new biomarker for cellular aging and implicate a functional role for annexin A5 in cellular senescence. ß 2012 Elsevier Ireland Ltd. All rights reserved.

Key words: Cellular aging Senescence Annexin A5 Human skin Biomarker Cell cycle

1. Introduction Cellular senescence is a state of irreversible growth arrest that somatic cells enter as a result of replicative exhaustion through telomere erosion or in response to oncogenic stress (Harley et al., 1990; Serrano et al., 1997). In cell culture systems, many primary human cells can pass 50–80 population doublings (PD) until the senescence phenotype is fully established (Hayflick and Moorhead, 1961; Hayflick, 1965). In contrast to quiescence, senescent cells do not exit the cell cycle but enter a state of ‘active arrest’ at very advanced points of the G1, G1/S and G2 phase of the cell cycle (Blagosklonny, 2011). Although senescent cells are unable to proliferate and are resistant to mitogenic stimuli (Phillips et al., 1984), they are metabolically active and can be maintained in culture for several months (Matsumura et al., 1979). The concept of cellular senescence is now established as a general mechanism that triggers an irreversible proliferation arrest of cells caused by various stresses, including

* Corresponding author at: Leibniz Institute for Age Research – Fritz Lipmann Institute, Beutenbergstr. 11, 07745 Jena, Germany. Tel.: +49 03641 656262; fax: +49 03641 656310. E-mail addresses: kklement@fli-leibniz.de (K. Klement), [email protected] (C. Melle), umurzik@fli-leibniz.de (U. Murzik), diekmann@fli-leibniz.de (S. Diekmann), [email protected] (J. Norgauer), phemmer@fli-leibniz.de (P. Hemmerich). 0047-6374/$ – see front matter ß 2012 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.mad.2012.06.003

oxidative damage, telomere dysfunction, DNA damage and several chemotherapeutic drugs (Ben-Porath and Weinberg, 2005). Short or dysfunctional telomeres generate a persistent DNA damage response (DDR), which initiates and maintains the senescence-associated growth arrest (d’Adda di Fagagna et al., 2003; Karlseder et al., 2002; Takai et al., 2003; Herbig et al., 2004; Aubert and Lansdorp, 2008). However, a non-telomeric DNA damage response appears to play an equivalent role in triggering cellular senescence, both in vitro and in vivo (Sedelnikova et al., 2004, 2008; Nakamura et al., 2008; Le et al., 2010). Consistent with this idea, the DNA double strand break (DSB)-induced cell cycle arrest after g-irradiation represents the prototype of stress induced premature cellular senescence (SIPS) (Di Leonardo et al., 1994; Toussaint et al., 2000). Chemotherapeutic drugs, oncogenes, oxidative stress and inappropriate culture conditions can also induce SIPS, in most cases by triggering a telomereindependent DDR (Toussaint et al., 2002; d’Adda di Fagagna, 2008). DNA damage-induced premature cellular senescence is a hallmark of tumor cells in vitro and in vivo (te Poele et al., 2002). DNA damage induced senescence functionally correlates with the appearance of senescence-associated persistent g-H2AX foci (SDF) at which several key DDR proteins such as ATM, 53BP1, NBS1 and MDC1 accumulate (d’Adda di Fagagna et al., 2003; Herbig et al., 2004; Sedelnikova et al., 2004). SDF originate at eroded or dysfunctional telomeres as well as at non-telomeric sites (Takai et al., 2003;

K. Klement et al. / Mechanisms of Ageing and Development 133 (2012) 508–522

Sedelnikova et al., 2008; Nakamura et al., 2008; Wang et al., 2009). Signals generated at SDF are amplified by the kinases ATM and ATR which activate CHK1 and CHK2. These kinases communicate the damage signals to the cell cycle machinery by phosphorylation and activation of several key cell cycle proteins (von Zglinicki et al., 2005). The senescence-associated cell cycle arrest is mainly implemented and maintained through the p53/p21 and p16/pRB pathways (Shay et al., 1991; Tahara et al., 1995; Hara et al., 1996; Kulju and Lehman, 1995; Stein et al., 1999). In vivo, the cell cycle arrest triggered by oncogene-induced cellular senescence (OIS) limits cancer progression (Krizhanovsky et al., 2008; Prieur and Peeper, 2008). Senescent cells are prevalent in some pre-malignant tumors arguing that cellular senescence represses cancer by imposing a cell-autonomous block to the proliferation of oncogenically activated cells (Serrano and Blasco, 2007; Collado and Serrano, 2010). The number of senescent cells as detected by expression of the aging markers SDF, p16 or senescence-associated b-galactosidase (SA-b-Gal) activity increases during aging in various mouse and human tissues (Zindy et al., 1997; Nielsen et al., 1999; Melk et al., 2004; Sedelnikova et al., 2004; Krishnamurthy et al., 2004; Herbig et al., 2006; Ressler et al., 2006; Jeyapalan et al., 2007; Sedelnikova et al., 2008; Liu et al., 2009; Wang et al., 2009; Geng et al., 2010). These observations are compatible with the potentially deleterious effects of senescent cells on tissue microenvironments in agerelated diseases (Vijg and Campisi, 2008; Jeyapalan and Sedivy, 2008; Kuilman et al., 2010; Rodier and Campisi, 2011). There is now compelling evidence that senescent cells also accumulate in aging tissues (Jeyapalan and Sedivy, 2008; Hornsby, 2010) where they are causally implicated in the generation of age-related phenotypes and in tumor immune surveillance (Baker et al., 2011; Kang et al., 2011). Removal of senescent cells can prevent or delay tissue dysfunction and extend healthspan (Baker et al., 2011). In recognition of the pathophysiological relevance of cellular senescence (Campisi and d’Adda di Fagagna, 2007), it is essential to reliably detect senescent cells in vitro and in vivo (Kuilman et al., 2010; Sikora et al., 2011). Specific morphological alterations of senescent cells in vitro include a flat and enlarged cell shape (Wang and Gundersen, 1984). Beside the cell cycle arrest, aged cells are primarily characterized by their inability to replicate DNA in vitro as well as in vivo (Schneider et al., 1979). Consequently, cell cycle related factors such as p16, p21 or p53 are instrumental as biomarkers of aging (Cristofalo and Pignolo, 1996; Jansen-Du¨rr, 1998; Lawless et al., 2010). The most commonly used senescence marker is expression of SA-b-Gal (Dimri et al., 1995). Although some limitations have been reported (Severino et al., 2000), increased SA-b-Gal activity evolved as a reliable biomarker for the course of replicative senescence if used under standardized conditions (Maier et al., 2007; Debacq-Chainiaux et al., 2009). Related to telomere dysfunction and DNA damage, detection of SDF has also been established as a cellular aging marker (Wang et al., 2009; Lawless et al., 2010). During cellular senescence, specific chromatin alterations occur (Funayama and Ishikawa, 2007), among which the formation of senescence-associated heterochromatin foci (SAHF) has been established as a senescence marker in some (Narita et al., 2003; Zhang et al., 2005) but no all settings (Kosar et al., 2011; Di Micco et al., 2011). Additional cellular and molecular signatures of cellular senescence include lipofuscin accumulation in the cytoplasm (von Zglinicki et al., 1995), free radicals (Lu and Finkel, 2008), inflammatory-type secreted molecules (SASP/SMS) (Shelton et al., 1999; Kuilman and Peeper, 2009; Coppe´ et al., 2010), an elevated number of promyelocytic leukemia (PML) nuclear bodies (Jiang and Ringertz, 1997), DNASCARS (DNA segments with chromatin alterations reinforcing senescence) (Rodier et al., 2011) and chromosomal instability (Mosieniak and Sikora, 2010).

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Despite this rich toolkit of methods, unequivocal identification of senescent cells has significant limitations: (i) no single of the aging markers can reliably identify senescent cells in isolation, (ii) most markers have been studied only in a limited number of tissues, (iii) epiphenomenological markers such as SA-b-Gal, may be restricted to specific settings, (iv) many biomarkers have only been tested in vitro, and (v) biomarkers may not be specific for cellular senescence (Shmookler Reis, 1988; Cristofalo and Pignolo, 1996; Kuilman et al., 2010; Sikora et al., 2011). Therefore, the identification and application of new senescence markers will improve the diagnosis of aged cells in vitro and in vivo. We report here that the accumulation of annexin A5 at the nuclear envelope (SA-ANX5) is a unique feature of senescent cells in vitro and in vivo. SA-ANX5 is equally well suited to detect senescent cells as SA-bGal and p21 at the single-cell level in vitro and we also detected SAANX5 specifically in pATM-positive/Ki-67-negative cells of human skin. Depletion of annexin A5 accelerates DNA damage induced senescence in primary human fibroblast cultures via p38MAP kinase signaling indicating an important role for annexin A5 in the cellular stress response associated with cellular senescence. 2. Materials and methods 2.1. Cell culture Primary human lung and foreskin fibroblasts MRC-5, WI-38 and BJ, Hela and U-2 OS cell lines were obtained from ATCC (LGC Promochem GmbH, Wesel, Germany). Fibroblasts were cultured as recommended in Dulbecco’s Modified Eagle’s Medium DMEM with L-glutamine (PAA Laboratories, Pasching, Austria) supplemented with 10% fetal calf serum (PAA Laboratories, Pasching, Austria) until the Hayflick limit was reached. At their arrival, the primary cells had passed already until the following population doublings: MRC-5 (22 PD), WI-38 (24 PD), and BJ (21 PD). During passaging, re-freezing and re-thawing was not performed to minimize induction of premature senescence due to culture stress. The cells were passaged in a 1:4 or 1:2 ratio. The exact PD was calculated by using the equation PD = PD at plating + ln(no.harvest/no.seeded)/ln 2. For immunofluorescence analysis cells seeded onto 15 mm cover slips (Saur Laborbedarf, Reutlingen, Germany) and grown for 48 h for attachment. Premature senescence was induced by incubation of young cells with normal growth medium supplemented with 50 mM 5-bromodesoxyuridine (BrdU) (Becton Dickinson, Heidelberg, Germany) and 10 mM distamycin A (DMA) (Sigma–Aldrich, Taufkirchen, Germany) for six to twelve days. 2.2. Preparation of nuclear fractions Isolation of nuclei was performed as described by Remboutsika et al. (1999). Briefly, cells from sub-confluent cultures were dissolved in buffer N (0.3% NP-40, 15 mM Tris– HCl pH 7.5; 60 mM KCl, 15 mM NaCl2, 5 mM MgCl2, 1 mM CaCl2, 1 mM Dithiotreitol, 2 mM Na-Vanadate, and 250 mM sucrose) containing complete-protease inhibitor (Roche) for 5 min on ice. After centrifugation, the supernatant was transferred into a new tube. The remaining pellet (nuclei) was washed two times in buffer N without NP40 and subjected to SDS-PAGE, Western blotting and mass spectrometry. 2.3. Protein identification by peptide fingerprint mapping and mass spectrometry Isolated nuclei were resuspended in lysis buffer (100 mM NaH2PO4 pH 7.5; 5 mM EDTA, 2 mM MgCl2, 3 mM 2-b-mercaptoethanol, 0.1% CHAPS, 0.5 mM Leupeptin, and 0.1 mM PMSF), separated by SDS-PAGE and isolated single bands were identified by SELDI-MS (surface enhanced laser desorption/ionization-mass spectrometry) as described previously (Murzik et al., 2008). Briefly, excised gel bands were trypsin digested at 37 8C over night and supernatants applied to gold arrays (Bio-Rad). After addition of the energy absorbance matrix (CHCA), peptide fragment masses were analyzed in a ProteinChip Reader (series 4000 instrument; Bio-Rad) by SELDI-MS. Proteins were identified using the fragment masses searching in a public domain available database (http://prowl.rockefeller.edu/prowl-cgi/profound.exe). The search revealed annexin A5 as the best candidate with an estimated Z-score of 2.43. 2.4. Western blotting 1xPBS washed cells (whole cell extracts), cytoplasmic fractions and nuclei were resuspended in loading buffer and incubated at 100 8C for 5 min. After SDS-PAGE proteins were transferred onto nitrocellulose transfer membranes (Whatman) using a Fastblot B34 (Biometra). Membranes were blocked with 5%-dry milk solution (Roth). After incubation with appropriate dilutions of primary antibodies for 1 h membranes were incubated with secondary antibodies coupled to horse radish peroxidase (HRP) (1 h). Enzymatic light detection was performed using the ECL system (GE Healthcare). The following antibodies were used for Western blot detection: annexin A5 (Sigma–Aldrich, A8604; mouse, 1:1000), a-tubulin

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(Sigma–Aldrich, T.9026; mouse; 1:5000), p16 (clone G175-1239; mouse; 1:250), p21 (Calbiochem, CALBO68; mouse; 1:1000), p53 (Calbiochem, Ab-6; mouse; 1:400), lamin A/C (Sigma–Aldrich, HPA006660; rabbit 1:100), NUP160 (a kind gift from Volker Cordes (Max-Planck Institute of Biophysical Chemistry, Go¨ttingen, Germany) (guinea pig; 1:200), Ki67 (abcam, ab15580; rabbit; 1:200), Rb (BD Pharmingen, 554136; mouse; 1:50), p38MAPK (Cell Signaling; rabbit; 9212; 1:50), pT180/Y182p38MAPK (Cell Signaling, rabbit; #9211S; 1:50) and HRP-coupled antirabbit or mouse IgG (Jackson Immuno Research Lab). 2.5. Immunofluorescence and confocal microscopy For indirect immunofluorescence, cells were fixed in 4% paraformaldehyde/ 1  PBS for 10 min followed by permeabilisation in 0.25% Triton-X 100/1  PBS for 3 min. Primary and secondary antibodies were incubated on cover slips for 1 h at room temperature. DNA was visualized with 40 -6-diamidine-2-phenyl indole (DAPI) containing mouting medium (Prolong1 Gold antifade reagent, Invitrogen). Confocal microscopy was carried out using a Zeiss LSM710 microscope with a 63/ 1.40 oil immersion objective as described previously (Hemmerich et al., 2008). The optical thickness per confocal section according to the microscope software is 0.317 mm. The following antibodies were used: annexin A5 (Sigma–Aldrich, A8604; mouse; 1:200), annexin A2 (Santa cruz, sc-47696; mouse; 1:100), annexin A4 (Sigma–Aldrich, HPA007393; rabbit; 1:20), lamin A/C (Sigma–Aldrich, HPA006660; rabbit; 1:100), Ki-67 (Abcam, ab15580; rabbit; 1:500), pATM (pS1981) (Abcam, ab81292; rabbit; 1:100), p16 (BD Pharmingen, clone G175-1239; mouse; 1:50), p21 (Santa cruz, sc-756; rabbit; 1:50) and gH2AX (Upstate clone JBW301; mouse; 1:400). Secondary fluorescence labeled antibodies were anti-mouse-Alexa488 (Molecular Probes); anti-rabbit-Cy2 (Dianova), anti-rabbit-Cy3 (Jackson Immuno Research Lab) and anti-mouse-Cy3 (Jackson Immuno Research Lab). 2.6. SA-b-Gal assay The SA-b-Gal assay was performed as described by Dimri et al. (1995). Cells were fixed in 4% paraformaldehyde and incubated 4–16 h at 37 8C in staining solution: 1 mg/ml X-Gal, 8 mM citric acid/sodium phosphate pH 6.0, 5 mM K3Fe(CN)6, 5 mM K4Fe(CN)6, 150 mM NaCl, and 2 mM MgCl2 (Sigma/Aldrich). Afterwards, the cells were mounted with DAPI-containing Prolong1 Gold antifade reagent (Invitrogen). 2.7. Tissue collection Skin biopsy donors (8 female, 6 male, and age range: 18–86 years; age groups: <30 years (n = 2), 30–70 years (n = 6) and >70 years (n = 6)) were recruited from the Department of Dermatology, Friedrich Schiller University, Jena, Germany, in conformity with the Declaration of Helsinki over a period of 1.5 years. The study was further approved by the Ethics commission of the Friedrich Schiller University Jena. All individuals were informed about the procedure and the risks and gave written, informed consent. Frozen tissue samples were mounted in cryo-medium at arrival and cutted into 12 mm thin slices. The sections were fixed in 4% paraformaldehyde for 15 min and permeabilized with 0.25% Triton-X 100/PBS for 10 min. Sections were blocked using 5% goat-serum for 1 h. Antibody-staining was carried out as described above. Quantitation of SA-ANX5 was done independently by two researchers (K.K. and P.H.), yielding the same results. When sections were analyzed by combined DAPI/DIC detection, no significant changes in cell density between the biopsies were observed. 2.8. Silencing RNA-mediated knockdown Experiments were performed using siRNA from Dharmacon (ABgene Ltd., Surrey, UK) against annexin A5 (L-011209-00-0005) and control RISC-free (D001220-01-05). DEPC-treated sterile water, RNase free tips and tubes (Starlab) were used for the work with siRNA. The lyophilized RNAi (5 nmol) was dissolved in 250 ml siRNA-Puffer to yield a concentration of 20 mM. Cells were seeded at least one day before siRNA transfection for young cells and two weeks for senescent cells. Transfection of siRNA was performed according to the manufacture’s instructions using lipofectamine RNAiMAX (Invitrogen, Germany). 2.9. Statistical evaluation P values were determined using the t-test or the Mann–Whitney test in Origin software.

foreskin fibroblast cell strain, BJ. In our hands, these cells had a maximum number of population doublings (PD) of 76, 60 and 67, respectively (Fig. 1A). The onset of replicative senescence of these cells was monitored by senescence-associated b-galactosidase (SA-b-Gal) activity staining (Fig. 1B and C). All three fibroblast types also developed typical molecular markers of cellular senescence, including upregulation of p16, p21 and p53 protein (Fig. 1D). As an alternative to replicative senescence we also employed stress-induced premature senescence (SIPS) (Toussaint et al., 2000). 5-Bromodeoxyuridine (BrdU) accelerates aging in D. melanogaster, mice and rats (Potapenko et al., 1982; Anisimov, 1994) and induces senescence in primary and immortal mammalian cells (Michishita et al., 1999). The senescence-inducing activity of BrdU can be potentiated by co-administration of ATrich DNA binding compounds such as distamycin A (DMA) (Suzuki et al., 2002). In MRC-5, WI-38 and BJ cells, senescence was efficiently induced by BrdU/DMA treatment within one week as judged by SA-b-Gal activity assays (Fig. 1E and F). At the concentrations used, the drugs had little or no effect on HeLa or U-2 OS cells, respectively (Fig. 1F). Similar to replicative senescent cells but on a much shorter time scale, BrdU/DMA also induced the expression of p16, p21, p53 and SAHF (Fig. 1G and H). By interfering with DNA replication and structure respectively, BrdU and DMA may induce senescence through DNA damage induction. In contrast to untreated cells which showed very little gH2AX reactivity (Fig. 1I), the number of cells containing several DNA damage foci increased to more than 80% after 12 days of BrdU/DMA treatment (Fig. 1J). These gH2AX foci also contained 53BP1, phosphorylated NBS1 and MDC1 (Fig. S1). These observations strongly suggest that BrdU/DMA treatment can induce cellular senescence in human fibroblasts by triggering a DNA damage response. To identify potentially new aging markers, protein lysates of senescent fibroblast nuclei were analyzed by SDS-PAGE. Most prominently, the amount of core histones was substantially reduced in B/D treated and replicative senescent cells, consistent with the previous demonstration that both, DNA damage and replicative aging reduces histone levels (O’Sullivan et al., 2010). This analysis reproducibly revealed a protein band at 35 kDa, the signal of which was reduced in senescent MRC-5 and Wi-38 cells (Fig. 2A and data not shown). This protein band represented annexin A5 as revealed by mass spectrometry. Database searches revealed annexin A5 as the best candidate with an estimated Zscore of 2.43. A Western blot of nuclear extracts of MRC-5 cells at different population doublings confirmed the decrease of annexin A5 protein levels in nuclear fractions during replicative senescence (Fig. 2B). Annexin A5 is present in the cytoplasm as well as in the nucleoplasm (Barwise and Walker, 1996). To examine the relative amounts of annexin A5 in these subcellular compartments, Western blots were performed after fractionation of MRC5 cells. This analysis showed that while a subfraction of annexin A5 co-purified with nuclear proteins, the majority was present in the cytoplasmic fraction (Fig. 2C). However, Western blots of whole cell fractions showed only little change in annexin A5 protein levels during replicative senescence except for a slight increase in MRC-5 cells (Fig. 2D). While the overall amount of annexin A5 is slightly increased, its presence in the nuclear fraction appears to decrease during replicative senescence of human diploid fibroblasts.

3. Results 3.2. Annexin A5 accumulates at the nuclear envelope of senescent cells 3.1. Annexin A5 expression during cellular senescence of primary human fibroblasts In the present study we used two primary human lung fibroblast cell strains (MRC-5 and WI-38), and the primary human

To further characterize cellular annexin A5 expression, confocal microscopy was performed. In young fibroblasts, annexin A5 immunofluorescence was mainly detected in the cytoplasm. A diffuse, but less intense staining was also observed in some nuclei

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Fig. 1. Replicative and BrdU/DMA-induced senescence. (A) MRC-5, WI-38 and BJ human primary fibroblasts were passaged until all cells ceased proliferation. Graphs show one representative PD measurement for each cell type. Independent measurements resulted in similar proliferation curves (data not shown). (B) SA-b-Gal assays were performed on MRC-5 cells at the indicated population doublings (PDs). Scale bar: 50 mm. (C) The number of SA-b-Gal positive cells was determined for MRC-5, WI-38 and BJ cells as a function of PDs. Graphs show mean values and standard deviations (n = 100 cells) from at least three independent measurements. (D) Expression of p16, p21 and p53 during replicative senescence was analyzed by Western blotting of whole cell extracts from MRC-5 cells at different PDs as indicated. Numbers on the left side indicate the position of protein molecular weight standards. (E) SIPS in MRC-5 cells treated with bromo-deoxy-uridine (BrdU) and distamycin A (DMA) for 6 days was monitored by SA-bGal staining. Scale bar: 50 mm. (F) Kinetics of BrdU/DMA induced SA-b-Gal staining in different cell lines. Graphs show mean values and standard deviations (n > 100 cells each) from at least three independent measurements. (G) Induction of SAHF during BrdU/DMA induced senescence in MRC-5 cells. Scale bar: 5 mm. (H) Expression of p16, p21 and p53 was analyzed by Western blotting of whole cell extracts of MRC-5 cells at different time points during BrdU/DMA induced senescence. Numbers on the left side indicate the position of molecular weight standards (I) Detection of gH2AX foci in MRC-5 fibroblasts before and 12 days after cultivation in BrdU/DMA containing medium. Scale bar: 5 mm. (J) Kinetics of g-H2AX-foci formation during BrdU/DMA induced senescence in MRC-5 cells. Graphs show mean values and standard deviations (n > 100 cells each) from at least three independent measurements.

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Fig. 2. Soluble levels of nuclear annexin A5 decrease during cellular senescence. (A) Young (PD 35), BrdU/DMA-treated (B/D) and replicative senescent (PD 75) MRC-5 cells were subjected to nucleo/cytoplasmic fractionation. Purified nuclear fractions were resolved by SDS-PAGE followed by Coomassie-Blue staining. A protein band at 35 kDa with decreased intensity in senescent cells was identified by mass spectrometry as ANX5 (asterisk). The relative signal intensity (RSI) of this band was determined using Metamorph software. RSI values are indicated below the respective bands. (B) Western blot analyses of nuclear fractions of MRC-5 cells at different population doublings using anti-ANX5 and anti-lamin A/C antibodies. RSI of antibody labeling was determined using Metamorph software. RSI values are indicated below the respective bands. (C) Western blot analysis of whole cell extracts (WCE), cytoplasmic (C) and nucleoplasmic (N) fraction of MRC-5 cells. Fractionation was monitored by detection of a-tubulin and nuclear pore protein (NUP) 160 as cytoplasmic and nuclear markers, respectively. (D) Cellular annexin A5 protein levels were determined during replicative senescence of MRC-5, WI-38, and BJ fibroblasts in Western blots of whole cell extracts of equivalent cell numbers (3  104 cells per lane) at different population doublings.

excluding nucleoli (Fig. 3A, left panels). In mid-aged cells the overall cellular distribution of annexin A5 did not change but the fluorescence intensity in the nucleus was increased compared to young cells (Fig. 3A, middle panels). At high population doublings, annexin A5 was found to accumulate at the nuclear periphery in many cells (Fig. 3A, right panels). High-resolution confocal microscopy at mid-nuclear sections of single cells confirmed the redistribution behavior of annexin A5 during cellular senescence of MRC-5 cells (Fig. 3B). Accumulation of annexin A5 at the nuclear periphery of senescent cells was also observed in senescent WI-38 and BJ fibroblasts (Fig. 3C). Strong colocalization with lamin A/C immunofluorescence staining confirmed that annexin A5 accumulates at the nuclear envelope (Fig. 3D). In order to quantitatively assess annexin A5 relocalization during cellular senescence, three different phenotypes were defined. The first phenotype (P1) included cells in which the intensity of nuclear anti-annexin A5 immunostaining was very low (Fig. 3E–G, blue bars). The second phenotype (P2) consisted of cells in which nuclear anti-annexin A5 fluorescence was substantially stronger than in the cytoplasm without accumulation at the nuclear envelope (Fig. 3E–G, green bars). Cells of the third phenotype (P3) showed a strong annexin A5 staining of the nuclear envelope (Fig. 3E–G, red bars). At low population doublings the majority of MRC-5 cells showed the P1 phenotype. In mid-aged MRC-5 cultures, P1 cells decreased and cells displaying P2 or P3 increased in number. At late passage, more than 90% of MRC-5 cells exhibited P3 (Fig. 3E). Similar dynamics of annexin A5 redistribution were also observed during replicative senescence of WI-38 and BJ fibroblasts (Fig. 3F and G). The P3 phenotype was observed in more than 90% of senescent MRC-5 and WI-38 cells, and 70% of senescent BJ fibroblasts. Accumulation of annexin A5 at the nuclear envelope was also observed during BrdU/DMA-induced senescence (Fig. S2A and B), showing that this phenotype occurs in both, replicative and stress-induced senescence. A line scan of high

resolution confocal images confirmed that the fluorescence signals of ANX5 and lamin A/C fully colocalize in senescent fibroblasts (Fig. S2C). The redistribution of annexin A5 to the nuclear envelope during senescence appears contradictory to our Western blot data which showed decreasing amounts of annexin A5 in nuclear fractions of senescent cells (Fig. 2B). A reasonable explanation for this result is an alteration of the biochemical solubility of annexin A5 in aging cells. Alternatively, or in addition, annexin A5 may alter its subcellular localization during fractionation. To test whether other annexin family members also redistribute to the nuclear envelope, we determined the cellular localization of annexins A2 and A4 in senescent MRC-5, WI-38 and BJ fibroblasts. Confocal microscopy revealed that ANX2 and ANX4 do not accumulate in the nuclear periphery of senescent fibroblasts although they were clearly present in nuclei (Fig. S3). Thus, accumulation at the nuclear envelope of senescent cells is not a general phenomenon of annexin family members. Taken together, these results identify accumulation of annexin A5 in the nucleus and at the nuclear envelope as a specific feature of cellular senescent human fibroblasts. We propose the name SA-ANX5 for this novel biomarker of aging. 3.3. Cross-validation of SA-ANX5 with other biomarkers of senescence We then compared the expression of the SA-ANX5 phenotype during replicative senescence of human fibroblasts with wellestablished biomarkers of cellular senescence on the single cell level by confocal microscopy (Fig. 4A). As expected, the number of p21, p16 and SAHF positive MRC-5 cells increased during serial passaging (Fig. 4B). Quantitative comparison of these markers and SA-b-Gal with respect to SA-ANX5 revealed that this phenotype is a relatively late event during replicative senescence in MRC-5 cells (Fig. 4B). The onset of the SA-ANX5 phenotype was observed at 62 PD, while p16,

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Fig. 3. Annexin A5 accumulates at the nuclear envelope during replicative senescence. (A) MRC-5 and WI-38 cells grown on cover slips at different PDs were processed for immunofluorescence staining to detect annexin A5 (red) and DNA (green). All images represent confocal recordings of the mid-nucleus section. Panels on the right side show ant-annexin A5 fluorescence in monochrome. Scale bar; 20 mm. (B) Nuclear regions of MRC-5 cells at indicated PDs were analyzed at higher resolution by confocal microscopy. Annexin A5 staining is bright and intense in the cytoplasm in low passage fibroblasts, but weak staining was also detected in nuclei (PD 28). In mid-aged cells (PD 62), annexin A5 staining increased in the nucleus. In replicative sensescent cells, annexin A5 was found to accumulate strongly at the nuclear envelope (PD 70). Most of these cells also contained senescence associated heterochromatin foci (SAHF, green). Scale bar: 5 mm. (C) Annexin A5 accumulation at the nuclear envelope (red) was also observed in replicative senescent WI-38 and BJ fibroblasts. Scale bars: 5 mm. (D) Colocalization of annexin A5 with the nuclear envelope (red) was confirmed by co-immunostaining using an anti-lamin A/C antibody (green). Scale bar: 5 mm. (E–G) Quantification of alterations of annexin A5 localization in different human primary fibroblast cell lines during replicative senescence. P1 phenotype (blue bars): cells with no or weak nuclear staining; P2 phenotype (green bars): cells with strong nuclear staining but no or little nuclear envelope decoration; P3 phenotype (red bars): cells with strong annexin A5 accumulation at the nuclear envelope. Bars show mean values and standard deviations (n > 100 cells at each PD per measurement) from at least three independent measurements.

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Fig. 4. Comparison of SA-ANX5 with to other markers of cellular senescence. (A) MRC-5 fibroblasts were fixed at different time points during serial passaging and processed to detect p21 (red), p16 (red), and DNA (green) by confocal microscopy. Scale bar: 50 mm in top and middle images, 5 mm in bottom images. (B) Quantification at single-cell level of experiments shown in (A) along with SA-bGal and SA-ANX5. Data points show mean values (without standard deviations to avoid confusion) (n > 100 cells at each PD per measurement) from at least three independent measurements. (C–E) Pairwise incidence plots for assays as shown in (A) for MRC-5, WI-38 and BJ cells. The incidence plots compared the occurrence of one marker with another marker in % of cells. The increase along the x and y achses are equivalent to PDs. The incidence of SA-ANX5 was plotted against the occurrence of known cellular senescence markers for MRC-5, WI-38 and BJ fibroblasts as indicated. Data points show mean values and standard deviations (n > 100 cells per measurement) from at least three independent measurements.

p21 and SA-b-Gal started to increase already at PD 50 (Fig. 4B). SAANX5 was as efficient as SA-b-Gal and p21 to detect senescent cells during late passage, while p16 expression and SAHF formation occurred in only 60% of senescent MRC-5 cells (Fig. 4C). In contrast to MRC-5 cells, kinetics of SA-ANX5, SA-b-Gal and p21 expression were almost identical in aging WI-38 fibroblasts (Fig. 4D). SA-ANX5 was as efficient as SA-b-Gal in detecting senescent WI-38 cells, while only 80% of late passage WI-38 fibroblasts were positive for p21 (Fig. 4D). Again, 40% of late passage WI-38 cells were negative for p16 or SAHF (Fig. 4D). One hundred percent of replicative senescent BJ cells expressed p21 and were SA-b-Gal positive, while 30% of these cells did not display the SA-ANX5 phenotype (Fig. 4E). Only 40% of senescent BJ cells were positive for p16 and none of them

developed SAHF (Fig. 4E). These observations indicate that the senescence markers studied here develop in a cell type specific manner and with cell type-specific kinetics. Compared to SA-b-Gal, p21 and SA-ANX5, the p16 and SAHF markers appear to be less suitable diagnostic tools to safely identify replicative senescent fibroblasts in the single cell assay. Finally, with the exception of BJ cells, SA-ANX5 is as efficient in detecting replicative senescent fibroblasts as SA-b-Gal or p21. 3.4. SA-ANX5: a potential marker for senescent epidermal cells We next asked whether the SA-ANX5 phenotype also occurs in vivo. Skin tissue sections from healthy human individuals of

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different ages were immuno-stained against annexin A5 (Fig. 5A). The dermis showed a strong cellular annexin A5 immunoreactivity in the majority of cells independent of age (Fig. 5A). In contrast, the epidermis of young individuals showed a much weaker staining, while a subset of cells displayed strong annexin A5 immunoreactivity in tissue samples from aged donors (Fig. 5A). In dermal cells of all ages, annexin A5 staining was detected in the cytoplasm and diffusely in the nucleus (Fig. 5A, enlargements). Due to the high expression level of annexin A5 in almost all cells of the dermis

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it was not possible to detect accumulation of annexin A5 in the periphery of the cell nuclei. Therefore the frequency of the SAANX5 expressing cells could not be reliably assessed. However, cells clearly displaying the SA-ANX5 phenotype were negative for Ki-67 (Fig. 5B) and positive for pATM foci (Fig. 5C), suggesting that senescent dermal cells showing SA-ANX5 exist in the human dermis. Although the number of annexin A5 expressing cells was already high in the dermis of all donors, a donor age-dependent increase was observed for dermal fibroblasts (Fig. 5C).

Fig. 5. Expression of annexin A5 in human dermis. (A) Representative sections of skin biopsies from three donors of different age (18, 66, and 77 years in upper, middle, and bottom panel, respectively) were immunostained against anti-annexin A5 antibody (red) and counter-stained with DAPI (green). Sections were analyzed by confocal microscopy in the epidermis and the demis (dotted blue line). Annexin A5 staining is also shown in monochrome (middle panels). Panels on the right side show enlargements of annexin A5 staining in the dermis. Scale bar: 5 mm. (B) Confocal images of fibroblasts in the dermis of skin biopsy sections of different age immuno-stained with antibodies against annexin A5 (green) and Ki67 or pATM, as indicated (red). DNA was counterstained with DAPI (blue). Scale bar: 5 mm. (C) Quantification of the amount of annexin A5 expressing cells in human skin tissue from donors of different age groups. Bars represent mean values  SD (n = 300 cells) for each age group. P values were determined using ttest and Mann–Whitney test in Originlab software. n.s., not significant.

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Fig. 6. SA-ANX5 expression in the epidermis correlates with markers of cellular senescence. (A) A skin tissue section from an old donor was immuno-stained against annexin A5 (red), and the epidermis was analyzed by confocal microscopy. DNA was counter-stained with DAPI (green). Scale bar: 5 mm. Cells with different levels of annexin A5 expression were detected in the epidermis. (B) The same staining as in (A) was applied on epidermal sections of different age. (C) The number of epidermal cells with high expression of annexin A5 was quantified and plotted as a function of donor age groups. Bars represent mean values  SD (n = 300 cells) for each age group. P values were determined using t-test and Mann–Whitney test in Originlab software. (D) Frequently, annexin A5 accumulated at the nuclear envelope (NE) as well as the cell membrane (CM) of epidermal cells in skin sections of old individuals. Scale bar: 5 mm. (E) The epidermis of a skin biopsy section from an old individual was immunostained against lamin A/C (green) and annexin A5 (red) and analyzed by confocal microscopy. Nuclei were counter-stained with DAPI (blue). Accumulation of annexin A5 at the nuclear envelope of the epidermal cell is evident form the colocalization with lamin A/C staining. Scale bar: 5 mm (F) Skin tissue sections from donors of different ages were co-immunostained to detect annexin A5 (green) and Ki67 (red). DNA was counterstained with DAPI (blue). Confocal images were taken in the epidermis at mid-nuclear optical sections. Scale bar: 5 mm (G) Stained samples were quantitated at the single-cell level with respect to cells positive for the SA-ANX5 phenotype (green), Ki67 (red), or both (black) as a function of skin donor age. Bars represent mean values  SD (n = 300 cells) for each sample. Statistical significance is shown for data at 18 years compared to the other ages. (H) Skin tissue sections from donors of different ages

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The level of annexin A5 expression in epidermal cells of old samples varied between low and high (Fig. 6A). Single cell analysis revealed a clear relationship between strong annexin A5 expression in the epidermis and donor age (Fig. 6B). The fraction of strongly stained cells increased from <2% in young individuals (age <30 years) to 12% in skin of elderly (age >70 years) (Fig. 6C). A subset of epidermal cells in aged skin tissue showed strong accumulation of annexin A5 at the nuclear periphery (Fig. 6D) which colocalized with the lamin A/C signal (Fig. 6E), again resembling the SA-ANX5 phenotype. To further characterize SA-ANX5-positive epidermal cells, double-labeling experiments against Ki-67 and pATM were performed (Fig. 6F–I). We found more than 20% Ki-67-positive epidermal cells in young skin. This number dropped below 10% in samples from old donors (Fig. 6B). The number of SA-ANX5 positive epidermal cells increased with age and these cells never expressed Ki-67, indicating that these cells are post-mitotic (Fig. 6G, black bars). The number of pATM-positive (>two large foci) epidermal cells increased with age (Fig. 6H, black bars). Although the number of pATM-positive cells was usually higher than the number of SAANX5-positive cells, all epidermal cells displaying the SA-ANX5 phenotype also contained elevated numbers of pATM foci indicating an increased DNA damage response in these cells. Taken together, the absence of Ki-67 and the presence of increased numbers of pATM foci displaying SA-ANX5 suggest that this phenotype may serve as a novel marker for aged skin cells in vivo. 3.5. Annexin A5 depletion accelerates cellular senescence through the p38MAPK pathway The SA-ANX5 phenotype observed during in vitro and in vivo cellular aging raised the question if annexin A5 may be involved in senescence pathways. To address this question siRNA-mediated knockdown of annexin A5 was performed. Annexin A5 expression was efficiently repressed in MRC-5 cells after six to ten days of siRNA treatment (Fig. 7A). Annexin A5 knockdown induced p21 expression in almost all cells as judged by immunofluorescence analysis (Fig. 7B). The individual fluorescence intensity varied greatly with many cells showing low but clearly detectable antip21 immunoreactivity (Fig. 7B, yellow arrowheads). Furthermore, annexin A5 depleted fibroblast cultures showed an elevated number of SA-b-Gal positive cells in comparison to control cells (Fig. 7C). We then analyzed the impact of annexin A5 downregulation during BrdU/DMA-induced senescence. This approach revealed that cells with reduced annexin A5 levels developed the senescence marker SA-b-Gal earlier than control cells (Fig. 7D). In the absence of BrdU/DMA, annexin A5 depletion induced an increase of p21-positive cells from 50% to more than 80% (Fig. 7D). In contrast, the number of p16 positive cells as determined by immunofluorescence was unaffected (Fig. 7D). Induction of SAHF during BrdU/DMA-induced senescence was not altered by annexin A5 depletion (Fig. 7D). Interestingly, BrdU/ DMA-induced senescence lead to an upregulation of annexin A5 protein level in whole cell lysates (Fig. 7E), which was not observed at this magnitude during replicative senescence (Fig. 2D). This observation suggests that increased annexin A5 expression is a specific DNA damage response. Consistent with a senescence-associated block of proliferation, Western blots of MRC-5 fibroblasts showed that depletion of annexin A5 leads to a downregulation of the proliferation marker Ki-67 (Fig. 7E). As expected, BrdU/DMA treatment induced the

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expression of p16, p21 and p53 (Fig. 7E). Annexin A5 depletion resulted in slightly increased levels of p21 in Western blots (Fig. 7E) consistent with the immunofluorescence results. In contrast, while the number of p16 positive cells was not further increased as judged by immunofluorescence (Fig. 7D), its protein level increased in Western blots of whole cell populations (Fig. 7E). These observations underline the conclusion that both, single cell analysis by immunofluorescence and Western blot analysis in parallel are instrumental to reliably monitor expression of senescence markers (Herbig et al., 2004). Consistent with an upregulation of p16, we also observed dephosphorylation of pRB in BrdU/DMA treated fibroblasts (Fig. 7E). In order to obtain first insights into the mechanism by which annexin A5 depletion induces an acceleration of stress-induced senescence we analyzed senescence-associated signaling pathways. JAK/STAT signaling is triggered during DNA damage induced cellular senescence, including BrdU and DMA induced genotoxicity (Hubackova et al., 2010). As expected, STAT1 expression was induced upon BrdU/DMA treatment of MRC-5 fibroblasts but depletion of annexin A5 did not further upregulate STAT1 levels or its phosphorylation (Fig. 8A). By contrast, activation of p38MAPK by BrdU/DMA was markedly enhanced in MRC-5 cells lacking annexin A5 (Fig. 8B). p38MAPK is activated during the senescence response to genotoxic stress (Freund et al., 2011) and is part of a signaling cascade that induces reactive oxygen species (ROS) production. The elevated ROS levels are believed to maintain a sustained DNA damage response as part of a positive feedback loop with p21 (Passos et al., 2010). Consistent with this hypothesis, we found increased levels of gH2AX and pATM in annexin A5 depleted, premature senescent MRC-5 fibroblasts (Fig. 8B). Confirming observations were made during BrdU/DMA induced senescence of BJ fibroblasts (Fig. 8C). The p38MAPK-driven ROS/DDR feedback loop is still active in late passage human fibroblasts (Passos et al., 2010). When annexin A5 was depleted by siRNA in MRC-5 fibroblasts at PD 63, we still observed enhanced phosphorylation of p38MAPK concomitant with an increase in phosphorylation of H2AX and ATM with and without BrdU/DMA treatment (Fig. 8D). Collectively, these observations suggest that unscheduled loss of annexin A5 triggers and accelerates cellular senescence by activation of the p38MAPK senescence pathway and an increased DNA damage response.

4. Discussion In direct comparison to established replicative senescence markers, SA-ANX5 was as efficient as SA-b-Gal staining and p21 immunoreactivity in cultured fibroblasts (Fig. 4). SA-b-Gal activity is the most widely used biomarker in studies of cellular senescence in vitro and in vivo (Debacq-Chainiaux et al., 2009). Because the SAANX5 phenotype is similarly efficient, it may be used in the future in combination with other senescence markers, in particular DNA damage foci, p21 or p16 in order to more reliably identify senescent cells. The advantage of this approach is, in contrast to SA-b-Gal, detection of senescent cells in tissue sections using standard immuno-histochemistry techniques, thus avoiding enzymatic assays. Consistent with this idea, we found SA-ANX5 exclusively in pATM foci-positive/Ki-67-negative epidermal cells of aged human skin. SA-ANX5 may therefore support the latter marker combination which already provides a reliable assessment of the frequency of senescent cell in vitro and in vivo (Lawless et al., 2010). SA-ANX5 may also complement the SA-b-Gal assay in some

were co-immunostained to detect annexin A5 (green) and pATM (red). DNA was counterstained with DAPI (blue). Confocal images were taken in the epidermis at mid-nuclear optical sections. Scale bar: 5 mm (I) Stained samples were quantitated at the single-cell level with respect to cells positive for the SA-ANX5 phenotype (green), pATM (red), or both (black). Bars represent mean values  SD (n = 300 cells) for each sample. Statistical significance is shown for data at 18 years compared to the other ages.

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Fig. 7. Annexin A5 depletion induces and accelerates cellular senescence. (A) Down-regulatin of annexin A5 in MRC-5 fibroblasts. Whole cell extracts of MRC-5 cells cultured without siRNA (ctrl, 6 days), control siRNA (siCON, 6 days) or siRNA against annexin A5 (siANX5, 6 or 10 days) were processed for immunodetection by Western blots of annexin A5 (ANX5) and a-tubulin (a-tub) protein levels. Numbers on the left side indicate the molecular weight of protein size markers run in the same gel. (B) Confocal microscopy analysis of control (siCON) and annexin A5 siRNA-treated (siANX5, 6 days) early passage MRC-5 fibroblasts. Cells were fixed and immuno-stained to detect p21 (green) and annexin A5 (red). DNA was counterstained with DAPI (blue). Cells with very low but clearly detectable anti-p21 immunoreactivity are indicated by a yellow arrowhead. Scale bar: 20 mm. (C) SA-b-Gal assay of early passage MRC-5 cells cultured for 6 days in the presence of control siRNA or annexin A5 siRNA. DNA was counterstained with DAPI (blue). Scale bar: 50 mm. (D) Annexin A5 depletion accelerates drug-induced senescence. Early passage MRC-5 fibroblasts were left untreated (black graphs) or treated with control siRNA (blue graphs) or annexin A5 siRNA (red graphs). After 6 days the culture medium was supplemented with BrdU/DMA for another 12 days to induce cellular senescence. Cover slips taken at the indicated time points after BrdU/DMA addition were processed to detect SA-bGal, p21, p16 or SAHF. Data points show mean values  SD (n > 100 cells) for each measurement. (E) Western blot detection of indicated proteins in MRC-5 fibroblasts treated or not with annexin A5 specific siRNAs for 6 days followed by incubation for another 6 days in the absence () or presence (+) of BrdU/DMA. Numbers on the left side indicate the molecular weight of protein size markers run in the same gels.

settings, for example when analyzing archival tissue section samples. We also described limitations of the new senescence marker. SA-ANX5 was observed in more than 90% of senescent MRC-5 and Wi-38 cell, but only in 67% of senescent BJ fibroblasts, indicating cell type-specific penetrance of this new marker. Another limitation was observed in the dermis where high expression levels of annexin A5 prevented unambiguous assessment of the frequency of SA-ANX5-positive fibroblasts. Nevertheless, dermal fibroblasts displaying a clear SA-ANX5 phenotype were always Ki-67 negative and pATM foci positive, indicating a correlation between SA-ANX5 and potentially senescent fibroblasts in vivo. Our observation that BrdU/DMA treatment also induces SA-ANX5 indicates that this phenotype may represent an additional member of DNA damage induced senescence markers

with potential applications in vitro and in vivo (Jiang et al., 2008). It remains to be tested if detection of annexin A5 upregulation and SA-ANX5 will also assist in other tissues and in cancer staging during the study of OIS (Collado et al., 2005). Annexins are a family of 13 ubiquitous phospholipid and Ca2+binding proteins which can assemble on negatively charged membrane surfaces. Annexins are involved in the organization of membranes and membrane-cytoskeleton interactions, as well as in the regulation of exo/endocytosis and ion flux (Gerke et al., 2005; Lemmon, 2008). A Ca2+ binding-induced conformational change leads to annexin oligomerization, high affinity binding to phospholipids, and reversible attachment to vesicle, nuclear and plasma membranes (Gerke and Moss, 2002). Mice lacking annexin A5 are viable and fertile (Brachvogel et al., 2003) but endogenous

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Fig. 8. Annexin A5 knockdown activates p38MAPK and DDR during stress-induced senescence. (A and B) Western blot detection of indicated proteins in MRC-5 fibroblasts at PD 48 treated or not with annexin A5 specific siRNAs for 6 days followed by incubation for another 6 days in the absence () or presence (+) of BrdU/DMA. (C) Western blot detection of indicated proteins in BJ fibroblasts at PD 46 treated or not with annexin A5 specific siRNAs for 6 days followed by incubation for another 6 days in the absence () or presence (+) of BrdU/DMA. Numbers on the left side indicate the molecular weight of protein size markers run in the same gels. (D) Western blot detection of indicated proteins in MRC-5 fibroblasts at PD 62 treated or not with annexin A5 specific siRNAs for 6 days followed by incubation for another 6 days in the absence () or presence (+) of BrdU/DMA. Numbers on the left side of each panel indicate the molecular weight of protein size markers run in the same gels.

annexin A5 influences phagocytosis of necrotic cells and modulates the immune response towards allogeneic cells (Frey et al., 2009). DT40 chicken cells lacking annexin A5 are resistant to Ca2+dependent apoptosis indicating an involvement in cell fate decision after stress (Hawkins et al., 2002). RNAi mediated knockdown experiments have revealed that invasion capacity, a main feature of tumors, is at least partially regulated by annexin A5 in oral carcinoma (Wehder et al., 2009). These observations point to a prominent role of annexin A5 in cancer which might be related to its potential function in stress-induced cellular senescence as indicated in the present report. Some annexin family members, including annexin A5, have been found to shuttle between the cytoplasm and the nucleus. During quiescence, annexin A5 is absent from the nucleus and does not accumulate at the nuclear envelope of primary human fibroblasts (Barwise and Walker, 1996), while we observed in the present report its nuclear accumulation during cellular senescence. Colocalization analyses with lamin A/C showed full overlap between annexin A5 and the nuclear lamina (Fig. S2C). However, electron or superresolution light microscopy is required to assess the precise localization of annexin A5 at the nuclear envelope of senescent cells. Treatment of human foreskin cells with Ca2+ ionophores to raise intracellular Ca2+ concentration results in relocation of intranuclear annexin A5 to the nuclear envelope (Raynal et al., 1996), phenotypically similar to the SA-ANX5 phenotype in senescent cells. Because young and senescent human fibroblasts have similar intracellular Ca2+-levels and exhibit similar changes in cytosolic Ca2+-fluxes following growth factor stimulation (Takahashi et al., 1992; Brooks-Frederich et al., 1993, and our own data), the SA-ANX5 phenotype is not induced by shifts in Ca2+ concentrations but rather represents a senescence-specific alteration. In vitro, annexin A5 as well as other annexins form voltage-gated channels which are selective for divalent cations (Rojas et al., 1990). Therefore, SAANX5 may impact on ion homeostasis at the nuclear envelope. Annexin A5 is also able to form 2-dimensional scaffolding platforms on membranes (Pigault et al., 1994). At the nuclear envelope,

such assemblies of annexin A5 might alter the physico-chemical properties of the membrane, such as stabilization of certain membrane structures and/or in changing membrane curvature and therefore nuclear shape. Annexin A5 is also present in a secreted form (Reutelingsperger et al., 1985) where it plays a role in apoptosis, phagocytosis and formation of plasma membranederived microparticles (van Genderen et al., 2008). Annexin A5 may therefore not only function as a signaling molecule within senescent cells but also extracellularly where it could contribute to the inflammatory effects of SASP/SMS. In this respect it is noteworthy that therapeutic annexin A5 administration attenuates vascular inflammation and remodeling which improves endothelial function in mice (Ewing et al., 2011). It will be interesting to test whether annexin A5 is a member of the senescence-associated secretome (SASP/SMS) in vitro and in vivo. We observed that depletion of annexin A5 triggers and accelerates cellular senescence in human primary fibroblasts. This effect correlated with upregulation of p21, p16 and p53 and dephosphorylation of pRB. These data strongly suggest that an acute stress imposed by unscheduled annexin A5 depletion in proliferating fibroblasts can trigger an p21- and/or p16-induced cell cycle arrest. Further analysis of potentially affected senescence pathways showed that annexin A5 knock-down also induces enhanced activation (phosphorylation) of p38MAPK and the DDR markers gH2AX and pATM. Recently, a signaling cascade involving p21, p38MAPK and transforming growth factor (TGF) b has been identified which links telomere-dependent and -independent DDR and ROS production in primary fibroblasts. It is believed that this p38MAPK-dependent pathway provides a senescence-promoting positive feedback loop (Passos et al., 2010). Our observation that lack of annexin A5 induces p38MAPK activation suggests a coregulation of this feedback loop through annexin A5. A p16-driven feedback loop with a similar outcome has recently been described: the p16/Rb pathway is linked to mitogenic signaling that increases ROS formation. In human fibroblasts such elevated ROS levels activate PKCd which promotes further generation of ROS.

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Sustained activation of ROS–PKCd signaling irreversibly blocks the cell cycle in fibroblasts (Takahashi et al., 2006). Since PKCd activation and signaling is regulated by annexin A5 through direct interaction (Kheifets et al., 2006) it is possible that annexin A5 is also involved in the p16–ROS–PKCd feedback loop. Cellular expression levels of annexin A5 in cultured human fibroblasts were slightly upregulated during replicative senescence and substantially upregulated in drug-induced senescence. Consistent with these observations, we found that annexin A5 expression increases with age in the dermis and epidermis of human skin. Previously, the protein level of annexin A5 was also found to be increased during replicative senescence of human umbilical vein endothelial cells (HUVECs) (Eman et al., 2006). Furthermore, metadata analysis of transcript profiles revealed that the annexin A5 gene is transcriptionally upregulated in a variety of aged mammalian tissues (de Magalha˜es et al., 2009). Collectively, these observations support the idea that annexin A5 may be part of a common signature of aging (de Magalha˜es et al., 2009). Upregulation of annexin A5 during cell culture aging appears counter-intuitive given the fact that its down-regulation in fibroblasts induced accelerated senescence. However, we think that these apparently inconsistent observations must be interpreted individually. Unscheduled down regulation of annexin A5 in cultured fibroblasts is an experimental intervention in vitro that is physiologically unrelated to the in vivo situation of fibroblasts in the skin. Most significantly, in cell culture most fibroblasts proliferate while in the dermis most fibroblasts are cell cycle arrested (quiescence). Therefore, not all in vitro data obtained on fibroblasts, particularly those related to cell proliferation, must necessarily recapitulate fibroblast behavior in vivo, and vice versa. Senescence induction after knock-down suggests that annexin A5 may exert an antisenescence activity when upregulated. Such mechanisms have not been described yet for senescent cells in tissues. Recent in vitro evidence suggests the existence of signaling feedback loops involving the p21 or p16 pathways with the potential to ameliorate the phenotypic impact of senescent cells onto their microenvironment (Takahashi et al., 2006; Passos et al., 2010). It is possible that cellular mechanisms exist in vivo which are capable of counter-acting the potentially deleterious effects of senescent cells in tissues, for example the inflammation- and tumor-promoting activity of SASP (Coppe´ et al., 2010). Although highly speculative, we suggest that upregulation of annexin A5 in aging tissues may have an anti-senescence effect in tissues by interfering with senescence promoting mechanisms, such as the p21-ROS/DDR-p38MAPK (Passos et al., 2010). Whether increased expression of annexin A5 delays cellular senescence or counter-acts some of its harmful phenotypes remains to be determined experimentally. 5. Conclusion Unambiguous detection of senescent cells is an important issue in the research of in vitro and in vivo cellular aging. New biomarkers of cellular senescence will improve diagnosis and analysis of their function may improve our understanding of this important pathway. In this study we have described that annexin A5 accumulates at the nuclear periphery of aged cells. This new phenotype (termed SA-ANX5) is as efficient as or superior to previously used biomarkers of cellular aging and should be useful in future applications for safe detection of senescent cells. Our observation that loss of annexin A5 accelerates in vitro aging of human fibroblasts through triggering the p38MAP kinase pathway suggests that annexin A5 is a functional mediator connecting the cellular stress response to the senescence pathway.

Contributors K.K., S.D. and P.H. designed experiments. J.N. provided access to skin tissue samples and supervised ethical guidelines. C.M. and U.M. performed and analyzed mass spectrometry. K.K. performed all experiments and analyzed all data, except mass spectrometry. K.K. and P.H. evaluated all data and wrote the manuscript. All authors read and approved the final manuscript. Acknowledgments We greatly appreciate the help by Maik Baldauf for technical assistance during histology procedures and Dr. med. Mirjana Ziemer for collection of skin tissue samples. We thank Volker Cordes for providing anti-nuclear envelope protein antibodies. We acknowledge JenAge (http://www.jenage.de/) funding by the German Ministry for Education and Research (Bundesministerium fu¨r Bildung und Forschung – BMBF; support code: 0315581). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.mad.2012.06.003. References Anisimov, V.N., 1994. The sole DNA damage induced by bromodeoxyuridine is sufficient for initiation of both aging and carcinogenesis in vivo. Annals of the New York Academy of Sciences 719, 494–501. Aubert, G., Lansdorp, P.M., 2008. Telomeres and aging. Physiological Reviews 88, 557–579. Baker, D.J., Wijshake, T., Tchkonia, T., LeBrasseur, N.K., Childs, B.G., van de Sluis, B., Kirkland, J.L., van Deursen, J.M., 2011. Clearance of p16Ink4a-positive senescent cells delays ageing-associated disorders. Nature 479, 232–236. Barwise, J.L., Walker, J.H., 1996. Subcellular localization of annexin V in human foreskin fibroblasts: nuclear localization depends on growth state. FEBS Letters 394, 213–216. Ben-Porath, I., Weinberg, R.A., 2005. The signals and pathways activating cellular senescence. International Journal of Biochemistry and Cell Biology 37, 961–976. Blagosklonny, M.V., 2011. Cell cycle arrest is not senescence. Aging (Albany, NY) 3, 94–101. Brachvogel, B., Dikschas, J., Moch, H., Welzel, H., von der Mark, K., Hofmann, C., Poschl, E., 2003. Annexin A5 is not essential for skeletal development. Molecular and Cellular Biology 23, 2907–2913. Brooks-Frederich, K.M., Cianciarulo, F.L., Rittling, S.R., Cristofalo, V.J., 1993. Cell cycle-dependent regulation of Ca2+ in young and senescent WI-38 cells. Experimental Cell Research 205, 412–415. Campisi, J., d’Adda di Fagagna, F., 2007. Cellular senescence: when bad things happen to good cells. Nature Reviews Molecular Cell Biology 8, 729–740. Cristofalo, V.J., Pignolo, R.J., 1996. Molecular markers of senescence in fibroblastlike cultures. Experimental Gerontology 31, 111–123. Collado, M., Gil, J., Efeyan, A., Guerra, C., Schuhmacher, A.J., Barradas, M., Bengurı´a, A., Zaballos, A., Flores, J.M., Barbacid, M., Beach, D., Serrano, M., 2005. Tumour biology: senescence in premalignant tumours. Nature 436, 642. Collado, M., Serrano, M., 2010. Senescence in tumours: evidence from mice and humans. Nature Reviews Cancer 10, 51–57. Coppe´, J.P., Desprez, P.Y., Krtolica, A., Campisi, J., 2010. The senescence-associated secretory phenotype: the dark side of tumor suppression. Annual Review of Pathology 5, 99–118. d’Adda di Fagagna, F., Reaper, P.M., Clay-Farrace, L., Fiegler, H., Carr, P., Von Zglinicki, T., Saretzki, G., Carter, N.P., Jackson, S.P., 2003. A DNA damage checkpoint response in telomere-initiated senescence. Nature 426, 194–198. d’Adda di Fagagna, F., 2008. Living on a break: cellular senescence as a DNA-damage response. Nature Reviews Cancer 8, 512–522. de Magalha˜es, J.P., Curado, J., Church, G.M., 2009. Meta-analysis of age-related gene expression profiles identifies common signatures of aging. Bioinformatics 25, 875–881. Debacq-Chainiaux, F., Erusalimsky, J.D., Campisi, J., Toussaint, O., 2009. Protocols to detect senescence-associated beta-galactosidase (SA-betagal) activity, a biomarker of senescent cells in culture and in vivo. Nature Protocols 4, 1798–1806. Di Leonardo, A., Linke, S.P., Clarkin, K., Wahl, G.M., 1994. DNA damage triggers a prolonged p53-dependent G1 arrest and long-term induction of Cip1 in normal human fibroblasts. Genes and Development 8, 2540–2551. Dimri, G.P., Lee, X., Basile, G., Acosta, M., Scott, G., Roskelley, C., Medrano, E.E., Linskens, M., Rubelj, I., Pereira-Smith, O., Peacocke, M., Campisi, J., 1995. A novel biomarker identifies senescent human cells in culture and in aging skin in vivo.

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