Fluorescence-Based Detection and Quantification of Features of Cellular Senescence

CHAPTER 7

Fluorescence-Based Detection and Quantification of Features of Cellular Senescence Sohee Cho and Eun Seong Hwang Department of Life Science, University of Seoul, Seoul, Republic of Korea

Abstract I. Introduction II. Features Associated with Loss of Reproductive Cell Capability A. Shortened Telomeres in Replicative Senescence B. Telomere Dysfunction-Induced Foci (TIF) in Replicative Senescence C. Loss of the Doubling Capacity of Cells III. Cellular Hypertrophy IV. Changes Associated with Lysosomes A. Lipofuscin Accumulation B. Increased Lysosome Content C. Senescence-Associated b-Galactosidase (SA b-Gal) Activity V. Changes Associated with Mitochondria A. Increased Mitochondrial Mass B. Altered Structural Dynamics of Mitochondria C. Decreased Membrane Potential D. Decreased Autophagy VI. Changes in the Level of Reactive Oxygen Species (ROS) A. Changes in Level of Superoxide B. Changes in Level of Hydroxyl Radicals VII. Changes Associated with Nucleus and Chromosomes VIII. Use of Flow Cytometry for Analysis of Cellular Senescence IX. Conclusion Acknowledgments References

METHODS IN CELL BIOLOGY, VOL 103 Copyright 2011, Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-385493-3.00007-3

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Abstract Cellular senescence is a spontaneous organismal defense mechanism against tumor progression which is raised upon the activation of oncoproteins or other cellular environmental stresses that must be circumvented for tumorigenesis to occur. It involves growth-arrest state of normal cells after a number of active divisions. There are multiple experimental routes that can drive cells into a state of senescence. Normal somatic cells and cancer cells enter a state of senescence upon overexpression of oncogenic Ras or Raf protein or by imposing certain kinds of stress such as cellular tumor suppressor function. Both flow cytometry and confocal imaging analysis techniques are very useful in quantitative analysis of cellular senescence phenomenon. They allow quantitative estimates of multiple different phenotypes expressed in multiple cell populations simultaneously. Here we review the various types of fluorescence methodologies including confocal imaging and flow cytometry that are frequently utilized to study a variety of senescence. First, we discuss key cell biological changes occurring during senescence and review the current understanding on the mechanisms of these changes with the goal of improving existing protocols and further developing new ones. Next, we list specific senescence phenotypes associated with each cellular trait along with the principles of their assay methods and the significance of the assay outcomes. We conclude by selecting appropriate references that demonstrate a typical example of each method.

I. Introduction It is a known fact that normal cells have a finite capacity for proliferation while cancer cells have unlimited capacity for growth. The growth-arrest state of normal cells after a number of active divisions is termed cellular senescence or replicative senescence (Hayflick, 1965). Normal human somatic cells that are frequently cultured in vitro, such as endothelial cells, keratinocytes, chondrocytes, lymphocytes, and certain stem cells, all have low telomerase activity, and their serial division is accompanied with progressive shortening and failure in protection of telomeres at the ends of chromosomes, thus triggering DNA damage response and eventually a state of irreversible growth arrest (Campisi et al., 2001). Replicative senescence is only one type of cellular senescence. In fact, there are multiple experimental routes that can drive cells into a state of senescence. Normal somatic cells enter a state of senescence upon overexpression of oncogenic Ras or Raf protein (Serrano et al., 1997; Zhu et al., 1998). This phenomenon, termed premature senescence or oncogene-induced senescence, may be a manifestation of a cellular strategy to cope with oncogenesis (Campisi, 2007). A state of senescence is also induced in both normal and cancer cells by imposing certain kinds of stress, referred to as stress-induced senescence. DNA damage (e.g., by adriamycin, bleomycin, or actinomycin D) (Elmore et al., 2002; Linge et al., 2007; Robles and Adami, 1998; Toussaint et al., 2000), oxidative stress (e.g., by

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[(Fig._1)TD$IG]

Fig. 1 Induction of senescence in normal and cancer cells. (Left) Normal human fibroblasts at an early passage (P4) were either cultivated continuously until the cell number stopped increasing (P34) or treated with 0.5 mM adriamycin for 4 h and then chased for 9 days (P4 + Adr). Most of the cells at P34 and those treated with adriamycin appeared to be positive for SA b-Gal activity (panel A). (Right) HeLa cells were either mock-treated (mock) or treated to express the BPV1 E2 gene and undergo induced senescence (Pava1) (HeLa cells, a human cervical carcinoma line, express the E6 and E7 oncoproteins of human papillomavirus, which inactivate the p53 and Rb proteins, respectively). BPV1 E2 protein blocks the expression of the E6 and E7 genes and thereby relieves p53 and Rb to activate the growth arrest pathway (Goodwin et al., 2000). Regardless of the method of induction and cell type, the volume of cells and the number of cells positive for SA b-Gal activity evidently increased. In addition, an increase in lysosome content is another cellular trait associated with senescence, as demonstrated here through fluorescence imaging of lysosomes (panel B: by using LysoTrackerRed (left) and LysoTrackerGreen (right); panel C: by using an antibody specific to Lamp2a). The increase in lysosome content is also shown by histograms from flow cytometry of LysoTrackerRedstained cells (panel D). (Reprinted by permission from Park et al., 2007). (See plate no. 4 in the color plate section.)

H2O2 and t-butylhydroperoxide (t-BH)), or certain other chemicals such as bromodeoxyuridine (BrdU) (Suzuki et al., 2001) commonly activate tumor suppressor proteins, p53 or p16INK4a (Campisi and d’Adda di Fagagna, 2007, and see also Fig. 1, for examples). Activation of p53 also appears to be the major mechanism responsible for enforcing oncogene-induced senescence (Campisi, 2007). Therefore, both types of induced senescence are executed by cellular tumor suppressor function. Importantly, cells expressing certain senescence phenotypes are often detected in the tumor masses of animals (Mooi and Peeper, 2006). Considering this, oncogeneinduced senescence in normal cells and stress-induced senescence in cancer cells are regarded as evidence that cellular senescence is a spontaneous organismal defense mechanism against tumor progression that is raised upon the activation of oncoproteins or environmental stresses that must be circumvented for tumorigenesis to occur (Braig et al., 2005; Chen et al., 2005; Courtois-Cox et al., 2006; Michaloglou et al., 2005; Ohtani et al., 2009).

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Early on, Hayflick (1980) published a paper providing an extensive description of a variety of cellular changes associated with replicative senescence. Since then, a number of cytosolic and nuclear senescence-associated cellular traits have been newly recognized with details at the molecular or subcellular level, and their underlying mechanisms have been investigated (Hwang et al., 2009). Major cellular features associated with cellular senescence are listed along with their detection methods in Table I. Certain prominent features such as the presence of b-galactosidase activity Table I Summary of cellular traits associated with cellular senescence and major methods for their detection Senescence-associated cellular traits

Practical plausibilitya

Features associated with cell mortality Shortened telomeres in replicative senescence Decrease in the number of proliferating cells or in the doubling capacity Features associated with cell stasis Increase in cell volume Increase in Vimentin intermediate filaments Increase in focal adhesion Changes associated with lysosome Lipofuscin accumulation Increased lysosome content SA b-Gal activity Changes associated with mitochondria Altered structural dynamics of mitochondria Increased mitochondria content Decreased autophagy activity Changes associated with nucleus and chromosome Appearance of g -H2AX foci Appearance of TIF Appearance of heterochromatin foci (SAHF) Appearance of Hutchinson–Gilford progeria syndrome (HGPS)-like nuclear dysmorphism Increased ROS and oxidative damage adducts Increased mitochondrial superoxide level Increased mitochondrial hydroxyl radical level Increased cytosolic superoxide level Increased cytosolic hydroxyl radical level Increased protein oxidation Other features of cellular senescence Increased granule content (SSC) Increased glycogen granule

p p

p p

Major detection methodsb

GE (Harley et al., 1990), FC (Halaschek-Wiener, 2008), FISH FC with BrdU (Linge et al., 2007), [3H]-thymidine incorporation (Goodwin et al., 2000) CI (Chen et al., 2000) CI (Nishio et al., 2001) CI (Chen et al., 2000) FC (Sitte et al., 2001), FL, FM FC & CI (Park et al., 2007) IS (Dimri et al., 1995), FC (Kurz et al., 2000) CI (Yoon et al., 2006) FC (Lee et al., 2002), CI (Moiseeva et al., 2009) GE & CI (Kang and Hwang, 2009)

p p

p p p p p

CI (Sedelnikova et al., 2004) CI (Herbig et al., 2004) CI (Narita et al., 2003) CI (Scaffidi and Misteli, 2006)

FC (Moiseeva et al., 2009; Invitrogen, in press) FC (G€ orlach and Kietzmann, 2007; Invitrogen, 2010) FC (Moiseeva et al., 2009; Invitrogen, in press) FC (Moiseeva et al., 2009; Invitrogen, in press) GE (Conrad et al., 2000) FC IS (Seo et al., 2008)

GE, gel electrophoresis of nucleic acids or proteins; FC, flow cytometry; FISH, fluorescence in situ hybridization; CI, confocal imaging; FL, fluorometry; FM, fluorescence microscopy; IS, in situ staining. a b

Practical plausibility is based on both how frequently the phenotype is referred to and how reliably and easily it is assayed. A major method is referenced by a representative paper.

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at pH 6.0 (SA b-Gal), increase in autofluorescence and ROS, and senescence-associated heterochromatin foci (SAHF) have been well characterized and are being extensively utilized as markers for senescence and in the study of aging. One of the characteristics of cellular senescence regarding the phenotypes is the wide range of their expression levels among cells in a culture population. Due to such heterogeneity, analysis of the changes in certain cellular properties (e.g., change in the quantity of nucleic acids and proteins) in a senescent cell population could not only suffer from decreased sensitivity and specificity but also provide misleading results. A senescing cell population can be better analyzed by studying the individual cells or subpopulations sorted according to the levels of a certain phenotype. With the simple and easy nature of the procedure, single cell analysis by confocal microscopy and population analysis by flow cytometry, together or separately, are quite well suited for this purpose, and have become the dominant strategies in studies on cellular senescence as summarized in Table I. In this chapter, fluorescence methodologies including confocal imaging and flow cytometry that are frequently utilized to study a variety of senescence phenotypes are reviewed (any missed phenotypes are those that have not been assayed by the fluorescence methods). Key cell biological changes occurring during senescence are listed, and the current understanding on the underlying mechanisms of these changes is reviewed with the goal of improving current protocols and further developing new ones. And, specific senescence phenotypes associated with each cellular trait are listed with the principles of their assay methods and the significance of the assay outcomes are discussed along with pitfalls and limitations. Finally, references are selected that demonstrate a typical example of each method.

II. Features Associated with Loss of Reproductive Cell Capability In actively proliferating early-passage cells, the ends of telomeres form ‘‘t-loop’’ structures that are bound by multiple proteins. However, short telomeres in cells approaching the end of their replicative life span lack such protective structures (de Lange, 2002). The short and unprotected telomeres appear to be recognized as double-strand breaks and trigger the DNA damage response, which eventually activates DNA damage checkpoints executed by p53 and Rb (Deng et al., 2008). Since the shortened telomeres are kept unprotected, the DNA damage response is not extinguished and leads to a state of irreversible growth arrest, constituting the most basic phenotype of replicative senescence. Ongoing DNA damage response at these short telomeres has been demonstrated by the continuous presence of DNA damage response factors such as 53BP1, g -H2AX (phosphorylated histone H2AX), Rad17, ATM, and Mre11 in an ATM-dependent manner at the ends of chromosomes (Takai et al., 2003). Visualized structures are frequently called telomere dysfunction-induced foci (TIF, see below).

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Meanwhile, acute senescence induced in normal and cancer cells is not accompanied by telomere shortening. In these cells, growth arrest is dominantly enforced by p53/Rb or p16INK4a, but the reason for their activation is not completely understood. DNA damage induced by high-level reactive oxygen species (ROS) is likely involved (Campisi, 2007).

A. Shortened Telomeres in Replicative Senescence Telomere shortening is frequently demonstrated through Southern blotting analysis of telomere-restriction fragments (TRFs) using radio-labeled (CCCTAA)4 oligonucleotide, which hybridizes the ‘‘TTAGGG’’ repeat sequence of human telomeres (Harley et al., 1990). In this process, isolated genomic DNA is restrictiondigested (by Hinf1+RsaI, for example), and the lengths of DNA fragments that contain the entire telomeres are visualized by X-ray film exposed to the Southern blot. TRFs from a given population of cells form not a single band but instead a mass of multiple bands of different sizes, reflecting the various lengths of telomeres within and between cells in a population. Still, a gradual decrease in TRF size occurs as the number of population doublings increases (Harley et al., 1990). To numerically demonstrate telomere shortening, the lengths of telomeres in a cell population are determined and plotted. To determine the representative TRF length in a mass of multiple telomeric DNA bands, the lane is serially sectioned into same-sized squares, and the signal intensity of each square is determined by densitometric scanning. Since the signal intensity of a probed DNA band in a Southern blot is the combined outcome of the number of fragments and their lengths, the signal in each section should be normalized based upon the estimated MW of the fragment, and the MWof the section that gives the highest number (this represents the length of the most abundant TRF) is used as the representative TRF length. To assist, a program named Telorun was developed by Harley et al. (1995). A recent version of Telorun is posted at http://www4.utsouthwestern.edu/cellbio/shay-wright/ research/sw_lab_methods.htm. Recently, telomere length has been determined more easily and quickly using fluorochrome-labeled peptide nucleic acid probes (PNAs), which specifically and efficiently bind to telomere repeat sequences (Hacia et al., 1999). PNAs are oligonucleotide analogues in which the natural phosphodiester backbone is replaced by neutral amide linkages (of N-(2-aminoethyl)glycine), increasing their permeation through lipid bilayers (Corey, 1997; Nielsen, 1999). PNAs form a stable duplex with DNA at low salt concentrations without prior fixation with formamide, which causes reduced accessibility of target DNA sequences. Cy3conjugated (CCCTAACCCTAACCCTAA) PNA has been proven to bind to (TTAGGG) repeat sequences with high affinity and efficiently stain telomeres (Hamilton et al., 1997). This specific and quantitative binding allows easy estimation of the length of individual telomeres by flow cytometry (flowFISH (fluorescent in situ hybridization)) (Martens et al., 1998) or digital

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[(Fig._2)TD$IG]

Fig. 2 Fluorescence-based telomere length quantification by flow cytometry (flow-FISH). Cord blood T cells expressing either CD4+ (CB CD4+) or CD8+ (CB CD8+) or adult T cell subsets, naı¨ve, memory, or effector, were hybridized with FITCconjugated PNA-(C3TA2)5 probe (or such PNA probe as specific to sequences at the X chromosome as a control, X-probe) and applied to flow cytometry to obtain fluorescence histograms (A and B). In C and D, the mean telomere fluorescence levels from the different T cell subsets of 10 adult donors and from cord blood T cells were compared with each other. Telomere fluorescence of cord blood T cells is higher than that of adult naı¨ve cells (CD4 + RA+, in A and C or CD8 + RA + 27+, in B and D), which, in turn, is higher than that in the memory cells (CD4 + RO+, in A and C or CD8 + RA
27+, in B and D) or the effector cells (CD8 + RA
27+ in D). The numbers in C and D are the means of telomere fluorescence from the respective T-cell groups. The difference between the subsets of the adult T-cell population from an individual (linked by a line) likely reflects the difference in their replicative history in vivo. (Reprinted by permission from Rufer et al., 1998. Copyright 1998 Macmillan Publishers Ltd.)

fluorescence microscopy (Telomere/centromere-FISH (T/C-FISH)) (Perner et al., 2003; Poon et al., 1999). Flow-FISH can generate data with great ease and speed and also provide information on cell subpopulations. In addition, this method can produce data from quite a small number of cells (Fig. 2). Incorporation of an internal standard (cells of known telomere length) allows reasonable estimation of the mean length of telomeres in a test cell population. Meanwhile, T/C-FISH supported by software modules that determine the integrated fluorescence intensities and reference signal of telomeres also allows measurement of individual telomeres at a single chromosome arm (Perner et al., 2003). Baerlocher et al. (2002) described in detail the important parameters and possible pitfalls of the flow-FISH protocol. A modification of the flow-FISH protocol is presented in Chapter 8 of this Volume.

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B. Telomere Dysfunction-Induced Foci (TIF) in Replicative Senescence

g -H2AX, a DNA damage response factor, localizes en masse at the sites of DNA double-strand breaks (Rogakou et al., 1999) and is visualized as foci (called g -foci) in cells after staining with specific fluorescence-tagged antibody. Likewise, an ongoing DNA damage response can be demonstrated in situ by visualizing the presence of other DNA damage response factors such as 53BP1, Rad17, ATM, and Mre11 at chromosome lesions (Takai et al., 2003). At the same time, the telomeres of individual chromosomes can be visualized by FISH using either Cy3-conjugated PNA probes as mentioned above or immunofluorescence for TRF1, a telomere-associated protein. The telomere ends in association with the DNA damage response complex TIF, has been visualized by dual immunofluorescence for TRF1 and 53BP1 (or g -H2AX or others) or by an immuno-FISH technique (Takai et al., 2003) (Fig. 3), which combines immunofluorescence for g -H2AX (or 53BP1 or others) with telomere FISH (Herbig et al., 2004). The number of TIF has been found to be associated with cellular senescence. In one study, 20% of g -H2AX foci in a replicatively senescent fibroblast coincided with telomere signals, whereas such colocalization is found only occasionally in early-passage cultures (Sedelnikova et al., 2004). Therefore, TIF may be used as a marker for senescence induced by telomere shortening (i.e., replicative senescence). However, the same result also suggests a nontelomeric origin of the majority of the DNA damage foci in senescent cells. Importantly, a higher incidence of TIF in tissues isolated from old baboons (Jeyapalan et al., 2007) suggests its potential as a marker for in vivo aging. [(Fig._3)TD$IG]

Fig. 3

Telomere dysfunction-induced foci. (Left) Cells of a human fibroblast line whose telomeres were uncapped by knockdown of endogenous TRF2 activity, which is essential for telomere protection, were fixed and processed for immunofluorescence against 53BP1 (green) and TRF1 (a telomere-binding protein) (red). The left two images (1.0) are a single nucleus showing the fluorescence image of TRF1 alone and that of TRF1 and 53BP1 merged. The right enlarged images (1.5 and 2.0) show 53BP1 foci localized to telomeres, demonstrating that DNA damage-response machinery containing 53BP1 forms foci at unprotected telomeres. (Reprinted by permission from Takai et al., 2003. Copyright 2003 Elsevier.) (Right) Human fibroblasts at an early passage (p9), at a late passage (p37), in senescence (Sen) or immortalized by telomerase transduction (TERT) were processed by immunofluorescence to visualize g -H2AX foci (green) and by FISH using (C3TA2)3-Cy3-labeled PNA to visualize telomeres (red). Nucleus was counterstained with DAPI (blue). Arrows point to sites of colocalization of green and red fluorescence. The nuclei of senescent cells have more telomeres colocalized with the g -H2AX foci. (Reprinted from Herbig et al., 2004. Copyright 2004 Elsevier.) (See plate no. 5 in the color plate section.)

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C. Loss of the Doubling Capacity of Cells Senescence, by definition, is a state of irreversible cell cycle arrest. However, cell cycle arrest is hardly seen in all constituent cells in a culture that is replicatively senescent. This is in large part due to the heterogeneity of cells regarding doubling capacity. Such heterogeneity is, in turn, likely due to the difference in the timing of telomere-associated DNA damage signaling between individual cells (Martin-Ruiz et al., 2004; Von Zglinicki et al., 2003). The rate of telomere shortening differs from cell to cell (possibly due to different levels of ROS-induced telomere damage), which leads to different times at which DNA damage signaling is triggered (Martin-Ruiz et al., 2004). However, this heterogeneity is not observed with induced senescence, in which growth arrest occurs simultaneously in all cells of a culture. In many cases of induced senescence, cells appear to be arrested within the first 12 h (Elmore et al., 2002, and Sohee Cho, unpublished data). A growth curve is one way to convincingly demonstrate senescence. To construct a long-term growth curve of replicative senescence, cultures are continuously split (at an 1:4 ratio, for example). At each passage, cells are counted, and the number of population doubling (PD) (n) is calculated using the equation, n = log2 F/I, where I and F are the numbers of cells seeded at the beginning and obtained at the end of one passage, respectively. For induced senescence, a short-term growth curve should be sufficient to demonstrate a lack of population growth. In this case, it is important to verify that the absence of population growth is not due to cell death. A senescence-associated decrease in population doubling or an increase in the number of cells under growth arrest can be demonstrated by a significant decrease in DNA synthesis or in the number (or percentage) of cells synthesizing DNA in a given period of time. Quantitative DNA synthesis assay measuring the incorporation of [3H]-thymidine (Goodwin et al., 2000) and immunofluorescence or flow cytometry-mediated counting of the cells positive for nuclear incorporation of BrdU (Linge et al., 2007; Ota et al., 2006) have been the methods of choice in many studies. An increase in the number of cells in G1 phase (or G2/M in certain senescence models, such as those induced by adriamycin or bleomycin) and a concomitant decrease in the cells in S phase can also be used as a supportive measure of senescence induction. This ploidy analysis, although requiring only propidium iodide (PI) staining of the nucleus, does not always give rise to a dramatic result. In replicative senescence, the fraction of cells in S phase does decrease but rarely approaches zero (Linge et al., 2007; Ota et al., 2006).

III. Cellular Hypertrophy Increases in cell surface area and volume are features so prominent in cellular senescence that an experienced eye can easily recognize using only light microscopy. In the case of fibroblasts, cell volume increases at least several folds (Cristofalo and Kritchevsky, 1969; Greenberg et al., 1977). Furthermore,

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senescent cells are flat, which makes the increase in surface area appear even higher (Wang and Gundersen, 1984). The molecular mechanism underlying the increase in cell volume in senescence is under speculation. Doubling of all normal cells requires a growth factor (mitogen) for cell cycle progression and macromolecule synthesis. A major regulator of cellular protein synthesis is the mammalian target of rapamycin (mTOR), which, upon activation, stimulates translation initiation factor 4E binding protein 1 (4EBP1) to form the translation initiation complex at the 50 cap site of mRNAs (Mamane et al., 2006). During cellular senescence, cell cycle arrest occurs in the presence of mitogen, which maintains mTOR activity and overall protein synthesis (Blagosklonny, 2008). As a consequence, both cellular protein content and volume would increase. This is a plausible speculation supported by previous studies reporting upregulated activity of phosphatidylinositol 3-kinase (PI3K), an upstream activator of mTOR in fibroblasts undergoing premature senescence (Tu et al., 2002; Wang et al., 2004). However, there also is accumulated evidence indicating that the rate of protein synthesis is lower in senescent cells (Hayflick, 1980). It is possible that, until a certain point in the growth-arrest state in senescence, protein synthesis may be actively maintained and then turned down later. An easy and simple way to assess a change in cell size is through measurement of intensity of forward light scatter (FSC) by flow cytometry (Fig. 4). Comparison of the mean or peak values in a FSC histogram provides information on the fold change in cell surface area. The surface area and volume of individual cells can also be directly measured by mounting cells on a microslide field finder. The diameter of the cells can be determined by the grids of a field finder, and cell volume may be calculated based on the equation for a sphere: V = (4/3)pr3. Of note, there is a large heterogeneity in FSC levels among cells at or close to senescence. While an FSC histogram of early-passage human fibroblasts gives rise to a single sharp peak at low

[(Fig._4)TD$IG]

Fig. 4

Increased FSC and SSC of senescent human fibroblasts. Cells at an early passage (P4), those undergoing replicative senescence (P34), or those undergoing induced senescence by the treatment with adriamycin, as shown in Fig. 1 (left), were applied to flow cytometry without any treatment. The cells undergoing either type of senescence show an increase in both FSC and SSC. Meanwhile, the population approaching replicative senescence shows high heterogeneity in both FSC and SSC (the population of small FSC and SSC in the dot plot of P34 is likely of the cells fractured during trypsinization). (E. S. Hwang, unpublished data.)

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value, that of the cells in replicative senescence displays a broad curve with a small peak at high values. Such heterogeneity is less prominent in cells undergoing induced senescence in which growth arrest occurs rather synchronously in all cells of a culture, indicating again that the heterogeneity may be in large part related to the difference in the timing of the growth arrest pathway (Martin-Ruiz et al., 2004; Von Zglinicki et al., 2003) (Fig. 4). The morphological changes occurring during cell senescence have been recently analyzed by laser scanning cytometry and were shown to provide a sensitive biomarker of the ‘‘depth’’ of senescence of individual cells (Zhao et al., 2010).

IV. Changes Associated with Lysosomes As cells progress toward replicative senescence, the number and size of lysosomes increase (Comings and Okada, 1970; Park et al., 2007; and Fig. 1). Such a change in lysosome content appears to occur only in senescent cells cultured in vitro, since the lysosome content is not high in fibroblasts freshly isolated from aged individuals but increases after a number of passages in vitro (Robbins et al., 1970). The increase in lysosome content is probably due in large part to an increase in the number of secondary lysosomes that contain indigestible materials such as lipofuscins (see below). If true, it is likely that, in senescent cells, lysosomal enzymes may be wasted in nonfunctional lysosomes, as postulated by Terman et al. (2003). This downshift in lysosome activity would result in reduced turnover of cellular waste materials such as damaged mitochondria. This may be the major reason for the accumulation of dysfunctional mitochondria, which, as will be mentioned later on, produce ROS and thereby initiate a vicious pro-senescence cycle between dysfunctional mitochondria and ROS (Kurz et al., 2008). Meanwhile, the presence of unaltered primary lysosomes and an increase in the activity of certain lysosomal enzymes have also been observed in late-passage cells and senescence-induced cells (Johung et al., 2007; Knook and Sleyster, 1976; Robbins et al., 1970; Sanchez-Martin and Cabezas, 1997). However, the possibility of increased lysosome biosynthesis in senescence has not yet been systematically investigated. Meanwhile, the size of lysosomes also increases in postmitotic cells from aged animals and human subjects (De Priester et al., 1984; Porta et al., 1982; Schmucker and Sachs, 2002). A. Lipofuscin Accumulation Lipofuscins are produced mainly by peroxidation of unsaturated fatty acids in complex with proteins and are deposited as yellow brown pigments in aged tissues. Metals (mercury, aluminum, iron, copper, and/or zinc) are present in lipofuscins, and iron is especially known to be actively involved in lipofuscin genesis by initiating the peroxidation reaction (Brunk and Terman, 2002a). Lipofuscins are believed to be formed within lysosomes, where both transition metals and oxidized lipid peroxides

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are present in high concentrations and undergo Fenton-type reactions to produce hydroxyl radicals, which in turn cause peroxidation of lysosomal contents (Brunk et al., 1992). Lysosomes loaded with these indigestible materials may not turn over and thus remain in the cytosol to become residual bodies (Brunk and Terman, 2002b). As early as 1912, progressive accumulation of lipofuscins concurently with aging was recognized in animal tissues, and recently in various types of cells undergoing senescence (Sitte et al., 2001), and therefore, lipofuscins are considered a genuine marker for senescence and aging. However, the level of lipofuscins increases in cells during temporal growth arrest, whereas, in vivo, they mainly deposit in postmitotic cells such as neurons and cardiac myocytes (Collins and Brunk, 1976). These reports suggest the possibility that cells may continuously produce lipofuscins but then dilute them through cell division, which implies that lipofuscin accumulation may not be an exclusive indicator of cellular senescence.

1. Lipofuscin Autofluorescence in Fluorescence Microscopy Lipofuscins are autofluorescent possibly due to Schiff bases formed by reactions between carbonyls and amino compounds. Their detection in cells and fixed tissues by fluorescence microscopy is rather straightforward; under any excitation wavelength ranging from 360 to 647 nm, lipofuscins appear as irregular granules that emit yellow-orange fluorescence between 500 and 640 nm (Eldred et al., 1982; Eldred and Katz, 1988; Sohal and Brunk, 1989; Strehler, 1964). To obtain quantitative data, micrographic fluorescence images are digitized and quantified using appropriate image analysis software (Sheehy, 1996). Various conditions for fluorescence quantification of lipofuscin have been studied in detail by Thaw (1987). Despite the simplicity of the detection method, some cautionary notes should be mentioned when quantitative detection is attempted. First, fluorescence may also be emitted from normal cellular components, perturbing the detection of lipofuscin fluorescence. Such components include tryptophan (emission of 340 nm), pyridoxine (390 nm), flavins and riboflavins (550 nm), NADH and NADPH (470 nm), porphyrins (630/690 nm), and proteins containing these molecules (RichardsKortum and Sevick-Muraca, 1996). Ceroid pigments, which are similar to lipofuscins in chemistry, are produced under experimental or in vivo pathological conditions unrelated to aging and are known to accumulate in lysosomes emitting fluorescence between 360 and 430 nm (Porta et al., 1988). The background fluorescence contributed by these materials should be subtracted for proper quantification of lipofuscin. Usage of filters for excitation at 390–490 nm/emission at 515 nm can result in mostly yellowish lipofuscin autofluorescence at high intensities (Dowson and Harris, 1981). Second, the broad emission spectra of lipofuscin may cause inaccuracy in the fluorescence-mediated quantification of cellular proteins or ROS in senescent cells. However, no information is available thus far regarding the significance of the errors made by overlapping emission profiles of lipofuscin autofluorescence and commercial fluorophores. Third, exogenous fluorophores (such as phenol red present in most commercial media as a pH indicator) and

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fluorescent probes (such as LysoTrackers or MitoTrackers; see below) can perturb proper data acquisition for lipofuscin fluorescence by causing photobleaching that leads to shifts in the emission signal (Woodburn, 2001). Fourth, formaldehyde, which is frequently used to fix cells for fluorescence microscopy, cross-links various cellular components. Cross-linking of amine residues leads to formation of Schiff bases, which are the chemical base responsible for complex formation between lipid peroxides and proteins in lipofuscin (Ploem, 1971). Therefore, formaldehyde fixation can produce lipofuscin-like artifacts and should be avoided.

2. Flow Cytometric Analysis of Cellular Lipofuscin Levels A change in the level of lipofuscin can be easily determined through flow cytometric analysis. Cells, without fixation, are simply applied to a flow cytometer at an excitation wavelength of 488 nm, and fluorescence is collected by a 530/30 nm bandpass filter. Either the peak or mean fluorescence of a histogram is recorded and compared between different samples. The reported increase in lipofuscin in replicative senescence can be as big as one order of magnitude (Sitte et al., 2001). The basal levels and rates of lipofuscin accumulation in replicative senescence appear to differ between various cell lines. For example, both the level of lipofuscin in early passage cells and the rates of accumulation during progression to senescence are much higher in MRC-5 than BJ cells, which are both primary human fibroblast lines (Sitte et al., 2001). In induced senescence, the level of lipofuscin increases linearly with the duration of incubation (Goodwin et al., 2000; Fig. 5). B. Increased Lysosome Content To detect change in lysosome content, cells are stained live with commercially available acidotropic probes such as LysoTrackerTM (Molecular Probes, Eugene, OR, USA), which basically consists of fluorophores attached to amines. Amines, as weak bases, accumulate in acidic compartments such as late endosomes, secretory vesicles, and lysosomes by a process called pH partitioning (Kaufmann and Krise, 2007). In neutral pH, amines exist as free bases and permeate membranes, but they become protonated and trapped in acidic organelles. In addition, significant binding of amines to acidic polysaccharides and glycolipids, which are abundant in the lysosomal membrane, also localizes these probes to lysosomes (Bulychev et al., 1978; Kaufmann and Krise, 2007). These LysoTracker probes preferentially localize to acidic organelles more so than classical basic dyes such as neutral red and acridine orange (Allison and Young, 1969). However, the probes are still present in the cytosol at quantities high enough to cause background problems. Therefore, quantification of lysosome content based on acidotropic signals in fluorometry and flow cytometry could be unreliable. Meanwhile, immunofluorescence-based lysosome imaging or tagging does not suffer significantly from this background problem. For example, staining lysosomes with a fluorescently tagged antibody for Lamp2a, a lysosome membrane protein, gives rise to a lysosome-specific fluorescence pattern

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[(Fig._5)TD$IG]

Fig. 5 Increase in autofluorescence during senescence induced in HeLa. HeLa cells were induced to undergo senescence by expression of the BPV1 E2 gene as in Fig. 1 (right), further cultivated for the number of days indicated in B, and then applied to flow cytometry at an excitation of 488 nm. The peak fluorescence values of each time point were normalized to the peak value of the mock-control cells to yield a fold increase in autofluorescence (B). In (A), cells photomicrographed at 17 days post BPV1 E2 transduction (left) are highly fluorescent while those mock-treated are so only faintly (right). (Reprinted with permission from Goodwin et al., 2000. Copyright 2000 National Academy of Sciences, USA.)

similar to but more distinct than that of LysoTracker (Fig. 1A and B left). Imaging lysosomes with Lamp2a antibody also helps to eliminate the contribution of other acidic organelles stained with the acidotropic probes. C. Senescence-Associated b-Galactosidase (SA b-Gal) Activity Escherichia coli supplied with 5-bromo-4-chloro-indoly-b-D-galactoside (X-Gal) turns blue when grown on a neutral agar plate due to the activity of b-galactosidase, which cleaves the b-galactoside linkage of the chromophore. Meanwhile, eukaryotic cells express b-galactosidase, a lysosomal enzyme that is active at pH near 4.5 but not so when pH becomes neutral. SA b-Gal activity, which is responsible for cleaving X-Gal and staining cells blue at pH 6.0, is frequently found in cultured cells undergoing replicative and induced senescence

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(Dimri et al., 1995) (Fig. 1A and B). SA b-Gal activity is often observed in the tissues of a variety of aged animals from human beings (Dimri et al., 1995) to Caenorhabditis elegans (Dmitrieva and Burg, 2007) or zebrafish (Kishi et al., 2008) and, therefore, has become a critical marker of both senescence and aging. It turns out that SA b-Gal activity originates from lysosomal b-galactosidase activity, which increases high enough in senescent cells to be detected even at a suboptimal pH (6.0) (Lee et al., 2006). Meanwhile, high b-galactosidase activity at pH 6.0 has also been detected in cells under various conditions not related to senescence (summarized in Hwang et al., 2009), suggesting the possibility that SA b-gal activity may be an indicator of high lysosomal activity rather than an exclusive marker of cellular senescence. Conditions characterized by increased lysosomal activity have yet to be uncovered.

1. In situ Determination of SA b-Gal Activity The SA b-Gal positivity of a culture is usually determined by the assay originally developed by Dimri et al. (1995), which simply involves the incubation of cells in pH 6.0 buffer containing X-Gal. This assay has been cited in thousands of papers and is now the standard method for verifying senescent cells. In middle-passage cultures of human fibroblasts, SA b-Gal-positive cells do exist, albeit at a low frequency, and this number increases with additional passages. Meanwhile, even when the population has stopped doubling and is in a state of senescence, lower than 100% of the constituent cells are stained positive. Cells negative for SA b-Gal activity are usually small and slim, indicating that they are not quite senescent as far as morphology is concerned. This is another manifestation of heterogeneity in a cell population entering a senescent state. In addition, there is no consensus on the quantitative criteria regarding SA b-Gal activity of a culture in senescence. Therefore, it would be wise to track the number of positive cells in the test cultures since it may take many population doublings from a decent amount of positive cells to a very high amount. Considering the above, it may be a common occurrence that certain studies have used a cell population in which a large portion did not reach senescence yet. Meanwhile, in the case of induced senescence, it appears that SA b-Gal positivity remains low during the first several days after initiation of the treatment that causes senescence but increases rapidly thereafter (Sohee Cho, unpublished data).

2. Flow Cytometry for SA b-Gal-Positive Cells and Quantification of Activity There is room for controversy as to whether or not cells are sufficiently stained positive for SA b-Gal activity, due to the subjective nature of judging color. A method to objectively verify a change in SA b-Gal activity in a population of cells was developed by Kurz et al. (2000) based on the ‘‘FACS-Gal assay’’ initially designed by Nolan et al. (1988) and Fiering et al. (1991). This flow cytometric analysis detects the hydrolysis of 5-dodecanoylaminofluorescein di-b-D-galactopyranoside (C12FDG), a membrane-permeable and nonfluorescent substrate of

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b-galactosidase, which, upon cleavage, remains within the cytosol and emits green fluorescence. A histogram of the fluorescence levels allows estimation of the enzyme activity and also provides quantitative information on the population size of SA b-Gal-positive cells.

V. Changes Associated with Mitochondria Hundreds to thousands of mitochondria are present in a single mammalian cell. They undergo qualitative, quantitative, and morphological changes during senescence and aging, and under certain pathological conditions. Deterioration in mitochondrial respiratory chain function and accumulation of mutations and large deletions in mitochondrial DNA accompany senescence and aging as well as certain agerelated diseases (Boffoli et al., 1994; Lezza et al., 1994; Trifunovic et al., 2004). However, an increase in the size of individual mitochondria and total mitochondrial mass is one of the most prominent changes exhibited as cells progress toward replicative senescence or undergo induced senescence. Mitochondria suffer from a decrease in the activities of fusion and fission and thereby remain enlarged during senescence. A. Increased Mitochondrial Mass Mitochondrial mass as well as the lengths of individual mitochondria increase during replicative and oncogene or stress-induced senescence in human fibroblasts (Lee and Wei, 2001; Lee et al., 1998, 2002; Passos et al., 2007; Moiseeva et al., 2009; Yoon et al., 2006; and Fig. 6). Increases in mitochondrial protein and DNA content as well as mass have also been observed in tissues of aged animals (Beregi and Regius, 1987; O’Connell and Ohlendieck, 2009; Sachs et al., 1977). It seems that at least two different causes underlie the increase in mitochondrial mass: an increase in biogenesis and a decrease in mitochondrial turnover. First, cells may make more mitochondria in an attempt to compensate for the decline in mitochondrial function caused by ROS-induced damage (Lee et al., 1998; Moiseeva et al., 2009; Wei et al., 2001). Oxidative stress and inhibition of mitochondrial electron transport chain function have been shown to cause an increase in mRNA levels of mitochondrial proteins (Fu et al., 2008; Lee et al., 2000; Miranda et al., 1999). It also has been reported that DNA damage (including that caused by H2O2 treatment) activates AMP-activated protein kinase (AMPK), which in turn induces the mitochondrial biogenesis factors PGC-1a, NRF-1, and TFAM (Fu et al., 2008). The levels of TFAM and NRF-1 and their DNA binding activities were also found substantially increased in aged subjects (Lezza et al., 2001). Second, mitochondrial mass may also increase as a consequence of decreased mitochondria turnover, which is, in large part, mediated by autophagy (Bota and Davies, 2001). A decrease in autophagy activity has been observed in cells of aged animals (Cuervo and Dice, 2000) and likely contributes to the accumulation of

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Fig. 6 Increase in mitochondrial mass and formation of elongated mitochondria in senescent cells. Mv1Lu cells (a mink lung epithelial line) were induced to undergo senescence by treatment with 2 ng/mL of TGF-b1 (A and B) or 0.8 mM H2O2 followed by chase (C). Electron photomicrography (A and C) shows elongation and enlargement of mitochondria in cells at senescence caused by either treatment. (B) Progressive increase in mitochondrial mass in cells undergoing senescence is shown by flow cytometric analysis following staining with MitoTrackerTM Red. (Reprinted with permission from Yoon et al., 2006. Copyright 2006 John Wiley & Sons.)

altered mitochondria in aged tissues as well as senescent cells (Bergamini et al., 2007; Yen and Klionsky, 2008). The decline in autophagy activity may be caused by a decrease in autophagosome formation and/or functional lysosomes (Terman, 1995). In addition, the efficiency of autophagic removal of mitochondria (sometimes called mitophagy) also depends on the structure of the substrate mitochondria. For example, filamentous mitochondria would not be easily enwrapped by autophagosomes. Decreased mitochondrial fission has been observed in senescent cells (see below), and this may contribute to the increase in mitochondrial mass (Lee et al., 2007).

1. Quantification of Changes in Mitochondria Mass Cellular content of mitochondria can be quantified by measuring the fluorescence emitted from cells stained with a fluorescent probe specific to mitochondria by flow

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cytometry or fluorometry. Conventional mitochondrial probes such as rhodamine 123 and tetramethylrosamine are mostly lipophilic, and therefore permeate membranes, but they also contain one positive charge, and get retained in the inner mitochondrial membrane facing the negatively charged matrix. However, these probes are susceptible to photobleaching, and more importantly, are easily lost during fixation, which inevitably reduces the MMP (Chen, 1989). Meanwhile, a family of chemicals that contain a thiol-reactive chloromethyl moiety (CMXRos) has been developed (Poot et al., 1996). Like the conventional probes, these are taken up into mitochondria. However, they then interact with thiol groups of membrane proteins, which prevents their loss during fixation. Such probes have been commercially named ‘‘MitoTrackerTM’’ (Molecular Probes, Eugene, OR, USA), and they are most heavily used as fluorescent mitochondrial probes. Quantification of MitoTracker fluorescence in conjunction with changes in cellular FSC and side scattering (SSC) by flow cytometry provides reliable information on changes in the mitochondrial contents of cell subgroups during senescence. A comprehensive report on these mitochondrial probes by Poot et al. (1996) was published. It should be noted that most of these mitochondrial probes, including some of the MitoTracker families, are responsive to MMP. In other words, the quantity of fluorescence is subject to changes in the membrane potential of the particular mitochondrion. Therefore, to quantitatively analyze the mitochondrial content of cells in which a change in MMP is expected, the choice of mitochondrial probe should be made carefully. MitoTrackerTM Green FM is claimed by the manufacturer to not be affected by MMP (MitoTracker and Mitofluor Mitochondrion-Selective Probes, Molecular Probes). Meanwhile, 10-n-nonyl acridinium-orange chloride (NAO) binds to cardiolipin, a special type of phospholipid present only in the mitochondrial inner membrane, and thereby localizes to the inner membranes (Petit et al., 1992). NAO has been reported to not respond to MMP-altering drugs such as dinitrophenol (DNP) or carbonyl m-chlorophenylhydrazone (CCCP), and therefore it is considered a genuine probe for detection of mitochondrial mass (Benel et al., 1989; Ferlini et al., 1995; Ratinaud et al., 1988; Septinus et al., 1985). However, the MMP-independence of NAO has been challenged (Keij et al., 2000). It seems that NAO fluorescence may also be affected by changes in MMP, albeit to a much lesser degree compared to other probes. Furthermore, the emission of green fluorescence (through 530/30 nm band path filter, for example) by NAO may be less sensitive to MMP changes. Practically, mitochondrial content may be better determined by assaying both MTG and NAO fluorescence, although, in most cases, quantifications made using these two chemicals provide similar results (Fig. 7A). The best way, but not an easy way, to determine mitochondrial mass independent of MMP is to permeabilize cells and label mitochondria using a fluorescently tagged antibody specific for mitochondrial proteins, which would be detected by flow cytometry or fluorospectrometry. Other independent measures such as quantification of mitochondrial DNA by real-time PCR or protein bands of electron transport chain complexes by Western blotting can provide supplementary documentation of changes in mitochondrial mass (see Fig. 7A–C for an example).

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Fig. 7 Changes in cellular mitochondrial content demonstrated by several independent methods. Human fibroblasts were cultured for different durations in medium containing nicotinamide, which has been reported to cause a decrease in mitochondrial mass. (A) Cells were fixed, stained with either 10-nonylacridine-orange bromide (NAO) or MitoTrackerTM Red (MTR), and analyzed by flow cytometry. The mean fluorescence was divided by that of the mock-treated cells, and the relative values were plotted. Fluorescence of both chemicals rapidly decreased during the first 7 days. (B) Western blotting analysis was carried out for the proteins of the electron transport chain complexes of the cells incubated as in (A). (C) Total DNA from an equal number of cells was isolated and applied to real-time PCR for relative quantification of mitochondrial DNA. The levels of both mitochondrial proteins and DNA changed in a pattern similar to that of the mitochondrial fluorescence in (A), supporting the decrease in mitochondrial content in nicotinamide-treated cells. (D) Relative levels of mitochondrial protein mRNAs quantified by real-time PCR. (Reprinted with permission from Kang and Hwang, 2009. Copyright 2009 John Wiley & Sons.) (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of the chapter.)

B. Altered Structural Dynamics of Mitochondria In human cells undergoing replicative senescence or stress-induced senescence, mitochondria frequently change their morphology from short fragments or dots to long, thin, and irregularly shaped reticula (Jendrach et al., 2005; Lee et al., 2007; Yoon et al., 2006) (Fig. 6A and C). This filamentous degeneration is largely attributed to a decrease in the structural dynamics of mitochondria, especially fission activity (Lee et al., 2007; Yoon et al., 2006). In early-passage cells, mitochondria rapidly change their structure through frequent fission and fusion, which are mediated by special proteins; Fis1 and DrpI for fission and OpaI and MfnI (and II) for fusion (Hyde et al., 2010). However, the fission and fusion activities of late-passage HUVEC cells were substantially lower, and mitochondria existed as huge aggregates (Jendrach et al., 2005). Meanwhile, the expression of Fis1 and the fission activity in

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cellular senescence were found to be substantially downregulated (Yoon et al., 2006). And, subtoxic doses of H2O2 caused a state of cellular senescence while also decreasing mitochondrial fission and inducing elongated giant mitochondria (Yoon et al., 2006). Meanwhile, the alteration of mitochondrial structural dynamics may also play a causative role in senescence and aging. Inhibition of fission activity not only caused the formation of elongated mitochondria but also elevated the level of ROS and induced a state of senescence (Lee et al., 2007; Yoon et al., 2006). Conversely, overexpression of Fis1 delayed expression of the senescence phenotype (Yoon et al., 2006).

1. Visualization of Altered Mitochondria Structure The shift from short and fragmentary mitochondria to a thin reticulum is a change easily noticeable by confocal microscopy of cells after staining with a variety of the mitochondria-selective probes mentioned above (Jendrach et al., 2005; Yoon et al., 2006). Since most of these chemicals depend on high MMP and are washed out of the cells once MMP drops, they should be used without cell fixation. Meanwhile, the MitoTracker dyes based on CMXRos mentioned earlier can be used with other fluorescent probes that require cell fixation for coimaging of mitochondria. However, Minamikawa et al. (1999) raised a concern for using CMXRos in live cell imaging. CMXRos may cause loss of MMP, mitochondrial swelling, and release of cytochrome c. In addition, they warned that repeated laser scanning of CMXRosstained cells during confocal microscopy can damage mitochondria, leading to both substantial alteration in the image and MMP and severe phototoxity-induced cell death. It is noteworthy that staining of mitochondria using antibodies specific for mitochondrial proteins after fixing gives rise to a consistently better mitochondrial image (Sohee Cho, unpublished result). Meanwhile, visualization and quantification of mitochondrial fusion are possible with the use of mitochondrial matrix-targeted photoactivatable GFP (mtPA-GFP), which emits fluorescence upon irradiation with a laser at 750 nm (Twig et al., 2008). Mitochondrial fusion events can be deduced by determining the decrease in the GFP fluorescence intensity through the spread of fluorescence to mitochondria that are previously unlabeled.

C. Decreased Membrane Potential Mitochondria in senescent cells (Chen et al., 2005; Jendrach et al., 2005) and tissues of aged animals have low membrane potential (Navarro and Boveris, 2007). More precisely, the number of mitochondria with low MMP increases as cells are continuously passaged or induced to undergo senescence. The decrease in MMP may be attributed to the accumulation of defects in electron transport chain function, which is, in turn, brought about by either ROS-induced oxidative damage or expression of mutant proteins from damaged mtDNA (Lee and Wei, 2001).

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1. Quantitative Comparison of MMP The mitochondria matrix is negatively charged and certain lipophilic cations are trapped in the mitochondria inner membrane. Therefore, the extent of their uptake could reflect the potential across the membrane. By using such fluorescently labeled cations, a change in MMP can be either visualized in the individual mitochondria of a living cell via confocal microscopy or quantitatively presented by flow cytometry. Although there are a variety of fluorescent probes that respond to changes in MMP as mentioned above, JC-1 (5,50 ,6,60 -tetrachloro-1,10 ,3,30 tetraethylbenzimidazolylcarbocyanine iodide) is the most frequently used and favored probe since it is highly sensitive and responds consistently to MMP changes and is not sensitive to changes in plasma membrane potential (Salvioli et al., 1997). JC-1 exists as a monomer in the cytosol or mitochondria with low MMP, and emits green fluorescence (510/527 nm). However, JC-1 can accumulate in mitochondria with high MMP as J-aggregates emitting red fluorescence (585/590 nm) when excited at 490 nm (Cossarizza et al., 1993). Therefore, confocal imaging using JC-1 can identify mitochondria with high or low MMP (Kang and Hwang, 2009; Fig. 8) by

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Fig. 8 JC-1 staining of mitochondria. Human fibroblasts were stained with JC-1, washed with PBS, and visualized by confocal microscopy. To detect fluorescence from JC-1 monomers, samples were excited by an argon laser at 488 nm (green), and to detect fluorescence from JC-1 aggregates, a helium/neon laser at 543 nm (red) were used, respectively. The right bigger image is an overlap of the two left images. It is apparent that the green fluorescence localized mostly in the background of the cytosol, although some localized to form filaments, suggesting that a certain part of the filamentous mitochondria with low MMP was visualized. Meanwhile, all of the red fluorescence formed short filaments. This indicates that mitochondria in these cells are a mixed population of those with high MMP and others with low MMP. (Reprinted with permission from Kang and Hwang, 2009. Copyright 2009 John Wiley & Sons.) (See plate no. 6 in the color plate section.)

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utilizing two different optical filters (one for fluorescein and another for tetramethylrhodamine) for separate visualization of green and red emissions. For flow cytometric determination of the MMP level (but not an absolute voltage), both the FL1 (using a bandpass filter of 530/30) and FL2 (using a bandpass filter of 585/ 42) measures of individual cells are collected, after which the FL2 value is normalized to the FL1 value and the ratio is plotted to produce a histogram of the population. This way, possible error due to variation in the levels of JC-1 uptake or in the sizes among cells can be minimized. It is important to plot a histogram using the FL2/FL1 values from individual cells in order to obtain a mean or pik value (not of the population’s mean FL2 value normalized to the population’s mean FL1 value). A histogram of individual cell FL2/FL1 values can be drawn by running the raw data in a flow cytometry data analysis program such as Weasle (freeware developed by Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia). In addition, alteration in MMP is sometimes accompanied by a change in mitochondrial content (as in cells undergoing senescence). In such a case, the FL2/FL1 ratio might not correctly reflect changes in MMP. Therefore, mitochondrial quantity needs to be simultaneously but independently measured (by using NAO, for example) and used to normalize the mean FL2 values in place of the JC-1 FL1 values. A minimal concentration of JC-1 would most likely minimize red fluorescence emission from the cytosol, although an amount of about 1 mM is used in typical flow cytometric analysis. Treatment of cells with carbonyl cyanide m-chlorophenylhydrazone (CCCP), a potent uncoupler that causes rapid depolarization of mitochondria (Heytler and Prichard, 1962), reduces the emission of red fluorescence and therefore can serve as a good negative control. Tetramethylrhodamine ethyl ester (TMRE) and 2-(4-(dimethylamino)styryl)-1methylpyridinium iodide (DASPMI) are rapidly and reversibly taken up by live cells (Loew et al., 1994) and then used to demonstrate a shift in MMP. Such compounds are sequestered by functioning mitochondria in an MMP-dependent manner. Therefore, their relative intensities could highlight differences in MMP. Like JC-1, these voltage-sensitive fluorescence dyes should be applied to unfixed live cells. Quantitative data can also be produced by analyzing cells with digitalized confocal imaging. However, this method raises concerns in that mitochondria are not static or fixed structures. Mitochondria migrate and undergo rapid fusion and fission (Twig et al., 2008), resulting in movement out of the focal plane and decreased fluorescence intensity. Therefore, identification of changes in MMP based upon the fluorescence intensity of the confocal image requires costaining of mitochondria using an MMP-independent mitochondrial probe and determination of the ratio between the fluorescence intensities. Finally, it is important to remember that MMP measurement using these fluorescent probes is sensitive to the measurement conditions. Cells are to be measured at the same pH, temperature, and for the same time between the probe treatment and measurement.

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D. Decreased Autophagy The increase in mitochondrial mass in cells from aged individuals as well as in those undergoing senescence may be due to a decline in autophagy as mentioned above (Bergamini et al., 2007; Cuervo and Dice, 2000; Yen and Klionsky, 2008). The reason for this decline is not known in detail, but a decrease in autophagosome formation combined with a delay in autophagosome elimination may conceivably lead to decreased turnover of mitochondria and other damaged organelles. Recent evidence indicates that SIRT1, an NAD+-dependent deacetylase involved in a variety of physiological processes such as metabolism, cell survival, and senescence plays an active role in autophagosome formation by deacetylating and thereby activating several key autophagy proteins such as Atg5, Atg7, and Atg8 (Lee et al., 2008). A progressive decrease in SIRT1 activity during the cellular replicative life span has been reported (Sasaki et al., 2006), and this in combination with the degeneration of mitochondria may contribute to the senescence-associated decline in mitophagy. Meanwhile, autophagosome elimination may decline in part due to lysosomal dysfuction. Either lipofuscin overload or AGE-induced permeabilization of the lysosomal membrane (Patschan et al., 2008) may cause a decrease in lysosomal enzyme activity or autophagolysosomes formation.

1. Determination of Autophagy Activity LC3 protein is a key factor in the process of autophagosome formation. During the early stage of autophagosome formation, the C-terminus of LC3 is removed by the action of a cysteine protesase forming a smaller species (LC3-II), and the exposed glycine residue is conjugated to phosphatidylethanolamine on the autophagosome membrane. LC3-II is easily separated from the inactive full-length LC3 protein, and increases in its level can be monitored by SDS-PAGE to confirm autophagy activation. However, LC3 protein levels are difficult to predict due to variations between cells in their levels as wells as in the rates of autophagosome formation and elimination. Therefore, an increase in the ratio of LC3-II molecule/LC3 molecule rather than a change in LC3-II protein level alone can be considered a supportive evidence for autophagy activation (Fig. 9 and see Proikas-Cezanne et al., 2007 for an example). Frequently, GFP (or other types of fluorescence protein)-tagged LC3 is transfected into test cells, and increased formation of GFP puncta in the cytosol can be considered an indicator of increased autophagosome formation (and autophagy activity) (Proikas-Cezanne et al., 2007, for an example). Puncta formation is also observed with endogenous molecules by immunofluorescence (Kang and Hwang, 2009). LC3 puncta colocalization with or localization near mitochondria is also presented as supportive data for autophagy activation (Kim and Lemasters, 2011). However, LC3 proteins in autophagosomes are eventually removed through lysosome-mediated degradation, which makes detection of LC3 puncta formation unreliable. Sometimes, treatment with an inhibitor of autophagosome–lysosome fusion (bafilomycin A1) or lysosome acidification (monensin) results in higher levels of

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Fig. 9 WIPI-1 puncta formation in cells with active autophagy. Human G361 cells were treated with rapamycin or incubated in EBSS medium (as an amino acid-deprivation treatment) to induce autophagy in the presence or absence of wortmannin (an inhibitor of autophagy). (A) WIPI-1 puncta formation was detected in the rapamycin-treated cells by immunofluorescence (antiWIPI-1 antiserum/anti-rabbit IgG Alexa 488) and confocal microscopy. (B and C) The percentage of puncta-positive cells and the ratio of puncta/nonpuncta cells are presented. These graphs demonstrate that WIPI-1 puncta formation is attenuated by wortmannin treatment. (d) LC3-I and LC3-II proteins were detected by Western blotting using anti-LC3 antibody, and the LC3II/LC3-I ratios, which have been used as a major indication of autophagy activation, are presented in the bar graph. Different from WIPI-1 puncta formation, the increase in LC3-II/LC3-I ratio was not completely attenuated by wortmannin treatment. (Reprinted with permission from Proikas-Cezanne et al., 2007. Copyright 2007 Elsevier.)

LC3 puncta and helps to demonstrate autophagy activation. GFP-LC3 has been used in flow cytometry for quantification of autophagy activation. Different from LC3 puncta, GFP-LC3 fluorescence decreases during active autophagy due to rapid autophagolysis. In a study, the mean cellular GFP fluorescence decreased by 50% after 3 h of starvation (a known inducer of autophagy) in a manner sensitive to inhibitors of autophagy or lysosome activity (Shvets et al., 2008). Meanwhile, WDrepeat protein interacting with phosphoinositides (WIPI-1), another component of autophagosome, has been reported to accumulate upon autophagy stimulation, forming puncta that overlap with those of LC3 (Fig. 9A–C). WIPI-1 puncta appear to form more reliably than LC3 puncta do (Proikas-Cezanne et al., 2007), implying they may be better suited for quantitative analysis of autophagy.

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VI. Changes in the Level of Reactive Oxygen Species (ROS) Various ROS and nitrogen species (RNS) are produced in cells. Superoxide (O2
), hydroxyl (OH), peroxyl (RO2
), alkoxyl (RO), hydroperoxyl (HO2) radicals, and nonradical species such as hydrogen peroxide (H2O2), hypochlorous acid (HOCl), ozone (O3), singlet oxygen (1O2), and peroxynitrite (ONOO
) are those. ROS production in tissues increases with age (Perez-Campo et al., 1998; Sohal and Sohal, 1991) and in cells as they proliferate toward the end of their replicative life span (Hutter et al., 2002; Kang et al., 2006). ROS production also increases during Ras-induced senescence in human fibroblasts (Lee et al., 1999) and DNA damageinduced senescence in human fibroblasts and cancer cells (Song et al., 2005). ROS not only increase during senescence but also function as a common trigger of cellular senescence. Human fibroblasts grown in low oxygen proliferate for a longer time (Packer and Fuehr, 1977), whereas those grown in high oxygen have a shorter life span with accelerated telomere-shortening rates (Von Zglinicki et al., 1995). Treatment of primary fibroblasts with nonlethal doses of H2O2 induces the senescence phenotype (Chen and Ames, 1994). Meanwhile, Ras-induced senescence could be attenuated either by reducing ambient oxygen or by treatment with an antioxidant (Lee et al., 1999). Superoxide, hydrogen peroxide, and hydroxyl radical are directly associated with oxidative damage in cells. The levels of both superoxide and hydroxyl radical increase in mitochondria and cytosol during senescence (Fig. 10). Especially, a large quantity of superoxide anion is produced and serves as a major source of elevated levels of hydrogen peroxide and hydroxyl radical. Most superoxide anions are produced through electron leaking during oxidative phosphorylation in mitochondria, especially at complex I (NADH dehydrogenase) and complex III (ubiquinone– cytochrome c reductase) (Turrens, 1997). In proliferating human cells, approximately 0.1–4% of electrons are estimated to leak from the electron transport chain and interact with molecular oxygen to form superoxide anion (Batandier et al., 2002; Kudin et al., 2004). ROS production from mitochondria and its role in senescence and aging support the ‘‘mitochondria theory of aging’’ hypothesis (Linnane et al., 1989; Wei and Lee, 2002). Oxidative stress is an important contributor to mitochondrial dysfunction during aging and senescence. An increase in mitochondrial ROS production was shown to directly impair mitochondrial respiratory function (Melov et al., 1999). mtDNA, which lack protective histones, are direct targets of ROS attack (GarciaRuiz et al., 1995; Wei, 1998). And, damage-induced mtDNA mutations would lead to expression of defective proteins in electron transport chain complexes, which augments ROS production. Thereby, it is predicted that ROS are produced by dysfunctional mitochondria and aggravate the problem by causing more damage to mitochondria and mtDNA, thus leading to even more ROS production (Balaban et al., 2005; Harman, 1972). Although plausible, this hypothesis still lacks certain critical data (e.g., data that rule out association of mitochondrial dysfunction with other factors involved in aging or senescence).

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Fig. 10

Increase in levels of superoxide and hydroxyl radicals in mitochondria and the cytosol of senescent cells. MCF-7 cells were induced to undergo senescence by a pulse of 0.5 mM adriamycin as described in Fig. 1 (left). At day 6 postadriamycin pulse, cells were incubated with 0.1 mM MitoSox (Invitrogen), 5 mM DHE, 15 mM DHR123, or 10 mM DCF for 30 min, followed by flow cytometry to determine the levels of mitochondrial and cytosolic superoxide and of mitochondrial and cytosolic hydroxyl radical, respectively. Note that the increase in cytosolic and mitochondrial superoxide levels is more prominent than that of hydroxyl radical in this type of DNA damage-induced senescence. (Sohee Cho, unpublished data.)

It is not easy to correctly quantify the presence of ROS in cells due to their short lifetime and the presence of a variety of antioxidant chemicals and enzymes at different levels in cells (Khan et al., 1992). Quantitative analysis can be further hindered by the high intracellular concentration of glutathione, which can form thiyl or sulfinyl radicals while at the same time reducing oxygen species (Winterbourn and Metodiewa, 1994). In addition, metal ions that are present at variable concentrations can either promote or inhibit radical reactions (Khan et al., 1992). Among the many ROS, superoxide and hydroxyl radicals are exclusively assayed since they are the major species that cause the most damage to cellular macromolecules and due to their well-established methods for detection and quantification.

A. Changes in Level of Superoxide Hydroethidine (HE), more frequently called dihydroethidium (DHE), is one of the most widely used fluorogenic probes for detection of intracellular superoxide. DHE, a reduced form of ethidium (Etd+), is rapidly taken up by live cells and oxidized to

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Etd+, which then intercalates into nucleic acids and emits fluorescence at 610 nm when excited at 535 nm (Bucana et al., 1986)). However, DHE can be oxidized by H2O2 (Rothe and Valet, 1990) or by cellular processes involving peroxidases, oxidases, or cytochrome C to yield Etd+ (Benov et al., 1998; Bucana et al., 1986; Henderson and Chappell, 1993; LeBel et al., 1992; Zhu et al., 1994). Consequently, an increase in Etd+ fluorescence may not necessarily prove the increase in superoxide production. However, it was recently found that DHE oxidation by O2
is a two-step mechanism in which DHE is further oxidized by O2
to yield hydroxylated ethidium (HO-Etd+) (Zhao et al., 2003). HO-Etd+ is also a red fluorescent product, but it also has a distinct excitation wavelength (at 396 nm) (Robinson et al., 2006). Therefore, the cellular superoxide level can be specifically quantified and analyzed by fluorometry and flow cytometry (and visualized by fluorescence microscopy) by measuring the fluorescence intensity of HO-Etd+ using excitation and emission wavelengths of 400 nm and 590 nm, respectively. Recently, DHE was modified to allow detection of superoxide within mitochondria. DHE was covalently attached to hexyl triphenylphosphonium. The three lipophilic phenyl groups facilitate penetration of the molecule through the membranes, and the positive charge of phosphonium also facilitates its accumulation in the mitochondrial matrix, which has negative membrane potential (Ross et al., 2005). In mitochondria, this compound is then oxidized by superoxide and emits bright red fluorescence. This modified molecule has been commercially termed ‘‘MitoSOXTM Red’’ (Molecular Probes) and is used frequently to monitor changes in the O2
level in mitochondria (Fig. 10C). MitoSOXTM Red reagent was shown to be readily oxidized by superoxide but not by other ROS or RNS, and oxidation of the probe was shown to be prevented by superoxide dismutase (Janes et al., 2004).

B. Changes in Level of Hydroxyl Radicals In cells, hydroxyl radical is mostly derived from superoxide in a reaction called the Fenton reaction, which is catalyzed by Fe2+ or other transition metals (Chevion, 1988). The hydroxyl radical is very reactive and has a lifetime of about 2 ns in aqueous solution, and induces peroxidation of molecules present only in very close proximity. Consequently, quantitative detection of hydroxyl radical requires high sensitivity. Fluorogenic probes used for detection of cellular levels of hydroxyl radicals have been produced by exploiting a property common to fluorescent dyes such as fluorescein and rhodamine. The reduced ‘‘dihydro’’ forms of these molecules are nonfluorescent but freely permeate the lipid bilayer, similar to DHE (aforementioned). However, inside cells, these modified probes are readily oxidized by hydroxyl radicals to emit fluorescence. Nonfluorescent 20 ,70 -dichlorodihydrofluorescein diacetate (H2DCF-DA) crosses the cell membrane and is deacetylated by cellular esterases to 20 ,70 -dichlorodihydrofluorescein (H2DCF), which is then rapidly oxidized to highly fluorescent 20 ,70 -dichlorofluorescein (DCF) by the radicals (Brandt

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and Keston, 1965; LeBel et al., 1992; Zhu et al., 1994). The fluorescence intensity of DCF can be quantitatively detected and analyzed by fluorometry or flow cytometry or visualized by fluorescence microscopy at excitation and emission wavelengths of 480 nm and 530 nm, respectively (or with excitation sources and filters appropriate for fluorescein (FITC)) (Fig. 10). Carboxy or chloromethyl (CM) derivatives of H2DCF-DA (carboxy-H2DCF-DA or CM-H2DCF-DA) and a fluorinated derivative (H2DFFDA) have been developed for better retention and improved photostability, and they are currently commercially available (Molecular Probes). It should be noted that H2DCF is oxidized by peroxynitrite anion (ONOO–), horseradish peroxidase, or Fe2+ (in the absence of H2O2) (Myhre et al., 2003), which means it is not exclusively specific for hydroxyl radical. It is also important to note that regarding the use of these fluorogenic dyes, cells should be washed and resuspended in buffer free of phenol red or other colorimetric dyes prior to and during the measurement. Since the dyes are susceptible to photooxidation, work should be carried out in the dark. Dihydrorhodamine 123 (DHR123), like MitoSOXTM Red (aforementioned), freely passes through membranes and are then selectively trapped in mitochondria due its positive charge (Johnson et al., 1980) and oxidized to rhodamine 123, which emits fluorescence. DHR 123 reacts with H2O2 and peroxynitrite but poorly with superoxide (Henderson and Chappell, 1993). Therefore, DHR 123 provides information on the overall levels of nonsuperoxide ROS in mitochondria. A modified dihydrorhodamine (DHR 6G), whose oxidized form has a longer wavelength spectrum than rhodamine 123, is commercially available (Invitrogen) and can be used in multifluorescent probe analysis as well as for simultaneous measurement of ROS and autofluorescence (Wersto et al., 1996). All these reduced (and nonfluorescent) chemicals are slowly oxidized in air back to the parent fluorescent dyes and therefore should not be stored for too long, especially since this oxidation is accelerated by exposure to light.

VII. Changes Associated with Nucleus and Chromosomes The structure of the nucleus and chromatins also undergo conspicuous changes during cellular senescence. The appearance of SAHF is a well-documented nuclear trait of cellular senescence. This phenotype has not been addressed in a population study and can only be demonstrated in individual cells examined by confocal imaging. However, since SAHF is the most frequently referred to nuclear phenotype of senescence, it is mentioned here in detail. When actively proliferating or early-passage cells are treated with 40 ,6-diamidine20 -phenylindole dihydrochloride (DAPI), a fluorochrome that binds selectively to DNA, nuclei appear as bright ovals with relatively uniform distribution of fluorescence. However, when a senescent cell is stained, numerous bright foci appear in the fluorescent image of the nucleus. These foci represent severely condensed chromatin called SAHF (Narita et al., 2003). SAHF appear in cells in either replicative or

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[(Fig._1)TD$IG]

Fig. 11 Accumulation of heterochromatin foci in the nuclei of senescent cells. IMR90 cells at an early passage were either mock-treated or induced to undergo senescence (by the expression of H-rasV12 (Ras) or MEK1 Q56P (MEK)). High SA b-Gal activity confirms senescence. Along with these, IMR90 cells at a late-passage, those induced to quiescence (low serum) or those expressing both E1A and H-rasV12 (E1A/Ras) (E1A blocks oncogenic Ras-induced senescence) were treated with DAPI 6 days posttreatment. Enlarged images of DAPI-stained nuclei shown in the lower panels indicate that the nuclei in the control and nonsenescent cells produce a rather uniform staining pattern while those in the senescent cells are represented by small fluorescent puncta, which are referred to as senescence-associated heterochromatin foci. Scale bars are equal to 10 mm. (Reprinted with permission from Narita et al., 2003. Copyright 2003 Elsevier.)

induced senescence (Narita et al., 2003) (Fig. 11) and also marks senescent cells found in tumor masses in animal models (Braig et al., 2005). Due to the simplicity of the method (does not even need a confocal microscope), it has great potential to be utilized in the studies on senescence and aging. SAHF may be a mechanism underlying the irreversible growth arrest of senescent cells. In SAHF, proteins such as heterochromatin protein 1 (HP1) and histone H3 methylated on Lys 9 (H3K9m) bind to DNA en masse and seem to cause transcriptional incompetence (Narita et al., 2003; Zhang et al., 2005). Each SAHF is quite large and appears to cover a large portion of, if not the entire, chromosome. Genes that are embedded in SAHF include E2F-responsive genes, most of which are required for cell cycle progression (Narita et al., 2003). The molecular mechanism for SAHF formation is not well understood, although high-mobility group A (HMGA), nonhistone proteins, and a group of chaperones that deposit histones into nucleosomes have been shown to be involved (Narita et al., 2006; Zhang et al., 2005, 2007). It is noteworthy that SAHF formation in WI38 and IMR90 human fibroblasts is quite pronounced, but not in BJ human fibroblasts (Denoyelle et al., 2006; Narita et al., 2003). No solid explanation of this discrepancy has been provided, but this suggests that the activity of factors controlling SAHF formation is not modulated equally in all cell types. Meanwhile, although the occurrence of SAHF-positive cells in tissues of aged

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humans has not yet been shown, the possibility has been suggested by the presence of SAHF in the tissues of aging baboons (Herbig et al., 2006; Smith et al., 2002).

VIII. Use of Flow Cytometry for Analysis of Cellular Senescence As mentioned often above, the occurrence of senescence in a cell population is not a synchronous event and often occurs stochastically. Any culture has a fraction of cells that exhibit certain phenotype(s) of senescent cells. Meanwhile, it is often the case that not all cells exhibit senescence in a population that is actually presumed to be in senescence. In a population approaching the end of its replicative capacity, the exact size of the fraction of cells that exhibit senescence cannot be predicted. Certainly, handling a whole population in a culture en bloc and obtaining quantitative data for senescence would cause inaccuracy and confusion, especially for a first-time researcher of senescence. For example, an increase in the quantity of a phenotype in a population of cells undergoing senescence may occur in two different situations. The portion of the cells that are positive for the activity of interest may increase, or the activity may increase in individual positive cells without an increase in cell number. Increases in both the number of positive cells and the activity of interest in a limited number of cells would result in a similar increase in the mean value of the whole culture. These two cases cannot be easily resolved in assays such as Western blotting or fluorometry, which analyze the whole population. However, differentiation of multiple populations in a test culture in flow cytometry would allow an accurate quantitative assessment of any cell physiological processes including senescence. Second, determination of a culture of cells as being senescent is frequently dependent on a single time point assay for a senescence phenotype such as SA b-Gal activity. However, expression of a single senescence phenotype in a culture sometimes may not prove senescence. For example, SA b-Gal activity also appears in cells that are not in senescence (Coates, 2002; Cristofalo, 2005; Yang and Hum, 2005). Lipofuscin accumulation may also be used as a marker for senescence, but it is also known to occur in cells in a state of temporary growth arrest (Collins and Brunk, 1976). This emphasizes the necessity for assaying multiple phenotypes, which is, in many cases, possible through flow cytometry. Finally, senescence has not been defined quantitatively as of yet. The kinetics and quantitative measurement of changes in the levels of various cellular traits associated with senescence are rarely considered important or subjected to thorough investigation. However, such studies certainly would reveal the interrelationship between those changes and promote understanding of cell biological and molecular networks of senescence.

IX. Conclusion Recently, evidence supports that senescence is not simply a passive state arrived by exhaustion of doubling capacity but instead a condition actively affecting body

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function. Its role in tumor suppression is one example. Furthermore, senescent cells actively secrete cytokines and various molecules, collectively termed the secretome, which are found to be involved in regulation of immune function and tissue remodeling in wounds healing (Adams, 2009; Campisi and d’Adda di Fagagna, 2007; Krizhanovsky et al., 2008; Swann and Smyth, 2007). More physiological roles (and pathological ones as well) for senescence are expected to be uncovered in the future, and this will definitely be accelerated by the understanding of the underlying mechanisms of the various cellular phenotypes, which requires improvement of the detection methods in terms of both sensitivity and diversity. Especially, methods that allow quantitative analysis of cellular changes will be of utmost importance. Furthermore, new senescence markers and methods that allow for quantitative analysis can be utilized as diagnostic and prognostic tools and also potentially used as screening tools for senescence-inducing drugs as anticancer agents. Overall, both flow cytometry and confocal imaging analysis are greatly advantageous over biochemical and cell biological methods. In addition to easiness, they allow quantitative estimates of multiple different phenotypes expressed in multiple cell populations simultaneously. Utilization of flow cytometry and confocal imaging analysis equipped with various fluorogenic probes for a variety of proteins and organelles will dramatically increase in future studies on senescence. The imageassisted cytometric approaches such as provided by laser scanning cytometry (Pozarowski et al., 2005; Zhao et al., 2010) will additionally be expanding analytical capabilities probing populations of cells undergoing senescence. The image-assisted cytometric approaches such as provided by laser scanning cytometry (Pozarowski et al., 2005; Zhao et al., 2010) will additionally be expanding analytical capabilities probing populations of cells undergoing senescence.

Acknowledgments This work was supported by the Mid-Career Researcher Program through an NRF grant funded by the MEST (No. 2009-0086432) and the Center for Aging and Apoptosis Research (R11-2002-097-07002-0) funded by the MEST.

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