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Keywords:

  • Aging;
  • connective tissue;
  • mitochondria;
  • p16INK4a;
  • reactive oxygen species;
  • Sod2

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

The free radical theory of aging postulates that the production of mitochondrial reactive oxygen species is the major determinant of aging and lifespan. Its role in aging of the connective tissue has not yet been established, even though the incidence of aging-related disorders in connective tissue-rich organs is high, causing major disability in the elderly. We have now addressed this question experimentally by creating mice with conditional deficiency of the mitochondrial manganese superoxide dismutase in fibroblasts and other mesenchyme-derived cells of connective tissues in all organs. Here, we have shown for the first time that the connective tissue-specific lack of superoxide anion detoxification in the mitochondria results in reduced lifespan and premature onset of aging-related phenotypes such as weight loss, skin atrophy, kyphosis (curvature of the spine), osteoporosis and muscle degeneration in mutant mice. Increase in p16INK4a, a robust in vivo marker for fibroblast aging, may contribute to the observed phenotype. This novel model is particularly suited to decipher the underlying mechanisms and to develop hopefully novel connective tissue-specific anti-aging strategies.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

Although reactive oxygen species (ROS) are part of normal regulatory circuits, imbalance or loss of cellular redox homoeostasis during aging results in oxidative stress (Sies, 1986, 1991), causing damage to cellular components. Apart from permanent genetic damage involving protooncogenes and tumour suppressor genes, ROS activate cytoplasmic signal transduction pathways that are related to growth, differentiation, tissue homoeostasis and senescence. Therefore, ROS have been implicated to play a causal role in tissue degeneration and – if deviant in composition and concentration – also in cancer and aging. Increased ROS concentration occurs in senescent fibroblast in vitro and in vivo (Scharffetter-Kochanek et al., 1997; Allen et al., 1999; Hutter et al., 2002; Shin et al., 2005). Fibroblasts constitute the principal component of the connective tissue. As such, fibroblasts occur in virtually every tissue and organ of the body. The capacity of fibroblasts to synthesize and organize the extracellular matrix and to communicate with other cells and tissues makes them a so far underestimated central component in organ homoeostasis and aging. Notably, the incidence of aging-related disorders such as skin- and muscle atrophy, osteoporosis and delayed wound healing are particularly high in connective tissue-rich organs including skin, bone and muscle causing major disability in the elderly. Skin aging is characterized by the loss of the elastic and collagen fibre network due to dysfunctional fibroblasts (Fisher et al., 2008; Quan et al., 2010) leading to wrinkle formation, reduced tensile strength and impaired wound healing. With aging, a progressive loss of skeletal muscle mass and strength occurs, a condition termed sarcopenia or muscle atrophy. Similar to skin atrophy and the loss of bone mass in the elderly (osteoporosis), muscle atrophy is a universal characteristic of the aging process in several species from worm to human (Fisher, 2004; Hiona & Leeuwenburgh, 2008). In older individuals, loss of muscle and bone mass is highly predictive of falls (Newton-John & Morgan, 1970; Szulc et al., 2005), disability (Rantanen et al., 1999) and mortality (Metter et al., 2002). It leads to mobility disabilities with poor life quality and increased health care and social needs for the elderly (Iannuzzi-Sucich et al., 2002). These pathological states share unique features and are all characterized by a loss of collagen type I, dysregulated fibroblast-matrix interactions and impaired fibroblast interactions with organ parenchyma, mainly with organ-specific epithelial cells and muscle (Wenk et al., 1999, 2004; Krtolica & Campisi, 2002; Campisi, 2005; Labat-Robert & Robert, 2007; Treiber et al., 2009). A variety of genetic and environmental factors including increased concentration of ROS, mitochondrial dysfunction (Hiona & Leeuwenburgh, 2008), changes in autocrine, paracrine and endocrine release of hormones, growth factors (Perrini et al., 2010) and cytokines (Coppe et al., 2008) have been identified to contribute to skin aging, sarcopenia and osteoporosis in humans and rodents (Zofkova, 2003; Raisz, 2005; Ralston & de Crombrugghe, 2006; Hiona & Leeuwenburgh, 2008; Marzetti et al., 2009). Research on the regulation of connective tissue organization by enhanced release of ROS from mitochondria during fibroblast aging is a matter of increasing interest and relevance as it may provide ultimate clues for mechanisms underlying disruption of connective tissue homoeostasis in aging-related skin atrophy, sarcopenia and osteoporosis.

To protect against injury, aerobic cells, fibroblasts in particular have evolved a multilayered interdependent antioxidant system that includes enzymatic and non-enzymatic components. Among these Manganese superoxide dismutase (Sod2), a homotetramer, has been the subject of particular interest because it is located in the mitochondria and represents the first line of defence against superoxide radical mainly produced as by-product of oxidative phosphorylation. It converts superoxide radical into hydrogen peroxide, which is further detoxified to water by glutathione peroxidase and other peroxidases. Sod2 has earlier been identified as part of a phylogenetically conserved signalling pathway controlling aging and longevity at least in lower organisms (Kops et al., 2002). Also, defined polymorphisms in the Sod2 gene correlate with longevity and heart failure in elderly humans (Valenti et al., 2004; Stessman et al., 2005). However, due to early lethality in Sod2 complete knock-out mice (Li et al., 1995; Lebovitz et al., 1996) and unchanged lifespan in mice heterozygous for the Sod2 deficiency (Van Remmen et al., 2003), the role of Sod2 and its specific requirements in distinct histogenetic compartments like the mesenchyme-derived connective tissues to prevent aging phenotypes have not yet been analysed. This information would be particularly important as connective tissue consists of resident mesenchyme-derived cells (e.g. fibroblasts) and extracellular matrix proteins, which are essential for organ function, its homoeostasis and most likely also for organismal aging. Most of the earlier studies investigate the effect of antioxidant enzymes deficiency or overexpression in all cells of transgenic mice for aging-related phenotypes (Muller et al., 2007; Van Remmen & Jones, 2009). Even though these studies have provided valuable insight into aging-related disorders, they do not allow specifically investigating the impact of pro-oxidant connective tissue cells on skin, muscle and bone aging. This question is particularly interesting as fibroblasts revealed an accumulation of ROS during senescence in vitro and in vivo (Hutter et al., 2002; Shin et al., 2005) and given the ubiquitous distribution of fibroblasts in connective tissue stromas of all organs, it is likely that fibroblasts influence organ and organismal aging.

We have used a connective tissue-specific strategy to delete Sod2 resulting in mutant mice. In contrast to mice with heterozygous Sod2 deficiency in all organs (Van Remmen et al., 2003; Strassburger et al., 2005), we found a complex premature aging phenotype with skin atrophy, osteoporosis, muscle degeneration and a significantly reduced lifespan in the mutant mice. Fibroblasts in the skin of mutant mice revealed an increase in the expression of p16INK4a, but did not undergo enhanced apoptosis, suggesting that atrophy in the skin, and most likely also in other organs, is due to the instalment of an oxidative damage-induced senescence programme.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

Generation of the mutant phenotype

To circumvent early lethality of Sod2-deficient mice (Li et al., 1995; Lebovitz et al., 1996; Strassburger et al., 2005) and to address the impact of connective tissue-specific Sod2 deficiency on aging of individual organs and lifespan, we have used a connective tissue-specific strategy with the introduction of loxP sites flanking exon 3 of the Sod2 gene (Strassburger et al., 2005). Deletion of exon 3 completely abrogates Sod2 activity, as exon 3 codes for the Sod2 tetramer formation domain (Li et al., 1995). Mice carrying the floxed Sod2 allele (Sod2f/f) (Strassburger et al., 2005) were bred with Col1α2-Cre+Sod2+/+ transgenic mice (Florin et al., 2004) to generate Col1α2-Cre+Sod2+/f mice. The connective tissue-specific Cre expression was achieved by the activation of α2(I) collagen promoter. While the above cross (first mating) resulted in a connective tissue-specific Cre expression, the second mating (Col1α2-Cre+Sod2+/f x Col1α2-CreSod2f/f) from the offspring of the first mating, due to transient Cre activation during gametogenesis in offspring of the first mating, resulted in mutant mice (mut) with connective tissue-specific homozygous deficiency for Sod2 and a heterozygous deficiency for Sod2 in all other organ parenchyma (Col1α2-Cre+Sod2−/f; Data S1 and Fig. S1, Supporting information). To assess specifically the impact of the connective tissue-specific homozygous lack of Sod2 on aging, mutant mice (mut) were compared with mice revealing a heterozygous deficiency for Sod2 in all organs (h) and to Sod2-competent mice (co) with wild type levels of Sod2 activity. As we never observed differences between Sod2+/+, Col1α2-CreSod2+/f, Col1α2-CreSod2f/f and Col1α2-Cre+Sod2+/+ in phenotype nor in Sod2 activity, these genotypes were comparably used for experiments and were referred to as Sod2-competent control mice (co). Also, no difference in the Col1α2-CreSod2−/f and Sod2+/− was observed, and these genotypes were referred to as Sod2 heterozygous mice (h).

Mutant mice reveal a progeroid phenotype and a decrease in lifespan

The mutant mice had a normal appearance until the age of 10–20 days, when a smaller size and reduced weight were first noted. As the mice got older, the kyphosis and decrease in size and weight became very obvious (Fig. 1a–c). Mutant mice have a progeroid appearance with a typical prominent forehead compared with Sod2-competent (co) mice (Fig. 1a). Reduction in length and weight occurred to a similar extent in female and male mutant mice (Fig. 1b,c) compared with Sod2-competent mice. A decrease in body weight was reported in human aging (Kalu, 1995) and is also well known in mice older than ∼1.5 years (Haines et al., 2001). The probability of survival decreased sharply during the first 4 weeks of life and thereafter decreased to a lesser extent (Fig. 1d), a finding most likely due to the mixed genetic background (Sv129 × C57BL/6J) of the mutant mice. No difference in lifespan was observed in Sod2 heterozygous mice (h) compared with Sod2-competent mice (co, Fig. 1d), confirming previous data (Van Remmen et al., 2003). The early dying mice can clinically be distinguished from late dying mutant mice by their almost complete immobility, their lack of feeding as controlled by metabolic cages (Fig. S2, Supporting information) and an increased rate of apoptosis in the hearts (Fig. S3, Supporting information). In contrast, the late dying mutant mice feed comparable quantities as control mice (Fig. S2, Supporting information) and did not reveal significantly increased apoptosis in the heart muscle (Fig. S3, Supporting information). In the following experiments, we have included only late dying mutant mice (> 100 days), as the early dying mutant mice (< 100 days) may not represent a convincing aging model, but may be more pertinent to a disease model.

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Figure 1.  Progeroid gross appearance and decrease in lifespan in mutant mice. (a–c) The mutant mice (mut) at an age of 150 days revealed kyphosis and a prominent progeroid forehead compared with littermates of Sod2-competent (co) mice (a), reduced body size (b) and weight (c). Female controls (f co) vs. female mutant mice (f mut), *= 0.005, male controls (m co) vs. male mutant (m mut) mice, **= 0.003 (after Bonferroni correction). All error bars indicate standard deviation. (d) Significantly reduced survival time of mutant mice (median 444 days) compared with Sod2-competent control mice (median 784 days) (***< 0.0001, log-rank test stratified for gender). No statistically significant difference was found between Sod2-competent (co) and Sod2 heterozygous (h) mice (= 0.65). The maximal lifespan for mutant mice was 837 days compared with 971 days in Sod2-competent mice and 932 days in Sod2 heterozygous mice.

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Accelerated aging phenotype in muscle, bone and skin of mutant mice

A motor phenotype with progressive limb clasping consistently occurred upon tail suspension in mutant mice (mut, 30/32) compared to no clasping in Sod2-competent (co, 0/40) and Sod2 heterozygous mice (h, 0/40) (Fig. 2a). This limb clasping was progressive and more severe in older animals. Testing motor coordination and balance by means of the rotarod test, mutant mice revealed a severe phenotype with only 50% performance (5/5) compared with mice of either Sod2 heterozygous or Sod2-competent genotype (data not shown). Histology revealed a large reduction in the diameters of muscle fibres and high variability of fibre size indicating a severe muscle cell atrophy in mutant mice (mut, 5/5, 150 days) compared with aged-matched Sod2-competent (co, 0/5) and Sod2 heterozygous littermates (h, 0/5) (Fig. 2b, c). Variability of fibre size and reduction in diameter of muscle fibres were also found in intrinsically aged control mice of 900 days (old) (Fig. 2c). In addition, transverse sections of skeletal muscle from Sod2 heterozygous and competent mice did not reveal gross differences in morphology (Fig. 2b,c). The ratio of quadriceps mass (weight) to body weight revealed a reduction in muscle mass in mutant (150 days) and intrinsically aged control mice (900 days), but not in heterozygous-deficient mice (150 days) and littermate control mice of the same age (Fig. 2d). These data indicate that the reduced muscle fibre diameter is due to muscle atrophy and not simply to overall reduced body weight. Notably, similar to intrinsically aged control mice (900 days), a severe osteoporosis-like phenotype was found in mutant mice (150 days) as quantitative assessment with X-ray densitometry of femur bones revealed a dramatic decrease in the mineral density in mutant mice (mut, 5/5) compared with Sod2-competent mice (co, 0/5) of the same age (Fig. 3a). In addition, severe skin atrophy was detected in mutant (150 days) and old mice (900 days), but not in Sod2 heterozygous-deficient mice compared with Sod2-competent mice at the same age of 150 days (Fig. 3b and Fig. S4, Supporting information). Histologically, mutant mice showed severe skin atrophy in the dermal and the subcutaneous compartment with reduced vertical diameter (5/5) compared with no or less atrophy in Sod2 heterozygous-deficient (5/5) and control mice (5/5) (Fig. 3b and Fig. S4, Supporting information). In addition, also the muscular layer of the panniculus carnosus in the skin is atrophic with reduced vertical diameter (5/5). Similar results were found in intrinsically aged mice of > 900 days (Fig. 3b and Fig. S4, Supporting information).

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Figure 2.  Motor phenotype and muscle atrophy in mutant mice. (a) Mutant mice at ages ranging from 100 days to > 600 days revealed clasping of hind limbs (indicating deficits in motor coordination) (mut, 30/32), whereas Sod2-competent mice (co, 0/40) at the same age exhibited normal hind limb extension upon tail suspension. (b) Representative haematoxylin-eosin stained sections of the quadriceps muscle revealed muscle atrophy in mutant mice (mut, 150 days), and intrinsically aged mice (old, 900 days), Compared with the normal muscle architecture in Sod2 heterozygous-deficient (h, 150 days) and Sod2-competent mice (co, 150 days). (c) Diameters of skeletal muscle derived from mutant mice (mut) and intrinsically aged mice (old, 900 days) were significantly reduced compared with Sod2-competent (co) and Sod2 heterozygous (h) mice at an age of 150 days. Observations and the mean value are shown (co/h vs. mut, ***< 0.001, co vs. old **< 0.01 after Bonferroni correction). Scale bars, 20 μm. (d) Ratio of quadriceps muscle weight to body weight (co/h vs. mut, ***< 0.001, co vs. old **< 0.001, after Bonferroni correction).

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Figure 3.  Osteoporosis-like phenotype and skin atrophy in mutant mice. (a) X-ray analysis (left panel) and bone density (right panel) from dissected femur bone of Sod2-competent mice (co), Sod2-heterozygous mice (h), mutant mice (mut) and intrinsically aged mice (old) revealed an osteoporosis-like phenotype in mutant mice (co/h vs. mut, ***< 0.001; after Bonferroni correction) and old mice (co vs. old, ***< 0.001; after Bonferroni correction). Scale bars, 0.5 cm. co, h, mut mice were studied at an age of 150 days and intrinsically aged mice at an age of 900 days (old). (b) Mutant mice at an age of 150 days revealed a severe atrophy of the dermis, the subcutaneous fat tissue and the muscle fibres of the panniculus carnosus compared with Sod2-competent mice. Only minor changes in Sod2 heterozygous-deficient mice at the same age were evident compared with Sod2-competent mice. Intrinsically aged mice of 900 days (old) revealed a similar phenotype as mutant mice. Representative haematoxylin-eosin stained sections of the skin. E, epidermis; D, dermis; H, hair follicle; SC, subcutaneous fat tissue; PC, panniculus carnosus. Scale bars, 20 μm. (c) Representative Masson’s trichrome staining revealed an altered organization and reduced thickness of collagen fibres and the dermal connective tissue in all mutant mice (mut, 6/6) at an age of 150 days and intrinsically aged (old, 900 days, 3/3) mice compared with the normal architecture of the dermis in Sod2-competent controls (co, 5/5) and heterozygous-deficient (h) mice at an age of 150 days. Collagen fibres of Sod2-competent (co), heterozygous (h), mutant (mut) and intrinsically aged (old) mice are shown at higher magnification (lower left panel). Higher magnification of Masson’s trichrome staining of dermal collagen fibres in Sod2-competent control (co) and Sod2 heterozygous (h)-deficient mice revealed densely packed thick fibres. In contrast, in mutant and old mice, collagen fibrils are thinner and loosely organized. E, epidermis; D, dermis; SC, subcutaneous fat tissue; PC, panniculus carnosus. Scale bars, 20 μm for overview and 5 μm for magnified view. (d) Western blot analyses of skin lysates with antibodies directed against pro-collagen type I (upper panel) revealed a significant decrease in pro-collagen type I in mutant mice (mut) at different ages (50, 94, and 189 days) compared with age-matched Sod2-competent controls (co). Actin served as a loading control. Semi-quantitative RT-PCR of pro-collagen type I mRNA isolated from the dermis of mutant mice (mut) and Sod2-competent mice (co) at different ages (lower panel) as indicated above. GAPDH served as housekeeping gene and loading control.

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To study the impact of Sod2 deficiency of fibroblasts in the skin, deposition of collagen and collagen fibril organization were assessed. Using Masson’s trichrome staining for collagen, the atrophy of the dermis as indicated by reduced vertical thickness of the dermis in the mutant mice (mut) (Fig. 3c, left middle panels) compared with Sod2-competent (co) littermates was confirmed (Fig. 3c left upper panel). Mice with a Sod2 heterozygous deficiency (h) (Fig. 3c, upper right panel) did not differ from control mice (co). In addition, skin of intrinsically aged (old) mice of 900 days (Fig. 3c, middle right panel) revealed a similar atrophy of the dermis as found in mutant mice. The lower panel (Fig. 3c) depicted that virtually no difference in thickness and density of collagen fibres occurred in control mice when compared with Sod2 heterozygous-deficient mice. Notably, smaller collagen fibres loosely arranged in less density were observed both in mutant mice and in old mice. Western blot analysis of skin lysates of mice at different ages revealed a significant decrease in collagen type I [α12)2I] to almost undetectable levels in mutant mice (Fig. 3d). Interestingly, specific collagen mRNA levels did not differ in mutant and Sod2-competent mice suggesting that the decrease in collagen deposition occurs at the translational and/or posttranslational level (Fig. 3d).

Deletion of Sod2 is fibroblast-specific in different organs of mutant mice

Crossing the Sod2 floxed mice with ROSA26-LacZ reporter mice, revealed β-galactosidase expression only in interstitial fibroblasts in different organs (Fig. S5, Supporting information). To show specific Sod2 deficiency in the connective tissue resident fibroblasts in skin and skeletal muscle, we performed double staining with an antibody against Sod2 and an antibody directed against a fibroblast-specific intracellular epitope of reticulin (Er-TR7) which previously has been successfully used to define the microarchitecture of organs (Petrie & Van Ewijk, 2002; Kim et al., 2009). Expectedly, skin and muscle (Fig. S6, Supporting information) from control mice revealed double staining exclusively in fibroblasts when merged, while all other cells in skin and muscle fibres stained for Sod2. In contrast, in the skin of mutant mice, exclusive ER-TR7 positive staining was observed in fibroblasts, while no staining for Sod2 was detected. The epidermis, hair follicles and the muscle layer of the panniculus carnosus of the skin, as well as skeletal muscle fibres revealed a clear staining for Sod2 in the mutant mice. These data indicate a connective tissue-specific lack of Sod2 in fibroblasts studied in the skin and skeletal muscle (Fig. S6, Supporting information) and other organs (data not shown).

Mutant mice reveal oxidative damage and a premature increase in aging-related in vivo biomarkers

Using Western blot analysis, we confirmed the lack of Sod2 in mutant fibroblasts (Fig. 4a). The lack of Sod2 in mutant fibroblasts led to an increase in superoxide anion concentration (Fig. 4b) (P < 0.05), as measured by indirect fluorescence with the mitochondrion-specific superoxide anion indicator, MitoSOX. Adenovirally transduction of the HyPer construct was used to visualize specifically changes in H2O2 levels in the mitochondria of Sod2-deficient and competent fibroblasts. A reduction of mitochondrial H2O2 in Sod2-deficient fibroblasts compared with Sod2-competent fibroblasts was found (Fig. 4c) (P < 0.05). Lack of Sod2 in fibroblasts led to severe structural changes in 80% of the mitochondria with loss of cristae and degeneration, with > 60% intact mitochondria in Sod2 heterozygous and 95% intact mitochondria in Sod2-competent fibroblasts (Fig. 4d). Oxidative damage of protein as detected by carbonylation, was consistently increased in lysates of dermal fibroblasts from mutant mice, compared with Sod2-competent fibroblasts (Fig. 4e) and heterozygous fibroblasts (data not shown).

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Figure 4.  Oxidative damage, accumulation of superoxide anion and decrease in hydrogen peroxide in skin fibroblasts of mutant mice. (a) Representative Western blot analysis of skin fibroblast lysates derived from the Sod2-competent (co), heterozygous (h) and mutant mice (mut) were probed with an anti Sod2 antibody. Actin was used as loading control. (b) Superoxide anions in fibroblasts were quantified using indirect fluorescence with MitoSOX. MitoSOX is mitochondrion-specific, and the reaction of superoxide anions with this dye caused fluorescence emission. Fibroblasts from mutant mice revealed a significantly higher fluorescence emission compared with Sod2-competent mice, indicating an accumulation of superoxide anion in the mitochondria of mutant mice. Data are expressed as percentage of mean ± standard error of mean, (*< 0.05 co vs. mut). The experiment shown here is representative of three independent experiments. (c) Fibroblasts from control and mutant mice were adenovirally transduced with the construct of mitochondrially targeted HyPer (Ad Mito-HyPer) and subjected to FACS analysis for quantification of HyPer oxidation in the mitochondria as a measure of defined H2O2 levels. Fluorescence intensities of AdMito-HyPer in mutant fibroblasts and Sod2-competent fibroblasts were measured following activation by 488 and 405 nm and expressed as fluorescence ratio 488/405 as indicated in Materials and Methods, comparing fluorescence ratio of mutant fibroblasts with Sod2-competent fibroblasts. Data indicate that H2O2 levels are significantly reduced in mutant compared with Sod2-competent fibroblasts (*< 0.05). The data are expressed as mean ± standard error of mean. Three independent experiments were performed in triplicates with a representative experiment shown here. (d) Transmission electron microscopy and assessment of 100 mitochondria revealed a severely disturbed structure with loss of cristae and degeneration of intramitochondrical structure in > 80% of all assessed mitochondria in mutant fibroblasts (mut) compared with 40% damaged mitochondria in Sod2 heterozygous (h) and only 5% in Sod2-competent fibroblasts (co). Similar data were found in three additional mice of each genotype in the dermis. Representative mitochondria are shown for each genotype. (e) Oxyblots were performed with lysates from dermal fibroblasts of mutant (mut) and Sod2-competent mice (co) with significantly enhanced general protein carbonylation indicative of oxidative damage in all mutant mice (mut, 3/3) compared with less carbonylation in Sod2-competent mice (co, 3/3). Two independent lysates of the competent and mutant genotype are shown.

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Increase in superoxide anion concentrations, as is the case in mutant mice, may induce oxidative DNA damage and DNA double strand breaks. This can activate cell cycle check points through p21 or p53, and subsequently results in growth arrest of cells harbouring DNA damage. Alternatively, a cellular senescence programme can be activated by DNA double strand breaks, resulting in irreversible p16-mediated cellular growth arrest. Cell cycle arrest thus prevents that the damaged DNA becomes mutagenic for daughter cells (for review see Campisi, 2005). We did not find any upregulation of p21 and p53 (data not shown). However, fibroblasts in the skin of mutant mice at ages ranging from 150 to 550 days revealed high expression of p16INK4a (Fig. 5a, b), a cell cycle inhibitor, which not only mediates the DNA damage response to the senescence state with induction of growth arrest, but also constitutes a robust in vivo biomarker for aging of different organs including the skin (Krishnamurthy et al., 2004; Ressler et al., 2006). Enhanced p16INK4a expression was also found in the skin of intrinsically aged control mice of 700 and 900 days (Fig. 5b). Western blot analysis showed a slight increase in p16INK4a in the skeletal muscle of mutant mice at the age of 300 days compared with Sod2 control mice at the same age (Fig. 5b). In contrast, apoptosis of resident fibroblasts in the dermis of 150 days old mutant mice did not differ significantly compared with Sod2-competent mice, as revealed by ‘Terminal deoxynucleotidyl’ transferase dUTP nick end labelling (TUNEL) and active caspase-3 immunostaining (Fig. 5c, d). Also Western blot analysis did not show any difference in the activation of caspase-3 in skin lysates of mutant mice at an age of 150 and 550 days (data not shown), indicating that apoptosis is not enhanced in the dermis of mutant mice.

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Figure 5.  Oxidative damage resulted in the induction of the senescence marker p16INK4a in mutant mice. (a) Immunostaining with antibodies against p16INK4a, an established aging marker, was significantly increased in the dermal connective tissue of all mutant mice at an age of 550 days (mut, 5/5) compared with Sod2 heterozygous (h, 0/5) and Sod2 compentent (co, 0/5) at the same age of 550 days. p16INK4a was stained in red (Alexa 555) and nuclei in blue (DAPI). Immunostaining of the skin and the mean value of p16INK4a positive cells are shown (co vs, mut, ***< 0.001, h vs, mut **< 0.01 after Bonferroni correction). E, epidermis; D, dermis; H, hair follicle. Scale bars, 20 μm. (b) Upper left panel shows Western blot analysis of p16INK4a in skin lysates of 150 days old co, h and mut mice. Upper right panel shows Western blot analysis of p16INK4a in skin lysates of 150, 700 and 900 days old control mice. Left third panel shows Western blot analysis of p16INK4a in skeletal muscle lysate of 150 days old co, h and mut mice and right third panel shows Western blot analysis of p16INK4a in skeletal muscle lysates of 300 days old co, h and mut mice. Actin served as loading control for the p16INK4a Western blot. (c) ‘Terminal deoxynucleotidyl’ Transferase dUTP nick end labelling (TUNEL) assay (left panel) of skin sections and quantification of TUNEL positive cells (right panel) did not reveal any significant difference in apoptosis in the skin of Sod2 mutant (150 days) compared with control mice of the same age. (d) Immunostaining with antibodies against cleaved (active) caspase-3 indicative of apoptotic cells was performed on skin sections (left panel) and was quantitatively assessed (right panel). No significant difference was observed between Sod2-competent (co) and mutant mice (mut) at an age of 150 days.

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Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

Here, we have shown for the first time that connective tissue-specific lack of superoxide anion detoxification resulted in a reduced lifespan and premature onset of aging-related phenotypes, such as weight loss, skin atrophy, kyphosis, osteoporosis and muscle degeneration. This is most likely due to oxidative damage-induced activation of a senescence programme with overexpression of p16INK4a and irreversible growth arrest of connective tissue-associated fibroblasts with substantially reduced collagen deposition and impaired interaction with interstitial collagen fibrils, the major structural component of the extracellular matrix in the skin, bone and muscle.

Our major finding is that selective loss of redox balance in connective tissue resident fibroblasts is sufficient to drive such a complex aging phenotype in different organs and to shorten lifespan. In fact, the mice with a heterozygous Sod2 deficiency did not reveal this severe progeroid phenotype in all studied aspects. This is further supported by few independent publications, which did not report a reduced lifespan, bone or muscle phenotype in mice with heterozygous Sod2 deficiency (Van Remmen et al., 2003; Strassburger et al., 2005). We observed a bimodal shape of the lifespan, with early dying mice during the first weeks after birth and late dying mutant mice, which revealed a reduced lifespan compared with control mice. Several reasons including segregation of modifier genes or replicative segregation of mutant mitochondrial DNA or a combination thereof may contribute to the early death of the mutants and to the bimodal shape of the lifespan curve in the mutant mice. The studied mutant mice are on a mixed background derived from C57BL/6J and SV129/ola inbred strains. Interestingly, genetic modifiers of lifespan of mice deficient in Sod2 have earlier been reported (Huang et al., 2006). The gene encoding nicotinamide nucleotide transhydrogenase (Nnt) was found to be one of such modifiers in the C57BL/6J mice, where this gene is mutated and functional Nnt protein could not be detected. Nnt is a nuclear encoded mitochondrial inner membrane protein which couples the generation of NADPH to proton transport and provides NADPH for the generation of two important antioxidants (glutathione and thioredoxin) in the mitochondria. If non-functional as in the C57BL/6J background, the mutant Nnt could contribute to the lifespan shortening in Sod2-deficient mice (Huang et al., 2006). To test the possibility that the early dying mutants in our study reveal a mutation in the Nnt gene, we systematically screened for this mutation. However, we did not detect any correlation with lifespan shortening and the Nnt mutation in the early dying mutants (data not shown). This does not, however, exclude the segregation of other so far unknown modifier genes responsible for shortening of the lifespan in early dying mutant mice. In addition, the stochastic and quantitative nature of mitochondrial DNA genetics is known to result in a highly variable inheritance and expression of heteroplasmic mutations (Wallace et al., 1999; Wallace, 2010) and thus may contribute to the bimodal shape of the lifespan curve in the herein described mutant mice.

The mutant mice with a complete Sod2 deficiency in sarcolemmal fibroblasts in close proximity to the muscle fibres with a heterozygous Sod2 deficiency, showed a severe muscle atrophy in terms of decreased fibre diameter, increased variability and a significantly decreased muscle mass to body weight ratio, accepted hallmarks of muscle atrophy in intrinsically aged rodents and humans (Larsson, 1978; Usuki et al., 2004). In contrast, mice with a heterozygous Sod2 deficiency throughout all organs including sarcolemmal fibroblasts and muscle fibres themselves did not show any muscle atrophy and could not be distinguished from muscle fibres of Sod2-competent littermate controls. Therefore, given that homozygosity for the Sod2 deficiency in fibroblasts represents the only difference between Sod2-deficient heterozygous and mutant mice, it is tempting to suggest a scenario where the complete Sod2 deficiency in fibroblasts of the sarcolemmal connective tissue, which ensheat each muscle fibre, may significantly contribute to the observed severe muscle atrophy. The muscle fatigability and the observed pathological motor phenotypes in mutant mice as suggested by the data from rotarod and tail suspension test, mostly likely originate from the impairment of the muscle architecture. These observed changes have earlier been described in old rodents (Usuki et al., 2004) and human muscle (Larsson, 1978). However, we cannot exclude that disruption of the neuromuscular junction may have some influence on the observed motor phenotype of the mutant mice as previously published in mice deficient for the Cu, Zn superoxide dismutase (Jang et al., 2010). Serial sections of the spinal cord with α-motor neurons and neurons from the cerebral cortex did not reveal any morphological changes (data not shown). This, in conjunction with the observed severe muscle atrophy, provides evidence that the mutant mice rather suffer from a progressive aging-related muscle atrophy.

This is further supported by preliminary data on Tamoxifen inducible connective tissue-specific Sod2 deletion (data not shown), where the Sod2 deficiency exclusively occurred in fibroblasts, but not in muscle fibres. Several reasons may contribute to the effects of Sod2-deficient fibroblasts on the development of severe muscle atrophy observed in the mutant mice. Accordingly, Sod2−/− fibroblasts from mutant mice may affect skeletal muscle cells by changes in the quality and quantity of extracellular matrix proteins, soluble paracrine factors including catabolic cytokines, growth factors (Coppe et al., 2008) or long-lived toxic lipid peroxidation products due to enhanced superoxide anion generation. Given the fact that fibroblasts are in close proximity to skeletal muscle cells, their impact on the microenvironment of skeletal muscle cells is critical. Age-related oxidative damage, lipid peroxidation and mitochondrial protein damage in murine skeletal muscle have previously been reported (Lass et al., 1998). Irrespective of the underlying mechanisms, we have shown that Sod2 homozygous deficiency in sarcolemmal fibroblasts in close proximity to muscle fibres can tilt the biological outcome to muscle atrophy. This may be of relevance to human aging and aging-related pathologies as, similar to the here reported mutant model, senescent human fibroblasts reveal increased superoxide anion concentrations (Fig. S7, Supporting information).

Similarly, osteoporosis and skin atrophy belong to universal aging-related phenotypes in mice and humans leading to major disability in the elderly. We have found a severe decrease in mineral density in femur and vertebrae bone in mutant mice (150 days) and intrinsically aged mice (900 days), but not in Sod2 heterozygous-deficient and Sod2-competent control mice at an age of 150 days. Consistent with our data, a similar decrease in mineral density in both vertebrae and femur bone was earlier reported in intrinsically aged mice (Kalu, 1995). Kyphosis (curvature of the spine), as observed in the mutant and intrinsically aged mice, may be linked to osteoporosis with loss of trabecular bone in vertebral bodies of the middorsal spine or to muscle atrophy (Gerhard & Kasales, 2003) or a combination thereof, as most likely in our model. In humans, osteoporosis causes kyphosis through the deformity of the vertebral bodies (Cummings & Melton, 2002). The current concept suggests that osteoporosis represents a continuum with multiple underlying mechanisms contributing to loss of bone mass and mineral density and overall microarchitectural deterioration of bones (Ferrari et al., 2004; Tasker et al., 2004; Bodine et al., 2005; Horowitz & Lorenzo, 2007; Perrini et al., 2010). These factors are causally related to increased risk of falls, contributing to high incidence of fragility fractures in osteoporotic patients (Raisz, 2005; Ralston & de Crombrugghe, 2006).

Similar to the bone and muscle atrophy, also the skin is atrophic in mutant mice and intrinsically aged mice. A comparable skin atrophy in intrinsically aged mice and humans has repeatedly been reported (Oikarinen, 1994; Scharffetter-Kochanek et al., 1997; Kaya et al., 2006). As expected, dermal fibroblasts from mutant mice revealed an increase in superoxide anion concentration and a decrease in hydrogen peroxide concentration in the mitochondria. This led to the development of oxidative damage and finally to the degeneration of mitochondria. Both collagen deposition and fibril thickness are severely reduced in the dermis of the skin. Collagen type I, the most abundant matrix protein in the skin, was severely reduced at the protein level in the skin of mutant mice. As the specific collagen type I mRNA concentrations are similar in mutant mice compared with control mice, either increased collagen degradation by matrix-metalloproteases or reduced biosynthesis at a post-transcriptional level are responsible for the observed reduction in collagen deposition in the skin of mutant mice. Both reduction in collagen synthesis and induction and activation of matrix-degrading metalloproteases are critical in human skin aging (Ma et al., 2001; Shin et al., 2005).

The connective tissue-specific Sod2-deficient model sharing murine and human aging-related pathologies is particularly interesting to unravel the underlying mechanisms. Due to Sod2 deficiency and a lack of superoxide anion detoxification, the concentration of superoxide anion increased in mutant fibroblasts as shown herein. Superoxide anions and intermediates thereof like peroxynitrite can severely damage macromolecules, and most importantly, they can induce DNA double strand breaks and intercrosslinks in the DNA (Szabo et al., 2007). Such lesions usually promote apoptosis or induce a senescence programme with permanent growth arrest contributing to organ atrophy and most likely to organismal aging (Niedernhofer et al., 2006; van der Pluijm et al., 2007; Garinis, 2008; Garinis et al., 2008). We did not find induced apoptosis in the connective tissue of Sod2 mutant mice compared with Sod2-competent wild type control. Notably, we found that the cell cycle inhibitor p16INK4a time-dependently increased in the dermal and muscle connective tissue of mutants, while there was significantly less p16INK4a expression in Sod2 heterozygous-deficient and Sod2-competent mice. p16INK4a constitutes a major player in the instalment of a ROS/DNA damage-induced senescence programme (Dai & Enders, 2000; Kim & Sharpless, 2006). In this regard, p16INK4a functions as an inhibitor of CDK4 and CDK6, the D-type cyclin-dependent kinases that initiate the phosphorylation of the retinoblastoma tumour suppression protein RB. Thus, p16INK4a has the capacity to arrest cells in the G1 phase of the cell cycle implementing irreversible growth arrest. The fact that the expression of the p16INK4a increases markedly with aging in many tissues of rodents and humans (Krishnamurthy et al., 2004, 2006; Molofsky et al., 2006; Ressler et al., 2006) very much suggests that our mutant mouse model distinctly shares mechanisms underlying intrinsic aging.

Our model with preferential oxidative damage of fibroblasts residing in the connective tissue of different organs may very well reflect the situation in patients subjected to ROS and DNA damage-inducing chemotherapy or radiotherapy (Grillari et al., 2007). While tissues with highly proliferative cells of epithelial origin mainly undergo apoptosis, connective tissue resident fibroblasts rather undergo a senescence programme with a metabolically changed phenotype, loss of tissue homoeostasis and aging. In fact, long-term survivors of chemotherapy and radiotherapy show evidence for premature aging (Grillari et al., 2007).

The herein reported mutant mice sharing universal murine and human age-related changes are particularly interesting to unravel the underlying mechanisms and to identify strategies to counteract accelerated aging.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

Generation of mutant mice

Mice carrying the homozygous floxed Sod2 allele (Sod2f/f; B6;129.Sod2tm1Ksk) (Strassburger et al., 2005) were bred with Col1α2-Cre+ transgenic mice (B6;129.TgN(Col-Iα2) (Florin et al., 2004) to generate Col1α2-Cre+Sod2+/f mice. In a second mating, this mouse (Col1α2-Cre+Sod2+/f) line was bred to homozygous Sod2f/f mice to obtain connective tissue-specific Sod2-deficient offspring. The resulting transgenic mice (B6;129.Sod2tm1.1Ksk) were of a mixed genetic background (Sv129 × C57BL/6J). All mice were maintained under specific pathogen-free conditions and only late dying mutants (> 100 days) were used for our analyses. All experiments were performed in compliance with the German Law for Welfare of Laboratory Animals.

Genotyping by Southern blot and PCR analysis

For Southern blot analysis, genomic DNA of tail biopsies, skin, muscle, kidney, spleen, liver, dispase separated epidermis and dermis (Sigma, Deisenhofen, Germany) and from macrophages and B-cells was isolated with Easy-DNA™ (Invitrogen, Karlsruhe, Germany). DNA (7.5 μg) was digested with BamHI yielding 2.6 kb fragments for the floxed Sod2 allele, 7.8 kb for the deleted Sod2 locus and 8.5 kb for the Sod2 wild type allele. Bands were detected using a 750 bp HindIII/BamHI fragment of intron2 of Sod2 allele as a probe. Primer sequences for the PCR-based genotyping for Sod2 are as follows: P1, 5′-TTA GTA CAT CAT GGC TTG ACG A-3′; P2, 5′-AGG AAA TGC TTT CCC AAC TG-3′ and P3, 5′-GAA AGT CAC CTC CAC ACA CAG A-3′. Primer sequences for the PCR-based genotyping for Cre-recombinase are Cre1, 5′-TTA GCA CCA CGG CAG CAG GAG GTT-3′; Cre2, 5′-CAG GCC AGA TCT CCT GTG CAG CAT-3′. Red-Taq® (Sigma) was used as Taq-Polymerase. PCR samples were subjected to 30 s 94 °C as pre-denaturation, 34 cycles [30 s as annealing (60 °C), 30 s as extension time (72 °C)] and 10 min, 72 °C for final extension.

Reverse transcriptase polymerase chain reaction (RT-PCR)

For RT-PCR, total RNA was isolated from mouse skin using the Trizol method, as described by the manufacturer (Life Technologies, Inc., Rockville, MD, USA). One μg RNA per sample was reverse transcribed using a first strand cDNA reverse transcription-polymerase chain reaction (RT-PCR) synthesis kit (MBI Fermentas, Vilnius, Lithuania). PCR samples were subjected to pre-denaturation (3 min at 94 °C), 21 cycles (30 s at 94 °C, 30 s at 60 °C, 1 min at 72 °C) for GAPDH and 23 cycles for pro-collagen(I) (45 s at 94 °C, 45 s at 60 °C, 1 min at 72 °C) and final extension (10 min at 72 °C). Primers for mouse GAPDH were used as an internal control (forward, 5′- AAG ATT GTC AGC AAT GCA TCC-3′; reverse, 5′- GCC CCT CCT GTT ATT ATG G – 3′) and mouse pro-collagen (I) (forward, 5′-TCG TGA CCG TGA CCT TGC G-3′; reverse, 5′-GAG GCA CAG ACG GCT GAG TAG-3′). PCR products were analysed on a 2.0% agarose gel, followed by quantification by densitometry analysis using TINA2.0 software (TINA; Raytest Isotopenmessgeräte, Straubenhardt, Germany).

Protein lysates and Western blot analysis

Mouse skin was homogenized in ice-cold lysis buffer (50 mm Tris–HCl, pH 7.4, 150 mm NaCl, 2 mm ethylenediamine tetraacetic acid (EDTA), 5 mm phenylmethanesulfonyl fluoride (PMSF), and 1 mm dithiothreitol (DTT), 1% Triton X-100) with freshly added protease inhibitor cocktail (Roche, Indianapolis, IN, USA). The homogenate was then centrifuged at 15 000 g for 30 min at 4 °C, supernatant was collected and aliquots were stored at −70 °C. The protein content in the lysate was determined by the Bradford assay. Equal amounts of protein were subjected to 10% and 12% Tris–Glycine SDS–PAGE gels (as applied), and then transferred onto PVDF membrane. The blot was subsequently blocked with blocking buffer (5% nonfat dry milk, 1% Tween-20 in 20 mm TBS, pH 7.6) for 1 h at room temperature and incubated with monoclonal anti-pro-collagen type I amino-terminal extension peptide antibody (1:1000, SP1.D8; Developmental studies Hybridoma Bank, Iowa, USA), and polyclonal anti-β-actin antibody (1:1000; Chemicon, Temecula, CA, USA), overnight at 4 °C. Thereafter, incubation with a secondary antibody conjugated to horseradish peroxidase (Amersham Life Science, Buckinghamshire, UK) for 1 h at room temperature and detection by ECL system (Amersham, Life Science, Buckinghamshire, UK) was performed. Western blots were also incubated with a primary antibody against Sod2 (FL-222, 1:200; Santa Cruz, CA, USA), actin (CP01, 1:10000; Calbiochem, Gibbstown, NJ, USA), p16INK4a (SC-1661, 1:1000; Santa Cruz) and cleaved (active) caspase-3 (#9664, 1:1000; Cell Signaling Technologies, Danvers, MA, USA) overnight at 4 °C. The secondary antibody goat anti-rabbit conjugated to horseradish peroxidase (for Sod2; IgG, 1:200; Santa Cruz; for actin; IgM, 1:10000; Calbiochem), goat anti-mouse conjugated to horseradish peroxidase (for p16INK4a, IgG, 1:10000; Jackson ImmunoResearch Laboratories, Weston, PA, USA) and goat anti-rabbit conjugated to horseradish peroxidase (for caspase-3, Jackson, IgG, 1:10000; ImmunoResearch Laboratories) was incubated for 1 h at room temperature, followed by detection with ECL system (Cell Signaling Technology, Danvers, MA, USA). The detection of carbonylated proteins was performed by OxyblotTM analysis (S7150; Chemicon) using standard protocols. In brief, protein lysates were derivatized with 2,4-dinitrophenyl hydrazine (DNPH) to add dinitrophenol residue (DNP) into the side chain carbonyl group. As a positive control, oxidized bovine serum albumin (BSA) and as a negative control, reduced BSA were used. Non-derivatized protein lysates were also included to assess the specificity of anti-DNP antibody. All samples were subjected to 10% SDS–PAGE and subsequently transferred onto nitrocellulose membrane and probed with anti-DNP antibody, following incubation with HRP-conjugated secondary antibody. The membrane was then treated with the ECL reagent to visualize the bands.

Fixation procedure for transmission electron microscopy

Mice were anaesthetized subcutaneously by means of Ketamine (100 mg kg−1) and Xylazine (5 mg kg−1) and transcardially perfused with a 2% paraformaldehyde, 2% glutardialdehyde, 0.1 m cacodylate solution buffered at pH 7.35 for 10 min. Thereafter, skin samples were immediately removed and post-fixed for further 6–8 h in the same fixative. The samples were then washed three times with 0.1 m cacodylate buffer and stored in this buffer at 4 °C until plastic embedding.

Transmission electron microscopy

Fixed skin samples were cut into small pieces containing both epidermis and dermis. The samples were postfixed with 2% osmium tetraoxide in 0.1 m cacodylate buffer for 2 h at 4 °C. After thorough rinsing in 0.1 m cacodylate buffer for 10 min, slices were dehydrated in a graded ethanol series, and infiltrated with and embedded in araldite. Sections of plastic-embedded specimens were cut with a glass knife for thin sections and a diamond knife for ultra-thin sections on a Reichert ultra-microtome. The 0.5-μm-thin slices were stained with methylene blue and investigated using a Zeiss Axiophot (Zeiss, Oberkochen, Germany). Further contrasting with uranyl acetate–lead citrate was performed and ultrathin 60 nm sections were examined using an electron microscope (902A; Fa. Zeiss).

Histology

Tissues were dissected from the mice and either deep-frozen in O.C.T.TM (Tissue-TEK, Torrance, CA, USA) or fixed overnight in 4% paraformaldehyde in PBS at 4 °C for paraffin embedding. Paraffin sections of 5 μm thickness were used for haematoxylin-eosin and Masson’s trichrome staining.

Immunostaining

Cryosections were fixed in 70% ethanol for 10 min at −20 °C and thereafter treated with 0.1% Triton X-100 for 10 min at room temperature. Blocking was performed in 5% BSA for 1 h at 37 °C. Anti-p16INK4a (SC-1661, 1:200; Santa Cruz), Er-TR7 (NB100-64932, 1:200, Novus Biologicals, Cambridge, UK), Sod2 (06-984, 1:200; Millipore, Schwalbach, Germany) and cleaved (active) caspase-3 (#9664, 1:200; Cell Signaling Technologies) as primary antibody were used overnight at 4 °C. Incubation with the secondary antibody Alexa 488 goat anti-rat (for Er-TR7, 1:200), Alexa 555 goat anti-mouse (for p16INK4a, 1:200) and Alexa 555 goat anti-rabbit (for Sod2 and caspase-3 1:500) (Invitrogen) was performed for 45 min at room temperature. Isotype control sections were incubated with respective IgG as primary antibodies.

Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining

For in situ detection of apoptosis, TUNEL staining was performed using a commercially available kit (In Situ Cell Death Detection Kit, POD, Roche, Manheim, Germany) according to the manufacturer’s protocol. Briefly, paraffin-embedded skin sections were prepared on glass slides, deparaffinized, rehydrated and treated with Proteinase K. Skin sections were then incubated with labelling solution containing recombinant Terminal deoxynucleotidyl Transferase (TdT) and fluorescein-labelled nucleotides. After washing, sections were counterstained with DAPI and mounted in fluorescence mounting medium (DAKO, Hamburg, Germany) prior to examination by fluorescence microscope (Axio Imager M1; Zeiss) equipped with a green filter.

LacZ reporter mice and X-gal staining

Col1α2-Cre+ transgenic mice (B6;129.TgN(Col-Iα2) were bred with ROSA26-LacZ reporter mice (Florin et al., 2004) and the resulting Cre recombinase positive pups (Col1α2-Cre+; ROSA26-LacZf/+) were used to study the site/region of Cre expression. The LacZ gene in ROSA26 locus was kept inactive by placing one ‘STOP’ codon (flanked by loxP sites on both ends) between the ubiquitous promoter (CMV) and LacZ coding region. Activation of Cre recombinase deleted the ‘STOP’ codon and thus initiated the expression of β-galactosidase from LacZ gene. Therefore, β-galactosidase would only be expressed in those cells, where cell-specific promoter driven Cre recombinase expression occurred. The mice were first perfused with phosphate-buffered saline (pH 7.4) and later by 4% paraformaldehyde (in phosphate buffered saline, pH 7.4). Different organs, including the skin, intestine, kidney, skeletal muscle, were dissected out and washed with phosphate-buffered saline (pH 7.4) and subjected to mild fixation with 4% paraformaldehyde. The mildly fixed organs were placed in X-gal staining solution (6 mm potassium ferricynide, 6 mm potassium ferrocynide, 2 mm Magnesium chloride, 2.5 mm X-Gal, 0.02% Igepal in phosphate-buffered saline, pH 7.4) and incubated overnight in 37 °C. The stained organs were washed briefly with phosphate-buffered saline (pH 7.4), again fixed with 4% paraformaldehyde and finally processed for paraffin embedding. 5 μm thick paraffin sections were prepared and after counter staining with Nuclear Fast Red (Sigma), the sections were mounted and examined under microscope (Axio Imager M1; Zeiss).

Tail suspension test

This test was performed to assess the motor coordination in mice. Physiologically, when mice are suspended by the tail, they prepare safe landing by actively spreading their hind limbs. Mice were suspended by their tails for 10 s and looked for clasping behaviour of the hind limbs and also the duration of clasping was noted. Hind limb clasping as observed in the mutant mice is indicative of a motor phenotype.

Rotarod

The rotarod test was used to assess motor coordination and balance in rodents (Brooks & Faulkner, 1988). Mice have to keep their balance on a rotating rod (Rotarod Version 1.2.0. MED Associates Inc., Vermont, VT, USA). The time (latency) it takes the mouse to fall off the rod rotating under continuous acceleration (from 4 to 40 rpm in 300 s) was measured. Each mouse had to perform three trials separated by 15 min inter-trial intervals. These trials were averaged.

Determination of muscle fibre diameter

The quadriceps muscle was prepared and subjected to cryopreservation. Cryosections of 10 μm thickness were stained with haematoxylin and eosin (HE). To determine the muscle fibre diameters (= 5 for each group), the standard method for myological diagnostics, the ‘lesser fibre diameter’ was taken as described previously (Brooke & Engel, 1969). More than 1000 fibres per muscle per mouse with 5000 fibres per genotype were measured at a 20-fold magnification using an Olympus BX51TF microscope equipped with a CCD camera (Soft Imaging System CC12; Olympus, Hamburg Germany) and the CellF imaging software (Olympus). The fibre measurements from each muscle section were pooled per animal for statistical analysis.

Murine dermal fibroblasts cultivation

New born mice were sacrificed, skin was removed, cut into small pieces digested in a trypsin solution. Single-cell suspension was centrifuged for 5 min at 800 rpm (Heraeus Multifuge 3S-R; Thermo Electron Corporation, Karlsruhe, Germany). The pellet was resuspended in Dulbecco’s modified Eagle’s medium (Life Technologies, Inc., Eggenstein, Germany) supplemented with 10% foetal calf serum (Biochrom, Berlin, Germany), glutamine (2 mm), penicillin (100 U mL−1) and streptomycin (50 μg mL−1) at 37 °C in a humidified atmosphere. Fibroblasts were cultivated with 3% oxygen and passaged at a 1:2 to 1:3 dilution every 3 days.

Isolation of B-cells

A single cell suspension was prepared from the spleen and B-cells were isolated by labelling with CD19 PE and anti PE Micro Beads (Miltenyi Biotec, Bergisch Galdbach, Germany) for positive selection according to the manufacturer’s protocol. Enrichment of > 86% was confirmed by flow cytometry (FACScan; BD Biosciences, Heidelberg, Germany) using an antibody directed against CD45R/B220 PerCP (clone RA3-6B2; BD Biosciences, Pharmingen, Heidelberg, Germany).

Bone marrow-derived macrophages

Bone marrow-derived macrophages (MF) were obtained from femurs as described previously (Sunderkotter et al., 1993). Briefly, bone marrow was flushed through with Dulbecco’s minimum essential medium (DMEM) (Biochrom). Following an osmotic-shock, 3 × 106 cells per 10 mL medium were grown in DMEM supplemented with 10% heat-inactivated FCS (PAA Laboratories, Pasching, Austria), 10% conditioned supernatant from L929 cells, 2%l-Glutamine, 1% penicillin/streptomycin (stock 10 000 U per 10 000 μg mL−1) and 1% non-essential amino acids (Biochrom) in Petri dishes (NUNC, Roskilde, Denmark) in a 7% CO2 atmosphere at 37 °C. After 6 days, mature macrophages were harvested for DNA extraction.

Bone mineral density (BMD)

High-resolution radiographs were performed on the femurs at 25 kV per 2.2 mA (43805 NX-Ray-System; Hewlett-Packard, Palo Alto, CA, USA) for the visualization of BMD. A quantitative determination was performed using μCT (Fan Beam μ-Scope; Stratec Medical, Pforzheim, Germany). Ten scans with a resolution of 30 μm and with a distance of 30 μm from each other were performed in the proximal right femur (centre of the femoral head). The average BMD was determined for each bone by calculating the mean values of the BMD from 10 scans.

Metabolic cage studies

To assess whether there is any difference in daily food and water consumption among different genotypes of mice, a controlled metabolic cage study was conducted. In this study, mice from each genotype, including mutant, Sod2 heterozygous-deficient and Sod2-competent control mice were caged individually in metabolic chambers (Scanbur, Karlslunde, Denmark) for 3 days under standard conditions. Prior to the actual study, the mice were kept in metabolic cages to adapt themselves. Thereafter, they were kept for further 48 h for the actual analysis. Mice had free access to water and food. The food and water in each cage were changed every day and the remaining food weight and water volume were measured and subtracted from initial values to assess the amount of food and water consumed per 24 h.

Intracellular superoxide anion concentration

Superoxide anions were quantified in fibroblasts using indirect fluorescence with MitoSOX (MitoSOX™ Cat. No. M36008; Invitrogen). MitoSOX is mitochondrial-specific and reaction of superoxide anion in the mitochondria with MitoSOX results in fluorescence emission at 510 nm if excited at 396 nm (Robinson et al., 2006). A concentration dependent curve of MitoSOX fluorescence with increasing concentrations of rotenone in Sod2-competent fibroblasts served as a reference. Rotenone is a specific inhibitor of complex I and, thus, causes increased generation of superoxide anion in the mitochondria (Kushnareva et al., 2002). In brief, 1 × 105 fibroblasts were seeded into six well plates and stained with 5 μm MitoSOX for 30 min in the presence and absence of rotenone. After incubation, fibroblasts were washed with HBSS, dislodged by accutase (GmbH; PAA Laboratories, Cölbe, Germany), suspended in 200 μL of HBSS and subjected to fluorescence measurement using a Perkin-Elmer fluorometer for murine dermal fibroblasts (LB 50; Perkin Elmer, Waltham, Massachusetts, USA) or FACSCanto II (BD, Franklin Lakes, NJ, USA) for human dermal fibroblasts.

Detection of mitochondrial H2O2 using adenoviral transduction of the genetically encoded fluorescence indicator

The plasmid construct pHyPer-dMito coding for a derivative of the H2O2-specific OxyR protein were purchased from Evrogen (Cat.# FP941; Moscow, Russia) (Belousov et al., 2006). The expression cassette was cloned into the adenovirus vector plasmid backbone pGS66 for generation of a replication-deficient E1-deleted adenovirus vector by standard procedures (Schiedner et al., 2000). Cloning details can be obtained upon request. In brief, the vectors were serially amplified on N52.E6 cells and purified by double CsCl-banding. The titres of the vector preparations were determined by a vector genome-based slot-blot procedure (Kreppel et al., 2002) and genome integrity was confirmed by restriction analysis. The replication-deficient adenoviral vector AdMito-HyPer expresses the sensor construct targeted to mitochondria due to the presence of a mitochondrial signal peptide sequence. 1 × 106Sod2-deficient mutant fibroblasts or control fibroblasts were transduced at a multiplicity of infection of 500. Twenty-four hours posttransduction, the mean fluorescence intensity of gated events was measured at 488 nm for FITC and 405 nm for AmCyan by flow cytometry (FACS Canto II; BD Biosciences) and evaluated considering both the autofluorescence (AF) and the number of events in each gate (Ev) as given in the formula below. Transduced fibroblasts were also analysed with a fluorescence microscope (Axiovert 200M; Zeiss).

Statistical calculations

The exact two-sample Wilcoxon test or one-way anova were used to compare independent measurements between control, heterozygous, mutant and old mice. For repeated comparison at different time points, a correction of the P-values according to Bonferroni (i.e. multiplication of each P-value by the number of time points) was performed. For comparisons of survival times, the log-rank test was applied, optionally stratified for gender. All tests were two-sided. Calculations were performed using the sas software, version 9.1 (SAS Institute Inc., Cary, NC, USA).

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Acknowledgments

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

AL, SI, MB and KS-K are supported by the German Research Foundation within the Clinical Research Group KFO142: ‘Cellular and Molecular Mechanisms of Ageing – From Mechanisms to Clinical Perspectives’. MW and KS-K are supported by the European Union through the Integrated Project PROTEOMAGE (FP6-518230). AS and KS-K received a grant from the German Research Foundation within the Collaborative Research Centre SFB 497: ‘Signals and Signal Processing during Cellular Differentiation’. JHC is supported by the Korea Science and Engineering Foundation (KOSEF) through the Centre for Aging and Apoptosis Research at Seoul National University (R11-2002-097-06001-0). We are grateful to HJ Fehling for providing the RFP indicator mice and to P. Angel for providing the Col1α2-Cre mice.

Author contributions

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

Nicolai Treiber generated and characterized the phenotypes of mutant mice. Pallab Maity, Karmveer Singh characterized the phenotype of the mutant mice and were involved in data collection, analysis, interpretation and assembly. Matthias Kohn analysed the genetic basis of the mutant mice. Florentina Ferchiu and Lea Sante contributed data on enhanced superoxide anion levels in senescent human fibroblasts. Meinhard Wlaschek, Wilhelm Bloch, Sebastian Iben, Anca Sindrilaru, Mark Berneburg, Albert Ludolph contributed to the concept and design and approval of the manuscript. Alexander F. Keist. Sebastian Frese, Michael Ohnmacht, Josef Högel, Jin Ho Chung, Min Jung Lee, York Karmenich, Kerstin Braunstein contributed to data collection, and analysis. Lutz Claes, Anita Ignatius, Thorsten Nikolaus, Anne-Dorte Sperfeld added knowledge on translation of our murine data to human diseases. Karlis Briviba, Florian Kreppel and Stefan Kochanek provided the techniques for mitochondrial-specific H2O2 quantification, data analysis and interpretation. Karin Scharffetter-Kochanek is responsible for the concept, design, collection and assembly of data, data interpretation, manuscript writing and final approval of the manuscript.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgments
  8. Author contributions
  9. References
  10. Supporting Information

Fig. S1 Generation of the mutant mice with severe aging phenotype.

Fig. S2 Analysis of feeding and drinking pattern of early and late dying mutant mice.

Fig. S3 Analysis of apoptosis in the hearts of early and late dying mutant mice.

Fig. S4 Analysis of vertical diameter or thickness of different layers of skin.

Fig. S5 Specificity of Col1α2-Cre using ROSA26 LacZ reporter mice.

Fig. S6 Fibroblast specific deletion of Sod2 in the skin and skeletal muscle.

Fig. S7 Enhanced superoxide anion generation in replicative senescent human dermal fibroblasts.

Data S1 Generation of the mutant phenotype.

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