Pleiotropic age-dependent effects of mitochondrial dysfunction on epidermal stem cells
Edited by Philip C. Hanawalt, Stanford University, Stanford, CA, and approved July 15, 2015 (received for review March 21, 2015)
Significance
Mitochondrial damage can accelerate features of aging, including impaired tissue regeneration, but little is known about how aging and this damage interact to impair tissue renewal. We show that mitochondrial oxidative stress in the epidermis alters wound healing depending on age. Epidermal mitochondrial damage accelerated wound closure in young mice; however, in older mice, this damage limited epidermal cell proliferation and reduced epidermal stem cell numbers, leading to delayed wound closure. Our findings uncover a surprising beneficial effect of mitochondrial dysfunction at young age (accelerated wound closure), and a potential mechanism for the reduced epidermal regeneration at older ages (stem cell depletion).
Abstract
Tissue homeostasis declines with age partly because stem/progenitor cells fail to self-renew or differentiate. Because mitochondrial damage can accelerate aging, we tested the hypothesis that mitochondrial dysfunction impairs stem cell renewal or function. We developed a mouse model, Tg(KRT14-cre/Esr1)20Efu/J × Sod2tm1Smel, that generates mitochondrial oxidative stress in keratin 14-expressing epidermal stem/progenitor cells in a temporally controlled manner owing to deletion of Sod2, a nuclear gene that encodes the mitochondrial antioxidant enzyme superoxide dismutase 2 (Sod2). Epidermal Sod2 loss induced cellular senescence, which irreversibly arrested proliferation in a fraction of keratinocytes. Surprisingly, in young mice, Sod2 deficiency accelerated wound closure, increasing epidermal differentiation and reepithelialization, despite the reduced proliferation. In contrast, at older ages, Sod2 deficiency delayed wound closure and reduced epidermal thickness, accompanied by epidermal stem cell exhaustion. In young mice, Sod2 deficiency accelerated epidermal thinning in response to the tumor promoter 12-O-tetradecanoylphorbol-13-acetate, phenocopying the reduced regeneration of older Sod2-deficient skin. Our results show a surprising beneficial effect of mitochondrial dysfunction at young ages, provide a potential mechanism for the decline in epidermal regeneration at older ages, and identify a previously unidentified age-dependent role for mitochondria in skin quality and wound closure.
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Stem and progenitor cells are crucial for tissue homeostasis, repair, and regeneration. In response to injury, they proliferate and differentiate to replace damaged or dysfunctional cells (1, 2). In the skin, epidermal basal cells differentiate to form distinct epidermal layers: the stratum basale (SB), stratum spinosum (SS), stratum granulosum (SG), and stratum corneum (SC) (3). In the SB layer, epidermal basal cells are identified by nuclei that stain strongly with hematoxylin and eosin (H&E). These cells differentiate to form the SS layer, composed mainly of cells with lightly stained nuclei. SS cells differentiate into the SG layer, identified by cells with prominent cytoplasmic granules, which terminally differentiate to form the SC layer containing acidophilic anucleated cells.
Tissue homeostasis declines with age partly because stem/progenitor cells fail to self-renew or differentiate (4). Oxidative damage can contribute to this decline in compartments such as the hematopoietic system (5, 6). Aging is caused by intrinsic and extrinsic factors that cooperate to drive aging phenotypes (7). Mitochondrial dysfunction has been suggested to play a major role in intrinsic aging (8). Furthermore, mitochondrial damage is associated with extrinsic aging, particularly ultraviolet (UV) radiation-induced photoaging in the skin (9). Thus, mitochondrial damage may be a common link between intrinsic and extrinsic aging.
Mitochondrial stress can decrease life span and health span. Superoxide dismutase 2 (SOD2) scavenges mitochondrial superoxide to protect against oxidative damage. Sod2 deficiency decreases life span in several species. Mice with constitutive Sod2 deficiency are neonatally lethal on multiple genetic backgrounds, presenting with neurodegeneration, spongiform encephalopathy, cardiomyopathy, hepatic fat accumulation, and failure to thrive; cells from these mice exhibit impaired spare respiratory capacity, genomic instability, and mitochondrial functional defects (10–17). Sod2−/− cells also have a reduced proliferative capacity (17), consistent with the finding that constitutive Sod2 deficiency induces cellular senescence, a tumor-suppressive mechanism that irreversibly arrests cell proliferation (18), in mouse skin (19). Conversely, overexpression of mitochondrial antioxidants can partly rescue age-related pathologies (14, 20), increase organismal life span (21), and prolong stem cell replicative life span (22).
Interestingly, some studies have suggested that mild mitochondrial stress can be beneficial (23). Here we show that mitochondrial stress owing to Sod2 deficiency in epidermal cells can have positive or negative effects on skin regeneration, and that these effects depend on age. We show that epidermal Sod2 deficiency induces cellular senescence, which reduces proliferative capacity in the skin but stimulates the differentiation of epidermal stem/progenitor cells. This stimulation accelerates wound closure in young mice, but the proliferative decline drains stem cell pools with aging and retards wound closure. Our findings extend the concept of antagonistic pleiotropy, which stipulates that gene action can be beneficial at young ages but deleterious at older ages, to mitochondrial function in the skin.
Results
Mice with a Keratinocyte-Specific Mitochondrial Defect.
Because constitutive Sod2− deficiency is neonatal lethal (12), it is not possible to study age-specific effects in these mice. Consequently, we constructed an inducible, temporally regulated, tissue-specific mouse model of Sod2 deficiency. We created mice carrying a Sod2 gene into which we inserted LoxP sites (Sod2tm1Smel) into sequences flanking a region required for catalytic activity. We then crossed these animals to mice carrying a tamoxifen (TAM)-activated Cre recombinase (Cre-ERT) under control of the keratinocyte-specific keratin 14 promoter. This mouse, Tg(KRT14-cre/Esr1)20Efu/J x Sod2tm1Smel, herein designated K14S, allows deletion of critical Sod2 sequences in keratinocytes after TAM treatment. TAM, but not estrogen, causes nuclear translocation of Cre-ERT, a fusion protein comprised of Cre and a mutant estrogen receptor ligand-binding domain, thus allowing Cre to excise sequences between the LoxP sites (Fig. S1A). Because the K14 promoter is active primarily in epidermal stem/progenitor cells (24, 25), TAM deletes Sod2 mainly in epidermal, but not dermal, cells (Fig. S1 B and C). As expected, TAM deleted Sod2 in dorsal skin and skin-containing tails and toes, but not in the heart, liver, intestine, or lungs, of K14S mice (Fig. S1D). TAM did not delete Sod2 in Sod2tm1Smel mice (designated S), which lack Cre recombinase (Fig. S1D).
Fig. S1.
We measured the copy numbers of recombined and total Sod2 alleles by quantitative PCR (qPCR) (Fig. S1E). After TAM treatment, >92% of Sod2 genes showed a deletion in the epidermis, but not the dermis, of K14S mice. No recombination occurred in the epidermis or dermis of Tg(KRT14-cre/Esr1)20Efu/J × B6-Gt(ROSA)26Sortm1Sor/J mice (designated K14R), which lack LoxP sites. Accordingly, qPCR showed that Sod2 mRNA levels were substantially lower in the epidermis (Fig. S1F), but not the dermis (Fig. S1G), of TAM-treated, but not vehicle-treated, K14S mice. As expected, Sod2 mRNA levels were comparably high in the epidermis and dermis of vehicle- and TAM-treated K14R mice. This finding indicates that TAM reduced Sod2 expression in epidermal cells of K14S mice, confirming successful generation of an inducible keratinocyte-specific Sod2-deficient mouse and overcoming the impediment of neonatal lethality caused by constitutive Sod2 deficiency.
To determine the SOD2 expression pattern in skin, we used immunohistochemistry on whole mounts of tail epidermis from vehicle- or TAM-treated K14S mice, counterstaining for CD49f, a keratinocyte marker to visualize epidermal compartments. SOD2 was detectable in the sebaceous glands and epidermal layers, with highest expression in the middle of the hair follicle or isthmus region (Fig. 1A) in which stem/progenitor cells reside (26). Cells in the bulb area expressed lower SOD2 levels. TAM reduced epidermal SOD2 levels markedly, consistent with Sod2 deletion in the targeted tissue.
Fig. 1.
Constitutive Sod2 loss in mice severely reduces mitochondrial complex II activity, but not complex IV activity, and decreases mitochondrial spare respiratory capacity without altering the basal respiration rate (12, 19). To determine whether keratinocyte-specific Sod2 loss has similar effects, we stained K14S skin for succinate dehydrogenase (SDH) (complex II) and cytochrome c oxidase (COX) (complex IV) activities. TAM-treated, but not vehicle-treated, K14S mice had significantly less SDH activity in the hair follicles, but not in the underlying skeletal muscle (Fig. 1B and Fig. S1I). As expected (19), mitochondrial complex IV (COX) activity was similar in the TAM- and vehicle-treated K14S mice (Fig. S1 H and I). These results confirm that the mitochondrial dysfunction was confined to epidermal keratinocytes in TAM-treated K14S mice.
Epidermal Sod2 deficiency did not significantly increase morbidity in all cohorts tested up to ∼16 mo after TAM treatment of 4-mo-old mice. Thus, epidermal deletion of Sod2 did not noticeably compromise the health of mice up to 20 mo of age.
Epidermal Sod2 Deficiency Accelerates Wound Closure in Young Mice.
To determine the consequences of epidermal Sod2 deficiency, we assessed vehicle- and TAM-treated K14S mice for wound healing after a skin biopsy. We treated 4-mo-old mice, and assessed wound healing 4 mo later (at 8 mo of age). Surprisingly, TAM-treated, but not vehicle-treated, mice showed accelerated wound closure (Fig. 2A). TAM-treated mice also had an unusually thick layer of epidermal cells near the wound edges (Fig. 2B), but no difference in collagen formation (Fig. S2A), suggesting rapid epithelialization. Indeed, the number of epidermal cells near the wound edges was greater in TAM-treated K14S mice compared with vehicle-treated K14S mice. There was a trend toward a lower percentage of proliferating cells in the wound areas of TAM-treated mice, but this difference was not statistically significant (Fig. 2C and Fig. S2 B and C).
Fig. 2.
Fig. S2.
To explore the mechanism behind the accelerated wound closure in young mice, we isolated RNA from wound areas of vehicle- and TAM-treated K14S mice at 2 and 6 d after injury, and quantified mRNA levels of genes associated with cell proliferation, differentiation, and inflammation. Genes associated with cell cycle progression (Ccna2, Ccnb1, and Ccnd1, encoding cyclins A2, B1, and D1), and growth inhibition (Lrig1, encoding leucine-rich repeats and immunolobin-like 1 domains) (27), were expressed at similar levels in vehicle- and TAM-treated K14S wounds (Fig. S2D), consistent with comparable PCNA staining (Fig. 2C and Fig. S2B). Genes expressed predominantly in epidermal basal cells, such as delta-like 1 (Dll1) and integrin-alpha 6 (Itga6), were also at comparable expression levels (Fig. S2D). Genes associated with epidermal stem/progenitor cells, such as leucine-rich repeat-containing G protein-coupled receptor 6 (Lgr6) (28), and with hyperproliferation, such as keratin 6B (Krt6b) (29), showed significantly lower expression in TAM-treated K14S mice at 2 d after wounding (Fig. 2D). Thus, it is unlikely that increased proliferation is responsible for the accelerated wound closure.
We also assessed apoptosis in the wound sites. Both cohorts had few TUNEL-positive cells, suggesting that apoptosis does not contribute to the difference in wound closure (Fig. S2E). Finally, because transient inflammation is important for wound healing (30), we assessed mRNA levels of genes encoding the proinflammatory cytokines IL-1α (Il1a), IL-1β (Il1b), and tumor necrosis factor (Tnf), which remained similar in vehicle- and TAM-treated K14S wounds (Fig. S3A). Thus, epidermal loss of Sod2 does not have a significant affect on the inflammatory response during wound healing.
Fig. S3.
Increased Epidermal Differentiation in Young Mice.
In contrast to proliferation- and inflammation-associated genes, genes associated with differentiation, expressed predominantly in suprabasal cells, were elevated at wound sites of TAM-treated K14S mice (Fig. 2E). These genes include loricrin (Lor), S100 calcium-binding protein A3 (S100a3), and, to a lesser extent, Krüppel-like factor 9 (Klf9), keratin 10 (Krt10), and CD36, suggesting that epidermal Sod2 deficiency enhances keratinocyte differentiation. Therefore, we examined the healing wounds by histology. At 6 d after injury, wounds in vehicle-treated mice had numerous cells in the SB layer and a negligible SG layer (Fig. 2F and Fig. S3B). In contrast, wounds in TAM-treated mice had few cells in the SB layer and a substantial SG layer (Fig. 2F and Fig. S3B). There were no apparent differences in the SC and SS layers.
To confirm the prominent SG layer in TAM-treated K14S mice, we immunostained wounds for loricrin (LOR), which was more prominent in the TAM-treated animals (Fig. 2G). These wounds also contained higher Lor mRNA levels (Fig. 2E). Because SB cells differentiate into SS and SG layers, the increased SG layer in wounds of TAM-treated K14S mice suggests that Sod2 deficiency accelerates epidermal differentiation during wound healing. This acceleration was transient; at 10 d after wounding, epidermal stratification was similar in the TAM- and vehicle-treated mice (Fig. S3C). Thus, increased differentiation, rather than cell proliferation, appears to drive the more rapid wound closure in young TAM-treated K14S mice.
Delayed Wound Closure and Rapid Epidermal Thinning in Old Mice.
The accelerated wound closure in young keratinocyte-specific Sod2-deficient mice suggests that mitochondrial dysfunction improves skin repair, contradicting the free radical theory of aging. To explore this possibility, we monitored wound closure in older (age 11 and 14 mo) mice treated with TAM or vehicle at age 4 mo. Although keratinocyte-specific Sod2 deficiency accelerated wound closure in young mice (Fig. 2A), this acceleration was lost in 11-mo-old mice (Fig. S4A). Furthermore, Sod2 deficiency delayed wound closure in 14-mo-old mice (Fig. 3A). Thus, the Sod2 deficiency promoted epidermal differentiation and wound closure in young mice, but delayed wound closure in older mice. We also monitored epidermal thickness, which declines with age in humans and mice (31, 32). We treated K14S mice with vehicle or TAM at age 4 mo, and measured epidermal thickness 4, 7, and 10 mo later (at age 8, 11, and 14 mo). Vehicle-treated mice showed a significant decline in epidermal thickness at 10 mo after treatment, whereas TAM-treated animals exhibited this decline at 7 mo after treatment (Fig. 3B and Fig. S4B). Thus, epidermal Sod2 deficiency accelerated age-associated thinning of the epidermis.
Fig. 3.
Fig. S4.
Depletion of Epidermal Stem Cells in Old Mice.
One possible reason for the phenotypes of older TAM-treated K14S mice is a reduced capacity of stem/progenitor cells to repopulate the tissue. We quantified mRNA levels of genes expressed in K14-positive stem cells, involucrin (Ivl)-committed progenitor cells, total basal interfollicular epidermal (IFE) cells, and suprabasal IFE cells, as described previously (33). Epidermal Sod2 deficiency did not significantly alter gene expression associated with K14-positive stem cells or Ivl-committed progenitor cells in young mice (Fig. 3C and Fig. S4 C and D), but significantly decreased the expression of genes associated with K14-positive stem cells in older mice (Fig. 3C). These genes include kinesin family member 11 (Kif11), Ccnb1, centromere protein E (Cenpe), cell division cycle 20 (Cdc20), Cdca5, and Kif14. Because expression of these proproliferative genes declined in the TAM-treated, but not vehicle-treated, K14S epidermis at the older age but not the younger age, our data suggest that aging exacerbates the effects of the Sod2 deficiency.
mRNAs associated with Ivl-committed progenitor cells increased with age, but only the ceramide synthase 4 (Lass4) mRNA significantly declined in the old Sod2-deficient epidermis (Fig. S4 C and D). mRNA levels of genes expressed in basal and suprabasal cells showed no change (Fig. S4 C and D). Our data indicate that the Sod2 deficiency depletes K14-positive stem cells, but not Ivl-committed progenitor or total basal and suprabasal IFE cells, in aged mice.
To verify these phenotypes, we analyzed epidermal stem cell populations by flow cytometry. Using well-characterized markers (CD49f, CD34, and Sca1), as described previously (34), we found a marked reduction in K14-positive stem cells in the junctional zone and bulge area in 14-mo-old TAM-treated, but not vehicle-treated, K14S skin (Fig. 3D and Fig. S5). Because junctional zone and bulge area stem cells are partly responsible for repopulating the epidermis after injury, their depletion in TAM-treated K14S mice suggests that mitochondrial dysfunction exhausts these cells. This exhaustion takes time and manifests as delayed wound healing only in older mice.
Fig. S5.
Cellular Senescence and Reduced Regeneration.
Mitochondrial dysfunction can induce cellular senescence (19, 35). We have shown that constitutive Sod2 deficiency causes the chronic presence of senescent keratinocytes in the epidermis (19). We also have shown that cutaneous wounding transiently induces senescence in fibroblasts and endothelial cells, which promotes wound healing (36). To determine whether senescent cells reside in the skin of keratinocyte-specific Sod2-deficient mice, we treated young (4 mo old) K14S mice with TAM or vehicle and stained the skin for senescence-associated β-galactosidase (SA-βgal), an established senescence marker (37), 4 mo later. Relative to vehicle-treated mice, TAM-treated mice had ∼2.5-fold higher SA-βgal activity in the epidermis (Fig. 4A and Fig. S6A). Interestingly, the staining was most prominent in the stratum corneum or more differentiated layers; staining in hair follicles was nonspecific, as reported previously (37). We used qPCR to assess the expression of p16INK4a, an additional senescence marker (36). Untreated 8-mo-old K14S mice have low to undetectable levels of epidermal p16INK4a mRNA. After TAM treatment, however, p16INK4a mRNA was detectable in most (>70%) K14S skin samples (Fig. 4B). p16INK4a mRNA persisted even 10 mo after TAM treatment (Fig. S6B), confirming that Sod2 deficiency induced persistent senescence in the epidermis.
Fig. 4.
Fig. S6.
Platelet-derived growth factor-A (Pdgfa) is secreted by senescent cells during wound healing (36), but showed no difference in wounds of 8-mo-old vehicle- and TAM-treated K14S mice, treated at age 4 mo (Fig. S6C). We also assessed mRNA levels of other factors that could contribute to wound healing, including Pdgfb, transforming growth factor-beta 1–3 (Tgfb1, Tgfb2, and Tgfb3), fibroblast growth factors 2 and 7 (Fgf2 and Fgf7), and vascular endothelial growth factor (Vegf), none of which differed between the two groups (Fig. S6C). Thus, it is unlikely that growth factors contribute to the altered wound healing in epidermal Sod2-deficient mice.
The senescence growth arrest could reduce the ability of tissues to regenerate. To determine the effect of Sod2 deficiency on epidermal proliferative capacity, we treated K14S mice with the proproliferative agent 12-O-tetradecanoylphorbol-13-acetate (TPA). TPA promoted epidermal thickening (arrows) in vehicle-treated, but not in TAM-treated, K14S mice (Fig. 4C). Furthermore, it caused epidermal lesions and SC layer separation in TAM-treated K14S mice (Fig. 4C), which occurs when old mice are treated with TPA (38). Moreover, whereas TPA enhanced epidermal proliferation in vehicle-treated K14S mice, as determined by Ki67 staining, it induced less proliferation in the TAM-treated epidermis (Fig. 4D and Fig. S6D). Thus, forced epidermal proliferation by TPA accelerated epidermal thinning in young Sod2-deficient mice, phenocopying the reduced proliferative capacity observed at older ages.
Mitochondrial Dysfunction in Human Keratinocytes.
To test the idea that mitochondrial dysfunction reduces proliferation and increases differentiation in epidermal cells, we treated primary human keratinocytes with the mitochondrial electron transport chain complex I inhibitor rotenone. We previously showed that these cells senesce in response to rotenone (19). Rotenone also increased p16INK4a mRNA (Fig. 4E) and cell size (Fig. S6E), a characteristic of senescent cells, and decreased mRNA levels of the proproliferation genes CCNA2 and CCNB1 (Fig. 4F), supporting the idea that persistent mitochondrial dysfunction halts keratinocyte proliferation by inducing cellular senescence. Notably, rotenone increased the differentiation-associated mRNAs CD36 and KLF9 and, to a lesser extent, S100A3 (Fig. 4F), consistent with mitochondrial dysfunction promoting keratinocyte differentiation. We conclude that mitochondrial dysfunction can accelerate wound closure in young Sod2-deficient mice by increasing keratinocyte differentiation, whereas persistent mitochondrial dysfunction during aging can deplete dividing keratinocytes.
Discussion
Our results demonstrate a previously unidentified age-dependent contribution of mitochondria to stem cell and skin function. The use of K14S mice allowed us to study mitochondrial dysfunction in K14+ epidermal cells in young and old animals, leading to important and surprising conclusions. First, mitochondrial dysfunction (owing to Sod2 deficiency) can accelerate wound closure in young mice. This unexpected response was associated with increased reepithelialization and epidermal differentiation. Second, mitochondrial dysfunction can deplete the stem cell reservoir in older mice, leading to decreased epidermal thickness and delayed wound closure. Third, mitochondrial dysfunction induces cellular senescence in many K14+ epidermal keratinocytes. Finally, forced epidermal cell proliferation by TPA limited the ability of keratinocytes to proliferate and populate the epidermis in young Sod2-deficient mice, mimicking the decline in tissue regenerative potential at older ages. To our knowledge, this is the first report demonstrating markedly disparate age-dependent effects of mitochondrial dysfunction.
Mitochondrial oxidative stress can potentiate stem/progenitor cell differentiation in Drosophila hematopoietic cells (39). Our findings suggest that this is also true in murine skin. The signaling pathways associated with this phenomenon are incompletely understood. However, mitochondrial oxidative stress was shown to promote epidermal differentiation in mice (40) and in culture (41), as did uncoupled mitochondria (42). Our studies of constitutive Sod2-deficient mice also showed increased keratinocyte differentiation (19), but early lethality complicated interpretation of this result. An interesting question for future study is whether a mitochondrial signal is needed for wound closure in humans.
Mitochondrial dysfunction caused by epidermal Sod2 deficiency depleted epidermal stem cells, but with phenotypic effects only later in life. Some stem cells, including those in the epidermis, reportedly are resistant to oxidative stress and refractory to age-related decline in mice (43, 44). Because rapidly dividing transit-amplifying (TA) cells arising from stem cells are responsible for epidermal expansion (45), we speculate that the senescence of TA cells increases the demand for stem cells. Thus, whereas epidermal stem cells can resist oxidative damage, the increased senescence resulting from mitochondrial oxidative stress can exhaust the stem cell pool (46), causing epidermal thinning and delayed wound healing but only at late ages. Therefore, although epidermal stem cells might not be a direct target of oxidative damage, senescence in TA cells can drive stem cell exhaustion with aging.
Constitutive whole-body Sod2 deficiency decreased epidermal thickness by postnatal day 17 (19), yet keratinocyte-specific Sod2 deficiency caused epidermal thinning after age 7 mo. This milder phenotype suggests that other tissues might contribute to epidermal thinning in constitutive whole-body Sod2-deficient mice. Likewise, the numerous tissues that are affected in constitutively Sod2-deficient mice (10, 12) might synergize through cell nonautonomous and/or systemic factors to accelerate the aging phenotypes in these mice.
Finally, senescent cells accumulate with age in multiple tissues, including the skin, where they are thought to contribute to impaired tissue homeostasis and regeneration (18, 47, 48). On the other hand, senescent cells are induced after skin or liver injury; in the skin, they appear to accelerate wound closure, and in both tissues they appear to limit fibrosis (36, 49, 50). Senescent cells also occur during embryonic development, where they apparently fine-tune morphogenesis (51, 52). We speculate that mitochondrial signals might contribute to the pleiotropic effects of senescent cells, but these effects might depend strongly on age.
Materials and Methods
Animal Experiments.
All animal studies complied with protocols approved by the Institutional Animal Care and Use Committee of the Buck Institute for Research on Aging. Sod2tm1Smel mice were generated by inserting LoxP sites 3′ of exon 1 and 5′ of exon 4 (Fig. S6F and SI Materials and Methods). Tg(KRT14-cre/Esr1)20Efu/J × Sod2tm1Smel (K14S) mice were generated by crossing Sod2tm1Smel (S) mice with Tg(KRT14-cre/Esr1)20Efu (K14) mice (Jackson Laboratory). K14R mice were generated by crossing B6-Gt(ROSA)26Sortm1Sor/J mice (Jackson Laboratory) with K14 mice. Genotyping was performed by PCR (SI Materials and Methods; primer sequences listed in Table S1). TAM (50 mg/kg body weight) or vehicle (corn oil) was given to 4-mo-old male mice by i.p. injection. Wounds were delivered to dorsal skin using an 8-mm biopsy punch and measured with a caliper. Wound areas were collected after biopsy as described and processed for staining and/or qPCR. Skin samples were also collected after topical treatment with TPA (9.87 µg/mouse/3 mm2) or vehicle (acetone) for 48 h.
Table S1.
| Gene name | Forward primer sequence | Reverse primer sequence |
|---|---|---|
| Neo | 5′-AGGCTATTCGGCTATGACTGGGG-3′ | 5′-TGGATACTTTCTCGGCAGGAGC-3′ |
| Bl6 | 5′- TCTGGACAAACCTGAGCCCTAAG-3′ | 5′-CCTGAACACATTCTCTATTCCTCCC-3′ |
| Cre | 5′-CCCGCAGAACCTGAAGATGTT-3′ | 5′-CGGCTATACGTAACAGGGTG-3′ |
| Gdf | 5′-AAGCCCTCAGTCAGTTGTGC-3′ | 5′-AAAACCATGAAAGGAGTGGG-3′ |
| LoxP | 5′-GCGGTCGTGTAAACCTCAATAGAG-3′ | 5′-AAAAAACCAACAATCGGGGC-3′ |
| Total control | 5′- GAGGGGCCCTGATTACTCC-3′ | 5′- GAAACCCTGGAGACTTTCCTC-3′ |
Tissue Staining.
Tissues were fixed in 10% (vol/vol) buffered formalin, embedded in paraffin, cut into 7-µm-thick sections, and processed for immunofluorescence (19), H&E (12), and picrosirius red staining (SI Materials and Methods). Epidermal thickness was measured using ImageJ software. Skin samples were also embedded in frozen optimal temperature cutting medium (OCT), cut, and processed for immunofluorescence, SA-βgal activity (19), and SDH or COX activity staining (12). For mouse tail whole mounts, epidermal sheets were isolated, fixed, permeabilized, and stained as described previously (53). Antibody conditions are described in SI Materials and Methods.
Keratinocyte Analysis.
Mouse keratinocytes were isolated from dorsal skin and processed for genotyping, RT-PCR and flow cytometry (SI Materials and Methods). PCR primer sets are listed in Tables S2 and S3. For flow cytometry, cell types were identified by gating procedures as described previously (34) and analyzed using FlowJo analysis software.
Table S2.
| Gene name | Forward primer sequence | Reverse primer sequence | UPL probe no. |
|---|---|---|---|
| Actb | 5′-CTAAGGCCAACCGTGAAAAG-3′ | 5′-ACCAGAGGCATACAGGGACA-3′ | 64 |
| Ccna2 | 5′- TGCAAACTGTAAGGTTGAAAGC-3′ | 5′- TGTAGAGAGCCAAGTGGAAGG-3′ | 67 |
| Ccnb1 | 5′-TGCATTTTGCTCCTTCTCAA-3′ | 5′-CAGGAAGCAGGGAGTCTTCA-3′ | 45 |
| Ccnd1 | 5′-GAGATTGTGCCATCCATGC-3′ | 5′-CTCCTCTTCGCACTTCTGCT-3′ | 67 |
| Cd36 | 5′- TTGAAAAGTCTCGGACATTGAG-3′ | 5′- TCAGATCCGAACACAGCGTA-3′ | 6 |
| Cdc20 | 5′- GAGTGCTGTGGATGTGCATT-3′ | 5′- TCAGCTCCTTATAGTGGGGAGA-3′ | 58 |
| Cdca5 | 5′- TCAAGACTTGTAGTGTCCCTGGTA-3′ | 5′- CTCCAGAGTCAGGCTCAACA-3′ | 67 |
| Cdkn2a (p16) |
5′-AATCTCCGCGAGGAAAGC-3′ | 5′-GTCTGCAGCGGACTCCAT-3′ | 91 |
| Cenpe | 5′- CGCCTCCCAGTCTGTTGT-3′ | 5′- TCACTGTCCCGTGTGGAAG-3′ | 58 |
| Cenpp | 5′- GCAGAATGCTGCAAAACG-3′ | 5′- CAAAGATTCCCACTCCTCAGA-3′ | 7 |
| Dgat2 | 5′- GCTGGTGCCCTACTCCAAG-3′ | 5′- GCTTGGGGACAGTGATGG-3′ | 9 |
| Dll1 | 5′- GGGCTTCTCTGGCTTCAACT-3′ | 5′- CACTTGGCACCGTTAGAACA-3′ | 103 |
| Dsc1 | 5′- TGGTCCACCTTTTCAGTTCC-3′ | 5′- ATGGCACGTTTACCATCCTG-3′ | 11 |
| Elovl7 | 5′- CAGTGTCCCCCAGGTAAGTG-3′ | 5′- CACAAACCCTACAACCAGTGAC-3′ | 1 |
| Il1a | 5′- TTGGTTAAATGACCTGCAACA-3′ | 5′- GAGCGCTCACGAACAGTTG-3′ | 52 |
| Il1b | 5′- TGTAATGAAAGACGGCACACC-3′ | 5′- TCTTCTTTGGGTATTGCTTGG-3′ | 78 |
| Il1r2 | 5′-GCAAGAAGCAGCAAGGTACA-3′ | 5′-CCGCACCAACTTCCTGAG-3′ | 1 |
| Itga2 | 5′- AGGGGAGCAAATATTCAGCA-3′ | 5′- CCAACTTGTGCCATTTCCAT-3′ | 63 |
| Itga3 | 5′- GAGCTGTGGTTGGTGCTTG-3′ | 5′- GCACTTCCACAAGAGGAGGAT-3′ | 45 |
| Itga6 | 5′- GACCAGTGGATGGGAGTCAC-3′ | 5′- TGCACACGTCACCACTTTG-3′ | 21 |
| Itgb1 | 5′- CTGCTTCTAAAATTGAGATCAGGA-3′ | 5′- TCCATAAGGTAGTAGAGATCAATAGGG-3′ | 41 |
| Ivl | 5′- GGATCTGCCTGATCAAAAGTG-3′ | 5′- CAGCTGCTGCTTTTGTGG-3′ | 71 |
| Kif11 | 5′- ACAATGGAAGGTGAAAGGTCA-3′ | 5′- TGGAATTATACCAGCCAGAGGA-3′ | 52 |
| Kif14 | 5′- GGGCAGAGGTCTCTGAACTG-3′ | 5′- GCAAGCACGGTAGTTCTCCT-3′ | 58 |
| Klf9 | 5′- CTCCGAAAAGAGGCACAAGT-3′ | 5′- GCGAGAACTTTTTAAGGCAGTC-3′ | 76 |
| Krt1 | 5′- TTTGCCTCCTTCATCGACA-3′ | 5′- GTTTTGGGTCCGGGTTGT-3′ | 62 |
| Krt10 | 5′- CGTACTGTTCAGGGTCTGGAG-3′ | 5′- GCTTCCAGCGATTGTTTCA-3′ | 95 |
| Krt6b | 5′- CACATTTGGTGCTTCATGCT-3′ | 5′- CCAGGCAACTCAGAGACAGA-3′ | 52 |
| Lass4 | 5′- TTCCCAGTGGCTCTGGTC-3′ | 5′- GGCAAGGCCACAAATCTCT-3′ | 76 |
| Lgr6 | 5′- TGTGCCAACAGCTGCCTA-3′ | 5′- AGGCTGGGTAACTCCTCGAT-3′ | 1 |
| Lor | 5′-GGTTGCAACGGAGACAACA-3′ | 5′- CATGAGAAAGTTAAGCCCATCG-3′ | 11 |
| Lrig1 | 5′- ACAGCTGCCCCACATACAAC-3′ | 5′- GGGATGGTAGGCTGTGTCA-3′ | 21 |
| Pdgfa | 5′-GTGCGACCTCCAACCTGA-3 | 5′-GGCTCATCTCACCTCACATCT-3′ | 52 |
| Pdgfb | 5′-CGGCCTGTGACTAGAAGTCC-3′ | 5′-GAGCTTGAGGCGTCTTGG-3′ | 32 |
| S100a3 | 5′- AGCAACAGCAGCAGTGTGAG-3′ | 5′- TGGAAGGTGCACACGATG-3′ | 1 |
| Sod2 | 5′-CCATTTTCTGGACAAACCTGA-3′ | 5′- GACCCAAAGTCACGCTTGATA-3′ | 67 |
| Tgfb1 | 5′-TGGAGCAACATGTGGAACTC-3′ | 5′-CAGCAGCCAATTACCAAG-3′ | 72 |
| Tgfb2 | 5′-CTTACCCTAAGCGAGAAAGTGC-3′ | 5′- GCACCCTTCCCTAGCTTCTC-3′ | 31 |
| Tgfb3 | 5′- GCAGACACAACCCATAGCAC-3′ | 5′- GGGTTCTGCCCACATAGTACA-3′ | 1 |
| Tgm1 | 5′- GCCCTTGAGCTCCTCATTG-3′ | 5′- CCCTTACCCACTGGGATGAT-3′ | 10 |
| Tnf | 5′- TCTTCTCATTCCTGCTTGTGG-3′ | 5′- GGTCTGGGCCATAGAACTGA-3′ | 49 |
| Tuba1a | 5′-CTGGAACCCACGGTCATC-3′ | 5′-GTGGCCACGAGCATAGTTATT-3′ | 88 |
| Vegf | 5′-AAAAACGAAAGCGCAAGAAA-3′ | 5′-TTTCTCCGCTCTGAACAAGG-3′ | 1 |
Table S3.
| Gene name | Forward primer sequence | Reverse primer sequence | UPL probe no. |
|---|---|---|---|
| ACTB | 5′- CCAACCGCGAGAAGATGA -3′ | 5′- TCCATCACGATGCCAGTG -3′ | 64 |
| CCNA2 | 5′- CCATACCTCAAGTATTTGCCATC -3′ | 5′- TCCAGTCTTTCGTATTAATGATTCAG-3′ | 67 |
| CCNB1 | 5′- CCAGTGCCAGTGTCTGAGC-3′ | 5′- TGGAGAGGCAGTATCAACCA-3′ | 21 |
| CD36 | 5′- CCTCCTTGGCCTGATAGAAA-3′ | 5′- GTTTGTGCTTGAGCCAGGTT-3′ | 9 |
| CDKN2A (p16) |
5′- GAGCAGCATGGAGCCTTC -3′ | 5′- CGTAACTATTCGGTGCGTTG -3′ | 67 |
| DGAT2 | 5′- GAGGGGTCTGGGAGATGG-3′ | 5′- TTGGACCTATTGAGCCAGGT-3′ | 55 |
| KLF9 | 5′- CTCCGAAAAGAGGCACAAGT-3′ | 5′- CGGGAGAACTTTTTAAGGCAGT-3′ | 76 |
| LGR6 | 5′- TGTGCCAGCTTCTTCAAGG-3′ | 5′- GGGCCTTTTTGAAGACTCCT-3′ | 41 |
| S100a3 | 5′- TCTGGTTCAGGTTCCTGACTG-3′ | 5′- TGGAAGGTGCACACGATG-3′ | 50 |
| TUBA1A | 5′- CTTCGTCTCCGCCATCAG -3′ | 5′- TTGCCAATCTGGACACCA -3′ | 58 |
Cell Culture.
Human keratinocytes (AG21837) from the Coriell Cell Repository were cultured in keratinocyte growth medium (CnT-07; Zenbio) with penicillin-streptomycin (Invitrogen) in 20% O2. Media were replaced every 2 d. Cells were treated with vehicle (DMSO) or rotenone (Sigma-Aldrich) at 100 nM for 7 d, then collected for PCR analysis (SI Materials and Methods).
SI Materials and Methods
Generation of Transgenic Mice.
We contracted Ozgene (www.ozgene.com) to generate a LoxP flanked Sod2 allele. We modified the endogenous Sod2 locus in mouse embryonic stem cells by inserting a PGK/neo cassette between exons 3 and 4, which facilitated selection of the correctly modified allele. LoxP sites were inserted 3′ of exon 1 and 5′ of exon 4, and Cre-mediated recombination removed exons 2 and 3 (Fig. S6F). We chose this region for deletion because exon 3 is crucial for manganese binding and homotetramer formation. Thus, recombination completely inactivates Sod2. We confirmed the correctly modified locus by sequencing, and generated mice with the modified allele in a C57BL/6J background. The allele was registered (www.informatics.jax.org/mgihome/nomen/strains.shtml), and is termed Sod2tm1Smel (S).
Genotyping.
Genotyping was done using Platinum Blue PCR SuperMix (Life Technologies). For Sod2, we used primer pairs for neo (0.4 µM) and bl6 (0.4 µM). PCR conditions were 95 °C for 5 min; 34 cycles at 95 °C for 1 min, 59.5 °C for 1 min, and 72 °C for 30 s; and 72 °C for 5 min. For K14 mice, we used the primer pairs Cre (0.4 µM) and GDF (0.28 µM) and the following qPCR conditions: 94 °C for 2 min; 34 cycles at 94 °C for 1 min, 58.5 °C for 1 min, and 72 °C for 30 s; and 72 °C for 10 min. We detected the recombined Sod2 gene by qPCR using SensiFast Probe (Bioline) and EvaGreen (Biotium; catalog no. 31000-T). Genotyping for Sod2 deletion used the primer pairs LoxP (0.5 µM) and total control sequence (1 µM), and the following qPCR conditions: 95 °C for 10 min, and 40 cycles at 95 °C for 10 s, 60 °C for 30 s, and 72 °C for 10 s. Melting curves of PCR products were monitored to ensure the absence of primer dimers. The copy number of recombined Sod2 alleles was normalized to the copy number of the total control sequence.
Picrosirius Red Staining.
Samples were fixed in 10% buffered formalin, embedded in paraffin, and cut into 7-µm sections. Sections were then deparaffinized, stained with Weigert's hematoxylin for 8 min, and washed before being stained with picrosirius red solution [0.1% Direct Red 80 (Sigma-Aldrich) in saturated (1.3%) picric acid] for 1 h. Sections were washed with 0.5% acetic acid and dehydrated before mounting on coverslips. Slides were viewed under polarized light using a Nikon Eclipse E800 microscope. To capture the complete wound site, multiple photomicrographs of different sections were taken at 4× magnification, then tiled in Adobe Photoshop.
Immunofluorescence.
Skin was embedded in frozen OCT and processed as described. Wound site specimens were fixed in 10% buffered formalin, embedded in paraffin, and cut into 7-µm sections. Sections were deparaffinized, microwaved, blocked, and immunostained. Primary antibodies were rabbit anti-Ki67 antibody (Novus Biologicals, NB110-89717; 1:750 for OCT; Thermo Fisher Scientific, RM-9106-S0; 1:500 for paraffin-embedded), rabbit anti-PCNA antibody (Cell Signaling, B00177; 1:1,000), rat anti-CD49f (BD Pharmingen, 555734; 1:750) and rabbit anti-loricrin (Covance, PRB-145P; 1:1,000). Secondary antibodies were Alexa Fluor 555 donkey anti-rabbit (Molecular Probes; 1:750) and Alexa Fluor 488 donkey anti-rat (Molecular Probes; 1:750). Multiple photomicrographs of wound sections were taken under 10× magnification and tiled in Adobe Photoshop.
Tail Whole Mounts.
Epidermal sheets were permeabilized [0.25% fish skin gelatin (Sigma-Aldrich), 0.5% Triton X-100, and 20 mM Hepes, pH 7.2] for 30 min at room temperature, then sequentially incubated with primary antibodies at 4 °C overnight and secondary antibodies at 4 °C overnight with 4-h washes between incubations using 0.2% Tween 20 in PBS. Primary antibodies were rabbit anti-SOD2 (Assay Designs; 1:1,000) and rat anti-CD49f (BD Pharmingen; 1:1,000). Secondary antibodies were Alexa Fluor 555 donkey anti-rabbit (Molecular Probes; 1:750) and Alexa Fluor 488 donkey anti-rat (Molecular Probes; 1:750). Samples were mounted in Prolong Gold with DAPI (Life Technologies).
Keratinocyte Isolation.
Shaved mouse skin was collected, and the fat layer was removed. Skin was then washed and decontaminated with Hibiclens (Fisher Scientific) and Ca/Mg-free HBSS containing 5× antibiotics (Life Technologies). Skin was floated on 25 U/mL dispase mix (BD Biosciences) in Ca/Mg-free HBSS containing 50 µg/mL gentamicin and incubated at 4 °C overnight. Epidermal cells were scraped from the dermis and processed for PCR assays. For flow cytometry, epidermal cells were incubated with TrypLE for 8 min at 37 °C. After neutralization with 10% chelexed FBS (BioRad) in Ca/Mg-free HBSS, cells were collected by centrifugation (300 × g for 5 min), and floating cells were aspirated, filtered through a 70-µm strainer and washed twice with Ca/Mg-free HBSS. Keratinocytes were resuspended in CnT-07 (Zenbio) for flow cytometry.
RT-PCR.
Skin samples were homogenized in QIAzol Lysis Reagent (Qiagen), and RNA was isolated using the RNeasy Mini Kit (Qiagen). cDNA was synthesized using the High- Capacity cDNA Reverse Transcription Kit (Life Technologies). Then 1 μg of RNA was amplified at 95 °C for 7 min and 50 cycles of 95 °C for 5 s and 60 °C for 30 s. Transcripts were quantified by qPCR using SensiFast Probe (Bioline), normalized to β-actin and α-tubulin, and reported as relative levels. Probes were obtained from the Roche Universal Probe Library system. Primer sets were used at 0.1 μM (Tables S2 and S3).
Flow Cytometry.
Epidermal cells were incubated with purified rat anti-mouse CD16/CD32 Fc block (BD Pharmingen) at 1:100 for 20 min on ice, followed by incubation for 20 min on ice with the following primary antibodies: Sca1-FITC (e-Bioscience) at 1:100, CD49f/ItgA6-RPE (AbD Serotec) at 1:50, and CD34-Alexa Fluor 660 (e-Bioscience) at 1:40. After a wash with PBS containing 1% BSA, cells were fixed with 1% formaldehyde in PBS before flow cytometry analysis with a BD FACSAria (BD Biosciences). Corresponding isotype controls were used for compensation correction. Cell types were analyzed using FlowJo analysis software.
Data Analysis.
Data are presented as mean ± SEM and were subjected to statistical analysis using Student’s t test or two-way ANOVA (with Bonferroni post hoc analysis), as indicated in the figure legends. P < 0.05 was considered statistically significant.
Acknowledgments
We thank the Buck Morphology Core for processing tissues, San Francisco VA Medical Center’s Flow Cytomtery Core for flow cytometry, Ethan Sarnoski and Isaac Daviet for help with immunostaining, Nuno Luis for instruction on processing whole mounts, Kevin Perrott and Elvira Rafikova for blind ranking of activity staining, Sally D. Pennypacker for instruction on separating epidermis from dermis, and Leila Mashouf for measuring epidermal thickness. We especially thank Pierre-Yves Desprez for critically reading the manuscript. This work was funded by National Institutes of Health Grants AG009909 (to J.C.), AG18679 (to S.M.), AG025901 (to S.M. and J.C.), and AG041221 (to M.C.V.).
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Published online: August 3, 2015
Published in issue: August 18, 2015
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Acknowledgments
We thank the Buck Morphology Core for processing tissues, San Francisco VA Medical Center’s Flow Cytomtery Core for flow cytometry, Ethan Sarnoski and Isaac Daviet for help with immunostaining, Nuno Luis for instruction on processing whole mounts, Kevin Perrott and Elvira Rafikova for blind ranking of activity staining, Sally D. Pennypacker for instruction on separating epidermis from dermis, and Leila Mashouf for measuring epidermal thickness. We especially thank Pierre-Yves Desprez for critically reading the manuscript. This work was funded by National Institutes of Health Grants AG009909 (to J.C.), AG18679 (to S.M.), AG025901 (to S.M. and J.C.), and AG041221 (to M.C.V.).
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This article is a PNAS Direct Submission.
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The authors declare no conflict of interest.
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