Human mesenchymal stem cell (hMSC) aging may lead to a reduced tissue regeneration capacity and a decline in physiological functions. However, the molecular mechanisms controlling hMSC aging in the context of prelamin A accumulation are not completely understood. In this study, we demonstrate that the accumulation of prelamin A in the nuclear envelope results in cellular senescence and potential downstream regulatory mechanisms responsible for prelamin A accumulation in hMSCs. We show for the first time that ZMPSTE24, which is involved in the post-translational maturation of lamin A, is largely responsible for the prelamin A accumulation related to cellular senescence in hMSCs. Direct binding of miR-141-3p to the 3′UTR of ZMPSTE24 transcripts was confirmed using a 3′UTR-luciferase reporter assay. We also found that miR-141-3p, which is overexpressed during senescence as a result of epigenetic regulation, is able to decrease ZMPSTE24 expression levels, and leads to an upregulation of prelamin A in hMSCs. This study provides new insights into mechanisms regulating MSC aging and may have implications for therapeutic application to reduce age-associated MSC pool exhaustion.
Advancing age is a major risk factor for developing many chronic diseases, but the mechanisms that regulate the aging process remain largely unknown. Recent studies have revealed that aging has been negatively correlated with the number and functional activities of tissue stem cells. Adult stem cells present in mammalian organs are essential for tissue generation, maintenance and injury repair throughout adult life. Similar to other somatic cells, adult stem cells experience lifelong exposure to intrinsic and extrinsic factors, throughout the lifetime of the organism, which lead to an age-associated decline in their number and function (Janzen et al., 2006; Kasper et al., 2009; Rao and Mattson, 2001; Trosko, 2008).
It has been previously reported that the premature aging disease Hutchinson–Gilford Progeria syndrome (HGPS) is caused by a mutation in the LMNA gene (the corresponding protein is known as lamin A). HGPS seems to mainly affect mesenchymal cell lineages, and HGPS induces progerin-mediated mesenchymal stem cell pool exhaustion (Halaschek-Wiener and Brooks-Wilson, 2007) and mesenchymal lineage differentiation defects (Scaffidi and Misteli, 2006). These studies have shown that HGPS fibroblasts have a short replicative lifespan compared with their wild-type counterparts. In HGPS fibroblasts, premature senescence is due to the toxic accumulation of progerin, a mutant form of prelamin A (Bridger and Kill, 2004). In particular, introduction of progerin, a mutant form of lamin A, into hMSCs accelerated cellular senescence and caused adult stem cell dysfunction (Scaffidi and Misteli, 2008). This study showed that hMSCs expressing progerin display marked defects in growth, nuclear abnormalities and accelerated senescence, which closely resemble the defects observed in cells from HGPS patients.
Lamin A, synthesized as a precursor named prelamin A, undergoes a multi-step maturation process, which includes ZMPSTE24-mediated cleavage of the last 15 amino acids (Corrigan et al., 2005). Failure to cleave the last 15 amino acids of prelamin A because of disruptions in either prelamin A or ZMPSTE24 causes nuclear structural abnormalities, a shortened lifespan and multiple age-related phenotypes (Bergo et al., 2002; Pendás et al., 2002). Some reports have demonstrated that ZMPSTE24 expression levels are downregulated in aged or senescent human vascular smooth muscle cells and fibroblast cells (Ragnauth et al., 2010; Ukekawa et al., 2007), but the molecular mechanisms that regulate ZMPSTE24 expression during hMSC aging have not been identified (Maraldi et al., 2011).
MiRNAs, small RNAs of ∼22 nucleotides in length, are known to perform important regulatory roles through the repression of target mRNA translation by complementary binding to the 3′ untranslated region (UTR) (Bartel, 2004). More recent evidence has shown that several miRNAs (miR-371, miR-369-5p, miR-29c, miR-499 and let-7f) are involved in the regulation of cellular senescence (Wagner et al., 2008). However, the effects of these miRNAs on ZMPSTE24 expression during in vitro cellular senescence of hMSCs are unknown.
The dynamic balance between epigenetic elements, such as acetylation, methylation or phosphorylation, is important for the maintenance of stemness and the regulation of cellular senescence because these elements control chromatin modification and transcriptional regulation (Bibikova et al., 2008; Narita, 2007). Histone deacetylase (HDAC) and DNA methyltransferase (DNMT) inhibitors induce senescence of hMSCs by affecting p16INK4A and p21CIP1/WAF1 expression (Jung et al., 2010; So et al., 2011). Moreover, non-coding RNAs as well as protein-coding RNAs can be regulated by epigenetic modifications (Delcuve et al., 2012).
In the study reported here, we assessed the impact of downregulation of ZMPSTE24 on the proliferation defects and DNA damage responses observed in hMSCs as they progress toward replicative senescence. Furthermore, our results indicate that ZMPSTE24 expression can be directly regulated by miR-141-3p during hMSC senescence. We anticipate that these results might provide clues to understanding the biological relevance of ZMPSTE24 in hMSCs senescence and the aging process of adult stem cells.
Senescent hMSCs show increased prelamin A accumulation and progeria-like abnormalities in the nucleus
First, we evaluated the direct effects of replicative- and progerin-induced senescence on senescence-associated-β-galactosidase (SA-β-gal) activity and hMSC proliferation. Both senescent conditions significantly increased SA-β-gal activity and decreased hMSC proliferation (Fig. 1A,B). Replicative and progerin-induced senescence clearly increased expression of p16INK4A, a senescence marker, in hMSCs (Fig. 1C; supplementary material Fig. S1B). To determine whether prelamin A accumulation is related to the physiological aging of hMSCs, we investigated the accumulation levels of prelamin A in senescent cells. Replicative senescence significantly increased prelamin A accumulation compared with negative controls (Fig. 1C). To support this observation, we subsequently investigated whether replicative and progerin-induced senescence were correlated with abnormal nuclear morphology of hMSCs. We found that replicative and progerin-induced senescence increased the incidence of severely wrinkled nuclei in hMSCs compared with negative controls (Fig. 1D). To assess the age-related accumulation of DNA damage under a prelamin A accumulation condition, we examined whether replicative and progerin-induced senescence had any effect on the expression of γH2AX as a general biomarker of DNA damage. As illustrated in Fig. 1E, γH2AX expression was increased significantly under both senescent conditions.
ZMPSTE24 inhibition induces cellular senescence with DNA damage accumulation
We found that expression of the LMNA gene was not significantly altered by serial in vitro passages (supplementary material Fig. S1A). As illustrated in Fig. 1F and supplementary material Fig. S1C, the expression of ZMPSTE24, which converts prelamin A into mature lamin A, was clearly decreased by replicative senescence in hMSCs. To further elucidate the role of ZMPSTE24 as a regulator of prelamin A accumulation during cellular senescence, hMSCs were transfected with ZMPSTE24 siRNA. We found that transfection with ZMPSTE24 siRNA efficiently increased prelamin A accumulation, which was followed by an increase in p16INK4A and γH2AX expression (Fig. 1G). Furthermore, ZMPSTE24 knockdown increased SA-β-gal activity (Fig. 1H) and decreased proliferation (Fig. 1I; supplementary material Fig. S1D) compared with negative controls, and these effects coincided with increased prelamin A-accumulation and γH2AX expression (Fig. 1J).
HDAC regulates ZMPSTE24 expression during replicative cellular senescence
The activities of DNMT and HDAC, regulatory factors of epigenetic states and transcriptional activities, decreased during cellular senescence in our present (Fig. 2A) and previous studies (Jung et al., 2010; So et al., 2011). To evaluate whether these epigenetic factors are involved in the regulation of ZMPSTE24 expression, we determined the effects of the DNMT inhibitor [5-azacytidine (5-AzaC); 2 µM] and two HDAC inhibitors [valproic acid (VPA; 4 mM) and sodium butyrate (NB; 2 mM)] on the induction of senescence in hMSCs. As shown in Fig. 2B–E, inhibition of DNMT and HDAC induced cellular senescence and increased p16INK4A expression. However, only HDAC inhibition decreased ZMPSTE24 expression, which resulted in an increase in prelamin A expression (Fig. 2E) and nuclear abnormalities (Fig. 2F,G). Knockdown of HDAC1 and HDAC2 decreased ZMPSTE24 expression, resulting in prelamin A accumulation (Fig. 2H), and treatment with specific inhibitors for HDAC1 and HDAC2 activities increased p16INK4A expression and SA-β-gal activity (Fig. 2H,I). We also found that suppression of HDAC1 and HDAC2 activity by specific inhibitors decreased proliferation (Fig. 2J; supplementary material Fig. S1E) and that these effects coincided with increased nuclear abnormalities (Fig. 2K). Taken together, these results indicate that the inhibition of HDAC suppresses ZMPSTE24 expression, which in turn induces prelamin A-mediated cellular senescence in hMSCs.
miR-141-3p regulates cellular senescence through the regulation of ZMPSTE24 expression
Epigenetic regulations, such as DNA methylation and histone modifications, can affect the expressions of miRNAs as well as protein-coding RNAs (Delcuve et al., 2012). Moreover, according to our previous study, the decrease of DNMT activity affected the expression of polycomb-gene-targeting miRNAs in replicative senescent hMSCs (So et al., 2011). Therefore, we hypothesized that the decrease of HDAC activity would affect the expressions of ZMPSTE24-targeting miRNAs in replicative senescent hMSCs. Using three periodicity and correlation algorithms (miRanda, TargetScan5.0 and PicTar), we identified three miRNAs (miR-124, miR-141-3p and miR-182) as the most likely regulatory miRNAs affecting ZMPSTE24 expression (Fig. 3A). Among the predicted miRNAs, miR-141-3p significantly reduced firefly luciferase activity in the reporter containing the ZMPSTE24-3′UTR compared with the control Renilla luciferase activity, suggesting that miR-141-3p could directly bind to the 3′UTR of the ZMPSTE24 RNA (Fig. 3B,C). To support this observation, we further investigated whether miR-141-3p regulates ZMPSTE24 expression, prelamin A accumulation and nuclear abnormalities. The transfection of hMSCs with miR-141-3p decreased ZMPSTE24 expression, resulting in increased expression of prelamin A, p16INK4A and γH2AX. Conversely, these effects were reversed by miR-141-3p knockdown (Fig. 3D). In the miR-141-3p-transfected cells, MTT assays and cell cycle analyses revealed that the growth rate was reduced 48 hours post-transfection (supplementary material Fig. S3A,B). In contrast, the transfection of hMSCs with anti-miR-141-3p yielded an increase in proliferation. The correlation between the expression of ZMPSTE24 and miR-141-3p was confirmed in five other hMSC cell lines (supplementary material Fig. S2A), and the cells treated with miR-141-3p showed changes in their nuclear morphology, with many of the cells displaying folds in the nuclear envelope (Fig. 3E). To determine whether these observations could be extended to hMSCs derived from different sources, we analyzed the effects of miR-141-3p on the nuclear morphology of hMSCs derived from bone marrow and adipose tissue (supplementary material Fig. S2B). The overexpression of miR-141-3p also increased SA-β-gal activity and γH2Ax expression (Fig. 3F,G) in these cells.
To demonstrate the long-term effect of miR-141-3p, we infected hMSCs with a GFP-tagged miR-141-3p virus. The virally infected hMSCs showed strong GFP expression, with significantly elevated levels of miR-141-3p, as confirmed by real-time qPCR (Fig. 4A,B). Using the MTT assay and a cell cycle analysis, we also confirmed that proliferation was inhibited after 5 days of miR-141-3p overexpression (supplementary material Fig. S3D,E). To examine the role of miR-141-3p in vivo, we delivered the miR-141-3p virus into mice by intraperitoneal (i.p.) injection. Previously, it was reported that using this method of delivery the organ with the highest level of viral particles was the liver, presumably because this organ has a relatively high percentage of blood-derived cells (Delgado et al., 2008; Dismuke et al., 2009). Therefore, we obtained liver tissue 7 days after i.p. injection and performed immunohistochemistry and western blot analyses. Because the viral vector was designed to express GFP together with miR-141-3p, we used GFP fluorescence to detect the overexpression of miR-141-3p in liver tissue (Fig. 4C). Accordingly, we found that the expression of miR-141-3p was increased and ZMPSTE24 expression was reduced in the miR-141-3p-virus-injected liver (Fig. 4D,E). These results indicate that ZMPSTE24 is the direct target of miR-141-3p both in vitro and in vivo. Collectively, these results support the hypothesis that miR-141-3p modulates cellular senescence through the regulation of ZMPSTE24, which correlates with the change in phenotype observed in the siZMPSTE24-treated cells.
Activation of a histone marker at the miR-141-3p promoter increases miR-141-3p expression
We hypothesized that miR141-3p may play a role in cellular senescence of hMSCs by controlling ZMPSTE24 expression. The expression levels of miR-141-3p were significantly increased with consecutive passages (Fig. 5A). We subsequently investigated whether the increased expression levels of miR-141-3p were correlated with changes in the histone acetylation and methylation of the promoter region of miR-141-3p during cellular senescence by using a chromatin immunoprecipitation (ChIP) assay. In late passage cells, the acetylation of histone H4 and the trimethylation of histone H3 at the lysine 4 residue (H3K4) increased in the putative coding region of miR-141-3p (Fig. 5B), whereas histone H3 trimethylation at lysines 9 (H3K9)z and 27 (H3K27) decreased. Because the transcription of miRNA is mediated by RNA polymerase II, we also tested whether RNA polymerase II was enriched at the miRNA coding region. RNA polymerase II binding to the putative coding region of miR-141-3p significantly increased during cellular senescence of hMSCs (Fig. 5C), whereas the expression of miR-106b, which was shown to not target ZMPSTE24 (see Fig. 3), was not altered by consecutive passages or VPA- or NB-induced senescence (supplementary material Fig. S4). To determine whether HDAC activity could regulate the transcription of ZMPSTE24-targeting miRNA, we investigated the expression of miR-141-3p using DNMT- and HDAC-inhibitor-induced senescence. 5-AzaC treatment did not affect the expression of miR-141-3p, whereas VPA and NB treatment clearly increased the miR-141-3p expression levels compared with negative controls (Fig. 5D–F). Moreover, inhibition of HDAC1 and HDAC2 significantly increased miR-141-3p expression in hMSCs (Fig. 5G). In the VPA- and NB-treated groups, the acetylation of histones H3, H4 and H3K4Me3 increased, whereas the acetylation of H3K9Me3 and H3K27Me3 decreased or remained relatively unchanged in the putative coding region of miR-141-3p (Fig. 5H). RNA polymerase II expression was induced around the miR-141-3p coding region by HDAC-inhibitor-induced cellular senescence, which suggests that the miR-141-3p coding region promotes transcription of miR-141-3p (Fig. 5I).
To identify whether the change in ZMPSTE24 expression caused by HDAC inhibitors is dependent on the miR-141-3p, we transfected anti-miR-141-3p into hMSCs after treating them with HDAC inhibitors. As expected, transfection of anti-miR-141-3p attenuated HDAC-inhibitor-induced effects on ZMPSTE24 expression (Fig. 6A). The upregulation of ZMPSTE24 expression further prevented HDAC-inhibitor-induced SA-β-gal activity and changes in hMSC nuclear morphology, with many cells exhibiting folds in the nuclear envelope (Fig. 6B–D). Taken together, these results indicate that miR-141-3p transcription is regulated by histone modification and RNA polymerase II activity at the miR-141-3p promoter region during replicative and HDAC-inhibitor-induced senescence.
hMSCs are essential for endogenous tissue generation, maintenance and injury repair throughout the adult life of mammals. Therefore, it has been proposed that MSC aging might lead to a reduction in tissue regeneration and a decline of physiological functions (Sethe et al., 2006). However, the potential molecular mechanisms controlling hMSC aging are not completely understood.
Prelamin A accumulated in hMSCs during cellular senescence, and the overexpression of prelamin A induced cellular senescence, suggesting that prelamin A is a causal factor in the initiation of hMSC aging rather than a consequence of it. It has been reported that aged vascular smooth muscle cells rapidly accumulate prelamin A, which coincides with reduced levels of ZMPSTE24, growth defects and nuclear abnormality (Ragnauth et al., 2010). These effects can be partially attenuated by the overexpression of ZMPSTE24 through suppressing expression of prelamin A (Candelario et al., 2008). Similarly, we observed a significant decrease in ZMPSTE24, which converts prelamin A into mature lamin A, during replicative senescence. These data suggest that ZMPSTE24 is largely responsible for prelamin A accumulation during aging. However, it remains to be determined whether the increase in p16INK4A in the siZMPSTE24-transfected cells is due to ZMPSTE24 itself or the accumulation of prelamin A.
The inhibition of HDACs induces the accumulation of acetylated histone forms, which modulate gene transcription by altering the chromatin structure and transcription factor complexes (Xu et al., 2007). In hMSCs, HDAC inhibitors decrease stemness and induce cellular senescence by affecting the expression of c-MYC, polycomb group genes and p16INK4A (Jung et al., 2010; Lee et al., 2009). In this study, we present direct evidence that HDAC inhibitors significantly inhibit ZMPSTE24 activity, leading to an accumulation of prelamin A after 6 days of treatment. We also demonstrate that, among the multiple targets of HDAC inhibitors, HDAC1 and HDAC2 can specifically affect the expression of ZMPSTE24 and prelamin A. Importantly, our observations in hMSCs are consistent with those of Miller et al., who recently reported that HDAC1 and HDAC2 are involved in the DNA damage response and are required for double-stranded break repair (Miller et al., 2010). However, in mouse Zmpste24−/− cells, treatment with HDAC inhibitors, including NB, promoted repair protein recruitment to DNA damage sites and substantially ameliorated aging-associated phenotypes both in vitro and in vivo (Krishnan et al., 2011). Further studies are required to identify which factor causes the different results in HDAC inhibitor treatment of Zmpste24−/− cells and MSCs. Epigenetic alterations of Zmpste24−/− cells, including a notable decrease in the acetylation status of histone H2B and H4, may be responsible for the different results.
Although some studies have reported that ZMPSTE24 expression decreases in senescent cells or tissues (Ragnauth et al., 2010; Ukekawa et al., 2007), the molecular mechanisms that modulate ZMPSTE24 expression during aging have not yet been elucidated. miRNA is a potent regulator of various target genes and the expression of nearly 60% of protein-coding RNAs are affected by miRNAs (Friedman et al., 2009). However, a significant number of miRNAs are regulated by epigenetic modulations. A short exposure of the breast cancer cell line SKBr3 to a HDAC inhibitor changed 40% of the miRNA population significantly (Scott et al., 2006). Thus, we focused on miRNA as a candidate regulator of ZMPSTE24, which is controlled by HDAC activity during senescence processes. To address this issue, we searched for miRNAs that target ZMPSTE24 during the aging process. Combining the results obtained using different predictive software prompted us to consider that transcriptional activation of miR-141-3p could be an important regulator of aging through suppression of ZMPSTE24 expression.
It has been reported that miR-200 family members (miR-141; miR-200a, b and c; and miR-429) inhibit p38α and that overexpression of miR-200s promotes human ovarian carcinogenesis and is linked to a poor overall prognosis (Mateescu et al., 2011). Furthermore, the miR-200 family has been identified as a marker for epithelial differentiation and a regulator of the epithelial-to-mesenchymal transition (EMT). The miR-200 family suppresses EMT by inhibiting translation of mRNA for the EMT activators ZEB1 and ZEB2, and ZEB factors control the miR-200 family, thus creating a feedback loop (Burk et al., 2008; Wellner et al., 2009). However, the role of miR-141 in the regulation of ZMPSTE24 expression has not been reported. Consistent with our hypothesis, we have found that miR-141-3p expression progressively increases during cellular senescence, and the transfection of hMSCs with miR-141-3p decreased ZMPSTE24 expression, which resulted in increased expression of prelamin A, p16INK4A and γ-H2AX. Moreover, we found that transcriptionally active histone forms were enriched in miR-141-3p-coding regions, and RNA polymerase II actively binds to the coding regions of miR-141 during replicative senescence. We also found that HDAC inhibitors induced histone acetylation in miR-141-3p-coding regions and increased miR-141-3p expression levels in hMSCs, suggesting that miR-141-3p expression is affected by HDAC activity during the aging process. The anti-miR-141-mediated maintenance of ZMPSTE24 expression levels resulted in suppression of an abnormal nuclear phenotype in the HDAC-inhibitor-treated cells.
In conclusion, prelamin A accumulates in hMSCs during cellular senescence in a ZMPSTE24-dependent manner, and HDAC inhibitors significantly inhibit ZMPSTE24 activity, leading to an accumulation of prelamin A. Moreover, miR-141-3p plays a role as a negative regulator of cellular senescence through the suppression of ZMPSTE24 expression, which in turn increases the accumulation of prelamin A as senescence progresses. This study provides new insights into the miR-141-3p–ZMPSTE24 signaling regulation of hMSC aging, which warrants further study, because there is great potential for pharmaceutical intervention to reduce age-associated MSC pool exhaustion.
Materials and Methods
Isolation and culture of hMSCs
Umbilical cord blood (UCB) samples were obtained with the written informed consent of the mothers, approved by the Boramae Hospital Institutional Review Board (IRB) and the Seoul National University IRB (IRB no. 0603/001-002-07C1), from the umbilical vein immediately following delivery. The hMSCs were isolated and cultured as previously described (Jung et al., 2010). Briefly, the UCB samples were mixed with Hetasep solution (StemCell Technologies, Vancouver, Canada) at a ratio of 5∶1 and then incubated at room temperature to deplete the erythrocytes. The supernatant was carefully collected, and the mononuclear cells were isolated using Ficoll density-gradient centrifugation at 1200 g for 20 minutes. The cells were washed twice in PBS and were seeded at a density of 2×105 to 2×106 cells/cm2 on plates in growth medium consisting of D-medium (formula no. 78-5470EF, Gibco BRL, USA) containing EGM-2 SingleQuot and 10% fetal bovine serum (Gibco BRL). After 3 days, the non-adherent cells were removed. For long-term culture, the cells were seeded at a density of 4×105 cells/10 cm plate, and the cells were subcultured upon reaching 80∼90% confluency.
Senescence-associated beta-galactosidase (SA β-gal) staining
The SA β-gal staining was performed as previously described (Yu et al., 2013). The hMSCs were seeded on 6-well plates at a density of 1×105 cells/well for late-passage cells and 5×104 cells/well for early-passage cells. The cells were incubated for 3 days until reaching the appropriate confluency. The cells were then washed twice with PBS and fixed with 0.5% glutaraldehyde in PBS (pH 7.2) for 5 minutes at room temperature. The cells were then washed with PBS containing MgCl2 (pH 7.2, 1 mM MgCl2) and stained with X-gal solution [1 mg/ml X-gal, 0.12 mM K3Fe(CN)6, 1 mM MgCl2 in PBS, pH 6.0] overnight at 37°C. The cells were washed twice with PBS, and the images were captured using a microscope (IX70, Olympus, Japan).
The proliferative potential of the cells was measured using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT, Sigma-Aldrich, USA) assay, which is based on the ability of live cells to convert a tetrazolium salt into purple formazan. hMSCs (20,000 per well) were seeded in 24-well plates. After 48 hours incubation, 50 ml MTT stock solution (5 mg/ml; Sigma) was added to each well, and the plates were further incubated for 4 hours at 37°C. The supernatant was removed, and 200 ml of DMSO was added to each well to solubilize the water-insoluble purple formazan crystals; the solution was then transferred to a 96-well microplate for reading. The absorbance at a wavelength of 540 nm was measured using an EL800 microplate reader (BIO-TEK Instruments, Winooski, VT, USA). All of the measurements were performed in triplicate.
Western blot analysis
Western blot analyses of lamin A, prelamin A, ZMPSTE24, HDAC1, HDAC2, DNMT1, DNMT3B, p16Ink4A, γ-H2AX and β-actin were performed as previously described (Jung et al., 2005). The hMSCs were lysed with 50 mM Tris-HCl buffer containing 0.1% Triton X-100 and freshly supplemented with a protease/phosphatase inhibitor cocktail. The proteins were then separated using 7.5–15% SDS-PAGE and transferred to nitrocellulose membranes at 350 mA for 5 hours. The primary antibodies used to detect each protein were as follows: lamin A (monoclonal, Abcam, 1∶2500); prelamin A (polyclonal, Santa Cruz, 1∶500); ZMPSTE24 (polyclonal, Abcam, 1∶500); HDAC1 (monoclonal, Upstate, 1∶2000); HDAC2 (monoclonal, Upstate, 1∶2000); DNMT1 (polyclonal, BD, 1∶1000); DNMT3A (polyclonal, Millipore, 1∶1000); DNMT3B (polyclonal, Abcam, 1∶1000); p16Ink4A (polyclonal, Abcam, 1∶1000); γ-H2AX (polyclonal, Abcam, 1∶1000) and β-actin (monoclonal, Cell-signaling, 1∶5000). All of the antibodies were used according to the manufacturer's instructions, and the protein bands were detected using an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, UK).
Viral packaging and cell infection
Cells expressing GFP-wt-lamin A and GFP-progerin were obtained from Addgene (Cambridge, MA; Plasmid 17662, 17663), and the miR-141 lentiviral vector was purchased from Genecopoeia (Rockville, MD; HmiR0181). Viral production and infection were performed as previously described (Yu et al., 2012b). Using the VSVG-based package system or the Mission lentiviral packaging mix (Sigma, Ronkonkoma, NY, USA) 6 µg of vectors were added to tubes containing the Fugene 6 transfection reagent (Roche, Basel, Switzerland). The plasmids were transfected into 293FT cells; after 48 and 72 hours, the viral supernatants were filtered through a 0.45 µm cellulose acetate filter and concentrated by centrifugation at 50,000 g for 90 minutes. The viral supernatants were then used to infect the human MSCs in the presence of polybrene at 5 µg/ml (Sigma).
The total cellular RNA was extracted from cells using the TRIzol reagent™ (Invitrogen, USA) according to the manufacturer's instructions. The cDNA was synthesized by adding the purified RNA and oligo(dT) primers to Accupower RT premix (Bioneer, Korea), according to the manufacturer's instructions. For the miRNA cDNA synthesis, the NCode VILO miRNA cDNA Synthesis Kit (Invitrogen, USA) was used.
Real-time quantitative PCR
Real-time qPCR analyses were performed using SYBR® Green (Applied Biosystems, USA) according to the manufacturer's protocol. For the miRNA real-time qPCR, the universal primers supplied with the NCode VILO miRNA cDNA Synthesis Kit (Invitrogen, USA) and miRNA-specific primers were used. RPL13A was used as an internal control. All of the amplicons were analyzed using the Prism 7000 sequence detection system 2.1 software (Applied Biosystems, USA). The primer set sequences used for this study are listed in supplementary material Table S1.
siRNA, mature miRNA and anti-miRNA transfection studies
Transient transfection assays were performed using specific, commercially available siRNAs for the inhibition of ZMPSTE24 (L-006104-00-0005), along with a non-targeting siRNA (D-001810-10; ON Target plus SMART pool, Dharmacon, USA). The inhibition or overexpression of the miRNAs was achieved by commercial antisense miRNAs or mature miRNAs for hsa-miR-141-3p with an appropriate miRNA precursor-negative control [mature miR-141-3p mimic-1 no. PM 10860 and anti-miR-141-3p inhibitor no. AM 10860, and miRNA precursor-negative control no. 1; Ambion, USA]. The siRNA, mature miRNA and anti-miRNA transfections were performed using the Dharmafect transfection reagent (Dharmacon) according to the manufacturer's instructions. Briefly, the cells were seeded at a concentration of 2×104 cells/well, and the siRNA, miRNA and anti-miRNA-containing media (without the addition of antibiotics) were added when the cells reached 50–60% confluence. The cells were incubated with 50 nM siRNA, 50 nM anti-miRNA or 50 nM mature miRNA for 48 hours. To investigate the long-term effects of inhibition, the cells were subcultured 48–72 hours following the siRNA, anti-miRNA or mature miRNA transfection. Subcultured cells were stabilized for 24 hours and incubated with siRNA, anti-miRNA or mature miRNA, at the same concentrations, for 48–72 hours. After inhibition, RNA extraction and the subsequent real-time qPCR or SA β-gal staining were performed for genetic or characteristic analyses, respectively.
Cultured cells were fixed in 4% paraformaldehyde and permeabilized with 0.2% Triton X-100 (Sigma Aldrich, USA). The cells were then incubated with 10% normal goat serum (Zymed Laboratories Inc., USA) and stained with antibodies against lamin A (monoclonal, Abcam, 1∶300), prelaimin A (polyclonal, Santa Cruz, 1∶200), ZMPSTE 24 (polyclonal, Abcam, 1∶200), p16Ink4A (polyclonal, Abcam, 1∶200), γ-H2AX (polyclonal, Abcam, 1∶200), followed by incubation for 1 hour with an Alexa-Fluor-488- or Alexa-Fluor-594-labeled secondary antibody (1∶1000; Molecular Probes, USA). The nuclei were stained with Hoechst 33258 (1 µg/ml; 10 minutes), and the images were captured using a confocal microscope (Eclipse TE200, Nikon, Japan).
Chromatin immunoprecipitation assays
The chromatin immunoprecipitation (ChIP) assays were performed according to the manufacturer's protocol (ChIP assay kit, Upstate Biotechnology, USA). To prepare for the ChIP testing, the hMSCs (1–2×107 cells per IP) were fixed with 1% formaldehyde for 10 minutes; the solution was then neutralized by the addition of 1/20 volume of 2.5 M glycine for 5 minutes. The cells were washed with ice-cold PBS and scraped with SDS lysis buffer (1% SDS, 10 mM EDTA and 50 mM Tris, pH 8.1) containing protease inhibitors. The lysates were sonicated to shear the DNA to lengths between 200 and 1000 base pairs, and the samples were centrifuged for 10 minutes at 15,000 g at 4°C to remove the insoluble material. The supernatant was diluted 10-fold in the ChIP dilution buffer [0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris-HCl (pH 8.1) and 167 mM NaCl], and the chromatin was immunoprecipitated using antibodies, according to the manufacturer's instructions. Real-time qPCR was performed at a final template dilution of 1∶50. The primer sequences used in the ChIP assays in this study are given in supplementary material Table S1.
Measurement of the proliferative potential and cell cycle distribution
The effects of replicative senescence, progerin, ZMPSTE24 inhibition and miR-141-3p inhibition on hMSC proliferation were measured using the MTT assay, as described above (Jung et al., 2005).
Flow cytometry cell cycle analyses using propidium iodide staining was also performed as previously described. Briefly, hMSCs in the exponential growth phase were transfected with siRNA or miRNA and then harvested by trypsinization. The cells were washed with ice-cold PBS and then fixed with 70% ethanol at −20°C and stained with 50 µg/ml of propidium iodide in the presence of 100 µg/ml RNase A for 30 minutes. The cell cycle distribution was analyzed using the FACS Calibur system (Becton Dickinson, Franklin Lakes, NJ, USA).
For miRNA target validation, the entire 3′UTR sequence of human ZMPSTE24 was amplified by PCR and cloned into a T-vector (Promega, Madison, WI, USA, no. A1360). The 3′UTR was subcloned into pmirGLO Dual-Luciferase vector (Promega, no. E1330) using the restriction enzymes, XhoI and SalI. The 293FT cells were seeded 24 hours prior to transfection in 24-well plates at 50% confluence. The control constructs and ZMPSTE24 3′UTR reporter constructs were co-transfected along with 50 nM of the miRNAs (Ambion) using Dharmafect, following the manufacturer's instruction. After 24 hours of transfection, the firefly and Renilla luciferase activities were measured using a luminometer with Dual-Glo Luciferase Assay System (Promega, E2920). The firefly luminescence was normalized to the Renilla luminescence.
All of the experiments were conducted at least in triplicate, and the results are expressed as the means ± s.d. Statistical analyses were conducted using an analysis of variance (ANOVA), followed by Duncan's multiple range tests or Student's t-test. A value of P<0.05 was considered significant (*P<0.05; **P<0.01).
K.-R.Y.: conception and design, collection and/or assembly of data, data analysis and interpretation, manuscript writing. S.H.L.: conception and design, collection and/or assembly of data, manuscript writing. J.-W.J.: conception and design, manuscript writing. I.-S.H.: data analysis and interpretation, manuscript writing. H.-S.K.: collection and/or assembly of data, data analysis and interpretation. Y.S., T.S.: data analysis and interpretation. K.-S.K.: conception and design, administrative support, final approval of manuscript, manuscript writing.
This work was supported by the Bio and Medical Technology Development Program [grant number MEST 2010-0020265]; a National Junior Research Fellowship from the National Research Foundation funded by the Ministry of Science, Information and Communications Technology and Future Planning, Korea [grant number 2012H1A8002383]; the Research Institute for Veterinary Science, Seoul National University; the Research Institute for Agriculture and Life Sciences, Seoul National University.