Upregulation of p16INK4A promotes cellular senescence of bone marrow-derived mesenchymal stem cells from systemic lupus erythematosus patients

Cellular Signalling 24 (2012) 2307–2314

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Upregulation of p16 INK4A promotes cellular senescence of bone marrow-derived mesenchymal stem cells from systemic lupus erythematosus patients Zhifeng Gu a,⁎, 1, Xiaolei Cao b, 1, Jinxia Jiang a, 1, Liren Li a, Zhanyun Da a, Hong Liu c, Chun Cheng d,⁎⁎ a

Department of Rheumatology, Affiliated Hospital of Nantong University, Nantong, China Department of Pathology, Medical College, Nantong University, Nantong, China Department of Hematology, Affiliated Hospital of Nantong University, Nantong, China d Department of Immunology, Medical College, Nantong University, Nantong, China b c

a r t i c l e

i n f o

Article history: Received 4 May 2012 Received in revised form 28 June 2012 Accepted 14 July 2012 Available online 20 July 2012 Keywords: Mesenchymal stem cells Systemic lupus erythematosus Senescence p16INK4A Immune regulation

a b s t r a c t Previous studies have indicated that bone marrow-derived mesenchymal stem cells (MSCs) from patients with systemic lupus erythematosus (SLE) exhibited impaired proliferation, differentiation, and immune modulation capacities. Thus, MSCs may be associated with the pathogenesis of SLE. The aim of this study was to determine whether MSCs from SLE patients were senescent and to determine the mechanism underlying this phenomenon. MSCs from both untreated and treated SLE patients showed characteristics of senescence. The expression of p16INK4A was significantly increased, whereas levels of CDK4, CDK6 and p-Rb expression were decreased in the MSCs from both untreated and treated SLE patients. Knockdown of p16INK4A expression reversed the senescent features of MSCs and upregulated TGF-β expression. In vitro, when purified CD4+ T cells were incubated with p16INK4A-silenced SLE MSCs, the percentage of regulatory T cells was significantly increased. Further, we have found that p16INK4A promotes MSC senescence via the suppression of the extracellular signal regulated kinase (ERK) pathway. p16INK4A knockdown up-regulated ERK1/2 activation. Our results demonstrated that MSCs from SLE patients were senescent and that p16 INK4A plays an essential role in the process by inhibiting ERK1/2 activation. Crown Copyright © 2012 Published by Elsevier Inc. All rights reserved.

1. Introduction Bone marrow mesenchymal stem cells (BM-MSCs) are widely studied as a cell source because of their ability to differentiate into a variety of cell types, including osteoblasts, chondrocytes, adipocytes and myoblasts [1–3]. MSCs can exert immune regulatory functions, both in vivo and in vitro, on a wide range of immunocompetent cells, such as T cells, B cells, natural killer (NK) cells, and dendritic cells [4–9]. The functional features of MSCs make them attractive as therapeutic agents for a number of autoimmune diseases, including graft-versus-host disease (GVHD) and systemic lupus erythematosus (SLE) [10–14]. SLE is a chronic autoimmune disease characterized by multi-organ involvement and a remarkable variability in clinical presentations [15]. Previous studies have found that allogenic MSC transplantation (MSCT) can be used successfully in mouse and human SLE [12–14,16–18]. However, Carrion et al. reported that autologous MSCT had no effect on disease activity in two SLE patients [19]. Previous studies revealed that MSCs from SLE patients showed ⁎ Corresponding author. Tel./fax: +86 513 81168512. ⁎⁎ Corresponding author. Tel./fax: +86 513 85051999. E-mail addresses: [email protected] (Z. Gu), [email protected] (C. Cheng). 1 These authors contributed equally to this work.

impaired capacities of proliferation, differentiation, secretion of cytokines and immune modulation [20–22]. Li et al. also reported that MSCs from SLE patients showed senescent behaviors [22]. These findings suggested that the senescence of MSCs from SLE patients may be a contributing factor to disease pathogenesis. Mostly, however, SLE patients from previous studies were treated with maintenance doses of corticosteroids and immunosuppressive agents during exacerbations; these included prednisolone, azathioprine, mycophenolate mofetil, and cyclophosphamide (CTX). These treated patients, particularly CTX-treated patients, may have confounded measurements of cell senescence [23]. To our knowledge, there have been no studies focused on the biology of MSCs in untreated patients. The signs of cellular senescence include a diminishing ability to undergo cell division, increased cell size, actin stress fibers, and senescence-associated β-galactosidase (SA-β-gal) activity in vitro. Proteins regulating cell cycles were reported to be involved in the cellular senescence process, among which p16 INK4A has been studied extensively [24]. p16 INK4A was identified as a causative factor for familial melanomas and, simultaneously, as an inhibitor for the cell cycle kinases CDK4 and CDK6, indicating its critical function as a tumor suppressor that negatively regulates the cell cycle. p16 INK4A has been recognized as a major molecule that induces premature cell senescence through a telomere-dependent or -independent

0898-6568/$ – see front matter. Crown Copyright © 2012 Published by Elsevier Inc. All rights reserved. doi:10.1016/j.cellsig.2012.07.012

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mechanism [25]. Subsequent investigations have suggested that expression of the p16 INK4A gene is closely associated with senescence of MSCs [26]. In the present study, we found that BM-MSCs from both treated and untreated SLE patients showed prominent features of senescence, characterized by increased SA-β-gal activity, disordered cytoskeleton distribution and reduced ability to upregulated regulatory T cells (Treg). We also found that the expression of p16 INK4A was significantly increased, while levels of CDK4, CDK6 and p-Rb expression were decreased in MSCs from both treated and untreated SLE patients. Knockdown of p16INK4A expression could reverse the senescent behavior of MSCs in SLE patients. Furthermore, activation of the ERK1/2 pathway was found to be involved in p16INK4A-mediated cellular senescence. 2. Materials and methods 2.1. Patients Eighteen female patients, including 9 treated SLE patients and 9 untreated SLE patients, age 14–42 years (mean 26.28 ± 8.24 years), were enrolled in the study. The patient detail summary is shown in Table 1. The SLE diagnosis was made based on the criteria proposed by the American College of Rheumatology. The Systemic Lupus Erythematosus Disease Activity Index (SLEDAI) was used to measure disease activity [27]. Using a cutoff SLEDAI score of 8, all patients were categorized as active. Twelve healthy subjects were included as normal controls. All patients were female, and their age distribution was similar to that of the cases. All patients and controls gave consent to the study, which was approved by the Ethics Committee of the Affiliated Hospital of Nantong University. 2.2. Isolation of MSCs from bone marrow and cell culture MSCs were isolated and expanded from BM taken from the iliac crest of the SLE patients and healthy subjects. Five milliliters of BM Table 1 Details of 18 SLE patients. Patient

Age

Disease duration

Current treated

1

14

2 years

2

26

2 years

3

39

10 months

4

37

3 years

5 6

34 26

4 years 18 months

7

20

1 years

8

19

1 years

9

21

16 months

10 11 12 13 14 15 16 17 18

42 28 21 32 20 24 22 15 33

3 1 2 2 1 2 3 1 4

HCQ 0.2/day Pred 15–25 mg/day Pred 5–7.5 mg/day CTX 0.4 g/2 weeks HCQ 0.2/day Pred 20 mg/day HCQ 0.2/day Pred 15–20 mg/day HCQ 0.2/day Pred 15 mg/day Pred 15 mg/day HCQ0.4/day Pred 10 mg/day CTX 0.6 g/3 weeks HCQ 0.2/day Pred 5–7.5 mg/day LEF 0.2 g/day HCQ 0.2/day Pred 5–7.5 mg/day LEF 0.2 g/day HCQ 0.2/day None None None None None None None None None

days week days days week days days day days

SLEDAI 9 8

11 10 8 13 12

18

14

9 17 19 16 21 18 17 8 12

Pred: prednisolone; HCQ: Hydroxychloroquine; CTX: cyclophosphamide; LEF: Leflunomide.

was mixed with an equal volume of phosphate-buffered saline (PBS). Then, the resuspended cells were layered over Ficoll solution (1.077 g/mL) and centrifuged at 2,000 rpm for 20 minutes at room temperature. The mononuclear cells were collected at the interface. Next, the cells were resuspended in low-glucose Dulbecco Modified Eagle Medium (L-DMEM) supplemented with 10% heat inactivated fetal bovine serum (FBS). The cell viability was determined by trypan blue exclusion. Then, the cells were counted and plated at a density of 2 × 10 7 cells per 25-cm 2 dish. The cultures were maintained at 37 °C in a 5% CO2 incubator, and the medium was changed after 48 hours and every three days thereafter. When the MSCs were confluent, the cells were recovered by the addition of 0.25% trypsin-EDTA. The cells were then replanted at a density of 1 × 10 6 cells per 25-cm 2 dish. Flow cytometric analysis showed that the cells were positive for CD29, CD44, CD105, and CD166, but negative for CD14, CD34, CD38, CD45 and HLA-DR [21]. After 3 passages, cells were used for the following studies. For cell cycle analysis, cells were fixed in 70% ethanol, rehydrated in PBS, treated with RNase A (50 μg/ml) (Boehringer Mannheim Corp, Mannheim, Germany), stained with 50 μg/ml of propidium iodide (Sigma) and kept on ice in the dark until flow cytometric analysis as previously described [28]. 2.3. SiRNAs and transfection Double-stranded RNA nucleotides targeted at human p16 INK4A and non-specific controls were obtained from Santa Cruz Biotechnology. Transfection of MSCs with duplex synthetic siRNA was performed using Lipofectamine 2000 reagents (Invitrogen) according to the manufacturer's instructions. Cells were assayed after 48 h of transfection. For mock transfection, all procedures listed above were performed in the absence of the siRNA duplex. 5'FAM-labeled non-silencing control siRNA was also used to further analyze the siRNA transfection efficiency. 2.4. CD4+ T cell isolation, co-culture and Treg cell assay Spleens were collected from the BALB/c mice, and single-cell suspensions were prepared by mechanical disruption in PBS. CD4+ T cells were isolated by magnetic sorting with dynabead-bound mouse CD4+ cells according to the manufacturer's directions (Dynal Biotech). Negatively selected cells contained an average of 99% CD4+ T cells as assessed by flow cytometric analysis with a CD4 monoclonal antibody. The cell cultures were performed in RPMI 1640 medium supplemented with 10% FCS, 1× nonessential amino acids and 1 mM sodium pyruvate. The following day, the CD4+ T cells and MSCs co-culture was transferred onto Transwell plates. The ratio of Treg among the CD4+ T cells was analyzed using a Treg assay kit. In brief, after 72 hours of co-culture, CD4+ cells were harvested and washed with PBS, resuspended in 100 μL staining buffer, divided into two aliquots (one for detection and another for hemotype control), and stained with antihuman fluorescein isothiocyanate -conjugated CD4/allophycocyanin-conjugated CD25 for 30 minutes at 4 °C in the dark. The stained cells were then washed with staining buffer. For Foxp3 expression, 1 mL fixation/permeabilization concentrate/diluent was added to each tube, which were then incubated for 60 minutes at 4 °C in the dark, washed with permeabilization buffer and blotted with intracellular phycoerythrin-conjugated anti-mouse Foxp3. PE-conjugated rat IgG2a isotype was used as a control. After incubation for 30 minutes at 4 °C in the dark, stained cells were washed with diluted permeabilization buffer twice and fixed with 1% paraformaldehyde in PBS. The data were acquired on a FACS Calibur and analyzed using Cell Quest software. 2.5. SA-β-gal assay The SA-β-gal assay was used to detect cell senescence. The SA-β-gal activity was determined using a kit from the Chemical

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Company following the manufacturer's instructions. In brief, cells were cultured on slips in the 24-well plates overnight and fixed with paraformaldehyde. After incubation with SA-β-gal overnight, the slips were washed and analyzed under the microscope.

2.6. Cell Proliferation Assay The MSCs proliferation assay was performed using the BrdU assay kit according to the manufacturer's protocol. Generally, cells were incubated with 100 μM BrdU labeling solution for 4 h at 37 °C. After removing the culture media, the cells were fixed and the DNA was denatured by FixDenat solution. The anti-BrdU-POD working solution and substrate solution were then added, and the absorbances of the samples were measured by an ELISA plate reader at 370 nm wavelength.

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2.11. Immunofluorescence assay of the skeleton of MSCs MSCs were washed once with PBS and fixed in 4% paraformaldehyde for 15 min. After permeabilization and bloking, they were incubated with fluorescein isothiocyanate-conjugated phalloidin. The stained cells were then examined by a Zeiss Confocal Laser Scanning Microscope.

2.12. Statistical analysis All the results were representative of three independent experiments. All data were expressed as the mean ± standard deviation (SD) and were analyzed by Student's t test with P values less than 0.05 considered statistically significant. All statistical analyses were performed using SPSS 11.0 software.

2.7. Flow cytometry For cell cycle analysis, cultured cells were collected and fixed in 70% ethanol for 30 minutes. After being washed with PBS and then treated with RNase A(50 μg/ml in Phosphate Buffered Saline, Sigma, USA) for 30 minutes, cells were incubated with propidium iodide (50 μg/ml, Sigma, USA) for 15 minutes and analyzed by the flow cytometry machine (BECKMAN-COULTER Co., U.S.A.). The data were acquired on a FACS Calibur (BD Immunocytometry Systems) and analyzed using Cell Quest software.

2.8. CDK4/6 activity assay in vitro The phosphorylation of histone H1 by activated Cdk4/6-associated kinase activity was detected by the level of phosphorylation of the substrate histone H1. Immune complex precipitated with anti-CDK4 antibody were washed two times with phosphate-buffered saline and resuspended in 50 μl of kinase buffer (50 mM Tris–HCl, pH 7.2, 10 mM MgCl2, 1 mM DTT) containing 100 μM ATP. One microgram of commercially purified histone H1 was then added to the immunoprecipitates, and the mixtures were incubated for 30 min at 30 °C. The reaction mixtures were resolved by SDS-PAGE, and phosphorylated histone H1 was detected by immunoblotting with anti-phosphohistone H1-specific antibody.

2.9. Detection of cytokine secretion using ELISA The concentrations of TGF-β released in the culture supernatants were measured by a specific TGF-β enzyme-linked immunosorbent assay (ELISA) according to the instruction of the manufacturer (BD PharMingen). Briefly, cells in 100 μl of medium were seeded onto 96-well plates. After 24 h, 100 μl of supernatants were harvested for ELISA assay.

3. Results 3.1. MSCs from SLE patients showed prominent feature of senescence. The primary culture of MSCs was successful in 18 cases of SLE patients and 12 cases of healthy donor. The proliferation of MSCs was measured with a BrdU assay. The result indicated that MSCs from both treated and untreated SLE patients grow more slowly than those from the control group (Fig. 1A). The growth curve is shown in Fig. 1B. On the basis of the growth curve, the population doubling time of MSCs from normal controls was faster than that of MSCs from both untreated and treated SLE patients, but there was no significant difference between the two subgroups of SLE patients. The MSCs from both treated and untreated SLE patients were larger than the control MSCs. They exhibited more numerous and longer podia. The F-actin distribution was disorderedly and assembled around the nuclear region in MSCs from both treated and untreated SLE patients (Fig. 1C). SA-β-gal was used to examine MSC senescence. We have found that the number of SA-β-gal-positive cells was notably increased in treated and untreated SLE MSCs. The cell count revealed that the number of SA-β-gal-positive cells from treated and untreated SLE patients were significantly higher than among the MSCs from control. However, we did not find a significant difference between treated and untreated SLE MSCs (Fig. 1D). Flow cytometry showed that there were more MSCs restricted in the G1 phase harvested from the treated and untreated SLE patients (74.64± 4.6%) than in the MSCs from normal persons (58.84 ± 4.8%). Further, the secretion of TGF-β was decreased in MSCs from both treated and untreated SLE patients as opposed to normal MSCs (Fig. 1E). After co-culture of MSCs with T cells for 72 hours, we have found that the ratio of Foxp3-positive Treg was decreased in both treated and untreated SLE MSCs compared with those co-cultured with normal MSCs (Fig. 1F). These data indicate that the MSCs from SLE patients were senescent cells.

2.10. Western blotting 3.2. p16 INK4A expression was up-regulated in MSCs from SLE patients Cells were washed twice with ice-cold PBS and extracted in lysis buffer for 45 minutes on ice. Equal amounts of protein were separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis and electrophoretically transferred to polyvinylidene difluoride (PDVF) membranes. Membranes were then blocked with TBST containing 5% non-fat dry milk for 1 hour and incubated with a mouse monoclonal antibody against p16 INK4A, p-Rb, ERK1/2 and phosphorylated ERK1/2 overnight. Membranes were then washed with TBST and incubated with horseradish peroxidase conjugated rabbit anti-mouse secondary antibody for 1 h. The blots were developed using an enhanced chemiluminescence kit.

Previous studies showed an essential role of p16 INK4A in cell cycle control, especially during the G1/S phase progression. During this process, p16 INK4A may inhibit CDK4/6-cyclin D1 complex expression. We found that the expression of p16 INK4A was up-regulated in the MSCs from both treated and untreated SLE patients (Fig. 2A), and the kinase activity of CDK4 and CDK6 were markedly decreased in MSCs from both treated and untreated SLE patients. Moreover, a reduced phosphorylation of Rb and decreased CDK activity in MSCs from both treated and untreated SLE patients were detected (Fig. 2B–C).

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Fig. 1. MSCs in SLE patients are senescent cells. (A) The cells were plated on six-well plates. The cell proliferation ratio was detected using the BrdU labeling assay. The absorbance was shown as the proliferation rate. MSCs from both treated and untreated SLE patients grow more slowly than those from the control group. (B)The cells were plated on six-well plates. After 1 to 4 days, cell numbers were counted. The population doubling time of MSCs from normal controls was faster than that of MSCs from both untreated and treated SLE patients. (C) MSCs were stained by FITC-Phalloidin. Immunofluorescence showed that the F-actin distribution was abnormal in MSCs from both treated and untreated SLE patients. (D) SA-β-gal was used to examine MSCs senescence. The number of SA-β-gal-positive cells obviously increased among treated and untreated SLE MSCs. There was no significant difference between treated and untreated SLE-derived MSCs. (E) The secretion of TGF-β was detected by ELISA, and the concentration of TGF-β was decreased in MSCs from both treated and untreated SLE patients. (F) After co-culture of MSCs with T cells for 72 h, the ratio of Foxp3-positive Treg cells was decreased in both treated and untreated SLE MSCs compared to the co-cultured normal MSCs. All data are expressed as the mean±SEM. Significant differences of MSCs from SLE patients compared with the normal controls are considered when ⁎ Pb 0.05.

3.3. Knockdown of p16 INK4A expression reversed the senescent features of MSCs from SLE patients In a previous study, we found that p16 INK4A was overexpressed in MSCs from both treated and untreated SLE patients, but there was no significant difference in the expression levels between the two subgroups. In the following experiment, MSCs from 2 treated SLE patients, 2 untreated SLE patients and 4 control subjects were considered. To further assess the role of p16 INK4A in MSCs senescence, we used MSCs transfected with p16 INK4A siRNA or a non-specific siRNA.

Nearly 95% of cells were transfected with the siRNA constructs. Additionally, p16 INK4A expression was considerably decreased in the p16 INK4A siRNA-transfected MSCs, although p16 INK4A was knocked down by only 90% (Fig. 3). We found that cell viability was not affected in the p16INK4A knockdown MSCs. There were less SA-β-gal-positive cells when p16INK4A expression was knocked down in MSCs from SLE patients (Fig. 4A). The cell proliferation assay showed that the proliferation rate of p16INK4A knockdown MSCs from SLE patients was restored to that of the normal MSCs (Fig. 4B). Cell-cycle analysis revealed that G1 phase arrest was reversed in p16 INK4A-knockdown MSCs from SLE

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Fig. 2. The expression of p16INK4A and its protein in MSCs from SLE patients. Cell lysates were separated by SDS electrophoresis and blotted using p16INK4A, p-Rb, and Rb antibodies. (A) The expression of p16INK4A was significantly increased in cells from SLE patients. (B) The expression of p-Rb in MSCs from SLE patients was decreased. (C) The kinase activities of CDK4 and CDK6 were determined by assessing the level of phosphorylation of the histone H1substrate. The kinase activities were decreased in MSCs from SLE patients. All data are expressed as the mean ± SEM. Significant differences of MSCs from SLE patients compared with the normal controls are considered when ⁎ P b 0.05.

patients (63.48 ±3.2%), but p16INK4A knockdown had no effect on the cell cycle distribution of BM-MSCs from normal person(58.73± 3.5%). Further, the disordered distribution of F-actin was effectively reversed after p16 INK4A knockdown in MSCs from SLE patients (Fig. 4C). Additionally, we detected increased TGF-β secretion upon p16INK4A knockdown (Fig. 4D). After co-culturing MSCs with CD4+ T cells for 72 hours, the amount of Treg was increased in p16INK4A knockdown senescent MSCs (Fig. 4E). These results implied that p16INK4A may play an essential role in MSC ageing in SLE patients. 3.4. ERK1/2 activation was critically involved in cellular senescence of MSCs from SLE patients The activation of ERK1/2 was inhibited, and ERK1/2 inhibition was reversible by p16 INK4A knockdown in senescent MSCs (Fig. 5A). When

MSCs were treated with PD98059, an ERK1/2 inhibitor, the percentage of SA-β-gal-positive cells was increased (Fig. 5B), but the cell proliferation rates and secretion of TGF-β were decreased (Fig. 5C–D). We also found that the Treg cell ratio was significantly decreased after co-culturing MSCs with T cells for 72 hours with PD98059 (Fig. 5E). These results indicated that the ERK1/2 pathway was involved in the p16 INK4A-mediated cellular senescence of SLE MSCs. 4. Discussion Cellular senescence, a state of permanent cell cycle arrest, is linked to organism aging and disease development [29]. Senescent cells display a characteristically enlarged and flattened morphology. These cells feature irreversible G1 growth arrest involving the repression of genes that drive cell cycle progression and the upregulation of

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Fig. 3. p16INK4A siRNA decreased p16INK4A expression in MSCs. Cells were transfected with p16INK4A siRNA for 24 h. RT-PCR (A) and immunoblot (B) analyses showed that p16INK4A expression was significantly decreased in MSCs.

cell cycle inhibitors such as p53/p21 and p16/Rb [30]. As shown in this study, the MSCs from SLE patients displayed prominent senescent characteristics, such as increased podia and spreading, and slower growth than those from healthy controls. These findings are similar to those of previous studies [20–22]. Many studies have described biological changes in senescent MSCs. For example, the growth rate of senescent MSCs is decreased [31]. More SA-β-galpositive cells are found among senescent MSCs [26,31–33]. We also found that the proliferation rate of MSCs from SLE patients was lower than that of MSCs from normal controls. More SA-β-gal-positive cells were found among MSCs from SLE patients than among those from normal MSCs. Senescent MSCs are reportedly larger than their younger counterparts, exhibit more podia that spread further and contain more actin stress fibers [34–36]. The F-actin was distributed in a disordered pattern and assembled mainly in the nuclear region of MSCs from SLE patients. Senescence-associated functional changes include the secretion of a variety of molecules, including proteases, cytokines, and growth factors, that can act at a distance within tissues, thereby altering the tissue microenvironment. More recently, modified cytokine/chemokine

secretion has been shown to be another key phenotype associated with senescence [30,37,38]. The secretion of TGF-β was decreased in MSCs derived from SLE patients. Compared with the co-culture with normal MSCs, the ratio of Foxp3-positive Treg cells was decreased after co-culture of the MSCs from SLE patients with T cells for 72 h. These data revealed that MSCs from SLE patients display prominent features of senescence. Thus, the senescence of MSCs may be a contributing factor to SLE pathogenesis. Interestingly, we have not found any distinct senescent characteristics in MSCs from untreated versus treated SLE patients. In addition, previous studies found that immunosuppressive therapy, particularly CTX, may lead to premature cellular senescence. In the current study, we did not detect any significant differences in senescence characteristics between MSCs from untreated versus treated SLE patients. These results indicate that MSCs from SLE patients show senescent behavior during early-stage lupus. p16 INK4A has been reported as one of the major factors that mediates the cellular senescence process. In senescent cells, the proliferation rates were decreased, which may be associated with the high level of p16INK4A expression. The binding of CDK4/6 to p16INK4A blocks the former complex's interaction with cyclin D and enhances its protein kinase activities. This kinase activity usually results in Rb protein phosphorylation and E2F activation, which may cause cell cycle arrest at G0/G1 [25]. This cell cycle regulator is not expressed in undifferentiated hESCs [39]. In the MSCs from SLE patients, the expression of p16INK4A was increased, the kinase activity of CDK4 and CDK6 were strongly decreased, and the phosphorylation of Rb was also decreased. It has been reported that p16INK4A is responsible for the senescent event in MSCs. Shibata et al. have report that p16INK4A is a key factor in regulating MSC growth [26]. A line of evidence indicated a tight association between the inactivation of p16INK4A and the transformation of MSCs. Targeting p16INK4A could delay growth arrest in MSCs. The MSCs immortalized with hTERT showed spontaneous transformation in vitro in association with the deletion of the p16INK4A gene [40]. The MSCs strain that transformed spontaneously in vitro lacked expression of the p16INK4A gene [41]. The Bmi1 protein, a polycomb group protein, represses the transcription of p16INK4A. Bmil-transduced immortalized hMSCs, which showed no detectable p16INK4A gene expression, became fully transformed after the introduction of an activated H-ras gene [42,43]. Wip1 (wild-type p53-inducible phosphatase-1), a well-studied

Fig. 4. p16INK4A knockdown reversed the ageing characteristics of MSCs from SLE patients. MSCs from SLE patients were transfected with p16INK4A siRNA for 24 h. (A) The SA-β-gal staining assay showed p16INK4A knockdown could decrease SA-β-gal activity in MSCs from SLE patients. (B) The BrdU assay showed that when p16INK4A expression was knocked down, the proliferation rate of SLE MSCs was similar to that of the normal MSCs. (C) Immunofluorescence showed that the abnormal distribution of F-actin in MSCs from SLE patients was reversed after p16INK4A knockdown. (D) The ELISA assay showed that the secretion of TGF-β was reversed in the MSCs from SLE patients. (E) Flow cytometry showed that p16INK4A knockdown increased the Treg cell ratio. Significant differences comparing the normal controls are considered when ⁎ Pb 0.05, and differences comparing the MSCs from SLE patients are considered when #Pb 0.05.

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Fig. 5. ERK1/2 activity may play an essential role in p16INK4A-induced senescence of MSCs. (A) Cells were transfected with p16INK4A siRNA and either treated or not treated with PD98059, an inhibitor of ERK1/2 signal. Western blot analysis showed that the activation of ERK1/2 was decreased in SLE MSCs and those treated with PD98059. When the expression of p16INK4A was knocked down, the activation of ERK1/2 was upregulated. (B) MSCs were treated with PD98059. β-gal staining showed that cellular senescence was increased in the presence of PD98059. (C) BrdU labeling showed that cell proliferation was suppressed in MSCs after treatment with PD98059. (D) After treatment with PD98059, the secretion of TGF-β was decreased in normal MSCs. (E) Flow cytometry showed that the Treg cell ratio was decreased in the co-culture model after treatment with PD98059. Significant differences of MSCs from SLE patients compared with the normal controls are considered when ⁎ Pb 0.05.

stress modulator, significantly lowered p16INK4A expression and apparently extended the life span of MSCs [44]. In MSCs from SLE patients, p16INK4A may play a similar role in the aging process. Jin et al. reported that hypoxic culture conditions maintained MSCs in an undifferentiated and non-senescent state through the down-regulation of p16INK4A and ERK [45]. Park found that the basal phosphorylation level of ERK was significantly increased in senescent hMSCs [31]. In the present study, we found that the ERK pathway was involved in the senescence of MSCs from SLE patients. When p16INK4A expression was knocked down, growth arrest was reversed, and the activation of ERK1/2 was increased. These data indicated that the expression of p16INK4A plays an essential role in MSCs growth arrest. Modification of the expression of specific genes may restore this biological behavior. Recently, unique immunologic properties of MSCs have been described, such as inhibition of the proliferation and cytotoxicity of NK cells, suppression of T-cell proliferation, and suppression of B-cell proliferation and differentiation [46]. Bocelli et al. reported that the MSCs from patients with autoimmune diseases, including 2 SLE patients, showed lower PBMC proliferation suppression abilities compared to MSCs from health donors [47]. Sun et al. reported that MSCs from NZBW/F1 lupus mice and SLE human patients had defective immunoregulatory function when compared with MSCs from healthy controls. CD4 + CD25+ regulatory T cells also play an essential role in maintaining immune homeostasis and preventing autoimmunity [48,49]. Therefore, defects in Treg development, maintenance or functioning have been found to be associated with several human autoimmune diseases, including SLE [50]. In this study, we found that

after co-culture of MSCs with CD4+ T cells, the Treg ratio among the MSCs from SLE patients was decreased compared to those from healthy controls. After p16 INK4A knockdown, the ratio of Treg was increased in co-cultured MSCs more than their normally expressing p16 INK4A counterparts obtained from SLE patients.In the MSCs from SLE patients, TGF-β secretion was down-regulated. This downregulation could be reversed by p16 INK4A knockdown. Although we have shown that MSCs from SLE patients and healthy controls could affect the ratio of Th1, Th2, and Th17 cells (data not shown), there were no significant differences between these two groups. Thus, these data suggested that p16INK4A may play an important role in modulating the immunoregulatory functions of MSCs. In summary, we have characterized the prominent senescent features of MSCs in both treated and untreated SLE patients. Our results indicate that p16 INK4A may play an essential role in the ageing process of MSCs from SLE patients by regulating cytokine secretion and the ERK1/2 signaling pathway.

5. Conclusion In the present study, our results have demonstrated the following: (1) MSCs from both treated and untreated SLE patients were senescent; (2) p16 INK4A plays an essential role in MSCs senescence in SLE patients by inhibiting ERK1/2 activation; (3) therefore, targeting p16 INK4A and ERK1/2 in MSCs may provide a new therapeutic strategy for treating SLE patients.

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Abbreviations MSCs mesenchymal stem cells SLE systemic lupus erythematosus MSCT MSCs transplantation CTX cyclophosphamide Pred prednisolone HCQ Hydroxychloroquine LEF Leflunomide SA-β-gal senescence-associated β-galactosidase SLEDAI Systemic Lupus Erythematosus Disease Activity Index BrdU bromodeoxyuridine ERK extracellular signal-regulated protein kinase

Conflict of interest There are no commercial affiliations or conflicts of interest to disclose. Acknowledgements The authors wish to thank Professor Lingyun Sun, Department of Rheumatology, Affiliated Drum Tower Hospital of Nanjing, and Professor Liwei Lv, Department of Pathology, University of Hong Kong, for their critical comments and advice during the preparation of this manuscript. This work was supported by grants from the Chinese National Natural Science Foundation (NO. 81172841), the Natural Science Foundation of Jiangsu Colleges and Universities Grant (09KJB320010); the “Top Six Types of Talents” Financial Assistance of Jiangsu Province Grant (No. 6); the project of Jiangsu Provincial Health Department (Z201005); the innovative project of Nantong University postgraduate students (13025043); and the Jiangsu province's Outstanding Medical Academic Leader Program (LJ201136). References [1] N. Saulnier, W. Lattanzi, M.A. Puglisi, G. Pani, M. Barba, A.C. Piscaglia, M. Giachelia, S. Alfieri, G. Neri, G. Gasbarrini, A. Gasbarrini, European Review for Medical and Pharmacological Sciences 13 (Suppl. 1) (2009) 71–78. [2] D.J. Prockop, Science (New York, N.Y.) 276 (1997) 71–74. [3] J. Sanchez-Ramos, S. Song, F. Cardozo-Pelaez, C. Hazzi, T. Stedeford, A. Willing, T.B. Freeman, S. Saporta, W. Janssen, N. Patel, D.R. Cooper, P.R. Sanberg, Experimental Neurology 164 (2000) 247–256. [4] A.J. Nauta, W.E. Fibbe, Blood 110 (2007) 3499–3506. [5] X.X. Jiang, Y. Zhang, B. Liu, S.X. Zhang, Y. Wu, X.D. Yu, N. Mao, Blood 105 (2005) 4120–4126. [6] P.A. Sotiropoulou, S.A. Perez, A.D. Gritzapis, C.N. Baxevanis, M. Papamichail, Stem Cells (Dayton, Ohio) 24 (2006) 74–85. [7] J.M. Ryan, F.P. Barry, J.M. Murphy, B.P. Mahon, Journal of Inflammation (London, England) 2 (2005) 8. [8] W.T. Tse, J.D. Pendleton, W.M. Beyer, M.C. Egalka, E.C. Guinan, Transplantation 75 (2003) 389–397. [9] A. Corcione, F. Benvenuto, E. Ferretti, D. Giunti, V. Cappiello, F. Cazzanti, M. Risso, F. Gualandi, G.L. Mancardi, V. Pistoia, A. Uccelli, Blood 107 (2006) 367–372. [10] T. Toubai, S. Paczesny, Y. Shono, J. Tanaka, K.P. Lowler, C.T. Malter, M. Kasai, M. Imamura, Current Stem Cell Research & Therapy 4 (2009) 252–259. [11] K. Le Blanc, F. Frassoni, L. Ball, F. Locatelli, H. Roelofs, I. Lewis, E. Lanino, B. Sundberg, M.E. Bernardo, M. Remberger, G. Dini, R.M. Egeler, A. Bacigalupo, W. Fibbe, O. Ringden, Lancet 371 (2008) 1579–1586. [12] J. Liang, H. Zhang, B. Hua, H. Wang, L. Lu, S. Shi, Y. Hou, X. Zeng, G.S. Gilkeson, L. Sun, Annals of the Rheumatic Diseases 69 (2010) 1423–1429. [13] Z. Gu, K. Akiyama, X. Ma, H. Zhang, X. Feng, G. Yao, Y. Hou, L. Lu, G.S. Gilkeson, R.M. Silver, X. Zeng, S. Shi, L. Sun, Lupus 19 (2010) 1502–1514.

[14] L. Sun, K. Akiyama, H. Zhang, T. Yamaza, Y. Hou, S. Zhao, T. Xu, A. Le, S. Shi, Stem Cells (Dayton, Ohio) 27 (2009) 1421–1432. [15] A. Rahman, D.A. Isenberg, The New England Journal of Medicine 358 (2008) 929–939. [16] J. Liang, F. Gu, H. Wang, B. Hua, Y. Hou, S. Shi, L. Lu, L. Sun, Nature Reviews. Rheumatology 6 (2010) 486–489. [17] K. Zhou, H. Zhang, O. Jin, X. Feng, G. Yao, Y. Hou, L. Sun, Cellular & Molecular Immunology 5 (2008) 417–424. [18] L. Sun, D. Wang, J. Liang, H. Zhang, X. Feng, H. Wang, B. Hua, B. Liu, S. Ye, X. Hu, W. Xu, X. Zeng, Y. Hou, G.S. Gilkeson, R.M. Silver, L. Lu, S. Shi, Arthritis and Rheumatism 62 (2010) 2467–2475. [19] F. Carrion, E. Nova, C. Ruiz, F. Diaz, C. Inostroza, D. Rojo, G. Monckeberg, F.E. Figueroa, Lupus 19 (2010) 317–322. [20] Y. Nie, C. Lau, A. Lie, G. Chan, M. Mok, Lupus 19 (2010) 850–859. [21] L.Y. Sun, H.Y. Zhang, X.B. Feng, Y.Y. Hou, L.W. Lu, L.M. Fan, Lupus 16 (2007) 121–128. [22] X. Li, L. Liu, D. Meng, D. Wang, J. Zhang, D. Shi, H. Liu, H. Xu, L. Lu, L. Sun, Stem Cells and Development (in press) PMID:22375903. [23] A. Palaniyappan, Biogerontology 10 (2009) 677–682. [24] J. Campisi, Cell 120 (2005) 513–522. [25] M. Serrano, G.J. Hannon, D. Beach, Nature 366 (1993) 704–707. [26] K.R. Shibata, T. Aoyama, Y. Shima, K. Fukiage, S. Otsuka, M. Furu, Y. Kohno, K. Ito, S. Fujibayashi, M. Neo, T. Nakayama, T. Nakamura, J. Toguchida, Stem Cells (Dayton, Ohio) 25 (2007) 2371–2382. [27] M.C. Hochberg, Arthritis and Rheumatism 40 (1997) 1725. [28] Q.L. Lam, C.K. Lo, B.J. Zheng, K.H. Ko, D.G. Osmond, G.E. Wu, R. Rottapel, L. Lu, International Immunology 19 (2007) 267–276. [29] J. Campisi, F. d'Adda di Fagagna, Nature Reviews. Molecular Cell Biology 8 (2007) 729–740. [30] S. Sethe, A. Scutt, A. Stolzing, Ageing Research Reviews 5 (2006) 91–116. [31] J.S. Park, H.Y. Kim, H.W. Kim, G.N. Chae, H.T. Oh, J.Y. Park, H. Shim, M. Seo, E.Y. Shin, E.G. Kim, S.C. Park, S.J. Kwak, Mechanisms of Ageing and Development 126 (2005) 551–559. [32] J.Y. Heo, K. Jing, K.S. Song, K.S. Seo, J.H. Park, J.S. Kim, Y.J. Jung, G.M. Hur, D.Y. Jo, G.R. Kweon, W.H. Yoon, K. Lim, B.D. Hwang, B.H. Jeon, J.I. Park, Stem Cells 27 (2009) 1455–1462. [33] Y. Chen, X. Xu, Z. Tan, C. Ye, Q. Zhao, Y. Chen, Archives of Medical Science 8 (1) (2012) 30–38. [34] M.A. Baxter, R.F. Wynn, S.N. Jowitt, J.E. Wraith, L.J. Fairbairn, I. Bellantuono, Stem Cells (Dayton, Ohio) 22 (2004) 675–682. [35] J.R. Mauney, D.L. Kaplan, V. Volloch, Biomaterials 25 (2004) 3233–3243. [36] K. Stenderup, J. Justesen, C. Clausen, M. Kassem, Bone 33 (2003) 919–926. [37] E.J. Moerman, K. Teng, D.A. Lipschitz, B. Lecka-Czernik, Aging Cell 3 (2004) 379–389. [38] I. Tsuboi, K. Morimoto, Y. Hirabayashi, G.X. Li, S. Aizawa, K.J. Mori, J. Kanno, T. Inoue, Experimental Biology and Medicine (Maywood, N.J.) 229 (2004) 494–502. [39] X. Zeng, M.S. Rao, Neuroscience 145 (2007) 1348–1358. [40] N. Serakinci, P. Guldberg, J.S. Burns, B. Abdallah, H. Schrodder, T. Jensen, M. Kassem, Oncogene 23 (2004) 5095–5098. [41] D. Rubio, J. Garcia-Castro, M.C. Martin, R. de la Fuente, J.C. Cigudosa, A.C. Lloyd, A. Bernad, Cancer Research 65 (2005) 3035–3039. [42] I.K. Park, S.J. Morrison, M.F. Clarke, The Journal of Clinical Investigation 113 (2004) 175–179. [43] Y. Shima, T. Okamoto, T. Aoyama, K. Yasura, T. Ishibe, K. Nishijo, K.R. Shibata, Y. Kohno, K. Fukiage, S. Otsuka, D. Uejima, T. Nakayama, T. Nakamura, T. Kiyono, J. Toguchida, Biochemical and Biophysical Research Communications 353 (2007) 60–66. [44] J.S. Lee, M.O. Lee, B.H. Moon, S.H. Shim, A.J. Fornace Jr., H.J. Cha, Stem Cells (Dayton, Ohio) 27 (2009) 1963–1975. [45] Y. Jin, T. Kato, M. Furu, A. Nasu, Y. Kajita, H. Mitsui, M. Ueda, T. Aoyama, T. Nakayama, T. Nakamura, J. Toguchida, Biochemical and Biophysical Research Communications 391 (2010) 1471–1476. [46] G.M. Spaggiari, A. Capobianco, H. Abdelrazik, F. Becchetti, M.C. Mingari, L. Moretta, Blood 111 (2008) 1327–1333. [47] C. Bocelli-Tyndall, L. Bracci, G. Spagnoli, A. Braccini, M. Bouchenaki, R. Ceredig, V. Pistoia, I. Martin, A. Tyndall, Rheumatology (Oxford, England) 46 (2007) 403–408. [48] S. Sakaguchi, Annual Review of Immunology 22 (2004) 531–562. [49] W.T. Hsu, J.L. Suen, B.L. Chiang, Clinical and Experimental Immunology 145 (2006) 513–519. [50] X. Valencia, C. Yarboro, G. Illei, P.E. Lipsky, Journal of Immunology 178 (2007) 2579–2588.