Rejuvenation of Cardiac Tissue Developed from Reprogrammed Aged Somatic Cells

Abstract

Induced pluripotent stem cells (iPSCs) derived via somatic cell reprogramming have been reported to reset aged somatic cells to a more youthful state, characterized by elongated telomeres, a rearranged mitochondrial network, reduced oxidative stress, and restored pluripotency. However, it is still unclear whether the reprogrammed aged somatic cells can function normally as embryonic stem cells (ESCs) during development and be rejuvenated. In the current study, we applied the aggregation technique to investigate whether iPSCs derived from aged somatic cells could develop normally and be rejuvenated. iPSCs derived from bone marrow myeloid cells of 2-month-old (2 M) and 18-month-old (18 M) C57BL/6-Tg (CAG-EGFP)1Osb/J mice were aggregated with embryos derived from wild-type ICR mice to produce chimeras (referred to as 2 M CA and 18 M CA, respectively). Our observations focused on comparing the ability of the iPSCs derived from 18 M and 2 M bone marrow cells to develop rejuvenated cardiac tissue (the heart is the most vital organ during aging). The results showed an absence of p16 and p53 upregulation, telomere length shortening, and mitochondrial gene expression and deletion in 18 M CA, whereas slight changes in mitochondrial ultrastructure, cytochrome C oxidase activity, ATP production, and reactive oxygen species production were observed in CA cardiac tissues. The data implied that all of the aging characteristics observed in the newborn cardiac tissue of 18 M CA were comparable with those of 2 M CA newborn cardiac tissue. This study provides the first direct evidence of the aging-related characteristics of cardiac tissue developed from aged iPSCs, and our observations demonstrate that partial rejuvenation can be achieved by reprogramming aged somatic cells to a pluripotent state.

Introduction

Aging is a complex phenomenon that occurs in nearly all organisms. The aging of mammals is primarily marked by genomic instability, telomere attrition, epigenetic alterations, loss of proteostasis, deregulated nutrient sensing, mitochondrial dysfunction, cellular senescence, stem cell exhaustion, and altered intercellular communication.^1

–5

Before the somatic cell reprogramming era, all of the endeavors to conquer aging to rejuvenate somatic cells appeared to be insufficient. The reprogramming of mouse and human somatic cells through forced expression of defined transcription factors demonstrated that adult somatic cells could be restored to pluripotency. Induced pluripotent stem cells (iPSCs) express markers exclusive to embryonic stem cells (ESCs), mimic ESC morphology and growth properties, and can differentiate into all three germ layers to produce chimeric mice.^6,7 iPSCs possess enormous therapeutic potential for aging and age-related diseases. Adult somatic cell reprogramming can be applied as a patient-specific model, and such models have been used in the study of disease pathogenesis and for the discovery and testing of potential new drugs.⁸

We noted that most of the previous studies on this topic were limited to in vitro observations, whereas only a few reports have addressed the in vivo aging characteristics of iPSCs derived from aged or senescent somatic cells.^5,9

–12 In the current study, we established iPSCs from bone marrow myeloid cells of both 18-month-old and 2-month-old C57BL/6 mice using the SCR510 | STEMCCA Constitutive Polycistronic (OKSM) Lentivirus Reprogramming Kit (Merck Millipore). The aging characteristics of the cardiac tissue derived from the 18-month-old iPSCs of chimeric newborns (18 M CA) were thoroughly investigated. Our observations showed that despite slight changes in mitochondrial ultrastructure, cytochrome C oxidase activity, ATP production, and reactive oxygen species (ROS) production in the cardiac tissue of chimeric newborns (both 18 M CA and 2 M CA), all of the evaluated aging-related characteristics of the 18 M CA were comparable with those of the 2 M CA.

Materials and Methods

Animals

C57BL/6-Tg (CAG-EGFP)1Osb/J mice were kindly provided by Dr. Rong Zhang (Division of Cancer Immunotherapy, Exploratory Oncology Research & Clinical Trial Center, National Cancer Center, Japan). ICR mice were purchased from HFK Bioscience Co., Ltd. All mice were maintained in the Animal Research Facility at the Second Xiang-ya Hospital of Central South University (China) under specific pathogen-free conditions and were used according to institutional guidelines. This study was carried out in strict accordance with the recommendations of the Regulations on Animal Experimentation of Central South University (China). The Animal Care and Use Committee of the Second Xiang-ya Hospital of Central South University (China) approved the protocol. All experiments were performed under anesthesia and were designed to minimize suffering. Mice were humanely sacrificed before tissue collection.

Cell culture

Bone marrow (BM) myeloid cells were generated according to the method of Inaba et al. and our previous protocol.^13,14 Briefly, BM cells were cultured in RPMI 1640 medium (10% fetal bovine serum [FBS]), 300 g/L L-glutamine, 100 U/mL penicillin, 100 mg/mL streptomycin, and 50 mM 2-mercaptoethanol (Sigma-Aldrich) containing 0.3% GM-CSF supernatant (from murine GM-CSF-producing Chinese hamster ovary cells, gifted by Professor Hengyi Xiao (West China School of Medicine/West China Hospital, Sichuan University). Floating and loosely adherent clustering cells were collected on day 4 and used as BM myeloid cells. SNL76/7 feeder cells (gifted by Dr. Hengyi Xiao, Sichuan University) were clonally derived from the STO cell line transfected with a G418R cassette and an leukemia inhibitory factor expression construct.¹⁵ All iPSCs were maintained via culture in KO-DMEM (Gibco) supplemented with 1% L-glutamine, 1% nonessential amino acid, 100 U/mL penicillin, 100 mg/mL streptomycin, 5.5 mM 2-mercaptoethanol, and 15% FBS at 37°C under 5% CO₂.

Reprogramming

Aged iPSCs (18 M) and 2 M BM-iPSCs were established following a modified version of the instructions of the SCR510 | STEMCCA Constitutive Polycistronic (OKSM) Lentivirus Reprogramming Kit (Merck Millipore). One day before transduction, 5 × 10⁵ BM myeloid cells (at 2 and 23 months) were briefly seeded on 1 × 10⁶ mitomycin C-treated SNL76/7 feeder cells in 6-cm dishes. After 24 hours, a virus-containing medium supplemented with 5 μg/mL polybrene was added. The BM myeloid cells were then incubated in the virus/polybrene-containing supernatants for 24 hours. This step was repeated once. After infection, the medium was replaced with the iPS medium every other day for up to 15–23 days. Clones were manually collected and expanded for further experimentation.

Chimera production and germ line transmission

The zona pellucida was removed from eight-cell embryos via brief exposure to Tyrode's solution. The denuded embryos were then washed with HEPES-buffered KSOM and transferred to four-well dishes in KSOM in a WOW system. Approximately 100 cells were subsequently selected and transferred to each well of the system for coculture with the embryos overnight. The resulting blastocysts were transferred to the uteri of pseudopregnant ICR mice at 2.5 dpc. Again, chimeras were identified based on the coat color of the pups at birth. The embryo transfer recipients were prepared by pairing mature ICR female mice with vasectomized ICR males overnight. Vaginal plugs were examined the following morning, and plugged mice were used as pseudopregnant recipients for embryo transfer. These mice were anesthetized through intraperitoneal injection of avertin (0.3 mg/g body weight), and 10–15 blastocysts were transferred to the tip of each uterine horn. Chimeras were subsequently selected and mated with ICR mice. The germ line transmission competence of iPSCs was determined based on the coat color of the resulting F1 pups (Table 1).

Table 1.

Table of Aggregation

iPSC colony used for aggregation	Transplanted aggregated embryos	GFP-positive chimera	Dead	Alive	Date of birth
18 M iPSCs Colony 1	86	4	1	3	November 5, 2014
2 M iPSCs Colony 1	142	6	2	4	March 19, 2015
18 M iPSCs Colony 3	153	6	2	4	March 20, 2015
2 M iPSCs Colony 2	133	5	2	3	May 17, 2015
18 M iPSCs Colony 4	122	6	3	3	May 26, 2015
2 M iPSCs Colony 4	126	8	6	2	June 13, 2015

Aggregation was performed using three iPSC clones in each group. For the 18 M iPSCs, colony 1, colony 3, and colony 4 were used for each of three aggregations. For the 2 M iPSCs, colony 1, colony 2, and colony 4 were used for each of three aggregations. During each aggregation, GFP-positive chimeras were selected for further examination. In total, 10 18 M chimeras and 9 2 M chimeras survived during aggregation.

iPSC, induced pluripotent stem cell.

Quantitative real-time PCR for telomere length analysis

Total genomic DNA was extracted from cardiac tissues obtained from newborn, adult (2 months), and aged (18–27 months) wild-type (WT) mice. The average telomere length was measured via quantitative real-time PCR and normalized to a single-copy gene; for this purpose, we chose the acidic ribosomal phosphoprotein PO (36B4) gene, which is well conserved and has previously been used for gene-dosage studies.¹⁶ Modified primer sequences were used for the telomeric portion of the assay. The forward and reverse mouse telomeric primer sequences were as follows: 5′ CGG TTT GTT TGG GTT TGG GTT TGG GTT TGG GTT TGG GTT 3′ and 5′ GGC TTG CCT TAC CCT TAC CCT TAC CCT TAC CCT TAC CCT 3′, respectively. The forward and reverse primer sequences for the mouse 36B4 gene were as follows: 5′ACT GGT CTA GGA CCC GAG AAG 3′ and 5′ TCA ATG GTG CCT CTG GAG ATT 3′, respectively. Each reaction for the telomere portion of the assay included 20 μL of SYBR Green PCR Master Mix and a 300 nM of each forward and reverse primer. An automated thermocycler (MX3005P) was used, with the following reaction conditions: 95°C for 10 minutes, followed by 30 cycles of data collection with 95°C denaturation for 15 seconds and 56°C annealing–extension for 1 minute. Each reaction for the 36B4 portion of the assay included 20 μL of SYBRGreen PCR Master Mix, 300 nM forward primer, and 500 nM reverse primer. The thermocycler reaction conditions were as follows: 95°C for 10 minutes, followed by 35 cycles of data collection with 95°C denaturation for 15 seconds, 52°C annealing for 20 seconds, and 72°C extension for 30 seconds.

Western blotting

Western blotting was performed as previously described.¹⁷ The primary antibodies used for these assays included mouse anti-p53 (1c12) mAb (Cell Signaling; #2524) and rabbit anti-p16 (M-156) (Santa Cruz Bio, Inc.; SC-1207). The secondary antibodies included ECLTM sheep anti-mouse IgG, horseradish peroxidase-linked whole antibody (GE Healthcare, UK Limited; lot 358934), and ECLTM donkey anti-rabbit IgG horseradish peroxidase-linked F(ab)2 fragment (GE Healthcare, UK Limited; lot 391954). Amersham ECL^™Prime was used to amplify the signal intensity, and the membranes were developed with the Western blot detection reagents (GE Healthcare, UK Limited; RPN 2232) solution A (lot 4626479) and solution B (lot 4626480).

Analysis of mitochondrial gene expression and mitochondrial DNA deletions

Total DNA was extracted from cardiac tissue samples using the phenol–chloroform–isoamyl alcohol method. The primers for the mitochondrial DNA deletion analysis were designed according to previously published reports.¹⁸ The primers used for the mitochondrial gene expression analysis and DNA deletion analysis are listed in Supplementary Table 1 (Supplementary Data are available online at www-liebertpub-com.web.bisu.edu.cn/rej).

Transmission electron microscopy

Cardiac tissue samples were sliced into 1 mm³ sections, prefixed via immersion in chilled 2.5% glutaraldehyde in phosphate buffer (pH 7.2) on ice for 1 hour, fixed in osmium tetroxide for 1 hour, and then dehydrated for 10 minutes each in a series of 50%, 70%, 80%, 90%, and 100% ethanol. Next, the samples were dehydrated three times with propylene oxide for 10 minutes each, infiltrated for 10 minutes with propylene oxide and epoxy resin (vol/vol = 1:1), embedded with EPON 812 epoxy resin, DDSA, DMP-30, and MNA resin, and aggregated for 24–48 hours at 60°C. After polymerization, 70 nm ultrathin sections were prepared using a diamond knife and Reichert-Nissei Ultracuts (Leica). Sections were then stained with uranyl acetate and lead staining solution (Sigma-Aldrich). The stained sections were photographed with a JEOL JEM-1400EX transmission electron microscope.

Cytochrome C oxidase activity analysis (COX staining)

Frozen cardiac tissue was sliced into 5 μm sections (prepared with a Leica CM 3050 S cryostat) before staining. The sections were then incubated in the following solution at 37°C for 6 hours: 4% sucrose, 0.01% cytochrome C, 0.05% DAB, 0.01% catalase C, and 50 mM Tris–HCL (pH 7.6), adjusted to a final pH of 7.4 with 1 N NaOH. The sections were subsequently rinsed three times with distilled water and postfixed for 5 minutes at room temperature in 4% PFA.

Measurement of ROS

The cardiac tissues were treated with liquid nitrogen and immediately stored at −80°C before ROS detection. ROS levels were determined using the OxiSelect In vitro ROS/RNS Assay Kit (Cell Biolabs, Inc.). This method uses 2′,7′-dichlorodihydrofluorescein DiOxyQ (DCFHDiOxyQ; a fluorogenic probe), which is deacetylated in the cytosol to the nonfluorescent 2′,7′-dichlorodihydrofluorescein (DCFH).

DCFH rapidly reacts with ROS and reactive nitrogen species (RNS) (predominantly NO and ONOO−) to form the fluorescent product 2′,7′-dichlorodihydrofluorescein (DCF). Thus, the total amount of ROS and RNS present is proportional to the intensity of the DCF fluorescence (λex = 480 nm, λem = 530 nm). Briefly, 30 mg of cardiac tissue was used and washed with ice-cold PBS. The tissues were cut into small pieces (0.1 mm3) and homogenized in 300 μL of 0.1% sodium dodecyl sulfate (SDS; dissolved in PBS) on ice. The lysates were adjusted to equal protein concentrations (range, 1–10 mg/mL) using PBS. Protein concentrations were analyzed using the Bradford assay, and ROS/RNS levels were assayed using the OxiSelect In vitro ROS/RNS Assay Kit using a calibration curve obtained from standard solutions of DCF in PBS. Absorbances were measured on a Multimode Plate Reader (EnSpire™ Multimode Plate Reader; PerkinElmer). The experiment was performed in triplicate.

Detection of ATP content

The cardiac tissues were treated with liquid nitrogen and immediately stored at −80°C before ATP detection. The ATP content of the cardiac tissues was evaluated using a luciferase ATP assay (ATP Assay Kit, S0026. Beyotime) according to the manufacturer's instructions. Thirty milligrams of cardiac tissue was used and washed with ice-cold PBS. Then, the tissue was cut into small pieces (0.1 mm³) and homogenized in ice-cold ATP lysis buffer. The homogenates were collected and centrifuged at 12,000 g for 7 minutes at 4°C, and then, the supernatant was collected and the RLU value was detected using the Multimode Plate Reader (EnSpire Multimode Plate Reader; PerkinElmer) to determine the ATP concentration using a standard curve from reference standards. The ATP content of the tissues was determined by comparing the RLU value of the samples with that of the reference standards. The experiment was performed in triplicate.

Statistical analysis

The data were analyzed using SPSS 15.0, and p < 0.05 was considered statistically significant. Grayscale values of Western blot bands were obtained with BandScan v5.0, and the data were analyzed with Student's t test. Mitochondrial diameters were measured using a JEOL JEM-1400EX transmission electron microscope, and the data were analyzed using Student's t-test. COX activity was analyzed with the chi-square test. Real-time PCR was performed a minimum of three times for each sample. Standard curves were generated according to the relative standard curve protocol supplied with the Applied MX3005P thermocycler, and the input amount of each test sample was calculated using the same protocol. For each sample, the relative input amount of telomere DNA was divided by the relative input amount of 36B4 DNA, generating the telomere-to-36B4 ratio. The average ratio for each sample was reported as the average telomere length ratio (ATLR).

Results

18 M-iPSCs and 2 M-iPSCs showed comparable pluripotent marker expression and in vitro and in vivo abilities to differentiate into three layers

We established and expanded three colonies in each group, designated 18 M-iPSC colony 1, 18 M-iPSC colony 3, 18 M-iPSC colony 4, 2 M-iPSC colony 1, 2 M-iPSC colony 2, and 2 M-iPSC colony 4.

To compare the pluripotency of 2 M-iPSCs and aged iPSCs, the expression of the pluripotent markers pou5f-1 and SSEA-1 in each cell line was determined via immunofluorescence. Similar pluripotent marker expression was observed in each cell line, as shown in representative images (Supplementary Fig. S1A). The in vitro differentiation ability of 18 M-iPSCs and 2 M-iPSCs was analyzed through hanging drop EB formation, followed by spontaneous differentiation. An endoderm marker (alpha-fetoprotein), ectoderm marker (neurofilament-H), and mesoderm marker (beta-actin) were used to determine the three germ layers derived from spontaneous differentiation. A comparable in vitro differentiation ability was observed in each cell line (Supplementary Fig. S1B). Furthermore, we compared the in vivo teratoma formation ability of 18 M-iPSCs and 2 M-iPSCs, and the results showed that both cell lines could generate teratomas containing three germ layers (Supplementary Fig. S1C). Since the 18 M-iPSCs showed a similar pluripotency to that of 2 M-iPSCs, we further concentrated on observing the senescence characteristics of each cell line in vitro and in vivo.

Aged iPSCs showed higher SA-beta-gal staining rates and slightly decreased proliferation compared with 2 M-iPSCs during in vitro differentiation into three germ layers

SA-beta-gal staining was performed to analyze the senescence of 2 M-iPSCs and 18 M-iPSCs. The cells were collected at several time points (D0, D7, D14, and D21) during in vitro differentiation. The cells exhibiting positive SA-beta-gal staining were counted, and we found that each cell line showed an increasing SA-beta-gal staining rate, and the rate was higher in the BM-18 M-iPSC lines than in BM-2 M-iPSCs at each time point (Fig. 1A, left). The cell numbers of each cell line were counted using trypan blue staining. According to the obtained proliferation curves, although the same number of cells was used at the beginning of differentiation, we found that the increase in the number of 18 M-iPSCs was slower than that of 2 M-iPSCs. Furthermore, after day 15, the 18 M-iPSCs showed a slight decrease in the cell number, while 2 M-iPSCs continued to proliferate (Fig. 1A, right).

FIG. 1.

In vitro differentiation and senescence assay. (A) SA-β-gal assay and proliferation curve during in vitro differentiation. SA-beta-gal staining was performed during the in vitro differentiation of each iPSC colony. Cells were collected at several time points (D0, D7, D14, and D21) during differentiation. The cells exhibiting positive SA-beta-gal staining were counted and analyzed. The columns on the left side show that each cell line showed an increasing SA-beta-gal staining rate, and the rate was higher in the BM-18 M-iPSC lines (dark gray) than in BM-2 M-iPSCs at each time point (light gray). The cell numbers of each cell line were counted using trypan blue staining at each time point and analyzed. The proliferation curves on the right show that although the same number of cells was used at the beginning of differentiation, the increase in the number of BM-18 M-iPSCs (dark gray) was slower than that of BM-2 M-iPSCs (light gray). After day 15, the 18 M-iPSCs showed a slight decrease in cell number, whereas the 2 M-iPSCs continued to proliferate. (B) p16 and p19 expression during differentiation. p16 and p19 expression was determined during the in vitro differentiation in each cell line. p16 showed low expression in BM-18 M-iPSCs but was not expressed in BM-2 M-iPSCs at the beginning of differentiation (D0). On day 17 of differentiation, p16 was expressed in both groups and was decreased in BM-2 M-iPSCs but not in BM-18 M-iPSCs on day 22. Regarding p19 expression, a similar pattern was observed only at the beginning of differentiation when both groups showed low expression of p19; however, only aged iPSCs expressed p19 on day 22. iPSC, induced pluripotent stem cell.

p16 and p19 expression increased in both 18 M-iPSCs and 2 M-iPSCs during the beginning stage of in vitro differentiation and decreased after day 22 in 2 M-iPSCs, but not in 18 M-iPSCs

Since p16 and p19 are considered to be related to cellular senescence and aging, their expression in each group during in vitro differentiation was determined through Western blotting.¹⁹ p16 showed low expression in BM-18 M-iPSCs but was not expressed in BM-2 M-iPSCs at the beginning of differentiation. On day 17 of in vitro differentiation, p16 was expressed in both groups and then decreased in BM-2 M-iPSCs, but not in BM-18 M-iPSCs on day 22. Regarding p19 expression, a similar pattern was observed only at the beginning of differentiation, when both groups showed low expression of p19, and only aged iPSCs expressed p19 on day 22 (Fig. 1B).

Chimeras could be derived using both BM-2 M-iPSCs and BM-18 M-iPSCs via aggregation with ICR embryos

To further analyze the aging characteristics of BM-18 M-iPSCs, chimeras were generated via aggregation of BM-2 M-iPSCs and BM-18 M-iPSCs with 8-cell-stage ICR mouse embryos. After several attempts, we failed to produce full germ line-transmitted chimeras. Although we did not produce full germ line-transmitted chimeric mice in either the BM-2 M-iPSC or BM-18 M-iPSC group, we did produce live chimeras for further observations (Table 1 and Fig. 2A). After analyzing GFP expression to determine transmission in several organs in both 2 M CA and 18 M CA newborns, we selected chimeric newborns exhibiting high expression of GFP in cardiac tissue for further investigation (Fig. 2B). The next question that we asked was whether the cardiac tissues of 18 M CA and 2 M CA possess similar aging characteristics. To address this important issue, we first analyzed the expression of aging-related proteins in the cardiac tissue from both groups and then performed comparisons with several WT C57BL/6-Tg (CAG-EGFP)1Osb/J mice of different ages (newborn, 2 months, 23 months). Consistent with previous studies, the expression of both p16 and p53 increased with cardiac aging, but we did not observe any increase in p16 and p53 expression in either 2 M CA or 18 M CA (Fig. 3A).

FIG. 2.

Characteristics of iPSC-derived chimera newborns. (A) Chimera newborns were derived by aggregation of iPSCs with ICR embryos. All of the surviving newborns were imaged under UV light to examine their GFP expression. Left: Representative picture showing a strong GFP signal was detected in this newborn derived by aggregation of BM-18 M-iPSCs with ICR embryos. Right: Representative picture showing a 5-month-old chimera derived by aggregation of BM-18 M-iPSCs with ICR embryos. The chimera possess the coat color of both the (CAG-EGFP)1Osb/J and ICR mice. (B) The vital organs, including the intestine (upper left), heart (upper right), lung (middle left), muscle (middle right), liver (lower left), and kidney (lower right), were collected from chimera newborns and were used for further examinations. Parts of the organs were fixed with OCT compound and sectioned to detect their GFP expression. Representative images are shown. Color images available online at www-liebertpub-com.web.bisu.edu.cn/rej

FIG. 3.

Aging-related investigation in iPSCs derived from chimera newborns. (A) p16 and p53 expression in cardiac tissue of newborn chimera mice. Age-related p16 and p53 expression was detected in the cardiac tissues of chimera newborns and compared with that of WT C57BL/6-Tg (CAG-EGFP)1Osb/J mice of the following ages: newborns (WT NB), 2-month-old (WT 2 M), and 23-month-old (WT 23 M) mice. The expression of p16 was positive in both the B6 2 M and B6 23 M, whereas p53 expression was positive only in B6 23 M. Neither the WT nor the chimera newborns expressed p16 or p53. All the cardiac tissues were GFP positive. (B) Age-related changes in the average telomere length within the cardiac tissues of the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WT NB), 2 M CA, and 18 M CA. Left: Total genomic DNA was isolated from the cardiac tissues and analyzed using real-time quantitative PCR. The single copy PO 36B4 gene was used to normalize the samples. Standard curve of 36B4 (gray) and telomere (black). Right: For each sample, the relative input amount of telomere DNA was divided by the relative input amount of 36B4 DNA. Each sample was run three times, and the results are reported as the ATLR. The ATLRs of the cardiac tissues from the WT newborns (WT NB; white), 2 M-iPSC-derived chimera newborns (2 M CA; gray) and 18 M-iPSC-derived chimera newborns (18 M CA; black) are shown. NS, nonsignificant; n = 3. ATLR, average telomere length ratio.

Telomere lengths were comparable in cardiac tissue from 2 M CA and 18 M CA newborns

Real-time PCR was applied to determine telomere lengths. Telomere shortening has traditionally been considered an important factor in aging.^20
–22 To measure age-related changes in telomere length in cardiac tissue from aged iPSC chimeras, we isolated genomic DNA from cardiac tissues collected from WT C57BL/6-Tg (CAG-EGFP)1Osb/J mice, 2 M CA and 18 M CA. The telomere length was analyzed via quantitative real-time PCR. The single-copy acidic ribosomal phosphoprotein PO 36B4 gene (36B4) was used to standardize our results (Fig. 3B, left). Although there were slight variations in telomere length between groups, we found no significant age-related change in the average telomere length in the aged iPSC chimera group (Fig. 3B, right).

Cardiac tissue derived from 18 M CA newborns exhibited no significant age-related changes

Mitochondrial gene expression and mitochondrial DNA deletions were analyzed using PCR, and the results showed that changes in mitochondrial gene expression (Mt-Nd1, Mt-co1, Mt-cytb) and mitochondrial DNA deletions (PL51/PL52 and PL85/PL86) in cardiac tissue could be tested during aging. The PL51/PL52 primers were used to determine the deletion of the CoIII and Nd3 sites, whereas the PL85/PL86 primers were used to determine the deletion of the 16sRNA site. We observed an increase in the prevalence of COIII and ND3 deletions in cardiac tissue isolated from adult WT mice relative to the WT newborns. The prevalence of these deletions increased further in the aged WT mice. An increase in the prevalence of 16sRNA deletions in the aged WT mice was also observed. Interestingly, we did not observe significant differences in mitochondrial gene expression or mitochondrial DNA deletions between the mitochondrial 2 M CA and 18 M CA, which indicated that at the nuclear level, cardiac tissue developed from rejuvenated 18 M iPSCs showed comparable mitochondrial DNA integration to that developed from 2 M iPSCs (Fig. 4A).

FIG. 4.

Mitochondrial gene expression, mitochondrial genes, mitochondrial DNA deletions, mitochondrial deletion, mitochondrial ultrastructure, cytochrome C oxidase activity, ATP production, and ROS production in chimera newborns. (A) Age-related changes in mitochondrial gene expression and mitochondrial DNA deletions within the cardiac tissues of WT C57BL/6-Tg (CAG-EGFP)1Osb/J mice at different ages, 2 M-iPSC-derived chimera newborns (2 M CA), and 18 M-iPSC-derived chimera newborns (18 M CA). Left: Expression of Mt-Nd1, Mt-co1, and Mt-cytb in the cardiac tissue of WT (newborn, 2-month old, and 23-month old), 2 M CA, and 18 M CA. Right: Deletions of the 16S rRNA (PL85/PL86) and cytochrome C subunit (PL51/PL52) genes in the cardiac tissues of WT (newborn, 2-month old, and 23-month old), 2 M CA, and 18 M CA. (B) Age-related changes in cytochrome C activity within the cardiac tissues of the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WT NB), 2 M CA, and 18 M CA. Upper: COX staining (brown) of the cardiac tissues isolated from the WT newborns (upper left, WT NB), 2 M-iPSC-derived chimera newborns (upper middle, 2 M CA) and 18 M-iPSC-derived chimera newborns (upper right, 18 M CA). Lower: Graphic display of the percentage of COX-positive cells in the cardiac tissues isolated from the WT newborns (light gray, WT NB), 2 M-iPSC-derived chimera newborns (dark gray, 2 M CA), and 18 M-iPSC-derived chimera newborns (black, 18 M CA). (C) Age-related changes in mitochondrial ultrastructure within the cardiac tissues of the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WT NB), 2 M-iPSC-derived chimera newborns (2 M CA), and 18 M-iPSC-derived chimera newborns (18 M CA). Left: Cardiomyocyte mitochondria from the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WT NB). Middle: Cardiomyocyte mitochondria from the 2 M-iPSC-derived chimera newborns (2 M CA). Right: Cardiomyocyte mitochondria from the 18 M-iPSC-derived chimera newborns (18 M CA). (D) Age-related changes in mitochondrial ultrastructure within the cardiac tissues of the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WT NB), 2 M-iPSC-derived chimera newborns (2 M CA), and 18 M-iPSC-derived chimera newborns (18 M CA). Left: Cardiomyocyte mitochondria from the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WTNB). Middle: Cardiomyocyte mitochondria from the 2 M-iPSC-derived chimera newborns (2 M CA). Right: Cardiomyocyte mitochondria from the 18 M-iPSC-derived chimera newborns (18 M CA). (E) ROS production (left) and ATP production (right) in cardiac tissues from the WT C57BL/6-Tg (CAG-EGFP)1Osb/J newborns (WTNB), the 2 M-iPSC-derived chimera newborns (2 M CA), and the 18 M-iPSC-derived chimera newborns (18 M CA). ROS, reactive oxygen species. Color images available online at www-liebertpub-com.web.bisu.edu.cn/rej

We next sought to observe cytochrome C oxidase activity, which is associated with oxygen consumption and the aging process.^23
–25 COX staining was performed as described previously. The number of COX-positive cells was counted and normalized to total cell numbers. Subsequent analysis revealed no significant differences in cytochrome C oxidase activity between 2 M CA and 18 M CA, but some slight differences in activity could be observed between 2 M CA and 18 M CA and wild-type newborns (Fig. 4B). The ultrastructure of the mitochondria in newborn C57BL/6-TG (CAG-EGFP)1OSB/J, 2 M CA and 18 M CA mice was observed under a transmission electronic microscope. The long and short axes of the mitochondria were measured and analyzed as described in previous reports.²⁶ We found that although cardiac tissue from 2 M CA and 18 M CA newborns showed a similar ultrastructure, the shape was slightly more rounded than that of C57BL/6-TG (CAG-EGFP)1OSB/J newborn cardiac tissue (Fig. 4C, D). ATP production and ROS production were slightly decreased in the cardiac tissue of chimera newborns (both 2 M CA and 18 M CA) compared with the levels in WT newborns (Fig. 4E). The mild changes in COX activity, ATP production, and ROS production in the cardiac tissue of chimeras suggested that there were still some differences in oxygen consumption conditions between the chimera and wild-type cardiac tissue. The results were synchronized with observations of mitochondrial ultrastructure.

Discussion

Although accumulating data have indicated that telomeres are elongated and mitochondria are reorganized in iPSCs, there are controversial results indicating that the reprogramming efficiency declines with age, while somatic cells generated from iPSCs tend toward premature senescence during in vitro differentiation.^27

–31 Hence, comprehensive testing of iPSCs and their potential aging signature should be conducted. In the current study, we sought to thoroughly investigate the aging characteristics of iPSCs derived from aged somatic cells. We found that 18 M iPSCs exhibited comparable pluripotent marker expression, pluripotency, and teratoma formation ability to those of 2 M iPSCs.^7,14,32 Increased senescence marker levels were observed during in vitro differentiation, possibly due to the increase in cell death and apoptosis during the differentiation process.³³ We did not observe any changes in p53 expression during the differentiation process (data not shown). This unique and interesting phenomenon of inconsistent aging marker expression reminds us that the key signaling pathways that are involved in senescence and function during differentiation are not exactly the same as those involved in the senescence and aging that occur during development. In addition, it should be noted that all of the observations made in the experiments only provide indirect evidence of whether the 18 M iPSCs were rejuvenated, as aging and senescence can be induced under certain culturing conditions.^34

–37

To determine whether reprogramming of aged somatic cells could lead to rejuvenation, we sought to aggregate 18 M iPSCs and 2 M iPSCs with 8-cell-stage ICR mouse embryos to derive chimeric newborns. Aggregation was performed using three iPSC clones in each group. Development and germ line transmission were investigated. The mice in the F0 18 M CA and F0 2 M CA groups that remained alive showed comparable life spans to those of WT mice (Table 1 and Fig. 2A).

In human cardiac tissue, telomeres are reported to be important in the aging process.^1,38 For example, Sahin et al. demonstrated that Tert^−/− mice exhibit early aging and heart failure.³⁹ It was observed in our experiments that telomere length was not significantly different between cardiac tissues harvested from newborn, adult, and aged mice, which may have been caused by maintenance of telomerase activity in aged rodents.⁴⁰

p53 and p16 are cell cycle regulators that cause cell cycle arrest through different, but interacting pathways, and both are upregulated by telomere shortening and cell stress.^41
–43 We determined that p53 and p16 both exhibited upregulated expression with age, but no significant differences between 2 M CA and 18 M CA were observed. In addition to its roles in DNA damage repair, cell cycle, and apoptosis, p53 has been shown to regulate cellular respiration and metabolism, while modulating the balance between mitochondrial respiration and glycolytic pathways.^44
–46 Since p53 expression was comparable between 2 M CA and 18 M CA, we further sought to investigate the age-related changes in mitochondrial gene expression and deletion as well as the mitochondrial ultrastructure and cytochrome C oxidase activity.

Mt-Nd1 has been associated with mitochondrial encephalomyopathy and stroke-like episodes.^47,48 Mt-co1 encodes a cytochrome C oxidase subunit.⁴⁹ Mt-cytb or mitochondria-encoded cytochrome B is also known as the bc1 complex or ubiquinol–cytochrome C reductase.⁵⁰ All of these mitochondrial genes showed comparable expression between 2 M CA and 18 M CA. Mitochondrial DNA integrity is compromised by aging and stress.^23,51,52 Nakada et al. reported a correlation between functional and ultrastructural abnormalities of the mitochondria in mouse hearts carrying a pathogenic mtDNA deletion mutant.⁵³ We evaluated the age-related changes in mitochondrial DNA integrity within cardiac tissue. The PL51/PL52 primers were used to determine the deletion of the CoIII and Nd3 sites, while the PL85/PL86 primers were used to determine the deletion of the 16sRNA site. We observed an increase in the prevalence of COIII and ND3 deletions in cardiac tissue isolated from the adult WT mice relative to WT newborns. The prevalence of these deletions increased further in the aged WT mice. An increase in the prevalence of 16sRNA deletions in aged WT mice has also been observed. None of these deletions was detected in 2 M CA and 18 M CA newborns, which indicated comparable mitochondrial integrity in 2 M CA and 18 M CA.

Drahota et al. demonstrated that cytochrome C oxidase (COX) activity in cardiomyocytes is an important regulator of mitochondrial respiration.⁵⁴ It has previously been found that COX activity gradually increases from birth to 6 months of age, with a peak activity occurring at 18 months of age and decreasing thereafter.²⁶ Since cardiac mitochondria generate 90% of the ATP in the heart, the metabolic activity of chimera cardiac tissue was also observed by conducting an ATP content assay.²³ Previous studies showed that ROS are a major cause for aging. An increase in ROS production or a decrease in the defense against ROS appears to be associated with the decrease in life span.⁵⁵ In the current study, we found that COX activity, ATP production, and ROS production decreased slightly in both 2 M CA and 18 M CA newborns, which implied that cellular respiration was slightly decreased in chimeric newborns. Nakada et al. reported that the mitochondrial ultrastructure could be shortened and underdeveloped due to decreased COX activity.⁵³ In the current study, the mitochondria were found to be elongated in WT newborns and slightly shortened in both 2 M and 18 M CA newborns, which may have been caused by the relatively underdeveloped mitochondrial network, in addition to low mitochondrial and COX activity in chimeric newborns.

The somatic cell reprogramming technique gave rise to completely new insights in the study of aging and senescence. It was proposed by Rando et al. that reprogramming somatic cells to a pluripotent state may be an effective strategy for rejuvenating an aged organism.⁵⁶ Alejandro Ocampo et al. proved that partial reprogramming by short-term cyclic expression of Oct4, Sox2, Klf4, and c-Myc (OSKM) ameliorates cellular and physiological hallmarks of aging and prolongs life span in a mouse model of premature aging.¹⁰ Carlos López-Otín and colleagues reported that DOT1 L inhibition in vivo extends the life span and ameliorates the accelerated aging phenotype of progeroid mice.^11,12 In chimera experiment, we found that the generated cardiac tissue showed slight differences in COX activity and mitochondrial ultrastructure compared with those of WT newborns. Nevertheless, compared with the cardiac tissue of wild-type mice and 2 M CA newborns, the cardiac tissue of 18 M CA newborns showed greater restoration of the mitochondrial structure and mitochondrial gene expression. Other chimera organs have also been further studied (data not shown). For the first time, we observed the detailed characteristics of cardiac tissue aging in aged iPSC chimeric mice. Our observations may provide new evidence related to the question of whether aged somatic cells can be completely rejuvenated through somatic reprogramming. Although we did not observe identical aging characteristics of cardiac tissue in 18 M CA newborns compared with WT newborns, we confirmed that there were no significant differences between 2 M CA and 18 M CA newborns. The controversial results from in vivo observations remind us that differentiation can only partially reflect the pluripotency of iPSCs and is not sufficient for determining the aging characteristics of aged iPSCs. More aged iPSC lines, and particularly those that can produce germ line-competent chimeric mice, should be used to produce more conclusive results in further studies.

Ethics Approval and Consent to Participate

This study was carried out in strict accordance with the recommendations of the Regulations on Human and Animal Experimentation of Central South University (China). All experimental protocols were approved by the Ethics Committee of the Second Xiang-ya Hospital of Central South University (China).

Funding

This work was supported by the National Natural Science Foundation of China 81400093.

This work was supported by the National Natural Science Foundation of China 81400343.

This work was supported by the National Natural Science Foundation of China 81470323.

Authors' Contributions

The work presented here was performed in collaboration with all authors.

Zhao Cheng, Xian-ming Fu, and Guang-sen Zhang conceived and designed the work, analyzed the data, and interpreted the results.

ZhaoCheng and Guang-sen Zhang drafted the manuscript.

Hongling Peng and Rong Zhang provided important materials and interpreted the data.

Guang-sen Zhang and Xian-ming Fu financially supported the study.

Xian-ming Fu and Guang-sen Zhang are co-corresponding authors of this article.

All authors reviewed and approved the final manuscript.

Footnotes

Acknowledgments

The authors thank Professor Hengyi Xiao (West China School of Medicine/West China Hospital, Sichuan University, P.R. China) and Professor Ken-ichi Iosbe (Department of Immunology, Nagoya University Graduate School of Medicine, Japan) for their kind support of this study.

Author Disclosure Statement

The authors declare that they have no competing interests.

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