Transplantation of mesenchymal stem cells improves age-related ovarian dysfunction via regulating the local renin-angiotensin system

doi:10.21203/rs.3.rs-4569611/v1

Background: the context and purpose of the study

Methods: how the study was performed and statistical tests used

Results: the main findings

Conclusions:

Age-related reproductive aging is a natural and irreversible physiological process, and delaying childbearing is increasingly common all over the world. Transplantation of umbilical cord-derived mesenchymal stem cells (MSCs) is considered a new and effective therapy to restore ovarian function, but the relevant mechanisms remain unclear. In recent years, it has been found that there is a local renin-angiotensin system (RAS) in human ovary and it plays a key role.In our research, local RAS of ovary, which is independent of circulating RAS, is affected by age and related to ovarian function. Furthermore, the in vivo(mice) and in vitro (KGN cells) experiments were designed to confirm that transplantation of MSCs improves age-related ovarian dysfunction by the local RAS. Together, our findings indicate that a novel possible mechanism to explain how stem cells restore age-related ovarian dysfunction.

Biological sciences/Stem cells/Ageing

Health sciences/Medical research/Stem-cell research

Renin-angiotensin system

Mesenchymal stem cells

Age-related ovarian dysfunction

Inflammation

Oxidative stress

According to the World Population Outlook 2022 issued by the United Nations, women's mean age of childbearing around the world has generally increased [1]. The age-related reproductive aging is a natural and irreversible physiological process, especially for oocytes [2]. Apart from strong advocation of age-appropriate reproduction, few redeemable therapies are effective in the age-related ovarian dysfunction. Fortunately, the rapid development of regenerative medicine brings promising hope in recent years, and the transplantation of mesenchymal stem cells (MSCs) is considered a new and effective therapy [3]. Searching on ClinicalTrials.gov, as of 30 August 2023, 25 clinical trials on SCs therapy for the treatment of premature ovarian failure/insufficiency (POF/POI) have been registered. Among them, the umbilical cord-derived MSCs are considered as an ideal source, furthermore, they have been proved effective by a large number of scientific and clinical research [4]. However, the relevant mechanisms remain unclear.

Ovary as the second largest source of renin in human, the ovarian RAS has been confirmed by a large number of studies. The local RAS emphasizes a complex network of precursors, peptides, enzymes and receptors [5], which is involved in regulating ovarian function, oogenesis and ovarian diseases or abnormalities via paracrine and autocrine [6, 7]. Local RAS has been proved to be the key to regulating repair and regeneration in epidermal and vasculogenesis related progenitor cells [8, 9]. Based on the above, the present study was devised to verify the role of local RAS in MSCs transplantation improves age-related ovarian dysfunction. Furthermore, the possible mechanisms of inflammation and oxidative stress were discussed. According to our results, there are reasons to believe that MSCs transplantation MSCs improves age-related ovarian dysfunction through regulating the local RAS on inflammation and oxidative stress.

Inclusion criteria

Women who received IVF merely because infertility caused by male factors in the First Affiliated Hospital of Soochow University, Jiangsu Province, China from January 2021 to February 2022.

Exclusion criteria

①The body mass index (BMI) less than 18.5 kg/m² or greater than 23.9 kg/m²; ②In the past 3 months, used drugs that are prohibited or with caution in pregnant women, such as anti-psychotic, anti-epileptic, anti-tubercular, anti-tumor drugs, etc.; ③In the past 6 months, received allogeneic blood transfusion, transplantation, stem cell therapy and immunotherapy; ④Patients with ovarian diseases or abnormalities, such as polycystic ovarian syndrome, premature ovarian failure/premature ovarian insufficiency, ovulatory dysfunction, luteinized unruptured follicle syndrome, etc.; ⑤Patients with uterine diseases or abnormalities, such as endometriosis, adenomyosis, hysteromyoma, endometrial polyps, thin endometrium, etc.; ⑥Patients with tubal or pelvic diseases, abnormalities or inflammation, such as hydrosalpinx, salpingemphraxis, salpingitis, pelvic inflammatory disease, etc.; ⑦Chromosome number and/or structural abnormalities in one of the spouses; ⑧Patients with cancer; ⑨Patients with ovarian hyperstimulation syndrome (OHSS) during controlled ovarian stimulation (COS); ⑩Incomplete information.

COS protocols and IVF

Selection of COS protocols and collection of follicular aspirates were performed strictly according to the latest China expert consensus and guide. Briefly, women under-went individualized COS according to age, AFC and FSH levels. Leuprolide acetate or cetrorelix acetate are used to block estrogen feedback to the hypothalamus, leaving the body at a high estrogen level. Recombinant FSH and/or highly purified human urinary gonadotropin are used for ovarian stimulation at step-down doses starting with 225 to 300 IU, followed by ultrasound monitorization. When at least one dominant follicle up to 18 mm or two up to 17 mm in diameter is found, recombinant human chorionic gonadotrophin (r-hCG) 250 μg by intramuscular injection. Follicles were aspirated about 35h after hCG injection and the oocytes were identified in a culture dishusing a stereomicroscope. Insemination was performed by conventional IVF or ICSI, based on the presence and location of two pronuclei, and then oocyte fertilization was assessed after 18–20 h. According to the current cleavage-stage scoring criteria for embryo morphology, the features of cell number, degree of fragmentation, equality of size and shape of blastomeres and multinucleationsize were observed under high magnification (×200) on the third day after insemination, and then that were classified as transplantable embryos and high-quality embryos.

Collect the clear, yellowish and bloodless follicular fluid for the first puncture, that from follicles 18-22 mm in diameter and the oocyte retrieval successful. Immediately centrifuge at 4 ℃ for 3,000 r/min for 15min, and then use ELISA or test kit from Jiangsu Meimian Industrial Co.,Ltd. (Jiangsu, China) to determine the levels of RAS components, inflammation and oxidative stress indexes, sex hormone. Including renin, ACE, ACE2, AngⅡ, Ang(1-7), IL-6, IL-10, ROS, MDA, SOD and GSH. At the same time, information was collected for research from medical records that does not contain personal identification. The information includes ①General clinical data: age, duration of infertility, BMI, dosage of gonadotropins (Gn) used, length of stimulation; ②Basic serum sex hormone level: E₂, FSH, LH, P, PRL, T; ③AMH; ④AFC; ⑤IVF laboratory outcomes: the number of oocyte retrieval count and the rate of metaphase Ⅱ oocyte maturation, 2 pronucleus embryos, transplantable embryos, high-quality embryos.

Isolation, culture and identification of MSCs

MSCs isolation: The full-length umbilical cords from healthy full-term pregnant women who underwent cesarean sections were selected and placed in a biosecure transport box for transportation at a low temperature of 2~8 ℃. The umbilical cord was sectioned into small segments of approximately 5~8 cm in length, thoroughly rinsed with PBS buffer, and two umbilical arteries and one umbilical vein were excised, followed by the separation of the epidermis. The tissues were cut into fragments of about 1 cm3 in size and cultured in DMEM/Low Glucose culture medium supplemented with 10% FBS and 1% Penicillin-Streptomycin under the conditions of 37 ℃ and 5% CO₂. The culture medium was replaced every 3~4 d, with continuous monitoring of cell growth.

Identification of cellular immunophenotype: 0.25% trypsin digestion, centrifugation at 800-1000 rpm, and resuspension in PBS to achieve a concentration of 5~10 ×10⁶ cells/cm². 100 μl of the cell suspension was dispensed into flow tubes, to which about 5 μl of each of the antibodies CD29, CD44, CD90, CD14, CD34, and HLA-DR were added. The cells were then incubated at ambient temperature, protected from light, for 15~30 min. Following this, the cells were washed 2~3 times with PBS buffer, collected by centrifugation, and finally resuspended in 500 μl of PBS.

Osteogenic-induced differentiation: Cells were collected using the same trypsinization method as described above and then seeded in gelatin-coated six-well plates at a density of 2×10⁴ cells/cm². After this, 2 ml of osteogenic induction medium was added, and the fluid was changed every 3~4 d. Calcium nodules were closely monitored, and alizarin red staining was performed after approximately 2~4 weeks.

Adipogenic-induced differentiation: Cells were collected using the same trypsinization method as described above and then seeded in gelatin-coated six-well plates at a density of 2×10⁴ cells/cm². The cells were cultured in DMEM/Low Glucose medium until they reached 100% confluence or over-confluence under the microscope. At this point, the medium was replaced with lipidogenic induction medium solution A. After 3 d, the medium was replaced with solution B and then after a further 24 hours, it was returned to solution A. This cycle of alternating induction was repeated 3-5 times. Cell culture was then continued with solution B for 4~7 d before being stained with Oil Red O.

MSCs transplantation

Using the same trypsinization method as described above, cells were collected. Mice were then intravenously injected in the tail vein with a cell suspension containing 1×10⁶ of hUC-MSCs. This transplantation was carried out in four separate passages, each separated by 1 week.

Animals

A total of 24 inbred SPF grade female C57BL/6 mice were used, including 8 at 6 weeks of age (17~19 g) and 16 at 40 weeks of age (26~30 g). The above experimental animals were purchased from Hangzhou Ziyuan Experimental Animal Technology Co., Ltd. (Zhejiang, China), with license No. SCXK (Zhejiang) 2019-0004 and certificate of conformity No. 20210802Abzz0105000375. The animals were housed in the SPF grade laboratory animal room of Soochow University, Jiangsu Province, China with the indoor temperature ranging from 20 to 26 ℃, the daily temperature difference less than 4 ℃, the relative humidity maintained at 40~70%, the noise less than 60 dB, and the light time from 8:00 to 20:00. The mice had free access to water and were fed with standard mouse chow, ensuring an adequate water supply and changing the bedding every 2 to 3 days. This experiment meets the requirements and conditions of SPF grade animal feeding management.

Acquisition of mouse serum and ovarian tissue

The animals were weighed on an electronic balance and the weights were recorded. After the mice were executed and placed on ice, the left and right ovarian tissues were rapidly dissected and removed. Connective tissues such as fat and fascia were removed on filter paper moistened with PBS buffer, then the tissues were rinsed 2~3 times with PBS and blotted dry. The weight of each ovarian tissue was measured on a microbalance and recorded.

Hematoxylin-eosin staining

Mouse ovarian tissues were completely immersed in 4% polymerized formaldehyde solution and fixed for 24 h. Gradient alcohol dehydration was performed:

75% C₂H₅OH	1 h (35 ℃)
85% C₂H₅OH	1 h (35 ℃)
95% C₂H₅OH Ⅰ	1 h (35 ℃)
95% C₂H₅OH Ⅱ	1 h (35 ℃)
100% C₂H₅OH Ⅰ	30 min (35 ℃)
100% C₂H₅OH Ⅱ	30 min (35 ℃)

Xylene transparent was performed:

C₈H₁₀ Ⅰ	20 min (35 ℃)
C₈H₁₀ Ⅱ	20 min (35 ℃)

Tissues are dipped in wax for 3-5 hours and then paraffin embedded and sectioned. Dewaxing was performed:

C₈H₁₀ Ⅰ	15 min
C₈H₁₀ Ⅱ	15 min

Rehydration was performed:

100% C₂H₅OH Ⅰ	5 min
100% C₂H₅OH Ⅱ	5 min
95% C₂H₅OH Ⅰ	5 min
95% C₂H₅OH Ⅱ	5 min
85% C₂H₅OH	5 min
75% C₂H₅OH	5 min
H₂O	3-5 min

Hematoxylin staining for 5-10 min, hydrochloric acid alcohol differentiation for 3-5 s, eosin staining for 2-3 min. Dehydration and sealing were performed:

75% C₂H₅OH	5 min
85% C₂H₅OH	5 min
95% C₂H₅OH Ⅰ	5 min
95% C₂H₅OH Ⅱ	5 min
100% C₂H₅OH Ⅰ	5 min
100% C₂H₅OH Ⅱ	5 min
C₈H₁₀ Ⅰ	5-10 min
C₈H₁₀ Ⅱ	5-10 min
Neutral mounting medium to seal the slide	Long-term preservation

The morphology of the tissue was visualized microscopically and all levels of follicles were counted, including primordial, primary, secondary, antral/mature and atretic follicles.

qRT-PCR

Mouse ovary tissue RNA was extracted and reverse transcribed to cDNA. Primers were designed as follows:

TARGET GENE	PRIMER	NUCLEOTIDE SEQUENCE
m-Renin	F	GCCCTCTGCCACCCAGTAA
m-Renin	R	CAAAGCCAGACAAAATGGCCC
m-Agt	F	GGTCTCTTTCTACCTTGGATCC
m-Agt	R	GACCTTGTGTCCATCTAGTCG
m-Ace	F	CTACCCCCAAGCATCTATACAG
m-Ace	R	CCACTCCTGGTTATAGTTCTCC
m-Ace2	F	GTGTACAAAGGTCACAATGGAC
m-Ace2	R	CTTCATTGGCTCCGTTTCTTAG
m-Agtr1a	F	CACTGTTTGCGCTTTTCATTAC
m-Agtr1a	R	AGCCTTCTTTAGAGCTTTCCAT
m-Agtr1b	F	CCCTGGCTGATTTATGCTTTTT
m-Agtr1b	R	GCGATCTTACATAGGTGATTGC
m-Agtr2	F	CTGTCTCAAAGAAGGAATCCCT
m-Agtr2	R	ATGTTGGCAATGAGGATAGACA
m-Mas	F	CCACTTGTCCATTGCTGATATC
m-Mas	R	GTCACCGATAATGTCACGATTG
m-Gapdh	F	AAAATGGTGAAGGTCGGTGTG
m-Gapdh	R	TGAGGTCAATGAAGGGGTCGT

The Power Up SYBR Green Master Mix premix, forward primer, reverse primer and ddH₂O were added in the proportions shown in the table below:

COMPONENTS	10 μl SYSTEM
2× Power Up SYBR Green Master Mix	5 μl
Forward primer	0.5 μl
Reverse primer	0.5 μl
cDNA	1 μl
ddH₂O	Make up to 3 μl

Add 9 μl of reaction solution and 1 μl of sample cDNA to each well of the 96-well plate, and do 3 replicates for each sample. The PCR instrument parameters were set according to the table below:

NAME	TEMPERATURE	TIME (min:sec)	REMARK
Holding	95 ℃	5:00
Cycling (40 cycles)	95 ℃	0:10	Record
Cycling (40 cycles)	60 ℃	0:30	Record
Melt curve stage	95 ℃	0:15	Record
	60 ℃	1:00
	95 ℃	0:15

The program was run, the data was exported when it was finished and a record of the experiment was kept.

Western blot

Protein lysate was prepared by mixing RIPA with PMSF (100 mM) at a ratio of 100:1. Then, 20 μl of the protein lysate was added to 1 mg of tissue for grinding. The lysate was kept on ice for 30 min and centrifuged at 15,000 rpm for 10~15 min at 4 °C. The supernatant was collected. SDS loading buffer was added and the system was metal bathed at 100 °C for 15~20 min. The gel was dispensed and the electrophoresis tank was assembled with the prepared protein samples. Electrophoresis was performed at 80 V for 30~50 min, followed by a change to 100~120 V for 2 h. The wet transfer system was then assembled and electrified at 200 mA at a constant current for 2 h. The membrane was blocked in a solution of 5-10% skimmed milk on a shaking bed for 1~2 h at ambient temperature, followed by incubation with the primary antibody at ambient temperature for 12 h with shaking. Depending on the properties of the primary antibody, the membrane was then incubated with the corresponding secondary antibody for 1 h at ambient temperature with shaking. Protein bands on the membrane were visualized using an ECL luminescent agent in a dark room.

Cell counting kit-8

The cells were collected using the same trypsinization method as described above, and inoculated into 96-well plates at a density of 2000 cells/100 μl/well. They were then incubated at 37 ℃ in a 5% CO₂ environment for 12 hours. Media with a gradient concentration of AngⅡ or Ang(1-7) were added to the wells and incubated at 37 ℃ and 5% CO₂ for varying durations, as per the experimental design. Each incubation time was repeated five times. Each well was then incubated with 10 μl of CCK-8 solution for 0.5~4 h. The absorbance at 450 nm (OD) was detected by an enzyme counter and recorded.

Enzyme linked immunosorbent assay

The standard was diluted according to the table below:

HUMAN	STANDARD 1	STANDARD 2	STANDARD 3	STANDARD 4	STANDARD 5
Renin (pg/ml)	10	20	40	80	160
ACE (ng/L)	20	40	80	160	320
ACE2 (ng/L)	4	8	16	32	64
AngⅡ (ng/L)	2.5	5	10	20	40
Ang(1-7) (ng/L)	3	6	12	24	48
E₂ (pmol/L)	3	6	12	24	48
P (pmol/L)	100	200	400	800	1600

MOUSE	STANDARD 1	STANDARD 2	STANDARD 3	STANDARD 4	STANDARD 5
E₂ (pmol/L)	12.5	25	50	100	200
FSH (IU/L)	1	2	4	8	16
LH (pg/ml)	150	300	600	1200	2400
AMH (pg/ml)	11.25	22.5	45	90	180
Renin (pg/ml)	15	30	60	120	240
ACE (ng/L)	10	20	40	80	160
ACE2 (ng/L)	5	10	20	40	80
AngⅡ (ng/L)	30	60	120	240	480
Ang(1-7) (ng/L)	22.5	45	90	180	360

Blank control wells were prepared without any sample or standard. In the standard wells, 50 μl of each standard 1~5 was added respectively. 10 μl of each group of samples was added to the test wells, with at least 3 replicates created for each sample. They were then incubated at 37 ℃ for 30 min and subsequently washed 3-5 times. After 50 μl of enzyme reagent was added to each well, they were again incubated at 37 ℃ for 30 min, followed by another wash. 50 μl each of colorants A and B were added to every well, and after incubating in the dark for 10 min at 37 ℃, 50 μl of termination solution was added. The zero point was set using the blank wells, and the absorbance (OD) of each well at 450 nm was measured and recorded.

Reactive oxygen species by DCFH-DA assay

he DCFH-DA staining solution was prepared and added in the dark to the six-well plate inoculated cells. The plate was then incubated at 37 ℃ with 5% CO₂ for 20 min. After incubation, the wells were washed 2-3 times in the dark using FBS-Free medium and then observed under a fluorescence microscope in the dark. Images were taken from three randomly selected, non-overlapping fields of view in each quadrant of each well.

Senescence and acid β-galactosidase staining

The fixative was added to the six-well plate inoculated with cells and fixed at ambient temperature for 15 min, then washed three times with PBS buffer. The Acid-β-Gal staining solution was prepared according to the following table:

Acid β-Gal staining A	10 μl
Acid β-Gal staining B	10 μl
Acid β-Gal staining C	930 μl
X-Gal staining	50 μl

The SA-β-Gal staining solution was prepared according to the following table:

Senescence-Associated β-Gal staining A	10 μl
Senescence-Associated β-Gal staining B	10 μl
Senescence-Associated β-Gal staining C	930 μl
X-Gal staining	50 μl

According to the experimental design, 1~2 ml of Acid-β-Gal staining solution or SA-β-Gal staining solution was added to each well, and incubated at 37 ℃ for 12 h. Observations were made under an ordinary light microscope, and images were taken from three randomly selected, non-overlapping fields of view in each quadrant of each well.

Mitochondrial membrane potential by JC-1 assay

JC-1 staining solution was prepared, added in the dark to the six-well plate inoculated with cells, and incubated at 37 ℃ with 5% CO2 for 20 min. Then, JC-1 staining buffer was added in the dark and the plate was washed 2-3 times. After that, the plate was observed under a fluorescence microscope in the dark. Images were taken from three randomly selected, non-overlapping fields of view in each quadrant of each well.

Quantification and statistical analysis

The correlation between hFF-RAS and age was analyzed by simple linear regression, and multivariate linear regression was used to further analyze the correlation between hFF-RAS and ovarian function, inflammation, oxidative stress indexes and IVF laboratory outcomes. General clinical data did not obey normal distribution and were statistically described in quartiles (p25, median, p75).

The organ coefficient of ovaries = ovarian weight (mg) / body weight (g). The formula for analyzing the qRT-PCR results of mRNA transcription levels can be referred to as follows: 2^-^△△^ct=2^{- [(Average ct value of the target gene in the test group)-(Average ct value of the reference gene in the test group)]- [(Average ct value of the target gene in the control group)-(} ^{Average ct value of the reference gene in the control group)].} The Western blot results for protein translation levels were analyzed using Image J software to measure the grayscale values. The CCK-8 detection results were statistically analyzed using the reference formula: The cell proliferation rate = [(Experimental well absorbance - Blank well absorbance) / (Control well absorbance - Blank well absorbance) × 100%. The average fluorescence intensity of Immunofluorescence images was analyzed by Image J software. The positive cell count for β-Gal staining was performed using Image J software. Data that followed a normal distribution were described statistically as mean ± standard deviation (x ± s). For data that followed a normal distribution and had homogeneous variances, a one-way analysis of variance (ANOVA) was used for between-group comparisons, and the least significant difference (LSD) method was applied for multiple comparisons. If the data did not have homogeneous variances, Tamhane's T2 method in a nonparametric test was chosen. A p-value less than 0.05 indicated a statistical difference, while a p-value less than 0.01 indicated a statistically significant difference.

Ovarian RAS is influenced by age and independent of circulating RAS

According to the inclusion and exclusion criteria of experimental design, 139 human follicular fluid (hFF) samples available for analysis were obtained (Fig.S1A). Refer to Table 1 for statistical description of general clinical data, and simple linear regression was used for correlation analysis. There was a linear relationship between age and renin, angiotensin converting enzyme (ACE), angiotensin Ⅱ (AngⅡ), ACE/ACE2, AngⅡ/Ang(1–7) in hFF respectively, but there was a correlation between age and renin, ACE, ACE2, AngⅡ, Ang(1–7), AngⅡ/Ang(1–7) in plasma respectively (P < 0.05). In addition, in order to further explore the relationship between ovarian RAS and circulating RAS, each component was analyzed in one-to-one correspondence. A significant correlation between the above both was merely found in renin (P < 0.01) (Fig. 1A).

Table 1

Statistics of general clinical characteristics of 139 hFF samples
Charactristics	p25	Median	p75
Age (years old)
Wife	28.0	31.0	34.0
Husband	30.0	32.0	35.0
Duration of infertility (years)	1.0	3.0	5.0
Wife's BMI (kg/m²)	20.7	22.0	23.8
AMH (ng/ml)	2.16	3.15	4.49
AFC (counts)	10.0	14.0	19.0
Basal sexhormone
FSH (IU/L)	6.82	7.94	9.10
LH (IU/L)	3.40	4.32	5.23
E₂ (pg/ml)	24.80	32.10	39.80
P (ng/ml)	0.37	0.52	0.70
PRL (ng/ml)	10.48	12.98	20.30
T (ng/ml)	0.35	0.46	0.58
RAS level in hFF
Renin (pg/ml)	147.30	161.40	181.10
ACE (ng/L)	330.80	380.00	528.50
ACE2 (ng/L)	59.16	65.09	71.71
AngⅡ (ng/L)	41.74	49.31	71.41
Ang(1–7) (ng/L)	41.80	45.17	50.11
Dose of Gn used (IU)	1800.0	2100.0	2700.0
Length of stimulation (days)	8.0	9.0	10.0
IVF laboratory outcome
Oocyte retrieval (counts)	4.0	7.0	10.0
MⅡ maturation (%)	71.4	77.8	88.9
2PN embryos (%)	75.0	100.0	100.0
Transplantable embryos (%)	0.0	20.0	50.0
High-quality embryos (%)	0.0	0.0	100.0
General clinical characteristics with non-normal distribution are statistically described by quartile. p25, the value at the 25% position, namely 1st quartile (Q1); p75, the value at the 75% position, namely 3rd quartile (Q3); BMI, body mass index; AMH, anti-müllerian hormone; AFC, antral follicle counting; FSH, follicle stimulating hormone; LH, luteinizing hormone; E₂, estradiol; P, progesterone; PRL, prolactin; T, testosterone; RAS, renin-angiotensin system; ACE, angiotensin converting enzyme; ACE2, angiotensin converting enzyme 2; AngⅡ, angiotensin Ⅱ; Ang(1–7), angiotensin (1–7); Gn, gonadotropins; IVF, in vitro fertilization; MⅡ, maturation oocyte (metaphase Ⅱ); 2PN, bipronuclear embryos.

In the analysis of RAS components one by one, there was a significant negative correlation between age and renin (r=-0.3133, P < 0.01), ACE (r=-0.1836, P < 0.05), AngⅡ (r=-0.2186, P < 0.01), while there was a nonlinear correlation between age and ACE2, Ang(1–7) in hFF (P > 0.05). In addition, further analysis based on the physiological function of RAS shows that there was a significant negative correlation between age and ACE/ACE2 (r=-0.3192, P < 0.01), AngⅡ/Ang(1–7) in hFF (r=-0.2243, P < 0.01). Analogously, there was a significant negative correlation between age and renin (r=-0.5768, P < 0.01), ACE (r=-0.2617, P < 0.01), ACE2 (r=-0.3551, P < 0.01), AngⅡ (r=-0.3552, P < 0.01), Ang(1–7) (r=-0.5213, P < 0.01), and there was a significant positive correlation between age and AngⅡ/Ang(1–7) (r = 0.1814, P < 0.05) in plasma. At last, there was a nonlinear correlation between ovary and circulation in most RAS components, including renin, ACE, ACE2, AngⅡ, Ang(1–7), ACE/ACE2 and AngⅡ/Ang(1–7) (P > 0.05). However, the correlation between ovarian and circulating RAS was only manifested in renin (r = 0.2521, P < 0.01) (Fig. 1B). Refer to Fig.S1B for negative results or more details.

Ovarian RAS is related to ovarian function, IVF laboratory outcomes, inflammation and oxidative stress indexes

The multiple regression model constructed by ovarian RAS and age can explain the variation of indexes partly, including 53.62% follicle stimulating hormone (FSH), 9.74% luteinizing hormone (LH), 64.07% antral follicular count (AFC), 17.94% anti-Müllerian hormone (AMH), 77.52% oocyte retrieval counting (ORC), 43.01% oocyte metaphase Ⅱ ratio (MⅡ%), 36.59% bipronuclear embryos ratio (2PN%), 50.28% transplantable embryo ratio (tE%), 49.75% high-quality embryo ratio (hqE%), 76.06% reactive oxygen species (ROS), 45.45% malondialdehyde (MDA), 9.03% interleukin-6 (IL-6) and 40.70% superoxide dismutase (SOD) (P < 0.05). Nevertheless, this model cannot explain more containing estradiol (E₂), progesterone (P), prolactin (PRL), testosterone (T), glutathione (GSH), interleukin-10 (IL-10) (P > 0.05) (Fig. 2A). The residuals of the above multiple regression model were independent (D-W tests is close to 2), and there was no multicollinearity in this model (VIF < 5) (Fig.S2A).

Detailedly speaking, FSH was related to age (B = 0.3585, P < 0.01), ACE (B=-0.0060, P < 0.01), ACE2 (B = 0.0342, P < 0.05), AngⅡ (B = 0.0388, P < 0.01) and Ang(1–7) (B=-0.0519, P < 0.05); LH was related to AngⅡ (B = 0.0325, P < 0.01) and Ang(1–7) (B=-0.0623, P < 0.01); AFC was related to age (B=-0.8689, P < 0.01), ACE2 (B=-0.0824, P < 0.01) and Ang(1–7) (B = 0.0761, P < 0.05); ORC was related to age (B=-0.6023, P < 0.01), renin (B = 0.0276, P < 0.01) and ACE (B = 0.0060, P < 0.01); MⅡ% was related to age (B=-1.8360, P < 0.01), AngⅡ (B = 0.1686, P < 0.05) and Ang(1–7) (B = 0.2910, P < 0.05); ROS was related to age (B = 0.1070, P < 0.01), renin (B = 0.0032, P < 0.05) and AngⅡ (B = 0.0049, P < 0.01); MDA was related to age (B = 0.0474, P < 0.01) and AngⅡ (B = 0.0035, P < 0.05); SOD was related to age (B=-0.4819, P < 0.01) and Ang(1–7) (B = 0.0711, P < 0.05) (Fig. 2B). Refer to Fig.S2B for negative results or more details.

Transplantation of MSCs after isolation, culture and identification improves ovarian function indexes in mice

The human umbilical cord-derived MSCs were successfully obtained according to our own previous research (Fig. 3A) [10]. After the Wharton's Jelly of umbilical cord was cultured for about 7 days, the fibroblast-like spindle cells growing on culture dish were observed under microscope (Fig. 3B). After about 2 weeks, the cells entered the logarithmic growth phase, and a large number of cells were clustered into “clone spheres” (Fig. 3C). Stable cell population can be observed after 3–5 passages, and the cell morphology of the primary generation can still be maintained even 15 passages later.

The obtained cells can directionally differentiate into osteoblasts in osteoblastic induction medium, and a large number of calcium deposition on the cell surface can be dyed reddish brown by the von Kossa staining (Fig. 3D). At the same time, the obtained cells can directionally differentiate into adipocytes in adipogenic induction medium, and a large number of intracellular lipid droplets can be dyed bright red by the Oil Red O staining (Fig. 3E). Furthermore, the immunophenotyping of the stable cell population were detected by flow cytometry, showing positive expression of CD 29, CD 44 and CD 90 and negative expression of CD14, CD34, and HLA-DR (Fig. 3F).

Depending on the timing (35 years old) of human reproductive aging, 10-month-old mice were correspondingly selected [11], and modeling was performed according to the protocol (Fig. 3G). Briefly, the MSCs transplantation includes four courses of treatment each one week apart, and the complete modeling was one week after the fourth transplantation. The serum sex hormone are a series of major indexes to reflect ovarian function. The Old group had increased FSH and LH levels, and decreased E₂ and AMH compared to the Control group (P < 0.01). After MSCs transplantation, there was a decrease in FSH and LH levels, and an increase in E₂ and AMH to some extent (P < 0.05) (Fig. 3H).

MSCs improves ovarian histomorphology and inflammation & oxidative stress in mice

The ovary was a pair of oval solid organs with an unclear boundary between the medulla and cortex. A large number of nerves, blood and lymphatic vessels were mainly distributed in the medulla, and the cortex contained follicles at different developmental stages. According to morphological changes, the follicles can be divided into primordial, primary, secondary, mature and atretic follicles (Fig. 4A 1–5). The ovarian structure and function have obvious changes with age. The Old group had decreased the number of primordial, primary, secondary and mature follicles, and increased atretic follicles compared to the Control group (P < 0.05) (Fig. 4B). In addition, the overall ovarian structure has shrunk, and the cortex has become thinner, so the ovary/body weight has decreased significantly (P < 0.01) (Fig. 4C). After MSCs transplantation, there was an increase in the number of primordial, secondary and mature follicles, and a decrease in atretic follicles (P < 0.01). Forthermore, the MSCs group had increased the ovary/body weight (P < 0.01).

Age-related reproductive aging is a natural and irreversible physiological process, and inflammatory and oxidative stress play an irreplaceable role in it. The Old group had increased ROS, MDA and IL-6 levels, and decreased SOD, GSH and IL-10 in ovarian homogenate compared to the Control group (P < 0.01). The results from the mouse ovary were consistent with hFF, which showed that inflammatory and oxidative stress indexes increased significantly with age. In the MSCs group, there were a decrease in ROS, MDA and IL-6 levels, and an increase in SOD, GSH and IL-10 compared to the Old group (P < 0.05) (Fig. 4D). Stem cells significantly restored the level of inflammatory and oxidative stress indexes in aging mouse ovary, which provided the premise and clues for further research.

Role of ovarian local RAS in regulating repair of MSCs

Firstly, we analyzed the circulating RAS components. Although the Old group had all decreased renin, ACE, ACE2, AngⅡ and Ang(1–7) in serum compared to the Control group (P < 0.05), there were no statistical difference in the above components in the MSCs group (P > 0.05), just as the previously mentioned human circulating RAS (Fig. 5A). Meanwhile, we analyzed the ovarian local RAS. In the Old group had decreased renin, ACE and AngⅡ (P < 0.01), but merely renin was increased after MSCs transplantation (P < 0.05) (Fig. 5B). In addition, we analyzed the RAS components ratio according to physiological function. In the Old group, circulating AngⅡ/Ang(1–7) increased while ovarian AngⅡ/Ang(1–7) decreased compared to the Control group (P < 0.05) (Fig. 5C). It is further confirmed that the ovarian RAS was independent of circulating RAS.

Further research on the level of transcription and translation were conducted. Note that different descriptions were based on the level of mRNA and protein: the AngⅡ type 1 receptor was described as AGTR1 at mRNA and AT1R at protein; the AngⅡ type 2 receptor was described as AGTR2 at mRNA and AT2R at protein. Moreover, MasR is the academic name of Ang(1–7) receptor. Although the Old group had increased AGTR1a and AGTR1b, and decreased renin, ACE and AGTR2 mRNA, while merely AGTR1a and AGTR1b were decreased after MSCs transplantation (P < 0.05) (Fig. 5D). Similarly, there were an increase in AT1R and a decrease in renin and AT2R protein in the Old group, but only AT1R was decreased in the MSCs group (P < 0.05) (Fig. 5E).

RAS regulate inflammatory and oxidative stress of KGN cells co-cultured with MSCs

The results showed that H₂O₂ inhibited the KGN cells in manner of concentration- and time-dependent. It was the closest to half maximal inhibitory concentration (IC50) when KGN was treated with 10^− 4 M H₂O₂ for 4 h (49.98%) (Fig. 6A). At the same time, the cellular viability of KGN was inhibited by AngⅡ or Ang(1–7) in manner of concentration- and time-dependent, such as 65.36% and 39.55% cells were inhibited by AngⅡ and Ang(1–7) with 10^− 4 M for 36 h, respectively. For treatment of 10^− 4 M AngⅡ, the cellular viability was similar whether 24 h (68.69%) or 36 h (65.36%). When KGN was treated with 10^− 4 M Ang(1–7) for 24 h (53.46%), the cellular viability was the closest to IC50 (Fig. 6B). Based on the above results, the optimal concentration and time of AngⅡ and Ang(1–7) were determined as 10^− 4 M for 24 h in subsequent experiments.

After intervention on KGN cells according to the above results (Fig. 6C), the culture medium was tested. In the Old group had all decreased the production of E₂ and P by KGN compared to the Control group, while but E₂ and P levels were increased after co-cultured with MSCs (P < 0.01). Further adding AngⅡ or Ang(1–7) into the co-culture system, P level increased and the effect of Ang(1–7) was more significant (P < 0.05). Nevertheless, there was no statistical difference in E₂ level no matter which one was added (P > 0.05) (Fig. 6D). Meanwhile, the Old group had increased levels of ROS, MDA and IL-6, and decreased SOD, GSH and IL-10 in culture medium compared to the Control group, but all of these indexes recovered after co-cultured (P < 0.01). There were an increase in MDA and a decrease in SOD, GSH and IL-10 after adding AngⅡ in the co-culture system (P < 0.05). In addition, there were an increase in IL-10 and a decrease in ROS after adding Ang(1–7) (P < 0.05) (Fig. 6E).

In addition to the culture medium, further research was performed at the intracellular level. The intracellular ROS (Fig. 7A) and the proportion of oxidative stress damaged KGN cells (Fig. 7B) increased, and the mitochondrial membrane potential (MMP) (Fig. 7C) decreased in the Old group (P < 0.01). After co-culture with MSCs, the intracellular ROS and the proportion decreased, and MMP increased (P < 0.01). Additionally, the intracellular ROS and the proportion further decreased, and MMP further increased when add Ang(1–7) into the co-culture system (P < 0.05). Nevertheless, there was no statistical difference in the intracellular ROS, the proportion and MMP after adding AngⅡ (P > 0.05).

Not only were RAS components detected in hFF, but ovarian RAS was proved to be independent of circulating RAS. At first, it was thought that RAS was just an endocrine system composed of substrates or enzymes from different tissues [12]. Later, it was gradually discovered that relatively independent RAS in almost all organs and tissues can play a local role in many organs and tissues through paracrine/autocrine [13–19]. Of course, this also includes the reproductive system [6–7, 20–21]. In the human ovary, local RAS is involved in many important physiological processes such as supporting oogenesis, regulating ovulation function, synthesizing steroid hormones, and mediating gonadotropin function [7, 21–23]. It is an extremely complex process for oocytes from maturation to expulsion, and a constantly changing intrafollicular microenvironment is crucial [24–25]. Up till now, two key polypeptides in RAS as well as their enzymes and receptors constitute two core pathways in this system, ACE-AngⅡ-AT1/2R and ACE2-Ang(1–7)-MasR, which restrict each other and jointly maintain the relative stability of the internal environment [26–28].

It should be pointed out that the positive results of renin dose not refute the independence of ovarian RAS. The reason is that the ovary is the second largest source of circulating renin, and ovarian stimulation activates local RAS to produce more renin [29]. This actually means that the local RAS can affect circulating RAS under special circumstances, just as ovarian hyperstimulation syndrome (OHSS). The RNA-seq results of tissue-specificity of renin protein-coding gene show that the ovarian RPKM is second only to kidney (BioProject: PRJEB2445). There are a large number of renin original substrates AGT in hFF with healthy oocytes [30–31], and the content ranks 11th-14th [32]. As an enzyme located in the initiation link of RAS, renin regulates the activity of the whole system. The latest cohort study shows that the increase of prorenin related to larger oocyte, faster cleavage and better implantation [33]. Renin levels in hFF also show differently in a variety of ovarian diseases or abnormalities according to a large number of proteomic analyses [34–36], and all above have provided the premise for the important physiological role of ovarian RAS.

We have comprehensively analyzed the relationship between ovarian RAS and age, ovarian function indexes, IVF laboratory outcomes. The results showed that the level of hFF-RAS is influenced by age, and the AngⅡ axis is more significantly affected by age than the Ang(1–7) axis. It is suggested that renin and AngⅡ axis in follicles of young women have high activity, while Ang(1–7) axis activity is stable at a certain level. A negative correlation was found between the ratio of AT1R/AT2R and age in a study on human granulosa cells (GCs) [37], which provided detail for studying the relationship between ovarian AngⅡ axis and age. Moreover, the increased activity of circulating AngⅡ axis / Ang(1–7) axis may be a factor that predisposes elderly women to vascular risk. Meanwhile, hFF-RAS has a good correlation with gonadotropin and AFC, and higher AngⅡ and Ang(1–7) in hFF predicted more oocyte retrieval and higher maturation rate. Therefore, correct recognition of reliable biomarkers in hFF will be conducive to clinical assessment of oocyte quality and prediction of IVF outcome [38].

Although several RAS components changed in older ovary, MSCs merely improved the expression of AT1R. It is fresh and bold speculation that the effectiveness of stem cell therapy is achieved by the local RAS, based on its activity in many organs or tissues and even cancers. This conjecture was first verified in cardiovascular progenitor cells which have the closest relationship with RAAS [39]. Then what is getting more and more attention is that cancer stem cells, which are considered the fundamental cause of metastasis and recurrence, are similarly regulated by the local RAS [40–42]. As the core of ACE-AngⅡ-AT1/2R axis, AngⅡ has important effects on follicle growth and oocyte maturation, which has been verified in many mammals [43–47]. A recent study has provided new evidence, which showed that the serum AT1R autoantibody of female infertility increased [48]. Moreover, ACE I/D and AT1R 1166A > C gene polymorphisms were associated with the occurrence of PCOS and POI, which may involve the direct effects of gene mutations and the indirect effects of RAS signaling pathway [49–52].

Inflammation and oxidative stress proved to be the key to ovarian RAS after MSCs transplantation. Because the position of RAS is relatively upstream, involving multiple potential targets, we want to find the most possible specific mechanism of stem cells therapy. Professor Yufang Shi, an Academician of the European Academy of Sciences, has long-term exchanges and cooperation with us. In their research, MSCs exert their therapeutic effects largely through their paracrine actions, and their communication with the inflammatory microenvironment is an essential part of this process [53–55]. The age-related ovarian dysfunction is essentially a concrete manifestation of reproductive aging, the relationship between inflammation, oxidative stress and reproductive aging is more credible [56–57]. At the same time, during severe inflammatory physiology and diseases, the RAS is an important part of inflammatory response regulation [58–59]. By considering the above evidence from multiple sources, it is confirmed that inflammation and oxidative stress are breakthrough in the further research.

The role of Ang(1–7) is revealed without the control of the gonadal axis, which can enhance MSCs to improve inflammation and oxidative stress of GCs. Undoubtedly, the gonadal axis is decisive for the regulation of ovary, so the role of other branches is discovered to be difficult, and the in vitro experiment is considered as an ideal model. We observed that RAS can affect the inflammatory and oxidative stress indexes in the culture medium of co-culture system and Ang(1–7), as the core of ACE2-Ang(1–7)-MasR axis, showed the prominent role at the intracellular level. Since it was first proved that the Ang(1–7) axis exists in the human ovary in 2011 [22], the most valuable research result is the discovery that Ang(1–7) is positively correlated with oocyte maturation rate [60]. Animal models provide more evidence that Ang(1–7) through MasR has effects on the establishment of dominant follicles [61], response to gonadotropin [62–63], the regulation of local steroid production and ovulation [64–66].

Although the relationship between ovarian RAS and circulating steroid hormone has not been observed, it is demonstrated that ovarian RAS merely promotes GCs to produce P. In fact, the relationship between ovarian RAS and reproductive endocrine has been studied for a long time, whereas this is still a dilemma [67–68]. There was a previous study on AngⅡ was consistent with our results, however Ang(1–7) was not verified when the importance of Ang(1–7) axis has not been realized [69]. A study provides a new clue for this, which shows that GCs differentiation and steroidogenic enzyme enzymes are regulated by AngⅡ [70]. Another verified that the expression of AT II was decreased after adding E₂ in ovarian follicle in vitro maturation [71]. In addition, gonadotropin showed a significant correlation with hFF-RAS in our results. This is consistent with the results of previous studies, which clarify that gonadotropin mediates the activation of ovarian RAS [60, 62], and local RAS may be an intermediate for the gonadotropin work in ovary [63].

Limitations of the study

The real relationship between RAS and steroid hormones may be covered up or confused, because of the hFF collected after ovarian stimulation. The IVF laboratory outcomes of embryos were not unexpected, as uncontrolled male factors confounded the regression models from the moment of fertilization. We can not collect the FF from healthy couples, after all, they can successfully conceive naturally. In addition, the in vitro experiment may exaggerate the role of stem cells in inflammation and oxidative stress, because H₂O₂ was used to construct the aging model of KGN cells. Ovarian RAS is a large, complex, precise and dynamic system. Although we have observed gratifying results, the relationship between the AngⅡ axis and Ang(1–7) axis has not been fully clarified, so it may be more meaningful to analyze the ovarian RAS as a whole probably. In the future, a large number of high-quality multi-center, large-sample, prospective clinical cohort studies and animal/cells model studies are still needed to further clarify the biological process of local ovarian RAS. At the same time, it is necessary to design a multi-center randomized controlled clinical trial to explore effective and feasible transplantation opportunities, and to provide scientific basis for the early application of stem cell technology in clinics.

In the observational clinical studies, local RAS of ovary, which is independent of circulating RAS, is affected by age and related to ovarian function, inflammation, oxidative stress indexes and in vitro fertilization laboratory outcomes. The in vivo experiments in mice were designed to confirm that transplantation of human umbilical cord mesenchymal stem cells improves age-related ovarian dysfunction by the local RAS. Furthermore, the role of RAS in regulating inflammation and oxidative stress in the co-culture system was explored in in vitro experiments on KGN cells (Fig. 8). Together, our findings indicate that a novel possible mechanism to explain how stem cells restore age-related ovarian dysfunction.

Acknowledgements

We thank Prof. Yufang Shi, an Academician of the European Academy of Sciences, for his guidance in this study, and KGN cells obtained from the kind gift of Professor Li Chen, Clinical School of Medical College, Nanjing University.

Author contributions

L.W., L.B., H.G. and C.M. conceived the project. L.W. wrote the main manuscript text and prepared figures and tables. L.W., L.B. and C.L. performed most of the experiments and analyzed the data. N.Y. and H.G. optimized stem cells culture and performed experiments with MSCs. L.B. and A.Z. collected the human follicular fluid. L.W. and X.L. collected the medical history information. W.J. and F.Q. performed image analyses. L.W., L.B. and C.M. supervised the project. All authors commented on previous versions of the manuscript and all the authors have read and approved the final manuscript.

Funding

This work was funded by grants from the National Natural Science Foundation of China (81671535), Postgraduate Research & Practice Innovation Program of Jiangsu Province (KYCX24_3340), Jiangsu Key Discipline of Human Assisted Reproduction Medicine Foundation (FXK202149), Suzhou Major Project Research (20220901).

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Ethics approval and consent to participate

This study was performed in line with the principles of the Declaration of Helsinki, and written informed consent was obtained from all individual participants before being enrolled in the study. The study design, protocol and informed consent were approved and adopted by the Ethics Committee of the First Affiliated Hospital of Soochow University. The project “Sample collection of human follicular fluid” was approved on August 16, 2022 with approval number of 2022(323), and the project “Sample collection of human umbilical cord” was approved on August 16, 2022 with approval number of 2022(324).

Consent for publication

Not applicable.

Competing interests

All authors declare no competing interests.

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Fig.S1.tif
Supplementary figure 1. Correlation analysis of age and RAS components in follicular fluid or circulating plasma (negative results) (A) The flow chart showing the study identification and selection process. (B) Details of simple linear regression analysis with statistical significance, including renin, ACE, ACE2, AngⅡ, Ang(1-7), ACE/ACE2 and AngⅡ/Ang(1-7). RAS, renin-angiotensin system; hFF, follicular fluid; human ACE, angiotensin converting enzyme; ACE2, angiotensin converting enzyme 2; AngⅡ, angiotensin Ⅱ; Ang(1-7), angiotensin (1-7).
Fig.S2.tif
Supplementary figure 2. Correlation analysis of RAS components and ovarian function indexes, inflammation & oxidative stress indexes (negative results) (A) Collinearity diagnostics of multiple linear regression analysis. (B) Details of multiple linear regression analysis with statistical significance, including AMH, E₂, P, PRL, T, 2PN%, tE%, hqE%, IL-6, GSH and IL-10. AMH, anti-müllerian hormone; E₂, estradiol; P, progesterone; PRL, prolactin; T, testosterone; 2PN%, bipronuclear embryos ratio; tE%, transplantable embryo ratio; hqE%, high-quality embryo ratio; IL-6, interleukin-6; GSH, glutathione; IL-10, interleukin-10; B, regression coefficient; 95%CI, 95% confidence interval; ACE, angiotensin converting enzyme; ACE2, angiotensin converting enzyme 2; Ang Ⅱ, angiotensinⅡ; Ang(1-7), angiotensin (1-7).
Fig.S3.tif
Supplementary figure 3. Western blot gels of ACE
Fig.S4.tif
Supplementary figure 4. Western blot gels of ACE2
Fig.S5.tif
Supplementary figure 5. Western blot gels of AGT
Fig.S6.tif
Supplementary figure 6. Western blot gels of Renin
Fig.S7.tif
Supplementary figure 7. Western blot gels of AT1R
Fig.S8.tif
Supplementary figure 8. Western blot gels of AT2R
Fig.S9.tif
Supplementary figure 9. Western blot gels of MasR
Fig.S10.tif
Supplementary figure 10. Western blot gels of GAPDH

Transplantation of mesenchymal stem cells improves age-related ovarian dysfunction via regulating the local renin-angiotensin system

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