Category Archives: Stem Cells

Ethical approval

Experimental protocols and procedures were approved by Chang Gung Memorial Hospitals Institutional Animal Care and Use Committee (No. 2019062002), Institutional Board Review (IRB: 201800954B0) and funded by the National Science and Technology Council Grants (MOST 107-2314-B-182A-103-). The study period took place from August 1st, 2018 to January 31st, 2020.

All procedures involving humans were carried out in accordance with relevant guidelines and regulations, and approved by Institutional Board Review Chang Gung Memorial Hospital. Informed consent was obtained from all participants/donor.

All experimental procedures were performed under the supervision of a licensed veterinarian, in a manner consistent with the regulations of the National Institute of Health of Taiwan. All animal related procedures were approved by the Institutional Animal Care and Use Committee of Chang Gung Memorial Hospital (IACUC Approval No.: CGMH2019062002). All methods involving animals are reported in accordance with ARRIVE guidelines.

The hAFSCs were obtained from freshly collected amniotic fluid by routine amniocentesis from healthy pregnant donors at 1520 gestational weeks. Cells were cultured in StemPro MSC Serum free medium supplemented with 10% fetal bovine serum (Invitrogen, Carlsbad,CA) and incubated at 37C with 5% carbon dioxide. Culture medium was changed every 34days. The specific surface antigens of hAFSCs were characterized using flow cytometry analyses. The cultured cells were trypsinized and stained with phycoerythrin (PE)-conjugated antibodies against CD90 (BD PharMingen,CA). The cells were analyzed using the Calibur flow cytometer (Becton Dickinson, Heidelberg, Germany). Passage 4 to 6 hAFSCs were collected and prepared to a final concentration of 3106 cells/0.3mL Phosphate Buffer Solution (PBS). Thereafter, 3106 hAFSCs were seeded on a sterile mesh-scaffold and cultured for 3days prior transplantation. This is accordance to the previous work by Liang et al.12.

Three types of absorbable materials were compared, comprising AlloDerm RTM; PLGA mesh (VICRYL) and PDS mesh. The characteristic of each mesh is displayed in Table 1. Cell line from amniotic fluid stem cells was cultivated with basic fibroblast growth (bFGF). To measure the cells ability to proliferate, EdU Assay (Click-iTEdU Assay, Invitrogen, Life Technologies Corporation, Carlsbad, CA, USA) is incubated with hAFSC, and meshes-harvested with hAFSc. EdU Assay (5-ethynyl-2-deoxyuridine) works as a nucleoside analog of thymidine and is incorporated into DNA during active DNA synthesis. Procedure was performed in accordance to protocol (Supplementary, S1). DNA staining is performed for imaging and analysis. hAFSCs seeded mesh-scaffold were incubated at 37C in 5% CO2 for 60min, followed by Dulbeccos modified Eagle medium. MTS (5-(3-carboxymethoxyphenyl)-2-(4,5-dimethyl-thiazoly)-3-(4-sulfophenyl) tetrazolium, inner salt assay)13 colorimetric assay test were conducted on day 7 and day 14 for cell metabolic activity (Fig.1). DAPI (Santa Cruz Biotechnology, Santa Cruz, CA, USA) Stain for immunofluorescence imaging were also conducted on day 14. The most suitable mesh with hAFSC growth was determined by immunofluorescence assay and scanning electron microscopy (SEM) via LIVE/DEAD Viability/cytotoxicity Kit on day 7 and day 14. SEM enables direct microscopic imaging of the material properties on the surface sample, that offers adjustable magnification and large field depth. In this study, spot charge-coupled device color digital camera (Olympus DP72, Tokyo, Japan) was used to obtain immunohistochemistry images under 20objective (Olympus BX-51, Tokyo, Japan) and immunofluorescence under Leica TCS SP8X confocal laser scanning microscope (Leica Microsystem, Heidelberg, Germany) with appropriate filters for DAPI. Camera was interfaced with Image-Pro Plus Software (Media Cybernetics, Silver Spring, MD, USA). This is in accordance as previous study conducted by Liang et al.14.

Total of 28 SpragueDawley rats, with the mean age 12.31.7weeks old and weighing 298.227.1g were treated and cared for under the supervision of a licensed veterinarian, in a manner consistent with the regulations of the National Institute of Health of Taiwan. All animal related procedures were approved by the Institutional Animal Care and Use Committee of Chang Gung Memorial Hospital (IACUC Approval No.: CGMH2019062002).

Seven groups were assigned: [1] sham control group with fascia operation; [2] AlloDerm implant; [3] PDS implant; [4] PLGA implant; [5] AlloDerm harvest with hAFSC (AlloDerm-SC); [6] PDS harvest with hAFSC (PDS-SC); and [7] PGLA harvest with hAFSC (PGLA-SC).

Rats were anesthetized with 2% Isoflurane mask inhalation. An abdominal midline incision of 4cm and subcutaneous blunt dissection to muscle were made. A 1.01.0cm full-thickness abdominal muscle fascia resected. Mesh measuring 2.02.0cm was fixated with continuous, absorbable suture (Polygactin, Vicryl 3/0) to cover the defect. Skin was then closed with running subcuticular absorbable sutures (Vicryl 3/0) (Fig.2). Sham control group with fascia operation underwent abdominal muscle fascia resection without mesh implantation. In the present study, the number of control rats was reduced in compliance with IACUCs recommendation. Total of 12 rats (week 1), 4 rats (week 2) and 12 rats (week 12) were sacrificed at respective weeks. The tissues were harvested at 12weeks, and meshes were retrieved for tensile properties characteristic and immuno-histological examination.

The mechanical properties of three absorbable mesh with and without AFSC meshes were estimated utilizing tensile test equipment (Lloyd, Ametek, Berwyn, PA, USA) (Fig.3). The maximum strengths of three absorbable mesh with and without AFSC were compared. The stretching speed was set at 100mm/min and the ultimate load and deformation were recorded.

Tensiometry for mechanical properties of the scaffolds.

The rats were euthanized with 3% isoflurane and then decapitated, in accordance and manner consistent with the regulations of the National Institute of Health of Taiwan and Institutional Animal Care and Use Committee of Chang Gung Memorial Hospital.

Sample size calculation was done by using crude method based on law of diminishing return with the equation of E=total number of animals-total number of groups. After the calculation with (7 groups6 rats/group) (7groups)=35, suggesting the sample size for this study was adequate, and 6 rats were used for each group15. The data were analyzed and expressed as meanSD for continuous variables. Continuous data were compared among the groups by using one-way analysis of variance. To evaluate the effect of hAFSC among groups, chi-square test was performed with Fishers exact test. Probability value of<0.05 are statistically significant.

All procedures involving humans were carried out in accordance with relevant guidelines and regulations, and approved by Institutional Board Review Chang Gung Memorial Hospital. Informed consent was obtained from all participants/donor. All experimental procedures were performed under the supervision of a licensed veterinarian, in a manner consistent with the regulations of the National Institute of Health of Taiwan. All animal related procedures were approved by the Institutional Animal Care and Use Committee of Chang Gung Memorial Hospital (IACUC Approval No.: CGMH2019062002). All methods involving animals are reported in accordance with ARRIVE guidelines.

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The properties of absorbable scaffold harvested with human amniotic fluid stem cells on rat model: an innovation for ... - Nature.com

For some, a cure for their blood disorder may be just a swab away.

HAMPTON, Va. Those of African descent with blood disorders like leukemia, lymphoma, or sickle cell are less likely to receive the lifesaving blood stem cells they need compared to those from other cultural backgrounds.

This is hopefully about to change as NMDP (formerly National Marrow Donor Program) hosted its Be the Match blood stem cell donation drive Saturday in Hampton at the Boo Williams Sportsplex.

Attendees were entered in drawings to receive gift cards and were educated about donorship and the disparities in the available pool (of possible donors), said Michelle Bradstock, a friend of John Slade, a Hampton native who is in desperate need of a stem cell match and transplant to help cure his leukemia.

A cure exists, but first, those affected must find a donor. Registering with Bethematch.org, which connects people with blood disorders to those who may be a genetic match, makes finding a genetic match that much easier for those who need one.

Swabbing a cheek at an event like Saturdays, is also a great way to help. Today were swabbing people and were trying to help people get on the registry to help save lives, said Amanda Slade, John Slades daughter and an organizer and volunteer at the event. For more ways to support or donate, visit bethematch.org.

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Blood stem cell donation drive held in Hampton - 13newsnow.com WVEC

Stem cells play a vital role in repairing damaged tissue, whether its a scraped knee or a scarred uterus following pregnancy. New stem cell research has identified the molecules that the cells produce to promote the healing process. The finding could pave the way for the development of new, more effective drugs for injuries or various diseases, including conditions related to reproductive health such as Asherman syndrome, a gynecologic condition in which the uterus scars and becomes fibrotic.

Scientists believed in the past that stem cells served as backup cells that repaired tissues by differentiating into new cells that repopulated the site of injury. Now, they have learned that it is rare for stem cells to completely replace injured tissue. But they still dont fully understand how the cells are able to help damaged areas regenerate.

We found the molecules that stem cells make to help heal and repair tissue, and we hope that understanding this will be potentially useful as a medication in the future.

In the uterus, stem cells play a number of roles, including helping it to expand during pregnancy and to regenerate and repair after childbirth. This new study identified several microRNAs (miRNAs) secreted by the stem cells that helped drive the growth and proliferation of cells in uterine tissue. The researchers published their findings in Stem Cell Research & Therapy on May 1.

We found the molecules that stem cells make to help heal and repair tissue, and we hope that understanding this will be potentially useful as a medication in the future, says Hugh Taylor, MD, chair and Anita OKeeffe Young Professor of Obstetrics, Gynecology & Reproductive Sciences at Yale School of Medicine and the studys principal investigator.

Exosomes are extracellular vesicles, which contain various bioactive molecules and allow cells to communicate with one another. In their new study, Taylors team isolated exosomes secreted by stem cells from human bone marrow. They then used RNA sequencing to characterize all of the miRNA contained in the vesicles and identified those that were most abundant. Then researchers took the most prominent miRNAs and introduced them into human uterine tissue.

The team found that the miRNAs significantly increased the growth and proliferation of the uterine cells. They also studied their effect on the cells decidualization in the endometrium. (Decidualization is the differentiation process uterine cells undergo that prepares the uterus to support an embryo.) The study showed that the miRNAs blocked decidualization.

In a uterus, once a cell becomes differentiated to support pregnancy, it can no longer repair and regenerate. Its permanently locked in that state and often is shed through menstruation later on, says Taylor. By blocking this process, it allows the cells to focus on proliferating and turns on these reparative processes.

The study offers insight into how stem cells promote reparative processes without replacing the tissue itself. Taylor hopes that as researchers continue to gain a greater understanding about how miRNAs work, they could one day be used as drugs for repairing various damaged tissue.

Asherman syndrome, for example, typically occurs after pregnancy, when the supply of stem cells may not be adequate to help the organ heal properly, which can hinder fertility in the future. The idea is that these miRNAs could be used as a medication that is much more readily available and practical, says Taylor. We could potentially deliver them to help prepare the uterus in the critical window when it is damaged and may be vulnerable.

The finding could also have significance beyond the uterus. In future stem cell research, Taylors team plans to study how miRNAs respond to other types of traumatic tissue injury in animal models. We studied the uterus, but the implications are beyond reproduction, potentially including many other conditions where stem cells are involved in repair and regeneration, whether thats injury due to trauma or degenerative diseases, says Taylor.

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Stem Cell Research Uncovers Clues to Tissue Repair That Could Help Heal the Uterus and More - Yale School of Medicine

For some people with blood cancers who need a stem cell transplant, finding a donor who is an excellent match can mean the difference between life and death.

Unfortunately, even though there are more than 40 million potential donors in the national registry, finding a perfect match isnt always possible, especially in underrepresented racial and ethnic groups.

But a new approach using an old chemotherapy drug,cyclophosphamide, isis opening up new possibilities for people with cancers like leukemia, lymphoma, and multiple myeloma. Researchers have found that by administering the drug several days after transplantation, people receiving blood stem cells from unrelated, partially matched donors can have survival rates comparable with those who received exactly matched cells.

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This innovative approach can greatly expand patient access to safe and effective stem cell transplant, regardless of matching degree with the donor, says lead coauthor Monzr M. Al Malki, MD, a hematologist and oncologist and director of the Unrelated Donor BMT program at City of Hope, a cancer research and treatment organization with locations across the United States.

Thats exciting because it means more patients will be able to receive this potentially life-extending therapy, says Dr. Al Malki.

Donor compatibility is determined by a set of protein markers on blood cells called HLAs (human leukocyte antigens), says David Miklos, MD, a professor of medicine and chief of Stanford BMT and Cell Therapy Program at Stanford Medicine in California. Stanford was one of the medical sites of the trial, though Dr. Miklos is not a coauthor of the research.

[2]

[3]

Why was an exact match needed? Anything less increased the likelihood of a graft failure, as well as graft-versus-host disease meaning the transplanted cells attack the patients own, which can cause serious or even fatal complications, explains Miklos.

About a decade ago, researchers started using cyclophosphamide to destroy the parts of a persons immune system that would reject the transplant. That breakthrough allowed researchers to not only have better outcomes in fully matched donors, it also opened the door for successful transplants between people who were only partial matches.

[4]

The new study looked at cyclophosphamide treatment in patients receiving peripheral blood stem cell transplantation meaning healthy stem cells are harvested from a donors bloodstream, and then administered via infusion to the person with cancer.

Blood stem cell transplantation has largely replaced bone marrow transplantation, according to researchers.It's an easier way of collecting stem cells from donors, and its a little safer, because donors dont need to be under anesthesia as they would in bone marrow transplantation, says Al Malki.

For this part of the study, the researchers examined data from 70 adults who were 65 years old on average, all with advanced blood cancers. Participants received a reduced-intensity conditioning regimen to somewhat suppress their immune system to prepare them for transplantation, followed by an infusion stem cells from unrelated, partially matched donors.

The researchers reported an overall high survival rate of 79 percent at one year which is comparable to survival rates seen with fully matched donors.

The main side effect or risk of transplantation is graft-versus-host disease, says Al Malki. After one year, 51 percent of participants were free of the disease and had not relapsed, which is also comparable to what would be seen with fully matched donors, he says.

Historically, barriers in access to transplant have existed due to the low availability of matched, related sibling donors, as well as the substantial variance of matched, unrelated donor availability, especially for patients with diverse ancestry, says study coauthor Steven M. Devine, MD, chief medical officer of NMDP (formerly known as the National Marrow Donor Program and Be The Match).

These findings advance our ability to offer more options to patients without a fully matched donor, many of whom are ethnically diverse and have been underserved in receiving potentially lifesaving cell therapy, says Dr. Devine.

These findings are incredibly important and critical in the effort to improve existing inequities, says Miklos.

In the past, we could not bring some patients forward to receive this lifesaving therapy because they didnt have a compatible donor, but with the new approach of using post-transplant cyclophosphamide, all patients have donors now, he says.

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Cancer Patients Who Need Stem Cell Transplants May Have New Donor Options - Everyday Health

Ru(II) complexes containing 2-thiouracil derivatives display potent cytotoxicity in cancer cell lines

The cytotoxic effects of complexes 1 and 2 were evaluated on a panel of 23 cancer cell lines (HepG2, HCT116, NB4, THP-1, Jurkat, K-562, HL-60, KG-1a, MDA-MB-231, MCF-7, 4T1, HSC-3, CAL 27, SCC-25, SCC4, SCC-9, A549, PANC-1, OVCAR-3, DU 145, U-87 MG, A-375, and B16-F10) and three noncancerous cells (PBMC, MRC-5, and BJ) using the Alamar Blue assay after 72h of treatment (Fig. 1B and Table S1). Both complexes displayed potent cytotoxicity against all cancer cell lines, with half-maximal inhibitory concentration (IC50) values ranging from 2.4M in OVCAR-3 ovarian cancer cells to 17.5M in A549 lung cancer cells for complex 1 and from 1.6M in OVCAR-3 ovarian cancer cells to 10.5M in A549 lung cancer cells for complex 2. Doxorubicin was used as a positive control and showed cytotoxicity in all cell lines.

In noncancerous cells, complex 1 had an IC50 of 17.7M in MRC-5 pulmonary fibroblasts, 7.0M in BJ foreskin fibroblasts and 14.2M in PBMCs. In comparison, complex 2 presented IC50 values of 15.2M in MRC-5 pulmonary fibroblasts, 7.8M in BJ foreskin fibroblasts and 11.7M in PBMCs. The selectivity indices (SIs) were calculated by the following formula: SI=IC50 ([noncancerous cells]/IC50 [cancerous cells]). Figure 1C and Table S2 present the calculated SI. Curiously, both complexes showed an SI>2 for many of the cancer cells investigated.

To study the anti-HCC potential of these complexes, the HCC cell line HepG2 was used in further experiments. Therefore, the viability of HepG2 cells treated with complex 1 at concentrations of 2, 4, and 8M and complex 2 at concentrations of 1.5, 3, and 6M was determined by trypan blue assay after 12, 24, 48, and 72h of incubation. Both complexes reduced HepG2 cell viability in a concentration- and time-dependent manner (Fig. S1A1D). After 72h of incubation, complex 1 reduced cell viability by 46.5, 72.2, and 95.8%, while complex 2 inhibited cell viability by 39.4, 61.4, and 93.5%, respectively.

To determine whether complexes 1 and 2 can act against liver CSCs, we first performed a long-term colony formation assay to determine whether these complexes affect the clonogenic ability of HCC HepG2 cells. Clonogenic assays are well-known methods for evaluating the stemness of CSCs since a single CSC can form clonogenic colonies [16, 17]. Interestingly, treatment with both complexes significantly decreased the clonogenic viability of HepG2 cells in a concentration- and time-dependent manner (Fig. 2A, B).

A Representative images and (B) quantification of the number of colonies formed from HepG2 cells after treatment with complexes 1 and 2. (C and D) Quantification of CD133 expression on HepG2 cells after 24h of incubation with 8M complex 1 or 6M complex 2, as determined by flow cytometric analysis. (E and F) Quantification of CD44high in HepG2 cells after 24h of incubation with 8M complex 1 or 6M complex 2, as determined by flow cytometric analysis. The vehicle (0.2% DMSO) was used as a negative control (CTL). The data are expressed as the meanS.E.M. of three biological replicates carried out in duplicate. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test.

Next, we quantified the expression of two biomarkers of liver CSCs, CD133 [18] and CD44 [19], in HepG2 cells treated with complexes 1 and 2. Likewise, both complexes reduced the percentage of HepG2 CD133-positive cells (Fig. 2C, D), while complex 2 reduced the percentage of HepG2 CD44high cells (Fig. 2E, F).

In a new set of experiments, we measured the effects of complexes 1 and 2 on three-dimensional (3D) tumorspheres formed from HepG2 cells since multicellular 3D tumor spheroids are well-known cell culture systems that can enrich cells with CSC characteristics [20, 21]. Both complexes reduced HepG2 tumorsphere growth (Fig. S2 and S3) and caused cell death (Fig. S4), corroborating that these molecules may inhibit CSCs in HCC HepG2 cells.

A series of cellular and molecular analyses were performed to examine the mechanism of cell death in HepG2 cells treated with complexes 1 and 2. HepG2 cells that were treated with complexes 1 and 2 for 24, 48, and 72h showed cell morphology changes that were associated with apoptosis, including a reduction in cell volume, chromatin condensation, and fragmentation of the nuclei, as observed in May-Grunwald-Giemsa-stained cells (Fig. S5).

Light scattering characteristics measured by flow cytometry were used to analyze cellular parameters such as size and complexity/granularity in HepG2 cells treated with complexes 1 and 2 (Fig. S6AS6F). Forward light scattering (FSC) was employed as a cell size metric in this experiment, while side scattering (SSC) was used to determine cell complexity/granularity. Treatment with these complexes caused cell shrinkage, as indicated by a decrease in the FSC, accompanied by an increase in the SSC, probably due to nuclear condensation. Both morphological changes are associated with cellular apoptosis, corroborating the findings observed in cells stained with May-Grunwald-Giemsa.

Internucleosomal DNA fragmentation and cell cycle distribution were evaluated in HepG2 cells after 24, 48, and 72h of incubation with complexes 1 and 2 via a DNA content-based flow cytometry assay (Fig. 3AG). All DNA of subdiploid size (sub-G0/G1) was considered fragmented. Both complexes induced DNA fragmentation in a time- and concentration-dependent manner. After 72h of incubation, complex 1, at concentrations of 2, 4, and 8M, caused DNA fragmentation by 12.3, 26.7, and 43.1%, respectively, while complex 2, at concentrations of 1.5, 3, and 6M, induced DNA fragmentation by 18.6, 39.1, and 72.9%, respectively (against the 5.6% detected in the control). The cell cycle phases G0/G1, S and G2/M decreased proportionally in HepG2 cells treated with complexes 1 and 2. Doxorubicin, used as a positive control, also caused DNA fragmentation.

Representative flow cytometric histograms of the cell cycle distribution of HepG2 cells after treatment with complexes 1 and 2 after 24 (A), 48 (B) and 72 (C) h of incubation. The percentages of cells in the sub-G0/G1 (D), G0/G1 (E), S (F) and G2/M (G) phases were quantified via flow cytometric analysis. The vehicle (0.2% DMSO) was used as a negative control (CTL), and doxorubicin (DOX, 1M) was used as a positive control. The data are expressed as the meanS.E.M. of three biological replicates carried out in duplicate. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test.

Annexin V-FITC/propidium iodide (PI) double staining was also applied to HepG2 cells treated with complexes 1 and 2 for 24, 48 and 72h to quantify phosphatidylserine exposure and cell membrane integrity, which are markers of apoptosis and necrosis, respectively. Both complexes induced a significant increase in the percentage of apoptotic cells in a time- and concentration-dependent manner, and no significant increase in the percentage of necrotic cells was detected (Fig. 4AF). After 72h of incubation, complex 1, at concentrations of 2, 4, and 8M, increased apoptosis by 9.0, 48.9, and 76.1%, respectively, while complex 2, at concentrations of 1.5, 3, and 6M, increased apoptosis by 9.1, 40.7, and 76.9%, respectively (against 4.8% found in the control). Treatment with doxorubicin, which was used as a positive control, also led to apoptosis.

Representative flow cytometric dot plots of HepG2 cells stained with annexin V-FITC/PI after treatment with complexes 1 and 2 after 24 (A), 48 (B) and 72 (C) h of incubation. The percentages of viable (annexin V-FITC-/PI- cells) (D), apoptotic (early apoptotic [annexin V-FITC+/PI- cells] plus late apoptotic [annexin V-FITC+/PI+ cells]) (E) and necrotic (annexin V-FITC-/PI+ cells) (F) cells were quantified. The vehicle (0.2% DMSO) was used as a negative control (CTL), and doxorubicin (DOX, 1M) was used as a positive control. The data are expressed as the meanS.E.M. of three biological replicates carried out in duplicate. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test.

As mitochondrial dysfunction and PARP cleavage are well-known events in apoptotic cell death, mitochondrial transmembrane potential and PARP (Asp214) cleavage were also determined by flow cytometry. Significant mitochondrial depolarization (Fig. 5A) and increased levels of PARP (Asp214) cleavage (Fig. 5B, C) were found in HepG2 cells treated with complexes 1 and 2, corroborating that these complexes can cause cell death via apoptosis. Moreover, the BAD KO SV40 MEF cell line, as well as its parental cell line, WT SV40 MEF, were used to assess the involvement of the proapoptotic protein BAD in the cell death caused by complexes 1 and 2 (Fig. 5D, E). On the other hand, these complexes cause cell death independent of the protein BAD.

A Quantification of mitochondrial membrane depolarization in HepG2 cells after 24h of incubation with complex 1 or 2, as determined by flow cytometry. B and C Quantification of PARP (Asp214) cleavage in HepG2 cells after 24h of incubation with complex 1 (8M) or 2 (6M), as determined by flow cytometric analysis. MFI: Mean fluorescence intensity. D Survival curves of WT SV40 MEFs and BAD KO SV40 MEFs upon treatment with complexes 1 and 2 and 5-fluorouracil (5-FU, used as a positive control). The curves were obtained from at least three biological replicates carried out in duplicate using the Alamar Blue assay after 72h of incubation. E DNA fragmentation (sub-G0/G1 cells) and cell cycle distribution (G0/G1, S and G2/M phases) of WT SV40 MEFs and BAD KO SV40 MEFs after 48h of incubation with complexes 1 (10M) and 2 (10M) or 5-FU (40M). The vehicle (0.2% DMSO) was used as a negative control (CTL). The data are expressed as the meanS.E.M. of three biological replicates carried out in duplicate. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test.

To investigate the molecular mechanism of action of complexes 1 and 2, we analyzed the transcripts of 82 target genes using a qPCR array (Fig. 6A, B and Table S3). Among the altered gene transcripts, genes related to NF-B (the NFKB1 gene with RQ=0.45 for complex 1), PI3K/Akt/mTOR (the PIK3CA gene with RQ=0.44 for complex 2; the MTOR gene with RQ=0.39 for complex 1) and oxidative stress (the GSTP1 gene with RQ=0.49 for complex 1 and RQ=0.27 for complex 2; the TXN gene with RQ=0.37 for complex 2; and the TXNRD1 gene with RQ=0.35 for complex 2) were downregulated in HepG2 cells treated with complexes 1 and 2.

A, B Genes up- and downregulated in HepG2 cells after 12h of treatment with complexes 1 (8M) and 2 (6M). The vehicle (0.2% DMSO) was used as a negative control (CTL). The data are expressed as the relative quantification (RQ) compared to the CTL data. The genes were upregulated if RQ2 (red bars) and downregulated if RQ0.5 (green bars). Quantification of the levels of phospho-NF-B p65 (S529) (C, D), Akt1 (E, F), phospho-Akt (S473) (G, H), phospho-mTOR (S2448) (I, J), and phospho-S6 (S235/S236) (K, L) in HepG2 cells after 24h of incubation with complexes 1 (8M) and 2 (6M), as determined by flow cytometry. The vehicle (0.2% DMSO) was used as a negative control (CTL). The data are expressed as the meanS.E.M. of three biological replicates carried out in duplicate. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test. MFI: Mean fluorescence intensity.

Next, the protein levels of several elements of the NF-B and Akt/mTOR signaling pathways were quantified. The levels of phospho-NF-B p65 (S529) (Fig. 6C, D), Akt1 (Fig. 6E, F), phospho-Akt (S473) (Fig. 6G, H), phospho-mTOR (S2448) (Fig. 6I, J), and phospho-S6 (S235/S236) (Fig. 6K, L) were reduced in complex 1-treated HepG2 cells. In contrast, the levels of Akt1 (Fig. 6E, F), phospho-Akt (S473) (Fig. 6G, H), and phospho-mTOR (S2448) (Fig. 6I, J) were reduced after treatment with complex 2, indicating that these complexes interfere with NF-B and Akt/mTOR signaling. The levels of phospho-PI3K p85/p55 (T458/T199) (Fig. S7A and S7B), phospho-Akt (T308) (Fig. S7C and S7D), phospho-4EBP1 (T36/T45) (Fig. S7E and S7F), and phospho-elF4E (S209) (Fig. S7G and S7H) were not affected by treatment with these complexes.

As complexes 1 and 2 downregulate the level of phospho-mTOR (S2448), a negative regulator of autophagy [22], the effect of these complexes on the induction of autophagy was investigated. On the other hand, none of them caused autophagy, as assessed by quantification of p62/SQSTM1 expression levels in HepG2 cells treated with complexes 1 and 2 (Fig. S8AS8C).

Since both complexes reduced the proportion of liver CSC markers in HepG2 cells and liver CSCs are directly associated with cell migration and invasion [23], we hypothesized that these complexes could reduce HepG2 cell motility. Initially, noncytotoxic concentrations of complexes 1 and 2 were selected (Fig. S9) and tested in the wound healing assay. Both complexes reduced HepG2 cell migration after 72h of incubation at noncytotoxic concentrations (0.5M for complex 1 and 0.3M for complex 2) (Fig. 7A, B). Similarly, both complexes, at the same concentrations, also reduced motility in a transwell cell migration assay (Fig. 7C, D) using HepG2 cells.

A Representative images and (B) quantification of HepG2 cell migration in the wound healing assay after 72h of incubation with complexes 1 and 2. C Representative images and (D) quantification of HCT116 cell migration in the transwell migration assay after 24h of incubation with complexes 1 (8M) and 2 (6M). Quantification of vimentin (E, F) and E-cadherin (G, H) expression in HepG2 cells after 24h of incubation with complexes 1 (8M) and 2 (6M), as determined by flow cytometry. The vehicle (0.2% DMSO) was used as a negative control (CTL). The data are expressed as the meanS.E.M. of three biological replicates carried out in duplicate. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test. MFI: Mean fluorescence intensity.

Next, the epithelialmesenchymal transition (EMT) markers vimentin and E-cadherin were evaluated in HepG2 cells treated with complexes 1 and 2 after 24h of incubation. Vimentin (Fig. 7E, F) was reduced, and E-cadherin (Fig. 7G, H) was increased by treatment with complex 2, indicating that this molecule can modulate EMT.

The in vivo antitumor activity of complexes 1 and 2 was investigated in C.B-17 SCID mice grafted with HepG2 cells. The animals were treated with 2 or 4mg/kg of both complexes intraperitoneally once a day for 21 consecutive days. Both complexes inhibited the growth of HepG2 cells in mice (Fig. 8A, B). At the end of treatment, the mean tumor weight in the negative control group was 981mg, while it was 665mg in the doxorubicin-treated group. In complex 1-treated animals, the mean tumor weights were 635 and 455mg, corresponding to 35.3 and 53.6% tumor inhibition, respectively. In complex 2-treated animals, the mean tumor weights were 358 and 340mg, corresponding to 63.6 and 65.4% tumor inhibition, respectively. Doxorubicin reduced the tumor weight by 32.2%.

A, B In vivo antitumor activity of complexes 1 and 2 on C.B-17 SCID mice inoculated with HepG2 cells. The animals were treated with complexes 1 and 2 at doses of 2 or 4mg/kg intraperitoneally once a day for 3weeks. C Representative photomicrographs of HepG2 tumors from animals treated with complexes 1 and 2. Histological sections were stained with hematoxylin-eosin and analyzed by light microscopy. The asterisks indicate areas of tissue necrosis. Scale bar=50m. The vehicle (5% DMSO) was used as a negative control (CTL), and doxorubicin (DOX, 1mg/kg) was used as a positive control. The data are expressed as the meanS.E.M. from 8 animals. *P<0.05 compared to CTL by one-way analysis of variance (ANOVA) followed by Dunnetts multiple comparisons test.

The tumors presented histological characteristics compatible with hepatocellular carcinoma, such as intense cellular and nuclear pleomorphism, hyperchromatism, atypical mitotic figures, and hepatocyte-like cells (Fig. 8C). The histological grading of the tumors varied from poorly to moderately differentiated in all the experimental groups. The tumor cells were organized in nodules or cords surrounded by a poorly vascularized collagen matrix. Areas of coagulative necrosis were frequent, especially in more central tumor regions. In addition, an infiltrate of inflammatory cells, predominantly mononuclear, was observed mainly adjacent to the necrotic areas. Areas of dystrophic calcification were observed in some of the tumors in the negative control, doxorubicin and complex 2 (4mg/kg) groups. Furthermore, invasion fronts in the muscular tissue were observed in the control groups.

The toxicity parameters of animals treated with complexes 1 and 2 were also examined. No significant changes in body weight or organs (liver, kidney, lung, or heart) were detected in the animals treated with these complexes (P>0.05) (Fig. S10AS10F). Histopathological analysis of the kidneys (Fig. S11), livers (Fig. S12), and lungs (Fig. S13) of mice treated with complexes 1 and 2 revealed some alterations that were minor and/or reversible, indicating little damage to normal tissues. No significant changes were observed in the hearts of the animals treated with complexes 1 and 2 (data not shown).

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Ru(II)-based complexes containing 2-thiouracil derivatives suppress liver cancer stem cells by targeting NF-B and Akt ... - Nature.com

Animal model and tissue processing

Twenty-six 3-month-old male small-tailed sheep with body weights ranging from 25 to 27kg were used in this study under a research protocol approved by the Ethics Committee of Tianjin Stomatological Hospital (approval code: Tjskq2013001). All experiments were performed in compliance with the Animal Management Regulations and Administrative Measures on Experimental Animals, and are reported in accordance with the ARRIVE guidelines. The animals were housed in a laboratory animal facility with adequate facility management services and specialised nursing care and husbandry practices, similar to our previous study8. The animals underwent unilateral TMJ surgery, involving the removal of two-thirds of the articular disc and severe damage to the articular fossa to induce bony ankylosis, following the protocol in our previous publication8. The anaesthesia, analgesia and euthanasia methods used were the same as those described in a previous study26.

Three sheep per time point were sacrificed for tissue specimen collection via euthanasia (120mg/kg of pentobarbitone sodium administered via intravenous injection) on Days 1, 4, 7, 9, 11, 14 and 28 postsurgery, and five were sacrificed on Day 14 after TMJ surgery for subsequent isolation and culture of MSCs. The TMJ complexes were removed en bloc with a band saw, and newly formed tissue within the joint space was bluntly dissected from the surrounding soft tissue using periosteum separators.

After fixation with 10% formalin, the collected tissue was routinely dehydrated and embedded and then cut into 5m-thick sections using a microtome for subsequent staining experiments. Successive slices were taken to ensure that the haematoxylin and eosin (HE), immunohistochemistry (IHC) and multiplex immunohistochemistry (mIHC) staining were performed on the same area.

To observe the histological manifestations of the early stages of traumatic bony ankylosis of the TMJ, paraffin sections of tissues collected on Days 1, 4, 7, 9, 11, 14 and 28 post -operation were stained with HE (Sigma, USA).

IHC was used to detect YAP and Runt-related transcription factor 2 (RUNX2) expression positions in TMJ ankylosis formation. Antibodies against RUNX2 was used to label the osteogenically active regions. After routine dewaxing, hydration, heat repair and antibody blocking of the tissue sections, a nonblocking kit (ZSGBBIO, China) was used to prepare the sections for staining with an anti-YAP antibody (rabbit monoclonal, 1:1000, ProteinTech Group, USA) and an anti-RUNX2 antibody (mouse monoclonal, 1:1000, Abcam, USA), to which a matching secondary antibody was then added, followed by incubation at 37C for 30min. Diaminobenzidine (DAB, 1:20, ZSGBBIO, China) was used for the final immunological colour development, and the nuclei were stained with haematoxylin (Sigma, USA). Image acquisition was performed using a microscope (Nikon, Japan).

We used mIHC to further analyse the expression level of YAP in the early stages of traumatic bony ankylosis of the TMJ. A specialised tyramine signal amplification-immunohistochemistry (TSA-IHC) multitarget immunofluorescence staining kit (Bruno, China) was used for this experiment. Different primary antibodies were applied sequentially, followed by horseradish peroxidase-coupled secondary antibody incubation and tyramine signal amplification (TSA). After each TSA, the slides were processed for antigen elution. After labelling YAP and RUNX2, cell nuclei were stained with 4', 6-diamidino-2-phenylindole (DAPI) (Bruno, China). The images were acquired using a panoramic microscope camera system (Jinan Tangier Electronics Co., China), and the digital scanning and viewing software used was CaseViewer 2.4 (3DHISTECH, Hungary).

All collected tissues were initially viewed at100 for each section, in which functioning YAP was stained red, RUNX2 was stained green, and nuclei were stained blue. Subsequently, 20micrographs were acquired and three fields of view were obtained for each specimen. The percentage of cells exhibiting positive coexpression of RUNX2+YAP (i.e., number of positive coexpression cellstotal number of cells) was calculated using the Halo (Indica Labs, USA) whole-slide image analysis platform. The fluorescence intensity measured as the integrated density (Int Den) and area were measured using ImageJ (National Institutes of Health, USA), and the mean grey value (mean) of each image was then calculated.

Primary cells were isolated using the tissue attachment method27. The specific isolation, incubation, culture and purification processes applied to the MSCs were the same as described in a previous study13. Third-passage cells were harvested for subsequent experimental procedures.

The third-passage cells were digested with 0.25% trypsin (Solarbio, China) to obtain a single-cell suspension. The cells were then resuspended with phosphate buffered saline (PBS) solution supplemented with 1% foetal bovine serum (FBS) after centrifugation. For 30min at 4C in the dark, 1106 cells were incubated with the corresponding commercial monoclonal antibodies (CD44, Immunostep, clone 25.32, 1:10; CD29, Biolegend, clone TS2/16, 1:20; CD31, AbD Serotec, clone CO.3E1D4, 1:10; CD45, AbD Serotec, clone 1.11.32, 1:10), and then measurements were made using a FACSCanto (BD Biosciences, USA) flow cytometer. Flow cytometric analyses were performed using FACSDiva (BD Biosciences, USA) and FlowJo software (TreeStar, Ashland, Oregon).

Third-passage cells were seeded at a density of 1105/ml in a six-well plate (Corning, USA). When the cells reached 6070% confluence, the medium was removed, and the cells were washed with PBS three times. Osteogenic induction (OI) medium (the formula used for this osteogenic induction medium is described in an earlier study13) was added to the OI group, while normal control (NC) group was still added complete medium (-minimal essential medium (-MEM) plus 10% foetal bovine serum, 100 U/ml penicillin, 100g/ml streptomycin and 2.5g/ml amphotericin B).

The cells were stained with 1% Alizarin red (pH=4.3, Solarbio, China) after undergoing OI for 7days and, subsequently, 14days. Mineralised nodules were then dissolved in 10% cetylpyridinium chloride (Solarbio, China) for semiquantitative analysis by examining the absorbance at 562nm. Alkaline phosphatase (ALP) activity was tested using an ALP colour development kit (Beyotime, China) after the cells underwent OI for 7days and, subsequently, 14days. The mRNA expression levels of YAP, RUNX2, the Sp7 transcription factor (Osterix) and osteocalcin (OCN) were measured via real-time polymerase chain reaction (PCR) after induction for 7 and 14days.

To detect the intracellular localisation of YAP during the OI of MSCs, third-passage cells were inoculated into confocal culture dishes (NEST, China) at a density of 1103 cells/ml, the OI medium was cultured for three days, and immunofluorescence staining was performed as described in a previous study13. The monoclonal antibody used was anti-mouse YAP1 (clone: 3A7A9, ProteinTech Group, 1:800) with 594-conjugated goat anti-mouse lgG. Fluorescently labelled cells were photographed using an inverted fluorescence microscope (Nikon, Japan).

Total cellular RNA was extracted using a universal RNA extraction kit (TaKaRa, Japan), and the RNA concentration was determined. cDNA was extracted via reverse transcription using a synthesis kit (Promega, USA), and quantitative real-time PCR was performed with FastStart Universal SYBR Green Master Mix (Roche, Switzerland, Cat. #04,913,850,001) using a LightCycler 480 II (Roche, Switzerland). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as an internal reference. The reaction system and PCR cycling parameters were the same as those described in a previous study28. The sequences of the primers used for PCR amplification are listed in Table 1.

A YAP-silenced MSC model was constructed using shRNA plasmid transfection technology (RuiboBio, Guangzhou, China). The target sequences are presented in Table 2. Lipofectamine 3000 (Thermo Fisher Scientific, Waltham, USA) was used to transfect the MSCs, according to the manufacturers instructions. Third-passage cells were briefly inoculated into 35mm dishes (Corning, USA) at a density of 1105 cells/ml after transient transfection with shYAP1, shYAP2, shYAP3 and shRNA plasmid empty vectors. The transfection efficiency was tested after 24h of culture to determine the highest transfection efficiency of the shRNA plasmid. The MSCs were then reinoculated in 35mm dishes following the previously described density and method, and divided into three groups: The blank group (the blank control group, in which the shRNA plasmid was not transfected), the shRNA-YAP-NC group (the negative control group, in which the shRNA plasmid was transfected with an empty vector), and the shRNA-YAP group (the silencing group, in which the shYAP plasmid was transfected with the highest efficiency). After one day of transfection, the stock solutions of the three groups were discarded, after the following experimental procedures: (1) The stock medium was replaced with OI medium to detect the effect of YAP silencing on the osteogenic ability of the MSCs; the procedures for Alizarin red staining, ALP activity detection and real-time PCR were the same as those described earlier. (2) Continue to use complete medium to detect the effects of YAP silencing on the proliferation and migration ability of MSCs.

EdU assay, cell formation assays and cell counting kit-8 (CCK-8) assays were used to investigate whether YAP silencing affects the proliferative capacity of MSCs.

A BeyoClick EdU-488 Cell Proliferation Assay Kit (Beyotime, China) was used to determine the cell proliferative capacity. Third-passage MSCs were seeded in 24-well plates (Corning, USA) at a density of 2,000 cells /well. After transfection using the preceding method, the three groups of cells (Blank, shRNA-YAP-NC and shRNA-YAP) were stained following the protocol of the kit. Fluorescence detection was performed on each well at a 495nm wavelength (10) using an inverted fluorescence microscope (Nikon, Japan). Quantitative analysis of EdU proliferation was performed using ImageJ; the percentage of EdU-positive=cells was calculated as (number of EdU-positive cells divided by the total number of cells)100%.

Third-passage cells were inoculated in 6-well plates (Corning, USA) at a density of 500 cells/well. After transfection using the method described earlier, the three groups of cells (Blank, shRNA-YAP-NC and shRNA-YAP) were fixed with 4% paraformaldehyde and stained with 5% Giemsa (Hydrogen, China) for 40min on Day 7 of culture, and colonies containing>50 cells were counted under the microscope (Nikon, Japan). To clearly observe cell proliferation under a microscope, the number of colonies was analysed using ImageJ.

Third-passage cells were seeded in four 96-well plates (Corning, USA) at a density of 2,000 cells/well. After transfection following the method described earlier, cell proliferation tests were performed on the three groups of cells (Blank, shRNA-YAP-NC and shRNA-YAP) using a CCK-8 kit (Beyotime, China) as described in a previous study13, and the detection time points were 1day, 3days, 5days and 7days.

Wound healing and Transwell migration assays were used to investigate whether YAP silencing affects the migratory capacity of MSCs.

Third-passage MSCs were inoculated in 6-well plates (Corning, USA) at a density of 1105 cells /well. When the cells reached to 90% confluence after transfection, a straight line was drawn in the cell layer with a sterile 200l pipet tip. Images were acquired under a microscope (Nikon, Japan) at 0h, 6h, 24h and 48h. The distance migrated by the cells was analysed using ImageJ. The relative wound closure rate was calculated as the ratio of the wound distance at 6h, 24h and 48h to the wound distance at 0h.

A 24well plate with Transwell chambers (Beyotime, China) with an 8.0m pore size was used for this experiment. Serumfree -MEM was added to the upper chambers, and a medium containing 20% FBS was added to the lower chamber. After transfection, the cells of the three groups (Blank, shRNA-YAP-NC, shRNA-YAP) were inoculated in the upper chamber at a cell density of 3104/well. After 24h of incubation at 37C, the migratory cells on the membrane were fixed in 4% paraformaldehyde and stained with 0.1% crystal violet (Beyotime, China) for 30min. The membrane was air-dried and then photographed under a light microscope (Nikon, Japan) at 10magnification. The number of migrated cells was quantified using ImageJ.

The results of the experiments were analysed using unpaired t tests between two groups, while comparisons among three or more groups were performed using one-way ANOVA combined with Bonferronis multiple comparisons test (GraphPad Prism 9.0, USA); P<0.05 was considered to indicate statistical significance. The DAgostino-Pearson test was used to assess normality. Lines and error bars in all figures denote of the mean and standard error mean (SEM), respectively.

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YAP promotes the early development of temporomandibular joint bony ankylosis by regulating mesenchymal stem cell ... - Nature.com

ARID1B plays an essential role in regulating adult mouse incisor growth and tissue homeostasis

To investigate the role of ARID1B in regulating MSC fate commitment and mesenchymal tissue homeostasis, we evaluated the expression pattern of ARID1B in the proximal region of the mouse incisor. We found that ARID1B is widely expressed in the dental mesenchyme near the NVB where MSCs reside, as well as in odontoblasts, dental pulp cells, and epithelial cells, but less in the TAC region and pre-odontoblasts (Fig.1a, b). Previous study has shown that GLI1+ cells are MSCs surrounding the NVB19. To find out whether ARID1B is expressed in these GLI1+ MSCs, we co-stained GLI1 and ARID1B and found that ARID1B+ cells in the NVB region overlap with a sub-population of GLI1+ cells (Fig.1c, d). Thus, we hypothesized that ARID1B plays a role in regulating MSC commitment and tissue homeostasis in the adult mouse incisor.

ad Immunostaining of ARID1B (red) in control (a, b) mouse incisor and co-immunostaining of ARID1B (red)/-galactosidase (-gal) (green) in Gli1-LacZ mouse incisor (c, d). b, d represent the high-magnification image of the box in (a, c). White dotted lines outline the cervical loop. White arrows indicate ARID1B+ cells. Yellow arrows indicate the ARID1B+/GLI1+ cells. ej Notch movement assay in control (eg) and Gli1-CreER;Arid1bfl/fl (hj) mice. Yellow dotted lines show the notch position. Blue lines show the gingival margin. TMX, tamoxifen; wpt, week post-tamoxifen injection. Illustration below (ek) created with BioRender.com released under a Creative Commons Attribution-NonCommercial-NoDerivs 4.0 Internatioanl license. k Quantification of the notch movement every other day from D2 to D10. Data are meanSEM, control n=5, Gli1-CreER;Arid1bfl/fl n=7, unpaired two-tailed Students t-test. D2, p=0.0083; D4, p=0.0103; D6, p=0.0001; D8, p=0.0003; D10, p=0.0001. lo MicroCT and HE staining of control (l, m) and Gli1-CreER;Arid1bfl/fl (n, o) mouse incisors at 3 months after tamoxifen induction. l, n MicroCT of control (l) and Gli1-CreER;Arid1bfl/fl (n) mouse incisors. White arrow indicates the narrowed dental pulp. m, o HE staining of control (m) and Gli1-CreER;Arid1bfl/fl (o) mouse incisors. Yellow arrows indicate the initiation of odontoblast polarization. Green arrow indicates the disordered alignment of odontoblasts. White two-way arrows indicate the dentin thickness. Black asterisk indicates stacked and distorted dentin. Boxes in m and o are shown at higher magnification on the right. p, q Dspp (red) in situ hybridization in control (p) and Gli1-CreER;Arid1bfl/fl (q) mouse incisors. White dotted lines outline the cervical loop. Yellow dotted lines show the cervical loop bending point to the odontoblast initiation distance. Unfilled arrows indicate the distance between the yellow dotted lines. n=3. r Quantification of the dental pulp cavity. Data are meanSEM, n=3, unpaired two-tailed Students t-test. p=0.0113. s Quantification of the distance of Dspp+ cells to the cervical loop. Data are meanSEM, n=3, unpaired two-tailed Students t-test. p<0.0001. Source data are provided as a Source Data file. Scale bars: 2mm (ej, l, n); 100m (other images).

GLI1+ MSCs located surrounding the NVB support the mouse incisors growth and replenishment throughout the lifespan. Using the Gli1-CreER line, we generated Gli1-CreER;Arid1bfl/fl mice, in which Arid1b was inactivated in the GLI1+ lineage after tamoxifen induction at one month of age, and confirmed that Arid1b was effectively deleted from the incisor mesenchymal and epithelial cells (Supplementary Fig.1ad). To evaluate the impact of the loss of Arid1b in MSCs on the mouse incisor growth, we first performed an incisor growth assay by comparing the movement of a notch made in the incisor enamel above the gingival margin in both control and Arid1b mutant mice. We found that the growth rate was significantly slower in Arid1b mutant mice than in the control across all measurement time points (Fig.1ek). This result indicated that the loss of Arid1b in the GLI1+ lineage impairs the adult mouse incisor growth.

Next, we assessed the long-term impact following the loss of Arid1b. At 2 months post-tamoxifen induction, the dental pulp cavity was narrower in Gli1-CreER;Arid1bfl/fl mice than in the control, as shown by microCT (Supplementary Fig.1e, h). Histologically, the polarization of odontoblasts (Supplementary Fig.1f, i) and the expression of odontoblast differentiation marker Dspp (Supplementary Fig.1g, j) were initiated more proximally to the cervical loop in Arid1b mutant mice than in the control mice. The cervical loop also appeared to be smaller in the Arid1b mutant mice. The odontoblast alignment was affected, and the dentin appeared thicker in the Arid1b mutant mice. Moreover, the phenotype of the Arid1b mutant mice became more severe at 3 months post-tamoxifen induction, with a limited dental pulp cavity and stacked dentin at the proximal end of the incisor (Fig.1lo, r). The odontoblasts were well aligned, and their differentiation was marked by the organized alignment of nuclei along the basement membrane in the control; in contrast, the organization of the odontoblasts was abnormal, and the nuclei were in the opposite position in Arid1b mutant mice. The odontoblasts were premature in the proximal region in the Arid1b mutant mice, as confirmed by Dspp marker staining (Fig.1p, q, s). These results indicated that ARID1B plays a role in maintaining adult mouse incisor tissue homeostasis.

Furthermore, to investigate the cellular changes underlying the reduced growth rate and abnormal dentin formation in Arid1b mutant incisors, we evaluated the potential of TACs to differentiate into odontoblasts after the loss of Arid1b. We labeled TACs in the DNA synthesis phase using EdU injection and harvested the tissue 48h later to assess the TAC differentiation. The overlap between Dspp+ odontoblasts and EdU-labeled cells represented the TAC differentiation ability, and the overlap length indicated the migration rate of these differentiated cells during the preceding 48h. We observed a reduced overlap length of EdU+/Dspp+ cells in Arid1b mutant mice compared to controls at 1 week post-induction, indicating that loss of Arid1b caused the compromised migration rate of the differentiated TACs (Supplementary Fig.1kn, q). To understand the cause of abnormal dentin formation, we administered calcein and alizarin red S dual fluorescence injections at different time points to dynamically compare the odontoblast migration rate and dentin deposition rate. Using this approach, the fluorescence precipitation represents dentin formation at the time of injection. We compared the odontoblast migration length and dentin deposition depth over a period of 5 days. Statistical analysis revealed a significantly shorter odontoblast migration length in Arid1b mutant mice compared to controls, while there was no significant difference in the depth of dentin deposition (Supplementary Fig.1op, r). These findings indicated that the loss of Arid1b impairs the odontoblast migration rate, leading to abnormal stacked dentin.

GLI1+ cells contribute to the mesenchymal and epithelial lineages in the mouse incisor19 and ARID1B was effectively deleted in both tissues in the Gli1-CreER;Arid1bfl/fl mice. To clarify whether the loss of Arid1b in the dental epithelium has any effect on incisor tissue homeostasis, we used the Sox2-CreER line to specifically delete Arid1b in the epithelial lineage by generating Sox2-CreER;Arid1bfl/fl mice. Again, we induced Cre activity with tamoxifen at 1 month of age and harvested the incisor at 2 months post-induction. There were no signs of either premature odontoblasts or a differentiation defect in Sox2-CreER;Arid1bfl/fl mice as compared to the control (Supplementary Fig.2af). This result indicated that the loss of Arid1b specifically in the dental mesenchyme impairs the incisor growth and tissue homeostasis.

The continuous growth of the mouse incisor is induced by the progression of MSCs giving rise to TACs, along with proliferation and differentiation in the proximal region. Due to the overlap between ARID1B+ cells and a subpopulation of GLI1+ cells in the proximal region, we investigated the potential impact of Arid1b loss on GLI1+ MSCs. We generated Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mice to compare the numbers of GLI1+ cells in the incisors of these mutants and Gli1-LacZ control mice. At 5 days post-induction, the number of GLI1+ cells was significantly reduced in the Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mice compared to Gli1-LacZ mice (Fig.2ae). These MSCs are quiescent cells that undergo slow-cycling self-renewal and reside in the proximal region of the mouse incisor. Thus, we investigated the impact of ARID1B on MSCs quiescence based on their label-retaining ability. Since incisor mesenchyme turnover takes about 1 month19, we injected control and Gli1-CreER;Arid1bfl/fl mice with EdU for a 1-month period beginning from postnatal day 5 and analyzed the cells after another month. In this case, the label retaining cells (LRCs) detected by EdU staining would be the quiescent cells. We found the number of LRCs was significantly reduced in Gli1-CreER;Arid1bfl/fl mice compared to controls (Fig.2fj), confirming that loss of Arid1b impairs mouse incisor MSC quiescence.

ad Immunostaining of -gal (green) in Gli1-LacZ and Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mouse incisors. b, d represent the high-magnification images of the boxes in (a, c). White dotted lines outline the cervical loop. Yellow arrows point to the positive cells. dpt, day post-tamoxifen injection. e Quantification of GLI1+ cells in dental mesenchyme. Data are meanSEM, Gli1-LacZ n=4, Gli1-CreER;Arid1bfl/fl;Gli1-LacZ n=3, unpaired two-tailed Students t-test. p=0.0007. fi EdU staining of the LRCs (green) of incisors from control (f, g) and Gli1-CreER;Arid1bfl/fl (h, i). g, i represent the high-magnification images of the boxes in(f, h). White dotted lines outline the cervical loop. Yellow arrows point to the positive cells. Schematics of tamoxifen induction and EdU labeling protocol under (ad, fi), created with BioRender.com released under a Creative Commons Attribution-NonCommercial-NoDerivs 4.0 Internatioanl license. j Quantification of LRCs in dental mesenchyme. Data are meanSEM, n=3, unpaired two-tailed Students t-test. p=0.0238. k UMAP visualization of integrated scRNA-seq and the subclusters in dental mesenchymal cells from control and Gli1-CreER;Arid1bfl/fl mouse incisors. MSC, mesenchymal stem cell; PDF, proximal dental follicle; TAC, transit-amplifying cell; DP, dental pulp; Pre-Od, pre-odontoblast. l Cell number percentage of dental mesenchymal cell clusters in control and Gli1-CreER;Arid1bfl/fl samples based on scRNA-seq data. m GeneSwitches comparison of pseudotime along the MSC to TAC trajectory between control and Gli1-CreER;Arid1bfl/fl mouse incisor samples. Green dots indicate the pseudo-timepoints of genes that have been switched off, while orange dots indicate the genes that have been switched on. The green and orange lines connect the same genes. nq Co-immunostaining of LRCs (green) and Ki67 (red) in control (n, o) and Gli1-CreER;Arid1bfl/fl (p, q) mouse incisors. o,q represent high-magnification images of boxes in (n, p). White dotted lines outline the cervical loop. Yellow arrows point to the co-stained positive cells. r Quantification of the percentage of LRC+/Ki67+ cell number in LRC+ cells from control and Gli1-CreER;Arid1bfl/fl mouse incisors. Data are meanSEM, n=3, unpaired two-tailed Students t-test. p=0.0333. Source data are provided as a Source Data file. Scale bars: 100m.

To understand the reason for the reduction in GLI1+ MSCs following the loss of Arid1b, we first conducted TUNEL assay to compare apoptosis levels between control and Arid1b mutant mice. The results showed that there was no obvious increase in apoptosis at 5 days post-induction, suggesting that the decrease in MSCs was not caused by an increase in cell death (Supplementary Fig.2gj). To further investigate the underlying causes for MSC changes, we performed single-cell RNA sequencing (scRNA-seq) on the proximal incisor tissue from both control and Gli1-CreER;Arid1bfl/fl mice 4 days post-tamoxifen induction. We integrated the scRNA-seq data from the control (9210 cells) and Gli1-CreER;Arid1bfl/fl (8915 cells) mice, and identified a large group of dental mesenchymal cells along with other cell types via unsupervised clustering and marker analysis (Supplementary Fig.3a, b). We then focused on the dental mesenchymal cells and subclustered them into five distinct populations (Fig.2k). We identified the specific markers for each cluster and validated their expression patterns in the mouse incisor (Supplementary Fig.3ce). To investigate the effects of Arid1b loss at the single-cell level across different cell clusters, we compared the relative percentage of cells in each cluster between control and Arid1b mutant samples. We observed a decrease in the number of MSCs in the Arid1b mutant sample, consistent with the in vivo marker analysis, alongside an increase in the TAC number (Fig.2l). The comparison at the single-cell level suggested that the loss of Arid1b may impact MSC quiescence by inducing MSC proliferation.

To validate our hypothesis, we utilized scRNA-seq data to construct a computational model of change across time between control and Arid1b mutant mouse incisor mesenchymal cells using Monocle3 pseudotime analysis (Supplementary fig.4a, b). This model confirmed the sequence of cell differentiation, showing that MSCs are the earliest (least differentiated) cells, followed by TACs and dental pulp cells, which are later and more differentiated cells. This data aligns with the in vivo dental mesenchymal cell differentiation axis. To gain further insights into the stem cell fate transition of dental mesenchymal cells, we conducted GeneSwitches analysis to identify the order of genes that are activated or repressed in the specific lineages30. We acquired sets of genes that switched on/off along the MSC to TAC trajectory in both control and Arid1b mutant samples (Supplementary Fig.4c, d). This analysis helped us identify cluster-specific markers, such as Igfbp6 for MSCs and Pclaf for TACs, which we compared between control and Arid1b mutant samples (Supplementary Fig.4e, f). Furthermore, we visualized and compared the patterns of common markers switching on/off along the MSC-TAC pseudotime trajectory between control and Arid1b mutant samples. Interestingly, the majority of switched-off genes displayed slightly earlier pseudo-timepoints in the Arid1b mutant, whereas the genes switched-on in TACs exhibited significantly earlier activation in the Arid1b mutant. This was especially true of proliferation-related genes, such as Mki67, Top2a, and Cdk1 (Fig.2m). This data indicated that loss of Arid1b may induce MSC proliferation. To confirm that the loss of Arid1b stimulates the proliferation of slow-cycling MSCs in vivo, we performed co-immunostaining of LRCs and Ki67 in the mouse incisor. The results revealed an increased number of LRC+ cells co-labeled with Ki67+ in Arid1b mutant mice compared to controls (Fig.2nr), indicating that the loss of Arid1b leads to the proliferation of MSCs. Taken together, these results suggested that the loss of Arid1b disturbs MSC quiescence and leads to the proliferation of MSCs.

To investigate the downstream mechanisms of ARID1B in regulating the MSC population, we performed bulk RNA-seq to compare transcriptional profiles between control and Gli1-CreER;Arid1bfl/fl mouse incisors. Hierarchical clustering and volcano plots revealed distinct transcriptional profiles between the two groups, identifying 155 upregulated and 55 downregulated genes (false discovery rate [FDR] 0.1; fold change<1.5 or >1.5) in Gli1-CreER;Arid1bfl/fl mouse incisors (Fig.3a, Supplementary Fig.5a). We then conducted a comprehensive analysis and listed the top 20 differentially expressed genes and significantly changed signaling pathways based on the upregulated and downregulated genes (Fig.3b, Supplementary Fig.5b, c). Using the scRNA-seq data, we plotted the top 20 changed genes(Supplementary Fig. 5d). Among these genes, Bcl11b showed specific enrichment in the MSC and proximaldental follicle (PDF)clusters, colocalizing with Gli1 expression (Fig.3c). We compared its expression level within dental mesenchymal cells and observed its upregulation in Arid1b mutant mouse incisors (Fig.3d). To confirm the expression pattern of Bcl11b in vivo, we colocalized Bcl11b with GLI1+ cells and found that Bcl11b is expressed in the proximal region of the mouse incisor surrounding the NVB and colocalized with GLI1+ cells (Fig.3e, h). Furthermore, we confirmed the upregulation of Bcl11b expression in Arid1b mutant mouse incisors compared to the control (Fig.3f, g, ik). Taken together, these findings gave a strong indication that Bcl11b might act as a functionaldownstream target of ARID1B to regulate MSC quiescence. Additionally, considering that Bcl11b encodes a subunit of the BAF complex, it is essential to investigate the inter-regulation among the BAF subunits.

a Heatmap of bulk RNA-seq data for the proximal region of control and Gli1-CreER;Arid1bfl/fl mouse incisor at 4 days post-tamoxifen induction. b Top 7 signaling pathways identified by GO analysis using upregulated genes identified from bulk RNA-seq analysis. c FeaturePlot of Bcl11b and Gli1 expression in the control mouse incisor sample. The red arrow points to the co-localized positive cells. d ViolinPlot of Bcl11b expression levels in control and Gli1-CreER;Arid1bfl/fl mouse incisor mesenchymal cells. e, h In situ hybridization of Bcl11b (red) and immunostaining of -gal (green) in Gli1-LacZ mouse incisor. h represents a high-magnification image of the box in (e). The white dotted line outlines the cervical loop. White arrows point to the positive cells. n=3. f, g, i, j Comparison of Bcl11b (red) expression in control (f, i) and Gli1-CreER;Arid1bfl/fl (g, j) mouse incisors at 4 days post-tamoxifen induction. i,jrepresent high-magnification images of the boxes in (f,g), respectively. The white dotted line outlines the cervical loop. Yellow arrows point to the positive signals. k Quantification of the Bcl11b expression level per cell in (i, j). Data are meanSEM, n=3, unpaired two-tailed Students t-test. p=0.0142. Source data are provided as a Source Data file. Scale bars, 100m.

To investigate how ARID1B suppresses Bcl11b expression, we conducted single-cell transposase-accessible chromatin sequencing (scATAC-seq) using the proximal region of incisors from both control and Gli1-CreER;Arid1bfl/fl mice 4 days after tamoxifen induction. By analyzing and integrating the sequencing data from control and Arid1b mutant mice using Signac and marker analysis, we identified distinct cell clusters (Fig.4a, Supplementary Fig.6). Gli1 putative gene activity was found to be significantly enriched in the MSC population (Fig.4b). To compare the differences in chromatin accessibility specifically within the proximal mesenchyme cells between the control and Arid1b mutant mice, we categorized the cells into Control-MSC, Mutant-MSC, and other clusters (Fig.4c). We generated the normalized signals to visualize the DNA accessibility and annotated peaks for Bcl11b. The scATAC-seq results revealed that the Mutant-MSC cluster had higher peaks compared to the Control-MSC cluster at the promoter region of Bcl11b, indicating increased chromatin accessibility at the promoter region following the loss of Arid1b (Fig.4d). These results provided evidence that ARID1B functions as a suppressor of Bcl11b gene expression in the mouse incisor.

a UMAP visualization of integrated scATAC-seq data from control and Gli1-CreER;Arid1bfl/fl mouse incisors at 4 days post-tamoxifen induction. b FeaturePlot of Gli1 putative gene activity of scATAC-seq data. c Re-clustered UMAP visualization with control-MSC, mutant-MSC, and other clusters. d Peak calling from scATAC-seq at Bcl11b gene. The annotated open chromatin of the changed intron region and promoter region of Bcl11b are enlarged in the box below. e CHIP-qPCR primers and CRISPRi gRNA design at the targeted intron region of Bcl11b. Black arrows indicate the position and direction of forward and reverse primers. Red bolded line indicates the gRNA position. f CHIP assay with ARID1B antibody (or immunoglobulin G [IgG]), followed by qPCR with designed primers at the intron region of Bcl11b. Data are meanSEM, n=3, unpaired two-tailed Students t-test. p=0.0007. g RT-qPCR analysis of Bcl11b expression following CRISPRi treatment with vectors (generated by Origene Technologies) containing control and Bcl11b gRNA. Data are meanSEM, n=4, unpaired two-tailed Students t-test. p=0.0141. hm -gal (green) immunostaining of incisors from Gli1-LacZ (h, k), Gli1-CreER;Arid1bfl/fl;Gli1-LacZ (i, l), and Gli1-CreER;Arid1bfl/fl;Bcl11bfl/+;Gli1-LacZ (j, m) mice 5 days post-tamoxifen induction. k, l, m represent high-magnification images of boxes in (hj), respectively. White dotted lines outline the cervical loop. Yellow arrows point to the positive cells. Scale bars, 100m. (n) Quantification of the GLI1+ cells in dental mesenchyme of incisors from Gli1-LacZ, Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, and Gli1-CreER;Arid1bfl/fl;Bcl11bfl/+;Gli1-LacZ mice. Data are meanSEM, n=3, unpaired two-tailed Students t-test. Gli1-LacZ vs Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, p=0.0039; Gli1-CreER;Arid1bfl/fl;Gli1-LacZ vs Gli1-CreER;Arid1bfl/fl;Bcl11bfl/+;Gli1-LacZ, p=0.0078. Source data are provided as a Source Data file.

To investigate whether ARID1B directly binds to the cis-regulatory elements of Bcl11b to regulate its expression, we compared the peaks between control and Arid1b mutant scATAC-seq data, and identified a reduced peak call at Chr12: 107964756-107965631, corresponding to the Bcl11b third intron region in Arid1b mutant sample (Fig.4d). To identify whether ARID1B can bind to this 865 base-pair (bp) region, we designed primers within this region and performed ARID1B chromatin immunoprecipitation followed by quantitative PCR (CHIP-qPCR) (Fig.4e). The CHIP-qPCR results revealed significantly higher DNA binding of ARID1B CHIP compared to the IgG control in the +101bp to +262bp binding region (as shown in Fig.4e between the black arrows), suggesting that ARID1B can directly bind to the intron of Bcl11b to suppress its expression (Fig.4f). To further validate the functional relevance of this binding site, we performed CRISPRi to specifically target the binding region and assess its effect on the transcriptional activity. It is known that the dCas9 for CRISPRi exerts its binding activity within its gRNA target position with a range of 150bp31,32. Accordingly, we designed a gRNA around the qPCR-targeted region with high targeting efficiency and low off-target rate (Fig.4e). Following CRISPRi treatment of primary cells, we observed a significant increase in the Bcl11b expression level in the group treated with the vector containing Bcl11b gRNA when compared to the group treated with the control vector (Fig.4g), indicating that the expression level of Bcl11b was upregulated when there was interference with the ARID1B binding site.

Furthermore, to confirm the role of Bcl11b as a key downstream mediator of ARID1B in regulating MSC homeostasis, we generated Gli1-CreER;Arid1bfl/fl;Bcl11bfl/+;Gli1-LacZ mice. By comparing the numbers of GLI1+ MSCs with Gli1-LacZ and Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mice, we aimed to assess the impact of Bcl11b expression on MSC populations in vivo. Through immunostaining, we observed that the reduction of Bcl11b expression in Gli1-CreER;Arid1bfl/fl;Bcl11bfl/+ mice led to therestoration of GLI1+ cells (Fig.4hn). These compelling findings strongly supported the notion that Bcl11b serves as the functional downstream target of ARID1B, playing a crucial role in regulating MSC dynamics. Moreover, our data unveiled the direct inter-regulatory interactions between ARID1B and BCL11B during their functional regulation.

Using the differentially expressed genes identified from bulk RNA-seq, we investigated the significant changes in signaling pathways following the loss of Arid1b in the mouse incisor. Through gene ontology (GO) analysis, we identified highly enriched upregulated and downregulated signaling pathways based on the genes that showed significant increased or decreased expression in Arid1b mutant mice, respectively (Fig.3b, Supplementary Fig.5c). Among these pathways, we observed significant changes in angiogenesis and its related p53 pathway, as well as FGF, Wnt, Endothelin, and EGF signaling, with both upregulated and downregulated genes. This suggested that these signaling pathways may not be directly regulated by the loss of Arid1b. TGF- signaling exhibited significant upregulation in Arid1b mutant mice, indicating that this could be a downstream signaling pathway that undergoes substantial modulation following the loss of Arid1b. We therefore explored the role of TGF- signaling and sought to determine the specific ligands and receptors involved in maintaining MSC homeostasis.

Previous studies on Bcl11b-deficient mice have shown downregulation of TGF- family ligands during embryonic incisor development, including BMP4 and activin, leading to disruption of TGF- signaling33. We thus proposed that TGF- signaling superfamily may exert as a downstream regulatory effect of BCL11B. To test our hypothesis, we compared the expression patterns and levels of ligands and receptors within the TGF- superfamily, including TGF- signaling, Activin/Inhibin signaling, and BMP signaling, using integrated scRNA-seq and bulk RNA-seq data. Our analysis did not reveal significant changes in BMP signaling (Supplementary Fig.7). However, we observed significant upregulation of Tgfbr1 and Inhba following the loss of Arid1b in the mouse incisor from the bulk RNA-seq data. Inhba encodes a subunit for the Activin signaling ligand, activin A. Notably, when we plotted these ligands and receptors in the scRNA-seq data, we observed that Inhba was predominantly enriched in the PMC and dental follicle clusters, which coincided with Bcl11b expression (Fig.5a, Supplementary Fig.8a, b). However, Tgfbr1 exhibited widespread expression in the dental mesenchyme, and according to the JASPAR transcription factor (TF) binding sites prediction tool integrated into the UCSC Genome Brower, there is no predicted binding site for BCL11B at the promoter region of Tgfbr1 (Supplementary Fig.8b, c). This indicated that the increase of Tgfbr1 may not be directly regulated by BCL11B. Thus, we turned our focus to Inhba as a potential target of BCL11B within the TGF- superfamily. To further investigate the expression of Inhba and its colocalization with Bcl11b in vivo, we performed in situ hybridization. The staining revealed the colocalization of Inhba and Bcl11b within the MSC region and the dental follicle of the control incisors (Fig.5bd), indicating BCL11B may directly regulate Inhba expression.

a FeaturePlot shows the colocalization of Bcl11b and Inhba in the control scRNA-seq data. Red arrows point to the co-localized positive cells. bd in situ hybridization of Bcl11b (green) and Inhba (red) in the proximal region of the mouse incisor. c and d represent high-magnification images of boxes in (b). White dotted line outlines the cervical loop. Yellow arrows point to the co-stained positive cells. n=3. e UCSC binding prediction of BCL11B binding motif to the promoter region of Inhba. The red box emphasizes the binding site with the highest score. f Peak calling from scATAC-seq at Inhba. The annotated open chromatin at the promoter region of Inhba are enlarged at the right-side box. g CHIP assay with BCL11B antibody (or immunoglobulin G [IgG]), followed by qPCR with primers designed for the promoter region of Inhba. Data are meanSEM, n=3, unpaired two-tailed Students t-test. p=0.0337. hm Immunostaining of Activin A (red) in the incisors from control (h, k), Gli1-CreER;Arid1bfl/fl (i, l), and Gli1-CreER;Arid1bfl/fl;Blc11bfl/+ (j, m) mice 5 days post-tamoxifen induction. k, l,m represent high-magnification images of boxes in (hj), respectively. White dotted lines outline the cervical loop. Yellow arrows indicate positive cells. n=3. n Schematic drawing of the proximal ends of mouse incisors in explant culture created with BioRender.com released under a Creative Commons Attribution-NonCommercial-NoDerivs 4.0 Internatioanl license. oq -gal (green) immunostaining of incisor explants of Gli1-lacZ treated with IgG beads (o), Gli1-CreER;Arid1bfl/fl;Gli1-lacZ treated with IgG beads (p), and Gli1-CreER;Arid1bfl/fl;Gli1-lacZ treated with anti-ACTIVIN A beads (q). White dotted lines outline the cervical loop. White arrows indicate positive cells. Yellow boxes indicate the area for quantification. n=3. r Quantification of the GLI1+ cells in dental mesenchyme of incisor explants. Data are meanSEM, n=3, unpaired two-tailed Students t-test. Gli1-LacZ - IgG vs Gli1-CreER;Arid1bfl/fl;Gli1-LacZ - IgG, p=0.0176; Gli1-CreER;Arid1bfl/fl;Gli1-LacZ - IgG vs Gli1-CreER;Arid1bfl/fl;Gli1-LacZ - ACTIVIN A, p=0.0125. Source data are provided as a Source Data file. Scale bars, 100m (b, hm, oq); 200m (c, d).

To elucidate the mechanism by which BCL11B, acting as a transcription factor, mediates the regulation of Inhba expression, we investigated the direct binding of BCL11B to the promoter region of Inhba. Utilizing the JASPAR TF binding sites prediction, we identified two potential binding sites located within 1.5kb upstream of the transcription start site (TSS) in the promoter region (Fig.5e). We specifically targeted the binding site closer to the TSS, as it exhibited a higher predicted binding score according to JASPAR prediction. We first compared the scATAC-seq data for Inhba and observed higher peaks in the Mutant-MSC cluster compared to the Control-MSC cluster at the promoter region of Inhba (Fig.5f). This data indicated increased chromatin accessibility at the Inhba promoter region in the Arid1b mutant mice. Furthermore, we designed primers according to the predicted binding motif and performed BCL11B ChIP-qPCR in control mouse incisors. The result revealed significantly higher DNA binding of BCL11B CHIP than the IgG control at the promoter region of Inhba, indicating that BCL11B directly binds to the promoter region and regulates Inhba gene expression (Fig.5g). These findings provided evidence supporting the direct regulation of Inhba expression by BCL11B.

Furthermore, we examined the ligand activin A level through immunostaining and found that it was primarily deposited in the TAC region, odontoblasts, and dental pulp cells near the odontoblasts in control incisors. We noticed an ectopic upregulation of activin A in the MSC region following the loss of Arid1b, which was restored to normal after reducing Bcl11b level in Arid1b mutant mice (Fig.5hm). These findings indicated that BCL11B functionally and directly regulates the expression of the activin A subunit Inhba. Activin A, as the downstream target of BCL11B, undergoes ectopic upregulation in the MSC region within the Gli1-CreER;Arid1bfl/fl mouse incisor. To further explore the impact of ACTIVIN A upregulation in Arid1b mutant mice and its potential effect in the reduction of GLI1+ MSCs, we conducted an explant culture experiment. Using IgG or neutralizing ACTIVIN A antibody-loaded beads, we treated Gli1-LacZ and Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mouse incisor explants (Fig.5n). Immunostaining confirmed the decrease of GLI1+ MSCs in the Arid1b mutant mouse incisors compared to controls treated with IgG (Fig.5o, p, r). Notably, treatment of Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mouse incisor explants with the ACTIVIN A antibody restored the normal level of GLI1+ MSCs, in comparison to IgG treatment (Fig.5pr). These findings strongly suggested that ACTIVIN A serves as a functional downstream factor capable of maintaining MSC homeostasis in mouse incisors affected by the loss of Arid1b.

Activin A, the ligand that activates the Activin signaling pathway, belongs to the TGF- superfamily. Activin A binds to the extracellular domain of a type II receptor, ActRIIA or ActRIIB, then forms a complex with the type I receptor, ActRI or TGFBR1. The type I receptor phosphorylates downstream signaling molecules, leading to the activation of the Activin signaling pathway. Upon ligand binding, TGF-/Activin receptors activate intracellular signaling pathways such as the canonical Smad-dependent pathway via Smad2/Smad3, and non-Smad pathways including p38 MAPK, JNK, ERK, AKT, NF-B, and COFILIN signaling34,35. These signaling pathways act in a context-dependent manner, leading to different cellular responses. To investigate changes in Activin signaling activity in the Gli1-CreER;Arid1bfl/fl mouse incisor compared to the control, we initially conducted western blot to assess the changes in the signaling pathways. Interestingly, we observed that p-JNK, p-ERK, and p-COFILIN signaling were elevated following the loss of Arid1b, while p-SMAD2, the readout of canonical TGF-/Activin signaling, exhibited no significant changes (Fig.6a, Supplementary Fig.9). To further corroborate these findings and explore where these signaling pathways were activated in vivo, we performed immunostaining for p-JNK, p-ERK, and p-COFILIN in control and Arid1b mouse incisors. Our data revealed that p-COFILIN and p-JNK were specifically activated within the MSC region, as evidenced by their co-localization with GLI1+ cells, and exhibited heightened expression upon Arid1b loss (Fig.6bm). Interestingly, p-ERK, which was not activated in the MSC region of the control incisor, exhibited ectopic activation in the MSC region after loss of Arid1b (Fig.6nu). Our data provided robust evidence suggesting that non-canonical Activin signaling pathways play a pivotal role in the regulation of MSC homeostasis.

a Western blot of ARID1B, p-SMAD2, SMAD2, p-AKT, AKT, p-P38, P38, p-ERK, ERK, p-JNK, JNK, p-COFILIN, COFILIN, p-NF-B, NF-B, and -ACTIN in proximal incisor mesenchyme from control and Gli1-CreER;Arid1bfl/fl mice. bg Immunostaining of p-COFILIN (red) with -gal (green) at Gli1-LacZ mouse incisor (b, e) and comparison of p-COFILIN expression in control (c, f) and Gli1-CreER;Arid1bfl/fl (d, g) mouse incisors. eg represent high-magnification images of boxes in (bd), respectively. White dotted lines outline the cervical loop. Yellow arrows indicate positive cells. n=3. hm Immunostaining of p-JNK (red) with -gal (green) in Gli1-LacZ mouse incisor (h, k) and comparison of p-JNK expression in control (i, l) and Gli1-CreER;Arid1bfl/fl (j, m) mouse incisors. km represent high-magnification images of boxes in (hj), respectively. White dotted lines outline the cervical loop. Yellow arrows indicate positive cells. n=3. no Immunostaining of p-ERK (red) with -gal (green) in Gli1-LacZ. o represents the high-magnification image of (n). White dotted line outlines the cervical loop. The asterisk in (o) represents the absence of co-stained cells. pu Immunostaining of p-ERK (red) in control (pr) and Gli1-CreER;Arid1bfl/fl (su) mouse incisors. q, r represent the high-magnification image of boxes in (p),t, u represent the high-magnification image of boxes in (s). White dotted lines outline the cervical loop. Yellow arrows indicate positive cells. n=3. Scale bars, 100m.

Additionally, we investigated where p-SMAD2 was activated in both control and Gli1-CreER;Arid1bfl/fl mouse incisors. Our results demonstrated that p-SMAD2 was primarily activated in dental pulp cells and adjacent to GLI1+ cells (Supplementary Fig.10ah). This finding identified that Smad-dependent TGF-/Activin signaling is confined within the differentiated dental pulp compartment, delineating the boundary between the MSCs and dental pulp cells in the mouse incisor. This spatial segregation remained unaltered following the loss of Arid1b at 5 days post-tamoxifen induction.

To determine whether the elevated non-canonical Activin signaling was the cause of the incisor defect observed in the Gli1-CreER;Arid1bfl/fl mice, we generated Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+ mice as a rescue model. This approach was based on the significant upregulation of Tgfbr1 following the loss of Arid1b (Supplementary Fig.8b), and the notion that TGFBR1 plays a crucial role in TGF-/Activin signaling by forming receptor dimers with TGFBR2 and ActRII, transporting signals from intercellular to intracellular. Through in situ hybridization, we detected Tgfbr1 signal widely expressed in the dental mesenchyme, which was upregulated in Arid1b mutant mouse incisors compared to the control (Supplementary Fig.10in). Tamoxifen was injected at 1 month of age to induce Cre activity. At 3 months post-tamoxifen induction, microCT images revealed that the narrowed dental pulp cavity observed in Gli1-CreER;Arid1bfl/fl mice were completely rescued in Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+ mouse incisors. Moreover, HE staining further confirmed that the incisor defects in the Arid1b mutant mice, such as stacked dentin at the cervical loop, misaligned odontoblasts, and thicker dentin, were also rescued in Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+ mice. The odontoblast marker Dspp showed that the premature differentiation of odontoblasts was eliminated in the rescue mouse incisor as well (Fig.7ai). We assessed the numbers of GLI1+ cells in Gli1-LacZ, Gli1-CreER;Arid1bfl/fl;Gli1-LacZ and Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+;Gli1-LacZ mice 5 days after tamoxifen induction and confirmed that the reduced number of GLI1+ MSCs observed in the Gli1-CreER;Arid1bfl/fl;Gli1-LacZ mouse incisors was restored in the Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+;Gli1-LacZ mice (Fig.7jp).

ai The reduction of Tgfbr1 rescues the phenotypes observed in Gli1-CreER;Arid1bfl/fl mouse incisors. MicroCT (a, d, g), HE staining (b, e, h), and Dspp (red) (c, f, i) in situ hybridization of incisors from control (ac), Gli1-CreER;Arid1bfl/fl(df), and Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+ (gi) mice at 3months post-tamoxifen induction. White arrow indicates the narrowed dental pulp. Boxes in (b, e, h) are shown at higher magnification on the right. Yellow arrows indicate the initiation of odontoblast polarization. White two-way arrows indicate the dentin thickness. Black asterisk indicates stacked and distorted dentin. Yellow dotted lines show the distance between the cervical loop bending point and the initiation of the odontoblast. Unfilled arrows indicate the distance of the yellow dotted lines. n=3. jo Immunostaining of -gal (green) in Gli1-LacZ (j, m), Gli1-CreER;Arid1bfl/fl;Gli1-LacZ (k, n), and Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+;Gli1-LacZ (l, o) mouse incisors. m, n,o represent high-magnification images of the boxes in (jl), respectively. White dotted lines outline the cervical loop. Yellow arrows indicate positive cells. n=3. p Quantification of the GLI1+ cells in the dental mesenchyme of incisors from Gli1-LacZ, Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, and Gli1-CreER;Arid1bfl/fl;Tfgbr1fl/+;Gli1-LacZ mice. Data are meanSEM, n=3, unpaired two-tailed Students t-test. Gli1-LacZ vs Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, p=0.0102; Gli1-CreER;Arid1bfl/fl;Gli1-LacZ vs Gli1-CreER;Arid1bfl/fl;Tfgbr1fl/+;Gli1-LacZ, p=0.013. q Western blot of ARID1B, p-JNK, p-COFILIN, p-ERK, and -ACTIN in the proximal incisor mesenchyme from control (C), Gli1-CreER;Arid1bfl/fl (M), and Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+ (T) mice. rw Immunostaining of -gal (green) in Gli1-LacZ (r, s), Gli1-CreER;Arid1bfl/fl;Gli1-LacZ (t, u), and Gli1-CreER;Arid1bfl/fl;Erk2fl/+;Gli1-LacZ (v, w) mouse incisors. s, u, w represent high-magnification images of the boxes in (r, t, v), respectively. White dotted lines outline the cervical loop. Yellow arrows indicate positive cells. n=3. x Quantification of GLI1+ cells in the dental mesenchyme of incisors from Gli1-LacZ, Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, and Gli1-CreER;Arid1bfl/fl;Erk2fl/+;Gli1-LacZ mice. Data are meanSEM, n=3, unpaired two-tailed Students t-test. Gli1-LacZ vs Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, P=0.0037; Gli1-CreER;Arid1bfl/fl;Gli1-LacZ vs Gli1-CreER;Arid1bfl/fl;Erk2fl/+;Gli1-LacZ, p=0.0055. Source data are provided as a Source Data file. Scale bars: 2mm (a, d, g); 100m for the rest of the images.

Furthermore, to investigate whether the dysregulated non-canonical Activin signaling had been restored in the Tgfbr1 rescue mouse model, we performed western blot analysis of p-JNK, p-ERK, and p-COFILIN on control, Gli1-CreER;Arid1bfl/fl, and Gli1-CreER;Arid1bfl/fl;Tgfbr1fl/+ mouse incisors. Notably, only the p-ERK signaling showed obvious restoration, suggesting that it plays a key role in the non-canonical Activin signaling associated with the loss of Arid1b in the mouse incisors (Fig.7q). To validate that the p-ERK signaling pathway is the functionally downstream, we generated Gli1-CreER;Arid1bfl/fl;Erk2fl/+;Gli1-LacZ mice. By comparing GLI1+ cell populations among Gli1-LacZ, Gli1-CreER;Arid1bfl/fl;Gli1-LacZ, and Gli1-CreER;Arid1bfl/fl;Erk2fl/+;Gli1-LacZ mouse incisors, we observed the restoration of GLI1+ cells in the Erk2 rescue mouse model (Fig.7rx). Ultimately, the loss of Arid1b led to the activation of p-ERK signaling in MSCs following activin A ligand binding to TGFBR1-associated receptors. The aberrant p-ERK signaling disrupted MSC homeostasis and diminished their population.

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Enhancing prime editing in hematopoietic stem and progenitor cells by modulating nucleotide metabolism - Nature.com

"Animal Stem Cell Therapy Market 2024,"

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Animal Stem Cell Therapy Market Size, Share, Industry, Forecast and outlook (2024-2031) - openPR