Category Archives: Stem Cells

Millions of pregnant women get the pitch through their OB-GYN: Put a bit of your newborns umbilical cord on ice, as a biological insurance policy. If your child one day faces cancer, diabetes or even autism, the precious stem cells in the cord blood could become a tailor-made cure.

Many families are happy to pay for the assurance of a healthy future. More than 2 million umbilical cord samples sit in a handful of suburban warehouses across the country. Its a lucrative business, with companies charging several thousand dollars upfront plus hundreds more every year thereafter. The industry has grown rapidly, bolstered by investments from medical device companies, hospital partnerships and endorsements from celebrities such as Drew Barrymore and Chrissy Teigen.

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Cord Blood Registry, in Tucson, Ariz., on June 26, 2024. The company stores more than one million samples, double the number it had in 2014. (Rebecca Noble/The New York Times)

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Renee Johnson in Scottsdale, Ariz., on July 7, 2024. Johnson banked her sons cord blood with ViaCord in 2014 and learned years later the cells were infected with E. coli. (Rebecca Noble/The New York Times)

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Anna Lazos at her home in Egg Harbor Township, N.J., on June 27, 2024. After spending thousands of dollars on cord blood storage, Lazos asked to withdraw a sample to enroll her son in an autism clinical trial. The company told her that the cells were contaminated with E. coli. (Hannah Yoon/The New York Times)

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Jenna Edwards with her son at home in Parkland, Fla., on June 24, 2024. When Edwards tried in 2017 to withdraw her sons cord blood cells for a clinical trial to treat his cerebral palsy, she learned that the company had found bacteria in the sample, but still charged her for the next two years. (Eva Marie Uzcategui/The New York Times)

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Read more from the original source:
Cord Blood Banks Sold Families on False Hope - myheraldreview.com

Faced with repairing a major nerve injury to the craniofacial region, surgeons can use a nerve from an arm or leg to restore movement or sensation. This approachknown as an autograftis the standard of care, but it can take a toll on a previously uninjured body part, and the procedure doesnt always result in complete and functional nerve regrowth. Dr. Anh Le, Chair and Norman Vine Endowed Professor of Oral Rehabilitation in the Department of Oral and Maxillofacial Surgery at Penn Dental Medicine, is pioneering a different approach.

Le and collaborators are coaxing gingival mesenchymal stem cells (GMSCs)stem cells from gum tissueto produce nerve-supportive cells that facilitate nerve regrowth.

We wanted to create a biological approach and use the regenerating ability of stem cells, said Le. To be able to recreate nerve-supportive cells in this way is really a new paradigm.

For more than a decade, Les lab has explored the use of GMSCs to regenerate different types of craniofacial tissues and to treat osteonecrosis of the jaw, which can occur when a patient takes bisphosphonate, a drug used to treat metastatic cancer or prevent bone loss in osteoporosis. Her lab team was able to apply their previous understanding of GMSCs to facilitate their conversion into Schwann-like cells, the pro-regenerative cells of the peripheral nervous system that make neural growth factors and myelin, the insulating layer around nerves.

To move the work forward, Le collaborated with bioengineer D. Kacy Cullen of Penn Medicine, an expert in creating and testing nerve scaffold materials. Together they showed that infusing a collagen scaffold with these cells and using them to guide the repair of facial nerve injuries in animals was just as effective as an autograft procedure. Although the repaired gap was small, the team is continuing to refine the method to repair larger ones that often result from trauma or tumor-removal surgeries.

Le notes that this approach would enable patients with oral cancer or facial trauma to use their own tissue to recover motor function and sensation following a repair.

While Les group focuses on the head and neck, further work on this model could translate to nerve repair in other areas of the body as well.

Im hopeful we can continue moving this forward toward clinical application, she said.

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Penn Dental Medicine Researchers Coaxing Stem Cells from Gum Tissue to Repair Nerves - Dentistry Today

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"My focus will be on people and scrutinizing processes so that they better support the diversity of needs of our members across identities including geographies and career stages," Dr. Greco said."In turn this will increase opportunities for professional growth of our members and augment our collective impact. As I commit to this vision, I equally commit to speak with openness about the struggles that we have and will experience in order to make this vision a reality.

Credit: Yale School of Medicine

The ISSCR is thrilled to announce Valentina Greco, Yale School of Medicine, Genetics Department andYale Stem Cell Center USA, as its President. Her term began at the ISSCR 2024 Annual Meeting held in Hamburg, Germany that concluded on Saturday, 13 July 2024.

I am honored to be taking on the role of ISSCR President for the coming year, Dr. Greco said. Building on Amander Clarks efforts, my focus will be on people and scrutinizing processes so that they better support the diversity of needs of our members across identities including geographies and career stages. In turn this will increase opportunities for professional growth of our members and augment our collective impact. As I commit to this vision, I equally commit to speak with openness about the struggles that we have and will experience in order to make this vision a reality, Dr. Greco added.

Valentina Greco was born in Palermo, Italy on 3 September 1972. She earned an undergraduate degree in Molecular Biology at the University of Palermo, Italy (1996) where she studied the role of tumor suppressors in cell cycle using in vitro system in the lab of Aldo Di Leonardo (1995-1998), her first academic research experience. She was accepted by Suzanne Eaton and the EMBL/MPI-CBG PhD program, Germany (1998-2002) and fell in love with microscopy and the power of developmental biology using fly wing imaginal disc as a model system to understand epithelial cell communication. Dr. Greco subsequentially completed a post-doc training in the Fuchs lab at the Rockefeller University (2003-2009) where she learned about the mammalian skin hair follicle as model system for stem cell driven regeneration. She was then hired as an Assistant Professor in the Genetics department at Yale School of Medicine by Richard Lifton and Haifan Lin (1 August 2009).

Dr. Greco is currently the Carolyn Walch Slayman Professor of Genetics as well as the Co-chair of Status of Women in Medicine (SWIM) at the Yale School of Medicine. She and members of her lab feel excited about visual driven research to study how cells behave in a living mouse. The team understands cell behaviors as an expression of the architectures and principles that govern the tissues thesecells inhabit, much like human behaviors are an expression of the systems and structuresin whichthey are embedded(e.g. a lab, anorganization). The Greco lab is passionate about identifying the mechanisms that govern communication and cooperation to sustain function over a lifetime.

Dr. Greco has served in numerous leadership roles in the scientific community including many within the ISSCR over the last decade. She also serves on numerous additional boards including President Elect for the Society of Investigative Dermatology (SID), SID Board member 2016-2020, Member of the National Arthritis and Musculoskeletal and Skin Diseases Advisory Council (NAMSAC (2022-2024)), Member of the Yale Ciencia Academy Advisory Committee, Member of the 2030STEM Leadership Team and Secretary of Board of Directors of the Life Science Editors Foundation (2020-2023).

Greco lab research has been recognized by numerous accolades awarded to both lab members and Dr. Greco. She in particular has received the 2014 Women in Cell Biology Junior Award (WICB) for Excellence in Research from the American Society of Cell Biology (ASCB), the 2014 ISSCR (International Society for Stem Cell Research) Outstanding Young Investigator Award, the 2015Robertson Stem Cell Investigator Award from the New York Stem Cell Foundation (NYSCF), the 2015 Mallinckrodt Scholar Award, the 2016 Early Career Award from the American Society of Cell Biology (ASCB), the 2016 HHMI Faculty Scholar Award, the 2017 Glenn Foundation Award, the 2017 Class of 61 Award by the Yale Cancer Center, the 2019 NIH DP1 Pioneer Award and the 2021 ISSCR Momentum Award. Dr. Greco finds it particularly meaningful to have received the 2018 Yale Mentoring Award in the Natural Sciences, the 2019 Yale Genetic Department Mentoring Award, the 2019 Yale Post-doc Mentoring Award.

The ISSCR is equally pleased to announce Hideyuki Okano, MD, PhD, Keio University, Japan is President-elect and will serve as president officially starting 1 July 2025. Lorenz Studer, MD, founding director of the Center for Stem Cell Biology and member of the Developmental Biology Program, Memorial Sloan Kettering Cancer Center, USA, is the new Vice President.

The following three members are newly elected to the ISSCR Board of Directors and beginning their three-year term: Jacqueline Barry, PhD, Cell and Gene Therapy Catapult, UK, Tenneille E. Ludwig, PhD, WiCell, USA, and Thomas A. Rando, MD, PhD, University of California, Los Angeles, USA.

The following members are starting their second, three-year term as a result of the 2024 election: Melissa Carpenter, PhD, Carpenter Consulting Corporation, USA, Malin Parmar, PhD, Lund University, Sweden, and Mitinori Saitou, MD, PhD, Kyoto University, Japan.

Learn more about the ISSCR Board of Directors.

About the International Society for Stem Cell Research With nearly 5,000 members from more than 80 countries, the International Society for Stem Cell Research is the preeminent global, cross-disciplinary, science-based organization dedicated to stem cell research and its translation to the clinic. The ISSCR mission is to promote excellence in stem cell science and applications to human health. Additional information about stem cell science is available at AboutStemCells.org, an initiative of the Society to inform the public about stem cell research and its potential to improve human health.

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Valentina Greco takes on new position as President of the ISSCR - EurekAlert

ORANGE PARK, Fla. Dr. Eric Weiss, a certified plastic and reconstructive surgeon with a thriving practice on the outskirts of Jacksonville, Florida, still remembers the moment he realized the transformative effect umbilical cord stem cells could have on those with autism and other immune disorders.

First introduced to stem cell treatment by his wife Christine, who was searching for medical help for their autistic son, Marston, Weiss became intrigued by its possibilities.

Compelled by the research, and the subsequent positive impact stem cell treatment had on his son a journey chronicled in a book he and his wife subsequently co-authored Weiss soon incorporated it into his practices.

Studies show that people with autism have neuroinflammation, similar to other immune disorders, which utilize stem cells as a treatment.

With the advance of technology and a better understanding of autoimmune disorders and autism, regenerative medicine has been a proven solution to these seemingly unsolvable health concerns.

While many people are hesitant to pursue this treatment because of the excessive cost and lack of FDA approval, Weiss says, I understand these fears because not everyone gets better. But the role of neuro-diseases is changing and this treatment is proven to help patients recover on a cellular level.

When addressing the FDA concerns, Weiss stressed that the FDA regulates drug companies, not health care, and that stem cell blood has been used in treatment for over 60 years.

The stem cells in the umbilical cord hold abundant powerful cells that help the human bodys healing capabilities. Stem cells work by sending chemical signals to old, damaged or injured cells to restore them.

However, stem cell research has faced substantial opposition from various cultural and religious groups around the world. The controversies stem from differing beliefs regarding the beginnings of life, the moral status of the embryo and the ethical implications of manipulating human cells.

Most controversial of all has been human embryonic stem cell research, because it involves the destruction of human embryos.

Regarding its efficacy, there have been multiple studies demonstrating the positive results of stem cell treatments in both children and adults. These treatments include remedies for neuro-related health concerns such as:

Weiss ultimate goal is to restore wholeness to patients by using stem cells to treat multiple health conditions. He is one of the only physicians in the country to utilize this form of treatment.

As the human body ages, it works harder to keep you healthy. Those with conditions like autism or other ailments are constantly struggling to find solutions.

As we age, our stem cells begin to die off at an alarming rate:

While stem cells can be found in various places throughout the body, the cells in the umbilical cord are the most useful. It is the least invasive form of stem cell gathering and as a bonus, the umbilical cord is chock full of them according to Weiss.

Today, patients come from all over the world to receive this treatment.

Weiss now dedicates two days a week of his practice to administering stem cell treatment.

There is still much to study regarding stem cell treatment for autism and other neurological diseases, but Weiss points those who are wary back to the science and the literature.

There have been lab studies, animal studies and human studies that show positive results with this treatment, so I want to do everything I can to help these patients get better, he said.

With the hope that this treatment becomes a standard form of care, Weiss is learning more every day and helping families and patients who need it most.

I thought to myself, why wouldnt this work for autism? Luckily it has, and now Im able to help families who have been through the same struggles as my own, Weiss said.

You can reach us at [emailprotected] and follow us on Facebook

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Surgeon Turns to Stem Cells to Help Treat Autism and Immune Disorders - The Well News

Bryan Johnson, the billionaire tech CEO whose sole goal in life seems to be to live forever, underwent an experimental longevity treatment in the Bahamas last month that involved pumping himself up with 300 million stem cellsand documented it all for his YouTube channel. Johnson, 46, flew out to the Albany, a high-end Bahamas golf resort co-owned by the likes of Tiger Woods and Justin Timberlake, for the procedure. The stem cells, sourced from bone marrow by a company called Cell Colars Clinical, target a patients joints in the hopes of rejuvenating them, Johnson explained in a June 25 vlog. Those healthy young Swedish cells should multiply in my body, future-proofing all of my major joints and taking me one step closer to age 18, he said. The entrepreneur, who all but abandoned Silicon Valley for biohacking and a strict $2-million-a-year regimen he calls Project Blueprint, claimed earlier this year to have reversed his epigenetic age by a little over five years.

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Ellen DeGeneres Says Shes Done With Fame After Netflix Special Drops - The Daily Beast

Mice

Lrig1 constitutive knockout mice were generated by crossing Lrig1CreERT2 (Jackson Strain No. 018418, Lrig1tm1.1(cre/ERT2)Rjc/J) heterozygous mice on the 129-Elite strain background (Charles River Canada, strain code 476) together to produce homozygous knockout mice as described in ref. 14. Genotypes were confirmed post-weaning from ear notch genomic DNA (isolated with DirectPCR Lysis Reagent (Ear, Cat # 401-E)) using PCR primers and Jackson Protocol 26202 exactly as described. For all experiments female and male mice were randomly assigned to experimental groups and analyzed at ages of 8-weeks, 15-weeks and 24-weeks. Specific timepoints are noted in the figure legends and captions. Mice used in this work all followed a 12h light/dark cycle and had ad libitum access to water and rodent chow. Mice did not demonstrate any visible signs of being immunocompromised or display any obvious behavioural phenotypes. All animal work conducted followed policies formed by the Canadian Council of Animal Care and approved by the Local Animal Care Committee at the University of Toronto and The Hospital for Sick Children.

Frozen tissue sections were dried for 1h at room temperature or 30min at 37C, then rehydrated in phosphate buffered saline (PBS, 137mM NaCl, 2.7mM KCl, 10mM Na2HPO4, 1.8mM KH2PO4) for 10min. Subsequently, sections were blocked and permeabilized in a solution of 5% BSA and 0.3% Triton X-100 in PBS. Following blocking, sections were incubated in primary antibody diluted in 1:1 PBS to blocking solution overnight, at 4C in a humidified chamber. The following day sections were washed in PBS three times for 5min, then incubated in diluted secondary antibody for 2h at room temperature. Afterwards, sections were washed three times in PBS for 5min per wash. Finally, sections were counterstained for 5min at room temperature in 0.5g/mL Hoechst 33258 (Sigma-Aldrich), washed with PBS and then mounted with PermaFluor mountant (Thermo Scientific).

5-ethynyl-2-deoxyuridine (EdU, Toronto Research Chemicals Cat #: E932175) was dissolved in PBS and injected intraperitoneally three times two hours apart at a dose of 50mg/kg to 15-week-old Lrig1 WT and KO mice. After 3 weeks brains were collected and prepared for IHC as described below. EdU+ cells were detected using the Click-it EdU Alexa 488 kit (Invitrogen, Cat #: C10337) following the manufacturers instructions and then stained with Hoechst 33258. EdU+ cells were counted from five sections per brain.

Tissue sections from 15-week-old Lrig1 WT and KO mice were prepared as described below, blocked and incubated overnight with 1:100 anti-LRIG1 and anti-BMPR-1B (1:100 see Supplementary TableS3) antibodies as described above for IHC. The next day sections were washed three times for 10min with Wash Buffer A (0.01M Tris, 0.15M NaCl and 0.05% Tween-20) and then incubated with the Duolink In Situ PLA Probe Anti-Goat

PLUS (Sigma-Aldrich) and Duolink In Situ PLA Probe Anti-Mouse MINUS Affinity purified Donkey anti-mouse IgG (H+L) (Sigma-Alrich, See Supplementary TableS4) for 1hour at 37 C in a humidified chamber. Following washing of the PLA probes, Ligation and Amplification was carried out as described by the manufacturer using the Duolink In Situ Detection Reagents kit (Sigma-Aldrich) and nuclei stained for Hoechst 33258.

The periventricular area containing the V-SVZ from N=2 female and N=2 male Lrig1 KO and N=3 female and N=1 male WT 8-week old brains was dissected as described in ref.18 from both hemispheres of each brain and stored at 80C until needed. One hemisphere sample was used to isolate RNA as described below. The other hemisphere sample was lysed and western blotted for LRIG1 and pEGFR as described previously14.

Lrig1 KO and WT mice were anesthetized with 23% inhaled isoflurane and then perfused transcardially with PBS followed by 4% paraformaldehyde (PFA). Brains were then dissected, post-fixed overnight in 4% PFA and then cryoprotected in 30% sucrose in PBS for 24h. Tissues were then embedded with O.C.T (Fisher Healthcare Tissue-Plus O.C.T. Compound, Cat # 23-730-571) and sections were cut 18m thick in the coronal plane using a Thermo Fisher Scientific HM525 NX cryostat at 20C. Sections were collected on glass slides (Fisherbrand Superfrost Plus Microscope Slides, Cat #1255015) coated with gelatin and stored frozen until use.

All primary and secondary antibodies used for western blot are listed in Supplementary TablesS1 and S2, while those used for IHC/PLA are listed in Supplementary TablesS3 and S4.

For all experiments, plasmid constructs were used following endotoxin-free maxipreps using a Qiagen EndoFree Plasmid Maxi Kit or a ZymoPURE II Plasmid Maxiprep Kit. Plasmid DNA concentration was determined using a NanoDrop 2000 (ThermoFisher). All plasmids used in this study are listed in Supplementary TableS5.

Images were collected using a Zeiss Spinning Disk confocal microscope system or a Zeiss AxioImager M2 microscope system with a Calibri LED light source. Images were acquired using Z-stacks (with the apotome engaged in the case of the AxioImager) and staked tiles imaging set-up using Zen Blue software. All images were acquired with Z-stack sizes ranging between 15-25 slices depending on the dataset analyzed. Images shown were produced using the orthogonal projection feature implemented in Zen Blue.

For all cell count analysis on acquired images from V-SVZ, counting was done using ImageJ. For cell counts in the V-SVZ, only visibly immunostained (positive) cells along the dorsal and ventral portions of the lateral wall (LW), closest to the ventricle (periventricular area) were counted. To differentiate between the dorsal and ventral areas of the LW, these regions were measured using the line drawing and measurement tools in ImageJ along the length of the entire LW. To assess proliferation, neuronal progeny and pEGFR-positive cells in the V-SVZ, the top 1/3rd of the length of the LW was considered the dorsal portion. The bottom 2/3 measurement of the LW was considered the ventral portion. To obtain a cell count value for each individual brain, positive cells counted from three anatomically matched 18 m thin coronal sections containing the V-SVZ LW region were totaled and averaged. For the proportion of proliferating (Ki67+) cells, a percent cell count was used to represent the data by dividing the number of SOX2+GFAP+Ki67+/SOX2+GFAP+ cells. For the number of DCX-positive and pEGFR-positive cells the total number of cells counted per ventricle area was used to represent these data. For cell counts in the OB, positive cells in the GCL, GL and MCL were counted using three anatomically 18 m thick coronal sections of the whole OB from each mouse brain. To create a cell count value for each individual brain, counts from all three sections were totaled and averaged. For the number of CalB or CalR-positive interneurons, the total number of cells in each layer were counted and used to represent these data. pSmad2 and pSmad1/5/9 positive cells only in the V-SVZ were counted on the entire ventral portion of the LW of the LV. Three coronal sections from each brain which were anatomically matched in order of contain the LW area of the V-SVZ and were used for cells count analysis, where the total positive cells from each coronal section were then averaged.

Following RNA isolation using a RNeasy Plus Mini Kit (Qiagen, Cat#:74134) from tissue isolated as described above, polyA selected mRNA next generation sequencing libraries were prepared using the NEBNext Ultra II DNA Library Prep Kit and sequenced on one lane of an Illumina Novaseq SP flow cell achieving ~4050 million reads per sample. Following sequencing, FASTQ files were generated with bcl2fastq2 v2.20. Library preparation, sequencing and FASTQ file production were done by The Centre for Applied Genomics (TCAG) at the Hospital for Sick Children. FASTQ files were then used as input for Salmon19 for alignment to the mouse genome (Gencode m29) and for read quantification. Deseq2 as implemented in R and was used for normalization and differential gene expression analysis20.

Immunoprecipitation (IP) assays were done similar to those described in ref. 21. Neuro-2a cells (N2a) (ATCC Cat No: CCL-131) were used for all IP experiments which were cultured in Dulbeccos modified Eagles medium (DMEM) containing high levels of glucose supplemented with 1% penicillin-streptomycin and 10% fetal bovine serum (FBS). Transfections of N2a cells were carried out using Poly-Jet reagent (SignaGen Laboratories; Cat# SL100688) 24h after plating 400,000 cells per well of 6 well plate. Flag-tagged pCMV expression vectors containing either the TGFR1, TGFRII and BMPRI were used to co-transfect N2a cells with Lrig1 pCMV-overexpression plasmid using PolyJet In Vitro DNA Transfection Reagent (SignaGen Laboratories; Cat# SL100688). A mock condition with no plasmids was used as a negative control and BMPRI was used as a positive control21. N2a cells were lysed after 48h in TNTE buffer containing 0.5% Triton-X-100 (150mM NaCl, 50mM Tris pH 7.4, and 1mM EDTA)21. Anti-Flag Magnetic beads (Selleckchem Cat#: B26101) were used to carry out all immunoprecipitations. The beads and sample mixtures were incubated on a tube rotator at 4C for 2h. Supernatant was then discarded and the beads were washed three times with wash buffer (TNTE buffer containing 0.1% Triton-X100) on a magnetic separation rack. Following the final wash, magnetic beads were eluted using 1x SDS-PAGE loading buffer heated at 70C for 10min. 10% of the samples prior to immunoprecipitation were collected for a load condition in 5x SDS Sample Loading Buffer (10% SDS, 500mM DTT, 50% Glycerol, 250nM Tris-HCl, 0.5% bromophenol blue dye, pH 6.8). Western blotting was performed as described previously14 using anti-flag antibody (DYKDDDDK Tag Antibody Cell Signalling Technology Cat#: 2368) to detect the TGFR1, TGFRII and BMPRI receptors and LRIG1 antibody (R&D Systems, Cat#:3688) to detect LRIG1. GAPDH antibody was used as an internal reference to normalize protein expression levels.

LRIG1 ectodomain (ECD, Cat#: 3688-LR-050 from R&D Systems) concentrations of 2.5g/mL, 0.25g/mL, and 0.025g/mL were immobilized on a nitrocellulose membrane in duplicate. 2.5% Bovine serum albumin and 1xPBS were used as negative controls. Once dried, the blot was incubated at room temperature (RT) in blocking buffer (2% BSA in PBS-T (1xPBS containing 0.1% Tween)) for 30min. 1.2 ug of biotinylated human TGF1 (Avi- Tag, Biotin-Labeled, BPS Bioscence Cat #. 100843) was then added to blocking buffer on the blot and incubated for one hour at RT. A 1:5000 dilution of Streptavidin-HRP in blocking buffer was added for 30min followed by 35min and 315-minute washes with PBS-T. The blot was then developed with Bio-Rad Clarity Western enhanced chemiluminescence (ECL) Detection Reagent on a Syngene G:Box chemiluminescence imager.

The binding affinity of LRIG1 extracellular domain (ECD) to TGF1 was measured using a BLItz instrument. LRIG1 ECD in 1xPBS was used as the analyte and biotinylated-TGF1 was used as the bait. Streptavidin-coated sensors were hydrated in BLI rehydration buffer (PBS, 0.5mg/mL BSA, and 0.02% (v/v) Tween-20). The biotinylated bait protein was diluted in kinetics buffer (PBS, 0.5mg/mL BSA, 0.02% (v/v) Tween-20) to a final concentration of 12.4g/mL. The bait was immobilized on a streptavidin-coated biosensor tip for 30s. Next, the analyte concentrations were diluted in kinetic buffer in order to obtain concentration of LRIG1 ECD of 1M, 750nM, 500nM, 100nM. The binding association was measured over a period of 120s with subsequent dissociation measured after immersion of the biosensor tip into kinetic buffer for another 120s. The data were analyzed and sensorgrams were step corrected, reference corrected and globally fit to a 1:1 binding model. Dissociation constants (KD) were calculated using BLItz Pro Version 1.1.0.16 and the average of two independent determinations is reported here.

N2a cells were transfected with guide RNAs #1, #2, or LacZ control contained in the pU6-(Bbsl)_CBh_Cas9_T2A_mCherry plasmid (as described previously14 in using Lipofectamine STEM, according to manufacturers protocol in 6 well plates. Two days following transfection, cells were replated into 96 well plates at a concentration of 0.5 cells/200l media in each well. Cells were allowed to grow for 3 weeks and wells containing individual colonies that expressed red fluorescence from mCherry were selected and propagated further to produce clonal cell lines. Three control lines and four LRIG1 KO lines were selected following western blotting for LRIG1 to confirm loss of the LRIG1 protein and used for the experiments described here.

In the case of TGF1 and BMP4 treated non-transfected Ctrl N2a clonal cell lines and Lrig1 KO N2a clonal cell line experiments, cells were plated at 90,000 cells/well in DMEM media (described above) in 24-well plates. Two days later the media was changed to DMEM containing 0.1% FBS for serum starvation overnight. The following day, 10ng/ml TGF1 (Cedarlane; Cat# 781802) or BMP4 (Peprotech Cat#315-27) ligands suspended in DMEM containing 0.1% FBS. The later of which on its own was used as the control. Following 30min at 37C, the cells were collected using 50L of lysis buffer containing, tris-buffered Saline, pH 7.4, 0.5% b-octyl-D-glucopyranoside, 0.5% Triton X-100, 1mM NaF, 1mM -glycerophosphate, 1mM Na3VO4 and 1 cOmplete, mini, EDTA-free protease inhibitor tablet per 10ml. Protein assay was done with the samples using the DC Assay (Bio-Rad) and then equal amounts of protein were used for western blots to assess the pSmad2/3, total Smad2, pSMAD1/5/9 and total SMAD1 along with GAPDH (as a loading control) and LRIG1 (to confirm genotypes).

Since N2a cells express low levels of TGFRII, we used TGFRI and TGFRII constructs to express these receptors in Ctrl and Lrig1 KO clonal N2a cells followed by treatment with TGF1 to assess the response of this signalling pathway in the context of loss of LRIG1. To do so, 1g of DNA per well, using PolyJet of TGFRI and TGFRII were transfected one day after plating 90,000 cells per well of each of the Ctrl and Lrig1 KO clonal N2a cell lines. 24h after transfection, the cells were transitioned into DMEM containing 0.1% FBS media. The following the Lrig1 KO and WT clonal cell lines were treated with 10ng/ml per well of TGF1 for 30min at 37C, lysed, and protein concentrations determined as above. Western blotting for pSmad2/3, total Smad2 and GAPDH were then performed.

Neurospheres were grown essentially as described in ref. 18 using NeuroCult Mouse&Rat Basal Medium containing NeuroCult Proliferation Supplement (STEMCELL Technologies), FGF2 (Corning), EGF (STEMCELL Technologies), 0.2% heparin (Sigma-Aldrich, Cat #: H4784) and 1% penicillin-streptomycin from brains of 15 week old Lrig1 WT and KO mixed male and female mice (N=2 male and N=1 female KO mice, N=2 male and N=2 female WT mice). Following seven days of growth of secondary spheres, spheres from each brain were gently distributed into two wells of a 24-well plate and treated with BMP4 as described above without changing the culture media. Cells were then collected by centrifugation and western blots were performed as described above.

Data previously published by Mizrak et al.22, from lateral wall cells of 810 week old mice was obtained from NCBI GEO accession number: GSE109447. Specifically, the 13,055 cell data set was reduced to just the lateral wall cells and then was processed using Seurat (version 4.4.0) as implemented in R (version 4.2.1). Cells with less than 200 genes expressed in at least three cells were removed. The data were then normalized and scaled, principal components were computed and clustering was performed with 25 principal components and a resolution of 0.8. UMAP plots were created using the DimPlot and FeaturePlot functions as implemented in Seurat.

GraphPad Prism Version 9.3.1 software was used to perform all statistical analyses and create graphical representations. For statistical analyses in Figs.13, a two-way ANOVA and Tukeys Honestly Significant Difference (HSD) post-hoc test were used to compare data between the different experimental groups present in each paradigm (ex. KO vs. WT), and the area of the LW being analyzed (ex. dorsal vs ventral). In Fig.5, a two-tailed Students t-test in order to compare the differences between Lrig1 KO and WT groups. In Fig.7 and Supplementary Fig.S3, the results from the four conditions were averaged from two independent experiments and normalized to total SMAD2/3 for the TGFR transfections or total SMAD1 for the BMP experiments. Subsequently, a one-way ANOVA followed by a dk post-hoc test was used to compare groups. In all figures, values were reported as means. To create error bars, means were represented as the standard error of the mean (S.E.M). Experimental data was denoted as statistically significant at P values less than 0.05 (P<0.05). In all the figures, *p<0.05, **p<0.01, ***p<0.001 and ns = not significant.

a Immunostaining of the lateral wall of 15-week old Lrig1 KO and WT mice for GFAP, SOX2, Ki67 and merged with Hoechst to label nuclei. Sox2/GFAP-double positive cells allows identification of NSCs. Ki67/SOX2/GFAP-triple positive cells represent proliferating NSCs. Arrowhead indicate triple positive cells. Quantification of the number of SOX2/GFAP-double positive NSCs (b) and proportion of proliferating Ki67-positive NSCs (c) along the dorsal and ventral portions of the lateral wall in 15-week old Lrig1 KO and WT mice. d Immunostaining of the lateral wall of 24-week old Lrig1 KO and WT mice for GFAP, SOX2, Ki67 and Merged with Hoechst. Quantification of the number of SOX2/GFAP-double positive NSCs (e) and proportion of proliferating Ki67-positive NSCs (f) along the dorsal and ventral portions of the lateral wall in 24-week old Lrig1 KO and WT mice. Scale bars represents 25m. Error bars indicate S.E.M. For (ac, N=5 per group, and for df, N=3 per group. Source data for graphs are included in Supplementary Data2.

a Immunostaining of coronal sections through the lateral ventricle of 15-week old Lrig1 KO and WT mice for DCX to mark new-born neurons/neuroblasts and GFAP to label ventricular cells merged with Hoechst. b Quantification of the number of DCX-positive cells in the dorsal and ventral portions of the lateral wall. c Representative immunostaining of Olfactory Bulb (OB) sections with antibodies for Calretinin (CalR) and Calbindin (CalB) and merged with Hoechst at low magnification (left, indicating the Granule Cell Layer (GCL), the Mitral Cell Layer (MCL) and the Glomerular Layer (GL)) and at high magnification (right) from Lrig1 and KO mice. d Quantification of the number of CalB-positive cells in each of the granule cell (GCL), mitral cell (MCL) or glomerular layers (GL), (e) Quantification of the number of CalB-positive cells in the GCL, MCL and GL of the OB. f, g Quantification of the number of EdU-positive cells in the olfactory bulb (f) and representative staining of EdU-positive cells with Hoechst from an Lrig1 WT brain. Scale bars represent 50m for (a), 100m for (c) and 20m for (g). Error bars represent S.E.M. For (a, b, N=5 per group and for cf, N=3 per group. Source data for graphs are included in Supplementary Data2.

a Immunostaining of coronal sections along the lateral wall of Lrig1 KO and WT mice with antibodies for phosphorylated EGFR (pEGFR) and merged with Hoechst. b Quantification of the number of pEGFR-positive cells in the dorsal and ventral portions of the lateral wall of the V-SVZ. c Western blots using antibodies for LRIG1, pEGFR and GAPDH (loading control) from periventricular tissue dissected from one hemisphere per animal from 8-week old Lrig1 KO and WT mice See also Supplementary Fig.S2d. d Quantification of the western blot data from (c) for pEGFR relative to GAPDH. Scale bar represents 50um. Error bars represent S.E.M. For (a, b, N=5 per group and for c, N=4 per group. Source data for graphs are included in Supplementary Data2.

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LRIG1 controls proliferation of adult neural stem cells by facilitating TGF and BMP signalling pathways - Nature.com

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Derivation of spermatogenic cells from human embryonic stem cells (hESCs)

To determine the spermatogenic differentiation potential of non-manipulated hESCs, MUSIe002-A were subjected to a spermatogenic differentiation procedure. At the end of culture, the hESC-derived spermatogenic cells expressed several human spermatogenic marker proteins, including the pan-germ cell marker (VASA), spermatogonia stem cell markers (PLZF, CD90 and GPR125), and a spermatocyte marker (PIWIL1) (Fig.1A) compared to non-differentiate hESCs on culture day 0 (Fig. S1). In accordance with their protein expression, hESC-derived spermatogenic cells were subjected to transcriptional analysis by qRT-PCR. Up-regulation of SOX17 and TFAP2C was observed, which indicated the development toward germ cell lineages35. Furthermore, an early PCG marker BLIMP1, the post-migratory gonocyte PIWIL4 and the germline-specific gene VASA were up-regulated on day 5 and were maintained to the end of culture. Spermatogonia markers, including PLZF, GFRA, and NANOS3, were found to be highly expressed on day 5 and to remain throughout the culture period, while expressions of GRP125, ID4, and c-Kit were found to be upregulated on day 5 after induction (Fig.1B). Furthermore, the expression levels of the meiotic marker PIWIL2, and the post-meiotic haploid marker ACR, in differentiated hESCs also increased steadily and reached their highest levels at the end of culture (Fig.1B).

Generation of hESC-derived spermatogenic cells. (A) Representative immunofluorescence micrographs show the expression of germ cell markers VASA, the hSSC markers PLZF, CD90, GPR125, and the spermatocyte marker PIWIL1 in hSSLC. Non-differentiated hESCs on culture day 0 and IgG staining served as a control. Scale bar: 100m in the first and middle panel, 20m in the last panel. The micrographs shown in the last column are the zoomed-in of the area indicated in red box (B). The graphs show the expression levels of the early germ cell markers (STELLA, TFAP2C, BLIMP1, SOX17, PIWIL4 and VASA), hSSC markers (PLZF, GFRA, GPR125, ID4, NANOS3 and c-Kit), spermatocyte (PIWIL2) and spermatid marker (ACR) during spermatogenic differentiation of hESCs relative to those on culture day 0. Data are presented as meanSD of 3 experiments. (C) Expression of CD90+ cells (top row), PLZF+ cells in the sorted CD90+ cell population determined by flow cytometry (middle row) and immunofluorescent staining (bottom row). The picture in the right panel showed the zoomed-in area indicated in red. (D) Analysis of DNA content by flow cytometry shows the percentages of hESC-derived haploid spermatid-like cells at the end of spermatogenic differentiation. Non-differentiated hESCs on culture day 0 serves as a negative control, while mature human sperm serves as a positive control. (E) Representative immunofluorescence micrographs show the expression of the haploid germ cell marker Acrosin in human sperm cells (top row), Acrosin and TNP1 in hESC-derived spermatid-like cells. (F) Representative micrographs show a single copy of chromosome 13, 21, X and Y chromosome in hESC-derived spermatid-like cells determined by fluorescent in situ hybridization (FISH). Non-differentiated hESCs, which served as a control, had two copies of these chromosomes.

Next, we determined the percentage of hESC-derived hSSLCs after spermatogenic differentiation using the cell surface marker CD90, which has previously been used to identify SSLC in human and mouse models20,36. The results show that 66% of differentiated hESCs expressed CD90 (Fig.1C; top row). We also found that more than 90% of the CD90+hSSLCs also co-expressed PLZF, another widely used SSLC marker, as determined by flow-cytometry and immunofluorescent staining (Fig.1C; middle and bottom rows, respectively). These results confirm that hESC-derived hSSLCs are present in the culture after spermatogenic differentiation and could be identified with a combination of CD90 and PLZF markers.

To determine whether the differentiated hESCs generated any haploid spermatid-like cells, a DNA content analysis was performed. The results show that 6.70.72% of the differentiated hESCs are haploid cells, while no haploid cell was detected in non-differentiated hESCs (Figs.1D and S2). The haploid spermatid-like cells derived from hESCs expressed human spermatid markers, Acrosin and TNP1 (Figs.1E and S2). Furthermore, fluorescent in situ hybridization (FISH) analysis confirmed that our sorted haploid spermatid-like cell derived from hESCs has a single copy of chromosome 13, 21 and sex chromosome compared to two copies of the same chromosomes in non-differentiated hESCs (Fig.1F). These results demonstrated that the spermatogenic differentiation procedure used in this study successfully generated haploid spermatid-like cells from hESCs. The pattern of marker expression during in vitro spermatogenic differentiation closely matched that of human testicular germ cells that underwent in vivo spermatogenesis (Supplementary Fig. S3AG)34. Collectively, these results confirm that we could derive male germ cells from hESCs in vitro using the protocol previously described by Easley et al.21. Next, we determine whether Hippo/YAP is expressed in human testicular germ cells. Analysis of scRNA sequencing data showed YAP expression during spermatogenesis (Supplementary Fig. S3H,I). Like PLZF, the expression level of YAP was higher in undifferentiated human spermatogonia compared to the later stages of human spermatogenic differentiation (Supplementary Fig. S3H,I). This result suggested that YAP might play a crucial role during the early stages of human spermatogenesis and could be involved in the expression of the undifferentiated spermatogonia marker PLZF.

To investigate the role of YAP during human spermatogenesis, the isogenic YAP-depleted hESCs (YAP-KD cells) were established by transducing hESCs (MUSIe002-A cells) with the YAP-targeted CRISPR/Cas9 plasmid33. Western blot analysis showed a 50% reduction in the YAP level in YAP-KD cells compared to their wild-type control (WT cells) (Fig.2A). The YAP-KD cells retained a typical morphology of non-differentiated wild-type hESCs even after being expanded for more than 25 passages (Figs.2B and S4). After spermatogenic differentiation, the YAP-KD cells generated several clusters of cells, which increased in size and number toward the end of the culture (indicated by black arrows in Figs. 2C and S5). These cell clusters were also observed in differentiated WT cells, but their number was less than in the differentiated YAP-KD cells (Figs. 2C and S5).

Spermatogenic differentiation of YAP knockdown hESCs (YAP-KD cells). (A) Western blot shows the level of YAP relative to -ACTIN in YAP-KD cells compared to wild-type hESCs (WT). (B) Representative micrograph showing the morphology of a non-differentiated YAP-KD cell colony. Scale bar: 200m. (C) Representative micrographs show the morphology of differentiated YAP-KD cells during their spermatogenic differentiation. Wild-type hESCs cultured under the same conditions (WT) serve as controls. Scale bar: 200m in micrographs, 50m in zoomed areas. (D) Representative immunofluorescence micrographs show the expression of the SSC marker PLZF in YAP-KD cells during their spermatogenic differentiation compared to WT cells. Scale bar: 200m. (E) Representative dot plots show the percentages of PLZF+ hSSLCs in YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. (F) Graphs show the number of PLZF+ hSSLCs derived from YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. Data are presented as meanSD from 3 experiments. ****P<0.0001. (G) Western blot shows the levels of the YAP and PLZF proteins relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. (H) Graphs show the expression level of YAP protein relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to that of WT cells. Data are presented as meanSD of 3 experiments. *P<0.05, **P<0.01, ****P<0.0001. (I) Graphs show the expression level of PLZF protein relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to that of WT cells. Data are presented as meanSD of 3 experiments. **P<0.01, ***P<0.001. (J) Correlation analysis between YAP protein levels and hESC-derived PLZF+ hSSC numbers during spermatogenic differentiation of WT and YAP-KD cells. The uncropped gels associated with Fig.2A are shown in Supplementary Fig. S8.

YAP protein is highly expressed in spermatogonia residing in testicular germ cells10,11. Similar to WT cells, differentiated YAP-KD cells highly expressed PLZF, a spermatogonia marker. Immunofluorescence shows that the level of PLZF expression in differentiated YAP-KD cells increased steadily throughout the entire culture period and was much higher than that of WT cells (Fig.2D). Consistent with the level of PLZF expression, the percentages of PLZF-expressing hSSLCs (PLZF+ cells) derived from the differentiated YAP-KD cells increased steadily throughout the differentiation period and were 23 times higher than those derived from WT cells throughout the culture period (Figs. 2E and S6). Consistent with these data, when the absolute number of PLZF+ cells was determined by BD Truecount, the YAP-KD cell also generated a higher number of PLZF+ cells than the WT cells, resulting in an approximately threefold increase in the number of PLZF+ cells at the end of the culture (8.61105cells vs 2.19105 cells on culture day 12) (Fig.2F). This result indicated that the depletion of YAP in hESCs enhances their spermatogenic differentiation toward PLZF+ hSSLCs.

Next, we quantified the expression levels of YAP and PLZF proteins at different time points during the spermatogenic differentiation of YAP-KD cells compared to WT cells. The expression level of YAP in WT cells was initially downregulated on culture day 5, bounced back on culture day 10, and continued to increase towards the end of the culture (Fig.2G,H). Interestingly, YAP-KD cells, whose YAP expression was 50% lower than that of WT cells at the beginning of culture, rapidly increased their YAP expression on culture day 5 (Fig.2G,H). Similar to WT cells, the expression level of YAP in YAP-KD cells continued to increase toward the end of the culture. At the end of the culture (day 12), the expression level of YAP in YAP-KD cells is similar to that of WT cells. However, unlike WT cells, down-regulation of YAP during the early phase of spermatogenic differentiation (day 5) was not observed in differentiated YAP-KD cells (Fig.2G,H).

The up-regulation of YAP in WT and YAP-KD cells coincided with the expression of PLZF, whose level also increased steadily towards the end of the culture (Fig.2G,I). However, the differentiated YAP-KD cells expressed PLZF earlier than WT cells (day 5 vs day 10), and their levels of PLZF were significantly higher than those of WT cells throughout the culture period (Fig.2G,I).

Furthermore, the level of YAP protein in both WT and YAP-KD cells was positively correlated with the expression level of PLZF+ hSSLCs generated during their spermatogenic differentiation (R2=0.8577, P=0.001) (Fig.2J). These results suggested that depletion of YAP at the beginning of spermatogenic differentiation leads to higher and earlier expression of PLZF, resulting in a higher number of hESC-derived hSSLCs.

As shown above, early depletion of YAP in hESCs increased the percentage and absolute number of hSSLCs during spermatogenic differentiation. Next, we determine whether hESC-derived hSSLCs express c-Kit, a differentiated spermatogonia marker. We performed a flow cytometric analysis for the expression of c-Kit in hSSLCs derived from WT and YAP-KD cells at different time points during their spermatogenic differentiation. The results showed that both the percentage and the absolute number of c-Kit+differentiated spermatogonial cells were up-regulated in the WT and YAP-KD groups as differentiation progressed towards the end of the culture. In which the percentages and absolute numbers of c-Kit+cells in the YAP-KD groups are significantly higher than those of WT on days 7 and 12 of culture (Fig.3A). This pattern of c-kit expression is similar to that of PLZF expression and therefore confirms that YAP depletion increases the spermatogenic differentiation of hESCs.

Derivation of haploid spermatid-like cells from YAP-KD cells. (A) Representative dot plots show the percentages of c-Kit+ cells in differentiated WT and YAP-KD cells during their spermatogenic differentiation (left). The graph shows the absolute number of c-Kit+ differentiated spermatogonia derived from YAP-KD cells during their spermatogenic differentiation compared to those of WT cells (right). Data are presented as meanSEM from 3 replicates. *P<0.05, **P<0.01 (B) Western blot shows the levels of VASA proteins relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. (C) Graphs show the expression level of VASA protein relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to that of WT cells. Data are presented as meanSD of 3 experiments. *P<0.05, ****P<0.0001. (D) Western blot shows the levels of Acrosin proteins relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. (E) Graphs show the expression level of Acrosin protein relative to -ACTIN in YAP-KD cells during their spermatogenic differentiation compared to that of WT cells. Data are presented as meanSD of 3 experiments. **P<0.01, ****P<0.0001. (F) Analysis of DNA content by flow cytometry shows the percentages of haploid spermatid-like cells derived from YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. (G) The graphs show the percentages of haploid spermatid-like cells derived from YAP-KD cells during their spermatogenic differentiation compared to those of WT cells. Data are presented as meanSD of 3 experiments. *P<0.05. The uncropped gels associated with Fig.3A,C are shown in Supplementary Fig. S9.

We then investigated the effect of YAP depletion during the early stages of spermatogenic differentiation by measuring the expression level of VASA, an early germ cell marker. We found that YAP-KD cells expressed a significantly higher level of VASA protein than WT cells throughout the entire culture period (Fig.3B,C).

Next, we investigated the effect of YAP depletion during the later stages of spermatogenic differentiation by measuring the expression level of Acrosin, a specific marker of haploid spermatids, in differentiated YAP-KD cells. Similar to PLZF, differentiated YAP-KD cells up-regulated their acrosin expression earlier than WT cells (day 5 vs day 10) and their acrosin levels were significantly higher than those of WT cells on day 5 and day 10 of culture (Fig.3D,E). Consistent with the level of acrosin expression, the percentages of haploid spermatid cells in differentiated YAP-KD cells were significantly higher than those of their wild-type counterparts on day 5 (4.40.05% vs. 1.80.68%) and day 10 of culture (6.22.09% vs. 20.10%) (Fig.3F,G). These results suggest that YAP depletion at the beginning of spermatogenic differentiation could accelerate the progression of hESC-derived hSSLCs into haploid spermatid-like cells. However, we found that acrosin+spermatid-like cells could not be maintained in culture, since the level of acrosin in differentiated YAP-KD cells on day 12 of culture was lower than on day 10 (Fig.3D).

Due to the heterozygous nature of the mutated YAP gene in our YAP-KD cells, up-regulation of YAP expression was observed during the later stages of spermatogenic differentiation (days 512) (Fig.2G,H). This pattern of YAP expression in YAP-KD cells suggests that although YAP depletion at the beginning of spermatogenic differentiation appears to promote the germline specification of hESCs, up-regulation of YAP during the later stages of spermatogenic differentiation could be crucial for the expression of germ cell markers and the derivation of both hSSLCs and haploid spermatid-like cells.

To prevent the up-regulation of YAP in YAP-KD cells during the later stages of spermatogenic differentiation, a short hairpin RNA (shRNA) targeting the YAP transcript37 was transfected into YAP-KD cells to establish a YAP double knockdown hESCs (YAP-DKD cells). YAP-DKD cells showed a further reduction in YAP (Fig.4A; Supplementary Table S2) and YAP target genes, CTGF and CYR61, compared to YAP-KD cells (Fig.4B). Like YAP-KD cells, YAP-DKD cells exhibited a typical morphology of non-differentiated wild-type hESCs (Supplementary Fig. S4).

Spermatogenic differentiation of YAP double knockdown hESCs (YAP-DKD cells) (A). Western blot shows the level of YAP relative to -ACTIN in YAP-DKD cells compared to that of YAP-KD cells and wild-type hESCs (WT). ****P<0.0001. (B) Graphs show the expression levels of YAP target genes, CTGF and CYR61, in YAP-DKD cells compared to those of YAP-KD cells and WT. Data are presented as meanSD of 3 experiments. **P<0.01, ***P<0.001. (C) Western blot shows YAP, PLZF, VASA and acrosin protein levels relative to -ACTIN in WT, YAP-KD, and YAP-DKD cells during their spermatogenic differentiation. (D) Graphs show the level of PLZF protein relative to -ACTIN in YAP-DKD cells during their spermatogenic differentiation compared to YAP-KD cells and WT. (E) Graphs show the expression levels of spermatogenic cell markers (VASA, PLZF, GFRA1, GPR125 and ACR) during spermatogenic differentiation of YAP-DKD, compared to those of YAP-KD cells and WT. Data are presented as means SD of 3 experiments. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001. (F) Representative micrographs show the morphology of YAP-DKD cells during their spermatogenic differentiation. White dots indicate areas of dead cells. Scale bar: 200m for the upper panel and 100m for the lower panel. (G) Western blot shows the level of the apoptotic marker cleaved Caspase3 relative to -ACTIN in WT and YAP-DKD cells during their spermatogenic differentiation. The uncropped membranes associated with this figure are provided in Supplementary Fig. S10.

When subjected to spermatogenic differentiation, different expression patterns of YAP and early germ cell markers of each cell type were observed. Unlike YAP-KD cells, the level of YAP expression in YAP-DKD cells was barely detected on day 10 and day 12 of culture (Fig.4C). This result confirmed that the shRNA successfully prevented the up-regulation of YAP in YAP-DKD cells during the later stages of their spermatogenic differentiation. Further analysis found that constitutive depletion of YAP in YAP-DKD inhibited the expression of the early germ cell marker VASA (Fig.4C), the SSC marker PLZF (Fig.4C,D), and the spermatid marker acrosin (Fig.4C). These results suggest that the up-regulation of YAP level during the later stages of spermatogenesis might be critical for the expression of germ cell markers and the derivation of hSSLCs and haploid spermatid-like cells from hESCs.

Consistent with the protein expression study, differentiated YAP-DKD cells expressed lower transcript levels of germ cell marker (VASA), hSSC marker genes (PLZF, GFRA1, and GPR125), and meiotic germ cell markers (ACR) compared to WT (Fig.4E). In contrast, YAP-KD cells, whose YAP was depleted only at the beginning of their spermatogenic differentiation, expressed higher levels of hSSC markers, PLZF, GPR125, and GFRA1 compared to WT cells (Fig.4E). Taken together, these results suggest that depleting YAP at the beginning of spermatogenic differentiation promotes germline specification of hESCs, and increasing the level of YAP during the later stages of spermatogenic differentiation is critical for the derivation and survival of hESC-derived hSSLCs and haploid spermatid-like cells.

Furthermore, we also found that the differentiated YAP-DKD cells contained several large areas of dead cells (dotted areas in Figs. 4F and S5). Consistent with this, western blots also showed a high level of cleaved-caspase 3 in YAP-DKD cells at the later stages of spermatogenic differentiation (D5-12) compared to D0 (Fig.4G; YAP-DKD), while the expression of cleaved-caspase 3 in WT did not show a sharp contrast compared to its control D0. These results confirm the higher level of apoptosis in differentiated YAP-DKD cells during the later stages of spermatogenesis.

To confirm that an initial depletion followed by an up-regulation of YAP promotes spermatogenic differentiation of hESCs, we modulated the expression level of YAP at specific time points during spermatogenic differentiation of wild-type hESCs using two small molecules. Dobutamine hydrochloride (DH), which has previously been shown to promote YAP phosphorylation and suppress YAP activity38,39, was used to inhibit YAP expression, while lysophosphatidic acid (LPA) activated YAP expression in wild-type hESCs. DH-treated hESCs showed a 50% reduction in YAP level compared to untreated hESCs (Supplementary Fig. S7B,C), and also expressed lower levels of YAP target genes, CTGF and CCND1, compared to untreated hESCs (Supplementary Fig. S7D). Our results demonstrate that DH treatment successfully inhibits the expression of YAP and its target genes in wild-type hESCs. Although LPA-treated hESCs increased CTGF, CCND1, and CYR61 expression levels compared to untreated hESCs (Supplementary Fig. S7D), their YAP protein level did not increase compared to the untreated cells (Supplementary Fig. S7B,C). These results suggest that although LPA did not increase YAP level in hESCs, it might enhance YAP activity resulting in higher levels of YAP target genes.

We then supplemented DH and LPA at various time points during spermatogenic differentiation of wild-type hESCs (Fig.5A). Consistent with the results generated from YAP-KD cells, the highest number of hESC-derived hSSLCs was observed under the condition that DH was added to inhibit YAP expression during the first 5days of spermatogenic differentiation before being removed for the rest of the culture period (Fig.5B, D12: DH-ET in Group 2). On the contrary, hESCs treated with LPA to increase YAP activity during the first 5days of their spermatogenic differentiation generated a lower number of hSSLCs than both D12: DH-ET and untreated cells (Fig.5B, D12: LPA-ET in Group 2).

Modulation of YAP expression during the various stages of spermatogenic cell differentiation of wild-type hESCs by using DH and LPA. (A) Diagrams show the timeline of DH and LPA supplementation during spermatogenic differentiation of wild-type hESCs. (B) Graphs show the total number of differentiated hESCs and the number of PLZF+ hSSCs derived from various conditions described in (A) at the end of the culture. Data are presented as meanSD of 3 experiments. **P<0.01, ***P<0.001, ****P<0.0001 and ### P<0.001 indicated the statistically significant of spermatogonia number from DH treated cells between each group. (C) Representative immunofluorescence micrographs show the level of PLZF expression in DH-ET and LPA-ET cells compared to that of their untreated counterpart. Scale bar: 200m. (D) Western blot shows the expression of YAP, PLZF, Acrosin, and -ACTIN proteins in DH-ET cells during their spermatogenic differentiation compared to those of DH-AT cells, untreated control cells, and non-differentiated hESCs. Red numbers indicate the expression level of each sample relative to -ACTIN. (E) Analysis of DNA content by flow cytometry shows the percentage of haploid spermatid-like cells derived from DH-ET cells at the end of their spermatogenic differentiation compared to that of LPA-ET and untreated cells. (F) The graph shows the percentage of haploid spermatid-like cells derived from DH-ET cells at the end of their spermatogenic differentiation compared to that of LPA-ET and untreated cells. Data are presented as meanSD of 3 experiments. **P<0.01, ***P<0.001. The uncropped membranes associated with Fig.5D are provided in Supplementary Fig. S11.

Unlike YAP suppression during the early stage of spermatogenic differentiation (days 05), suppressing YAP during the middle phase (days 510) of spermatogenic differentiation reduced the number of hESC-derived hSSLCs compared to untreated hESCs (Fig.5B, D12: DH-MT in Group 3). Moreover, suppressing YAP during the middle stage (DH-MT in Group 3) or throughout the spermatogenic differentiation (DH-AT in Group 1) also reduced the total number of differentiated hESCs in culture (Fig.5B), which could result in the decrease in the number of hESC-derived hSSLCs at the end of culture. This is consistent with our previous result with YAP-DKD cells showing that continued suppression of YAP during the later stages of spermatogenic differentiation increased the apoptotic level of differentiated hESCs (Fig.4F,G).

Furthermore, our immunofluorescence study shows that the level of PLZF expression in DH-ET cells on day 12 was higher than that of the untreated control, while the level of PLZF expression in LPA-ET cells was lower than that of the untreated control (Fig.5C). Western blot analysis further confirmed that DH-ET cells expressed higher levels of PLZF than the untreated control at the end of the culture (Fig.5D), while LPA-ET cells expressed lower levels of PLZF than the untreated control at the end of the culture (Supplementary Fig. S7E,F). Unlike suppression of YAP during the early stage of spermatogenic differentiation (DH-ET), suppression of YAP throughout spermatogenic differentiation (DH-AT) decreased the expression level of PLZF compared to the untreated control (Fig.5D). Although DH-ET treatment produces a higher number of haploid germ cells than the control, the level of Acrosin expression in the DH-ET group is lower than that of the control (Fig.5D). Because the level of Acrosin is associated with spermatid maturity, it is possible that while DH-ET treatment increases the number of hESC-derived spermatids, these spermatids may be less mature than those derived from the untreated control.

We further investigated haploid germ cell production using cell cycle analysis to observe DNA content of the cells. The DH-ET cells also generate significantly higher percentages of haploid spermatid-like cells at the end of culture than their untreated counterpart (14.162.61% vs 8.302.86%) (Fig.5E,F). These results support our hypothesis that inhibition of YAP activity during the early stage of spermatogenic differentiation facilitates the derivation of hSSLCs and spermatid-like cells from hESCs. In contrast, inhibition of YAP activity during the later stages of spermatogenic differentiation reduced the number of hESC-derived hSSLCs and spermatid-like cells, possibly by increasing the apoptotic rate of hSSLCs and preventing their maturation into haploid spermatid-like cells.

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The dynamic expression of YAP is essential for the development of male germ cells derived from human embryonic stem cells - Nature.com

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S.BIOMEDICS Dopamine Cell Therapy for Parkinsons Disease with TED-A9 Shows Promising Results at 12 months in Phase I/IIa Clinical Trial

S.BIOMEDICS(KOSDAQ: 304360) announced positive one-year assessment data from the first 3 participants (first low dose cohort) of Phase I/IIa clinical trial for Parkinsons disease with TED-A9, hESC (human embryonic stem cell)-derived midbrain dopaminergic progenitors. The data demonstrates that 3 participants showed safety and efficacy of TED-A9 in a one-year follow-up.

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Dong-Wook Kim, Professor at Yonsei University College of Medicine and CTO of S.BIOMEDICS (Photo: S.BIOMEDICS)

The clinical trial was conducted at Severance Hospital of Yonsei University in South Korea led by Prof. Jin-Woo Chang, a neurosurgeon and Prof. Phil Hyu Lee, a neurologist.

For the 3 participants who received initial low dose (3.15 million cells), MRI and CT scans after one year revealed no adverse effects related to the cell transplantation or surgery.

Moreover, the MDS-UPDRS Part III (off) evaluation, which objectively measures motor functions, showed a mean decrease of 12.7 points, from baseline 61.7 to 49.0 one-year post treatment. The MDS-UPDRS Part III (off) evaluation indicated significant improvement in their motor abilities. Improvement was also observed in symptoms such as wearing off and freezing of gait.

DAT brain imaging (FP-CIT-PET), conducted one-year post transplantation, revealed an increase in dopamine transporters (DAT), suggesting the potential engraftment of dopamine neurons, which correlated with improvements in the patients' Parkinson's symptoms.

Although this clinical evaluation only targets the first 3 low-dose patients, but not all 12 subjects, no adverse issues related to transplant surgery or cell safety were observed in one-year post transplantation. Importantly, the clinical results demonstrated very promising efficacy, said Prof. Dong-Wook Kim at Yonsei University College of Medicine and CTO of S.BIOMEDICS. The results are believed to align closely with the findings from our preclinical in vitro and in vivo studies. We are excited that TED-A9 could be a fundamental treatment that directly replaces dopaminergic neurons lost in patients with Parkinson's disease.

Assessment data of further participants (first high dose cohort) will be announced in September or October of 2024 after a one-year follow-up.

About TED-A9 and Phase I/IIa clinical trial

TED-A9 is an investigational cell therapy designed to replace ventral midbrain-specific dopaminergic cells lost in patients with Parkinson's disease. These ventral midbrain-specific dopaminergic cells are derived from hESCs by exclusively utilizing small molecules only. TED-A9 represents a significant advancement in the field, offering highly purified dopamine cells derived from hESCs. Through surgical procedure, these hESC-derived dopaminergic progenitor (precursor) cells (TED-A9) are transplanted to three segments of the putamen; the anterior, middle, and posterior sections, with three tracks per each putamen. Bilateral putamina received cell transplantation in a single surgical procedure, with cells injected at three points within each track. After transplantation, the expectation is that the transplanted dopaminergic progenitor cells will mature into dopaminergic neurons which will supply the dopamine that Parkinsons patients are lacking, restoring the motor function of patients.

The Phase I/IIa clinical trial is conducted on 12 participants who have been diagnosed with Parkinson's disease for more than 5 years and exhibited motor complications such as wearing-off, freezing of gait or dyskinesia. The participants age was between 50 and 75 years old. TED-A9 were administered to 6 participants in the low-dose group (3.15 million cells) and to another 6 participants in the high-dose group (6.30 million cells).

An initial cohort of three patients was enrolled at a low dose to assess initial safety, including dose-limiting toxicity (DLT) evaluation, over a period of up to 3 months post-transplantation. There were no safety concerns within this timeframe. Thus, an additional 3 patients were enrolled at a high dose for evaluation over another 3-month period post-transplantation. As no safety issues arose during this extended period, the clinical trial continued by adding three further patients to each of the low-dose and high-dose groups, totaling 12 patients. The final participant was administered with TED-A9 on February 2024.

The primary objective of the Phase I/IIa trial is to assess the safety and exploratory efficacy of TED-A9 transplantation over two years post-transplant. Safety will be monitored for additional 3 years, making it a total 5 years.

About S.BIOMEDICS

Established in 2005, S.BIOMEDICS is at the forefront of stem cell therapy, focusing on regenerative medicine powered by data-driven biology. Based on two core platform technologies, S.BIOMEDICS currently develops seven cell therapy programs, targeting non-curable diseases. Its leading programs are under clinical stage accelerating the journey of cell medicine as shown below:

For more information about S.BIOMEDICS, visit http://www.sbiomedics.com. S.BIOMEDICS is listed on the Korea Exchange (KOSDAQ: 304360) and is also the founder and controller of S.THEPHARM (www.sthepharm.com), a corporation specializing in anti-aging products such as HA-Filler. More Information about the Phase 1/2a clinical trial for Parkinsons disease is available at ClinicalTrials.gov (NCT05887466).

S.BIOMEDICS Jong-Wan Kim jwkim@sbiomedics.com

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