Global Stem Cell and Platelet Rich Plasma (PRP) Alopecia Therapies Market Recent Study Including Business Growth, Development Factors and Growth…

In its recently added report by MRInsights.biz with the title Global Stem Cell and Platelet Rich Plasma (PRP) Alopecia Therapies Market has provided a comprehensive analysis of the market structure which includes unique insights about the market for the given period. The report covers the competitive landscape and the conspicuous market players anticipated to lead the global Stem Cell and Platelet Rich Plasma (PRP) Alopecia Therapies market for the forecast period, 2019-2024. One of the main targets of this report is to classify the various dynamics of the market. The forenamed market is greatly transforming because of the moves of the key players and brands including developments, product launches, joint ventures, mergers and acquisitions that in turn change the view of the global face of the industry.

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The drivers and restraints are intrinsic factors while opportunities and challenges are extrinsic factors of the market. The research report is based on the integration, analysis, and interpretation of information gathered regarding the target market from various sources. The report analysts have assessed information and data information and data acquired using a mix of primary and secondary research efforts. The global economic conditions and other economic indicators and factors are analyzed to look at their respective impact on the global Stem Cell and Platelet Rich Plasma (PRP) Alopecia Therapiesmarket historically, as well as the current impact that will help to make informed forecasts about the scenarios in the future.

Market Insights of Competitive Landscape:

In the competition landscape section of the industry, our analysts provide an insight into the financial statements of all the major players along with its key developments product benchmarking and SWOT analysis. Company profiles cover the product offerings, key financial information, recent developments, SWOT analysis, and strategies employed by the major market players. Additionally, the market share of major players, along with the new projects and strategies adopted by players in the past five years (2014-2020) are also included.

List of some major players from a wide list of coverage used under the bottom-up approach is: Orange County Hair Restoration Center, Colorado Surgical Center & Hair Institute, Evolution Hair Loss Institute, Hair Sciences Center of Colorado, Hair Transplant Institute of Miami, Anderson Center for Hair, Virginia Surgical Center, Savola Aesthetic Dermatology Center,

The research provides information on opportunities available in the market. In terms of region, the market covers:

North America (United States, Canada and Mexico)

Europe (Germany, France, UK, Russia and Italy)

Asia-Pacific (China, Japan, Korea, India and Southeast Asia)

South America (Brazil, Argentina, Colombia)

Middle East and Africa (Saudi Arabia, UAE, Egypt, Nigeria and South Africa)

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Moreover, the report covers the ongoing as well as forecast trends likely to fuel the business graph of the global Stem Cell and Platelet Rich Plasma (PRP) Alopecia Therapiesmarket. Further, the report introduces a new project SWOT analysis, investment feasibility analysis, and investment return analysis. An overview of each market segment such as product type, application, end-users, and region are offered in the report. A comparative study between conventional and emerging technologies and the importance of technical developments in this market has been offered.

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Global Stem Cell and Platelet Rich Plasma (PRP) Alopecia Therapies Market Recent Study Including Business Growth, Development Factors and Growth...

Little Tissue, Big Mission: Beating Heart Tissues to Ride Aboard The ISS – Newswise

Newswise Launching no earlier than March 6 at 11:50 PM EST, the Johns Hopkins University will send heart muscle tissues, contained in a specially-designed tissue chip the size of a small cellphone, up to the microgravity environment of the International Space Station (ISS) for one month of observation.

The project, led by Deok-Ho Kim, an Associate Professor of Biomedical Engineering and Medicine at The Johns Hopkins University and the projects principal investigator, will hopefully shed light on the aging process and adult heart health, and facilitate the development of treatments for heart muscle diseases.

Scientists already know that humans exposed to space experience changes similar to accelerated aging, so we hope the results can help us better understand and someday counteract the aging process, says Kim.

The researchers also hope the study will demystify why astronauts in space have reduced heart function and are more prone to serious irregular heartbeat; these results could help protect astronauts hearts on long missions in the future, as well as provide information on how to combat heart disease.

Kim and his team used human induced pluripotent stem cells to grow cardiomyocytes, or heart muscle cells, in a bioengineered, miniaturized tissue chip that mimics the function of the adult human heart. While other researchers have studied stem cell-derived heart muscle cells in space before, these studies relied on cells cultured on 2D surfaces, or flat planes, that arent representative of how cells exist and behave in the body, and are therefore underdeveloped compared to their counterparts in adult humans.

The teams tissue platform gives the advantage of the cells residing in a 3D environment, which will allow for better imitation of how cell signals and actions develop as they would in the human body. This 3D environment is possible thanks to a new scaffold biomaterial, or support structure which holds the tissues together, that accelerates development of the heart muscle cells within. This will allow the scientists to collect data useful for understanding the adult human body. Scientists could someday use this data and platform to develop new drugs, among many other applications.

Using a motion sensor magnet setup, the team will receive real-time measurements of how the tissues on the ISS beat. After about one month in space, the tissues will return to Earth and will be analyzed for any differences in gene expression and contraction caused by the extended stay in microgravity. Some of these tissues will be cultured for an additional week on Earth for the researchers to examine any recovery effects. The team will also have identical heart tissues on Earth at the University of Washington to serve as controls.

We hope that this project will give us meaningful data that we can use to understand the hearts structure and how it functions, so that we can improve the health of both astronauts and those down here on Earth, says Kim.

"The entire team is excited to see the results we get from this experiment. If successful, we will embark on the second phase of the study where tissues will be sent up to the ISS once again in two years, but this time, we will be able to test a variety of drugs to see which ones will best ameliorate the potentially harmful effects of microgravity on cardiac function," says Jonathan Tsui, a postdoctoral fellow in the Department of Biomedical Engineering at The Johns Hopkins University and a member of Kims lab.

This project is funded by the National Center for Advancing Translational Sciences (NCATS) and the National Institute of Biomedical Imaging and Bioengineering (NIBIB) as part of the Tissue Chips in Space initiative in collaboration with the ISS U.S. National Laboratory.

Collaborators on this project include Eun Hyun Ahn of The Johns Hopkins University; Nathan Sniadecki and Alec Smith of The University of Washington; Peter Lee of Ohio State University; and Stefanie Countryman of Bioserve Space Technologies at the University of Colorado Boulder. For space flight the team has worked with BioServe Space Technologies to translate the ground platform into a space flight certified system.

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Little Tissue, Big Mission: Beating Heart Tissues to Ride Aboard The ISS - Newswise

With Over 280 Therapies Under Evaluation, the Stem Cell Therapy Market is Estimated to be Worth USD 8.5 Billion by 2030, Claims Roots Analysis – P&T…

The success of approved stem cell therapies has caused a surge in interest of biopharma developers in this field; many innovator companies are currently progressing proprietary leads across different phases of clinical development, with cautious optimism

LONDON, March 4, 2020 /PRNewswire/ -- Roots Analysishas announced the addition of "Global Stem Cells Market: Focus on Clinical Therapies, 20202030 (Based on Source (Allogeneic, Autologous); Origin (Adult, Embryonic); Type (Hematopoietic, Mesenchymal, Progenitor); Lineage (Amniotic Fluid, Adipose Tissue, Bone Marrow, Cardiosphere, Chondrocytes, Corneal Tissue, Cord Blood, Dental Pulp, Neural Tissue Placenta, Peripheral Blood, Stromal Cells); and Potency (Multipotent, Pluripotent))" report to its list of offerings.

There is a growing body of evidence supporting the vast applicability and superiority of treatment outcomes of stem cell therapies, compared to conventional treatment options. In fact, the unmet needs within this domain have spurred the establishment of many start-ups in recent years.

To order this 500+ page report, which features 185+ figures and 220+ tables, please visit this link

Key Market Insights

Over 280 stem cell therapies are under development, most of which are allogeneic products

More than 50% of the pipeline candidates are in the mid to late phase trials (phase II and above), and allogenic therapies (majority of which are derived from the bone marrow) make up 65% of the pipeline.

70% of pipeline candidates are based on mesenchymal stem cells

It is worth highlighting that the abovementioned therapies are designed to treat musculoskeletal (22%), neurological (21%) and cardiovascular (15%) disorders. On the other hand, hematopoietic stem cell-based products are mostly being evaluated for the treatment of oncological disorders, primarily hematological malignancies.

Close to 85% stem cell therapy developers are based in North America and Asia-Pacific regions

Within these regions, the US, China, South Korea and Japan, have emerged as key R&D hubs for stem cell therapies. It is worth noting that majority of the initiatives in this domain are driven by small / mid-sized companies

Over 1,500 grants were awarded for stem cell research, since 2015

More than 45% of the total amount was awarded under the R01 mechanism (which supports research projects). The NCI, NHLBI, NICHD, NIDDK, NIGMS and OD emerged as key organizations that have offered financial support for time periods exceeding 25 years as well.

Outsourcing has become indispensable to R&D and manufacturing activity in this domain

Presently, more than 80 industry / non-industry players, based in different regions across the globe, claim to provide contract development and manufacturing services to cater to the unmet needs of therapy developers. Examples include (in alphabetical order) Bio Elpida, Cell and Gene Therapy Catapult, Cell Tech Pharmed, GenCure, KBI Biopharma, Lonza, MEDINET, Nikon CeLL innovation, Roslin Cell Therapies, WuXi Advanced Therapies and YposKesi.

North America and Asia-Pacific markets are anticipated to capture over 80% share by 2030

The stem cell therapies market is anticipated to witness an annualized growth rate of over 30% during the next decade. Interestingly, the market in China / broader Asia-Pacific region is anticipated to grow at a relatively faster rate.

To request a sample copy / brochure of this report, please visit this link

Key Questions Answered

The USD 8.5 billion (by 2030) financial opportunity within the stem cell therapies market has been analyzed across the following segments:

The report features inputs from eminent industry stakeholders, according to whom stem cell therapies are currently considered to be a promising alternatives for the treatment of a myriad of disease indications, with the potential to overcome challenges associated with conventional treatment options. The report includes detailed transcripts of discussions held with the following experts:

The research covers brief profiles of several companies (including those listed below); each profile features an overview of the company, financial information (if available), stem cell therapy portfolio and an informed future outlook.

For additional details, please visit

https://www.rootsanalysis.com/reports/view_document/stem-cells-market/296.htmlor email sales@rootsanalysis.com

You may also be interested in the following titles:

Contact:Gaurav Chaudhary+1(415)800-3415+44(122)391-1091Gaurav.Chaudhary@rootsanalysis.com

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With Over 280 Therapies Under Evaluation, the Stem Cell Therapy Market is Estimated to be Worth USD 8.5 Billion by 2030, Claims Roots Analysis - P&T...

Biochemical and structural cues of 3D-printed matrix synergistically direct MSC differentiation for functional sweat gland regeneration – Science…

Abstract

Mesenchymal stem cells (MSCs) encapsulation by three-dimensionally (3D) printed matrices were believed to provide a biomimetic microenvironment to drive differentiation into tissue-specific progeny, which made them a great therapeutic potential for regenerative medicine. Despite this potential, the underlying mechanisms of controlling cell fate in 3D microenvironments remained relatively unexplored. Here, we bioprinted a sweat gland (SG)like matrix to direct the conversion of MSC into functional SGs and facilitated SGs recovery in mice. By extracellular matrix differential protein expression analysis, we identified that CTHRC1 was a critical biochemical regulator for SG specification. Our findings showed that Hmox1 could respond to the 3D structure activation and also be involved in MSC differentiation. Using inhibition and activation assay, CTHRC1 and Hmox1 synergistically boosted SG gene expression profile. Together, these findings indicated that biochemical and structural cues served as two critical impacts of 3D-printed matrix on MSC fate decision into the glandular lineage and functional SG recovery.

Mesenchymal stem cells (MSCs) hold great promise for therapeutic tissue engineering and regenerative medicine, largely because of their capacity for self-renewal and multipotent properties (1). However, their uncertain fate has a major impact on their envisioned therapeutic use. Cell fate regulation requires specific transcription programs in response to environmental cues (2, 3). Once stem cells are removed from their microenvironment, their response to environmental cues, phenotype, and functionality could often be altered (4, 5). In contrast to growing information concerning transcriptional regulation, guidance from the extracellular matrix (ECM) governing MSC identity and fate determination is not well understood. It remains an active area of investigation and may provide previously unidentified avenues for MSC-based therapy.

Over the past decade, engineering three-dimensional (3D) ECM to direct MSC differentiation has demonstrated great potential of MSCs in regenerative medicine (6). 3D ECM has been found to be useful in providing both biochemical and biophysical cues and to stabilize newly formed tissues (7). Culturing cells in 3D ECM radically alters the interfacial interactions with the ECM as compared with 2D ECM, where cells are flattened and may lose their differentiated phenotype (8). However, one limitation of 3D materials as compared to 2D approaches was the lack of spatial control over chemistry with 3D materials. One possible solution to this limitation is 3D bioprinting, which could be used to design the custom scaffolds and tissues (9).

In contrast to traditional engineering techniques, 3D cell printing technology is especially advantageous because it can integrate multiple biophysical and biochemical cues spatially for cellular regulation and ensure complex structures with precise control and high reproducibility. In particular, for our final goal of clinical practice, extrusion-based bioprinting may be more appropriate for translational application. In addition, as a widely used bioink for extrusion bioprinting, alginate-based hydrogel could maintain stemness of MSC due to the bioinert property and improve biological activity and printability by combining gelatin (10).

Sweat glands (SGs) play a vital role in thermal regulation, and absent or malfunctioning SGs in a hot environment can lead to hyperthermia, stroke, and even death in mammals (11, 12). Each SG is a single tube consisting of a functionally distinctive duct and secretory portions. It has low regenerative potential in response to deep dermal injury, which poses a challenge for restitution of lost cells after wound (13). A major obstacle in SG regeneration, similar to the regeneration of most other glandular tissues, is the paucity of viable cells capable of regenerating multiple tissue phenotypes (12). Several reports have described SG regeneration in vitro; however, dynamic morphogenesis was not identified nor was the overall function of the formed tissues explored (1416). Recent advances in bioprinting and tissue engineering led to the complexities in the matrix design and fabrication with appropriate biochemical cues and biophysical guidance for SG regeneration (1719).

Here, we adopted 3D bioprinting technique to mimic the regenerative microenvironment that directed the specific SG differentiation of MSCs and ultimately guided the formation and function of glandular tissue. We used alginate/gelatin hydrogel as bioinks in this present study due to its good cytocompatibility, printability, and structural maintenance in long-time culture. Although the profound effects of ECM on cell differentiation was well recognized, the importance of biochemical and structural cues of 3D-printed matrix that determined the cell fate of MSCs remained unknown; thus, the present study demonstrated the role of 3D-printed matrix cues on cellular behavior and tissue morphogenesis and might help in developing strategies for MSC-based tissue regeneration or directing stem cell lineage specification by 3D bioprinting.

The procedure for printing the 3D MSC-loaded construct incorporating a specific SG ECM (mouse plantar region dermis, PD) was shown schematically in Fig. 1A. A 3D cellular construct with cross section 30 mm 30 mm and height of 3 mm was fabricated by using the optimized process parameter (20). The 3D construct demonstrated a macroporous grid structure with hydrogel fibers evenly distributed according to the computer design. Both the width of the fibers and the gap between the fibers were homogeneous, and MSCs were embedded uniformly in the hydrogel matrix fibers to result in a specific 3D microenvironment. (Fig. 1B).

(A) Schematic description of the approach. (B) Full view of the cellular construct and representative microscopic and fluorescent images and the quantitative parameters of 3D-printed construct (scale bars, 200 m). Photo credit: Bin Yao, Wound Healing and Cell Biology Laboratory, Institute of Basic Medical Sciences, General Hospital of PLA. (C) Representative microscopy images of cell aggregates and tissue morphology at 3, 7, and 14 days of culture (scale bars, 50 m) and scanning electron microscopy (sem) images of 3D structure (scale bars, 20 m). PD+/PD, 3D construct with and without PD. (D) DNA contents, collagen, and GAGs of native tissue and PD. (E) Proliferating cells were detected through Ki67 stain at 3, 7, and 14 days of culture. (F) Live/dead assay show cell viability at days 3, 7, and 14. *P < 0.05.

During the maintenance of constructs for stem cell expansion, MSCs proliferated to form aggregates of cells but self-assembled to an SG-like structure only with PD administration (Fig. 1C and fig. S1, A to C). We carried out DNA quantification assay to evaluate the cellular content in PD and found the cellular matrix with up to 90% reduction, only 3.4 0.7 ng of DNA per milligram tissue remaining in the ECM. We also estimated the proportions of collagen and glycosaminoglycans (GAGs) in ECM through hydroxyproline assay and dimethylmethylene blue assay, the collagen contents could increase to 112.6 11.3%, and GAGs were well retained to 81 9.6% (Fig. 1D). Encapsulated cells were viable, with negligible cell death apparent during extrusion and ink gelation by ionic cross-linking, persisting through extended culture in excess of 14 days. The fluorescence intensity of Ki67 of MSCs cultured in 2D condition decreased from days 3 (152.7 13.4) to 14 (29.4 12.9), while maintaining higher intensity of MSCs in 3D construct (such as 211.8 19.4 of PD+3D group and 209.1 22.1 of PD3D group at day 14). And the cell viability in 3D construct was found to be sufficiently high (>80%) when examined on days 3, 7, and 14. The phenomenon of cell aggregate formation and increased cell proliferation implied the excellent cell compatibility of the hydrogel-based construct and promotion of tissue development of 3D architectural guides, which did not depend on the presence or absence of PD (Fig. 1, E and F).

The capability of 3D-printed construct with PD directing MSC to SGs in vitro was investigated. The 3D construct was dissolved, and cells were isolated at days 3, 7, and 14 for transcriptional analysis. Expression of the SG markers K8 and K18 was higher from the 3D construct with (3D/PD+) than without PD (3D/PD); K8 and K18 expression in the 3D/PD construct was similar to with control that MSCs cultured in 2D condition, which implied the key role of PD in SG specification. As compared with the 2D culture condition, 3D administration (PD+) up-regulated SG markers, which indicated that the 3D structure synergistically boosted the MSC differentiation (Fig. 2A).

(A) Transcriptional expression of K8, K18, Fxyd2, Aqp5, and ATP1a1 in 3D-bioprinted cells with and without PD in days 3, 7, and 14 culture by quantitative real-time polymerase chain reaction (qRT-PCR). Data are means SEM. (B) Comparison of SG-specific markers K8 and K18 in 3D-bioprinted cells with and without PD (K8 and K18, red; DAPI, blue; scale bars, 50 m). (C and D) Comparison of SG secretion-related markers ATP1a1 (C) and Ca2+ (D) in 3D-bioprinted cells with and without PD [ATP1a1 and Ca2+, red; 4,6-diamidino-2-phenylindole (DAPI), blue; scale bars, 50 m].

In addition, we tested secretion-related genes to evaluate the function of induced SG cells (iSGCs). Although levels of the ion channel factors of Fxyd2 and ATP1a1 were increased notably in 2D culture with PD and ATP1a1 up-regulated in the 3D/PD construct, all the secretory genes of Fxyd2, ATP1a1, and water transporter Aqp5 showed the highest expression level in the 3D/PD+ construct (Fig. 2A). Considering the remarkable impact, further analysis focused on 3D constructs.

Immunofluorescence staining confirmed the progression of MSC differentiation. At day 7, cells in the 3D/PD+ construct began to express K8 and K18, which was increased at day 14, whereas cells in the 3D/PD construct did not express K8 and K18 all the time (Fig. 2B and fig. S2A). However, the expression of ATP1a1 (ATPase Na+/K+ transporting subunit alpha 1) and free Ca2+ concentration did not differ between cells in the 3D/PD+ and 3D/PD constructs (Fig. 2, C and D). By placing MSCs in such a 3D environment, secretion might be stimulated by rapid cell aggregation without the need for SG lineage differentiation. Cell aggregationimproved secretion might be due to the benefit of cell-cell contact (fig. S2B) (21, 22).

To map the cell fate changes during the differentiation between MSCs and SG cells, we monitored the mRNA levels of epithelial markers such as E-cadherin, occludin, Id2, and Mgat3 and mesenchymal markers N-cadherin, vimentin, Twist1, and Zeb2. The cells transitioned from a mesenchymal status to a typical epithelial-like status accompanied by mesenchymal-epithelial transition (MET), then epithelial-mesenchymal transition (EMT) occurred during the further differentiation of epithelial lineages to SG cells (fig. S3A). In addition, MET-related genes were dynamically regulated during the SG differentiation of MSCs. For example, the mesenchymal markers N-cadherin and vimentin were down-regulated from days 1 to 7, which suggested cells losing their mesenchymal phenotype, then were gradually up-regulated from days 7 to 10 in their response to the SG phenotype and decreased at day 14. The epithelial markers E-cadherin and occludin showed an opposite expression pattern: up-regulated from days 1 to 5, then down-regulated from days 7 to 10 and up-regulated again at day 14. The mesenchymal transcriptional factors ZEB2 and Twist1 and epithelial transcriptional factors Id2 and Mgat3 were also dynamically regulated.

We further analyzed the expression of these genes at the protein level by immunofluorescence staining (figs. S3B and S4). N-cadherin was down-regulated from days 3 to 7 and reestablished at day 14, whereas E-cadherin level was increased from days 3 to 7 and down-regulated at day 14. Together, these results indicated that a sequential and dynamic MET-EMT process underlie the differentiation of MSCs to an SG phenotype, perhaps driving differentiation more efficiently (23). However, the occurrence of the MET-EMT process did not depend on the presence of PD. Thus, a 3D structural factor might also participate in the MSC-specific differentiation (fig. S3C).

To investigate the underlying mechanism of biochemical cues in lineage-specific cell fate, we used quantitative proteomics analysis to screen the ECM factors differentially expressed between PD and dorsal region dermis (DD) because mice had eccrine SGs exclusively present in the pads of their paws, and the trunk skin lacks SGs. In total, quantitative proteomics analyses showed higher expression levels of 291 proteins in PD than DD. Overall, 66 were ECM factors: 23 were significantly up-regulated (>2-fold change in expression). We initially determined the level of proteins with the most significant difference after removing keratins and fibrin: collagen triple helix repeat containing 1 (CTHRC1) and thrombospondin 1 (TSP1) (fig. S5). Western blotting was performed to further confirm the expression level of CTHRC1 and TSP1, and we then confirmed that immunofluorescence staining at different developmental stages in mice revealed increased expression of CTHRC1 in PD with SG development but only slight expression in DD at postnatal day 28, while TSP1 was continuously expressed in DD and PD during development (Fig. 3, A to C). Therefore, TSP1 was required for the lineage-specific function during the differentiation in mice but was not dispensable for SG development.

(A and B) Differential expression of CTHRC1 and TSP1in PD and back dermis (DD) ECM of mice by proteomics analysis (A) and Western blotting (B). (C) CTHRC1 and TSP1 expression in back and plantar skin of mice at different developmental times. (Cthrc1/TSP1, red; DAPI, blue; scale bars, 50 m).

According to previous results of the changes of SG markers, 3D structure and PD were both critical to SG fate. Then, we focused on elucidating the mechanisms that underlie the significant differences observed in 2D and 3D conditions with or without PD treatment. To this end, we performed transcriptomics analysis of MSCs, MSCs treated with PD, MSCs cultured in 3D construct, and MSC cultured in 3D construct with PD after 3-day treatment. We noted that the expression profiles of MSCs treated with 3D, PD, or 3D/PD were distinct from the profiles of MSCs (Fig. 4A). Through Gene Ontology (GO) enrichment analysis of differentially expressed genes, it was shown that PD treatment in 2D condition induced up-regulation of ECM and inflammatory response term, and the top GO term for MSCs in 3D construct was ECM organization and extracellular structure organization. However, for the MSCs with 3D/PD treatment, we found very significant overrepresentation of GO term related to branching morphogenesis of an epithelial tube and morphogenesis of a branching structure, which suggested that 3D structure cues and biochemical cues synergistically initiate the branching of gland lineage (fig S6). Heat maps of differentially expressed ECM organization, cell division, gland morphogenesis, and branch morphogenesis-associated genes were shown in fig. S7. To find the specific genes response to 3D structure cues facilitating MSC reprogramming, we analyzed the differentially expressed genes of four groups of cells (Fig. 4B). The expression of Vwa1, Vsig1, and Hmox1 were only up-regulated with 3D structure stimulation, especially the expression of Hmox1 showed a most significant increase and even showed a higher expression addition with PD, which implied that Hmox1 might be the transcriptional driver of MSC differentiation response to 3D structure cues. Differential expression of several genes was confirmed by quantitative polymerase chain reaction (qPCR): Mmp9, Ptges, and Il10 were up-regulated in all the treated groups. Likewise, genes involving gland morphogenesis and branch morphogenesis such as Bmp2, Tgm2, and Sox9 showed higher expression in 3D/PD-treated group. Bmp2 was up-regulated only in 3D/PD-treated group, combined with the results of GO analysis, we assumed that Bmp2 initiated SG fate through inducing branch morphogenesis and gland differentiation (Fig. 4C).

(A) Gene expression file of four groups of cells (R2DC, MSCs; R2DT, MSC with PD treatment; R3DC, MSC cultured in 3D construct; and R3DT, MSC treated with 3D/PD). (B) Up-regulated genes after treatment (2DC, MSCs; 2DT, MSC with PD treatment; 3DC, MSC cultured in 3D construct; and 3DT, MSC treated with 3D/PD). (C) Differentially expressed genes were further validated by RT-PCR analysis. [For all RT-PCR analyses, gene expression was normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) with 40 cycles, data are represented as the means SEM, and n = 3].

To validate the role of HMOX1 and CTHRC1 in the differentiation of MSCs to SG lineages, we analyzed the gene expression of Bmp2 by regulating the expression of Hmox1 and CTHRC1 based on the 3D/PD-treated MSCs. The effects of caffeic acid phenethyl ester (CAPE) and tin protoporphyrin IX dichloride (Snpp) on the expression of Hmox1 were evaluated by quantitative real-time (qRT)PCR. Hmox1 expression was significantly activated by CAPE and reduced by Snpp. Concentration of CTHRC1 was increased with recombinant CTHRC1 and decreased with CTHRC1 antibody. That is, it was negligible of the effects of activator and inhibitor of Hmox1 and CTHRC1 on cell proliferation (fig. S8, A and B). Hmox1 inhibition or CTHRC1 neutralization could significantly reduce the expression of Bmp2, while Hmox1 activation or increased CTHRC1 both activated Bmp2 expression. Furthermore, Bmp2 showed highest expression by up-regulation of Hmox1 and CTHRC1 simultaneously and sharply decreased with down-regulation of Hmox1 and CTHRC1 at the same time (Fig. 5A). Immunofluorescent staining revealed that the expression of bone morphogenetic protein 2 (BMP2) at the translational level with CTHRC1 and Hmox1 regulation showed a similar trend with transcriptional changes (Fig. 5B). Likewise, the expression of K8 and K18 at transcriptional and translational level changed similarly with CTHRC1 and Hmox1 regulation (fig. S9, A and B). These results suggested that CTHRC1 and Hmox1 played an essential role in SG fate separately, and they synergistically induced SG direction from MSCs (Fig. 5C).

(A and B) Transcriptional analysis (A) and translational analysis (PD, MSCs; PD+, MSCs with 3D/PD treatment; CAPE, MSCs treated with 3D/PD and Hmox1 activator; Snpp, MSCs treated with 3D/PD and Hmox1 inhibitor; Cthrc1, MSCs treated with 3D/PD and recombinant CTHRC1; anti, MSCs treated with 3D/PD and CTHRC1 antibody: +/+, MSCs treated with 3D/PD and Hmox1 activator and recombinant CTHRC1; and /, MSCs treated with 3D/PD and Hmox1 inhibitor and CTHRC1 antibody. Data are represented as the means SEM and n = 3) (B) of bmp2 with regulation of CTHRC1 and Hmox1. (C) The graphic illustration of 3D-bioprinted matrix directed MSC differentiation. CTHRC1 is the main biochemical cues during SG development, and structural cues up-regulated the expression of Hmox1 synergistically initiated branching morphogenesis of SG. *P < 0.05.

Next, we sought to assess the repair capacity of iSGCs for in vivo implications, the 3D-printed construct with green fluorescent protein (GFP)labeled MSCs was transplanted in burned paws of mice (Fig. 6A). We measured the SG repair effects by iodine/starch-based sweat test at day 14. Only mice with 3D/PD treatment showed black dots on foot pads (representing sweating), and the number increased within 10 min; however, no black dots were observed on untreated and single MSC-transplanted mouse foot pads even after 15 min (Fig. 6B). Likewise, hematoxylin and eosin staining analysis revealed SG regeneration in 3D/PD-treated mice (Fig. 6C). GFP-positive cells were characterized as secretory lumen expressing K8, K18, and K19. Of note, the GFP-positive cells were highly distributed in K14-positive myoepithelial cells of SGs but were absent in K14-positive repaired epidermal wounds (Fig. 6, D and E). Thus, differentiated MSCs enabled directed restitution of damaged SG tissues both at the morphological and functional level.

(A) Schematic illustration of approaches for engineering iSGCs and transplantation. (B) Sweat test of mice treated with different cells. Photo credit: Bin Yao, Wound Healing and Cell Biology Laboratory, Institute of Basic Medical Sciences, General Hospital of PLA. (C) Histology of plantar region without treatment and transplantation of MSCs and iSGCs (scale bars, 200 m). (D) Involvement of GFP-labeled iSGCs in directed regeneration of SG tissue in thermal-injured mouse model (K14, red; GFP, green; DAPI, blue; scale bar, 200 m). (E) SG-specific markers K14, K19, K8, and K18 detected in regenerated SG tissue (arrows). (K14, K19, K8, and K18, red; GFP, green; scale bars, 50 m).

A potential gap in MSC-based therapy still exists between current understandings of MSC performance in vivo in their microenvironment and their intractability outside of that microenvironment (24). To regulate MSCs differentiation into the right phenotype, an appropriate microenvironment should be created in a precisely controlled spatial and temporal manner (25). Recent advances in innovative technologies such as bioprinting have enabled the complexities in the matrix design and fabrication of regenerative microenvironments (26). Our findings demonstrated that directed differentiation of MSCs into SGs in a 3D-printed matrix both in vitro and in vivo was feasible. In contrast to conventional tissue-engineering strategies of SG regeneration, the present 3D-printing approach for SG regeneration with overall morphology and function offered a rapid and accurate approach that may represent a ready-to-use therapeutic tool.

Furthermore, bioprinting MSCs successfully repaired the damaged SG in vivo, suggesting that it can improve the regenerative potential of exogenous differentiated MSCs, thereby leading to translational applications. Notably, the GFP-labeled MSC-derived glandular cells were highly distributed in K14-positive myoepithelial cells of newly formed SGs but were absent in K14-positive repaired epidermal wounds. Compared with no black dots were observed on single MSC-transplanted mouse foot pads, the black dots (representing sweating function) can be observed throughout the entire examination period, and the number increased within 10 min on MSC-bioprinted mouse foot pads. Thus, differentiated MSCs by 3D bioprinting enabled exclusive restitution of damaged SG tissues morphologically and functionally.

Although several studies indicated that engineering 3D microenvironments enabled better control of stem cell fates and effective regeneration of functional tissues (2730), there were no studies concerning the establishment of 3D-bioprinted microenvironments that can preferentially induce MSCs differentiating into glandular cells with multiple tissue phenotypes and overall functional tissue. To find an optimal microenvironment for promoting MSC differentiation into specialized progeny, biochemical properties are considered as the first parameter to ensure SG specification. In this study, we used mouse PD as the main composition of a tissue-specific ECM. As expected, this 3D-printed PD+ microenvironment drove the MSC fate decision to enhance the SG phenotypic profile of the differentiated cells. By ECM differential protein expression analysis, we identified that CTHRC1 was a critical biochemical regulator of 3D-printed matrix for SG specification. TSP1 was required for the lineage-specific function during the differentiation in mice but was not dispensable for SG development. Thus, we identified CTHRC1 as a specific factor during SG development. To our knowledge, this is the first demonstration of CTHRC1 involvement in dictating MSC differentiation to SG, highlighting a potential therapeutic tool for SG injury.

The 3D-printed matrix also provided architectural guides for further SG morphogenesis. Our results clearly show that the 3D spatial dimensionality allows for better cell proliferation and aggregation and affect the characteristics of phenotypic marker expression. Notably, the importance of 3D structural cues on MSC differentiation was further proved by MET-EMT process during differentiation, where the influences did not depend on the presence of biochemical cues. To fully elucidate the underlying mechanisms, we first examined how 3D structure regulating stem cell fate choices. According to our data, Hmox1 is highly up-regulated in 3D construct, which were supposed to response to hypoxia, with a previously documented role in MSC differentiation (31, 32). It is suggested that 3D microenvironment induced rapid cell aggregation leading to hypoxia and then activated the expression of Hmox1.

Through regulation of the expression of Hmox1 and addition or of CTHRC1 in the matrix, we confirmed that each of them is critical for SG reprogramming, respectively. Thus, biochemical and structural cues of 3D-printed matrix synergistically creating a microenvironment could enhance the accuracy and efficiency of MSC differentiation, thereby leading to resulting SG formation. Although we further need a more extensive study examining the role of other multiple cues and their possible overlap function in regulating MSC differentiation, our findings suggest that CTHRC1 and Hmox1 provide important signals that cooperatively modulate MSC lineage specification toward sweat glandular lineage. The 3D structure combined with PD stimulated the GO functional item of branch morphogenesis and gland formation, which might be induce by up-regulation of Bmp2 based on the verification of qPCR results. Although our results could not rule out the involvement of other factors and their possible overlapping role in regulating MSC lineage specification toward SGs, our findings together with several literatures suggested that BMP2 plays a critical role in inducing branch morphogenesis and gland formation (3335).

In summary, our findings represented a novel strategy of directing MSC differentiation for functional SG regeneration by using 3D bioprinting and pave the way for a potential therapeutic tool for other complex glandular tissues as well as further investigation into directed differentiation in 3D conditions. Specifically, we showed that biochemical and structural cues of 3D-printed matrix synergistically direct MSC differentiation, and our results highlighted the importance of 3D-printed matrix cues as regulators of MSC fate decisions. This avenue opens up the intriguing possibility of shifting from genetic to microenvironmental manipulations of cell fate, which would be of particular interest for clinical applications of MSC-based therapies.

The main aim and design of the study was first to determine whether by using 3D-printed microenvironments, MSCs can be directed to differentiate and regenerate SGs both morphologically and functionally. Then, to investigate the underlying molecular mechanism of biochemical and structural cues of 3D-printed matrix involved in MSCs reprogramming. The primary aims of the study design were as follows: (i) cell aggregation and proliferation in a 3D-bioprinted construct; (ii) differentiation of MSCs at the cellular phenotype and functional levels in the 3D-bioprinted construct; (iii) the MET-EMT process during differentiation; (iv) differential protein expression of the SG niche in mice; (v) differential genes expression of MSCs in 3D-bioprinted construct; (vi) the key role of CTHRC1 and HMOX1 in MSCs reprogramming to SGCs; and (vii) functional properties of regenerated SG in vivo.

Gelatin (Sigma-Aldrich, USA) and sodium alginate (Sigma-Aldrich, USA) were dissolved in phosphate-buffered saline (PBS) at 15 and 1% (w/v), respectively. Both solutions were sterilized under 70C for 30 min three times at an interval of 30 min. The sterilized solutions were packed into 50-ml centrifuge tubes, stored at 4C, and incubated at 37C before use.

From wild-type C57/B16 mice (Huafukang Co., Beijing) aged 5 days old, dermal homogenates were prepared by homogenizing freshly collected hairless mouse PD with isotonic phosphate buffer (pH 7.4) for 20 min in an ice bath to obtain 25% (w/v) tissue suspension. The supernatant was obtained after centrifugation at 4C for 20 min at 10,000g. The DNA content was determined using Hoechst 33258 assay (Beyotime, Beijing). The fluorescence intensity was measured to assess the amount of remaining DNA within the decellularized ECMs and the native tissue using a fluorescence spectrophotometer (Thermo Scientific, Evolution 260 Bio, USA). The GAGs content was estimated via 1,9-dimethylmethylene blue solution staining. The absorbance was measured with microplate reader at wavelength of 492 nm. The standard curve was made using chondroitin sulfate A. The total COL (Collagen) content was determined via hydroxyproline assay. The absorbance of the samples was measured at 550 nm and quantified by referring to a standard curve made with hydroxyproline.

MSCs were bioprinted with matrix materials by using an extrusion-based 3D bioprinter (Regenovo Co., Bio-Architect PRO, Hangzhou). Briefly, 10 ml of gelatin solution (10% w/v) and 5 ml of alginate solution (2% w/v) were warmed under 37C for 20 min, gently mixed as bioink and used within 30 min. MSCs were collected from 100-mm dishes, dispersed into single cells, and 200 l of cell suspension was gently mixed with matrix material under room temperature with cell density 1 million ml1. PD (58 g/ml) was then gently mixed with bioink. Petri dishes at 60 mm were used as collecting plates in the 3D bioprinting process. Within a temperature-controlled chamber of the bioprinter, with temperature set within the gelation region of gelatin, the mixture of MSCs and matrix materials was bioprinted into a cylindrical construct layer by layer. The nozzle-insulation temperature and printing chamber temperature were set at 18 and 10C, respectively; nozzles with an inner diameter of 260 m were chosen for printing. The diameter of the cylindrical construct was 30 mm, with six layers in height. After the temperature-controlled bioprinting process, the printed 3D constructs were immersed in 100-mM calcium chloride (Sigma-Aldrich, USA) for 3 min for cross-linking, then washed with Dulbeccos modified Eagle medium (DMEM) (Gibco, USA) medium for three times. The whole printing process was finished in 10 min. The 3D cross-linked construct was cultured in DMEM in an atmosphere of 5% CO2 at 37C. The culture medium was changed to SG medium [contains 50% DMEM (Gibco, New York, NY) and 50% F12 (Gibco) supplemented with 5% fetal calf serum (Gibco), 1 ml/100 ml penicillin-streptomycin solution, 2 ng/ml liothyronine sodium (Gibco), 0.4 g/ml hydrocortisone succinate (Gibco), 10 ng/ml epidermal growth factor (PeproTech, Rocky Hill, NJ), and 1 ml/100 ml insulin-transferrin-selenium (Gibco)] 2 days later. The cell morphology was examined and recorded under an optical microscope (Olympus, CX40, Japan).

Fluorescent live/dead staining was used to determine cell viability in the 3D cell-loaded constructs according to the manufacturers instructions (Sigma-Aldrich, USA). Briefly, samples were gently washed in PBS three times. An amount of 1 M calcein acetoxymethyl (calcein AM) ester (Sigma-Aldrich, USA) and 2 M propidium iodide (Sigma-Aldrich, USA) was used to stain live cells (green) and dead cells (red) for 15 min while avoiding light. A laser scanning confocal microscopy system (Leica, TCSSP8, Germany) was used for image acquisition.

The cell-printed structure was harvested and fixed with a solution of 4% paraformaldehyde. The structure was embedded in optimal cutting temperature (OCT) compound (Sigma-Aldrich, USA) and sectioned 10-mm thick by using a cryotome (Leica, CM1950, Germany). The sliced samples were washed repeatedly with PBS solution to remove OCT compound and then permeabilized with a solution of 0.1% Triton X-100 (Sigma-Aldrich, USA) in PBS for 5 min. To reduce nonspecific background, sections were treated with 0.2% bovine serum albumin (Sigma-Aldrich, USA) solution in PBS for 20 min. To visualize iSGCs, sections were incubated with primary antibody overnight at 4C for anti-K8 (1:300), anti-K14 (1:300), anti-K18 (1:300), anti-K19 (1:300), anti-ATP1a1 (1:300), anti-Ki67 (1:300), antiN-cadherin (1:300), antiE-cadherin (1:300), anti-CTHRC1 (1:300), or anti-TSP1 (1:300; all Abcam, UK) and then incubated with secondary antibody for 2 hours at room temperature: Alexa Fluor 594 goat anti-rabbit (1:300), fluorescein isothiocyanate (FITC) goat anti-rabbit (1:300), FITC goat anti-mouse (1:300), or Alexa Fluor 594 goat anti-mouse (1:300; all Invitrogen, CA). Sections were also stained with 4,6-diamidino-2-phenylindole (Beyotime, Beijing) for 15 min. Stained samples were visualized, and images were captured under a confocal microscope.

To harvest the cells in the construct, the 3D constructs were dissolved by adding 55 mM sodium citrate and 20 mM EDTA (Sigma-Aldrich, USA) in 150 mM sodium chloride (Sigma-Aldrich, USA) for 5 min while gently shaking the petri dish for better dissolving. After transfer to 15-ml centrifuge tubes, the cell suspensions were centrifuged at 200 rpm for 3 min, and the supernatant liquid was removed to harvest cells for further analysis.

Total RNA was isolated from cells by using TRIzol reagent (Invitrogen, USA) following the manufacturers protocol. RNA concentration was measured by using a NanoPhotometer (Implen GmbH, P-330-31, Germany). Reverse transcription involved use of a complementary DNA synthesis kit (Takara, China). Gene expression was analyzed quantitatively by using SYBR green with the 7500 Real-Time PCR System (Takara, China). The primers and probes for genes were designed on the basis of published gene sequences (table S1) (National Center for Biotechnology Information and PubMed). The expression of each gene was normalized to that for glyceraldehyde-3-phosphate dehydrogenase and analyzed by the 2-CT method. Each sample was assessed in triplicate.

The culture medium was changed to SG medium with 2 mM CaCl2 for at least 24 hours, and cells were loaded with fluo-3/AM (Invitrogen, CA) at a final concentration of 5 M for 30 min at room temperature. After three washes with calcium-free PBS, 10 M acetylcholine (Sigma-Aldrich, USA) was added to cells. The change in the Fluo 3 fluorescent signal was recorded under a laser scanning confocal microscopy.

Cell proliferation was evaluated through CCK-8 (Cell counting kit-8) assay. Briefly, cells were seeded in 96-well plates at the appropriate concentration and cultured at 37C in an incubator for 4 hours. When cells were adhered, 10 l of CCK-8 working buffer was added into the 96-well plates and incubated at 37C for 1 hour. Absorbance at 450 nm was measured with a microplate reader (Tecan, SPARK 10M, Austria).

Proteomics of mouse PD and DD involved use of isobaric tags for relative and absolute quantification (iTRAQ) in BGI Company, with differentially expressed proteins detected in PD versus DD. Twofold greater difference in expression was considered significant for further study.

Tissues were grinded and lysed in radioimmunoprecipitation assay buffer (Beyotime, Nanjing). Proteins were separated by 12% SDSpolyacrylamide gel electrophoresis and transferred to a methanol-activated polyvinylidene difluoride membrane (GE Healthcare, USA). The membrane was blocked for 1 hour in PBS with Tween 20 containing 5% bovine serum albumin (Sigma-Aldrich, USA) and probed with the antibodies anti-CTHRC1 (1:1000) and anti-TSP1 (1:1000; both Abcam, UK) overnight at 4C. After 2 hours of incubation with goat anti-rabbit horseradish peroxidaseconjugated secondary antibody (Santa Cruz Biotechnology, CA), the protein bands were detected by using luminal reagent (GE Healthcare, ImageQuant LAS 4000, USA).

Total RNA was prepared with TRIzol (Invitrogen), and RNA sequencing was performed using HiSeq 2500 (Illumina). Genes with false discovery rate < 0.05, fold difference > 2.0, and mean log intensity > 2.0 were considered to be significant.

CAPE or Snpp was gently mixed with bioink at a concentration of 10 M. Physiological concentration of CTHRC1 was measured by enzyme linked immunosorbent assay (ELISA) (80 ng/ml), and then recombinant CTHRC1 or CTHRC1 antibody was added into the bioink at a concentration of 0.4 g/ml. The effect of inhibitor and activator was estimated by qRT-PCR or ELISA.

Mice were anesthetized with pentobarbital (100 mg/kg) and received subcutaneous buprenorphine (0.1 mg/kg) preoperatively. Full-thickness scald injuries were created on paw pads with soldering station (Weller, WSD81, Germany). Mice recovered in clean cages with paper bedding to prevent irritation or infection. Mice were monitored daily and euthanized at 30 days after wounding. Mice were maintained in an Association for Assessment and Accreditation of Laboratory Animal Careaccredited animal facility, and procedures were performed with Institutional Animal Care and Use Committeeapproved protocols.

MSCs in 3D-printed constructs with PD were cultured with DMEM for 2 days and then replaced with SG medium. The SG medium was changed every 2 days, and cells were harvested on day 12. The K18+ iSGCs were sorting through flow cytometry and injected into the paw pads (1 106 cells/50 l) of the mouse burn model by using Microliter syringes (Hamilton, 7655-01, USA). Then, mice were euthanized after 14 days; feet were excised and fixed with 10% formalin (Sigma-Aldrich, USA) overnight for paraffin sections and immunohistological analysis.

The foot pads of anesthetized treated mice were first painted with 2% (w/v) iodine/ethanol solution then with starch/castor oil solution (1 g/ml) (Sigma-Aldrich, USA). After drying, 50 l of 100 M acetylcholine (Sigma-Aldrich, USA) was injected subcutaneously into paws of mice. Pictures of the mouse foot pads were taken after 5, 10, and 15 min.

All data were presented as means SEM. Statistical analyses were performed using GraphPad Prism7 statistical software (GraphPad, USA). Significant differences were calculated by analysis of variance (ANOVA), followed by the Bonferroni test when performing multiple comparisons between groups. P < 0.05 was considered as a statistically significant difference.

Supplementary material for this article is available at http://advances.sciencemag.org/cgi/content/full/6/10/eaaz1094/DC1

Fig. S1. Biocompatibility of 3D-bioprinted construct and cellular morphology in 2D monolayer culture.

Fig. S2. Expression of SG-specific and secretion-related markers in MSCs and SG cells in vitro.

Fig. S3. Transcriptional and translational expression of epithelial and mesenchymal markers in 3D-bioprinted cells with and without PD.

Fig. S4. Expression of N- and E-cadherin in MSCs and SG cells in 2D monolayer culture.

Fig. S5. Proteomic microarray assay of differential gene expression between PD and DD ECM in postnatal mice.

Fig. S6. GO term analysis of differentially expressed pathways.

Fig. S7. Heat maps illustrating differential expression of genes implicated in ECM organization, cell division, and gland and branch morphogenesis.

Fig. S8. The expression of Hmox1 and the concentration of CTHRC1 on treatment and the related effects on cell proliferation.

Fig. S9. The expression of K8 and K18 with Hmox1 and CTHRC1 regulation.

Table S1. Primers for qRT-PCR of all the genes.

This is an open-access article distributed under the terms of the Creative Commons Attribution-NonCommercial license, which permits use, distribution, and reproduction in any medium, so long as the resultant use is not for commercial advantage and provided the original work is properly cited.

Acknowledgments: Funding: This study was supported in part by the National Nature Science Foundation of China (81571909, 81701906, 81830064, and 81721092), the National Key Research Development Plan (2017YFC1103300), Military Logistics Research Key Project (AWS17J005), and Fostering Funds of Chinese PLA General Hospital for National Distinguished Young Scholar Science Fund (2017-JQPY-002). Author contributions: B.Y. and S.H. were responsible for the design and primary technical process, conducted the experiments, collected and analyzed data, and wrote the manuscript. Y.W. and R.W. helped perform the main experiments. Y.Z. and T.H. participated in the 3D printing. W.S. and Z.L. participated in cell experiments and postexamination. S.H. and X.F. collectively oversaw the collection of data and data interpretation and revised the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors.

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Biochemical and structural cues of 3D-printed matrix synergistically direct MSC differentiation for functional sweat gland regeneration - Science...

Stem Cell Market Size Worth $17.9 Billion by 2027 | CAGR: 8.2%: Grand View Research, Inc. – PR Newswire UK

SAN FRANCISCO, March 2, 2020 /PRNewswire/ -- The global stem cell marketsize is expected to reach USD 17.9 billion by 2027, expanding at a CAGR of 8.2%, according to a new report by Grand View Research, Inc. Recent advances in tissue engineering strategies hold the potential to draw attention to the treatment of several chronic disorders. Moreover, automation in adult and cord blood processing and storage is expected to drive market growth significantly.

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The development of banking facilities and resultant enhancement of stem cell production, storage, and characterization are also expected to enhance the volumetric capabilities of this therapy, globally. This would lead to revenue generation in the global market.

Regenerative medicine and cellular therapies are considered to transform the healthcare industry in a few years. Therefore, key players are focused on developing advanced therapeutics for the treatment of Alzheimer's disease, Parkinson's disease, degenerative eye disorders, cancer, and stroke. Most of these therapies are under clinical trials and are expected to be launched soon.

Furthermore, the presence of a strong, diverse clinical pipeline is expected to accelerate revenue generation in the global market. Easy access to data through genomic, proteomic, and EHR databases has encouraged companies to investigate various therapies to aid in the treatment of previously untreatable conditions.

Grand View Research has segmented the global stem cells market on the basis of product, application, technology, therapy, and region:

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Grand View Research, U.S.-based market research and consulting company, provides syndicated as well as customized research reports and consulting services. Registered in California and headquartered in San Francisco, the company comprises over 425 analysts and consultants, adding more than 1200 market research reports to its vast database each year. These reports offer in-depth analysis on 46 industries across 25 major countries worldwide. With the help of an interactive market intelligence platform, Grand View Research helps Fortune 500 companies and renowned academic institutes understand the global and regional business environment and gauge the opportunities that lie ahead.

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Stem Cell Market Size Worth $17.9 Billion by 2027 | CAGR: 8.2%: Grand View Research, Inc. - PR Newswire UK

Avectas and CCRM partner on cell engineering – BioPharma-Reporter.com

The Solupore platform looks to support the burgeoning industry pipeline of cell and gene therapies by supporting the non-viral cell engineering of therapeutics.

At present, a shortage in capacity for the production of viral vectors, used in the development of cell and gene therapies, has resulted in waiting lists being established for companies to have access to the limited supply.

Avectas and CCRM (Centre for Commercialization of Regenerative Medicine) have partnered with the aim of addressing this need, by facilitating the Solupore platform into the clinic.

According to Avectas, its platform utilizes a membrane disruptive approach to deliver nucleic acids and proteins to cells. The Irish company is currently developing a closed continuous system for good manufacturing practice (GMP) manufacturing.

In terms of the benefits of its method of cell engineering, the company stated that unlike industry standards, Avectas technology is gentle to the cells allowing rapid recovery and functionality. There is no stall time in cell proliferation resulting in a faster cell engineering process.

The technology is also scalable and can be adjusted to the volume of cells needed for clinical therapies, suggested Avectas. The potential molecules that could be generated by this approach include mRNA, proteins and gene editing tools.

For its part, CCRM is a non-profit organization that is able to offer expertise in stem cell and biomaterials technologies through its strategic advisory board, which combines expertise from Canadian and US universities, amongst other global participants.

In addition, the CCRM has a network of industry companies, which features Amgen, Pfizer, Pall, and Roche.

Michael Maguire, CEO of Avectas, said: We are delighted to partner with CCRMto leverage their deep experience in cell manufacturing processes to support the translation of our Solupore platform towards clinical applications.

As well as looking to partner with immuno-oncology and gene editing business to produce their drug candidates, Avectas noted that it is also looking to in-license therapeutic molecules to build its own pipeline of potential products.

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Regenerative Medicine Market Analysis Growth Demand, Key Players, Share Size, and Forecast To 2025 – Monroe Scoop

Regenerative Medicine Market: Snapshot

Regenerative medicine is a part of translational research in the fields of molecular biology and tissue engineering. This type of medicine involves replacing and regenerating human cells, organs, and tissues with the help of specific processes. Doing this may involve a partial or complete reengineering of human cells so that they start to function normally.

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Regenerative medicine also involves the attempts to grow tissues and organs in a laboratory environment, wherein they can be put in a body that cannot heal a particular part. Such implants are mainly preferred to be derived from the patients own tissues and cells, particularly stem cells. Looking at the promising nature of stem cells to heal and regenerative various parts of the body, this field is certainly expected to see a bright future. Doing this can help avoid opting for organ donation, thus saving costs. Some healthcare centers might showcase a shortage of organ donations, and this is where tissues regenerated using patients own cells are highly helpful.

There are several source materials from which regeneration can be facilitated. Extracellular matrix materials are commonly used source substances all over the globe. They are mainly used for reconstructive surgery, chronic wound healing, and orthopedic surgeries. In recent times, these materials have also been used in heart surgeries, specifically aimed at repairing damaged portions.

Cells derived from the umbilical cord also have the potential to be used as source material for bringing about regeneration in a patient. A vast research has also been conducted in this context. Treatment of diabetes, organ failure, and other chronic diseases is highly possible by using cord blood cells. Apart from these cells, Whartons jelly and cord lining have also been shortlisted as possible sources for mesenchymal stem cells. Extensive research has conducted to study how these cells can be used to treat lung diseases, lung injury, leukemia, liver diseases, diabetes, and immunity-based disorders, among others.

Global Regenerative Medicine Market: Overview

The global market for regenerative medicine market is expected to grow at a significant pace throughout the forecast period. The rising preference of patients for personalized medicines and the advancements in technology are estimated to accelerate the growth of the global regenerative medicine market in the next few years. As a result, this market is likely to witness a healthy growth and attract a large number of players in the next few years. The development of novel regenerative medicine is estimated to benefit the key players and supplement the markets growth in the near future.

Global Regenerative Medicine Market: Key Trends

The rising prevalence of chronic diseases and the rising focus on cell therapy products are the key factors that are estimated to fuel the growth of the global regenerative medicine market in the next few years. In addition, the increasing funding by government bodies and development of new and innovative products are anticipated to supplement the growth of the overall market in the next few years.

On the flip side, the ethical challenges in the stem cell research are likely to restrict the growth of the global regenerative medicine market throughout the forecast period. In addition, the stringent regulatory rules and regulations are predicted to impact the approvals of new products, thus hampering the growth of the overall market in the near future.

Global Regenerative Medicine Market: Market Potential

The growing demand for organ transplantation across the globe is anticipated to boost the demand for regenerative medicines in the next few years. In addition, the rapid growth in the geriatric population and the significant rise in the global healthcare expenditure is predicted to encourage the growth of the market. The presence of a strong pipeline is likely to contribute towards the markets growth in the near future.

Global Regenerative Medicine Market: Regional Outlook

In the past few years, North America led the global regenerative medicine market and is likely to remain in the topmost position throughout the forecast period. This region is expected to account for a massive share of the global market, owing to the rising prevalence of cancer, cardiac diseases, and autoimmunity. In addition, the rising demand for regenerative medicines from the U.S. and the rising government funding are some of the other key aspects that are likely to fuel the growth of the North America market in the near future.

Furthermore, Asia Pacific is expected to register a substantial growth rate in the next few years. The high growth of this region can be attributed to the availability of funding for research and the development of research centers. In addition, the increasing contribution from India, China, and Japan is likely to supplement the growth of the market in the near future.

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Global Regenerative Medicine Market: Competitive Analysis

The global market for regenerative medicines is extremely fragmented and competitive in nature, thanks to the presence of a large number of players operating in it. In order to gain a competitive edge in the global market, the key players in the market are focusing on technological developments and research and development activities. In addition, the rising number of mergers and acquisitions and collaborations is likely to benefit the prominent players in the market and encourage the overall growth in the next few years.

Some of the key players operating in the regenerative medicine market across the globe areVericel Corporation, Japan Tissue Engineering Co., Ltd., Stryker Corporation, Acelity L.P. Inc. (KCI Licensing), Organogenesis Inc., Medtronic PLC, Cook Biotech Incorporated, Osiris Therapeutics, Inc., Integra Lifesciences Corporation, and Nuvasive, Inc.A large number of players are anticipated to enter the global market throughout the forecast period.

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Autologous Stem Cell And Non-Stem Cell Based Therapies Market 2020-2025 || Leading Players Fibrocell, Genesis Biopharma, Georgia Health Sciences…

Todays businesses call for the highly focused, comprehensive and detail-oriented information about the market so that they get a clear idea about the market landscape. The Autologous Stem Cell And Non-Stem Cell Based Therapies market research report is generated with a combination of detailed industry insights, and use of latest tools and technology. The study of this market research report covers a market attractiveness analysis, wherein each segment is targeted based on its market size, growth rate, and general attractiveness. The Autologous Stem Cell And Non-Stem Cell Based Therapies market research report plays a key role in developing the strategies for sales, advertising, marketing, and promotion.

TheGlobalAutologous Stem Cell and Non-Stem Cell Based Therapies Marketis expected to reach USD113.04 billion by 2025, from USD 87.59 billion in 2017 growing at a CAGR of 3.7% during the forecast period of 2018 to 2025. The upcoming market report contains data for historic years 2015 & 2016, the base year of calculation is 2017 and the forecast period is 2018 to 2025.

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Some of the major players operating in the global autologous stem cell and non-stem cell based therapies market areAntria (Cro), Bioheart, Brainstorm Cell Therapeutics, Cytori, Dendreon Corporation, Fibrocell, Genesis Biopharma, Georgia Health Sciences University, Neostem, Opexa Therapeutics, Orgenesis, Regenexx, Regeneus, Tengion, Tigenix, Virxsys and many more.

Market Definition:Global Autologous Stem Cell and Non-Stem Cell Based Therapies Market

In autologous stem-cell transplantation persons own undifferentiated cells or stem cells are collected and transplanted back to the person after intensive therapy. These therapies are performed by means of hematopoietic stem cells, in some of the cases cardiac cells are used to fix the damages caused due to heart attacks. The autologous stem cell and non-stem cell based therapies are used in the treatment of various diseases such as neurodegenerative diseases, cardiovascular diseases, cancer and autoimmune diseases, infectious disease.

According to World Health Organization (WHO), cardiovascular disease (CVD) causes more than half of all deaths across the European Region. The disease leads to death or frequently it is caused by AIDS, tuberculosis and malaria combined in Europe. With the prevalence of cancer and diabetes in all age groups globally the need of steam cell based therapies is increasing, according to article published by the US National Library of Medicine National Institutes of Health, it was reported that around 382 million people had diabetes in 2013 and the number is growing at alarming rate which has increased the need to improve treatment and therapies regarding the diseases.

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Market Segmentation:Global Autologous Stem Cell and Non-Stem Cell Based Therapies Market

Competitive Analysis:Global Autologous Stem Cell and Non-Stem Cell Based Therapies Market

The global autologous stem cell and non-stem cell based therapies market is highly fragmented and the major players have used various strategies such as new product launches, expansions, agreements, joint ventures, partnerships, acquisitions, and others to increase their footprints in this market. The report includes market shares of autologous stem cell and non-stem cell based therapies market for global, Europe, North America, Asia Pacific and South America.

Major Autologous Stem Cell and Non-Stem Cell Based Therapies Market Drivers and Restraints:

Introduction of novel autologous stem cell based therapies in regenerative medicine

Reduction in transplant associated risks

Prevalence of cancer and diabetes in all age groups

High cost of autologous cellular therapies

Lack of skilled professionals

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Autologous Stem Cell And Non-Stem Cell Based Therapies Market 2020-2025 || Leading Players Fibrocell, Genesis Biopharma, Georgia Health Sciences...

Why Aimmune Therapeutics Shares Fell 23.3% in February – Nasdaq

What happened

Shares of Aimmune Therapeutics (NASDAQ: AIMT) fell 23.3% in February, according to data provided by S&P Global Market Intelligence, as the launch of its new peanut allergy treatment took time to unfold.

Aimmune rose 40% last year as investors anticipated Palforzia, the firsttreatment approved by the Food and Drug Administration for peanut allergy in children. The regulatory agency approved the treatment on Jan. 31, and now investors are waiting to see how the first weeks of sales progress. The process is taking time because of the risk management procedure involved in the launch.

Image source: Getty Images.

Palforzia, an orally administered powder made of peanut protein, works by desensitizing patients to the allergen. In order to lower risk in case a new patient suffers a reaction to the therapy, the FDA has set up the Risk Evaluation and Mitigation Strategy (REMS) program. Physicians and patients must enroll in REMS and follow guidelines before treatment can begin. This process, along with the FDA's standard procedure of examining and releasing the first batches of biologic product, lengthened the timeline from approval to sales. Aimmune said during its recent earnings call that it expects to record the first Palforzia sales this month.

Getting physicians and patients on board in the next few months will be crucial for Aimmune. DBVTechnologies (NASDAQ: DBVT) is close behind, with an FDA decision on its peanut allergy drug expected in August. Aimmune now has the advantage of being first to market, before rivals enter with competing products. Aimmune is also expecting a decision on Palforzia from the Europeanregulatory agency in the fourth quarter, and said a decision in Switzerland could come in mid-2021.

All of these elements represent catalysts for the shares over the next year or so. If Aimmune can assure its position as market leader, the shares of this biotech company will benefit in the long term.

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Why Aimmune Therapeutics Shares Fell 23.3% in February - Nasdaq

Mapping the structure and biological functions within mesenchymal bodies using microfluidics – Science Advances

INTRODUCTION

In recent years, organoids have emerged as powerful tools for basic research, drug screening, and tissue engineering. The organoids formed in vitro show many features of the structural organization and the functional hallmarks of adult or embryonic anatomical structures (1). In addition, the formation of organoids alleviates the need to perform animal studies and provides an attractive platform for robust quantitative studies on the mechanisms regulating organ homeostasis and tissue repair in vivo (1). The formation of organoids usually starts with populations of stem cells. They are therefore expected to be heterogeneous because pluripotent stem cells [induced pluripotent stem cells (pscs) or embryonic stem cells] have been shown to dynamically and stochastically fluctuate from ground to differentiated state (2). In the same vein, LGR5+ intestinal stem cells are reported to contain several distinct populations (3). As such, the formation of organoids involves the inherent capacity of these heterogeneous populations to self-sort and self-pattern to form an organized three-dimensional (3D) architecture (4). However, the rules underlying organoid formation as well as the contribution of intrinsic population heterogeneity to the organoid self-assembly remain poorly understood (5). Consequently, there is a need for novel quantitative approaches at the single-cell level to reliably understand the mechanisms of spatial tissue patterning in 3D organoids, for which microfluidic and quantitative image analysis methods are well suited.

In this work, we use mesenchymal progenitors, alternatively named mesenchymal stromal cells (MSCs), which constitute a self-renewing population with the ability to differentiate into adipocytes, chondrocytes, and osteoblasts (5). Although human MSCs (HMSCs) express high levels of undifferentiation markers (e.g., CD105, CD44, CD73), they constitute a heterogeneous population of cells that exhibit considerable variation in their biophysical properties and epigenetic status, as well as the basal level of expression of genes related to differentiation, immunoregulation, and angiogenesis (6, 7). Nonetheless, their aggregation leads to the formation of highly cohesive 3D spherical structures [which we designate hereafter as mesenchymal bodies (MBs)] with improved biological activities in comparison to 2D cultures (8). However, little is known on how HMSCs self-organize or whether the intrinsic heterogeneity of the population regulates MB formation and individual cell functions in 3D.

The self-aggregation of HMSCs into MBs can recapitulate the early stages of mesenchymal condensation, and it promotes the secretion of paracrine molecules taking part in the process of ossification (9). During mesenchymal condensation in vivo, mesenchymal progenitors self-aggregate and form dense cell-cell contacts that lead to the initiation of bone organogenesis through endochondral (necessitating a chondrogenic intermediate) and intramembranous (direct osteogenic differentiation) ossification (10). In addition, the formation of these 3D MBs in vivo is associated with the secretion of important paracrine molecules such as prostaglandin E2 (PGE2) and vascular endothelial growth factor (VEGF), which participate in the recruitment of endogenous osteoblasts, osteoclasts, and blood vessels, leading to the initiation/restoration of bone homeostasis (11, 12). In these two ossification processes, the induction of nuclear factor B (NF-B) target genes, such as cyclooxygenase-2 (COX-2), and their downstream products (e.g., PGE2 and VEGF) plays a critical role as developmental regulators of ossification and bone healing (13). However, while mesenchymal condensation is critical for bone organogenesis, there is still a limited understanding on how the cellular spatial organization within 3D MBs regulates the individual cells endocrine functions (14).

In the present work, we interrogate the influence of phenotypic heterogeneity within a population of stem cells on the mechanisms of self-assembly and functional patterning within 3D organoids using HMSCs as a model of heterogeneous progenitor cell population. This is performed using a novel microfluidic platform for high-density formation of mensenchymal bodies, combined with the analysis of individual cells by quantitative image analysis. Our study reveals that the progenitor cell population self-assembles in a developmentally hierarchical manner. We also find that the structural arrangement in mensenchymal bodies is linked with the functional patterning in 3D, through a modulation of the activity of regulatory molecular signaling at a local scale. This study demonstrates the interplay between cell size and differentiation status, which mediates cellular spatial rearrangement in 3D, leading to the regionalized activation of unique biological functions while forming aggregates.

HMSCs are known to constitute a heterogeneous population (6, 7). In this study, fetal HMSCs were derived from the Whartons jelly of the umbilical cord (UC). UC-derived HMSCs are considered to be more primitive than HMSCs derived from adult bone marrow because of their higher proliferative capacity, their ability to form colony-forming unitfibroblast, as well as their lower degree of basal commitment (15). To examine the cellular diversity within the population, HMSCs were first characterized by their expression of membrane markers. Most of the HMSC population consistently expresses CD73, CD90, CD105, and CD146, but not CD31 (an endothelial cell marker), CD34 (a hematopoietic cell marker), CD14 (an immune cell marker), or human leukocyte antigenDR (HLA-DR) (a type of major histocompatibility complex II) (Fig. 1, A to F, and fig. S1, A to C). However, a deeper analysis of the flow cytometric data shows that the HMSC population contains cells of heterogeneous size [coefficient of variation (CV) = 33 to 37%] (Fig. 1, G and I), having a broad distribution in the expression of CD146 (Fig. 1F). Of note, the CD146 level of expression was linked to the size of the cells: The highest levels of CD146 were found for the largest cells (Fig. 1, H and J). Similar correlations with cell size were also observed for CD73, CD90, and CD105 (fig. S1, D to F). In addition, upon specific induction, the HMSC population used in this study successfully adopted an adipogenic (Fig. 1K), an osteogenic (Fig. 1L), or a chondrogenic (Fig. 1M) phenotype, demonstrating their mesenchymal progenitor identity.

(A) Percentage of positive cells for CD31, CD73, CD90, CD105, and CD146 (n = 3). Representative histograms of the distribution of the CD31 (B), CD73 (C), CD90 (D), CD105 (E), and CD146 (F) level of expression are shown. (G) Representative histogram of the forward scatter (FSC) distribution. (H) Correlation between cell size [FSC and side scatter (SSC)] and the level of CD146 expression. (I) Representative histogram of the cell projected area distribution. (J) Representative histogram of the size distribution of the CD146dim, CD146int, and CD146bright (ImageSteam analysis). (K) Representative images of hMSCs differentiated toward adipogenic lineage (Oil Red O staining). (L) Representative images of UC-hMSCs differentiated toward osteogenic lineage in (Alizarin Red S staining). (M) Representative images of UC-hMSCs differentiated toward chondrogenic lineage (Alcian Blue staining in 2D and cryosectioned micromass cultures). Scale bars, 50 m. The images were acquired using a binocular. FITC-A, fluorescein isothiocyanateA; APC-A, allophycocyanin-A.

To interrogate contribution of cellular heterogeneity (i.e., in terms of size and levels of CD marker expression) in the self-organization of HMSCs in 3D, MBs were formed at high density on an integrated microfluidic chip. This was done by encapsulating cells into microfluidic droplets at a density of 380 cells per droplet, with a CV of 24% (fig. S2, A and B). The drops were then immobilized in 250 capillary anchors in a culture chamber, as previously described (Fig. 2, A and B) (16). The loading time for the microfluidic device was about 5 min, after which the typical time for complete formation of MBs was about 4 hours (movie S1), as obtained by measuring the time evolution of the projected area (Fig. 2, C and D) and circularity of individual MBs (Fig. 2E and movie S2). The protocol resulted in the formation of a single MB per anchor (fig. S2C) with an average diameter of 158 m (Fig. 2F), when starting with a seeding concentration of 6 106 cells ml1. The diameter of the aggregates can easily be tuned by modulating the concentration of cells in the seeding solution (fig. S2, A and B). In addition, the complete protocol yielded the reproducible formation of a high-density array of fully viable MBs ready for long-term culture (for the images of the individual fluorescent channel, see Fig. 2G and fig. S2D), as described previously (16). Of interest, the CV of the MB diameter distribution was lower than the CV of the individual cell size and of the cell number in droplets (CV MB diameter = 13.3%, CV cell number per drop = 24%, and CV cell size = 35%), which demonstrates that the production of MBs leads to more homogeneous size conditions, compared with the broad heterogeneity in the cell population.

(A) Chip design. Scale bar, 1 cm. (B) Schematized side view of an anchor through the MB formation and culture protocol. (C) Representative time lapse of an MB formation. Scale bar, 100 m. (D and E) Measurement of the time evolution of the projected area (D) and circularity of each aggregate (E). n = 120 MBs. (F) Distribution of the MB diameter normalized by the mean of each chip (n = 10,072 MBs). (G) Top: Representative images of MBs after agarose gelation and oil-to-medium phase change. Bottom: The same MBs are stained with LIVE/DEAD. Scale bar, 100 m. (H) Representative images of MBs formed in the presence of EDTA, an N-cadherin, or a CD146-conjugated blocking antibody (Ab) (the red color shows the position of the CD146 brightest cells, and the dilution of the antibody was 1/100 and remain in the droplet for the whole experiment). Scale bar, 100 m. The images were acquired using a wide-field microscope.

To gain insight into the cellular components required to initiate the self-organization of HMSCs in 3D, the MB formation was disrupted by altering cell-cell interactions. This was first performed by adding EDTA, a chelating agent of the calcium involved in the formation of cadherin junctions, to the droplet contents. Doing so disrupted the MB formation, as shown in Fig. 2H, where the projected area of the cells increased and the circularity decreased in the presence of EDTA compared with the controls, as previously reported (17). The role of N-cadherins among different types of cadherins was further specified by adding a blocking antibody in the droplets before MB formation. This also led to a disruption of the MB formation, demonstrating that N-cadherin homodimeric interactions are mandatory to initiate the process of HMSC aggregation. CD146 [melanoma cell adhesion molecule (M-CAM)] plays important dual roles: as an adhesion molecule (that binds to Laminin 411) (18) and a marker of the commitment of HMSCs (19). We, thus, interrogate its contribution to MB formation. The addition of a CD146-conjugated blocking antibody also disrupted the formation of the MB (Fig. 2H), demonstrating that cell-cell interactions involving CD146 are also required during MB formation, as reported with other cell types (18). Of note, the brightest signal from the CD146-stained cells was located in the core of the cellular aggregates (Fig. 2H), suggesting that HMSCs self-organize relatively to their degree of commitment.

We found that the population of HMSCs constituted of cells of broad size and expressing different levels of undifferentiated markers [i.e., CD90, CD73, CD105, and CD146 are known to be down-regulated upon differentiation; (20)] and that the cells are capable of self-organizing cohesively in 3D. To better understand how the heterogeneous cells organized within the MBs, we measured how the different cell types composing the population self-assembled spatially in 3D by investigating the role of CD146. For this purpose, the CD146dim and CD146bright cells were separated from the whole HMSC population by flow cytometry (Fig. 3, A and B). The cells were then reseeded on a chip for the MB formation after fluorescently labeling the brighter and/or the dimmer CD146 populations. Image analysis revealed that the CD146bright cells were mostly located in the center of the cellular aggregates, while CD146dim cells were found at the boundaries of the MBs (Fig. 3, C to E, figs. S3A and S5A for confocal images, and movie S1). This organization was stable for a 3-day culture (fig. S3B).

(A) Representative dot plot of the hMSC population separation based on the level of CD146: The CD146dim constitutes 20% of the population expressing the highest levels of CD146; the CD146bright constitutes the 20% of the population expressing the lowest levels of CD146. (B) Fluorescence signal distribution in the CD146dim and CD146bright populations after cell sorting. (C) After cell sorting, the CD146bright or the CD146dim was stained with Vybrant Dil (red) or Vybrant DiO (green), remixed together and allowed to form MBs. Representative images of the CD146bright and CD146dim within the MBs. Scale bar, 100 m (n = 185 MBs). (D) The position of the CD146bright and CD146dim was quantified by correlating the fluorescence signal of the different stained cells as a function of their radial position within the MBs, after the staining of individual population with Vybrant Dil (CD146bright) = 500 and CD146dim (MBs; CD146bright = 85). Error bars show the SD. (E) Schematized representation of the structural organization of MBs. (F and G) RT-qPCR analysis of the relative RUNX-2, CEBP/, and SOX-9 expression to glyceraldehyde-3-phosphate dehydrogenase (GADPH) [Ct (cycle threshold) (D) and relative RNA expression (E)] in the CD146bright and CD146dim populations (n = 3). *P < 0.05; **P < 0.01; ***P < 0.001.

As we found that the CD146bright cells were larger than the CD146dim cells, the cells from the HMSC population were also separated on the basis of their relative size (a parameter that also discriminates the CD90, CD105, and CD73bright from the CD90, CD105, and CD73dim cells; fig. S1, D to F). After reseeding on the chip, the MBs were composed of large cells in the core, while the smallest cells were located at the boundaries, as expected from the previous experiments (fig. S3A). Moreover, we found that the speed of self-assembly of each population is not related to the rearrangement of CD146dim and CD146bright cells in 3D, because the mixing of dissociated cells or the fusion of aggregates made each population give rise to the same structural organization (21). It is well established that CD146bright defines the most undifferentiated HMSCs (20). The heterogeneity in level of commitment between the two subpopulations was therefore checked by reverse transcription quantitative polymerase chain reaction (RT-qPCR) analysis to quantify differences in the expression of differentiation markers. The analysis showed that the CD146dim cells expressed higher levels of osteogenic differentiation markers (i.e., RUNX-2) than the CD146bright cells (Fig. 3, F and G).

The level of RUNX-2 expression was also quantified at the protein level using immunocytochemistry and image analysis of the MBs on the microfluidic device by developing a layer-by-layer description of the MBs. This mapping was constructed by estimating the boundaries of each cell in the image from a Voronoi diagram, built around the positions of the cell nuclei stained with 4,6-diamidino-2-phenylindole (DAPI) (Fig. 4A) (22). These estimates were then used to associate the fluorescence signal from each cell with one of the concentric layers (Fig. 4B). Such a mapping provides better resolution for discriminating the spatial heterogeneity of protein expression than simply assigning a fluorescence signal to a defined radial coordinate (fig. S4). Moreover, the reliability of the measurements by quantitative image analysis was confirmed by performing several control experiments. In particular, we verified (i) the specificity of the fluorescence labeling, (ii) the absence of limitation for antibody diffusion, and (iii) the absence of the light path alteration in the 3D structure (fig. S5 and Materials and Methods). Consistent with the qPCR data, we found that HMSCs located at the boundaries of the MBs expressed higher levels of the protein RUNX-2 than the cells located in the core (see Fig. 4, C and D, and fig. S7 for individual experiments).

(A and B) The detection of nuclei within MBs enables the construction of a Voronoi diagram (A) that allows the identification of concentric cell layers (B) within the MBs. (C and D) Representative image (C) and quantitative analysis (D) (error bars represent the SD) of RUNX-2 staining within the cell layers of the MB (Nchips = 3 and nMBs = 458). N.S., nonsignificant. (E to H) Quantitative analysis (E) and representative images (F to H) of N-cadherin staining after methanol/acetone (F) (Nchips = 3 and nMBs = 405), after PFA/Triton X-100 fixation and permeabilization (G) (Nchips = 3 and nMBs = 649), and F-actin staining with phalloidin (H) (Nchips = 3 and nMBs = 421). Scale bars, 20 m. The images were acquired using a wide-field microscope. ***P < 0.001. (I) Schematized representation of the structural organization of MBs.

Thus, as CD146 defines the most undifferentiated and clonogenic cells as well as regulates the trilineage differentiation potential of HMSCs, the results indicate that HMSCs self-organize within MBs based on their initial commitment. The most undifferentiated and largest cells are found in the core (r/R < 0.8), while more differentiated cells positioned in the outer layers of the MBs (r/R > 0.8) (Fig. 4, C to E). In addition, these data reveal that HMSCs are conditioned a priori to occupy a specific location within the MBs.

The commitment of HMSCs is known to regulate their level of CD146 expression and the type of cell-cell adhesion molecules (23), which plays a fundamental role in the structural cohesion of the MBs (Fig. 2H). For this reason, we interrogated the organization of cell-cell junctions after the MB formation through measurements of the N-cadherin and F-actin fluorescence signal distribution. Two different protocols were used to discriminate several forms of N-cadherin interactions. First, paraformaldehyde (PFA) fixation and Triton X-100 permeabilization were used, because they were reported to retain in place only the detergent-insoluble forms of N-cadherin. Alternatively, ice-cold methanol/acetone fixation and permeabilization enabled the detection of all forms of N-cadherins (26). The results show a higher density of total N-cadherins in the core of the MBs (Fig. 4, E and F), while a higher density of F-actin was found in the cell layers located near the edge of the MBs (Fig. 4, E and H). The pattern of F-actin distribution was not related to the agarose gel surrounding the MBs (fig. S5B). These results are consistent with the theories of cell sorting in spheroids that postulate that more adhesive cells (i.e., expressing more N-cadherin or CD146) should be located in the core, while more contractile cells (i.e., containing denser F-actin) are located at the edge of the MBs (24). Moreover, our observations are in accordance with recent results demonstrating that HMSCs establishing higher N-cadherin interactions show reduced osteogenic commitment than HMSCs making fewer N-cadherin contacts, potentially through the modulation of Yap/Taz signaling and cell contractility (23).

In contrast, the most triton-insoluble forms of N-cadherins were located at the boundaries of the HMSC aggregates (Fig. 4, E and G), at the same position as the cells containing the denser F-actin. These results demonstrate that different types of cellular interactions were formed between the core and the edges of the MBs, which correlated with the degree of cell commitment that apparently stabilize the adherens junctions (Fig. 4I) (25).

We found above that the degree of commitment was linked with the pattern of HMSC self-organization in MBs (i.e., formation of adherens junctions), which may also regulate their paracrine functions (26). We therefore interrogated the functional consequences of the cellular organization in MBs by investigating the distribution of VEGF- and PGE2-producing cells.

The specific production of COX-2, VEGF, and two other molecules regulating bone homeostasis such as tumor necrosis factorinducible gene 6 (TSG-6) (27) and stanniocalcin 1 (STC-1) (28) was evaluated by RT-qPCR analysis. An increased transcription (20- to 60-fold) of these molecules was measured in 3D in comparison to the monolayer culture (Fig. 5, A and B). Consistent with this observation, while a very limited level of secreted PGE2 and VEGF was measured by enzyme-linked immunosorbent assay (ELISA) in 2D culture, they were significantly increased (by about 15-fold) upon the aggregation of HMSCs in 3D (Fig. 5C). In addition, to interrogate the specific role of COX-2 [the only inducible enzyme catalyzing the conversion of arachidonic acid into prostanoids; (29)] in PGE2 and VEGF production, indomethacin (a pan-COX inhibitor) was added to the culture medium. Indomethacin abrogated the production of PGE2, and it significantly decreased VEGF secretion (Fig. 5C), which suggests an intricate link between COX-2 expression and the secretion of these two molecules (30, 31).

(A and B) RT-qPCR analysis of the relative TSG-6, COX-2, STC-1, and VEGF expression to GADPH (Ct) (A) and relative RNA expression (B) in the 3D and 2D populations (n3D = 3 and n2D = 3). (C) Quantification by ELISA of the PGE-2 and VEGF secreted by hMSCs cultivated in 2D, as MBs or as MBs treated with indomethacin (nchips = 3 and n2D = 3). (D and E) Representative image (D) and quantitative analysis (E) of COX-2 (Nchips = 13 and nMBs = 2936) and (F) VEGF-A (Nchips = 3 and nMBs = 413) staining within the cell layers of the MBs (error bars represent the SD). Scale bars, 50 m. The images were acquired using a wide-field microscope *P < 0.05; ***P < 0.001; a and b: P < 0.05. (G) Schematized representation of the structural organization of MBs.

To further interrogate the link between the COX-2 and the VEGF-producing cells, their location was analyzed by quantitative image analysis at a layer-by-layer resolution. These measurements showed significantly higher levels of COX-2 in the first two layers, compared with the successive layers of the MBs (Fig. 5, D and E), with a continuous decrease of about 40% of the COX-2 signal between the edge and the core. This pattern of COX-2 distribution was not affected by the MB diameter (fig. S7B). Similar observations were made with VEGF (Fig. 5, D and F), demonstrating that cells at the boundaries of the MBs expressed both COX-2 and VEGF (Fig. 5G). Taken with the measurements of Fig. 5C, these results imply that COX-2 acts as an upstream regulator of PGE2 and VEGF secretion. Conversely, oxygen deprivation was unlikely to occur within the center of the MBs because no hypoxic area was detected through the whole MBs (fig. S5). Consequently, it is unlikely that hypoxia-inducible factor1 (HIF-1) signaling mediates the increase in VEGF expression at the boundaries of the MBs. Note that finding the link between these three molecules requires the 3D format, because the molecules are not detected in 2D. Here, the combination of population-scale measurements (Fig. 5C) and cell layer analysis (Fig. 5, E and F) provides strong evidence for this pathway.

Because variations of COX-2 and adherens junction distribution are colocalized within the MBs (Figs. 4, E to G, and 5, D and E), the results point to a link between the quality of cell-cell interactions and the spatial distribution of the COX-2high cells in 3D. The mechanisms leading to the spatial patterning of COX-2 expression in the MBs were therefore explored using inhibitors of the signaling pathways related to anti-inflammatory molecule production and of the molecular pathways regulating the structural organization (table S3): (i) 4-N-[2-(4-phenoxyphenyl)ethyl]quinazoline-4,6-diamine (QNZ) that inhibits NF-B, a critical transcription factor regulating the level of COX-2 expression (32); (ii) N-[N-(3,5-difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (DAPT) that inhibits the canonical Notch pathway, modulating cell-cell interactions and several differentiation pathways; (iii) Y-27632 (Y27) that inhibits ROCK involved in the bundling of F-actin (i.e., formation of stress fibers) to assess the role of actomyosin organization; and (iv) cytochalasin D (CytoD) that inhibits the polymerization of actin monomers.

While the addition of DAPT had virtually no effect on the ability of the cells to form MBs, Y27 led to MBs with more rounded cells, and both QNZ and CytoD strongly interfered with the MB formation process (Fig. 6, A to C). The results indicate that NF-B activation and the promotion of actin polymerization are critical signaling steps initiating the process of MB formation by HMSCs.

(A) Representative images of MBs formed 1 day after the droplet loading. Scale bar, 100 m. Inhibitors are added to the culture medium before the MB formation. (B and C) Quantitative analysis of the aggregates projected area (B) and shape index (C) in the presence of the different inhibitors. Red lines represent the mean value for each condition. (D and E) Representative images (D) (contrast is adjusted individually for a better visualization of the pattern; scale bar, 100 m; the images were acquired using a wide-field microscope) and quantitative analysis (E) of the COX-2 fluorescence signal intensity normalized by the control value with the different inhibitors. For these longer culturing times, QNZ and CytoD are only added during the phase change to allow the MB formation. Small dots represent one MB. Large dots represent the average normalized COX-2 fluorescence signal per chip. Each color corresponds to a specific chip. *P < 0.05. (F) Estimation of inhibitor effect in the cell layers with the COX-2 signal normalized by the control value. Control: Nchips = 11 and nMBs = 2,204; QNZ: Nchips = 6 and nMBs = 1215; DAPT: Nchips = 3 and nMBs = 658; Y27: Nchips = 4 and nMBs = 709; CytoD: Nchips = 3 and nMBs = 459. *P < 0.05; **P < 0.01; ***P < 0.001. (G) Proposed mechanisms regulating the MB formation and the patterning of their biological functions. (i) Regulation of the formation of MBs. (ii and iii) Spatial patterning of hMSC biological properties within MBs.

To assess the role of NF-B and actin polymerization in the pattern and the level of COX-2 expression in the MBs, QNZ and CytoD were added 1 day after the cell seeding, once the MBs were completely formed. In contrast, Y27 and DAPT were included in the initial droplets and maintained in the culture medium for the whole culture period. Typical images showing the COX-2 signal in these different conditions are shown in Fig. 6D (see also fig. S7 for quantification of the individual experiments). Of note, none of the inhibitors had an effect on Casp3 activation, indicating that they do not induce apoptosis at the concentration used in this study (fig. S8). The levels of COX-2 expression in MBs, after 3 days in culture, were significantly reduced with QNZ, also decreasing after the addition of CytoD (Fig. 6E). By contrast, Y27 and DAPT had no effect on the levels of COX-2 expression. As a consequence, the results demonstrate that a sustained NF-B activity after the MB formation is required to promote COX-2 expression. Moreover, the induction of actin polymerization in MBs constitutes a mandatory step to initiate COX-2 production.

To get a deeper understanding on the local regulation of these signaling pathways, we analyzed at the single-cell resolution the distribution of COX-2 within the MBs. The spatial mapping revealed that the COX-2 fluorescence intensity was mostly attenuated at the edge of the MBs treated with CytoD and QNZ, while more limited change in the pattern of its expression was observed in the presence of Y27 and even less so with DAPT (Fig. 6F). Consequently, the results revealed a strong link between cell phenotype, the capability to form functional adherens junctions, and the local regulation of NF-B and actin polymerization leading to the increased expression of PGE2 and VEGF that are mediated by COX-2 in 3D (Fig. 6G). Together, the results indicate that in 3D cell aggregates, the spatial organization has some implications on the specific activation of signaling pathways, resulting in local functional heterogeneity.

Understanding the mechanisms of the formation and the spatial tissue patterning within organoids requires a characterization at single-cell level in 3D. In this study, we used a novel microfluidic and epifluorescence imaging technology to obtain a precise quantitative mapping of the structure, the position, and the link with individual cell functions within MBs. The image analysis provided quantitative data that were resolved on the scale of the individual cells, yielding measurements on 700,000 cells in situ within over 10,000 MBs.

While the microfluidic technology developed here is very efficient for high-density size-controlled MB formation, the method is prone to some limitations. Chief among them, the cultivation in nanoliter-scale drops may subject the cells to nutrient deprivation and by-product accumulation under static culture conditions. This limits the duration of the culture to a few days, depending on the cell type and droplet size. To overcome this limitation, it is possible to continuously perfuse the chip with fresh culture medium after performing the oil-aqueous phase exchange, as we demonstrated previously (16). Alternatively, it is also possible to maintain the cells in liquid droplets (without using a hydrogel) by resupplying culture medium through the fusion of additional drops at later times. This operation requires, however, a new design of the anchors and additional microfluidic steps (21).

The second major drawback of the method emerges from the large distance between the MBs and the microscope objective, which requires the use of very large working distance objectives. This compounds the difficulty of applying different confocal techniques by limiting the fluorescence intensity of the images, which, in turn, reduces the throughput when 3D image stacks are required. Although we have shown above that wide-field imaging can be used to obtain spatial mappings of spheroid structure and cell functions, true single-cell measurements will need to overcome the limitations on imaging in the future.

A Voronoi segmentation was used to categorize the cells into concentric layers, starting from the edge of the MBs and ending with the cells in the central region (22), which allowed us to measure variations in the structural organization and in the protein expressions on a layer-by-layer basis within the 3D cultures. The MBs were found to organize into a core region of undifferentiated cells, surrounded by a shell of committed cells. This hierarchical organization results from the spatial segregation of an initially heterogeneous population, as is generally the case for populations of pluripotent and somatic stem cells (2, 3, 33). The process of aggregation of HMSCs obtained within a few hours takes place through different stages (Fig. 6G): The first steps of the aggregation of MBs are mediated by N-cadherin interactions. In parallel, NF-B signaling is activated, promoting cell survival by preventing anoikis of suspended cells (34, 35). At later stages, the formation of polymerized F-actin and, to a lesser extent, stress fibers mediates the MB compaction, mainly at the edge of the MBs where the cellular commitment helps the stabilization of adherens junctions. The formation of adherens junctions facilitates the cohesion of the 3D structure, probably through the enhanced - and -catenin availability in the CD146dim/RUNX-2+ cells (36, 37, 38), which are recruited in the CCC complexes of the adherens junctions to promote the stable coupling of the F-actin to the N-cadherin (39), which become more insoluble to Triton X-100 than unbounded N-cadherins.

A functional phenotype that correlates with this hierarchical segregation is an increase in endocrine activity of the cells located at the boundaries of the MBs. COX-2 expression is increased in the outer layers of the MBs, which also contain more functional adherens junctions as well as a sustained NF-B activity in this region. The promoter of COX-2 contains RUNX-2 and NF-B cis-acting elements (40). While RUNX-2 is required for COX-2 expression in mesenchymal cells, its level of expression does not regulate the levels of COX-2 (40). The increased COX-2 expression is, in turn, due to the unbundled form of F-actin (i.e., a more relaxed form of actin, in comparison to the dense stress fibers observed in 2D) near the edge of the MBs, which was reported to sustain NF-B activity (41) and to down-regulate COX-2 transcriptional repressors (42). Therefore, NF-B has a high activity in the outer layers of the MB, where it locally promotes COX-2 expression.

These results show that the 3D culture format may provide some insights to understand the mesenchymal cell behavior in vivo, because we found that the expression of key bone regulatory molecules is spatially regulated as a function of the structural organization of the MBs. The 3D structure obtained here recalls some of the conditions found at the initial steps of intramembranous ossification that occurs after mesenchymal condensation (i.e., no chondrogenic intermediate was found in the MBs). In the developing calvaria, the most undifferentiated mesenchymal cells (e.g., Sca-1+/RUNX-2 cells) are located in the intrasutural mesenchyme, which is surrounded by an osteogenic front containing more committed cells (e.g., Sca-1/RUNX-2+ cells) (43, 44). Similarly, we observed that undifferentiated HMSCs (i.e., CD146bright/RUNX-2 HMSCs) were surrounded by osteogenically committed cells (i.e., CD146dim/RUNX-2+ HMSCs), which also coexpressed pro-osteogenic molecules, namely, COX-2 and its downstream targets, PGE2 and VEGF. While the link between COX-2 and PGE2 is well established, there is also evidence that COX-2 can induce the production of VEGF in different cell types, e.g., colon cancer cells (45), prostate cancer cells (46), sarcoma (47), pancreatic cancer cells (48), retinal Mller cells (49), gastric fibroblasts (50), skin or lung fibroblasts (51). In these cases, the mechanism for VEGF production through COX-2 induction is thought to be linked to PGE-2, either in an autocrine/paracrine manner (52) or in an intracrine manner (53).

Beyond HMSCs, spatial organization related to the level of differentiation and cell size has been documented in growing embryoids and organoids, with more committed cells being positioned in the outer layers (54, 55, 56). Our results show that a similar hierarchical structure can also be obtained through the aggregation of a mixed population of adult progenitors. This suggests that cell sorting, based on the size and commitment, plays a dominant role in organizing stem cell aggregates. This data-driven approach of combining high-throughput 3D culture and multiscale cytometry (16) on complex biological models can be applied further for getting a better understanding of the equilibria that determine the structure and the function of cells within multicellular tumor spheroids, embryoid bodies, or organoids.

HMSCs derived from the Whartons jelly of the UC (HMSCs) [American Type Culture Collection (ATCC) PCS-500-010, LGC, Molsheim, France] were obtained at passage 2. Four different lots of HMSCs were used in this study (lot nos. 60971574, 63739206, 63516504, and 63739206). While the lots were not selected a priori, we found consistent results for COX-2 and CD146 distribution within MBs. HMSCs from the different lots were certified for being CD29, CD44, CD73, CD90, CD105, and CD166 positive (more than 98% of the population is positive for these markers) and CD14, CD31, CD34, and CD45 negative (less than 0.6% of the population is positive for these markers) and to differentiate into adipocytes, chondrocytes, and osteocytes (ATCC, certificates of analysis). HMSCs were maintained in T175 cm2 flasks (Corning, France) and cultivated in a standard CO2 incubator (Binder, Tuttlingen, Germany). The culture medium was composed of modified Eagles medium (-MEM) (Gibco, Life Technologies, Saint Aubin, France) supplemented with 10% (v/v) fetal bovine serum (FBS) (Gibco) and 1% (v/v) penicilin-streptomycin (Gibco). The cells were seeded at 5 103 cells/cm2, subcultivated every week, and the medium was refreshed every 2 days. HMSCs at passage 2 were first expanded until passage 4 [for about five to six population doublings (PDs)], then cryopreserved in 90% (v/v) FBS/10% (v/v) dimethyl sulfoxide (DMSO), and stored in a liquid nitrogen tank. The experiments were carried out with HMSCs at passages 4 to 11 (about 24 to 35 PDs, after passage 2).

HMSCs were harvested by scrapping or trypsinization from T175 cm2 flasks. Then, the cells were incubated in staining buffer [2% FBS in phosphate-buffered saline (PBS)], stained with a mouse anti-human CD146Alexa Fluor 647 (clone P1-H12, BD Biosciences), a mouse anti-human CD31Alexa Fluor 488 (BD Biosciences, San Jose, CA) antibody, a mouse anti-human CD105Alexa Fluor 647 (BD Biosciences, San Jose, CA), a mouse anti-human CD90fluorescein isothiocyanate (FITC) and a mouse anti-human CD73allophycocyanin (APC) (Miltenyi Biotec, Germany), a CD14-APC (Miltenyi Biotec), a CD34-FITC (BioLegend), and an HLA-DRAPC (BD Biosciences).

The percentages of CD73-, CD90-, CD105-, CD146-, CD31-, CD34-, and HLA-DRpositive cells were analyzed using a FACS LSRFortessa (BD Biosciences, San Jose, CA) or an ImageStream (Amnis) flow cytometer. To validate the specificity of the antibody staining, the distributions of fluorescently labeled cells were compared to cells stained with isotype controls: mouse immunoglobulin G1 (IgG1), k-PE-Cy5 (clone MOPC-21, BD Biosciences), and mouse IgG2a K isotype control FITC (BD Biosciences, San Jose, CA). Alternatively, HMSCs were sorted on the basis of their level of expression of CD146 or their size [forward scatter (FSC) and side scatter (SSC)] using a FACSAria III (BD Biosciences, San Jose, CA).

To induce adipogenic differentiation, UC-HMSCs were seeded at 1 104 cells/cm2 in culture medium. The day after, the culture medium was switched to StemPro Adipogenesis Differentiation medium (Life Technologies) supplemented with 10 M rosiglitazone (Sigma-Aldrich) for 2 weeks. To visualize the differentiated adipocytes, the cells were stained with Oil Red O (Sigma-Aldrich). As a control, UC-HMSCs were maintained in culture medium for 2 weeks and stained with Oil Red O, as above.

To induce osteogenic differentiation, UC-HMSCs were seeded at 5 103 cells/cm2 in culture medium. The day after, the culture medium was switched to StemPro Osteogenesis Differentiation medium (Life Technologies) supplemented with 2-nm bone morphogenetic protein 2 (BMP-2) (Sigma-Aldrich) for 2 weeks. To visualize the differentiated osteoblasts, the cells were stained with Alizarin Red S (Sigma-Aldrich). As a control, UC-HMSCs were maintained in culture medium for 2 weeks and stained with Alizarin Red S, as above.

To induce chondrogenic differentiation, UC-HMSCs were seeded at 1 106 cells/ml in a 15-ml conical tube to promote micromass culture. The medium consisted of StemPro Chondrogenic Differentiation medium (Life Technologies). After 3 weeks in culture, the pellets were fixed and cryosectioned and then stained for Alcian Blue 8GX (Sigma-Aldrich). As a control, UC-HMSCs were maintained in 2D using culture medium for 3 weeks and stained with Alcian Blue, as above.

The color images were acquired using a binocular (SMZ18, Nikon) equipped with a camera (D7500, Nikon).

Standard dry-film soft lithography was used for the flow-focusing device (top of the chip) fabrication, while a specific method for the fabrication of the anchors (bottom of the chip) was developed. For the first part, up to five layers of dry-film photoresist consisting of 50-m Eternal Laminar E8020, 33-m Eternal Laminar E8013 (Eternal Materials, Taiwan), and 15-m Alpho NIT215 (Nichigo-Morton, Japan) negative films were successively laminated using an office laminator (PEAK pro PS320) at a temperature of 100C until the desired channel height, either 135, 150, 165, or 200 m, was reached. The photoresist film was then exposed to ultraviolet (Lightningcure, Hamamatsu, Japan) through a photomask of the junction, the channels, and the culture chamber boundaries. The masters were revealed after washing in a 1% (w/w) K2CO3 solution (Sigma-Aldrich). For the anchor fabrication, the molds were designed with RhinoCAM software (MecSoft Corporation, LA) and were fabricated by micromilling a brass plate (CNCMini-Mill/GX, Minitech Machinery, Norcross). The topography of the molds and masters was measured using an optical profilometer (Veeco Wyco NT1100, Veeco, Mannheim, Germany).

For the fabrication of the top of the chip, poly(dimethylsiloxane) [PDMS; SYLGARD 184, Dow Corning, 1:10 (w/w) ratio of curing agent to bulk material] was poured over the master and cured for 2 hours at 70C. For the fabrication of the bottom of the chip, the molds for the anchors were covered with PDMS. Then, a glass slide was immersed into uncured PDMS, above the anchors. The mold was lastly heated on a hot plate at 180C for 15 min. The top and the bottom of the chip were sealed after plasma treatment (Harrick, Ithaca). The chips were filled three times with Novec Surface Modifier (3M, Paris, France), a fluoropolymer coating agent, for 30 min at 110C on a hot plate.

HMSCs were harvested with TrypLE at 60 to 70% confluence, and a solution containing 6 105 cells in 70 l of medium was mixed with 30 l of a 3% (w/v) liquid low-melting agarose solution (i.e., stored at 37C) (Sigma-Aldrich, Saint Quentin Fallavier, France) diluted in culture medium containing gentamicin (50 g/ml; Sigma-Aldrich) (1:3, v/v), resulting in a 100-l solution of 6 106 cells/ml in 0.9% (w/v) agarose.

HMSCs and agarose were loaded into a 100-l glass syringe (SGE, Analytical Science, France), while Fluorinert FC-40 oil (3M, Paris, France) containing 1% (w/w) PEG-di-Krytox surfactant (RAN Biotechnologies, Beverly, USA) was loaded into a 1- and 2.5-ml glass syringes (SGE, Analytical Science). Droplets of cell-liquid agarose were generated in the FC-40 containing PEG-di-Krytox, at the flow-focusing junction, by controlling the flow rates using syringe pumps (neMESYS Low-Pressure Syringe Pump, Cetoni GmbH, Korbussen, Germany) (table S1). After complete loading, the chips were immersed in PBS, and the cells were allowed to settle down and to organize as MBs overnight in the CO2 incubator. Then, the agarose was gelled at 4C for 30 min, after which the PEG-di-Krytox was extensively washed in flushing pure FC-40 in the culture chamber. After washing, cell culture medium was injected to replace the FC-40. All flow rates are indicated in table S1. Further operations were allowed by gelling the agarose in the droplets, such that the resulting beads were retained mechanically in the traps rather than by capillary forces (Fig. 2G). This step allowed the exchange of the oil surrounding the droplets by an aqueous solution, for example, to bring fresh medium for long-term culture, chemical stimuli, or the different solutions required for cell staining.

For the live imaging of the MB formation, the chips were immersed in PBS and then were incubated for 24 hours in a microscope incubator equipped with temperature, CO2, and hygrometry controllers (Okolab, Pozzuoli, Italy). The cells were imaged every 20 min.

2D cultures or MBs were washed in PBS and incubated with a 5 M NucView 488 caspase-3 substrate (Interchim, Montluon, France) diluted in PBS. After washing with PBS, HMSCs were fixed with a 4% (w/v) PFA (Alpha Aesar, Heysham, UK) for 30 min and permeabilized with 0.2 to 0.5% (v/v) Triton X-100 (Sigma-Aldrich) for 5 min. The samples were blocked with 5% (v/v) FBS in PBS for 30 min and incubated with a rabbit polyclonal antiCOX-2 primary antibody (ab15191, Abcam, Cambridge, UK) diluted at 1:100 in 1% (v/v) FBS for 4 hours. After washing with PBS, the samples were incubated with an Alexa Fluor 594conjugated goat polyclonal anti-rabbit IgG secondary antibody (A-11012, Life Technologies, Saint Aubin, France) diluted at 1:100 in 1% (v/v) FBS for 90 min. Last, the cells were counterstained with 0.2 M DAPI for 5 min (Sigma-Aldrich) and then washed with PBS.

The same protocol was used for the staining of VEGF-Aexpressing cells using a rabbit anti-human VEGF-A monoclonal antibody (ab52917, Abcam, Cambridge, UK), which was revealed using the same secondary antibody as above. RUNX-2positive cells were similarly stained using a mouse anti-human RUNX-2 monoclonal antibody (ab76956, Abcam, Cambridge, UK), which was revealed using an Alexa Fluor 488 goat anti-mouse IgG2a secondary antibody (A-21131, Life Technologies, Saint Aubin, France), both diluted at 1:100 in 1% (v/v) FBS.

To measure potential induction of hypoxia within the core of the MBs, the cells were stained with Image-iT Red Hypoxia Reagent (Invitrogen) for 3 hours and then imaged using a fluorescence microscope. As a positive control, the chips containing the MBs were immersed into PBS, incubated overnight in an incubator set at 37C under 3% O2/5% CO2, and lastly imaged as above.

To interrogate the contribution of signaling related to anti-inflammatory molecule production (COX-2 and NF-B) or molecular pathways regulated by the cell structural organization (Notch, ROCK, and F-actin), several small molecules inducing their inhibition were added to the culture medium (table S1). For all the conditions, the final concentration of DMSO was below 0.1% (v/v) in the culture medium.

The cell viability was assessed using LIVE/DEAD staining kit (Molecular Probes, Life Technologies). The MBs were incubated for 30 min in PBS containing 1 M calcein AM and 2 M ethidium homodimer-1, in flushing 100 l of the solution. The samples were then washed with PBS and imaged under a motorized fluorescence microscope (Nikon, France).

For the detection of the functional forms of N-cadherins (i.e., the N-cadherins closely linked to the actin network, which are PFA insoluble), the MBs were fixed with a 4% (w/v) PFA (Alpha Aesar, Heysham, UK) for 30 min and permeabilized with 0.2 to 0.5% (v/v) Triton X-100 (Sigma-Aldrich) for 5 min. Alternatively, the aggregates were incubated for 5 min with 100% cold methanol followed by 1 min with cold acetone, for the detection of total N-cadherins (i.e., the PFA-soluble and PFA-insoluble forms).

Then, the samples were blocked with 5% (v/v) FBS in PBS for 30 min and incubated with a rabbit polyclonal antiN-cadherin primary antibody (ab18203, Abcam, Cambridge, UK) diluted at 1:100 in 1% (v/v) FBS for 4 hours. After washing with PBS, the samples were incubated with an Alexa Fluor 594conjugated goat polyclonal anti-rabbit IgG secondary antibody (A-11012, Life Technologies, Saint Aubin, France) diluted at 1:100 in 1% (v/v) FBS for 90 min. Last, the cells were counterstained with 0.2 M DAPI for 5 min (Sigma-Aldrich) and then washed with PBS.

For the quantification of the polymerized form of actin (F-actin), the MBs were first fixed with a 4% (w/v) PFA (Alpha Aesar, Heysham, UK) for 30 min and permeabilized with 0.2 to 0.5% (v/v) Triton X-100 (Sigma-Aldrich) for 5 min. The samples were then blocked with a 5% (v/v) FBS solution and incubated for 90 min in a 1:100 phalloidinAlexa Fluor 594 (Life Technologies) diluted in a 1% (v/v) FBS solution. The cells were then counterstained with 0.2 M DAPI for 5 min (Sigma-Aldrich) and then washed with PBS.

To ensure the specificity of the antibody to COX-2 and N-cadherin, control UC-HMSCs were permeabilized, fixed, and incubated only with the secondary antibody (Alexa Fluor 594conjugated goat polyclonal anti-rabbit IgG), as above. The absence of fluorescence signal indicated the specific staining for intracellular COX-2 and N-cadherin.

Next, to validate that the distribution of the fluorescence intensity was not related to any antibody diffusion limitation, the MBs were fixed and permeabilized, as above. For this assay, the MBs were not subjected to any blocking buffer. The cells were incubated for 90 min with the Alexa Fluor 594conjugated goat polyclonal anti-rabbit IgG secondary antibody (A-11012, Life Technologies, Saint Aubin, France) diluted at 1:100 in 1% (v/v) FBS. Then, the cells were counterstained for DAPI, as above. Last, the MBs were collected from the chip, deposed on a glass slide, and imaged.

For the analysis of COX-2 expression by flow cytometry, the total MBs were recovered from the chip. The MBs were then trypsinized and triturated to obtain single-cell suspension. UC-HMSCs were stained for COX-2, as above. The percentage of COX-2positive cells was quantified on 5 103 dissociated UC-HMSCs using a Guava easyCyte Flow Cytometer (Merck Millipore, Guyancourt, France). The results were compared to the fluorescence intensity distribution obtained by image analysis.

To interrogate the influence of the MB opacity in the COX-2 and N-cadherin fluorescence signals, the samples were treated by the Clear(T2) method after immunostaining (57). Briefly, the MBs were incubated for 10 min in 25% (v/v) formamide/10% (w/v) polyethylene glycol (PEG) (Sigma-Aldrich), then for 5 min in 50% (v/v) formamide/20% (w/v) PEG, and lastly for 60 min in 50% (v/v) formamide/20% (w/v) PEG, before their imaging. The fluorescence signal distribution was compared with the noncleared samples.

The MBs were collected from the chip and then fixed using PFA, as above. The MBs were incubated overnight in a 30% sucrose solution at 4C. Then, the sucrose solution was exchanged to O.C.T. medium (optimal cutting temperature; Tissue-Tek) in inclusion molds, which were slowly cooled down using dry ice in ethanol. The molds were then placed at 80C. On the day of the experiments, the O.C.T. blocks were cut at 7 m using a cryostat (CM3050 S, Leica). The cryosections were placed on glass slides (SuperFrost Plus Adhesion, Thermo Fisher Scientific), dried at 37C, and rehydrated using PBS. The cryosections were permeabilized and stained for COX-2, as above. The slides were lastly mounted in mounting medium containing DAPI (Fluoromount-G, Invitrogen).

All the images used for the quantitative analysis were taken using a motorized wide-field microscope (Ti, Eclipse, Nikon), equipped with a CMOS (complementary metal-oxide semiconductor) camera (ORCA-Flash4.0, Hamamatsu) and a fluorescence light-emitting diode source (Spectra X, Lumencor). The images were taken with a 10 objective with a 4-mm working distance (extra-long working distance) and a 0.45 numerical aperture (NA) (Plan Apo , Nikon).

For control experiments, images were taken using a motorized (Ti2, Nikon) confocal spinning disc microscope equipped with lasers (W1, Yokogawa) and the same camera and objective as above. Alternatively, the samples were imaged with a multiphoton microscope (TCS SP8 NLO, MP, Leica). The objective was an HCX PL APO CS 10, 0.40 NA, working distance of 2.2 mm (Leica).

All immunostained samples were counterstained with DAPI, and most of the images (i.e., for N-cadherin, COX-2, VEGF-A, and F-actin) were taken using red light excitation that is known to penetrate deeper into the 3D objects than dyes emitting at lower weight length (e.g., DAPI, FITC). For wide-field microscopy, the focal plane was defined as the area containing the maximal number of DAPI-stained nuclei covering the focal area, while z stacks were taken for the whole in-focus planes containing DAPI-stained nuclei using spinning discs and two-photon confocal microscopy.

Wide-field imaging is sensitive for the emission of fluorescence from inside and outside the focal plane (i.e., from the out-of-focus upper and bottom planes of the spheroids) (58). Consistently, more DAPI signal from nuclei is emitted from the core than in the edges of MBs using epifluorescence microscopy (fig. S5I). We confirmed that our interpretation of the signal distribution from epifluorescence images was consistent with confocal and two-photon microscopy by comparing with images taken from the median z plane and the maximal z projection (fig. S5, N to P).

Consequently, the results unambiguously demonstrate that even if there are more cells in the z plane of the middle area of the MBs, the contribution of the out-of-focus signal from N-cadherin, COX-2, VEGF-A, and F-actin staining in this area of the MBs is minimal using wide-field imaging. Because of the higher throughput of wide-field microscopy, this method was chosen to quantitatively analyze the distribution of these immunolabeled proteins within MBs.

The culture supernatants of six-well plates were collected, while the total medium content of the chip was recovered by flushing the culture chamber with pure oil. A PGE2 human ELISA kit (ab133055, Abcam, Cambridge, UK) was used for the quantification of PGE2 concentration in the culture supernatant, following the manufacturers instructions. Briefly, a polynomial standard curve of PGE2 concentration derived from the serial dilution of a PGE2 standard solution was generated (r2 > 0.9). The absorbance was measured using a plate reader (Chameleon, Hidex, Finland).

A VEGF-A human ELISA kit (Ab119566, Abcam, Cambridge, UK) was used for the quantification of VEGF-A concentration in the culture supernatant of 2D cultures or from the chips. A linear standard curve of VEGF-A concentration derived from the serial dilution of a VEGF-A standard solution was generated (r2 > 0.9). The absorbance was measured using a plate reader (Chameleon, Hidex, Finland).

The total MBs of a 3-day culture period were harvested from the chips, as described above. Alternatively, cells cultured on regular six-well plates were recovered using trypsin after the same cultivation time; CD146dim and CD146bright isolated cells were immediately treated for RNA extraction after sorting. The total RNA of 1 104 cells were extracted and converted to complementary DNA (cDNA) using SuperScript III CellsDirect cDNA Synthesis System (18080200, Invitrogen, Life Technologies), following the manufacturers instructions. After cell lysis, a comparable quality of the extracted RNA was observed using a bleach agarose gel, and similar RNA purity was obtained by measurement of the optical density at 260 and 280 nm using a NanoDrop spectrophotometer (Thermo Fisher Scientific, Wilmington, DE) between total RNA preparations from 2D and on-chip cultures.

The cDNA was amplified using a GoTaq qPCR Master Mix (Promega, Charbonnieres, France) or a FastStart Universal SYBR Green Master Mix (containing Rox) (Roche) and primers (Life Technologies, Saint Aubin, France or Eurofins Scientific, France) at the specified melting temperature (Tm) (table S2) using a MiniOpticon (Bio-Rad) or a QuantStudio 3 (Thermo Fisher Scientific) thermocycler. As a negative control, water and total RNA served as template for PCR. To validate the specificity of the PCR, the amplicons were analyzed by dissociation curve and subsequent loading on a 2.5% (w/v) agarose gel and migration at 100 V for 40 min. The PCR products were revealed by ethidium bromide (Sigma-Aldrich) staining, and the gels were imaged using a transilluminator. The analysis of the samples not subjected to reverse transcription (RT) indicated negligible genomic DNA contamination (i.e., <0.1%), while no amplification signal was observed for the water template (no template control). The amount of TSG-6, COX-2, STC-1, VEGF-A, RUNX-2, CEBP-, and SOX-9 transcripts was normalized to the endogenous reference [glyceraldehyde-3-phosphate dehydrogenase (GADPH)], and the relative expression to a calibrator (2D cultures) was given by 2Ct calculation. At least five biological replicates of 2D and on-chip cultures were analyzed by at least duplicate measurements. The standard curves for GADPH, TSG-6, COX-2, and STC-1 were performed using a five serial dilution of the cDNA templates and indicated almost 100% PCR efficiency.

The image analysis allowed us to perform a multiscale analysis (16) of the MBs. For each chip, single images of the anchors were acquired automatically with the motorized stage of the microscope. The analysis was conducted on a montage of the detected anchors using a custom MATLAB code (r2016a, MathWorks, Natick, MA). Two distinct routines were used: one with bright-field detection and one for the fluorescence experiments.

For the bright field-detection described previously (16), the cells were detected in each anchor as pixels with high values of the intensity gradient. This allowed for each cell aggregate to compute morphological parameters such as the projected area A and the shape index SI that quantifies the circularity of an objectSI=4APwhere P is the perimeter. Shape index values range from 0 to 1, with 1 being assigned for perfect disk.

The MB detection with fluorescence staining (DAPI/Casp3/COX-2, DAPI/phalloidin, DAPI/N-cadherin, or LIVE/DEAD) was performed as described previously (16). First, morphological data were extracted at the MB level, such as the equivalent diameter of the MBs or the shape index. Also, the mean fluorescence signal of each MB was defined as the subtraction of the local background from the mean raw intensity.

At the cellular level, two different methods were used, both relying on the detection of the nuclei centers with the DAPI fluorescence signal. On the one hand, each cell location could be assigned to a normalized distance from the MB center (r/R) to correlate a nuclear fluorescence signal with a position in the MB, as previously described (16). On the other hand, the cell shapes inside the MBs were approximated by constructing Voronoi diagrams on the detected nuclei centers. Basically, the edges of the Voronoi cells are formed by the perpendicular bisectors of the segments between the neighboring cell centers. These Voronoi cells were used to quantify the cellular cytoplasmic signal (COX-2, F-actin and N-cadherin, VEGF and RUNX-2). In detail, to account for the variability of the cytoplasmic signal across the entire cell (nucleus included), the fluorescence signal of a single cell was defined as the mean signal of the 10% highest pixels of the corresponding Voronoi cell.

Image processing was also used to get quantitative data on 2D cultures, as previously described (16). Last, different normalization procedures were chosen in this paper. When an effect was quantified compared with a control condition, the test values were divided by the mean control value, and the significance was tested against 1. For some other data, the values were simply normalized by the corresponding mean at the chip level to discard the interchip variation from the analysis.

*P < 0.05; **P < 0.01; ***P < 0.001; NS, nonsignificant. Details of each statistical test and P values can be found in table S4.

Acknowledgments: C. Frot is gratefully acknowledged for the help with the microfabrication, and F. Soares da Silva is gratefully acknowledged for the help in flow cytometry. The group of Biomaterials and Microfluidics (BMCF) of the Center for Innovation and Technological Research as well as the Center for Translational Science (CRT)Cytometry and Biomarkers Unit of Technology and Service (CB UTechS is also acknowledged for the access to the microfabrication and flow cytometry platform at the Institut Pasteur). Funding: The research leading to these results received funding from the European Research Council (ERC) grant agreement 278248 Multicell. Author contributions: S.S., C.N.B., and A.C. conceived the experiments. S.S. performed the experiments. R.F.-X.T. wrote the image processing code and performed the image analysis. R.F.-X.T., S.S., G.A., and A.B. performed the image and data analyses. S.S., C.N.B., and A.C. discussed the results and wrote the manuscript. All authors discussed the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors.

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Mapping the structure and biological functions within mesenchymal bodies using microfluidics - Science Advances