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Scaffold-free cell-based tissue engineering therapies: advances, shortfalls and forecast

Andrea De Pieri, Yury Rochev, Dimitrios I. Zeugolis

npj Regenerative Medicine · 2021 · Review · CC BY

doi:10.1038/s41536-021-00133-3

Abstract

Cell-based scaffold-free therapies seek to develop in vitro organotypic three-dimensional (3D) tissue-like surrogates, capitalising upon the inherent capacity of cells to create tissues with efficiency and sophistication that is still unparalleled by human-made devices. Although automation systems have been realised and (some) success stories have been witnessed over the years in clinical and commercial arenas, in vitro organogenesis is far from becoming a standard way of care. This limited technology transfer is largely attributed to scalability-associated costs, considering that the development of a borderline 3D implantable device requires very high number of functional cells and prolonged ex vivo culture periods. Herein, we critically discuss advancements and shortfalls of scaffold-free cell-based tissue engineering strategies, along with pioneering concepts that have the potential to transform regenerative and reparative medicine.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 22, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

Cell-based therapy has gained tremendous interest in the past decades and holds promise for transforming treatments for a wide range of injuries and diseases. The market for cell therapy products is set to expand based on increasing investment from the industry and the implementation of advanced manufacturing technologies. In fact, the global cell therapy market is forecast to reach €7.24 billion by 2025 with compound annual growth rate of 14.9% from 2017, which would make it the fastest growing sector in the regenerative medicine industry.

Cells have enormous therapeutic potential, as they provide sophisticated tissue-specific mechanisms of actions that chemical compounds cannot imitate. Mesenchymal stem cells (MSCs), permanently differentiated cells and, more recently, induced pluripotent stem cells (iPSCs) have been used in preclinical and clinical trials with successful outcomes–. One critical aspect for therapeutic efficacy after cell transplantation is the delivery method (Fig. 1). The optimal cell delivery format should ensure high cell retention and survival rate, good tissue integration and zero to low side effects for the patient. Intra-venous/intra-arterial infusion or direct intra-tissue injection are the most common routes of cell transplantation. However, these approaches have shown limited success, mainly due to the poor cell localisation, retention and survival at the site of injury post-transplantation. In fact, numerous studies have shown that <5% of the injected cells persist at the site of injection in the first day(s) after transplantation, indicating a survival rate of as low as 1% (refs. –).

Scaffold-based tissue engineering was pioneered to overcome the limitations of direct cell suspensions, aiming not only to develop efficient cell delivery strategies, but also to produce elegant three-dimensional (3D) tissue analogues. Traditional scaffold-based tissue engineering strategies employ a cytocompatible, biodegradable and mechanically stable natural or synthetic in origin polymeric scaffold with a fully interconnected porous network for efficient transport and exchange of oxygen, nutrients and metabolites,. Although very many scaffold conformations (e.g., hydrogels,, sponges,, fibres,, films,) have been developed, and have demonstrated safety and efficacy in preclinical setting and phase I clinical trials as cell delivery vehicles, only a handful of them constitute a Food and Drug Administration (FDA)/European Medicines Agency (EMA) approved device (Table 1). This limited technology transfer from laboratory benchtop to clinical applicability has been attributed to component (e.g., limited understanding of the mechanism of action of the various device components; device components do not comply with regulatory frameworks; toxicity issues) and process (e.g., too complex to allow for large-scale efficient and reproducible manufacturing; too long to be profitable) limitations.

Considering that tissues are formed by cells and their secreted components with precision, efficiency, order and sophistication that is still unmatched by human-made devices, it made sense to develop means to exploit this inherent capacity of cells for the development of tissue analogues. In this case, the cell-secreted extracellular matrix (ECM) acts as carrier and protector of the transplanted cells. Further, as no artificial scaffold is used, the produced constructs are of superior biocompatibility and with less chances of foreign body response than any other technology that has been assessed to-date. Although the scaffold-free tissue engineering concept is far from new (the first scaffold-free device was developed in 1975 (ref. ), assessed in preclinical models in 1980 (ref. ) and assessed in humans in 1981 (ref. )), only a handful of products have been commercialised (Table 2). Herein, we critically discuss recent advancements and limitations that prohibit wide acceptance, clinical translation and commercialisation of scaffold-free cell-based tissue engineering strategies.

Cell sheet tissue engineering

Tissues and organs are comprised of different cell types that are surrounded by their secreted, tissue-specific ECM. This densely populated microenvironment allows for efficient cell–cell and cell–ECM communications, which determine cell fate and function,. Cell sheet tissue engineering takes advantage of the close cell–cell and cel–ECM interactions to autonomously engineer microtissues, utilising the temperature-responsive cell culture technology. The temperature-responsive polymer used [usually poly(N-isopropylacrylamide) (pNIPAM), although a variety of polymers with diverse properties have been developed and assessed over the years (Table 3)] undergoes a transition from hydrophobic to hydrophilic across its lower critical solution temperature (LCST) of 32 °C. At temperatures >32 °C, the surfaces are hydrophobic and allow for the culture of adherent cells as on normal tissue culture polystyrene at 37 °C. As the cells grow, they deposit ECM proteins that assemble into interconnected tissue-like structures. When the temperature is reduced <32 °C, pNIPAm molecules become highly hydrated, thus the pNIPAM grafted surfaces become hydrophilic. After this thermal transition, cultured cells almost spontaneously detach from the pNIPAm surface as a contiguous cell sheet with preserved cell–cell junctions and deposited ECM,. Since ECM proteins remain on the surface of the cell sheets, they are adhesive to biological surfaces and therefore can be transplanted to injured tissues without the need of sutures or external fixation. The deposited ECM also acts as a depot of numerous bioactive and trophic molecules, and also protects and localises the transplanted cells at the site of implantation–. This unique approach, with or without the use of a temperature-responsive polymer, has been used to develop implantable devices out of numerous human cells, and their safety and efficacy has been demonstrated in preclinical and clinical setting for a diverse range of clinical indications (Table 4).

Over the years, significant strides have been achieved in developing tissue analogues, with high levels of architectural biomimicry. For example, recapitulation of native anisotropic tissue topographies has been realised via the use of bi-directionally aligned temperature-responsive electrospun scaffolds, microcontact printing of aligned fibronectin patterns,, photolithography on non-cell adhesive anisotropic patterns–, grafting of temperature-responsive polymers onto micropatterned poly(dimethysiloxane) substrates, or unidirectional mechanical stimulation. With respect to the development of 3D tissue-like assemblies, multi-layered cell sheet stacking has been proposed, which has resulted in the formation of sophisticated microtissues in vitro (e.g., skeletal muscle-like tissue out of myoblasts,, myocardial-like tissue out of cardiomyocytes, annulus fibrosus-like tissue out of bone marrow MSCs, tubular neural-like tissue out of astrocytes and iPSC-derived neurons).

To further de-risk the technology and increase its scalability, automated systems have also been developed. For example, automated technologies have been utilised for the production of multi-layered tissue constructs, using robotic systems. Specifically, five layers of human skeletal muscle myoblast sheets were successfully stacked by a robotic apparatus within 100 minu, representing a cost-effective manufacturing system for the manipulation of cell sheets. Automated modular platforms have also been assembled for the sequential seeding, expansion and cell (e.g., skeletal myoblasts, articular chondrocytes and iPSCs) sheet preparation that have been shown to maintain aseptic conditions and to produce high quality cellular constructs, comparable to those produced in manual operations. Automated systems have been further advanced for the production of high number of cell sheet batches. For instance, 10 human oral mucosa epithelial cell sheets were simultaneously cultured into 5 separate fully closed culture vessels, automated with a circuit system, that yielded up to 50 cell sheets and satisfied the quality standards of manual procedures.

Considering that the thickness limitation of 3D constructs without vascular networks is ~40–80 μm (ref. ), co-culture cell sheet approaches have been proposed for the development of pre-vascularised networks,. For example, human umbilical vein endothelial cells co-cultured within human myoblasts sheets formed capillary-like structures within the construct and increased neovascularisation and graft survival after transplantation into the subcutaneous tissues of nude rats. However, in order to support the long-term culture of thick 3D tissue equivalents, the formation of functional mature blood vessels is required. To this end, bioreactor systems have been utilised in combination with engineered vascular beds, based on resected femoral muscles or synthetic collagen gels, which allowed continuous perfusion of culture media, formation of a functional vasculature and survival of 12-layer cell sheets,.

For the development of tubular tissues, such as blood vessels, tendons and neurons, cell and deposited ECM layers are rolled into tubular structures. Automated systems have also been designed, which utilise a wrapping device module composed of a fibrin tube holder and a sliding dish, capable of rolling three layers of cardiomyocyte sheets without any deformation. A pioneering study reported the fabrication of human biological tissue-engineered blood vessel composed of smooth muscle cells and skin fibroblast sheets, which were mechanically peeled off from the culture dishes and rolled onto a supportive tubular mandrel (polytetrafluoroethylene). After maturation (8 weeks under dynamic conditions) of the construct into a cohesive vascular structure, the supportive mandrel was removed and endothelial cells were seeded in the lumen, ultimately creating a well‐defined, three‐layered device with abundant ECM. The major limitation of this technology was the time required to produce functional grafts (~28 weeks), which resulted in the company closing down. Advanced anisotropic tubular assemblies have also been realised, using surface patterning to culture human astrocytes and human iPSC‐derived neurons, to create neural microtissues.

Limitations and way forward

Despite the significant advances that have been achieved in the field of scaffold-free tissue engineering, the technology is far for optimal, as evidenced by the limited number of clinically and commercially available products. Several interconnected and interdependent limitations must be addressed for this technology to become clinical standard.

Transitioning from 2D to 3D systems

The limiting factor in clinical translation and commercialisation of scaffold-free concepts is the high number of cells required to produce in a commercially relevant timeframe an ECM-rich and truly 3D tissue equivalent (Fig. 2A). Indeed, temperature-responsive surface derived single-layer cell sheets, as well as more elegant micro-stereolithography and electrochemical desorption for cell transfer systems,, require a substantial cell number and/or culture time to produce a barely 3D scaffold-free construct (Table 5). For example 104,000 cells/cm2 human endometrial gland-derived MSCs produced a 50 µm thick cell sheet after 5 days, 300,000/cm2 iPSCs-derived cardiomyocytes produced a 10 µm thick cell sheet after 27 days, 612,000/cm2 human corneal endothelial cells produced a 15 µm thick cell sheet after 28 days. To overcome these limitations, multi-layer cell sheet stacking has been proposed. Regrettably, even this approach has been found wanting, as again very high cell numbers and relatively long culture times are required to develop borderline 3D implantable devices (Table 6). For instance, five layers of 1,000,000/cm2 human skeletal muscle myoblasts grown on a temperature-responsive dish for 5 days produced a 50 μm thick device, three layers of 300,000/cm2 human adipose-derived MSCs grown on a temperature-responsive dish for 5 days produced a 20 μm thick device, nine layers of 200,000/cm2 human iPSCs-derived cardiomyocytes per layer grown on a temperature-responsive dish for 7–10 days produced a 359 μm thick device. Unfortunately, due to the absence of sufficient ECM, these high-density cultures are associated with poor nutrient and oxygen diffusion and waste accumulation in the middle layers that ultimately lead to cell necrosis, and delamination (Fig. 2B). To address these issues, multiple operations of up to three layers have been proposed; however, these multiple operations are associated with prolonged patient distress and high healthcare expenditure. To overcome the dimensionality limitation of scaffold-free systems, cell sheets (whole or in fragments) have been combined with various scaffold conformations (e.g., hydrogels, particles, tissue grafts)/fabrication systems (e.g., electrospinning,, 3D printing,), which have been summarised elsewhere recently.

Hypoxia

Considering the importance of ECM in regulating cell survival and tissue homoeostasis,, it is imperative to develop means to accelerate ECM synthesis and deposition. Physiological hypoxia (<10% O2) poses a biochemical cue crucial for the regulation of ECM synthesis and deposition. Indeed, hypoxia has been shown to increase mRNA levels of procollagen α1(I) in fibroblasts isolated from different tissues–. Hypoxia also regulates ECM homoeostasis through the activation of hypoxia-inducible transcription factor 1 (HIF-1). HIF-1 regulates collagen secretion and deposition by driving the transcription of prolyl 4-hydroxylase, which catalyses intracellularly the hydroxylation of proline and lysine residues and lysyl oxidase, which catalyses extracellularly collagen crosslinking–. Hypoxia therefore can be an ally in the fabrication of biomimetic tissue-engineered constructs.

Considerable efforts have been conducted to optimise oxygen levels of cultured cells, in order to control (stem) cell fate and promote ECM synthesis and deposition for the desired tissue engineering application (Table 7), such as skin, cartilage,, bone,, tendon, and heart. For example, low oxygen tension (5% O2) has been shown to retain undifferentiated and multipotent status of MSC cultures,. In addition, hypoxia modulates the paracrine activity of MSCs and enhances the secretion of soluble growth factors, especially pro-angiogenic factors, such as vascular endothelial growth factor (VEGF). In scaffold-free tissue engineering, only few studies have utilised low oxygen tension for the production of implantable cell constructs. For example, multi-layered human chondrocyte sheets fabricated in a co‐culture system with synoviocytes and cultured at 2% oxygen tension showed greater cell metabolic activity and proliferation compared to cells cultured at 21% oxygen tension. Furthermore, hypoxic conditions accelerated and enhanced the deposition of cartilage-specific ECM, mainly composed of proteoglycans and collagen type II (ref. ). Preconditioning of rabbit BMSCs or mouse cardiosphere-derived cell sheets under 2% oxygen tension, remarkably increased the expression of VEGF and significantly improved left ventricular function in myocardial infarction models in comparison to cells cultured under normoxia condition,.

Despite the importance of hypoxia in eukaryotic cell culture and in the development of functional cell therapies, routinely cell culture studies are performed under hyperoxic conditions (21% O2) that do not match physiological oxygen levels (e.g., 5–13% in blood, 2–9% in most tissues). Further, hyperoxic cell cultures lead to poor and slow ECM synthesis, cellular senescence and activation of stress pathways–. Although hypoxia chambers and incubators are utilised to perform hypoxic experiments, implementation of physioxia in industrial scale is expensive to purchase and maintain, thus of limited applicability despite the fact that studies have argued that hypoxia precondition should be a prerequisite for clinical translation of cell therapies,.

Mechanical stimulation

Another microenvironmental cue critical for optimal ECM synthesis and deposition is mechanical loading. Considering that uniaxial or multiaxial tensile, compressive or shear mechanical loads regulate ECM composition and function and tissue homoeostasis, mechanical stimulation of tissue-engineered constructs, with the use of bioreactors, is attracting growing interest in order to recapitulate the in vivo microenvironment of native (primarily musculoskeletal) tissues in in vitro setting (Table 8). For example, mechanical stimulation, in the form of shear force, hydrostatic pressure or compression, has been shown to promote ECM synthesis in human chondrocyte and MSC cultures, and to produce tissue‐engineered cartilaginous substitutes–. Similarly, in tendon engineering, mechanical loading has been shown to maintain tenocyte phenotype and to increase their proliferation and ECM synthesis,, and to direct MSCs towards tenogenic lineage–. In bone engineering, mechanical loading has been employed extensively to enhance mineralised matrix synthesis and deposition,.

With regards to scaffold-free tissue engineering, unidirectional stretching has been applied to induce the alignment of human iPSCs-derived cardiomyocyte sheets after detachment from temperature-responsive dishes. Two weeks after transplantation into the superficial gluteal muscle of athymic rats, the stretched cell sheets retained the unidirectionality of their myocardial fibres, holding great potential for heart engineering. While the application of mechanical loading has not been applied extensively in temperature-responsive systems, it has been successfully implemented in other scaffold-free tissue-engineered models. For example, cartilage constructs have been produced using high-density porcine chondrocytes, centrifugation and non-adhesive agarose substrates. Simultaneous application of cyclic unconfined compression and perfusion to the cartilage constructs increased the deposition of glycosaminoglycans and collagen type II in comparison to static control groups. Another study reported the fabrication of scaffold-free cartilage by applying high-amplitude compressive strain, using porcine chondrocyte seeded onto a hydroxyapatite carrier. Compression amplitude of 20% had the highest positive effect by inducing the synthesis of cartilage-specific ECM and enhancing the mechanical properties of the constructs.

An overwhelming amount of literature has demonstrated the positive effects of mechanical stimulation and/or conditioning on advanced tissue-engineered constructs. Nevertheless, the limited fundamental understanding of the molecular and cellular mechanisms, the lack of standardised protocols for mechanical stimulation and the very high costs of bioreactor systems, limit their scalability and use in commercial space.

Macromolecular crowding

Although various in vitro microenvironment modulators have been assessed over the years to control cell fate during in vitro culture, only marginally enhance and accelerate ECM synthesis and deposition. For example, mechanical stimulation and oxygen tension have been shown to increase by 2–5-fold ECM synthesis, and deposition both in permanently differentiated and stem cell cultures–, which although mathematically may be considered as an improvement, commercially, the associated expenditure does not justify the change in the process.

In recent years, macromolecular crowding (MMC) has emerged as a means to substantially increase and accelerate ECM deposition in vitro (e.g., up to 120-fold increase in collagen and associated ECM deposition within 4–6 days in differentiated– and stem– cell cultures). In tissues, the presence of numerous macromolecules, such as carbohydrates, proteins, lipids and nucleic acids, creates a crowded or confined microenvironment that affects the rate of biological and biochemical reactions,. The MMC’s mechanism of action is based on the theory of mutual excluded volume effect, which refers to the volume that is inaccessible in the system to new molecules as a result of pre-existing molecules; two molecules cannot be at the same place at the same time. Although the effect of MMC on protein folding and assembly–, DNA condensation and replication– and biochemical reactions, has been well established, it is still under investigation in cell culture context. Nonetheless, it is accepted now that in eukaryotic cell culture scenario, MMC accelerates the enzymatic processing of procollagen to collagen, resulting in enhanced collagen, and bound ECM, deposition (Fig. 3). Indeed, in standard cell culture setting, the conversion of water-soluble procollagen to insoluble collagen is relatively slow, since the proteinases required for the enzymatic cleavage of procollagen are dispersed in the dilute culture media. The addition of macromolecules to the culture media results in a more efficient volume occupancy, preventing their dispersion. MMC has been shown to drive the molecular assembly of collagen fibrils in vitro and to stabilise the formed matrix through enzymatic crosslinking,.

Several macromolecules, alone or in cocktail form, have been utilised as crowders to enhance and accelerate ECM deposition (Table 9). It has been demonstrated that negative charge and polydispersity are key regulators for ECM deposition. Indeed, negatively charged crowders cause a stronger volume-excluding effect due to electrostatic repulsion and polydispersity, indicative of the heterogeneity of sizes and/or shapes of molecules in a mixture, maximises the excluded volume effect through reduction in diffusion. These prompted the use mixed crowding molecules systems to achieve higher volume exclusion effect, reduced procollagen/proteinases diffusion and ultimately enhanced and accelerated ECM deposition. For instance, a cocktail of Ficoll™ 70 kDa/Ficoll™ 400 kDa/Ficoll™ 1000 kDa has been shown to lead to higher ECM deposition in dermal fibroblast cultures than the traditionally used Ficoll™ 70 kDa/Ficoll™ 400 kDa cocktail. Carrageenan, a naturally polydisperse and negatively sulfated polysaccharide has been shown to induce the highest volume exclusion effect, as judged by the highest and fastest ECM deposition in vitro,,. Although in traditional protein assembly investigations, for simplicity purposes, crowders are considered as inert macromolecules, eukaryotic cell culture experiments indicate that the chemistry of the crowder affects cell phenotype. For example, in corneal fibroblasts, dextran sulfate-induced myofibroblast trans-differentiation, while the Ficoll™ 70 kDa/400 kDa cocktail and carrageenan maintained their phenotype. Further, in stem cell cultures, non-sulfated polysaccharides have been shown to induce adipogenesis,,, while sulfated polysaccharides have been shown to induce chondrogenesis and osteogenesis,,.

In the field of scaffold-free tissue engineering, MMC has advanced the production of ECM-rich cell sheets, showing the possibility to dramatically speed up the production of implantable tissue equivalents,,. Interestingly, these studies demonstrated that the commercially available pNIPAM-based culture dishes were not able to induce detachment of intact cell sheets, due to the presence of abundant ECM produced under MMC conditions. Co-polymerisation of pNIPAM with the hydrophobic N-tert-butylacrylamide (NTBA) monomer, at an optimal ratio of 35%, allowed for first time the production of dense and cohesive cell sheets with intact cell–cell and cell–ECM junctions. The efficiency of the pNIPAM-NTBA copolymer to produce such ECM-rich construct was attributed to additional steric hindrance induced by the NTBA group, which decreased hydrogen bonding and consequently decreased protein adsorption, ultimately facilitating cell detachment. In addition, MMC has been used for the development of scaffold-free cell-derived matrices,, that have been used successfully for in vitro cell propagation– and for the generation of skin equivalents with complete stratification and a mature dermal–epidermal junction for either regenerative medicine, or drug discovery purposes–. To further boost the development of scaffold-free tissue equivalents, multifactorial approaches combining MMC and different in vitro microenvironment modulators have been explored. For example, the use of MMC in combination with low oxygen tension synergistically contributed to the development of ECM-rich tissue equivalents,,,. Similarly, the use of MMC simultaneously to mechanical stimulation facilitated the fabrication of tendon-like tissue constructs in vitro.

Despite the significant contribution and potential of MMC in tissue engineering, the optimal (with respect to maximum ECM deposition in the shortest period of time, while precisely controlling cell fate) crowding agent/cocktail remains elusive.

Roadmap to commercialisation

Although significant strides that have been achieved in the production of ECM-rich tissue equivalents within commercially relevant timeframes, a major roadblock in the commercialisation of cell-based strategies is the high costs of manufacturing solutions, which need to comply with current good manufacturing practices. A recent study analysed eight case studies in Europe using both autologous and allogeneic therapies (academic and other small-scale enterprises scale) and estimated manufacturing costs (i.e., materials, equipment, personnel and facility running costs) to be in the range of € 23,033 and € 190,799 per batch, with batch yield varying between 1 and 88 doses. With regards to scaffold-based strategies, another study reported the costs of stem cell-engineered airway transplants to range from US$ 174,420 to US$ 740,500 in three UK patients. It is worth noting that such regenerative medicine strategies are also associated with several risks (e.g., human errors, batch-to-batch variability, high risk of contamination), which further limit their availability. In order to overcome these limitations, automated platforms could significantly reduce the cost of goods up to 30% (ref. ), ensuring reliability and reproducibility across the life cycle of the product, from cell isolation and expansion to in‐line product quality assurance,. To further standardise the manufacturing process, the development of xeno-free, chemically defined media has been advocated to reduce the risk of pathogen sources and simplify the regulatory approval,.

Besides reducing the costs associated to manufacturing systems, it is also imperative to consider from outset both regulatory hurdles and reimbursement concepts to successfully translate cell-based concepts into blockbuster therapies. In fact, one should consider that no Advanced Therapy Medicinal Products (ATMPs) has yet achieved widespread reimbursement and access across the five biggest European countries (Germany, France, UK, Italy, Spain), and many have been restricted beyond the regulatory label. Both public and private healthcare systems are still under-equipped to absorb the high financial implications associated with cell therapies, also considering additional costs related to expensive hospital stays, procedures and rehabilitation, which come on top of the product price. Another factor which limits cellular therapies to secure reimbursement following market authorisation is the lack of comparative effectiveness data. Manufacturers must demonstrate that the product has incremental benefit against existing standard of care, not only from an economical perspective, but, most importantly, from a clinical one, ensuring long-term safety and efficacy. For example, ChondroCelect®, an autologous chondrocyte therapy licensed for the treatment of single symptomatic cartilage defects of the femoral condyle of the knee, despite being the first cell-based regenerative therapy to obtain a centralised marketing authorisation in Europe, has been withdrawn from the market, failing to established robust clinical efficacy and consequently not fulfilling reimbursement criteria.

Considering the unpredictability around the long-term efficacy and safety of regenerative medicines, new models for financing and reimbursement have been proposed to ensure patient access to such therapies. For instance, annuity or instalment payment models minimise high up-front single payment, allowing healthcare providers to amortise the cost of therapies over multiple years, and recognises the potential of single-administered cell therapies based on evidence that the treatment continues to be effective over a specified period of time. Another example are pay-for-performance models, where the reimbursement for a treatment depends on whether a specified clinical outcome is achieved. These models aim to partially shift financial risks from payers to manufacturers, which may prompt the interest of healthcare providers.

Overall, cell manufacturers need to address many challenges during product development, in order to gain approval from regulatory authorities and ensure that the proposed therapy will not be held back by reimbursement policies.

Conclusions

In vitro scaffold-free organogenesis has come long way since early 1980’s that was first appeared in the literature. Despite though the significant strides in in vitro, preclinical and clinical setting, only a handful of concepts have become clinical and commercial reality. Issues associated with required number of functional cells, dimensionality, production timeframe, automation, scalability (that affect reimbursement) and regulatory requirements/classification must be addressed for wide acceptance of this transformative and disruptive concept.

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    Dang JM, Leong KW. Myogenic induction of aligned mesenchymal stem cell sheets by culture on thermally responsive electrospun nanofibers. Adv. Mater. 2007;19:2775-2779.

    Not in our library10.1002/adma.200602159

  36. 36.

    Elloumi Hannachi I, et al. Fabrication of transferable micropatterned-co-cultured cell sheets with microcontact printing. Biomaterials. 2009;30:5427-5432.

    Not in our library10.1016/j.biomaterials.2009.06.033

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    Williams C, et al. Aligned cell sheets grown on thermo-responsive substrates with microcontact printed protein patterns. Adv. Mater. 2009;21:2161-2164.

    Not in our library10.1002/adma.200801027

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    Takahashi H, Nakayama M, Itoga K, Yamato M, Okano T. Micropatterned thermoresponsive polymer brush surfaces for fabricating cell sheets with well-controlled orientational structures. Biomacromolecules. 2011;12:1414-1418.

    Not in our library10.1021/bm2000956

  39. 39.

    Takahashi H, Shimizu T, Nakayama M, Yamato M, Okano T. Anisotropic cellular network formation in engineered muscle tissue through the self-organization of neurons and endothelial cells. Adv. Health. Mater. 2015;4:356-360.

    Not in our library10.1002/adhm.201400297

  40. 40.

    Kumashiro Y, et al. Rate control of cell sheet recovery by incorporating hydrophilic pattern in thermoresponsive cell culture dish. J. Biomed. Mater. Res A. 2014;102:2849-2856.

    Not in our library10.1002/jbm.a.34959

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    Backman, D. E., LeSavage, B. L., Shah, S. B. & Wong, J. Y. A Robust Method to Generate Mechanically Anisotropic Vascular Smooth Muscle Cell Sheets for Vascular Tissue Engineering. 17 (2017). 10.1002/mabi.201600434.

    Not in our library10.1002/mabi.201600434

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    Lin JB, et al. Thermo-responsive poly(N-isopropylacrylamide) grafted onto microtextured poly(dimethylsiloxane) for aligned cell sheet engineering. Colloids Surf. B Biointerfaces. 2012;99:108-115.

    Not in our library10.1016/j.colsurfb.2011.10.040

  43. 43.

    Homma J, Shimizu S, Sekine H, Matsuura K, Shimizu T. A novel method to align cells in a cardiac tissue-like construct fabricated by cell sheet-based tissue engineering. J. Tissue Eng. Regen. Med.–. 2020;14:944-954.

    Not in our library10.1002/term.3074

  44. 44.

    Takahashi H, Shimizu T, Nakayama M, Yamato M, Okano T. The use of anisotropic cell sheets to control orientation during the self-organization of 3D muscle tissue. Biomaterials. 2013;34:7372-7380.

    Not in our library10.1016/j.biomaterials.2013.06.033

  45. 45.

    Jiao A, et al. Regulation of skeletal myotube formation and alignment by nanotopographically controlled cell-secreted extracellular matrix. J. Biomed. Mater. Res A. 2018;106:1543-1551.

    Not in our library10.1002/jbm.a.36351

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    Haraguchi Y, Shimizu T, Yamato M, Kikuchi A, Okano T. Electrical coupling of cardiomyocyte sheets occurs rapidly via functional gap junction formation. Biomaterials. 2006;27:4765-4774.

    Not in our library10.1016/j.biomaterials.2006.04.034

  47. 47.

    Chuah YJ, et al. Scaffold-free tissue engineering with aligned bone marrow stromal cell sheets to recapitulate the microstructural and biochemical composition of annulus fibrosus. Acta Biomater. 2020;107:129-137.

    Not in our library10.1016/j.actbio.2020.02.031

  48. 48.

    Takahashi H, Itoga K, Shimizu T, Yamato M, Okano T. Human neural tissue construct fabrication based on scaffold-free tissue engineering. Adv. Health. Mater. 2016;5:1931-1938.

    Not in our library10.1002/adhm.201600197

  49. 49.

    Kikuchi T, Shimizu T, Wada M, Yamato M, Okano T. Automatic fabrication of 3-dimensional tissues using cell sheet manipulator technique. Biomaterials. 2014;35:2428-2435.

    Not in our library10.1016/j.biomaterials.2013.12.014

  50. 50.

    Kikuchi T, et al. A novel, flexible and automated manufacturing facility for cell-based health care products: tissue. Fact. Regen. Ther. 2018;9:89-99.

    Not in our library10.1016/j.reth.2018.08.004

  51. 51.

    Nishimura A, et al. Fabrication of tissue-engineered cell sheets by automated cell culture equipment. J. Tissue Eng. Regen. Med. 2019;13:2246-2255.

    Not in our library10.1002/term.2968

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    Shimizu T, et al. Polysurgery of cell sheet grafts overcomes diffusion limits to produce thick, vascularized myocardial tissues. FASEB J. 2006;20:708-710.

    Not in our library10.1096/fj.05-4715fje

  53. 53.

    Novosel EC, Kleinhans C, Kluger PJ. Vascularization is the key challenge in tissue engineering. Adv. Drug Deliv. Rev. 2011;63:300-311.

    Not in our library10.1016/j.addr.2011.03.004

  54. 54.

    Shimizu T. Cell sheet-based tissue engineering for fabricating 3-dimensional heart tissues. Circ. J. 2014;78:2594-2603.

    Not in our library10.1253/circj.cj-14-0973

  55. 55.

    Sasagawa T, et al. Design of prevascularized three-dimensional cell-dense tissues using a cell sheet stacking manipulation technology. Biomaterials. 2010;31:1646-1654.

    Not in our library10.1016/j.biomaterials.2009.11.036

  56. 56.

    Sekine H, et al. In vitro fabrication of functional three-dimensional tissues with perfusable blood vessels. Nat. Commun. 2013;4:1399.

    Not in our library10.1038/ncomms2406

  57. 57.
    In Vitro Engineering of Vascularized Tissue Surrogates · 2013
    No notice
  58. 58.

    L’Heureux N, Pâquet S, Labbé R, Germain L, Auger FA. A completely biological tissue-engineered human blood vessel. FASEB J. 1998;12:47-56.

    Not in our library10.1096/fasebj.12.1.47

  59. 59.

    Ni M, et al. Engineered scaffold-free tendon tissue produced by tendon-derived stem cells. Biomaterials. 2013;34:2024-2037.

    Not in our library10.1016/j.biomaterials.2012.11.046

  60. 60.

    Kubo H, Shimizu T, Yamato M, Fujimoto T, Okano T. Creation of myocardial tubes using cardiomyocyte sheets and an in vitro cell sheet-wrapping device. Biomaterials. 2007;28:3508-3516.

    Not in our library10.1016/j.biomaterials.2007.04.016

  61. 61.

    L’Heureux N, et al. Human tissue-engineered blood vessels for adult arterial revascularization. Nat. Med. 2006;12:361-365.

    Not in our library10.1038/nm1364

  62. 62.

    Enomoto J, et al. Engineering thick cell sheets by electrochemical desorption of oligopeptides on membrane substrates. Regen. Ther. 2016;3:24-31.

    Not in our library10.1016/j.reth.2015.12.003

  63. 63.

    Kobayashi Y, et al. Tailored cell sheet engineering using microstereolithography and electrochemical cell transfer. Sci. Rep. 2019;9:10415.

    Not in our library10.1038/s41598-019-46801-9

  64. 64.

    Sekine W, et al. Chondrocyte differentiation of human endometrial gland-derived MSCs in layered cell sheets. ScientificWorldJournal. 2013;2013:359109.

    Not in our library10.1155/2013/359109

  65. 65.

    Ishigami M, et al. Human iPS cell-derived cardiac tissue sheets for functional restoration of infarcted porcine hearts. PLoS ONE. 2018;13:e0201650.

    Not in our library10.1371/journal.pone.0201650

  66. 66.

    Sumide T, et al. Functional human corneal endothelial cell sheets harvested from temperature-responsive culture surfaces. FASEB J. 2006;20:392-394.

    Not in our library10.1096/fj.04-3035fje

  67. 67.

    Cerqueira MT, et al. Human adipose stem cells cell sheet constructs impact epidermal morphogenesis in full-thickness excisional wounds. Biomacromolecules. 2013;14:3997-4008.

    Not in our library10.1021/bm4011062

  68. 68.

    Komae H, et al. Three-dimensional functional human myocardial tissues fabricated from induced pluripotent stem cells. J. Tissue Eng. Regen. Med. 2017;11:926-935.

    Not in our library10.1002/term.1995

  69. 69.

    Takahashi H, Nakayama M, Shimizu T, Yamato M, Okano T. Anisotropic cell sheets for constructing three-dimensional tissue with well-organized cell orientation. Biomaterials. 2011;32:8830-8838.

    Not in our library10.1016/j.biomaterials.2011.08.006

  70. 70.

    Sekine W, Haraguchi Y, Shimizu T, Umezawa A, Okano T. Thickness limitation and cell viability of multi-layered cell sheets and overcoming the diffusion limit by a porous-membrane culture insert. J. Biochip Tissue Chip. 2011;S1:007.

    Not in our library

  71. 71.

    Haraguchi Y, et al. Thicker three-dimensional tissue from a “symbiotic recycling system” combining mammalian cells and algae. Sci. Rep. 2017;7:41594.

    Not in our library10.1038/srep41594

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    Kasai Y, Takeda N, Kobayashi S, Takagi R, Yamato M. Cellular events and behaviors after grafting of stratified squamous epithelial cell sheet onto a hydrated collagen gel. FEBS Open Bio. 2017;7:691-704.

    Not in our library10.1002/2211-5463.12213

  73. 73.

    Ma G, et al. Scaffold-based delivery of bone marrow mesenchymal stem cell sheet fragments enhances new bone formation in vivo. J. Oral. Maxillofac. Surg. 2017;75:92-104.

    Not in our library10.1016/j.joms.2016.08.014

  74. 74.
    Human Mesenchymal Stromal Cell Sheet Enhances Allograft Repair in a Mouse Model · 2017
    Not checked
  75. 75.

    Vaquette C, Saifzadeh S, Farag A, Hutmacher D, Ivanovski S. Periodontal tissue engineering with a multiphasic construct and cell sheets. J. Dent. Res. 2019;98:673-681.

    Not in our library10.1177/0022034519837967

  76. 76.
    A simvastatin-releasing scaffold with periodontal ligament stem cell sheets for periodontal regeneration · 2020
    Not checked
  77. 77.

    Bakirci E, Toprakhisar B, Zeybek M, Ince G, Koc B. Cell sheet based bioink for 3D bioprinting applications. Biofabrication. 2017;9:024105.

    Not in our library10.1088/1758-5090/aa764f

  78. 78.

    Cochis A, et al. 3D printing of thermo-responsive methylcellulose hydrogels for cell-sheet engineering. Materials. 2018;11:579.

    Not in our library10.3390/ma11040579

  79. 79.

    Zurina I, et al. Tissue engineering using a combined cell sheet technology and scaffolding approach. Acta Biomater. 2020;113:63-83.

    Not in our library10.1016/j.actbio.2020.06.016

  80. 80.

    Cox T, Erler J. Remodeling and homeostasis of the extracellular matrix: implications for fibrotic diseases and cancer. Dis. Model Mech. 2011;4:165-178.

    Not in our library10.1242/dmm.004077

  81. 81.

    Buchheit C, Weigel K, Schafer Z. Cancer cell survival during detachment from the ECM: multiple barriers to tumour progression. Nat. Rev. Cancer. 2014;14:632-641.

    Not in our library10.1038/nrc3789

  82. 82.

    Falanga V, et al. Low oxygen tension increases mRNA levels of alpha 1 (I) procollagen in human dermal fibroblasts. J. Cell Physiol. 1993;157:408-412.

    Not in our library10.1002/jcp.1041570225

  83. 83.

    Falanga V, et al. Hypoxia upregulates the synthesis of TGF-β1 by human dermal fibroblasts. J. Investig. Dermatol. 1991;97:634-637.

    Not in our library10.1111/1523-1747.ep12483126

  84. 84.

    Norman JT, Clark IM, Garcia PL. Hypoxia promotes fibrogenesis in human renal fibroblasts. Kidney Int. 2000;58:2351-2366.

    Not in our library10.1046/j.1523-1755.2000.00419.x

  85. 85.

    Tamamori M, Ito H, Hiroe M, Marumo F, Hata RI. Stimulation of collagen synthesis in rat cardiac fibroblasts by exposure to hypoxic culture conditions and suppression of the effect by natriuretic peptides. Cell Biol. Int. 1997;21:175-180.

    Not in our library10.1006/cbir.1997.0130

  86. 86.

    Weidemann A, Johnson RS. Biology of HIF-1α. Cell Death Differ. 2008;15:621-627.

    Not in our library10.1038/cdd.2008.12

  87. 87.

    Bentovim L, Amarilio R, Zelzer E. HIF1α is a central regulator of collagen hydroxylation and secretion under hypoxia during bone development. Development. 2012;139:4473-4483.

    Not in our library10.1242/dev.083881

  88. 88.

    Gilkes DM, Bajpai S, Chaturvedi P, Wirtz D, Semenza GL. Hypoxia-inducible factor 1 (HIF-1) promotes extracellular matrix remodeling under hypoxic conditions by inducing P4HA1, P4HA2, and PLOD2 expression in fibroblasts. J. Biol. Chem. 2013;288:10819-10829.

    Not in our library10.1074/jbc.m112.442939

  89. 89.

    van Vlimmeren MA, Driessen-Mol A, van den Broek M, Bouten CV, Baaijens FP. Controlling matrix formation and cross-linking by hypoxia in cardiovascular tissue engineering. J. Appl Physiol. (1985). 2010;109:1483-1491.

    Not in our library10.1152/japplphysiol.00571.2010

  90. 90.

    Erler JT, et al. Hypoxia-induced lysyl oxidase is a critical mediator of bone marrow cell recruitment to form the premetastatic niche. Cancer Cell. 2009;15:35-44.

    Not in our library10.1016/j.ccr.2008.11.012

  91. 91.

    Mohyeldin A, Garzón-Muvdi T, Quiñones-Hinojosa A. Oxygen in stem cell biology: a critical component of the stem cell niche. Cell Stem Cell. 2010;7:150-161.

    Not in our library10.1016/j.stem.2010.07.007

  92. 92.

    Mieremet A, et al. Human skin equivalents cultured under hypoxia display enhanced epidermal morphogenesis and lipid barrier formation. Sci. Rep. 2019;9:7811.

    Not in our library10.1038/s41598-019-44204-4

  93. 93.

    Duval E, et al. Molecular mechanism of hypoxia-induced chondrogenesis and its application in in vivo cartilage tissue engineering. Biomaterials. 2012;33:6042-6051.

    Not in our library10.1016/j.biomaterials.2012.04.061

  94. 94.

    Markway BD, Cho H, Johnstone B. Hypoxia promotes redifferentiation and suppresses markers of hypertrophy and degeneration in both healthy and osteoarthritic chondrocytes. Arthritis Res. Ther. 2013;15:R92.

    Not in our library10.1186/ar4272

  95. 95.

    Stiers P-J, van Gastel N, Carmeliet G. Targeting the hypoxic response in bone tissue engineering: a balance between supply and consumption to improve bone regeneration. Mol. Cell Endocrinol. 2016;432:96-105.

    Not in our library10.1016/j.mce.2015.12.024

  96. 96.

    Zhou Y, et al. Hypoxia induces osteogenic/angiogenic responses of bone marrow-derived mesenchymal stromal cells seeded on bone-derived scaffolds via ERK1/2 and p38 pathways. Biotechnol. Bioeng. 2013;110:1794-1804.

    Not in our library10.1002/bit.24827

  97. 97.

    Lee WY, Lui PP, Rui YF. Hypoxia-mediated efficient expansion of human tendon-derived stem cells in vitro. Tissue Eng. Part A. 2012;18:484-498.

    Not in our library10.1089/ten.tea.2011.0130

  98. 98.

    Zhang J, Wang JH. Human tendon stem cells better maintain their stemness in hypoxic culture conditions. PLoS ONE. 2013;8:e61424.

    Not in our library10.1371/journal.pone.0061424

  99. 99.

    Fehrer C, et al. Reduced oxygen tension attenuates differentiation capacity of human mesenchymal stem cells and prolongs their lifespan. Aging Cell. 2007;6:745-757.

    Not in our library10.1111/j.1474-9726.2007.00336.x

  100. 100.
    Long term culture of mesenchymal stem cells in hypoxia promotes a genetic program maintaining their undifferentiated and multipotent status · 2011
    No notice
  101. 101.

    Gaspar D, et al. Local pharmacological induction of angiogenesis: drugs for cells and cells as drugs. Adv. Drug Deliv. Rev. 2019;146:126-154.

    Not in our library10.1016/j.addr.2019.06.002

  102. 102.

    Kokubo M, et al. Characterization of layered chondrocyte sheets created in a co-culture system with synoviocytes in a hypoxic environment. J. Tissue Eng. Regen. Med. 2017;11:2885-2894.

    Not in our library10.1002/term.2192

  103. 103.

    Tanaka Y, et al. Autologous preconditioned mesenchymal stem cell sheets improve left ventricular function in a rabbit old myocardial infarction model. Am. J. Transl. Res. 2016;8:2222-2233.

    Not in our library

  104. 104.

    Hosoyama T, et al. Cardiosphere-derived cell sheet primed with hypoxia improves left ventricular function of chronically infarcted heart. Am. J. Transl. Res. 2015;7:2738-2751.

    Not in our library

  105. 105.

    Brahimi-Horn MC, Pouysségur J. Oxygen, a source of life and stress. FEBS Lett. 2007;581:3582-3591.

    Not in our library10.1016/j.febslet.2007.06.018

  106. 106.

    Parrinello S, et al. Oxygen sensitivity severely limits the replicative lifespan of murine fibroblasts. Nat. Cell Biol. 2003;5:741-747.

    Not in our library10.1038/ncb1024

  107. 107.

    Pomatto LCD, et al. Limitations to adaptive homeostasis in an hyperoxia-induced model of accelerated ageing. Redox Biol. 2019;24:101194.

    Not in our library10.1016/j.redox.2019.101194

  108. 108.

    Lee PJ, Choi AM. Pathways of cell signaling in hyperoxia. Free Radic. Biol. Med. 2003;35:341-350.

    Not in our library10.1016/s0891-5849(03)00279-x

  109. 109.

    Huang YC, Parolini O, Deng L, Yu BS. Should hypoxia preconditioning become the standardized procedure for bone marrow MSCs preparation for clinical use? Stem Cells. 2016;34:1992-1993.

    Not in our library10.1002/stem.2389

  110. 110.

    Tsiapalis D, Zeugolis DI. Hypoxia preconditioning of bone marrow mesenchymal stem cells before implantation in orthopaedics. J. Am. Acad. Orthop. Surg. 2019;27:e1040-e1042.

    Not in our library10.5435/jaaos-d-19-00044

  111. 111.

    Peroglio M, Gaspar D, Zeugolis DI, Alini M. Relevance of bioreactors and whole tissue cultures for the translation of new therapies to humans. J. Orthop. Res. 2018;36:10-21.

    Not in our library10.1002/jor.23655

  112. 112.

    Humphrey JD, Dufresne ER, Schwartz MA. Mechanotransduction and extracellular matrix homeostasis. Nat. Rev. Mol. Cell Biol. 2014;15:802-812.

    Not in our library10.1038/nrm3896

  113. 113.

    Jeon JE, et al. Effect of preculture and loading on expression of matrix molecules, matrix metalloproteinases, and cytokines by expanded osteoarthritic chondrocytes. Arthritis Rheum. 2013;65:2356-2367.

    Not in our library10.1002/art.38049

  114. 114.

    Meinert C, Schrobback K, Hutmacher DW, Klein TJ. A novel bioreactor system for biaxial mechanical loading enhances the properties of tissue-engineered human cartilage. Sci. Rep. 2017;7:16997.

    Not in our library10.1038/s41598-017-16523-x

  115. 115.

    Mouw JK, Connelly JT, Wilson CG, Michael KE, Levenston ME. Dynamic compression regulates the expression and synthesis of chondrocyte-specific matrix molecules in bone marrow stromal cells. Stem Cells. 2007;25:655-663.

    Not in our library10.1634/stemcells.2006-0435

  116. 116.

    Screen HRC, Shelton JC, Bader DL, Lee DA. Cyclic tensile strain upregulates collagen synthesis in isolated tendon fascicles. Biochem. Biophys. Res. Commun. 2005;336:424-429.

    Not in our library10.1016/j.bbrc.2005.08.102

  117. 117.

    Huisman E, Lu A, McCormack RG, Scott A. Enhanced collagen type I synthesis by human tenocytes subjected to periodic in vitro mechanical stimulation. BMC Musculoskelet. Disord. 2014;15:386.

    Not in our library10.1186/1471-2474-15-386

  118. 118.

    Youngstrom DW, LaDow JE, Barrett JG. Tenogenesis of bone marrow-, adipose-, and tendon-derived stem cells in a dynamic bioreactor. Connect Tissue Res. 2016;57:454-465.

    Not in our library10.3109/03008207.2015.1117458

  119. 119.

    Kuo CK, Tuan RS. Mechanoactive tenogenic differentiation of human mesenchymal stem cells. Tissue Eng. Part A. 2008;14:1615-1627.

    Not in our library10.1089/ten.tea.2006.0415

  120. 120.

    Thomopoulos S, et al. Fibrocartilage tissue engineering: the role of the stress environment on cell morphology and matrix expression. Tissue Eng. Part A. 2011;17:1039-1053.

    Not in our library10.1089/ten.tea.2009.0499

  121. 121.

    Fernandez-Yague MA, et al. Biomimetic approaches in bone tissue engineering: integrating biological and physicomechanical strategies. Adv. Drug Deliv. Rev. 2015;84:1-29.

    Not in our library10.1016/j.addr.2014.09.005

  122. 122.

    Ng J, Spiller K, Bernhard J, Vunjak-Novakovic G. Biomimetic approaches for bone tissue engineering. Tissue Eng. Part B Rev. 2017;23:480-493.

    Not in our library10.1089/ten.teb.2016.0289

  123. 123.

    Tran SC, Cooley AJ, Elder SH. Effect of a mechanical stimulation bioreactor on tissue engineered, scaffold-free cartilage. Biotechnol. Bioeng. 2011;108:1421-1429.

    Not in our library10.1002/bit.23061

  124. 124.

    Hoenig E, et al. High amplitude direct compressive strain enhances mechanical properties of scaffold-free tissue-engineered cartilage. Tissue Eng. Part A. 2011;17:1401-1411.

    Not in our library10.1089/ten.tea.2010.0395

  125. 125.

    Ku CH, et al. Collagen synthesis by mesenchymal stem cells and aortic valve interstitial cells in response to mechanical stretch. Cardiovasc Res. 2006;71:548-556.

    Not in our library10.1016/j.cardiores.2006.03.022

  126. 126.

    Waldman SD, Couto DC, Grynpas MD, Pilliar RM, Kandel RA. A single application of cyclic loading can accelerate matrix deposition and enhance the properties of tissue-engineered cartilage. Osteoarthr. Cartil. 2006;14:323-330.

    Not in our library10.1016/j.joca.2005.10.007

  127. 127.

    Lee A, et al. Hypoxia modulates the development of a corneal stromal matrix model. Exp. Eye Res. 2018;170:127-137.

    Not in our library10.1016/j.exer.2018.02.021

  128. 128.

    Falanga V, Zhou L, Yufit T. Low oxygen tension stimulates collagen synthesis and COL1A1 transcription through the action of TGF-β1. J. Cell Physiol. 2002;191:42-50.

    Not in our library10.1002/jcp.10065

  129. 129.

    Gaspar D, Fuller KP, Zeugolis DI. Polydispersity and negative charge are key modulators of extracellular matrix deposition under macromolecular crowding conditions. Acta Biomater. 2019;88:197-210.

    Not in our library10.1016/j.actbio.2019.02.050

  130. 130.

    Kumar P, et al. Macromolecularly crowded in vitro microenvironments accelerate the production of extracellular matrix-rich supramolecular assemblies. Sci. Rep. 2015;5:8729.

    Not in our library10.1038/srep08729

  131. 131.

    Satyam A, et al. Macromolecular crowding meets tissue engineering by self-assembly: a paradigm shift in regenerative medicine. Adv. Mater. 2014;26:3024-3034.

    Not in our library10.1002/adma.201304428

  132. 132.

    Lareu RR, et al. Collagen matrix deposition is dramatically enhanced in vitro when crowded with charged macromolecules: the biological relevance of the excluded volume effect. FEBS Lett. 2007;581:2709-2714.

    Not in our library10.1016/j.febslet.2007.05.020

  133. 133.

    Shendi D, et al. Hyaluronic acid as a macromolecular crowding agent for production of cell-derived matrices. Acta Biomater. 2019;100:292-305.

    Not in our library10.1016/j.actbio.2019.09.042

  134. 134.

    Graceffa V, Zeugolis DI. Carrageenan enhances chondrogenesis and osteogenesis in human bone marrow stem cell culture. Eur. Cell Mater. 2019;37:310-332.

    Not in our library10.22203/ecm.v037a19

  135. 135.

    Patrikoski M, et al. Effects of macromolecular crowding on human adipose stem cell culture in fetal bovine serum, human serum, and defined xeno-free/serum-free conditions. Stem Cells Int. 2017;2017:6909163.

    Not in our library10.1155/2017/6909163

  136. 136.

    Cigognini D, et al. Macromolecular crowding meets oxygen tension in human mesenchymal stem cell culture - a step closer to physiologically relevant in vitro organogenesis. Sci. Rep. 2016;6:30746.

    Not in our library10.1038/srep30746

  137. 137.

    Lee MH, et al. ECM microenvironment unlocks brown adipogenic potential of adult human bone marrow-derived MSCs. Sci. Rep. 2016;6:21173.

    Not in our library10.1038/srep21173

  138. 138.

    Ma L, et al. Comparative proteomic analysis of extracellular matrix proteins secreted by hypertrophic scar with normal skin fibroblasts. Burns Trauma. 2014;2:76-83.

    Not in our library10.4103/2321-3868.130191

  139. 139.

    Zeiger AS, Loe FC, Li R, Raghunath M, Van Vliet KJ. Macromolecular crowding directs extracellular matrix organization and mesenchymal stem cell behavior. PLoS ONE. 2012;7:e37904.

    Not in our library10.1371/journal.pone.0037904

  140. 140.

    De Pieri A, Rana S, Korntner S, Zeugolis DI. Seaweed polysaccharides as macromolecular crowding agents. Int. J. Biol. Macromol. 2020;164:434-446.

    Not in our library10.1016/j.ijbiomac.2020.07.087

  141. 141.

    Prewitz MC, et al. Extracellular matrix deposition of bone marrow stroma enhanced by macromolecular crowding. Biomaterials. 2015;73:60-69.

    Not in our library10.1016/j.biomaterials.2015.09.014

  142. 142.

    Minton AP. The influence of macromolecular crowding and macromolecular confinement on biochemical reactions in physiological media. J. Biol. Chem. 2001;276:10577-10580.

    Not in our library10.1074/jbc.r100005200

  143. 143.

    Zhou H-X, Rivas G, Minton AP. Macromolecular crowding and confinement: biochemical, biophysical, and potential physiological consequences. Annu. Rev. Biophys. 2008;37:375-397.

    Not in our library10.1146/annurev.biophys.37.032807.125817

  144. 144.

    Minton AP. Excluded volume as a determinant of macromolecular structure and reactivity. Biopolymers. 1981;20:2093-2120.

    Not in our library10.1002/bip.1981.360201006

  145. 145.

    Hatters DM, Minton AP, Howlett GJ. Macromolecular crowding accelerates amyloid formation by human apolipoprotein C-II. J. Biol. Chem. 2002;277:7824-7830.

    Not in our library10.1074/jbc.m110429200

  146. 146.

    van den Berg B, Ellis RJ, Dobson CM. Effects of macromolecular crowding on protein folding and aggregation. EMBO J. 1999;18:6927-6933.

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