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Abstract

Objectives

Although bone marrow mesenchymal stromal cells (BMSCs) are extensively used in biomedicine, they have yet to be used in the commercial development of a tissue engineered medicine. It has been argued that the major roadblock in their commercial deployment is the lengthy in vitro culture periods required for the development of implantable tissue surrogates. Macromolecular crowding (MMC) has been shown to enhance and increase extracellular matrix deposition in eukaryotic cell culture, allowing for the accelerated development of tissue facsimiles.

Methods

With these in mind, human BMSCs were cultured under MMC conditions and the developed tissue-engineered medicine was assessed in vitro and in vivo in a humanised athymic nude mouse excisional wound splinting model.

Results

Starting with basic cell function analysis, MMC did not significantly affect cell metabolic activity, viability and proliferation. Electrophoresis and immunofluorescence analyses revealed that MMC significantly increased collagen type I and collagen type IV deposition, without significantly affecting collagen type III deposition. Flow cytometry analysis demonstrated similar CD44, CD73, CD90, CD146, HLA-ABC, CD31, CD45, CD80 and CD86 expression between the without and the with MMC groups. Interestingly though the MMC group had higher CD105 and lower HLA-DR expression than the without MMC group. Preclinical analysis revealed similar wound closure, scar index and epidermal thickness between the without and the with MMC groups, largely attributed to issues encountered with the model.

Conclusions

Overall, this preliminary study demonstrates the potential of MMC in the accelerated development of functional and extracellular matrix-rich human BMSC-based tissue-engineered medicines.

Keywords

Advanced therapy medicinal products bone marrow mesenchymal stromal cells macromolecular crowding wound healing

Introduction

Bone marrow mesenchymal stromal cells (BMSCs) are by far the most studied adult stromal cell population in medicine (Source: PubMed; Term searched: bone marrow cells, 318,130 results, as opposed to 82,467 results for Term searched: adipose cells; Date: 27/09/2024), with numerous ongoing clinical trials (Source: clinicaltrials.gov; Term searched in Intervention/treatment: bone marrow cells, 2195 results, as opposed to 861 results for Term searched in Intervention/treatment: adipose cells; Date: 27/09/2024) for a diverse range of clinical indications. One should also note that their safety, efficacy and efficiency have been well-documented in the literature, in both preclinical and clinical settings for a diverse range of dermatological conditions. For example, in preclinical setting, intradermal injections have shown promise in full-thickness incisional acute cutaneous wounds in pigs¹ and in full-thickness burn wounds in rats.² Using a collagen sponge as a carrier, transplanted cells resulted in regeneration of subcutaneous tissue in nude mice and skin healing in 18 patients with intractable dermatopathies.³ In a randomised, blinded placebo-controlled trial, intradermal transplantation of autologous BMSCs was proven to be a safe and effective strategy to promote mechanical stretch induced skin regeneration.⁴ In a case report, where a decellularised allogeneic dermal matrix / autologous BMSCs / autologous split-skin graft was compared to a decellularised allogeneic dermal matrix / autologous split-skin graft, the BMSC transplant resulted in significantly less contraction in a patient who was severely burned.⁵ In another case study, allogenic BMSCs were shown to accelerate recovery of homeostasis and to promote healing of a patient with deep skin burns.⁶ Despite all these positive patient therapeutic outcomes, no BMSC-based therapy is currently available in the wound healing space.

Extracellular matrix (ECM) is a depot of bioactive molecules that through a spatiotemporal dynamic crosstalk with surrounding cells promotes tissue remodelling and wound healing post injury.^7–12 Commercially available living skin substitutes require prolonged ex vivo culture for ECM deposition (e.g. Apligraf^® 17–20 days,¹³ denovoSkin™ 21–31 days¹⁴). To avoid phenotypic drift, BMSC-based medicines are transplanted immediately after cell expansion without any cell deposited ECM. In the absence of cell deposited ECM, the secretome, and therefore, the therapeutic potential, of the BMSCs is not fully exploited, rendering the potential therapy inferior and more expensive to the state-of-the-art. To substantiate this one should consider that adipose derived MSC secretome has been shown to lead to diabetic wound healing in rats by modulating angiogenesis, scarring and inflammation.¹⁵ In a full-thickness rat skin excision model although both adipose-derived MSCs and adipose derived MSCs’ secretome enhanced early phase wound healing, the secretome group promoted fibroblast proliferation and migration and suppressed inflammation.¹⁶ In a diabetic rat splinted wound healing model, human foetal MSC secretome, delivered through poly lactic-co-glycolic acid particles, significantly enhanced wound healing via improved vascularisation and suppressed inflammation.¹⁷ In clinical setting, application of cultured (scraped cell layers as opposed to injected cell suspensions) autologous BMSCs resulted in complete healing and dermal rebuilding in three patients with chronic wounds that had not closed even after multiple applications of Apligraf^® or autologous skin grafting.¹⁸ It is evidenced that for superior patient therapeutic outcomes, the MSC secretome must be part of the intervention strategy. Upon this supposition, the concept of utilising macromolecular crowding (MMC) in tissue engineering was conceived.

In eukaryotic cell culture, MMC is defined as the addition of macromolecules in the culture media that by decreasing diffusion via restriction of molecular motion, favour protein–substrate interactions and result in enhanced and accelerated ECM deposition.^19–21 To-date, various MMC agents (e.g. carrageenan,^22–25 dextran sulphate,²⁶ polyvinylpyrrolidone,²⁷ polysucrose cocktails,^28–32 polysucrose / dextran sulphate cocktail³³) have been assessed in human BMSC cultures. Although MMC has shown promise in preclinical setting (one study with human adipose derived MSCs cultured with carrageenan,³⁴ one study with decellularised matrices derived from human BMSCs cultured with carrageenan³⁵ and one more study with decellularised matrices derived from human umbilical cord blood MSCs cultured with polysucrose³⁶), not one study has assessed in vivo a tissue-engineered medicine produced with human BMSCs under MMC conditions.

Herein, we ventured to assess the potential of a tissue-engineered medicine, composed of human BMSCs that were cultured under MMC conditions in wound healing context. We selected carrageenan as MMC agent, as, due to its negative charge and high polydispersity, it induces the highest ECM deposition in the shortest period of time.^37,38 A humanised athymic nude mouse excisional wound splinting model was used, as its suitability for human MSC transplantation is well-documented in the literature.^39–42

Materials and methods

Tissue culture consumables were purchased from Sarstedt (Ireland) and NUNC (Denmark). All chemicals, cell culture media and reagents were purchased from Sigma-Aldrich (Ireland), unless otherwise stated.

Cell isolation and culture

Fresh human bone marrow from the iliac crest was purchased from Lonza (UK) and human BMSCs were isolated and cultured in basal medium [α-minimal essential medium, GlutaMax™, ThermoFisher Scientific, Ireland) supplemented with 10% foetal bovine serum (FBS) and 1% penicillin / streptomycin], following established protocols.^22,24,30 A total of 50,000 human BMSCs per cm² were seeded at passage 4 in basal medium and after 24 h, the media were changed to media with 100 µM of l-ascorbic acid 2-phosphate sesquimagnesium salt hydrate without (-) / with (+) MMC (carrageenan at 75 µg/mL). The cells were cultured for 5 days for in vitro analysis and prior to in vivo transplantation as cell sheets.

Cell metabolic activity, viability and proliferation analyses

Using established protocols,^43–45 after 5 days of culture BMSCs-MMC and BMSCs + MMC were assessed for metabolic activity using the alamarBlue™ (Invitrogen, UK) assay; for viability using the Live/Dead™ assay with calcein AM (ThermoFisher Scientific, Ireland) and ethidium homodimer I (ThermoFisher Scientific, Ireland) stainings; and for proliferation using 4′,6-diamidino-2-phenylindole (DAPI, Invitrogen, Ireland) stained cell layers. Images were captured with an Olympus IX-81 (Olympus Corporation, Japan) inverted fluorescence microscope and processed with ImageJ (NIH, USA).

Collagen deposition analysis via electrophoresis and immunofluorescence

Collagen type I deposition at day 5 was assessed using non-reduced sodium dodecyl sulphate – polyacrylamide gel electrophoresis (SDS-PAGE), as per established protocol.⁴⁶ In brief, 100 μg/mL collagen type I standard (Symatese Biomateriaux, France) in pepsin in acetic acid (standard) and cell layers in pepsin in acetic acid were neutralised, denatured, loaded and separated on a Mini-Protean^® 3 system (Bio-Rad Laboratories, UK). Protein bands were stained with SilverQuest™ kit (ThermoFisher Scientific, UK) according to the manufacturer's protocol. Using ImageJ (NIH, USA), the densities of collagen α1(I) and collagen α2(I) chains of the cell layers were quantified and correlated to the densities of collagen α1(I) and collagen α2(I) chains of the standard.

At day 5, collagen type I, collagen type III and collagen type IV deposition was quantified using established immunofluorescence protocols.^47,48 In brief, cell layers were washed, fixed, blocked, incubated with mouse anti-collagen type I (NB600-408, Novus Biologicals, USA), rabbit anti-collagen type III (Ab7778, Abcam, Ireland) or rabbit anti-collagen type V (Ab7046, Abcam, Ireland) primary antibodies, washed, incubated with AlexaFluor^® 488 goat anti-rabbit (A11008, Invitrogen, Ireland) or AlexaFluor^® 488 goat anti-mouse (A10667, Invitrogen, Ireland) secondary antibodies, washed and stained with DAPI (Invitrogen, Ireland). Images were captured with an Olympus IX-81 (Olympus Corporation, Japan) inverted fluorescence microscope and processed with ImageJ (NIH, USA).

Flow cytometry analysis

Flow cytometry was conducted at day 5 using standard protocols^22,30 for positive [CD44 (BD Bioscience, Ireland), CD73 (BD Bioscience, Ireland), CD90 (BD Bioscience, Ireland), CD105 (BD Bioscience, Ireland), CD146 (Invitrogen, Ireland), HLA-ABC (ThermoFisher Scientific, Ireland)] and negative [CD31 (Invitrogen, Ireland), CD45 (Invitrogen, Ireland), CD80 (Invitrogen, Ireland), CD86 (BD Bioscience, Ireland) and HLA-DR (ThermoFisher Scientific, Ireland)] markers. Briefly, cells were detached, centrifuged and resuspended in 2% FBS in phosphate-buffered saline (PBS). Following straining, the cells were counted, diluted, stained with the fluorochrome-labelled antibodies for 30 min at 4°C, washed with PBS and resuspended in 2% FBS in PBS. Unstained cell samples were used to correct for background autofluorescence and SYTOX™ Blue Dead Cell Stain (Invitrogen, UK) was used to label and exclude dead cells. Single stained samples; fluorescence minus one controls; and isotype control antibodies were used to determine the level of spectral overlap between different fluorophores and for compensation; to determine gating boundaries; and to assess the level of background staining and non-specific binding, respectively. The analysis was conducted using a BD FACSCanto™ II cytometer (BD Biosciences, UK). The median fluorescence intensity was calculated using the FlowJo^® software v10 (TreeStar Inc., USA). For gating, a primary gate was placed on the area versus height signal of the forward scatter dot plot to discriminate for doublets and cell aggregates. The single cell population was identified by defining the gated population on a side scatter area signal versus a forward scatter area signal dot plot. Single parameter histograms were generated, overlayed with respective isotype controls and range gates were used to determine the % of cells expressing the individual surface markers.

Preclinical analysis

All animal experiments and procedures were approved by the Animal Care and Research Ethics Committee of the University of Galway and the Irish Health Products Regulatory Authority (Licence Number: AE 19125/P051 K) and were conducted as per Irish laws governing laboratory animal experimentation. The splinted wound healing model and subsequent analyses were selected based on previous publications.^34,39,49,50 In brief, female athymic nude mice (7 weeks old) were purchased from Charles River (Ireland). After one week acclimatisation, the animals were randomly assigned to the following groups: non-treated control, human BMSCs without MMC (BMSCs-MMC) and human BMSCs with MMC (BMSCs + MMC). The human BMSC groups were delivered (per wound) as a single dose and single cell sheet that was produced after 5 days of culture of 50,000 human BMSCs per cm². Two circular (5 mm in diameter) full-thickness (epidermis, dermis, subcutaneous tissue and panniculus carnosus muscle) wounds were created with a single puncture and a silicone splint was sutured around each wound to prevent contraction and promote healing by epithelisation. Each animal received the same treatment in both wounds and all animals received identical post-operative pharmacological treatment. Wound closure rate was determined by taking digital pictures of the wounds with a digital camera (Canon, Japan) at the different time points and processing them with ImageJ (NIH, USA). The % wound closure was calculated as follows: [(area of original wound – area of actual wound) / area of original wound]×100. Harvested skin tissue samples were fixed with 4% paraformaldehyde and paraffin embedded. After sectioning (5 μm in thickness), deparaffinisation (2 immersions in xylene) and re-hydrations in descending concentrations of ethanol (100%, 90%, 70% and 0% in distilled water), the sections were stained using haematoxylin-eosin stain and Masson–Goldner's trichrome stain (both Carl Roth, Germany). Images were captured using an Olympus VS120 virtual slide microscope and processed using the OlyVIA software (both Olympus Corporation, Japan). Deparaffinised sections were blocked, incubated with the primary rabbit anti-cytokeratin 5 (Abcam, UK) antibody, washed, incubated with the secondary biotinylated swine anti-rabbit (Dako, USA) antibody and treated with ABC horseradish peroxidase labelled Vectastain Elite ABC reagent (Vector, UK), diaminobenzidine (Dako, UK) and haematoxylin. Images were captured using an Olympus VS120 virtual slide microscope and processed using the OlyVIA software (both Olympus Corporation, Japan). Scar index [scar index (μm) = scar area (μm²) / average dermal thickness (μm)] and epidermal thickness were evaluated using ImageJ software (NIH, USA) on images from Masson–Goldner's trichrome stained histological sections.

Statistical analysis

Data are expressed as mean ± standard deviation. Number of replicates is indicated in each figure legend. Statistical analysis was performed using either MINITAB^® (Minitab LLC., USA) or Prism (GraphPad Software, USA). One-way analysis of variance was used for multiple comparisons and Tukey's post hoc test was used for pairwise comparisons after confirming the samples followed a normal distribution (Anderson–Darling test) and had equal variances (Bartlett's and Levene's test for homogeneity of variances). When either or both assumptions were violated, non-parametric analysis was conducted using Kruskal–Wallis test for multiple comparisons and Mann–Whitney test for pairwise comparisons. Statistical significance was accepted at p < 0.05.

Results

Cell metabolic activity, viability and proliferation analyses

Cell metabolic activity (Figure 1A), viability (Figure 1B) and proliferation (Figure 1C) analyses showed no significant (p > 0.05) differences between the groups.

Figure 1.

Quantitative analysis of % metabolic activity, viability and proliferation of human BMSCs after 5 days in culture without (−) and with (+) MMC. N = 4 for metabolic activity and N = 3 for viability and proliferation.

Collagen deposition analysis via electrophoresis and immunofluorescence

The SDS-PAGE and densitometry analyses showed that +MMC significantly (p < 0.05) increased collagen type I deposition (Figure 2). Immunofluorescence and fluorescence intensity analyses revealed that +MMC significantly (p < 0.05) increased collagen type I and collagen type IV deposition, without affecting (p > 0.05) collagen type III deposition (Figure 3).

Figure 2.

SDS-PAGE (A) and densitometry analysis (B) of collagen type I deposition of human BMSCs after 5 days in culture without (−) and with (+) MMC. # indicates significant (p < 0.05) difference. N = 3.

Figure 3.

Immunofluorescence (A) and normalised to cell number image intensity analysis (B) of collagen type I, collagen type III and collagen type IV deposition of human BMSCs after 5 days in culture without (−) and with (+) MMC. # indicates significant (p < 0.05) difference. Scale bar: 100 μm. N = 4.

Flow cytometry analysis

Flow cytometry analysis (Figure 4) revealed that over 92% of the BMSCs-MMC and the BMSCs + MMC were positive for the positive MSC markers CD44 (92.1% and 93.0%, respectively), CD73 (95.8% and 96.9%, respectively), CD90 (95.0% and 97.0%, respectively) and HLA-ABC (94.5% and 96.4%, respectively). The BMSCs-MMC and the BMSCs + MMC showed variable results for the positive MSC marker CD105 (10.6% and 51.2%, respectively, were positive) and less than 1% expressed the positive MSC marker CD146 (0.41% and 0.43%, respectively, were positive).

Figure 4.

Flow cytometry analysis of human BMSCs after 5 days in culture without (−) and with (+) MMC. Blue: isotype control. Red: antibody. N = 1.

Further, less than 2% of the BMSCs-MMC and the BMSCs + MMC were positive for the negative MSC markers CD31 (1.81% and 1.81%, respectively), CD45 (1.32% and 0.51%, respectively), CD80 (0.65% and 1.16%, respectively) and CD86 (0.23% and 0.23%, respectively). The BMSCs-MMC and the BMSCs + MMC showed variable results for the negative MSC marker HLA-DR (61.2% and 7.57%, respectively, were positive).

Wound closure, histological and immunohistochemical analyses

Neither infection nor necrotic tissues were found in any of the groups at any time point. The animals were removing the splints, which had to be re-sutured several times during the course of the study. Qualitative (Figure 5A) and quantitative (Figure 5B) wound closure analysis revealed no significant (p > 0.05) difference between the groups at a given time point. Haematoxylin–eosin staining showed complete re-epithelisation in all groups 14 days after injury and immunohistochemical analysis of cytokeratin 5 revealed that in all groups, protein expression was restricted to the epidermal layers and hair follicles (Figure 6). Further, at best case scenario, haematoxylin–eosin and cytokeratin 5 stainings revealed that the BMSC treated groups outperformed the empty defect control group (Figure 6). Masson–Goldner's trichrome staining revealed dense collagenous tissue formation in the BMSCs ± MMC, but not in the control group; scar index analysis revealed no significant (p > 0.05) differences between the groups; and epidermal thickness analysis revealed only the BMSCs + MMC group to have significantly (p < 0.05) higher epidermal thickness compared to the intact skin group (Figure 7). Further, at best case scenario, Masson–Goldner's trichrome staining revealed that the BMSC treated groups outperformed the empty defect control group (Figure 7).

Figure 5.

Qualitative (A) and quantitative (B) wound closure analyses at day 3, day 7 and day 14 of non-treated control group and treated with human BMSCs after 5 days in culture without (−) and with (+) MMC test groups. N = 5 wounds for all but BMSCs at day 14, for which N = 3 wounds.

Figure 6.

Qualitative haematoxylin–eosin (A, B) and cytokeratin 5 (C, D) stainings of intact skin and of day 14 post-surgery of non-treated control group and treated with human BMSCs after 5 days in culture without (−) and with (+) MMC test groups. Scale bars: 200 μm for intact skin, average haematoxylin–eosin and average cytokeratin 5 images; 500 μm for best haematoxylin–eosin and best cytokeratin 5 images. N = 5 wounds for all groups for haematoxylin–eosin. N = 1, 4, 6 and 5 wounds for intact skin, control, BMSCs-MMC and BMSCs + MMC, respectively, for cytokeratin 5.

Figure 7.

Qualitative Masson–Goldner’s trichrome staining (A, B) and quantitative scar index (C) and epidermal thickness (D) analyses of intact skin and of day 14 post-surgery of non-treated control group and treated with human BMSCs after 5 days in culture without (−) and with (+) MMC test groups. # indicates significant (p < 0.05) difference. Scale bars: 200 μm for intact skin and average Masson–Goldner's trichrome images; 500 μm for best Masson–Goldner's trichrome images. N = 1, 6, 6 and 5 wounds for intact skin, control, BMSCs-MMC and BMSCs + MMC, respectively, for Masson–Goldner's trichrome. N = 6, 6 and 5 wounds for control, BMSCs-MMC and BMSCs + MMC, respectively, for scar index. N = 5, 6, 6 and 5 wounds for intact skin, control, BMSCs-MMC and BMSCs + MMC, respectively, for epidermal thickness.

Discussion

Although human BMSCs have shown promise for a range of clinical indications, including wound healing, no BMSC-based tissue-engineered medicine has been commercial reality to-date. This limited technology transfer is attributed to the prolonged in vitro cultured periods required to develop an ECM / secretome-rich implantable construct that are associated with phenotypic drift and high manufacturing costs and therefore high cost of goods for healthcare providers. Although MMC has been shown to enhance and accelerate ECM deposition in various cell types, including BMSCs, a tissue-engineered construct based on BMSCs and MMC has yet to be assessed in preclinical setting. Herein, we assessed the therapeutic potential of human BMSCs, which were cultured -MMC and + MMC conditions, in a humanised athymic nude mouse excisional wound splinting model.

Cell metabolic activity, viability and proliferation analyses

Starting with basic cell function analysis, no differences were observed between the groups in metabolic activity, viability and proliferation. To-date carrageenan has been used extensively as MMC agent in a diverse range of cell populations (e.g. skin fibroblasts,^38,51,52 human tenocytes,^45,48,53 human BMSCs,²³ human adipose derived MSCs^23,47,54 and human umbilical cord MSCs⁴³) with no study reporting any negative effect. One should also note that carrageenan-based devices have long positive history in clinical setting,^55–57 further verifying its safety, efficiency and efficacy.

Collagen deposition analysis via electrophoresis and immunofluorescence

Extracellular matrix molecules are essential players in both physiological and pathological wound healing.⁵⁸ Therefore, the development of a tissue-engineered medicine, which accounts for physiological levels of ECM molecules, is crucial for functional tissue repair and regeneration. Collagen type I, collagen type III and collagen type IV are essential ECM molecules in wound healing. Collagen type III has various key roles (provides a temporary wound matrix, increases tensile strength, facilitates collagen type I fibrillogenesis) in the early stages of wound healing.⁵⁹ Collagen type I replaces collagen type III as the wound healing progresses and confers to the regenerated tissue mechanical stability.⁶⁰ Collagen type IV is essential for basement membrane formation.⁶¹ Under MMC conditions, a substantial increase in collagen type I and collagen type IV deposition was observed, whilst the amount of deposited collagen type III was not affected, as per previously published work.²³ It is interesting to note that other MMC agents (e.g. Ficoll^®,⁴⁵^62–64 hyaluronic acid^47,65) have resulted in increased collagen type III deposition. Although the mechanism that induces this selective collagen types increased deposition has yet to be elucidated, we consider the indifference in collagen type III deposition of therapeutic value, as increased collagen type III is often associated with fibrosis.^66–68

Flow cytometry analysis

Flow cytometry analysis made apparent that over 90% of the -MMC and + MMC BMSC populations expressed the positive MSC markers CD44, CD73, CD90 and HLA-ABC and less than 2% of the -MMC and + MMC BMSC populations expressed the negative MSC markers CD31, CD45, CD80 and CD86, as one would have expected.^69–72 Nonetheless, BMSCs-MMC (10.6% were positive) and, to a lesser extent, BMSCs + MMC (51.2% were positive) had low expression of the positive marker CD105. Although CD105 is widely accepted as MSC marker,^69,73 it is worth noting that CD105 negative populations have shown typical MSC differentiation potential and immunomodulatory capacity⁷⁴ and CD105 expression has been shown to depend on passaging and culture media composition.^75–78 With respect to the positive MSC marker CD146, both groups were almost entirely negative (less than 0.5% positive). Again, although CD146 is commonly considered as a positive MSCs marker^79,80 and CD146 positive cells have been shown to display greater therapeutic potential than CD146 negative cells in arthritis,⁸¹ several studies have shown similar differentiation potential for both CD146 negative and positive MSC populations,^82–85 with some differences though in modulating immunological and proliferation properties (improved for the CD146 positive cells).⁸⁶ With regards to the negative MSC marker HLA-DR, a fairly high % of the BMSCs-MMC were positive (61.2%) and a relatively low % of the BMSCs + MMC were positive (7.57%). Again, although MSCs should be negative for HLA-DR,⁶⁹ in vitro expression of HLA-DR has been reported to be donor- and culture media-dependent.^87–90 Some studies have even considered it an obsolete marker,⁸⁷ considering its expression informative, but not critical.⁹¹ Collectively, we believe that the implanted cells had maintained their MSC function and differences in the expression of surface markers may be attributed to donor and media variations between studies, as explained earlier. The differences between the -MMC and + MMC groups (for CD105, higher % of positive cells in the + MMC group, and HLA-DR, lower % of positive cells in the + MMC group) could be attributed to the enhanced ECM deposition that better protected the cells. To substantiate this one should consider that cells on stiff two-dimensional substrates (i.e. cells grown on tissue culture plastic under -MMC conditions) are more spread than cells on soft three-dimensional substrates (i.e. cells grown on deposited ECM under + MMC conditions)^92,93 and the rebound level of CD105 has been shown to be high when the cell spreading area is small (i.e. under + MMC conditions).⁹⁴ With respect to HLA-DR, it is known that its expression is associated with implant / cell rejection and immune response,^95–98 thus its reduction under + MMC conditions is welcome. In any case though, one should note that even clinical-grade BMSCs have been found positive for HLA-DR.⁹¹ Although the specific mechanism is unclear at this stage, considering that various growth factors have been shown to affect HLA-DR expression,^99–101 we speculate that this reduction under + MMC conditions may be due to growth factor retention in the deposited ECM.

Wound closure, histological and immunohistochemical analyses

The use of MSCs for skin tissue engineering has been advocated due to their immunomodulatory, anti-inflammatory, mitogenic and angiogenic capacities, along with their rich secretome.^102–105 Various studies have demonstrated human BMSCs-based^106,107 and MMC-based^34–36 therapies to result in improved healing; however, this was not witnessed herein. Indeed, preclinical analysis revealed similar wound closure, re-epithelisation, scar index and epidermal thickness between the groups. It should be noted that negative outcomes (e.g. infection, inflammation, necrosis) were not observed. Further, herein we transplanted a single cell sheet per wound that was produced after 5 days in culture (without or with MMC) using 50,000 human BMSCs per cm² as original seeding density; other studies that have demonstrated efficiency and efficacy of stem cell sheets used substantially higher cell numbers (e.g. 2,000,000 human BMSCs per cm²¹⁰⁸; 3 layers of 300,000 human adipose derived mesenchymal stromal cells per cm² per layer¹⁰⁹; ∼350,000 human amniotic fluid mesenchymal stromal cells per cm²¹¹⁰). This indifference between the groups may also be due to the animals removing the silicone rings, which resulted in healing by tissue contraction. Similar indifference between the groups has been reported previously.^111,112 The reader should note that over the years various modifications of the model (e.g. the use of Elizabethan collars and jackets with plastic wings¹¹³ or wound chambers¹¹⁴ or shape memory alloys as internal splints¹¹⁵) have been suggested. Regretfully, these approaches were not in our licence and therefore we were not able to implement them. Researchers should point out such crucial limitations of preclinical experimentation to inform future preclinical endeavours. Alternative models, such as a full-thickness Sprague–Dawley rat model of 1.0 cm by 1.0 cm wound area¹¹⁶ or a full-thickness athymic nude mouse, nu/nu, model of 2.5 cm by 2.5 cm wound area,¹¹⁷ should also be considered.

Conclusions

Despite the fact that human BMSCs are extensively used in tissue engineering and regenerative medicine, they have yet to constitute the building blocks of a commercially available tissue-engineered medicine. This limited technology transfer from research to development has been attributed to the protracted culture periods required to develop a tissue-engineered medicine that are associated with phenotypic drift, loss of therapeutic potential and very high manufacturing costs. Considering that MMC has the potential to accelerate the development of tissue-engineered medicines, herein human BMSCs were cultured under MMC conditions and the developed tissue-engineered medicine was assessed in vitro and in vivo. Our in vitro data indicate that MMC does not affect human BMSC function and enhances and accelerates ECM deposition. Regretfully, the in vivo data are inconclusive due to issues encountered with the model. In any case, this preliminary study sets the foundations for exploiting MMC in the development of functional and extracellular matrix-rich tissue-engineered medicines.

Footnotes

Acknowledgements

The authors would like to acknowledge the significant contribution of Dr Oonagh Dwane (University of Galway, Ireland) in the writing and management of all grants. The Graphical abstract was prepared using bioRender. No AI tools were used anywhere in the manuscript.

Authors contributions

Kyriakos Spanoudes, Laura Trujillo Cubillo, Stefanie H. Korntner and Diana Gaspar conducted experimental work, including data analysis, and contributed to the writing of the manuscript. Dimitrios I Zeugolis acquired and managed funding; supervised the work; conducted data analysis; and wrote, edited and finalised the manuscript. All authors have approved the final version of the manuscript.

ORCID iD

Dimitrios I. Zeugolis

Funding

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation programme, grant agreement No. 866126. This publication has emanated from research supported by Research Ireland under grant number 19/FFP/6982.

Declaration of conflicting interests

The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Data availability

Raw and processed data are available on request from Kyriakos Spanoudes, Laura Trujillo Cubillo, Stefanie H. Korntner and Diana Gaspar.

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A preliminary preclinical assessment of macromolecular crowding in tissue engineering

Abstract

Objectives

Methods

Results

Conclusions

Keywords

Introduction

Materials and methods

Cell isolation and culture

Cell metabolic activity, viability and proliferation analyses

Collagen deposition analysis via electrophoresis and immunofluorescence

Flow cytometry analysis

Preclinical analysis

Statistical analysis

Results

Cell metabolic activity, viability and proliferation analyses

Collagen deposition analysis via electrophoresis and immunofluorescence

Flow cytometry analysis

Wound closure, histological and immunohistochemical analyses

Discussion

Cell metabolic activity, viability and proliferation analyses

Collagen deposition analysis via electrophoresis and immunofluorescence

Flow cytometry analysis

Wound closure, histological and immunohistochemical analyses

Conclusions

Footnotes

Acknowledgements

Authors contributions

ORCID iD

Funding

Declaration of conflicting interests

Data availability

References