Matrix Vesicles-Containing Microreactors as Support for Bone-Like

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Biological and Medical Applications of Materials and Interfaces

Matrix Vesicles-Containing Microreactors as Support for Bone-Like Osteoblast Cells to Enhance Biomineralization Fabian Itel, Jesper Skovhus Thomsen, and Brigitte Städler ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b10886 • Publication Date (Web): 16 Aug 2018 Downloaded from http://pubs.acs.org on August 17, 2018

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ACS Applied Materials & Interfaces

Matrix Vesicles-Containing Microreactors as Support for Bone-Like Osteoblast Cells to Enhance Biomineralization Fabian Itel,a Jesper Skovhus Thomsen,b Brigitte Städlera,* a

Interdisciplinary Nanoscience Center (iNANO), Aarhus University, Gustav Wieds Vej 14, 8000

Aarhus, Denmark b

Department of Biomedicine, Aarhus University, Wilhelm Meyers Allé 3, 8000 Aarhus,

Denmark Keywords: Matrix vesicles, microreactors, alginate, droplet microfluidics, SaOS-2 cell spheroids, biomineralization, micro-computed tomography

ABSTRACT

Therapeutic cell mimicry aims to provide a source of cell-like assemblies, which exhibits the core structural or functional properties of their natural counterparts with broad envisioned applications in biomedicine. Bone tissue engineering (BTE) aims at promoting and inciting the natural healing process of for instance critically-sized bone defects. Microreactors designed to co-assemble with biological bone-forming osteoblast like SaOS-2 cells to kick-start

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biomineralization are reported for the first time. The alginate-based microparticles are equipped with active alkaline phosphatase-loaded artificial liposomes or SaOS-2 derived matrix vesicles (MVs). Spheroids assembled from SaOS-2 cells and microreactors not only exhibit higher cell viability, but also show enhanced biomineralization when MVs are present. The active biomineralization stimulation of the microreactors is illustrated by colorimetric calcium quantification and micro-computed tomography (µCT). These findings show the promising potential of applying cell mimicry in BTE.


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INTRODUCTION Therapeutic cell mimicry aims to provide a source of cell-like assemblies, which exhibit the core structural or functional properties of their natural counterparts with broad envisioned applications in biomedicine. Both top-down and bottom-up approaches are explored. In the former case, cell encapsulation in hydrogel particles is the dominating concept as recently reviewed by Grattoni and coworkers.1 Cell-laden microbeads are currently considered as injectable cell-based drug delivery systems because the sensitive cells can be protected from the harsh outside environment and can, thus, maintain their cellular functions to produce therapeutic drugs,2-4 for example encapsulating pancreatic cells for insulin delivery in diabetes.5 Furthermore, non-autologous engineered cell lines have been assayed to deliver therapeutic products in cancer therapy.6 When considering bone tissue engineering,

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cells have been

encapsulated into alginate hydrogel beads to engineer 3D synthetic extracellular matrix material for cell immobilization,8-10 cell transplantation,11 or stem cell differentiation12-13. On the other hand, bottom-up assembled cell mimics typically consist of polymers, lipids and protein building blocks (self-)assembled into functional microreactors.14 While the complexity of such microreactors (e.g., sub-compartmentalization,15 incorporation of extracellular vesicles16) has advanced in the past years, their interaction with biological cells remains little explored, especially when considering aspects beyond cell viability. In an early example, we illustrated that liposome-loaded core shell particles were able to preserve their biocatalytic activity in the presence of cells.17 Another interesting effort in this context has been reported by Tang et al.,18 who functionalized the surface of microparticles with membrane fragments of cardiac stem cells and loaded them with secretome to improved cell-microparticle interactions and support injured cells through the secretion of regenerative factors, respectively. In another attempt, we have


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shown that liposomes-loaded core-shell particles19 and alginate-based microreactors20 containing catalase could be integrated with proliferating liver-like HepG2 cells. In addition to structural support, the microreactors assisted the biological cells in removing cytotoxic hydrogen peroxide (H2O2). Microreactors carrying platinum nanoparticles as catalytic active entity were considered as to scavenge H2O2 and ammonia to improve the viability of neuroblastoma cells.21 While these initial efforts illustrated the potential of combining artificial and biological entities, the field is in its infancy and only limited cell and tissue types have been explored. Interaction of micro- or nanoreactors with bone cells would open up new possibilities in bone tissue engineering to address long-standing challenges including sustainable solutions for critically sized bone defects.22 Today’s gold standard treatment typically employs autologous bone grafts, suffering from drawbacks such as limited tissue supply, donor site morbidity, infections or poor integration.23 Bone tissue engineering aims to circumvent these disadvantages e.g., by implanting matured tissue consisting of patient’s own cells in combination with a scaffold material. Osteoconductive and osteostimulative materials (e.g., bioglass/ceramics, demineralized bone matrix) are widely explored or already FDA-approved for clinical use to improve mineralization in regenerating bone tissue.24 The cells of choice for bone regeneration are osteoblasts.25 These cells produce a collagen-rich extracellular matrix, where bone mineralization takes place, and show an increased tissue-nonspecific alkaline phosphatase (TNAP) activity. TNAP is an enzyme that catalytically cleaves inorganic pyrophosphates (PPi) into inorganic phosphate (Pi) in the extracellular fluid. Furthermore, osteoblasts secrete matrix vesicles (MVs) into the extracellular matrix to induce bone mineralization. MVs are 100 - 300 nm-sized extracellular vesicles (EVs), whose membrane is enriched with the negatively-charged lipid phosphatidylserine (PS) and that contain Ca2+ ions, Pi, TNAP and different Ca2+- and Pi-


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channel proteins.26-28 Further, the roles of other protein components of MVs in mineralization remain unclear and are still under investigation.28 However, MVs induce the primary crystallization of bone mineral, hydroxyapatite (HA), where Ca2+ and Pi accumulates inside the MV and precipitates.27,

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While Ca2+ - channels pump Ca2+ into the MV vesicles, TNAP

increases the local Pi concentration. As a consequence, the Ca2+ and Pi concentrations are above the critical solubility product and precipitate at the desired site. Furthermore, TNAP diminishes the mineralization inhibitor PPi in order to facilitate mineral precipitation and crystal growth and, thus, allowing bone mineralization.30-32 Beside the mineralization function of MVs, they are also able to mediate cell signaling.33 Bone morphogenic proteins (BMP-2 and BMP-4) on MVs may serve as stimulators for morphogenic information to nearby osteoblasts, and microRNA inside MVs serve as signaling molecules in cell-cell communication.34 Furthermore, TNAP, besides other proteins, has been reported to be involved in stimulating cell differentiation.33 MVs can be extracted from osteoblast cell cultures and utilized as crystallization cores since they facilitate the nucleation and formation of HA.35 Moreover, it has been reported that MVs even stimulate the differentiation of human bone marrow mesenchymal stem cells towards a mineralizing phenotype.36 In general, EVs are currently explored in clinical trials for therapeutic use, but have considerably lower therapeutic potential when administered systemically due to dilution effects and unwanted targeting, among others.37-38 Therefore, EVs are envisioned to possess great potential when encapsulated in hydrogel networks and delivered or placed at the targeting site. A sole example in this context was recently reported by the Steven’s group where they compared synthetic β-glucuronidase-loaded liposomes and beta-glucuronidase-loaded EVs in a hydrogel for local conversion of the prodrug curcumin-β-D-glucuronide into cytotoxic curcumin.16 Both


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hydrogel coatings performed similarly in terms of reducing the viability on mouse macrophages (RAW 264,7) in cultures apart from initial slightly enhanced activity when EVs were present. However, despite the anticipated potential for EVs in clinical applications, challenges like the standardization of the isolation process with respect to yield, reproducibility and purity need to be overcome.38-39 Therefore, artificially made mimics of MVs or EVs would provide an alternative because clinical grade liposomal carriers are already achievable today. Liposomes equipped with TNAP were reported as biomimetic- or artificial MVs (AMVs) to serve as a simplified alternative to MVs.40-45 Due to the osteoconductive property of certain calcium phophates,46-47 triggered mineralization inside the lumen of liposomes could be envisioned to induce an osteoconductive effect on bone cells. However, triggered mineralization within liposomes in presence of cells was never reported. Here, we report the generation of ~50 µm-sized alginate-based microreactors loaded with functional AMVs or MVs, which, when co-assembled with biological osteoblast-like SaOS-2 cells in a 3D spheroid, enhance biomineralization (Scheme 1). Specifically, i) the characteristics of spheroids consisting of SaOS-2 cells and empty microreactors were determined, ii) the TNAPactivity of AMVs and SaOS-2-derived MVs in solution and assembled as subunits in microreactors were compared (Scheme 1A), and iii) the time-dependent biomineralization of spheroids composed of SaOS-2 cells and microreactors (Scheme 1B) was assessed using calcium quantification and micro-computed tomography (µ-CT).


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Scheme 1. Schematic overview of the microreactor assembly (A) and the spheroid formation by combining

mammalian

SaOS-2

cells

with

microreactors

including

the

subsequent

biomineralization (B). A) Alginate-based microreactors were generated by droplet microfluidics (D-µF) and coated with positively-charged poly-L-lysine (PLL). Four different microreactors were employed: ME – PLL coated alginate particles without subunits; MAMV1 and MAMV2 microreactors loaded with artificial matrix vesicles (AMV) consisting of liposomes harboring tissue-nonspecific alkaline phosphatase (TNAP) inside and outside anchored to the membrane and only inside of the liposomes, respectively; MMV – microreactors equipped with SaOS-2derived matrix vesicles. B) Microreactors and SaOS-2 cells were co-cultured in ultra-low attachment round bottom well plates to form spheroids. In presence of osteoconductive media, TNAP hydrolyzes inorganic pyrophosphates (PPi) into inorganic phosphate (Pi) leading to calcium phosphate precipitates (CaP) i.e., biomineralization.


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MATERIALS AND METHODS Materials. Sodium alginate (Alg), poly(L-lysine) (PLL, MW of 30-70 kDa), sodium chloride (NaCl),

calcium

chloride

(CaCl2),

tris(hydroxymethyl)aminomethane

(TRIS),

4-(2-

hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES), phosphate buffered saline (PBS), Triton X-100, Span 80, Tween 80, mineral oil, 1H,1H,2H,2H-perfluorooctyltriethoxysilane (PFOTES),

dimethyl

sulfoxide

(DMSO),

ethanol,

chloroform

(purity

of

≥99.5%),

paraformaldehyde (PFA), phalloidintetramethylrhodamine B isothiocyanate (phalloidin), 6diamidino-2-phenylindole (DAPI), fluorescein diacetate (FDA), propidium iodide (PI, 1 mg mL-1 in H2O), 4-methylumbelliferyl phosphate (4-MUP), alkaline phosphatase (AP) from bovine intestinal mucosa (≥10 DEA units mg-1 solid), collagenase Type I from Clostridium histolyticum (≥125 CDU mg-1 solid), 2-amino-2-methylpropanol (AMP), o-cresolphthalein complexone (OCPC), and 8-hydroxychinoline were purchased from Sigma-Aldrich. Phoshphatidylinositolspecific phospholipase C (PHC-PI) and DyLight 633 maleimide were purchased from Fisher Scientific. 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), 1-palmitoyl-2-oleoyl-snglycero-3-phospho-L-serine

(POPS),

and

1-oleoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)

amino] hexanoyl]-sn-glycero-3 phosphocholine (NBD-PC) were purchased from Avanti Polar Lipids. Unless noted otherwise HEPES buffer consisting of 20 mM Hepes, and 130 mM NaCl (pH 7.5) was used. Other buffers were: receptor solution (20 mM Hepes, 50 mM CaCl2, 55 mM NaCl, pH 7.5), Ca-low-HEPES buffer (20 mM Hepes, 122.5 mM NaCl, 5 mM CaCl2, pH 7.5). Buffers were made using ultrapure water (monodistillation unit, GFL Corporation, Germany).


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For the fluorescent labeling of TNAP (TNAP-633), a stock solution of TNAP at 5 mg mL-1 in HEPES buffer was prepared. To this, a DyLight 633 maleimide stock solution in DMF was added dropwise at 10 molar excess of dye while stirring and incubated while stirring for 3 h at room temperature. The mixture was then dialyzed (MWCO 3.5 kDa) against 2 L of HEPES buffer for 24 h at room temperature., the dialyzed sample was put on a PD-10 Desalting Column (GE Healthcare Bio-Sciences) to further remove unreacted dye and eluent was collected in aliquots of 200 µL. The aliquots were analyzed for protein content by measuring the absorbance at λ = 280 nm using UV spectrophotometry (Nanodrop, ThermoScientific). Colocalization of TNAP with the liposomes inside the microreactors was quantified using the total Pearson correlation coefficient (PCC) determined (Coloc 2 plug-in in ImageJ) as recently described.48 Background subtraction (10 pixel ball pen size) was performed before the analysis for all of the images. Four images from one independent repeat were used. Catalytic active subunits. Artifical matrix vesicles AMV1 and AMV2. TNAP-containing liposomes were prepared by the film rehydration method. Briefly, POPC and POPS, dissolved in chloroform, were mixed in brown glass vial at a molar ratio of 4:1. The lipid mixture was dried under a gentle nitrogen stream, forming a smooth lipid film on the inside of the vial, followed by vacuum drying for 30 min. The lipid film was hydrated to a concentration of 5 mg mL-1 in a Ca-Hepes buffer (50 mM Hepes, 92.5 mM NaCl, 5 mM CaCl2, pH 7.5) containing 5 mg mL-1 TNAP. The sample was incubated with intermittent agitation for 15 min at room temperature and extruded 15× through a 200 nm track-etched filters (Nucleopore; Whatman). If required TNAP attached outside of the liposomes via a GPI-anchor, were removed by incubating the liposomes with 1 U mL-1 phosphatidylinositol-specific phospholipase C (PHC-PI) for 60 min at 37 °C on a thermoshaker.


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Extra-vesicular material was then removed and the buffer was exchanged for HEPES buffer on a gravity flow column using a 20 cm3 in-house prepacked size exclusion column (Sepharose 2B, Sigma Aldrich) yielding AMV1 (TNAP inside and attached to the membrane) and AMV2 (TNAP only inside). AMV1 and AMV2 were stored at 4 °C until further use. Liposome formation and size was confirmed by dynamic light scattering (DLS, Zetasizer Nano S90) using a material refractive index of 1.590 and a dispersant (water at 25 °C) refractive index of 1.330. Samples for DLS were diluted 10× in HEPES buffer at pH 7.0. Fluorescent AMV1 were obtained by adding 0.1 wt% NBD-PC to the lipid mixture and 10 wt% TNAP-633 to the 5 mg mL-1 TNAP during preparation. The morphology of AMVs and MVs were imaged using transmission electron microscopy (TEM) (Tecnai G2 Spirit, TWIN/BioTWIN, FEI Company). Carbon coated copper grids were glow discharged and 5 µL of 0.5 mg mL−1 sample was let to adsorb for 1 min. The grids were then washed 2 × 5 µL with ultrapure water and negatively stained with 2% uranyl acetate solution. Activity. Enzymatic activities of AMV1, AMV2 and SaOS-2-derived matrix vesicles (MV, see below for purification details) were determined using the fluorescent phosphatase substrate 4methylumbelliferyl phosphate (4-MUP). Activity measurements were performed in black 96well plates at final volumes of 200 µL per well. A volume of 2.0 µL vesicle stock solution was used in all measurements and diluted in HEPES buffer. The non-fluorescent substrate 4-MUP was added to a final concentration of 50 µM. A 10× concentrated 4-MUP stock solution in HEPES buffer was prepared just before the activity measurements in order to minimize selfhydrolysis of the substrate. Samples containing 0.1 wt% Triton-X in order to disintegrate the liposomes were used as controls. The time dependence of the conversion of 4-MUP into the


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fluorescent product 4-MU (λex = 370 nm, λem = 440 nm) was recorded using a multimode plate reader (EnSpireTM 2300 Multilabel Reader, PerkinElmer) at 37 °C. The activities of the different samples were determined by comparing the initial slopes, which are a direct measure for the relative reaction speed, as the increase in fluorescence intensity over time is directly linked to the increase in concentration of the fluorescent product 4-MU. Initial slopes were obtained by fitting an exponential rise equation to the obtained curves using Origin software (OriginLab, Northampton, MA). Microreactors. Assembly. Calcium crosslinked alginate (Alg) particles (microreactors) were prepared by droplet microfluidics (D-µF). A flow focusing microfluidic glass chip with wide channel crosssection dimensions of 100 µm × 300 µm (depth × width) was used (Dolomite Microfluidics). The channels were rendered hydrophobic by coating with PFOTES for 30 min at room temperature. The flow focusing design required two immiscible fluids to generate Alg droplets. The continuous phase consisted of mineral oil containing 5 wt% Span 80 and Tween 80 (9:1 weight ratio). The dispersed phase consisted of 1.5 wt% Alg solution dissolved in HEPES buffer. If incorporation of AMV1/2 or MV was required, a 3 wt% Alg solution was added dropwise while vortexing to the vesicle stock solution to yield a final Alg concentration of 1.5 wt%. The oil and solution for microreactor formation were pumped into the microchannels with digital syringe pumps (Harvard Apparatus). The flow rates for the oil phase and the aqueous phase were 1.0 mL h-1 and 0.05 mL h-1, respectively, and the microreactor production was visualized using a microscope. Gelation of the droplets was induced by collecting them in a receptor solution containing 50 mM CaCl2. Typically, microreactors were generated for 2 h at room temperature. The crosslinked microreactors in the receptor solution containing oil were washed 3× by


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sedimentation for 30 min at 4 °C. The microreactors settled at the bottom of a test tube and the supernatant, containing oil droplets, was removed. Before the PLL coating, the microreactors were settled again, the supernatant was removed and the buffer was exchanged with a Ca-lowHEPES buffer. The microreactors were then quantified in a hemocytometer for manual cell counting. The volume was adjusted to reach a particle concentration of ~400 particles µL-1. The microreactors were coated with PLL by mixing them with Ca-low-HEPES buffer containing 1.2 mg mL-1 PLL at a 1:1 (v:v) ratio and incubation with rotation for 5 min at room temperature. The tube was filled with Ca-low-HEPES buffer and let stand in the fridge until the microreactors settled. The supernatant was replaced with Ca-low-HEPES buffer. This washing procedure was repeated 3×. The final PLL-coated microreactors were counted again and the particle concentration was adjusted to ~400 particles µL-1. The microreactors were imaged using an inverted Olympus IX81 microscope in the bright field mode. Naming of the microreactors: empty microreactors: ME, microreactors loaded with AMV1: MAMV1, microreactors loaded with AMV2: MAMV2, and microreactors loaded with MV: MMV. Activity of microreactors. Enzymatic activities of MAMV1, MAMV2 and MMV were determined as outlined for the catalytic active subunits, using 104 microreactors per well. The microreactors were diluted in HEPES buffer containing additional 2 mM CaCl2. Kinetic measurements were performed at 37 °C. For long term activities, the microreactors were kept in the fridge until the measurement. Cell culture. SaOS-2 human osteosarcoma cells (European Collection of Cell Cultures) were used for all experiments. Cells were cultured and propagated in 75 cm2 cell culture flasks using McCoy’s 5A cell media (Sigma Aldrich) supplemented with 10% Fetal Bovine Serum (FBS), 1% penicillin/streptomycin, and 2 mM L-glutamine at 37 °C in a humidified chamber supplied


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with 5% CO2. Cells were grown until 80 – 90% confluency and 3 × 106 cells per flask were used for subcultures. SaOS-2-derived matrix vesicles (MV). MVs isolated from mineralizing SaOS-2 cells were purified according to a standard protocol with slight modifications.35 Briefly, SaOS-2 cells were cultured in two T75 culture flasks (25000 cells cm-2) with 15 mL mineralization buffer (cell media including 50 µg mL-1 ascorbic acid and 7.5 mM β-GP). Cells were incubated for 7 days at 37 °C (5 % CO2) and mineralization medium was exchanged every 3 days. After 7 days, cells were washed 3× with PBS. Then, 12.5 mL digestion buffer (200 U mL-1 collagenase Type I, 1 mM CaCl2 in Hank’s balanced salt solution (HBSS)) was added to each flask and incubated at 37 °C for 3 h. The cells were centrifuged at 600 × g for 15 min at 4 °C. The supernatant was centrifuged first at 20 000 × g for 20 min at 4 °C and then at 100 000 × g for 60 min at 4 °C. The resulting pellet contained the MVs and was carefully washed with 1 mL HEPES buffer. The pellet was then gently resuspended in 250 µL of HEPES buffer and kept at 4 °C until further use. Mineralization assay on MVs. A mineralization assay was performed according to a prior reported protocol.35 Briefly, 20 µL of AMV and 2 µL of MV stock solution was diluted in HEPES buffer containing 4 mM CaCl2 in a total volume of 100 µL and added to a 96-well plate. In order to start the mineralization, the same volume of HEPES buffer containing 6.84 mM Pi was added to the wells. After mixing the two solutions (1:1 (v/v)), a final CaCl2 concentration of 2 mM and Pi of 3.42 mM was obtained. DMSO was used as positive control (without MVs or AMVs) and added to final concentration of 4% (v/v). The plate was incubated at 37 °C and mineralization was determined by measuring the absorbance at λ = 340 nm in 30 min intervals for 3h on a multimode plate reader.


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Spheroid formation. ME were co-cultured with SaOS-2 cells at a cell-to-particle ratio of 20:1 in 96-well spheroid microplates with round well-bottom geometry and with ultra-low attachment surface coating (Corning, Inc) with 104 cells per well in 200 µL cell media. Spheroids with only cells (S-C) were seeded as controls in the same way. The cells and ME settled at the bottom of the wells and formed a stable spheroid. After 24 h, cell media was exchanged and experiments were started. Cell viability assay. For the Live/Dead staining assay, spheroids were washed with PBS, transferred into fresh spheroid microplate wells and stained with 20 µM FDA (8 µg mL-1) and 30 µM PI (20 µg mL-1) in PBS for 10 min at room temperature in the dark. The spheroids were washed with PBS and transferred into a drop of PBS on a microscopy glass slide for visualization on a LSM700 confocal laser scanning microscope (CLSM, Carl Zeiss) using a 20× objective. The dead-cell fraction was calculated based on 3 different CLSM images of day 3 and 7 per independent repeat. Since the number of live and dead cells could not be counted on single slices of the CLSM images, the area of the live (green channel) and dead cells (red channel) was calculated using the ImageJ software. In order to account for the area difference between live and dead cells (the stained nucleus of a dead cell is smaller than the stained area of a live cell) the area of the live fraction was divided by a factor of 3 (rough approximation). The dead-cell fraction was then calculated based on these areas. The sum of the total areas in the red and green channels were considered 100%. Biomineralization assay. Biomineralization was induced by adding osteogenic medium containing 7.5 mM β-glycerophosphate and 50 µg mL-1 ascorbic acid in 10% FBS/McCoy’s 5A cell medium. This cell media was exchanged every 2 – 3 days. Calcium deposition in the spheroids was quantified after 3, 7 and 14 days using the ortho-cresolphthalein complexone


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(oCPC) method. To extract the calcium, the spheroids were first washed with PBS, then transferred to a transparent 96-well plate and incubated in 100 µL 1.0 M HCl for at least 2 h at room temperature. Depending on the mineralization of the spheroids, 10 or 20 µL of sample or calcium standard were mixed with 2.0 or 4.0 µL of 5.0 M NaOH to neutralize the acid. 200 µL of calcium reagent solution (500 mM 2-Aminoamino-2-methylpropanol (AMP), 32 µM oCPC, 0.275 mM 8-hydroxychinoline, pH 10.5) was added and incubated for 10 min at 37 °C. The CaCl2 standards (0–100 µg mL-1) were generated by serial dilutions of a 100 mM CaCl2 stock solution. Subsequently, the absorbance was assessed in a multimode plate reader (λ = 575 nm). Micro-computed tomography (µCT). The spheroids were placed in custom made small cups made of poly-ether-imide. They were scanned in a desktop µCT scanner (µCT 35, Scanco Medical, Brüttisellen, Switzerland) with 1000 projections/180°, an isotropic voxel size of 3.5 µm, X-ray voltage of 45 kV, current of 88 µA, and an integration time of 800 ms. The threedimensional data sets were low-pass filtered using a Gaussian filter (σ = 0.8, support = 1) and segmented with a fixed threshold filter (487.6 mg HA/cm3). The volume of the spheroid was determined as the volume of the pixels above the filtering threshold and the density of each of these voxels were determined using the software supplied with the scanner. Quality assurance was performed by weekly (density) and monthly (geometry) scans of the solid-state calibration phantom provided with the scanner. Three-dimensional imaging were performed using Amira (version 5.6.0; FEI, Mérignac, France). Statistical analysis. At least three independent repeats were performed for all experiments. Data are displayed as mean ± standard deviation and indicating the number n of independent repeats. The statistical significance used to compare the distribution was determined using a one-


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way ANOVA followed by a Tukey’s multiple comparison posthoc test (* p < 0.05, **p < 0.01, ***p < 0.001).

RESULTS AND DISCUSSION SaOS-2 cells and alginate microparticles spheroids. The core requirement to employ artificial assemblies to support their biological counterpart is their integration in cell culture. In this context, we chose SaOS-2 human osteosarcoma cells as an established model with biomineralization ability49-50 and capability to secrete matrix vesicles.51 Although 2D cell cultures on surfaces represent the standard cell study system, a spheroid model was selected because the cells are prevented from attaching to the well surface and are forced to float, hence cell-to-cell and cell-to-particle interactions are promoted. Therefore, 3D cell clusters, spheroids, represent more realistic tissue models, which provide a simple way to mimic the physiologically 3D environment and the important cell-cell contacts crucial for cell fate and, thus, tissue formation.52-53 In addition, it is envisioned, that thousands of cell-specific spheroids can be fused to bioprint complex tissues. With the aim to assess the interaction of SaOS-2 cells and microreactors, alginate particles (ME) with a diameter of ~50 µm were produced via D-µF and employed as generic representatives (Figure 1A). Alginate is a popular scaffold material in bone tissue engineering,22 due to the benign cross-linking via calcium ions, its biocompatibility, biodegradability, non-immunogenecity and ability to form highly porous hydrogels to allow for transport of oxygen, nutrients and waste products.54 D-µF is an approach that facilitates controlled generation of diverse monodisperse particles with nearly-100% loading efficiency.55


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With the aim to determine if SaOS-2 cells co-assemble with uncoated ME (i.e. without positively-charged poly-L-lysine (PLL) coating, ME-PLL) into a spheroid, ME and SaOS-2 cells were mixed at a particle-to-cell ratio of 1:20 using 104 cells and cultured for 24 h in roundbottom well plates with ultra-low attachment surfaces. For comparison, SaOS-2 cells only spheroids (S-C) were made. While S-C formed stable spheroids (Supporting Information Figure S1A), only the cells formed a spheroid and all the ME-PLL were located next to the spheroid showing that the ME-PLL did not co-assemble with the cells (Supporting Information Figure S1B/C). An established possibility to promote cell adhesion is using positively charged surface modifications, which were shown to enhance cell differentiation, mineralization and TNAPexpression in osteoblast-like cells.56 Therefore, ME-PLL were coated with different concentrations of PLL i.e., approximately 200 ME-PLL µL-1 were incubated with PLL concentrations between 0.1 and 1 mg mL-1. The PLL concentration was limited to 1 mg mL-1 since higher PLL concentrations led to surface wrinkling,20 which would represent an additional parameter affecting the cell-to-particle attachment. Bright field microscopy images of spheroids containing PLL-coated ME revealed that stable spheroids formed after 24 h when the PLL concentration was ≥ 0.4 mg mL-1 (Figure 1B). Below that concentration, the spheroids showed patchy, unstable cell aggregates, suggesting weak cell-particle interactions (Supporting information Figure S2 and S3). Therefore, all subsequent spheroids were assembled using ME coated with 0.6 mg mL-1 PLL in a cell-to-ME ratio of 20:1 using 104 cells and are referred to as S-ME. In an attempt to quantify the effect of ME on the spheroid development, the spheroid diameter was measured for up to 14 days (Figure 1C) in comparison to S-C. In general, the spheroid diameters decreased within the first three days due to spheroid compaction, before it remained constant. Furthermore, the S-ME were significantly larger than the S-C on day 2 and 3, but on


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day 7 they were in size. This finding was probably due to a compression force inside the spheroid, which led to shrinkage of ME and thus, to a further decrease in S-ME size. Calculating the volumes of the spheroids by using the measured diameter and assuming a perfect sphere, the size of S-ME was ~1.8× larger than S-C after day 3. This difference in spheroid volume resulted in a calculated mean ME diameter of ~37 µm, which was ~20% smaller than the size of the assembled ME supporting the hypothesis of ME compression upon spheroid compaction. In order to visualize the integration of the ME into the spheroids, S-ME were fixed and stained for confocal laser scanning microscopy (CLSM) imaging. The pictures revealed close attachment of the cells to ME with their cytoskeleton surrounding the ME (Figure 1D). The cells formed a dense actin filament network throughout the entire spheroid and around ME, illustrating that the cells formed cell-particle contacts in addition to the cell-cell contacts yielding in the development of stable spheroids.

Figure 1. Characterization of S-ME. A) Representative bright-field image of ME. Scale bar is 100 µm. B) Representative bright-field image of S-ME after 6 h at a cell-to-ME ratio of 20:1. Scale bar is 500 µm. Inset: Integrated ME are visible at the edge of the spheroid. C) Time dependent


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change in S-ME and S-C diameter. Values are represented as mean ± SD, n = 3-9, *p < 0.05. D) Representative CLSM image of a fixed and stained S-ME (blue: DAPI-stained cell nucleus; red: phalloidin-stained actin filaments). Scale bar is 25 µm.

In a next step, the cell viability of S-C was compared to S-ME. Live-dead staining with fluorescein diacetate (FDA) and propidium iodide (PI), respectively, was used on spheroids after 3 d and 7 d in culture followed by visualization using CLSM. After 24 h, the growth media was changed to an osteoconductive media (i.e., cell media supplemented with 7.5 mM βglycerophosphate and 50 µg mL-1 ascorbic acid) to induce mineralization. As expected, S-C had viable cells merely in the periphery with a dense necrotic core, which increased from 3 d to 7 d (Figure 2A, top). In contrast, S-ME exhibited a less pronounced and more inhomogeneous necrotic core after 3 d and 7 d (Figure 2A, bottom). With the aim to quantify this aspect, the livecell fraction was estimated from the area of dead (red channel) to live (green channel) cells in the CLSM images taken ~20 µm inside the spheroids (Figure 2B). It has to be noted that it was impossible to take images deeper inside the spheroids due to loss of fluorescence intensity caused by the dense spheroid structure and mineralization. S-ME had a live-cell fraction of ~80% and ~50% on day 3 and 7, respectively, while S-C had significantly lower cell viability on both days (~60% and ~30% on day 3 and 7, respectively). The reduced cell viability of S-C was likely due to its dense structure, which inhibited diffusion of nutrients to the core. In contrast, the ~2× higher cell viability in S-ME might be explained by the presence of ME, which added low density areas that probably facilitated better access to nutrients, especially for the cells in the inner core of the spheroid. Taken together, the integration of ME might not only reduce the number of


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required cells to obtain a certain tissue volume/mass, but had also a beneficial impact on the mammalian cells by improving the short-term viability.

Figure 2. Cell viability. A) Representative CLSM images of live S-C and S-ME after 3 d and 7 d in culture. (green: FDA (live cells); red: PI (dead cells)). Scale bars represent 100 µm. B) Estimated live-cell fraction of S-C and S-ME after 3 d and 7 d. Values are represented as mean ± SD, n = 4-5, **p < 0.01.

Assembly of microreactors.


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Biocatalytic active subunits. The structurally beneficial ME were equipped with biocatalytic activity with the aim to enhance the biomineralization of the spheroids. Since MVs secreted from osteoblasts are the biologically relevant active moiety in bone mineralization, MVs isolated from SaOS-2 cell cultures were considered as the functional subunits embedded in the ME. Furthermore, we generated artificial matrix vesicles (AMVs) as an alternative active entity to MVs. Since TNAP is one of the major enzymes involved in bone mineralization and highly overexpressed in osteoblasts, we equipped liposomes with TNAP as a bottom-up approach. The artificial liposomal membrane was composed of a mixture of zwitterionic 1-palmitoyl-2-oleoylsn-glycero-3-phosphocholine (POPC) lipids and negatively-charged 1-palmitoyl-2-oleoyl-snglycero-3-phosphoserine (POPS) lipids at a molar ratio of 4:1. This lipid composition was chosen because POPS is enriched on the inner leaflet in biological MV membranes, and POPS binds to Ca2+ ions.27 In addition, TNAP naturally possesses a lipid anchor, the so-called glycosylphosphatidylinositol (GPI) linker, which causes the enzyme to attach automatically to lipid membranes. Therefore, the assembled artificial matrix vesicles (AMV1) possessed a TNAP-corona around the outer liposomal membrane in addition to the encapsulated enzymes. Furthermore, the GPI-anchor can be cleaved by a phoshphatidylinositol-specific phospholipase C enzyme in order to obtain artificial matrix vesicles with only encapsulated TNAP (AMV2).40, 57 In order to possibly achieve triggered mineralization inside AMV2 upon the enzymatically cleavage of the TNAP substrate β-GP to release Pi, the liposomes were further loaded with calcium. Employing more than 5 mM CaCl2 during assembly, the liposomes started to precipitate. The mean hydrodynamic diameters assessed by dynamic light scattering were similar for all three types of vesicles (~230 nm), but the polydispersity indices for AMV1 and AMV2 were much lower than that for MVs, indicating a narrower size distribution obtained by the


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extrusion process (Supporting Information Figure S4 and Table S1). The morphology of AMV1 (Figure 3Ai), AMV2 (Figure 3Aii) and MVs (Figure 3Aiii) were visualized by transmission electron microscopy (TEM) and confirmed the presence of vesicular structures. MVs showed less homogenous vesicles and other structures, most probably short collagen fibers, which were a side-product from the MV extraction. Subsequently, the catalytic activity of AMV1, AMV2 and MV was compared using the model substrate 4-methylumbelliferyl phosphate (4-MUP), which was converted into the fluorescent product 4-MU upon cleavage of the phosphate group by TNAP (Figure 3B and Supporting Information Figure S5). MVs showed the highest activity, followed by AMV1 and AMV2 exhibiting 24× and 60× lower conversion rates, respectively (Supporting Information Figure S5 and Table S2). Comparing the activity of AMV1 and AMV2 to each other showed that AMV1 had a ~2.5× higher activity likely due to the TNAP corona around the liposomal membrane. It should be noted that AMV1 and AMV2 had similar concentrations but MVs were likely higher concentrated, an issue originating from the different vesicles sources. Quantifying the MVs concentration is challenging since MV samples likely contained diverse lipids and proteins. Furthermore, when lysing AMV1 and AMV2 with 0.1% Triton-X (v/v) to release TNAP into solution, the AMV1 and AMV2 activity increased by ~5× and ~10×, respectively (Supporting Information Figure S6 and Table S2, top row). First, this finding confirmed the successful encapsulation of TNAP. In addition, the larger increase in catalytic activity of lysed AMV2 suggested that the TNAP was efficiently cleaved from the outer membrane. On the other hand, the activity of MVs before and after exposure to Triton-X was similar probably due to TNAP being largely attached to the outer lipid bilayer of the vesicles and/or better transport of the substrate across the lipid bilayer.


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Since the conversion of the model substrate 4-MUP to the fluorescent product 4-MU has only limited biological relevance, AMVs and MVs in solution were tested for their ability to spontaneously form mineral by turbidity measurements (Figure 3C).35 While mineral formation was observed for MVs in the presence of 2.0 mM Ca2+ and 3.42 mM Pi after 30 min, the turbidity of the MV solution without Ca2+ remained unchanged, illustrating the preserved mineral formation ability in the former case. As positive control, 4% DMSO in solution containing Ca2+ and Pi induced mineral precipitation as expected. On the other hand, AMVs did not exhibit any measurable turbidity change i.e., they were not able to induce mineral formation.

Figure 3. Characterization of the biocatalytic active subunits. A) Representative TEM images of AMV1 (i), AMV2 (ii) and MVs (iii). Scale bars represent 200 nm. B) Activity in solution of AMV1, AMV2 and MVs assessed by monitoring the time dependent conversion of 4-MUP into the fluorescent product 4-MU. C) Mineral formation in solution by MVs. Turbidity change over


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time was measured at 340 nm. Samples were incubated at 37 °C in mineralization assay buffer containing 3.42 mM Pi with or without 2 mM Ca2+. 4% DMSO in mineralization assay buffer was used as positive control to induce spontaneous precipitation.

Multi-compartmentalized microreactors. In the next step, the active subunits were encapsulated in the microparticles by mixing the vesicle stock solution with the alginate stock solution (1:1 vol. ratio) prior to the microreactor generation using D-µF yielding in MAMV1, MAMV2 and MMV. The morphology of these three microreactors was compared to ME using bright field microscopy (Figure 4A and Supporting Information Figure S7). While ME (Figure 4Ai) had a smooth morphology, MAMV1 and MAMV2 appeared grainy (Figure 4Aii/iii), likely caused by aggregated AMV1 and AMV2 originating from their mixing with the alginate solution, due to the high shear forces generated at this step.58 Even when the viscous alginate solution was slowly added to the liposome solution, aggregation was observed, which became progressively worse with time in the D-µF channels. However, this precipitation effect was not observed for MMV (Figure 4Aiv), probably due to different components in the MV stock solution, such as collagen fragments, etc., resulting from the MV extraction. In order to confirm the co-localization of TNAP and AMV1 in the microreactors, TNAP was fluorescently labeled with DyLight 633 and the liposomes were marked using the fluorescent lipid analogue NBD-PC (1-palmitoyl-2-(6-[(7nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl)-sn-glycero-3-phosphocholine) during assembly. Double-labeled AMV1 within the alginate microparticles were visualized by CLSM and the presence of both, the lipids and TNAP were confirmed (Supporting Information Figure S8). Further, the fluorescent signal from the lipids and TNAP overlapped, indicating that TNAP remained attached/entrapped to/in the liposomes. The co-localization of the two fluorescent


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channels from the CLSM images was illustrated by the total Pearson correlation coefficient of ~0.75. The performances of the microreactors were compared by assessing the reaction kinetics of 104 microreactors per well to convert 4-MUP to 4-MU (Figure 4B). First, all three types of microreactors were found to be active, illustrating that the encapsulation of the functional vesicles into the alginate particles was possible. However, the 4-MUP conversion rates of MAMV1 and MAMV2 were ~20× slower than that of MMV (Supporting information Figure S5). Specifically, MAMV1 and MAMV2 converted ~5% of the substrate after 5 h, while MMV already converted over 50% in the same time, which agreed with the higher activity of the vesicles in solution in the latter case. Overall, it was estimated that the activity of all types of vesicles was ~2 – 3× lower when encapsulated compared to the vesicles in solution (Supporting information Table S2). Furthermore, adding Triton-X to the microreactors released most of the enzymes from the alginate microparticles i.e., TNAP was released into the surrounding medium and became accessible for the substrate, illustrating the assembly of multi-compartmentalized microreactors (Supporting information Table S2). MAMV1 and MAMV2 exposed to Triton-X had ~10× higher activity comparted to the untreated counterparts. However, MMV incubated with Triton-X only exhibited a ~3× higher conversion. Taking into consideration that the activity of the MVs themselves was not affected by Triton-X, encapsulating them into alginate particles seemed to have affected them to a higher extent than AMV1 and AMV2. Nonetheless, (partially) structural integrity of the encapsulated MVs could be concluded. Since MMV were the most active microreactors, only they were further considered. The long-term activity of MMV was assessed for up to 10 days since that is the relevant time frame when aiming to kick-start biomineralization. The activity decreased to 50% within 4 days,


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but remained at this level until day 10 (Supporting Information Figure S9), showing that the MVs remained active and trapped in the microreactors. Due to the superior performance of MMV, both in terms of activity and mineralization ability of the subunits, MMV were selected for integration with SaOS-2 cells into spheroids with the aim to enhance biomineralization.

Figure 4. Characterization of multi-compartmentalized microreactors. A) Representative bright field microscopy images of ME (i), MAMV1 (ii), MAMV2 (iii), and MMV (iv). Scale bars represent 20 µm. B) Time dependent conversion of 4-MUP to the fluorescent product 4-MU of 104 ME, MAMV1, MAMV2, and MMV in solution (Vtot = 200 µL).

Active Microreactors in SaOS-2 cell spheroids. With the aim to confirm that MMV can impose a beneficial effect on biomineralization within spheroids, MMV and SaOS-2 cells were mixed at a particle-to-cell ratio of 1:20 using 104 cells and cultured in round-bottom well plates with ultra-low attachment surfaces. These spheroids are referred to as S-MMV. For comparison, S-C and S-ME were employed. The biomineralization was assessed via measuring the mineral content by quantifying the total calcium content per spheroid on days 3, 7 and 14 after the


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addition of osteoconductive media using the colorimetric o-cresolphtalein complexone (oCPC) method.59-60 At least three single spheroids per independent repeat were removed from the culture well plates and washed with Ca2+-free PBS in order to remove free Ca2+ from the solution. First, the spheroids were visualized by bright field microscopy (Figure 5A). All types of spheroids clearly mineralized in osteoconductive media, shown by their non-transparent appearances. The microreactors were still visible at the edges of S-ME and S-MMV even after 14 days of incubation (Figure 5Aii and iii). This observation confirmed that the microreactors were tightly integrated in the spheroids – the prerequisite for extended support of the SaOS-2 cells. Initial experiments to quantify Ca2+ of S-C grown in normal cell media (i.e. without osteoconductive additives), were analyzed and revealed at least 2× lower calcium concentrations compared to S-C grown in osteoconductive media. Therefore, the spheroids for biomineralization quantification were grown in osteoconductive media targeting for the most optimal outcome possible in our current set up. In order to quantify the Ca2+ content per spheroid, the mineralized matrix was dissolved in 1.0 M hydrochloric acid. After neutralizing with sodium hydroxide, the dissolved Ca2+ complexes to oCPC and the total calcium was calculated using a standard curve of known calcium concentrations (Supporting Information Figure S10). The total Ca2+ per spheroid of an independent repeat was obtained by calculating the mean value of at least three individual spheroids. The relative calcium content was then calculated for S-C on day 14 as the reference, representing the maximum mineralization without support of microreactors. First, the mineral content in S-C, S-ME and S-MMV increased over time. (Figure 5B). Second, the S-ME and S-C mineralized at the same rate within the first 7 days, but S-ME contained significantly more mineralized matrix after 14 days. We hypothesize that the higher number of viable cells in S-ME


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after 7 days of incubation largely contributed to the increased biomineralization of S-ME. On the other hand, the S-MMV already showed significantly increased calcium content on day 3 compared to S-ME, indicating that the MVs had an impact on the biomineralization process from the beginning. On day 14, there was ~50% and 20% more calcium present in S-MMV compared to S-C and S-ME, respectively, illustrating the sustained effect the MVs exhibited on the biomineralization. Further, the rate of biomineralization was found to be linear between day 3 and 14 (Supporting Information Figure S11).While the mineralized matrix in S-C increased at a rate of 8.3 ± 0.5% per day, S-ME and S-MMV resulted in increases of 10.5 ± 0.2% and 12.3 ± 0.4% per day, respectively (Supporting Information Table S3).

Figure 5. Calcium quantification of mineralized spheroids: A) Representative bright field images of i) S-C, ii) S-ME and iii) S-MMV grown for 14 days in osteoconductive media. Scale bars represent 200 µm. B) Relative total Ca2+ per spheroid measured by the o-CPC method after


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3, 7 and 14 d incubation time. The total Ca2+ content of S-C on day 14 was set as 100%. Values are represented as mean ± SD, n = 3-8, *p < 0.05, **p < 0.01, ***p < 0.001.

In order to further quantify the biomineralization, micro-computed tomography (µCT) was used to generate scans of the spheroids at a spatial resolution of 3.5 × 3.5 × 3.5 µm3 on day 7 and 14. To the best of our knowledge, this is the first reported analysis of mineralization inside artificially made spheroids using µCT. µCT measures the absorbance of X-rays of the sample and provides a 3D image of the spheroid with different absorbance values for each voxel. By doing so, the mineralized spheroid volume (SV) and at the same time the mean density of the spheroids could be determined. In order to calculate the SV, we set a lower threshold value to define at which absorbance value a given voxel could be considered as being mineralized. In the present study, a lower threshold of 487.6 mg HA cm-3 was applied, where HA stands for hydroxyapatite. For comparison, in a study of mice distal femoral metaphyses a threshold of 495.1 mg HA cm-3 was used to segment bone from bone marrow.61 In this way, the volume of calcified tissue can be obtained in the same way as for trabecular bone. Furthermore, by averaging the density of each voxel, which had a density above the threshold, it is possible to ascertain the mean spheroid density. Prior to assessment by µCT, the spheroids were washed in PBS and fixed in 4% PFA before being transferred to a sample holder, which contained small plastic cups with approximately 3.0 µL volumes, where the spheroids were placed in PBS for scanning by µCT. Two to three individual spheroids were recorded per independent repeat, and the SV was calculated from the single repeats. On day 7, the density of the spheroids was close to the detection limit of the defined threshold, illustrated by the µCT images showing a very low SV and low mineral density


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for S-C and S-ME (Supporting Information Figure S12). In contrast, spheroids on day 14 showed densely mineralized structures in all cases. While S-C were typically spherical (Figure 6Ai), the shape of S-ME (Figure 6Aii) and S-MMV (Figure 6Aiii) could differed from spheroid to spheroid, exhibiting often a slight elongation. Further, S-C and S-ME revealed low-density areas, indicating inhomogeneous biomineralization. µCT of S-MMV showed a homogeneously mineralized spheroid even within the core, demonstrating the beneficial effect of MMV. The SV obtained by µCT (Figure 6B) provided an alternative to quantify the biomineralization of the spheroids in addition to the Ca2+-quantification. As expected, SV increased over time for all the spheroids, but S-C and S-ME did not show any significant differences on day 7 and 14. However, S-MMV exhibited a significant increased SV compared to S-ME on both days. Furthermore, the SV of S-MMV on day 7 was already at the same level as for S-C on day 14. On day 14, S-MMV had a ~300% increased SV compared to S-C. Further analysis of the µCT data of the spheroids revealed that not only the SV but also the mean spheroid density is increased (Supporting Information Figure S13). The density was obtained based on the SV and the calculated mean X-ray absorption value for each voxel. As expected, the spheroid density confirmed the beneficial effect of MVs within the microreactors on the biomineralization rate in SaOS-2 spheroids. When comparing the findings from the calcium quantification and µCT, S-MMV performed significantly better in both cases, but there is a rather large discrepancy in relative changes between the two methods. Specifically, µCT suggested a higher impact of the MMV on the biomineralization than the o-CPC-based calcium quantification method. This difference might be explained by the fact, the SV obtained from µCT was based on a defined threshold, while there might be already mineralized parts inside the spheroids, which were not yet dense enough to be


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included in the SV. However, this low-density mineral might be detectable by the o-CPC-based calcium quantification method.

Figure 6. µCT analysis of spheroids. A) Representative µCT images of S-C (i), ME-S (ii) (cut in the middle plane) after 14 d incubation time. Scale bars represent 100 µm. B) Relative spheroid volume (SV) determined by µCT. S-C on day 14 was set as reference value (100%). Values are represented as mean ± SD, n = 3-8, **p < 0.01, ***p < 0.001.

CONCLUSION In summary, we have developed ~50 µm alginate-based microreactors with encapsulated MV as support for SaOS-2 cells to enhance biomineralization in co-assembled microreactor-cell spheroids. First, S-ME improved not only the viability of the SaOS-2 cells but also the biomineralization after 14 days compared to S-C. Next, we compared the performance of SaOS-


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2 derived MVs and their mimics AMV1/2 and found that only MVs allowed for the assembly of efficient microreactors with preserved function over at least 10 days. Finally, S-MMV had higher mineral content after 7 and 14 days compared to S-C and S-ME as assessed by the colorimetric oCPC-based method and µCT. Artificial cells as partners to their biological role models with a long and active residence time in a tissue offer a promising alternative in future biomedical technologies. With due further developments such as investigations into gene and protein expression (e.g., TNAP overexpression or phosphorylation status of osteopontin) of S-MMV during maturation and mineralization, microreactors and spheroids reported here might provide the basis for 3D bioprinted bone tissue as the next generation materials to be integrated in critical-sized bone defects.

ASSOCIATED CONTENT The Supporting Information is available free of charge: Detailed bright field images of spheroids, spheroid diameter change over time, DLS data, relative activities of active subunits and corresponding microreactors, kinetic of Triton-X lysed subunits, bright field images of microreactors, CLSM images of double-labelled subunits in alginate beads, long-term activity of MMV, calcium standard curve, biomineralization rate and calculated mineralization slopes, µCT images of 7 days spheroids, relative mean spheroid density (PDF). AUTHOR INFORMATION Corresponding Author *Corresponding author. E-mail address: [email protected] (BS).


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ORCID 0000-0002-4161-2383 (Fabian Itel) 0000-0001-9386-6679 (Jesper Skovhus Thomsen) 0000-0002-7335-3945 (Brigitte Städler)

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding Sources This work was supported by grants from the Aarhus University Research Foundation, Denmark and the Swiss National Science Foundation (P2BSP2_161912) (F.I.). ACKNOWLEDGMENT This work was supported by grants from the Aarhus University Research Foundation, Denmark and the Swiss National Science Foundation (P2BSP2_161912) (F.I.). REFERENCES 1.

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TOC


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Scheme 1 80x92mm (300 x 300 DPI)



Figure 1 80x79mm (300 x 300 DPI)


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Figure 2 80x137mm (300 x 300 DPI)



Figure 3 85x119mm (300 x 300 DPI)


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TOC 28x7mm (300 x 300 DPI)


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Matrix Vesicles-Containing Microreactors as Support for Bone-Like

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