Enzyme-induced mineralization of calcium carbonate in 3D printable granular hydrogels

1 Enzyme-induced mineralization of calcium carbonate in 3D printable granular hydrogels Francesca Bono1,2, Anna Puiggalí-Jou3, Lorenzo Lucherini1, Greta Cocchi1, Marcy ZenobiWong3, and Esther Amstad1,2* 1Soft Materials Laboratory, Institute of Materials, École Polytechnique Fédérale de Lausanne (EPFL), CH-1015 Lausanne (Switzerland) 2Swiss National Center for Competence in Research (NCCR) Bio-inspired Materials, University of Fribourg, Chemin des Verdiers 4, 1700 Fribourg, Switzerland 3Tissue Engineering + Biofabrication Laboratory Department of Health Sciences & Technology, ETH Zürich (Switzerland) * Corresponding author, email: [email protected] Keywords Biomineralization, Granular Hydrogels, 3D Printing, Calcium Carbonate Saffolds Abstract Many biological materials, such as bone, are organic-inorganic composites, made from a polymeric matrix that supports biomineralization under mild conditions. These materials are usually composed of a small set of abundant components like polysaccharides, proteins, and minerals and exhibit a remarkable combination of density normalized stiffness, toughness, and functionality. Producing bio-inspired synthetic porous composites with a similar combination of properties through energy-efficient processes still presents an unmet challenge. Some aspects of this challenge can be addressed using living bacteria that induce biomineralization. However, formulations containing such living bacteria limit biomedical applications especially in vivo, require careful handling, and are costly. To address these limitations, we introduce enzyme-containing granular precursors exclusively made from naturally sourced polymers. These precursors can be cast or direct ink written into cm-sized structures before they are mineralized under benign conditions to reach CaCO3 contents up to 92 wt% with a porosity of 56 vol%. The resulting mineralized scaffolds exhibit a specific compressive strength of 5.15 MPa·g-1·cm3 and a compressive modulus of 72 MPa·g-1·cm3. These values are similar to those of human trabecular bone, which has a similar porosity, despite that our synthetic composites are based on CaCO3 that is weaker than calcium phosphate, which constitutes the main component of human bone. The resulting biomineralorganic composites show low cytotoxicity. These findings highlight the potential of this approach to 3D print biocompatible CaCO3-based composites under mild conditions. We envisage this formulation to open up new possibilities for tissue engineering. 2 Introduction Many biological materials are made from a limited number of naturally sourced abundant components such as polysaccharides, proteins, and minerals through energy-efficient processes that are conducted under ambient conditions. Despite these severe limitations in material selection and processing conditions, many biological materials exhibit a fascinating combination of density normalized stiffness, toughness and functionality [1]. A prime example is nacre, which contains 95 wt% aragonite, a calcium carbonate (CaCO₃) polymorph that is surrounded by a proteinaceous organic matrix [2]. The brick-and-mortar architecture of nacre, composed of stiff aragonite platelets and a soft protein-based organic matrix, provides high stiffness and exceptional toughness—enhancing the latter by up to 1000-fold compared to pure aragonite crystals [2–5]. Similarly, spines of sea urchins exhibit well-defined structures spanning multiple length scales that render them damage tolerant. These spines are composed of the thermodynamically most stable calcium carbonate polymorph, calcite, amorphous calcium carbonate, and organic molecules [6,7]. Nature frequently uses such strong and tough mineral-based materials as defense shields[8], structural support[9], or for buoyancy regulation [10]. Calcium carbonate is one of the most abundant minerals: it constitutes up to 4 wt% of Earth's crust [11]. This mineral primarily forms through the sedimentation of skeletal constituents that originate from marine organisms, such as shellfishes [12], which contain up to 95 wt% CaCO3 [13].To reduce the density of minerals, which is essential for an energy-efficient locomotion of creatures, natural biominerals often contain pores. A paradigmatic example of porous biominerals are bones that incorporate pores of different sizes and shapes to minimize their density while maximizing their strength and torsion resistance [4,14]. Inspired by the excellent density normalized mechanical properties of such biological minerals, synthetic mineral-based composites have been introduced. Nacre-inspired examples, which display a high fracture toughness, include cement [15] or glass [16] platelets that have been infiltrated with synthetic polymers [15,16]. However, in many cases, these synthetic composites are produced through energy-consuming processes that include high temperature treatments or exposure to vacuum. This shortcoming can be addressed using foams or emulsions as templates [17], or through freeze-casting [18]. However, many of these methods still involve thermal treatments to rigidify the structures and those, that do not require thermal treatments involve exposure to vacuum or freezing steps that are also energy consuming, and far from the biogenic mechanism of living organisms. To address these limitations and inspired by nature, the mineralization of organic scaffolds has been triggered by living organisms like bacteria. For example, ureolytic bacteria such as Sporosarcina Pasteurii [19] have been used for microbially induced calcium carbonate precipitation (MICP) [20] for geo-environmental 3 [21,22] and construction applications[ 23,24] to form bulk clay [25] and CaCO3-based ceramics [26]. The mineral formation can be spatially confined by embedding bacteria in soft gels that can be cast into simple shapes [27]. If formulated as bacteria-loaded microgels, these materials can even be direct ink written into more complex 3D structures before being mineralized [28]. However, formulations containing living bacteria such as Sporosarcina Pasteurii (S. Pasteurii) are delicate to handle and costly, limiting biomedical applications and hampering in vivo implementations [29,30]. To address these limitations, bacteria can be replaced by enzymes such as urease, which are more cost effective. Urease has thus far mainly been used for sand cementation [31]. Recently, urease has been used to stiffen synthetic hydrogels by precipitating FeCO3 [32], amorphous[33] and crystalline CaCO3 [34,35] However, the processing of these structures was limited to casting. Here, we introduce a biocompatible granular ink that can be direct ink written into cm-sized intricate structures, which are subsequently mineralized under ambient conditions to result in composites containing up to 92 wt% CaCO3. These composites possess open pores whose volume accounts for up to 56 % of the entire structure, thereby resulting in low densities. The high mineral content combined with the high porosity results in a specific compressive strength of 5.15 MPa·g-1·cm3 and a specific modulus up to 72 MPa·g-1·cm3; these values are within the range of human trabecular bone that exhibits a similar porosity [36–38]. This combination of assets is achieved by formulating urease-loaded hydrogel microfragments exclusively from naturally-sourced polymers. These enzyme-containing fragments are jammed to form an ink that can be direct ink written in air and at room temperature before the scaffold is mineralized. The structure and morphology of the forming CaCO3 can be tuned with the composition of the microfragments. The resulting mineralized scaffolds exhibit low cytotoxicity towards osteoblasts, indicating their potential for tissue engineering applications. Results and discussion We use hydrogels as mineralization scaffolds. To ensure good biocompatibility of our formulation, we employ naturally sourced polymers, namely gelatin and κ-carrageenan. To enable their mineralization, we functionalize them with urease. Precursor solutions for gelatin and κ-carrageenan cannot be direct ink written close to room temperature if no additives[39– 42] or thermal gradients in the printer [43] are present. To enable 3D printing of these hydrogels at room temperature, we formulate them as microfragments. We produce microfragments from bulk hydrogels through cryo-milling to minimize risks for residues that might hamper cell viability, as shown in Fig. 1 i [44,45]. The microfragments are polydisperse with irregular shapes, as sketched in Fig. 1 ii. Fragments are firmly connected through a 4 second network formed by soaking them in phosphate-buffered saline (PBS) containing 4 wt% alginate (Fig. 1 iii). Alginate offers an additional benefit: it has a high affinity to Ca2+, such that it also serves as a Ca2+ source during mineralization [28]. The enzyme-loaded microfragments, which are surrounded by alginate, are jammed to obtain an ink that can be 3D printed in air at room temperature. The 3D printed structure is incubated in a solution containing 1.75 M urea and 1 M CaCl2. Urease contained within the microfragments catalyzes the hydrolysis of urea into ammonia (NH3) and CO2. Because the microfragments are immersed in aqueous solutions, these products partially transform into ammonium (NH4+) and carbonate ions (CO32) [22], which, in the presence of Ca2+, initiate the mineralization of the hydrogel, as graphically shown in Fig. 1 iv-vi. The mineralization is completed within 4 days, as demonstrated by the cm-sized model of a Nautilus half-shell in Fig. 1 vii and detailed in Fig. S1a-b. We employ gelatin as a hydrogel because it contains motifs that facilitate cell adhesion, migration, differentiation, and proliferation [46–48]. Gelatin is a denatured collagen derivative that is liquid at temperatures above 36 °C. We produce gelatin from an aqueous solution containing 25 wt% gelatin and 10 mg/g urease at 40 °C. If cooled to room temperature, gelatin structurally changes from coiled to a helix, resulting in its solidification [49]. 5 Fig 1 Schematic illustration of the fabrication of an enzyme-containing granular ink. (i) Urease is encapsulated into a bulk hydrogel (ii) that is cryo-milled to produce fragments. (iii) The fragments are suspended in an alginate solution and jammed. (iv) The enzyme-containing granular ink is 3D printed (v) before it is soaked in a solution containing urea and calcium chloride (vi) to mineralize the scaffold. Mineralization is triggered by the enzyme-induced hydrolysis of urea. (vii) Photograph of a 3D printed mineralized Nautilus half-shell. Scale bar is 2 cm To evaluate the influence of the mechanical properties of the bulk hydrogel on its degree of mineralization, we quantify the compressive modulus. The bulk gel comprising 25 wt% gelatin exhibits a compression modulus of 0.13 MPa, as shown in by the pink curve in Fig. 2a. To assess if the gel relaxes stress, we perform stress relaxation tests by subjecting the gel to 1% strain and quantifying the time required to decrease the stress by 50%, τ1/2. The relaxation time τ1/2 of gelatin amounts to 298 s, as shown in pink in Fig. 2b. This relaxation time is similar to that of various living tissues [50]. To enable direct ink writing of hydrogels, we cryo-mill them into fragments with an average equivalent diameter of 73 μm, as shown in Fig. 2c. These fragments are swollen in PBS containing 4 wt% alginate. To allow direct ink writing of the microfragments, they are jammed through centrifugation, as detailed in the Experimental Section. The resulting ink is shear thinning and displays a fast stress recovery, as shown in Fig. 2d-e. Moreover, the strain at the yield point, where the storage modulus G’ is equal to the loss modulus G’’ is 26%, as shown in Fig. 2f. This low yield point enables printing at low pressures that are readily accessible with commercially available bioprinters, such that these microfragment-based inks are wellsuited for direct ink writing [44,44,51–53], in good agreement with inks composed of jammed synthetic microfragments [44,54–61]. 6 Fig 2 Characterization of microgel fragments. a) Compression curves of bulk gelatin (pink) and κ-carrageenan (green) before fragmentation. The extracted compression moduli are shown in the inset. b) Stress relaxation of gelatin and κ-carrageenan bulk hydrogels. c) Optical microscopy image of (i) gelatin- (i) and (ii) κ-carrageenan-based fragments with the corresponding size distributions. d) Frequency sweeps of jammed gelatin and κ-carrageenan fragments. e) Shear recovery measurements showing the self-healing ability of the ink if 7 subjected to alternate low (0.3%, white areas) and high (80%, shaded areas) strains. f) Amplitude sweeps of the jammed gelatin and κ-carrageenan fragments To assess if the granular gelatin-based structure can be mineralized, we incubate it in a solution containing CaCl2 and urea and visualize the mineral formation with optical microscopy. Within 18 hours, we observe some minerals within the extruded filament. The density of minerals increases with incubation time, as shown in Fig. 3a. These minerals are crystalline, as indicated by their birefringence revealed with polarized optical microscopy in Fig. 3b. Calcium carbonate exists in different polymorphs that exhibit distinct mechanical properties [18,62,63]. To evaluate the structure of the formed CaCO3, we perform X-ray diffraction (XRD). Within 24 h of incubation, we observe diffraction peaks at 29.6°, 39.49°, 43°, and 47.6° and 48.6°, characteristic of calcite[26], as shown in Fig. 3a. This result is supported by scanning electron microscopy (SEM) analysis, which reveals cubic crystals, as shown in Fig. 3d. FTIR spectroscopy further confirms the calcite structure with peaks at 712 cm-1, 874 cm-1 and 1396 cm-1, as shown in Fig. 4a. Note that the peak at 1396 cm-1 is slightly shifted from the usual asymmetric stretch vibration at 1425 cm-1 possibly due to hydration.[64] After 4 days of mineralization, small diffraction peaks at 24.9°, 27.2°, 32.8°, and 36° characteristic of vaterite[26], appear in the XRD spectra, as shown in Fig. 3c. We hypothesize that small amounts of amorphous calcium carbonate (ACC) are present at any point during the mineralization of gelatin. Part of this ACC might transition into vaterite that possibly is stabilized by the organic phase [65,66]. These findings are in contradiction to those reported for S. Pasteurii bacteria containing gelatin-based microparticles, which formed almost the same quantity of vaterite and calcite after 1 day of mineralization, albeit after 4 days of mineralization, the majority of the mineral was also calcite [28]. A possible reason for this discrepancy during the early stages of mineral formation is that enzymes do not require a metabolic recovery period of 48–72 hours before they start forming the thermodynamically most stable polymorph, calcite [28]. 8 Fig 3 CaCO3 characterization. a) Optical microscopy images of extruded filaments composed of gelatin- (pink) and κ-carrageenan-based (green) fragments showing the growth of the minerals over time. All the scale bars are 500 μm. b) Polarized optical microscopy images of 9 CaCO3 minerals grown in the gelatin- (top) and κ-carrageenan-based granular scaffold (bottom) after 4 days of incubation. The birefringence is more evident in the gelatin-based scaffold where the CaCO3 crystals are bigger, as shown in the close-up. All scale bars are 200 μm. c) XRD spectra of CaCO3 grown in (i) gelatinand (ii) κ-carrageenan-based granular gels measured after 1 day (D1), 2 (D2), 3 (D3), and 4 (D4) days of mineralization. The gray shades indicate the peaks characteristic for calcite (29.6 °, 47.6°, 39.49°, 43°, and 48.6°), the blue shades those characteristic for vaterite (24.9°, 27.2°, 32.8°, and 36°). d) SEM image of CaCO3 crystals grown in the (i) gelatinand (ii) κ-carrageenan-based granular scaffold showing the cubic calcite, labeled with white arrows, and residues of dry polymer, highlighted by green arrows To quantify the amount of formed minerals, we perform thermogravimetry analysis (TGA). Dried samples that had been mineralized for one day contain 79 wt% minerals, as shown in Fig. 4b. The mineral content increases with increasing incubation time until it plateaus after 4 days of mineralization. Remarkably, if mineralized for at least 4 days, dried composites contain up to 92 wt% minerals, as shown in Fig. 4b. This mineral content is similar or higher than that achieved in previously published mineralized hydrogels, indicating the potential of our approach to fabricate highly mineralized composites. Functional groups present within the hydrogel can influence the CaCO3 formation and growth [62,67]. To assess the influence of the hydrogel composition on the mineral formation, we formulate microgels from κ-carrageenan. κ-carrageenan is a polysaccharide that is derived from red algae and has shown minimal cytotoxicity [68]. It contains one sulfate per repeat unit which can form complexes with different cations including calcium [69]. If dispersed in hot water, κ-carrageenan attains a coil configuration that transitions into a helix upon cooling, resulting in its gelation [70,71]. A bulk gel comprising 4 wt% κ-carrageenan exhibits a compression modulus of 0.1 MPa, a value similar to that of gelatin, as shown in the green curve in Fig. 2a. Yet, its relaxation time is with τ1/2 = 116 s significantly shorter than that of gelatin, as shown in Fig. 2b. Cryo-milled κ-carrageenan fragments are slightly larger than the gelatin-based ones with an equivalence diameter of 92 μm, as shown in Fig. 2c. As a result of this difference, jammed κ-carrageenan fragments display a strain at the flow point of 10%, as shown in Fig. 2f. This value is lower than that of gelatin-based counterparts such that these fragments can be printed at even lower pressures. To assess the influence of the hydrogel composition on the mineralization, we mineralized jammed, 3D printed urease-containing κ-carrageenan fragments and visualize the formation of the minerals with optical microscopy. By analogy to the gelatin fragments, minerals start to form within 24 h and continue to form for at least 48 hours, as shown in green in Fig. 3a. A close inspection of the optical micrographs reveals that minerals form within the κ-carrageenan filaments as well as at their surfaces. This rather homogeneous mineral distribution is in stark contrast to that observed for gelatin-based counterparts where minerals are initially preferably located at the filament surface, as shown in Fig. 3a. Remarkably, polarized optical microscopy 16 Fig 6 3D printing of the enzyme-containing granular inks. Photographs of a) gelatin-based fragments containing urease printed in air into (i) a brittle star that is incubated (ii) in an aqueous solution containing CaCl2 and urea to trigger mineralization. (iii, iv) Photographs taken after four days of mineralization in the (iii) wet and (iv) dry state. b) Mimicking the behavior of the brittle star fish, (i) the broken limb (ii-iii) can be re-attached by connecting the broken parts with new ink, (iii-iv) stained with red food dye for better visualization, and remineralizing. c) κ-carrageenan-based fragments containing urease printed in air into (i) a tower shell, where (ii) the cavities are maintained after complete mineralization and drying. The same 17 κ-carrageenan-based ink is used to 3D print a Stellaria solaris shell (iii, iv). All scale bars are 1 cm. d) Volumetric shrinkage of the CaCO3-gelatin (squares in pink palette) and CaCO3-κcarrageenan (circles in green palette) scaffolds as a function of the mineralization time We consider our samples to be potentially suitable for tissue engineering applications. To assess their biocompatibility, we culture MG-63 (osteoblasts-like cells) onto fully mineralized urease-containing microfragment-based scaffolds and characterize their viability and morphology over a 7-day culture period. Live/Dead staining reveals that CaCO3-gelatin and CaCO3-κ-carrageenan scaffolds support high cell viability at Days 1 and 7, as shown in Fig 7a. Cell viability is consistently above 80% for both scaffold types, with gelatin scaffolds showing slightly higher viability, as shown in Fig 7b. To further assess cellular morphology we perform scanning electron microscopy (SEM) on the samples.[83] At day 1, cells in gelatin and κ-carrageenan-based scaffolds exhibit a predominantly rounded shape, as indicated by the arrows in Fig 7a. As expected, by day 7, cells on gelatin scaffolds appear more spread and integrated into the calcite scaffold, resulting in larger cell areas and higher aspect ratios (ARs) whereas cells in κ-carrageenan scaffolds retain their spherical morphology with minimal changes, as shown in Fig 7d and Fig 7c. These results reveal good biocompatibility and higher cellular adhesion for the gelatin-based formulation, due to its cell-adhesive motifs. These results demonstrate the potential of our approach to design scaffolds compatible with cells and their environment without affecting their viability and potentially promoting their spreading and integration. 18 Fig 7 Cell viability and morphology of CaCO3-gelatin and CaCO3-κ-carrageenan scaffolds over time. a) Live/Dead staining (green: live cells (calcein), red: dead cells (propidium iodine)) and SEM micrographs of cells (pointed with white arrows) cultured in CaCO3-gelatin and CaCO3κ-carrageenan-based scaffolds on Day 1 and Day 7, showing differences in cell distribution, viability, and morphology. b) Quantification of cell viability (%) at day 1 and day 7, demonstrating higher viability in gelatin-based hydrogels (pink) compared to κ-carrageenanbased hydrogels (green) at both time points. c) Violin plots of cell area (μm²) indicating increased spreading of cells in CaCO3-gelatin (pink) scaffolds over time compared to CaCO3κ-carrageenan (green). d) Violin plots of cell aspect ratio (AR, arbitrary units), highlighting differences in cell elongation between the two scaffold types. Data are presented as mean ± SD and analyzed with unpaired ttest, significant differences are marked with **p < 0.01, ***p < 0.001, ****p < 0.0001 19 Conclusion We introduce urease functionalized gelatin and κ-carrageenan microfragments that, if jammed, can be direct ink written into cm-sized 3D structures in air and at room temperature. These structures, reach CaCO3 contents up to 92 wt% and a porosity up to 56 vol% if mineralized for 4 days. The high mineral content and porosity impart the resulting composites a specific compressive modulus up to 72 MPa∙g-1∙cm3 and a compressive strength up to 5.15 MPa∙g-1∙cm3. These values exceed those of previously reported 3D printed CaCO3-based porous scaffolds. Our ink is composed of naturally sourced reagents, namely gelatin and algae-derived κ-carrageenan, and contains enzymes that are much more cost-effective and less resource intense to cultivate and maintain than ureolytic bacteria. We envision that these assets, combined with their biocompatibility, render our ink a promising platform for tissue engineering, or to heal or repair broken minerals. Data Availability The raw and processed data required to reproduce these findings are available to download from Zenodo. Data Availability Data will be made available on request. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgements The authors acknowledge Albert Taureg and Lionel Pittet from PIXE Platform for the help with X-Ray μCT scannings and data analysis and David Wen Hua Bi for the help with the XRD experiments and data analysis. The authors would like to acknowledge the financial support of the bio-inspired Materials NCCR through the Swiss National Science Foundation (51NF40205603). The authors would like to thank Dr. Ran Zhao and Dr. Matteo Hirsch for the fruitful scientific discussions. Author Contributions 20 F.B. and E.A. designed the experiments and conceived the concept. A.P.J. and M.Z.W. designed the cell experiments. F.B. performed the experiments with the help of L.L. for the SEM imaging and G.C. for the TGA. A.P.G. performed the cell experiments. F.B. and E.A. analyzed the data and wrote the manuscript. A.P.J. and M.Z.W. analyzed the cell-related data and wrote the relevant section of the manuscript. All the authors discussed the results and approved the final version of the manuscript. Materials and Methods Materials. κ-carrageenan (ThermoFischer, 431590250), gelatin from porcine skin (gelatin Type-A from porcine skin (gel strength 300, Sigma-Aldrich, G2500), urea (Sigma-Aldrich, 51456), urease from Canavalia ensiformis (Jack bean) (Sigma-Aldrich, U1500-20KU), alginic acid sodium salt from brown algae (Sigma-Aldrich, A1112), phosphate buffered saline (PBS) (Gibco), calcium chloride (Carl Roth, CN93.1), Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco), fibroblast growth factor 2 (FGF-2, PeproTech), fetal bovine serum (FBS) (Gibco), MG63 cells (osteosarcoma cell line, ATCC #CRL-1427; ATCC, Wesel, Germany), Propidium Iodide (PI) (Fluka). Microfragments preparation. Microfragments were prepared by mechanical grinding via cryomilling gelatin and κ-carrageenan bulk hydrogels that were prepared as follows. A solution was obtained by dissolving 25 wt% gelatin at 40 °C or 4 wt% κ-carrageenan in water at 70 °C. The concentrations have been selected to obtain gels with similar rheological and mechanical properties.10 mg of urease was weighed per g of gelatin solution and dissolved in 100 μL of water just before use. The enzyme was poured into a beaker containing the gelatin or κcarrageenan solution (T = 40°C) and stirred. The solution was poured into a glass mold and cooled to room temperature to solidify the gelatin or the κ-carrageenan. The obtained bulk gel with the encapsulated enzyme was stored at 4°C. The bulk gels were fragmented with an oscillatory cryomill (Cryomill, Retsch) using 1 stainless steel ball (d = 25 mm). The cryomilling protocol is as follows: 3 min of precooling at 5 Hz; 1 milling cycle of 1 min at 30 Hz. The prepared fragments were freeze-dried and stored at 4°C. Ink preparation. Dry enzyme-loaded microfragments were resuspended in a 4 wt% alginate solution in PBS with a 1:4 volume ratio of dry fragments: alginate solution for 1 hour at 4 °C. To remove the supernatant the suspension was centrifuged for 10 minutes at 3700 rpm and 7 °C ( Mega Star 1.6R, VWR) . The obtained ink was stored at 4 °C and used for 3D printing and casting. Ink biomineralization. Once 3D printed or casted into the final shape, the ink was gelled by exposing it dropwise to a large excess of 1 M CaCl2 solution until submerged. After 30 minutes the ink was transferred into the incubation aqueous solution containing 1 M CaCl2 and 1.5 M 21 urea mixed in a 1:1 volume ratio. The solution was exchanged every 24 h for four days. After the fourth day, samples were removed from the incubation solution, soaked in water for 25 minutes, and dried in a vacuum chamber at room temperature overnight. Rheology measurements. Rheological measurements were performed on a DHR-3 TA Instrument rheometer with a 8 mm diameter parallel plate steel geometry. All measurements were performed at 25°C, with a 1000 μm gap and repeated on 3 distinct samples. Amplitude sweeps were performed at 10 rad∙s−1 for a strain range of 0.01 to 200%. Frequency sweeps were performed within the linear viscoelastic regime at a strain value of 0.5 % in a frequency range of 0.1-50 Hz. Shear recovery measurements were performed at 10 rad/s alternating 200 s at 0.3% with 200 s at 80% strain. Stress relaxation measurements on bulk hydrogels were performed at the constant strain of 1%, a value picked in the linear viscoelastic regime, for 16 minutes. The stress is normalized at 0.1 s. All the represented data were obtained from three independent samples and are reported as mean ± SD. XRD measurements. The regular theta-theta scans for all the samples were performed on a Panalytical Empyrean X-Ray polycrystalline diffractometer in Bragg-Brentano geometry, equipped with long-focused seal Cu X-Ray tube (λKα = 1.5418 Å), and PIXcel 1D X-Ray detector. The patterns were collected in continuous mode between 5 and 60 degrees (2θ), with the step-size of 0.02626 degree. The baseline removal and peak search were performed with the Peak Analyzer tool in OriginPro2021. Optical imaging. Optical microscopy images were obtained with a Nikon Eclipse TS100 microscope. The diameters of the microfragments were calculated as the average of the two diagonals measured with Image J. Polarized optical microscopy images were obtained with a polarizing microscope (ECLIPSE Ti-DH, Nikon). Micro computed tomography. X-Ray μCT was performed on an Ultratom micro tomography system (RX-SOLUTIONS). The dry samples were scanned at a voxel resolution of 1.05 mm, with a voltage of 45 kV and a current of 166 mA. Amira-Avizo v.2019.4 software was used for reconstruction, segmentation, and visualization. Image J is used to calculate the mineral, pore and gel fraction over the entire volume of each sample. All the results in Fig. S3 are reported as mean ± SD of all the obtained slices for each sample. All the measurements were performed on samples obtained by casting in a 4 mm x 2 mm mould. FTIR characterization. FTIR spectra were measured on a Spectrum 3 spectrometer (PerkinElmer) in the attenuated total reflectance (ATR) mode, and corrected for the 22 background and CO2 signals. Traces were acquired between 4000 and 650 cm−1 at a resolution of 4 cm−1. All the measurements were repeated on 3 distinct samples grinded in a fine powder. Volume shrinkage and density measurements. The volume shrinkage percentage (Vs) was calculated as follow: 𝑉𝑉 𝑠𝑠= 𝑉𝑉𝑖𝑖−𝑉𝑉𝑓𝑓 𝑉𝑉𝑖𝑖 x 100 Where Vi is the initial volume calculated by measuring diameter, thickness and mass of 3 distinct cylinders as prepared in a 6 mm x 2 mm mold, after removing the excess water from the surface, and Vf is the final volume of the same set of 3 samples measured in the same way after overnight drying in a vacuum chamber. Thermogravimetric analysis. Biomineralized samples were finely grinded in a mortar before testing with TGA (TGA 4000, PerkinElmer). The measurements were performed from 30 °C to 850 °C at a heating rate of 10 °C min-1 with an air flow rate of 20 mL min-1, and holding at 850 °C. The CaCO3 weight percentage (wt%CaCO3) was obtained with the following formula: 𝑤𝑤𝑤𝑤%𝐶𝐶𝐶𝐶𝐶𝐶𝐶𝐶3= 𝛥𝛥𝛥𝛥𝛥𝛥% 𝑚𝑚𝑚𝑚𝑚𝑚2 x 𝑚𝑚𝐶𝐶𝐶𝐶𝐶𝐶𝐶𝐶3 Where Δwt% is the weight percentage loss calculated between 600 °C and 850 °C, mCO2 is the molecular weight of CO2 (44.102 g∙mol-1) and mCaCO3 is the molecular weight of CaCO3 (100.1 g∙mol-1). All the samples were rinsed with water for 15 minutes and dried in a vacuum chamber overnight before testing. The represented data were obtained from at least three independent samples and are reported as mean ± SD. The full curves used for the calculations are reported in Figure S9. Compression measurements. Compression tests were performed on cylindrical samples obtained by casting the ink in 8 mm diameter and 6 mm tall molds. All the samples were rinsed with water for 15 minutes and dried in a vacuum chamber overnight before testing with a commercial machine (ZwickiLine, 5 kN load cell, Zwick Roell), compressed at a constant velocity of 5 mm/min until 80% strain was reached. The compression modulus was calculated as the slope of the initial linear region, from 5% to 12% strain. All the reported results were obtained from at least three independent samples and are reported as mean ± SD. 3D printing. Jammed gelatin-alginate or κ-carrageenan-alginate microfragments loaded with urease enzyme were loaded in a 3 mL Luer lock syringe. The syringe was centrifuged at 3500 rpm for 1 minute to remove trapped air bubbles at 7 °C. 3D printing was carried out with a 23 commercial 3D bioprinter (BIO X, Cellink) at 12 mm/s and 60 kPa or 70 kPa. The printing bed was maintained at 20 °C, unless stated differently. The granular ink was extruded through a blunt needle (20G, DT = 603 μm) using a pressure driven piston, unless stated differently. Cell Expansion and Seeding. MG-63 cells (osteosarcoma cell line, ATCC #CRL-1427; ATCC, Wesel, Germany) were expanded in a proliferation medium consisting of Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% (v/v) FBS, 100 µg/mL gentamicin (Gibco), and 5 ng/mL fibroblast growth factor 2 (FGF-2). Cells were maintained at 37 °C in a humidified atmosphere with 5% CO₂ and passaged upon reaching 80% confluency. Cells were trypsinized, resuspended in DMEM containing 10% (v/v) FBS and 10 µg/mL gentamicin, and collected by centrifugation (5 minutes at 500 × g). After counting, 100,000 cells were seeded on each composite construct (casted in a 4 mm x 2 mm mould) and cultured for 7 days. Live/Dead Imaging. Samples were incubated in DMEM supplemented with 1:1000 Calcein AM (Invitrogen) and 1:500 Propidium Iodide (PI) for 40 minutes without FBS, followed by three washes with phosphate-buffered saline (PBS). Imaging was performed using a Leica SP8 microscope (Leica) equipped with a 10× dry objective. Z-stacks were acquired from the sample surface at 4 µm intervals, extending 100 µm into the sample. The images presented in this study were generated using maximum intensity z-projection. Scanning Electron Microscopy (SEM). SEM imaging on dry mineralized scaffolds at the end of the incubation period and on dry gelatin/alginate and κ-carrageenan/alginate controls containing the same concentration of enzyme were performed on a Zeiss Gemini 300 at an operating voltage of 3 kV with 10 nm Au/Pd coating using a secondary electron detector. Samples seeded with cells, after the designated culture period, were washed with PBS and fixed with 4% paraformaldehyde (PFA) for 1 hour. 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