Dual carbon sequestration with photosynthetic living materials

Domain: earth_science
Published: 2025-04-23
Editorial outcome: major_revision

Abstract
Natural ecosystems efficiently sequester CO2 but containing and controlling living systems remains challenging. Here, we engineer a photosynthetic living material for dual CO2 sequestration that leverages biomass production and insoluble carbonate formation via microbially induced carbonate precipitation (MICP). To achieve this, we immobilize photosynthetic microorganisms within a printable polymeric network. Digital design and fabrication of the living structures ensures sufficient light access and nutrient supply to encapsulated cyanobacteria, enabling long-term culture for over a year. We showcase that photosynthetic living materials are able to sequester 2.2 ± 0.9 mg of CO2 per gram of hydrogel material over 30 days and 26 ± 7 mg of CO2 over 400 days. These findings highlight the potential of photosynthetic living materials for scalable, low-maintenance carbon sequestration with applications in carbon-neutral infrastructure and CO2 mitigation.

Introduction
Biological ecosystems, such as forests, aquatic systems, and wetlands, offer efficient pathways for carbon sequestration (storing carbon in a carbon pool) and conversion into carbon-based materials1. Natural systems operate under ambient conditions with sunlight and commonly available small molecules as their sole inputs. Further, living systems can sense, self-repair, and respond to their surroundings, making them resilient to environmental changes1,2. Biological carbon sequestration, for example via afforestation or the growth of marine phytoplankton and algae, is also cost-efficient and environmentally friendly3. In this context, natural carbon sequestration can serve as a passive, low-impact complement to industrial carbon sequestration, which normally requires specific, extreme, and energy-intensive conditions1,4 and proximity to large emission sources5. However, natural carbon sequestration is typically slower than industrial carbon sequestration, and the control of living systems outside of their native environments is often challenging5,6.

Strategies to engineer living systems for active CO2 sequestration would provide an additional approach to mitigate the accumulation of human-generated CO2 in the atmosphere. The CO2 concentrating mechanism of many photosynthetic microorganisms accumulates CO2 within the cell body up to 1000-fold above ambient levels6,7. Subsequently, concentrated carbon can be fixed in the form of biomass generated during growth8,9. In addition to biomass production, microbially-induced carbonate precipitation (MICP) in certain species can sequester CO2 irreversibly in the form of inorganic carbonate precipitates. MICP proceeds via multiple metabolic pathways, including ureolysis, sulfate reduction, and denitrification10,11. In some organisms, MICP can occur as a direct by-product of photosynthesis, whereby the inorganic precipitates effectively act as an additional carbon sink12, enabling dual carbon sequestration. In this context, immobilizing photosynthetic microorganisms, such as algae and cyanobacteria, within a support matrix may provide an approach to drive biological CO2 sequestration in the form of engineered photosynthetic living materials via dual carbon sequestration.

To date, engineered living materials have primarily been used for applications in biomedicine, sustainable materials production, and as living building materials13–17. For example, MICP has been exploited, primarily via ureolysis, to mechanically reinforce living materials based on the in situ formation of a stiff mineral phase18. Robust composites were produced via biomineralization using ureolytic MICP within a cellulose matrix19,20. Similarly, precipitates deposited in porous materials filled cracks and improved mechanical properties of composite building structures as well as consolidated soils21. Ureolytic MICP is attractive due to its short incubation period (typically 1–4 days), resistance to contamination, and rapid biomineralization; however, it poses substantial environmental concerns due to the associated production of large amounts (1–2 equimolar) of ammonia22. Further, ureolytic MICP requires a constant supply of urea and only proceeds in a narrow range of environmental conditions23–25. These challenges constrain the use of ureolytic MICP for long-term CO2 sequestration26. Many of these limitations can be addressed with photosynthetic MICP, which requires no additional feedstocks and produces no toxic by-products18,25. Recently, photosynthetic MICP was used to design living building materials that were reinforced by mineralization over time17. While photosynthetic living materials have been explored for carbon sequestration via reversible biomass accumulation9, they have not been explored for CO2 sequestration via biomass accumulation and irreversible MICP using atmospheric CO2 as the main carbon source and light as the sole source of energy.

In this work, we engineer photosynthetic living materials for dual CO2 sequestration by immobilizing MICP-capable photosynthetic Cyanobacterium Synechococcus sp. strain PCC 7002 within a printable Pluronic F-127 (F127)-based polymeric network. Dual carbon sequestration via biomass generation and insoluble carbonate formation proceeds over the lifecycle (beyond one year) of the bio-printed structures. The mineral phase mechanically reinforces the living materials and stores sequestered carbon in a more stable form. This work provides a strategy to engineer photosynthetic living materials as a scalable CO2 sequestration method to complement ongoing strategies to mitigate atmospheric CO2 accumulation.

Results
Engineering living materials for dual carbon sequestration
We engineered photosynthetic living materials for dual carbon sequestration by exploiting the carbon concentrating mechanism of cyanobacterium strain PCC 7002 (Fig. 1a). Dual CO2 sequestration proceeds via both reversible biomass accumulation and irreversible mineral precipitation. For carbon concentration to occur, CO2 from the surrounding environment dissolves in aqueous solutions and forms bicarbonate ions (HCO3−) that are then transported to the marine β-cyanobacteria 7002 carboxysome27. Within the carboxysome, carbonic anhydrase (CA) catalyzes the conversion of bicarbonate into CO2, which is then fixed by ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCo) during photosynthesis into two molecules of 3-phosphoglycerate. This product is enzymatically converted into sugars that support cell growth and biomass production, representing the reversible part of CO2 sequestration.

Irreversible CO2 sequestration as mineral precipitation occurs outside the cells, near the cyanobacteria. Negatively charged extracellular polysaccharides on the bacterial membrane combined with a suitable extracellular environment (alkaline pH, presence of divalent cations), facilitates the nucleation and formation of insoluble carbonates27–31. Thus, by culturing photosynthetic living materials in simulated seawater that contains Ca2+ and Mg2+, an increase in medium pH, which occurred in cyanobacteria-containing samples (Supplementary Fig. 1), allows CO32− in the medium to be fixed irreversibly into calcium or magnesium carbonates. Chemical equilibrium then favors the dissolution of additional atmospheric CO2 into the culture medium, continuously driving the dual carbon sink.

Cyanobacteria PCC 7002 were encapsulated in a hydrogel matrix to fabricate photosynthetic living materials capable of dual carbon sequestration (Fig. 1b, Supplementary Fig. 2). A synthetic polymeric hydrogel based upon Pluronic F-127 (F127) was chosen for the encapsulation of cyanobacteria due to its bioinert nature and its processing versatility. F127-based hydrogels have been used in the design of engineered living materials due to the facile diffusion of most small molecules through it32. Functionalized F127-bisurethane methacrylate (F127-BUM) can be photo-cross-linked either post-printing or processed directly via light-based additive manufacturing for long-term structural stability (Fig. 1c). In order to structure the photosynthetic living materials using digital fabrication, we designed a bioink (13.2 wt% F127 and 7.3 wt% F127-BUM) that maintained high viability of encapsulated PCC 7002 (Supplementary Fig. 3) and printability via direct ink writing and light-based additive manufacturing. Rationally designed hydrogel network allows the access of light into the scaffold, thereby enabling homogeneous bacteria growth throughout the entire structure.

Bioink transparency is required for efficient light transmission to drive photosynthesis28. Light attenuation due to absorption and scattering within the bioink is one of the main challenges to overcome during cultivation of encapsulated photosynthetic species33. The F127-based hydrogel transmitted light (76 ± 3%) over the entire visible wavelength range (400–750 nm); the BG11–ASNIII medium used to culture PCC 7002 had 95 ± 4% transmittance in the same range (Fig. 1d). The addition of the photoinitiator lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP), which absorbs light below 420 nm, reduced transmittance only at low wavelengths that are not relevant for PCC 7002 photosynthesis (Supplementary Fig. 4)34. The encapsulation of cyanobacteria decreased the transmittance of the final construct to 28 ± 8% and 31 ± 9% before and after cross-linking, respectively, due to scattering and productive absorption of light by the photosynthetic bacteria (Supplementary Fig. 5). Overall, the optical properties of the photosynthetic living material were suitable to enable cyanobacteria viability and growth when encapsulated within F127-based hydrogel.

To structure the photosynthetic living materials for enhanced carbon sequestration and longevity, we designed the F127-based hydrogel for both direct ink writing and light-based additive manufacturing (Supplementary Figs. 6 and 7). The ink exhibited shear-thinning (shear-thinning index n = 0.09, n < 1, Supplementary Table 1) and elastic recovery (~90%) after high shear, demonstrating its feasibly for extrusion-based printing (Supplementary Figs. 8 and 9). To stabilize printed structures for long-term use, the hydrogel mixture was photo-cross-linked (λ = 405 nm; I = 8 mW cm−2; t = 60 s) to increase the final storage modulus (G′) of the construct (Fig. 1c, e, Supplementary Figs. 10 and 11).

Dual CO2 sequestration capacity of photosynthetic living materials
Dual CO2 sequestration in the photosynthetic living materials was provided by PCC 7002 biomass growth and insoluble carbonate precipitate formation through MICP. To further investigate both aspects of carbon sequestration, uniform circular samples (Vsample = 40 µL, d = 10 mm, thickness l = 0.5 mm) were printed using 22 G conical nozzles (Ø = 0.41 mm) via direct ink writing and incubated for 30 days (Fig. 2a). During the incubation, BG11–ASNIII medium was changed every 5 days, and the dissolved inorganic carbon (DIC) was quantified in the collected samples (Supplementary Method 1). From day 5, the concentration of Ca2+ in the medium was set to 8.65 mM via CaCl2 addition to simulate natural seawater conditions35 that promote MICP. 40.4 mg L−1 (0.38 mM) of Na2CO3 was dissolved in the medium to simulate natural seawater. Analysis of the total DIC under equilibrium in the culture medium indicated that the DIC content was above or at the quantity of inorganic carbon initially dissolved in the form of CO32− (Supplementary Fig. 1). This increase in the DIC confirmed the dissolution of atmospheric CO2, which together with dissolved Na2CO3 in the simulated seawater medium, served as the carbon sources for CO2 sequestration.

PCC 7002 biomass growth within the living material for CO2 capture was evaluated by fluorescence microscopy (cell counts) and chlorophyll extraction (Fig. 2b). After 10 days of incubation samples exhibited visible clusters of PCC 7002 cells as well as many individual cells throughout the volume of the hydrogel matrix. The number of clusters and individual cells per unit volume of living material (written as mLgel−1) increased over time with cell concentrations reaching 5 × 109 cells mLgel−1 after 30 days (Fig. 2d, Supplementary Fig. 12). Further, the choice of initial OD730nm, within the range of 0.3–1.6, did not impact cell growth (Supplementary Fig. 13a). Moreover, the majority of the encapsulated bacteria remained within the hydrogel throughout the incubation in biotic samples (Supplementary Fig. 13b). Beyond 30 days, we expect the biomass accumulation to reach a pseudo-steady state (with similar cell death and growth rates) based on the plateau observed in the cell concentration beyond day 25.

To confirm CO2 sequestration via the production of insoluble carbonate precipitates via MICP, precipitates were visualized using calcium staining. In the presence of divalent ions such as Ca2+ and Mg2+, cyanobacteria go through the biomineralization process to form gypsum, calcite, and magnesite36. We confirmed the accumulation of Ca2+ in the hydrogels using Alizarin Red S staining (Fig. 2c). On day 0, abiotic and biotic samples were mainly orange after staining with green PCC 7002 cells visible in the biotic samples, indicating minimal Ca2+ accumulation initially. The abiotic samples remained orange throughout the 30-day incubation (Supplementary Fig. 14). Biotic samples stained red by day 10, indicating Ca2+ accumulation in the whole volume of the hydrogel and the dark red color was observed around growing bacteria clusters over time. From the analysis of Alizarin Red S staining, we observed that the amount of precipitates was equivalent to > 1 mg of CaCO3 mLgel−1 already after 10 days (Fig. 2e). It reached 4.8 ± 2 mg of CaCO3 mLgel−1 by the end of the 30-day period. Precipitates were also observed in the bottom of the incubation wells, and all analyses were carried out in fresh wells to avoid any influence of precipitation outside of the living material on our quantification (Supplementary Fig. 15). To assess the amount of biomass and precipitate formation, the mass of abiotic and biotic samples was measured and compared (Fig. 2e). The initial decrease in sample mass was attributed to the diffusion of unfunctionalized F127 polymer out of the printed discs, which also caused high variability in the data of day 1 and day 5. Over the incubation period of 30 days, the biotic samples had approximately 36% more dry mass than control abiotic samples with significant increase (P = 0.001) in dry mass observed from day 10 of incubation. In total, biomass and carbonate precipitates in biotic samples accounted for approximately 45% of the final sample mass (Supplementary Method 2).

To quantify the extent of precipitate generation, the organic biomass and polymer matrix were removed by thermal decomposition. As the temperature (T = 600 °C) was above the decomposition temperature of organic matter (polymer and biomass) and below the decomposition temperature of carbonate compounds37, only the insoluble carbonate precipitates remained after thermal decomposition. The mass of the inorganic precipitates corresponded to 50 µmol (2.2 ± 0.9 mg) of CO2 sequestered via MICP per gram of hydrogel (Supplementary Fig. 16 and Supplementary Method 2). CO2 sequestered via MICP has been stored in a more stable mineral form.

Composite materials form during the lifecycle
Having demonstrated that photosynthetic living materials are capable of CO2 sequestration via both biomass accumulation and carbonate formation, we further investigated the composition of the mineral phase produced during incubation. MICP results in the formation of crystalline carbonates that vary in stability and solubility with calcite being the most stable polymorph and, therefore, desirable for carbon sinking purposes (Fig. 3a)38. TGA analysis confirmed carbonate presence in the precipitates (Fig. 3b), which was additionally confirmed by FTIR (Supplementary Fig. 17). XRD analysis of precipitates in biotic samples revealed a highly crystalline structure with the main peaks corresponding to calcium–magnesium carbonates indicated by the position of the main diffraction peak representing the (104) plane (Supplementary Fig. 18). The observed shift likely indicated compositional variation due to the incorporation of magnesium in the carbonate structure as previously observed in marine environments and, in this case, due to the use of simulated seawater (ASNIII medium)39. The XRD was obtained after thermal decomposition (T = 600 °C, which is below the decomposition temperature of CaCO3)37 as the signal from the F127-based matrix was much stronger than the one of the precipitates prior to decomposition (Supplementary Fig. 19).

Precipitates were distributed throughout the polymeric matrix (Fig. 3c). Energy dispersive X-ray analysis (EDS) indicated mainly carbon and oxygen in the abiotic matrix, whereas peaks of calcium and magnesium were prominent in the areas with precipitates in biotic samples. Elemental mapping corroborated these data with calcium, magnesium, and oxygen enrichment in overlapping regions near clusters of cyanobacteria, suggesting the pericellular formation of carbonates (Fig. 3d and Supplementary Fig. 20). EDS measurements also indicated that Ca to Mg atom % ratio in the area of the precipitates was 78 ± 5% to 22 ± 5% (number of analyzed samples n = 10), which confirmed that both elements were embedded in the precipitates.

The formation of precipitates in hydrogel matrices via MICP can reinforce the mechanical properties of the material17,40,41. The storage moduli (G′) of the printed biotic samples on day 2 of the experiment were slightly lower compared with abiotic samples (5.4 ± 2 kPa and 6.7 ± 2 kPa, respectively). A significant increase (P = 0.05) in storage moduli (G′) in the case of biotic samples was observed already after 10 days (Supplementary Fig. 21). Over the course of 30 days, the modulus increased significantly to 10.1 ± 1 kPa (P = 0.01) in the biotic case while it did not change for abiotic samples (6.5 ± 1 kPa) (Fig. 3e). Comparable results were obtained for the Young’s moduli (E) of the samples using tensile tests of cast samples (Supplementary Fig. 22). We associated the increase in modulus with the formation of reinforcing inorganic precipitates42, though increases in biomass could also contribute to the change in Young’s moduli43,44. A similar trend was observed for the material toughness. After 30 days, the toughness of the biotic samples increased from 960 ± 110 J m−3 on day 2 to 1120 ± 200 J m−3 on day 30 (Fig. 3f, Supplementary Fig. 23).

3D-printed photosynthetic living structures for dual carbon sequestration
While simple discs served as a standardized shape for the systemic characterization of photosynthetic living materials, the geometry of the living structures was tailored via digital design and fabrication to improve the CO2 sequestration and long-term viability of the photosynthetic living materials. We designed 3D lattice structures with strut sizes between 0.15 mm and 0.70 mm to facilitate gas and nutrient transport within the printed constructs. The transport of gases and liquid medium as well as access to light are essential for the efficient functioning of biologic processes (including carbon sinking) in the photosynthetic living materials9,45. The decrease in light penetration over the course of the life cycle is expected due to increased light absorption and scattering with increased biomass, resulting in reduced proliferation in the volume fraction further from the surface (Supplementary Fig. 24). Drawing inspiration from cellular fluidics46, we employed a lattice design to engineer structures whereby growth medium was passively transported vertically through the construct via capillary forces (Fig. 4a, Supplementary Fig. 25). In this manner, the structure did not need to be fully immersed in the simulated seawater, minimizing medium use, and facilitating gas transport from the surrounding air to the living material. To achieve high resolution and defined internal porosity, light-based volumetric 3D printing was used to fabricate the lattice structures. Centimeter-scale objects were printed with complex geometries and an optical resolution of 28 × 28 µm within tens of seconds (Fig. 4b). The 3D-printed photosynthetic living structure was cultured for over a year, during which it continuously produced chlorophyll and performed dual carbon sequestration (Supplementary Fig. 26). After 30 days of incubation, the structure was able to stand vertically on a flat surface and liquid was actively drawn up via the construct’s internal structure. The printed sample stiffened further over prolonged incubation due to the accumulation of carbonate precipitates. This mechanical enhancement was observable through the structure’s increased capacity to maintain an upright position at day 30, 60, 120, and 365 and a final modulus of 111 ± 7 kPa (Supplementary Table 2). Over the incubation period of 400 days the living structures sequestered 26 ± 7 mg of CO2 per gram of hydrogel material in the form of carbonate precipitates (Supplementary Table 3). This prolonged culture of the photosynthetic living materials showed that dual carbon sequestration can take place beyond 30 days, especially within rationally designed 3D structures.

To demonstrate the scalable production of photosynthetic living materials, direct ink writing was used to print similar lattice structures at a larger scale. A body-centered cubic (BCC) lattice structure (unit cell length l = 5 mm, strut diameter d = 0.12 mm) was printed. The structure passively perfused medium to the top surface while only partially immersed in BG11–ASNIII medium (Supplementary Fig. 27a). After 60 days of incubation, the lattice was thermally decomposed (T = 600 °C). The remaining carbonate precipitates after thermal decomposition retained the shape of the porous lattice structure (Fig. 4c), indicating that carbon sequestration was homogeneous throughout the structure despite being partially immersed in medium.

This living material also has the potential to be applied on existing surfaces as coatings for dual carbon sequestration. Starting from bulk material, we observed 5 mm to be an optimal material thickness to maintain PCC 7002 viability (0.5 g cm−2material) (Supplementary Fig. 27b)9. To maximize the viable volume of the material per surface area, textured surface was designed as 3 × 3 pillar array on a 2 × 2 cm base to minimize self-shielding of light similarly to coral reef structures47. With this design we increased the printed gel volume by 150% while maintaining the same 1 cm2 footprint (to 0.75 g cm−2material) and without compromising bacteria viability compared with a bulk hydrogel (Fig. 4d, Supplementary Fig. 28). This result further highlight show the synergy between living materials and the design of living structures can increase the efficiency of dual carbon sequestration.

Methods
Bacterial culture and encapsulation
The cyanobacteria Synechococcus sp. PCC 7002 (Pasteur Institute, France) were cultured in BG11–ASNIII medium mixture (Supplementary Method 4, and Supplementary Tables 4 and 5) in Erlenmeyer flasks on a shaker plate with constant shaking (150 rpm) under full-spectrum white light (180 µmol photons m−2 s−1) with a 12-h day (on)/night (off) cycle. Optical density of the bacteria suspension at 730 nm (OD730nm) was measured using UV-visible light spectrophotometer (Lambda 35, PerkinElmer) to monitor bacterial culture growth phases. For encapsulation, cyanobacteria were collected at the early exponential growth phase.

Bioink preparation
Stock solutions of 18 wt% F127 in culturing medium, 30 wt% of F127-BUM (Supplementary Method 5 and Supplementary Fig. 31) in Milli-Q water, and 5 wt% LAP (Supplementary Method 6 and Supplementary Fig. 32) in Milli-Q water were mixed at 3:1:0.082 ratio to reach final concentrations of 13.2 wt% F127, 7.3 wt% F127-BUM, and 0.1 wt% LAP. This formulation was optimized to have the lowest polymer content and stable hydrogel after photo-cross-linking (Supplementary Figs. 6 and 10). LAP concentration was chosen to have a high cross-linking rate without compromising cell viability (Supplementary Figs. 33 and 34). For the bioink preparation, cyanobacteria suspension from the early exponential phase was centrifuged at 3500 × g for 5 min, the supernatant was discarded, and the cell pellet was resuspended in the prepared bioink mixture (liquefied on ice) via pipetting to reach a homogeneous solution with an equivalent optical density OD730nm = 0.8, as this was the value measured in our liquid culture at the start of the exponential growth phase. For volumetric printing, an equivalent optical density OD730nm = 0.3 was used to allow sufficient light transmittance for the fabrication. Choice of initial OD did not impact cell growth (Supplementary Fig. 13a). Biotic bioink OD730nm calculation is described in Supplementary Method 7 and Supplementary Fig. 35.

Bioink characterization
The bioink was characterized using strain-controlled shear rheometer (MCR502; Anton-Paar). The samples were loaded on a glass bottom plate with a light source (λ = 405 nm, I = 8 mW cm–2; M405L3, Thorlabs) underneath and measured using 20 mm plate-plate probe with 0.8 mm gap size at 25 °C. Cross-linking kinetics of the abiotic gels was evaluated by using oscillatory time sweep (ω = 10 rad s−1; γ = 0.1%). The samples were irradiated after 60 s of initial oscillation to induce photopolymerization of F127-BUM in the gel mixture until the plateau storage modulus G′ was reached. Bioink printability characterization is described in Supplementary Method 8.

UV-vis spectra of the medium and hydrogel samples were measured using a plastic cuvette (1 cm light path) with a UV-visible light spectrophotometer (Lambda 35, PerkinElmer). The transmittance was measured over the range of 800-400 nm.

Biofabrication
3D samples for the measurements were fabricated using a pneumatic-driven direct ink writing 3D printer (BioX, Cellink). The bioink was loaded into 3 mL cartridges with a 22 G conical nozzle (Ø = 0.41 mm) and the cartridge was heated to 37 °C for 5 min prior to printing. The printing was performed at a pressure range of P = 40–60 Pa and a speed of v = 5–10 mm s−1. Single layer circular disc-shaped samples (Ø = 1 cm, h = 150-350 µm) were modeled in Rhinoceros 3D and G-code generation was done via Slic3r. The more complex lattice structures were designed with Rhinocerous 3D (Rhino 7) and Grasshopper software. The printed disc-shaped samples were cross-linked for 1 min using an LED light source (λ = 405 nm light, I = 8 mW cm−2, Thorlabs GmbH, Germany), followed by a 10 min washing step in BG11–ASNIII medium. The washing medium was discarded, and fresh medium was added to the samples for bacteria growth.

For volumetric printing, a Tomolite tomographic bioprinter (Readily3D, Switzerland) was used64,65. All STL models were generated with Rhino 7 and Grasshopper software. The size constraints for the Tomolite vials were up to 25 mm in height and 14 mm in diameter. The lattice shown in Fig. 4a, b, is 21.85 mm tall and 11.85 mm in diameter, with a volume of 101.87 mm3 and a surface area of 2145.84 mm2. The lattice is radially arrayed into 16 sectors, in 3 concentric rings, and 20 vertical segments. This generated a lattice with 960 pores, with pore diameter from 0.5–1.0 mm, and strut diameters from 0.15–0.70 mm (median diameter 0.20–0.24 mm). The lattices were sliced into tomographical projection planes using a commercial software (Apparite, Readily3D, Switzerland). Bioink (4 °C) was loaded into autoclaved glass printing vials (outer diameter Ø = 20.0 mm, inner diameter Ø = 15.0 mm) and solidified at 25 °C for 5 min before printing. Cross-linking of the bioink was induced by a light source with a 405 nm wavelength with a pre-calibrated light dosage of 94 mJ cm−2 (Supplementary Method 9). The average light intensity during the printing process was set to be 8 mW cm−2. After printing, the vial was cooled to 4 °C to liquify the unpolymerized bioink. The retrieved polymerized structure was immersed in culturing medium to allow removal of excess LAP and residual bioink. The sample was incubated beyond one year with the above-mentioned culturing conditions.

Living material incubation conditions
The printed circular disc-shaped samples were cross-linked for 1 min using an LED light source (λ = 405 nm, I = 8 mW cm−2, Thorlabs GmbH, Germany), followed by a 10 min washing step in BG11–ASNIII medium. After that, printed samples were incubated under the same light conditions as liquid culture. The samples were incubated in BG11–ASNIII medium for 5 days with a change to fresh medium on day 2. The old medium was collected at each change for pH measurements. Fresh BG11–ASNIII medium with an additional 5 mM CaCl2 was added as the calcium-rich culturing medium. The medium was changed and collected every 5 days (on day 5, 10, 15, 20, and 25) until day 30. The collected medium pooled from individual samples (n = 20 samples, biological replicates) was used for pH measurements to evaluate total dissolved inorganic carbon as fully described in the Supplementary Method 1. The samples were inspected using optical microscope (Panthera Classic, Motic) equipped with Plan UC 10x/0.25, 40x/0.65 Ph2 and 100×/1.25 oil objectives and camera (Moticam S, Motic). The images were captured with Motic Images Plus 3.0 90 software.

Biomass quantification and cell viability characterization
To measure the sample mass change, 5 of the freshly printed (day 0) abiotic and biotic discs samples were washed in Milli-Q water for 10 min to remove salt deposited on the surface from the culturing medium. The samples were then dried under a constant laminar flow in a 1.5 mL microcentrifuge tube of known mass to evaporate the water entrapped within the material. The dried samples together with the microcentrifuge tube were weighed and the mass was used as the initial benchmark for characterization. On day 5, 10, 20 and 30, abiotic and biotic samples (n = 5; biological replicates) were randomly selected and washed with the same procedure. The samples were subsequently dried under the same condition before weighing. The mass was normalized to the average mass of the abiotic or biotic discs on day 0.

Cell viability and growth within the hydrogels were evaluated using confocal microscopy for day 1 to 5 and chlorophyll extraction for day 10 to 30 (Supplementary Fig. 36). Dead cells within the printed disc samples were stained using SYTOX Blue dead cell staining. Disc samples were first washed 3 times in 1 mL of MilliQ water then stained using 0.2 µM staining solution diluted with 1x ASNIII medium. The samples were then incubated in the dark for 15 min, washed 3 times with 1x ASNIII medium for 5 min, and stored in the same medium until imaging. The images of SYTOX Blue dead cell staining and cell autofluorescence were acquired with confocal microscopy (TCS SP8, Leica with a 506357 ∞/0.14-0.18/OFN25/E 40x, water objective, 26 slices, 2 µm each). A 405 nm laser was used to excite SYTOX Blue and emission was collected at 450–510 nm (HyD detector). Chlorophyll autofluorescence was excited with a 488 nm laser and was collected at 660–700 nm (HyD detector). Z-stacks of 50.4 µm were used for maximum intensity Z-projections using LAS X (Leica). Separate images (n = 4 for day 0, n = 5 for day 1, n = 3 for day 5–30) of dead cell and autofluorescence were imported in CellProfiler 4.2.5, (https://cellprofiler.org/). Gaussian filter was applied to the images followed by primary object identification using a built-in module (Adaptive Otsu threshold algorithm was applied with 3 classes where middle intensity was considered as background), resulting in a binary image and object count.

Chlorophyll extraction was done on randomly chosen disc samples (n = 5 randomly selected samples) every 5 days starting from day 10. The samples were fully immersed in 0.5–1 mL of isopropyl alcohol and homogenized (IKA T25 Ultra Turrax with S25N-8G-ST inset). The slurry was then sonicated for 85 min followed by centrifugation at 19,000 × g for 10 min. Technical triplicates of 100 µL supernatant were transferred into a 96-well plate and the absorption at 663 nm was measured by UV–Vis plate reader (HIDEX Sense). Suspension samples of different optical densities were used to calculate an equivalent cell concentration from the chlorophyll amount using exponential fitting (Supplementary Fig. 29). A non-linear fitting was chosen as we observed an increase in encapsulated cell concentration with chlorophyll extraction due to the interference of other cyanobacterial pigments, such as phycobiliprotein66–68.

For suspension and medium samples from the well plate, samples were collected every 5 days and the cell number was measured on a Multisizer 4e (Beckman Coulter) equipped with a 30 µm aperture tube (gain: 4, current: 600 µA). For each solution, 25 µL of media were diluted in 10 mL of 0.2-µm-filtered Isoton II solution within a 25 mL Accuvette (Beckman Coulter).

PCC 7002 encapsulation efficiency was quantified using the cell count and chlorophyll extraction results. The encapsulation efficiency was calculated as the percentage of encapsulated cell number to the total number of encapsulated cells and cells in the culture medium. Leakage was defined as the percentage of cells found in the culture medium to the total cell number of encapsulated cells and cells in the culture medium.

Inorganic precipitate characterization
Calcium-containing precipitate deposits in circular disc-shaped hydrogels were stained using 40 mM Alizarin Red S solution (pH = 4.2) in Milli-Q water. Each sample was fully immersed in 2 mL of Alizarin Red S solution for 20 s. The sample was then washed in 200 mL of Milli-Q water 3 times for 5 min and imaged using an optical microscope. To quantify the extent of precipitation, we extracted the Alizarin Red S stain from the samples using 10% acetic acid (0.5 mL per sample, n = 3, biological replicates) with vortexing for at least 1 h. The solutions were then neutralized using 1 M NaOH and 100 µL of the neutralized extraction solution was used to measure the absorbance at 405 nm using a UV-vis plate reader (HIDEX Sense). A calibration curve was generated using pure CaCO3 following the same method to quantify the amount of CaCO3 in the samples based on the Alizarin Red S absorbance (Supplementary Fig. 15a). Alizarin Red S staining was also performed on randomly chosen recovered wells (n = 3, biological replicates) of 24-well plate every 5 days. 0.5 mL of 40 mM Alizarin Red S solution was added to each well for 20 s. Each well was then washed with 0.5 mL Milli-Q water 3 times to collect the precipitates, which were then recovered by centrifugation at 12,300 × g for 10 minutes. The supernatant was then discarded and the remaining precipitate pellet was dissolved in 0.5 mL of 10% acetic acid by pipetting. The solution was neutralized and the absorbance was measured under the same conditions as those of the disc-shaped hydrogel samples.

The presence of carbonates in the precipitates was confirmed by thermogravimetric analysis using a TGA/DSC 3+ (Mettler Toledo). The sample was placed inside a 70 µL alumina crucible and heated from 25 to 900 °C in pure N2 under a constant flow rate of 150 mL min−1. Buoyancy effects were accounted for by performing measurements with an empty crucible. The sample was first heated to 400 °C and held at this temperature for 30 min to remove any bound water, before continuing heating to 900 °C at a heating rate of 5 °C min−1. The same measurement conducted with an empty crucible was taken as the baseline and was subtracted from the measured data.

The extent of carbonate precipitate formation within abiotic and biotic samples was further characterized by thermal decomposition at atmospheric pressure. 5 dry abiotic or biotic samples were loaded into a porcelain crucible and heated to 600 °C (ramp rate = 2 °C min−1) for 2 hours. The samples were then cooled to room temperature and the remaining mass was measured. In addition, the remaining precipitates were collected for attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR, Spectrum Two, PerkinElmer; 1 cm−1 resolution, 32 scans) as well as X-ray diffraction (XRD) analysis with a PANalytical Empyrean diffractometer (Cu Kα radiation, 45 kV and 40 mA) with an X’Celerator Scientific ultrafast line detector and Bragg-Brentano HD incident beam optics. XRD samples were measured over the 2θ range 10–90° for 1 h with a step increment of 0.016°. Results were compared with reference patterns from the ICDD database. XRD pattern baseline removal for analysis is described in the Supplementary Method 10.

Lyophilized samples (Supplementary Method 11) were used for Scanning Electron Microscope (SEM) characterization. Samples were sputtered with 10 nm of Pt/Pd 80/20 with a metal sputter coater (CCU-010, Safematic, Zizers, Switzerland). SEM imaging was done using a secondary electron detector with a scanning electron microscope JEOL JSM-7100F (accelerating voltage V = 12 kV, working distance WD = 10 mm) equipped with an Ametek-EDAX EDS detector (Si(Li) 20 mm). Element analysis was performed with the EDAX Team software.

Living material mechanical properties
Viscoelastic properties of the printed hydrogels at different time points (n = 3, biological replicates) during incubation were evaluated using the same rheometer as for the bioink characterization equipped with a Peltier stage and 8 mm sandblasted probe (gap size h = 0.15–0.35 mm, normal force FN = 0.1 N) at 25 °C. Samples were cut from disc-shaped samples using a metal punch (Ø = 8 mm) to match the rheometer probe geometry. Dynamic oscillatory frequency sweep measurements were performed in the range of 100 to 0.1 rad s−1 (γ = 0.3%). Young’s modulus was calculated as

E = 2G′(1 + ν) (1)

using storage value G′ at ω = 10 rad s−1 and an assumption for Poisson ratio of ν = 0.5.

To obtain homogeneous samples for tensile testing, abiotic and biotic bioink were molded into a rectangular-shaped mold (12 mm × 50 mm × 1 mm, width × length × thickness) and cross-linked for 2 min (λ = 405 nm light, I = 8 mW cm−2). Samples were then washed and incubated under the same conditions as the disc-shaped samples. Uniaxial tensile testing was performed on the samples (n = 5, biological replicates) incubated for 2 and 30 days using a tensile testing machine (Stable Micro Systems) equipped with two parallel metal screw-clamps. A 2 mm notch was cut with a razor blade on one side of the sample prior to the stress-strain measurement. The sample was placed securely between the two parallel clamps with an initial grip-to-grip distance of 300 mm. Stress-strain measurement was performed with a constant stretch speed of 0.5 mm s−1 until failure. The sample toughness was calculated by measuring the area under the stress–strain curve and Young’s modulus of the sample was calculated as the slope of the linear regime of the curve (Supplementary Fig. 12).

Compressive tests were conducted on the same tensile testing machine (Stable Micro Systems) equipped with a steel probe (Ø = 0.2 mm). The compression test was done at a constant deformation rate of 0.05 mm min−1 on a cylinder-shaped photosynthetic living material sample after 400 days of incubation (Ø = 10 mm, h = 3 mm). The compressive modulus was obtained by measuring the slope of the linear regime of the stress–strain curve (Supplementary Table 2).

Statistical analysis of the samples at different time points was performed using two-sided two-sample t-tests (OriginPro 2019 software). All values are shown as average with standard deviation or root mean square of standard deviations in cumulative cases.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The authors have performed several additional experiments. My comments are mostly related to the additional data:\n1. Figure S11a: The BG11-ASNIII medium was changed every 5 days, so additional carbon source is made available\n(potentially) each time that the medium is refreshed. How is that taken into consideration?\n2. Figure S11a: Why are the values for the DIC in the abiotic medium varying like this with a peak at 20 days? Please also\ninclude the number of replicates and error bars.\n3. The amount of CO2 sequestered by precipitation is less than 0.1% of the whole mass of the materials. Based on Figure\nS16 (biotic culture) and Supplementary text on “CO2 sequestration by biomass growth (pH change) in living materials”, the\nprecipitates’ weight is fairly low, so the dual carbon sequestration does not appear really efficient.\n4. Concerning the carbon sequestration, is it possible to provide a quantitative analysis of the carbon sequestration\ncontribution from each factor? (e.g., biomass accumulation, carbonate precipitation without the source of sodium bicarbonate\nin ASNIII-BG11).\n5. Figure S30: The CO2 consumption rate data is quite noisy, and a very low R2 value (0.15) is shown on panel b. Also, in\npanel b, is there a reason why the x-axis is from 0.5 to 1.4h? Please also include the number of replicates and error bars.\nMinor points:\n- The authors use expressions like carbon-negative and carbon-neutral infrastructure in the manuscript. To use such\nwording, it would require a quantitative cradle-to-grave CO2 emission analysis (LCA).\n- Caption of Figure S11a: Shouldn’t that be BG11-ASNIII medium?\n- Typo in Figure S11: mediuim (twice)\n- Typo in Figure 2a: medum", "reviewer_2": "After reading the authors’ Response to Reviewers, I appreciate the addition of cell counts and chlorophyll measurements to\nsupport the conclusions regarding biomass growth. However, I remain unconvinced that measuring extracellular pH is a\nvalid way to quantify uptake of inorganic carbon into biomass in this system, which is open to the atmosphere. The standard\nway to quantify photosynthesis in cyanobacteria is to measure O2 evolution using an oxygen electrode or by membrane inlet\nmass spectrometry.\nI understand that there are several references (cited in the Response to Reviewers) that describe the release of OH- ions by\ncarbonic anhydrase located in the carboxysome as part of a model for the cyanobacterial CCM and MICP, but this is an\noversimplification that is not biochemically correct. A paper on quantitative modeling of the CCM in beta-cyanobacteria\n(Mangan et al., 2016. PNAS 113: E5354-E5362) notes that the combined action of CA and Rubisco in the carboxysome\nactually produces a net H+, not an OH- ion.\nThe statements on p. 4 (“Here, carbonic anhydrase (CA) within the carboxysome converts the bicarbonate species into CO2,\nreleasing a hydroxyl ion (OH-) into the surrounding media7,33–35. The OH- is excreted to the exterior of the bacterium....”)\nand p. 7 (“Using HCO3- as a substrate, carbonic anhydrase generates equimolar amounts of OH- ions, which are excreted\ninto the surrounding medium, and CO2 molecules, which are integrated into bacterial metabolism.17,27”) and the depiction\nin Fig. 1 are incorrect.", "reviewer_3": "RESPONSE TO REVIEWERS: All responses are written in blue font and all changes to the main text or supplementary information have been highlighted in yellow.\nReviewer #1 (Remarks to the Author):\nThe manuscript of Dranseike and co-authors reports on a photosynthetic living material that sequesters CO2 via two routes: biomass accumulation and MICP. The living material is based on cyanobacteria encapsulated into a Pluronic F127-based hydrogel matrix. The use of Pluronic F127 enables them to 3D-print their living material. Over the course of 30 days of growth, the dry mass of the material is shown to increase. Such increase is then associated biomass growth and MICP. My most major concerns on this study relate to the novelty of this “dual sequestration” and to some technical aspects.\nWe appreciate the Reviewer’s consideration of our work. By addressing specifically the comments below, we believe that a revised version of our manuscript will articulate more clearly the ‘dual sequestration’ nature of our approach as well as clarify the technical aspects that were not sufficiently clear in the original manuscript.\n1. From what I understand, the main novel aspect is this dual sequestration obtained in a living material, i.e. combination of CO2 sequestration via biomass accumulation and photosynthetic MICP. This manuscript claims that such dual sequestration has not yet been explored in photosynthetic living materials. I am not sure that I agree with this. For example, the work of Reinhardt et al. Front. Bioeng. Biotechnol. 2023, which the authors also cite, reports on a living material that is using CO2 sequestration for biomass accumulation (cell growth) and photosynthetic MICP.\nWe appreciate the Reviewer noting the main novel aspect of the work, which is to tailor a living material specifically for dual carbon sequestration. We apologize that the original writing gave the impression that systems with both biomass generation and precipitation have not yet been explored. However, we maintain that, to our knowledge, no system has been engineered specifically for efficient dual carbon sequestration using photosynthetic living materials. The work of Reinhardt et al., and others, explored photosynthetic cyanobacteria primarily to reinforce materials via microbial-induced carbonate precipitation (MICP). This process also inherently involves biomass accumulation, but the focus of the materials design has been on the use of precipitation to augment the mechanical properties of the structure and the design was not tailored for efficient dual carbon sequestration.\nMore specifically, the manuscript by Reinhardt et al. encapsulated photosynthetic cyanobacteria within a printable mixture of alginate, methylcellulose, and sand. Their work focused on the change in mechanical properties of these living building materials with the aim of fabricating an environmentally benign alternative to the carbon emission intensive cement sector. The authors carried out a thorough analysis of the mechanical properties and cell viability of the living building materials and the materials were not engineered for efficient carbon sequestration specifically.\nIn the present work, we tailored the material design specifically to enable efficient dual carbon sequestration (via both biomass and precipitate generation) of the encapsulated cyanobacteria. This required rational material selection and 3D design that were combined with rigorous analysis of the developed photosynthetic living materials. That said, the work of Reinhardt et al. and others motivated the feasibility of our approach, and we have reframed the Introduction and Discussion to articulate these points more clearly.\nText in the original version of the Introduction:\n“Recently, photosynthetic MICP was used to design living building materials that mineralized over time17.”\n“Drawing inspiration from natural systems, we engineered photosynthetic living materials for dual CO sequestration by immobilizing MICP-capable photosynthetic microorganisms 2 within a printable polymeric network.”\nUpdated text in the revised version of the Introduction (page 3, 4):\n“Recently, photosynthetic MICP was used to design living building materials that were reinforced by mineralization over time17.”\n“Drawing inspiration from natural systems and recent advances in engineered living materials17, we engineered photosynthetic living materials for dual CO2 sequestration by immobilizing MICP-capable photosynthetic microorganisms within a printable polymeric network.”\nText in the original version of the Discussion:\n“The design of our photosynthetic living materials builds upon many recent advances in engineered living materials9,13,50,51. The diverse functionality of microorganisms has enabled the design and application of engineered living materials in various fields. For instance, bacteria-based biosensors are used to detect target molecules, such as glucose or specific pathogens52,53. Living skin patches adopt skin microbiomes to control bacterial outgrowth on the wounded area54. Utilizing the bacterial-mineralization process, microorganism-based living building materials can self-heal and regenerate16,17,27. Living materials have also been used to replace conventionally pollution-intensive processes, such as applications in the energy industry, the production of textiles or structural materials, due to their inherent bioremediation abilities, especially through the carbon sequestration capabilities of certain microorganisms55–59.”\nUpdated text in the revised version of the Discussion (page 14):\n“The design of our photosynthetic living materials builds upon many recent advances in engineered living materials9,13,50,51. The diverse functionality of microorganisms has enabled the design and application of engineered living materials in various fields. For instance, bacteria-based biosensors are used to detect target molecules, such as glucose or specific pathogens52,53. Living skin patches adopt skin microbiomes to control bacterial outgrowth on the wounded area54. Notably, bacterial mineralization processes have been exploited to design microorganism-based living building materials that can self-heal, strengthen over time, and regenerate16,17,27, demonstrating the potential to combine biomass accumulation and mineralization within a single living material. Living materials have also been used to replace conventionally pollution-intensive processes, such as applications in the energy industry, the production of textiles or structural materials, due to their inherent bioremediation abilities, especially through the carbon sequestration capabilities of certain microorganisms55–59.”\n2. The authors write that, in their living materials MICP is using atmospheric CO2 as the sole carbon source. I do not agree with this since they use the BG11 medium, which contains sodium carbonate.\nWe appreciate the Reviewer’s comment and agree that identifying the precise carbon source is important and tracking this would require further investigation, such as isotope labeling. Therefore, we revised the manuscript text to state that atmospheric carbon was the main source and include additional experimental data on the carbon balance and equilibrium in the culture medium throughout the sample incubation to support the claim that atmospheric CO is the main 2 source of carbon (see more details in the response to the next comment R1.3). We have also updated the composition table to include the full list of ions from ASN II and BG11 media in the Supplementary Information.\nText in original version (page 13):\n“Our photosynthetic living materials function at ambient conditions with atmospheric CO 2 as the sole carbon source, which highlights the ability to use photosynthetic living materials as a complementary approach in a more distributed manner.”\nText in the revised version (page 14):\n“Our photosynthetic living materials function at ambient conditions with atmospheric CO 2 as the main carbon source, [...]”\n3. The amount of sodium carbonate in BG11 should be considered in their calculations of CO2 sequestration, also considering that the medium is changed regularly, so more sodium carbonate in accessible to the living material.\nTo address the comment of the Reviewer, we further elaborated our analysis of the dissolved inorganic carbon equilibria in the Results section of the manuscript and clarified the information about the carbon source to avoid confusion for the reader. In total, our findings demonstrate that the dissolved inorganic carbon in the medium was always higher than the amount supplied by the sodium carbonate in the medium. Therefore, the majority of the CO taken up by the system 2 arose from the atmospheric carbon that equilibrates the system. As it is not possible to track each molecule specifically, we have softened the text in the results to state that the CO sequestration 2 was achieved “with atmospheric carbon as a primary carbon source” (page 7).\nWe analyzed the culture medium according to the established protocols for inorganic carbon equilibria used in seawater analysis, known as CO2SYS [ref. 1]. We also added the corresponding data and an extended version of the CO2SYS calculations in the Supplementary Information (page 7) and a brief version below. As can be seen in Figure S8, the dissolved inorganic carbon (both from the culture medium and atmosphere) is in equilibrium. The equilibria depend on medium conditions such as salinity, alkalinity, pH, and temperature. At the end of each 5-day incubation period, we recollected and pooled the media, measured its pH, total alkalinity, and salinity. From that, we were able to calculate the total amount of dissolved inorganic carbon under equilibrium. We can show that in the case of biotic samples, the total amount of dissolved inorganic carbon was greater than the total amount of CO 2- coming from 3 the medium components despite:\n1. the simultaneous occurrence of MICP and;\n3\n2. the increase in pH as biomass grows, which decreases the solubility of carbon species overall.\nDue to the mentioned equilibria of carbon species, total dissolved inorganic carbon in abiotic control samples was substantially higher than the CO 2- coming from BG11–ASNIII medium, 3 which did not decrease as much as in the biotic case because the pH is lower (due to the absence of biomass growth and bacterial metabolic activities). This highlights that substantial amounts of atmospheric carbon dissolve in the medium in our open culture system even with the moderate addition of dissolved carbonates in the BG11–ASNIII medium.\nOverall, as the total amount of dissolved inorganic carbon in the culture medium throughout the entire incubation period was more than the amount of carbon we supplemented through the addition of sodium carbonate, we are confident that a substantial portion of the carbon in the final system came from atmospheric carbon. Based on our data, the medium components define the initial amount of dissolved inorganic carbon in the medium and either remain or are replenished by atmospheric carbon. On day 25, the total dissolved inorganic carbon was slightly below (343 µmol/kg) the amount of inorganic carbon added to the medium in the form of Na CO 2 3 (381µmol/kg), which we associated with the high pH of the media and continuous MICP.\n1. Lewis, E. R., & D. W. R. Wallace (1998). Program developed for CO2 system calculations. No. cdiac: CDIAC-105. Environmental System Science Data Infrastructure for a Virtual Ecosystem (ESS-DIVE)(United States). https://www.osti.gov/biblio/1464255.\nPrevious text in Results section (page 7):\n“During the incubation, BG11–ASNIII medium was changed every 5 days. From day 5, the concentration of Ca2+ in the medium was set to 8.65 mM via CaCl addition to 2 simulate natural seawater conditions37 and to promote MICP. To highlight the CO 2 sequestration abilities of the living material, no additional HCO - was dissolved in the 3 medium. As such, all HCO - essential for cell metabolism came from the dissolution of 3 gaseous atmospheric CO .” 2\nUpdated text in Results section (page 7):\n“During the incubation, BG11–ASNIII medium was changed every 5 days and the dissolved inorganic carbon (DIC) was quantified in the collected samples (see Supplementary Information, Calculation of dissolved inorganic carbon in BG11- ASNIII medium with CO2SYS program). From day 5, the concentration of Ca2+ in the medium was set to 8.65 mM via CaCl addition to simulate natural seawater 2 conditions37 and to promote MICP. To highlight the CO sequestration abilities of the 2 living material, no additional HCO - was dissolved in the medium. 40.4 mg L-1 (0.38 3 mM) of Na CO was dissolved in the medium to simulate natural seawater; however, 2 3 CO 2- ions are not directly taken up by cyanobacteria as source of inorganic carbon. 3 Analysis on the total amount of DIC under equilibrium in the culture medium also showed similar or even higher value than the amount of carbon initially dissolved in the form of CO 2- (Figure S11a). The higher DIC capacity of the culture medium 3 confirms the dissolution of atmospheric CO and, therefore, the CO sequestration 2 2 ability of the living materials using atmospheric CO as a primary carbon source.” 2\nFigure S11. a) Total amount of dissolved inorganic carbon (DIC) under equilibrium in the collected abiotic (black) and biotic (green) medium. Gray dashed line indicates the amount of dissolved inorganic carbon in the form of Na CO that was used in simulated 2 3 seawater medium (ASNIII). b) pH changes of pooled biotic (green) and abiotic (black) medium every 5 days.\n4. The pH change was measured in the culture medium around the living material. Have the authors checked that the culture medium is free of cyanobacteria during the whole period of incubation? Cyanobacteria are likely to also grow in this culture solution and to affect the pH. Hence what is measured is (at least in part) the CO2 sequestration capability of the cyanobacteria in the liquid and not only in the hydrogel.\nWe appreciate the Reviewer’s comment and have added data on cell concentration in hydrogels and surrounding medium to the Supplementary Information. The ratio between encapsulated and escaped cells shows that less than 20% of the total number of the cells escaped. The retention efficiency, calculated based on 1 disc sample immersed in 2 mL of culture medium, was more than 90% for the first 25 days and more than 80% on Day 30 (Figure S13). Increase in bacteria leakage/growth on day 30 (18%) might be attributed to the space limitation within the gel as we also observed a slower increase in the number of encapsulated cells potentially reaching a steady state. Additionally, we observed that while the number of escaped cells was relatively constant, the increase in pH of the surrounding culture medium every 5 days was increasing. Furthermore, we exchanged the whole medium every 5 days to limit growth and accumulation of cyanobacteria outside of the hydrogel. Any contribution to the media’s pH change from escaped cyanobacteria would be marginal and we thus conclude that the major contribution to the increase in pH came from the encapsulated cells. Therefore, we do not anticipate that this contributes substantially to the overall analysis. The contribution would potentially be even lower in the samples that are only partially immersed in the medium.\nFigure S13. Cell count in the printed disc sample quantified with chlorophyll extraction (green) and cell count in the surrounding culture medium (pink) measured via Coulter Counter. The right axis indicates the percentage of cells that leaked into the surrounding culture medium (2 mL) normalized to the total number of cells in 1 printed disc sample (gray), as well as the encapsulation efficiency calculated as 100% – leakage % (brown).\nUpdated text in Results section (page 7):\n“The pH did not change in abiotic control samples throughout the incubation (Figure S11b) and the majority of the encapsulated bacteria remained within the hydrogel throughout the incubation in biotic samples (Figure S13).”\n5. Since there is a strong focus on carbon sequestration, I recommend a more rigorous analysis on CO2 sequestration values. For example, would it be possible to directly measure the evolution of CO2 used in the atmosphere during material growth? Given the previous point, it is preferable to do this without liquid at the base of the material, or only when there is freshly added culture media (without cyanobacteria).\nWe thank the Reviewer for the recommendation, and we think this is a great idea. We performed CO sequestration measurements in a closed system. Over the measurement period, we 2 observed a decrease in atmospheric CO level. In brief, a 0.25 mL sample with approximately 10 2 mL of culture medium was placed in a glass jar in an air-tight closed container (250 mL) with an embedded CO sensor (Sensirion, SEK-SCD41 Multiple Function Sensor) for 10 days. The 2 container was opened every day and was allowed to equilibrate with atmospheric air to replenish the CO within the closed system. The rate of change of CO concentrations were measured 2 2 during the 12-hour day cycle (Figure S30a). Starting from measurement day 3, the rate of change in CO concentrations within the closed container was below zero, indicative of CO 2 2 sequestration. The rate of change in CO concentration reached a value of approximately -30 2 ppm per hour beyond day 5. We also collected a representative snapshot of the CO 2 concentration within the closed container for one of the samples over the course of 1 hour (Figure S30b). Average atmospheric CO concentration decreased from approximately 947 ppm to 940 2 ppm during the measurement time.\nThe new Figure S30 is appended below:\nFigure S30. a) Rate of change of CO concentration in a 250 mL closed container 2 over a measurement period of 10 days with a 0.25 mL sample and 10 mL culture medium. CO sequestration was observed from day 3 and plateaued beyond day 5. 2 b) Representative snapshot of CO evolution (black) in a closed container and linear 2 interpolation of the measurement (blue).\nIn our future research, we plan to build a continuous measurement system to obtain more accurate read-outs of the rate of CO sequestration of the living materials. We believe that further 2 development on the experimental design will be a valuable area of research for the future work. More accurate analysis on direct CO measurement would require a tailored measurement setup 2 considering all the environmental parameters under dynamic equilibrium. In future work for upscaling the use of photosynthetic living materials, we plan to engineer controlled bioreactors that quantitatively control and monitor CO and O input/output, temperature, humidity, and the 2 2 supply of culture media. In this manner, we would be able to more quantitatively analyze the CO 2 sequestration directly during growth. We agree that this is an important area of research for future work, especially for the scale-up and translation of these concepts. We would like to emphasize again that in the present work, we relied on extensive post mortem analyses of our photosynthetic living materials to ensure robust quantification of the CO uptake and mineralization of the 2 material, which convincingly demonstrate the activity and importance of the present system.\nTo address the continued need for more robust quantification of CO uptake during the life cycle 2 of the photosynthetic living materials, we have added the following text to the Discussion.\nText in original version of the Discussion (page 13):\n“Implementing photosynthetic living materials in broad application still requires improved usability and upscaling of the material production. This can be achieved by taken advantage of recent advances in biofabrication for the creation of larger scale porous62 or granular63 structures.”\nUpdated text in the revised version of the Discussion (page 15):\n“Implementing photosynthetic living materials in broad application still requires improved usability and upscaling of the material production. This can be achieved by taking advantage of recent advances in biofabrication for the creation of larger scale porous62 or granular63 structures. Upscaling would provide the means to do more robust CO sequestration analysis using gas composition monitoring (Figure S30) as 2 well as control over carbon sources via media supply.”\n6. The pH reaches values above 9 in the growth medium. Was there also calcium precipitation observed in the dish containing the growth medium (outside the hydrogel)?\nWe observed calcium precipitates in the culture dish (24-well plates). However, the calcium precipitation in the well plates did not affect our measurements of the quantification of the CO 2 sequestration ability of the living materials as all samples were transferred to fresh, unused wells prior to any analysis.\nWhile there were minimal cyanobacteria observed in the growth medium outside of the hydrogel during the incubation period of 30 days (see R1.4), we did observe calcium precipitation in the well plates of biotic samples as the well plate surface was negatively charged and could serve as additional nucleation sites. We also quantified the difference in the amount of precipitates on the well plates (samples transferred every 5 days to new wells) between biotic and abiotic samples throughout the incubation period (Figure S15). Again, it should be noted that these additional precipitates were not counted in the quantification of the CO sequestration ability of 2 the living materials.\nFigure S15. a) Calibration curve to calculate equivalent CaCO amount from the 3 absorbance of Alizarin Red S staining. b) Equivalent CaCO amounts calculated as 3 difference in Alizarin Red S staining of the precipitates in the biotic and abiotic wells normalized per milliliter of hydrogel. The printed disc samples were transferred into new well plates every 5 days and Alizarin Red S quantification was done on the recovered well plates.\nTo quantify the CO sequestration, we carried out extensive post mortem analyses on all hydrogel 2 samples and that the calcium carbonate precipitates within the well plate or the growth medium did not affect the analysis of carbon sequestration within the photosynthetic living materials, as our samples were analyzed only after transfer to a new well thorough washing to remove any residual salts on the surface from the growth medium prior to burning the construct itself and measuring quantitatively the mineral phase within the material. The presence of additional carbonate precipitates in the surrounding environment of biotic samples, that were not accounted for in our calculations, suggests that the CO sequestration ability of the photosynthetic living 2 materials may even go beyond the amount of CO sequestered within the living material itself 2 depending on the local growth environment.\nSentence added to the revised version of the Results section of the manuscript (page 8):\n“Biotic samples stained red by day 10, indicating Ca2+ accumulation in the whole volume of the hydrogel and the dark red color was observed around growing bacteria clusters over time. From the analysis of Alizarin Red S staining, we observed that the amount of precipitates was equivalent to > 1 mg of CaCO (mL-1 of hydrogel) already 3 after 10 days (Figure 2e). It reached 4.8 ± 2 mg of CaCO (mL-1 of hydrogel) by the 3 end of the 30-day period. Precipitates were also observed in the bottom of the incubation wells, and all analyses were carried out in fresh wells to avoid any influence on precipitation outside of the living material on our quantification (Figure S15).”\nUpdated Methods section about Alizarin Red S staining quantification (page 18):\n“Calcium-containing precipitate deposits in circular disc-shaped hydrogels were stained using 40 mM Alizarin Red S solution (pH = 4.2) in Milli-Q water. Each sample was fully immersed in 2 mL of Alizarin red S solution for 20 s. The sample was then washed in 200 mL of Milli-Q water 3 times for 5 min and imaged using an optical microscope. To quantify the extent of precipitation, we extracted the Alizarin Red S stain from the samples using 10% acetic acid (0.5 mL per sample, n = 3) with vortexing for at least 1 hour. The solutions were then neutralized using 1 M NaOH and 100 μL of the neutralized extraction solution was used to measure the absorbance at 405 nm using a UV-vis plate reader (HIDEX Sense). A calibration curve was generated using pure CaCO following the same method to quantify the amount of CaCO in the 3 3 samples based on the Alizarin Red S absorbance (Figure S15a). Alizarin Red S staining was also performed on randomly chosen recovered wells (n = 3) of 24-well plate every 5 days. 0.5 mL of 40 mM Alizarin Red S solution was added to each well for 20 s. Each well was then washed with 0.5 mL Milli-Q water 3 times to collect the precipitates, which were then recovered by centrifugation at 12,300 rcf for 10 minutes. The supernatant was then discarded and the remaining precipitate pellet was dissolved in 0.5 mL of 10% acetic acid by pipetting. The solution was neutralized and the absorbance was measured under the same conditions as those of the disc-shaped hydrogel samples.”\n7. What is the viability of bacterial cells within gels after post-processing (e.g., printing, curing at 405 nm)? The authors use a 22G conical nozzle, which could have high shear stress exerted on cells. And the cells are exposed to UV during the curing.\nWe agree that this is a valuable set of data to include in the manuscript to guide the design of photosynthetic living materials, and therefore, we added data on cell viability after the printing process to the Supplementary Information. To confirm that the bacterial cells survive the printing process (extrusion and photo-cross-linking), we include viability data for day 0 and day 1 (Figure S2). The cells were viable (> 50%) after printing and the population recovered to > 85% viability by day 1. Given this, we can tailor the initial concentration of cyanobacteria to ensure a robust population of the sample at day 1 that remains viable and grows for over 1 year.\nWe also agree that the photo-cross-linking step using 405 nm light is likely one factor that impacts cell viability during printing, and therefore added the information on the impact of different light intensities on the reaction rate and cell viability to the Supplementary Information. We observed substantially compromised cell viability when using light intensity I = 12 mW cm-2 (Figure S10a). Therefore, we chose to use a light intensity of I = 8 mW cm-2 for our experiments to minimize photo-cross-linking time while achieving sufficient cell viability (Figure S10b).\nThe Supplementary Figures are appended below:\nFigure S2. a) Cell viability on day 0 and day 1 after bioprinting. b) Representative maximum intensity Z-projection images of dead cell staining (cyan) and cell autofluorescence (green). Scale bar, 100 μm.\nFigure S10. a) Cell viability on day 5 after photo-cross-linking the bioink at different light intensities (SYTOX Blue dead cell staining, cyan; cell autofluorescence, green; maximum intensity Z-projection). Scale bar, 100 μm. b) Bioink photo-cross-linking at different light intensities of 4, 8, and 12 mW cm-2 (λ = 405 nm; light turned on at t = 60 s).\nPrevious section in the Methods (page 16):\n“Biomass characterization\n5 of the freshly printed (day 0) abiotic and biotic disc samples were washed in Milli-Q water for 10 min to remove salt deposited on the surface from the culturing medium. The samples were then dried under a constant laminar flow in a 1.5 mL microcentrifuge tube of known mass to evaporate the water entrapped within the material. The dried samples together with the microcentrifuge tube were weighed and the mass was used as the initial benchmark for characterization.\nOn day 5, 10, 20 and 30, five of the abiotic and biotic samples were randomly selected and washed with the same procedure. The samples were subsequently dried under the same condition before weighing. The mass was normalized to the average mass of the abiotic or biotic discs on day 0.”\nUpdated Methods section (pages 17–18):\n“Biomass quantification and cell viability characterization\nTo measure the sample mass change, 5 of the freshly printed (day 0) abiotic and biotic disc samples were washed in Milli-Q water for 10 min to remove salt deposited on the surface from the culturing medium. The samples were then dried under a constant laminar flow in a 1.5 mL microcentrifuge tube of known mass to evaporate the water entrapped within the material. The dried samples together with the microcentrifuge tube were weighed and the mass was used as the initial benchmark for characterization. On day 5, 10, 20 and 30, 5 of the abiotic and biotic samples were randomly selected and washed with the same procedure. The samples were subsequently dried under the same condition before weighing. The mass was normalized to the average mass of the abiotic or biotic discs on day 0.\nCell viability and growth within the hydrogels were evaluated using confocal microscopy and chlorophyll extraction (Figure S34). Dead cells within the printed disc samples were stained using SYTOX Blue dead cell staining. Disc samples were first washed 3 times in 1 mL of MilliQ water then stained using 0.2 μM staining solution diluted with 1x ASNIII medium. The samples were then incubated in the dark for 15 min, washed 3 times with 1x ASNIII medium for 5 min, and stored in the same medium until imaging. The images of SYTOX Blue dead cell staining and cell autofluorescence were acquired with confocal microscopy (TCS SP8, Leica with a 506357 ∞/0.14 - 0.18/OFN25/E 40x, water objective, 26 slices, 2 μm each). A 405 nm laser was used to excite SYTOX Blue and emission was collected at 450–510 nm (HyD detector). Chlorophyll autoflorescence was excited with a 488 nm laser and was collected at 682– 720 nm (HyD detector). Z-stacks of 50.4 µm were used for maximum intensity Z- projections using LAS X (Leica). Separate images (n = 3–5) of dead cell and autofluorescence were imported in CellProfiler 4.2.5, (https://cellprofiler.org/). Gaussian filter was applied to the images followed by primary object identification using a built-in module (Adaptive Otsu threshold algorithm was applied with 3 classes where middle intensity was considered as background), resulting in a binary image and object count.\nChlorophyll extraction was done on randomly chosen disc samples (n = 5) every 5 days starting from day 10. The samples were fully immersed in 0.5–1 mL of isopropyl alcohol and homogenized (IKA T25 Ultra Turrax with S25N-8G-ST inset). The slurry was then sonicated for 85 min followed by centrifugation at 19000 rcf for 10 min. Technical triplicates of 100 μL supernatant were transferred into a 96-well plate and the absorption at 663 nm was measured by UV-vis plate reader (HIDEX Sense). Suspension samples of different optical densities were used to calculate an equivalent cell concentration from the chlorophyll amount (Figure S25).”\nFigure S25. a) Calibration curve fitting to relate chlorophyll absorption at OD to cell 663nm suspension OD (black) and calibration curve fitting to relate cell suspension OD and 730nm 730nm cell number measured via particle counting (black). b) Cell concentration in liquid suspension (particle counting, black) compared to average cell concentration in encapsulated samples obtained via confocal microscopy max projection (light green) and chlorophyll extraction (dark green). c) Comparison of equivalent OD of encapsulated cells (green) and actual OD 730nm 730nm of liquid suspension (black) from day 10.\n8. The modulus and the toughness of the material increased after 30 days, but the values are considerably lower than wood-based materials for example (another photosynthetic living materials with CO2 sequestration). It would be useful if the authors compare the values to other materials, including those based on ureolytic MICP for comparison.\nWe thank the Reviewer for this suggestion; however, we feel that a comparison of the mechanics of these materials to wood-based alternatives or any other material would distract from the central line of the paper. Our aim was solely to design a photosynthetic living material for long-term CO 2 sequestration, and the increase in mechanical properties was a “by-product” of the calcium carbonate precipitation pathway and further proof that the material was filling with a substantial mineral phase. This photosynthetic living material was not designed for load-bearing but specifically for efficient (compared with chemical processes), permanent (compared with wood materials), and environmentally-friendly (compared with ureolytic bacteria) CO sequestration 2 method. The work of Reinhardt et al., described above, nicely demonstrated how mineralization joining sand particles can be used to enhance mechanical properties and we have referred to this nice demonstration in several places. In order to focus our work on dual carbon sequestration and distinguish it from Reinhardt et al. and other studies in the field, we prefer to de-emphasize the mechanical reinforcement aspects of our work and further highlight the carbon sequestration.\n9. A ratio of 70% CaCO3 and 30% MgCO3 is obtained from the shift of the main diffraction peak representing the calcite (104) plane. To which level of accuracy can this approach measure ratios in carbonate precipitates?\nCa to Mg atom % ratios of EDX analysis (number of analyzed samples n = 10) was 78 ± 5 % to 22 ± 5 %, which indicates lower Mg amount in the mixture. Therefore, the 70% CaCO and 30% 3 MgCO ratio is potentially an underestimation of CaCO amount in the mixture. However, if we 3 3 compare the peak position of (104) plane to carbonates with increasing Mg content, the peak position is also similar to the pattern of dolomite, which has 50% Ca and 50% Mg. We used a ratio of 70% Ca and 30% Mg as a conservative estimate of the CaCO content given the EDX 3 quantification. We compared the difference in sequestered CO calculations between pure 2 CaCO and carbonate with 70% Ca and 30% Mg in the Supplementary Information, resulting in 3 a calculated difference of 5 %, indicating that the precise ratio of Ca:Mg did not have a substantial effect on the quantification.\nIn this work, we aimed to produce crystalline precipitates that are less soluble than amorphous ones to provide more stable carbon storage. We rephrased the Results section to emphasize this aspect instead of identifying a particular crystalline structure. We also include an extended comparison to potential components of the precipitates in Figure S18 also showing aragonite as another CaCO form that could be in the precipitates. 3\nFigure S18. XRD pattern of the precipitates after sample thermal decomposition at 600 °C and reference XRD patterns of aragonite, magnesite, dolomite and calcite7.\nText in the Results section of the original manuscript (page 9):\n“Having demonstrated that photosynthetic living materials are capable of CO 2 sequestration via both biomass accumulation and carbonate formation, we further investigated the composition of the mineral phase produced during incubation. MICP results in the formation of calcium carbonates that vary in stability and solubility with calcite being the most stable polymorph and, therefore, desirable for carbon sinking purposes (Figure 3a)41. XRD analysis of precipitates in biotic samples revealed a highly crystalline structure with the main peaks corresponding to calcite structure with a noticeable shift of the main diffraction peak representing the (104) plane (Figure 3b). The observed shift likely indicated compositional variation due to the incorporation of magnesium in the carbonate structure as previously observed in marine environments and, in this case, due to the use of simulated seawater (ASNIII medium) 42. The XRD was obtained after thermal decomposition (T = 600 °C, which is below the decomposition temperature of CaCO 40) as the signal from the F127-based matrix was 3 much stronger than the one of the precipitates (Figure S10).\nPrecipitates distributed throughout the polymeric matrix (Figure 3c). Energy dispersive X-Ray analysis (EDS) indicated mainly carbon and oxygen in the abiotic matrix whereas peaks of calcium and magnesium were prominent in the areas with precipitates in biotic samples. Elemental mapping corroborated these data with calcium, magnesium, carbon, and oxygen in overlapping regions near clusters of cyanobacteria, suggesting the pericellular formation of carbonates (Figure 3d).”\nText in the revised version of the Results section (page 10):\n“Having demonstrated that photosynthetic living materials are capable of CO 2 sequestration via both biomass accumulation and carbonate formation, we further investigated the composition of the mineral phase produced during incubation. MICP results in the formation of crystalline carbonates that vary in stability and solubility with calcite being the most stable polymorph and, therefore, desirable for carbon sinking purposes (Figure 3a)41. TGA analysis confirmed carbonate presence in the precipitates (Figure 3b), which was additionally confirmed by FTIR (Figure S17). XRD analysis of precipitates in biotic samples revealed a highly crystalline structure with the main peaks corresponding to calcium–magnesium carbonates indicated by the position of the main diffraction peak representing the (104) plane (Figure S18). The observed shift likely indicated compositional variation due to the incorporation of magnesium in the carbonate structure as previously observed in marine environments and, in this case, due to the use of simulated seawater (ASNIII medium)42. The XRD was obtained after thermal decomposition (T = 600 °C, which is below the decomposition temperature of CaCO 40) as the signal from the F127-based matrix was 3 much stronger than the one of the precipitates (Figure S19).\nPrecipitates distributed throughout the polymeric matrix (Figure 3c). Energy dispersive X-Ray analysis (EDS) indicated mainly carbon and oxygen in the abiotic matrix whereas peaks of calcium and magnesium were prominent in the areas with precipitates in biotic samples. Elemental mapping corroborated these data with calcium, magnesium, and oxygen enrichment in overlapping regions near clusters of cyanobacteria, suggesting the pericellular formation of carbonates (Figure 3d and S20). EDX measurements also indicated that Ca to Mg atom % ratio in the area of the precipitates was 78 ± 5 % to 22 ± 5 % (number of analyzed samples n = 10), which confirmed that both elements were embedded in the precipitates.”\nReviewer #2 (Remarks to the Author):\nThis manuscript describes a system in which Synechococcus sp. strain PCC 7002 cells are encapsulated in a hydrogel matrix, where they evidently grow via photosynthetic uptake of CO2 and simultaneously promote formation of insoluble carbonates (e.g. calcium carbonate). Although this is an interesting system, I am not convinced by the indirect measurements of CO2 sequestration via measurement of extracellular pH.\nWe are grateful for the Reviewer's remarks about our living material system for CO uptake. We 2 now provide more detailed information about the biomass growth as one of the carbon sequestration mechanisms based on imaging and cell viability data in the updated version of the manuscript, to support the measurements based on extracellular pH.\nMajor comments:\n1. On p. 4, the authors state “carbonic anhydrase (CA) within the carboxysome converts the bicarbonate species into a hydroxyl ion (OH-) and CO2.” On p. 7, this is restated as “Using HCO3- as a substrate, carbonic anhydrase generates equimolar amounts of OH- ions, which are excreted into the surrounding medium.” These statements are incorrect. CA catalyzes the conversion of bicarbonate (HCO3-) and H+ to CO2 and H2O. OH- is not a product of the reaction. Overall, there should be an uptake of H+ through CA activity, which will increase the pH, but this is a very indirect way to measure photosynthetic CO2 assimilation, and there are many other microbial processes that could affect the pH of the extracellular medium.\nWe thank the Reviewer for raising this point and we apologize for having not explicitly described the function of carbonic anhydrase in the context of marine β-cyanobacteria PCC 7002 clearly in the original version of the manuscript. The function of carbonic anhydrase (CA) within the carboxysome of PCC 7002 is indeed to convert HCO - to CO 1,2. However, according to the body 3 2 of literature describing carbon concentration mechanisms of cyanobacteria1-7, the species that perform MICP do it in this way:\n● CO uptake as HCO - into the cell occurs via active transport where CA enzymes convert 2 3 CO into HCO - at a uniport and then HCO - diffuses into the carboxysome. The 2 3 3 equilibrium of this reaction is, as described by the Reviewer, CO + H O ⇆ H+ + HCO −. 2 2 3\n● However in the carboxysome, CA converts HCO - into CO , liberating OH- ions that are 3 2 then released from the cell.\nAs a result, the pH increases in the sheath, which shifts the dissolved carbon equilibrium towards HCO - conversion to CO 2-, favoring CaCO nucleation within the sheath. 3 3 3\nDue to this mechanism in the carboxysome of PCC 7002—corroborated by the references below—we believe that it is reasonable to do a first order estimation of the amount of carbon integrated into biomass during growth based on the amount of hydroxyl ions released. To corroborate this approach, we conducted cell count with confocal microscopy as well as chlorophyll extraction to benchmark the increase in biomass throughout the 30 days. Cell count and chlorophyll extraction further supports the increase in biomass previously quantified by the increase in the pH due to biomass growth, which was positively correlated with the amount of CO sequestered via biomass accumulation. 2\n1. Riding, R. (2006). Cyanobacterial calcification, carbon dioxide concentrating mechanisms, and Proterozoic-Cambrian changes in atmospheric composition. Geobiology 4(4), 299-316.\n2. Kamennaya, N., Ajo-Franklin, C., Northen, T. & Jansson, C. (2012). Cyanobacteria as biocatalysts for carbonate mineralization. Minerals 2, 338–364.\n3. Reinhardt, O., Ihmann, S., Ahlhelm, M., & Gelinsky, M. (2023). 3D bioprinting of mineralizing cyanobacteria as novel approach for the fabrication of living building materials. Frontiers in Bioengineering and Biotechnology, 11, 1145177.\n4. Garg, R., Garg, R., & Eddy, N. O. (2023). Microbial induced calcite precipitation for self- healing of concrete: a review. Journal of Sustainable Cement-Based Materials, 12(3), 317-330.\n5. Achal, V., Mukherjee, A., Kumari, D., & Zhang, Q. (2015). Biomineralization for sustainable construction–A review of processes and applications. Earth-science reviews, 148, 1- 17.\n6. Dittrich, M., & Obst, M. (2004). Are picoplankton responsible for calcite precipitation in lakes?. AMBIO: A Journal of the Human Environment, 33(8), 559-564.\n7. Merz, M. U. E. (1992)The biology of carbonate precipitation by cyanobacteria. Facies 26, 81–101.\nWe adjusted the manuscript text to be more specific about the function of carbonic anhydrase in cyanobacteria PCC 7002 and release of OH- ions that contributes to the MICP process. In addition, we removed the illustrations of chemical reactions from Figure 2 to avoid confusion for the reader.\nText in the original manuscript:\n“For carbon concentration to occur, CO from the surrounding environment dissolves in 2 aqueous solutions and forms bicarbonate ions (HCO -) that are then transported to the 3 bacterial carboxysome33. Here, carbonic anhydrase (CA) within the carboxysome converts the bicarbonate species into a hydroxyl ion (OH-) and CO .” 2\nUpdated text in the revised version of the manuscript (page 4):\n“For carbon concentration to occur, CO from the surrounding environment dissolves in 2 aqueous solutions and forms bicarbonate ions (HCO -) that are then transported to the 3 marine β-cyanobacteria 7002 carboxysome33. Here, carbonic anhydrase (CA) within the carboxysome converts the bicarbonate species into CO , releasing a hydroxyl ion (OH-) 2 into the surrounding media7,33–35.”\nUpdated Figure 2:\nFigure 2. Dual carbon sequestration in photosynthetic living materials.\n2. One example of a microbial process that affects pH is the calcification reaction itself, which releases 2 H+ for each CaCO3 formed.\nWe thank the reviewer for this comment. We agree that any release of H+ into the growth medium from precipitation or other biological processes would affect the measurements based on pH change in the medium. However, in this context, release of H+ ions into the growth medium should be compensated by the release of OH- ions during the bacterial metabolic activities and we observed pH increase after each media change every 5 days. Therefore, the surrogate pH measurement conducted in the original manuscript is more likely to be an underestimation of the actual amount of biomass produced. To further support our findings on biomass growth, we have corroborated the original biomass quantification via pH measurement with additional quantification of biomass accumulation based on cell count with confocal microscopy and chlorophyll extraction in the revised manuscript (Figure S29). With these, we also observed an increase in cell number and high viability, which positively correlated with the amount of biomass production and CO sequestration. We still include the data of cumulative CO sequestration as 2 2 biomass calculated from pH measurements reported in the original Figure 2d as pH allowed us to estimate the order of magnitude of this process because it is not possible to quantify it from the cell number directly.\n3. Cyanobacterial growth was also assessed qualitatively by microscopy. Increases in cell number should be quantified by cell counting.\nWe thank the reviewer for this suggestion and agree that quantifying cyanobacteria growth would further support the conclusions around biomass accumulation, as described above. We performed individual cell counting in the hydrogels during days 0 and 1 of the incubation using autofluorescence images taken via confocal microscopy. However, the cells later grew in clusters (Figure S12) and accurate evaluation of their number was no longer possible after the first days of incubation. Therefore, after day 5, we quantified the cell number using chlorophyll extraction. We observed increasing cell number throughout the 30 day incubation (Figure S29b) with an initial rapid growth rate up to day 15 followed by a slower growth rate from day 15 to day 30. We have updated the Method section accordingly and we also added cell growth data to updated Figure 2d.\nSupplementary Figures S12 and S29 appended below:\nFigure S12. Representative sample maximum intensity Z-projection images at different time points (cell autofluorescence, green). Scale bar, 100 μm.\nFigure S29. a) Calibration curve fitting to relate chlorophyll absorption at OD to 663nm cell suspension OD (black) and calibration curve fitting to relate cell suspension 730nm OD and cell number measured via particle counting (black). b) Cell concentration 730nm in liquid suspension (particle counting, black) compared to average cell concentration in encapsulated samples obtained via confocal microscopy max projection (light green) and chlorophyll extraction (dark green). c) Comparison of equivalent OD 730nm of encapsulated cells (green) and actual OD of liquid suspension (black) from day 730nm 10.\nUpdated Figure 2 (panels d,e):\nFigure 2. d) An increase in cell concentration was observed throughout the 30-day incubation in the biotic samples, with an initial rapid growth rate up to day 15 followed by a slower growth rate from day 15 to day 30 (green). Biotic samples continuously sequestered CO over the 30-day incubation in the form of biomass accumulation 2 (~0.31 mg CO per g of gel, black). e) Both biomass growth and CaCO precipitation 2 3 contributed to the increase in dry mass of the biotic samples. The cumulative CO 2 sequestration by biotic samples was reflected by the difference in normalized dry mass between biotic and abiotic samples (black). Over the incubation period of 30 days, biotic samples accumulated 36% more dry mass than the abiotic samples. Alizarin red S staining on the biotic samples showed a total of 4.8 mg CaCO precipitation per g of 3 gel (red).\nUpdated Methods section (Page 17):\nBiomass quantification and cell viability characterization\n“<...> Chlorophyll extraction was done on randomly chosen disc samples (n = 5) every 5 days starting from day 10. The samples were fully immersed in 0.5– 1 mL of isopropyl alcohol and homogenized (IKA, T25 Ultra Turrax with S25N-8G-ST inset). The slurry was then sonicated for 85 min followed by centrifugation at 19000 rcf for 10 min. Technical triplicates of 100 μL supernatant were transferred into a 96-well plate and the absorption at 663 nm was measured by UV-vis plate reader (HIDEX Sense). Suspension samples of different optical densities were used to calculate an equivalent cell concentration from the chlorophyll amount (Figure S29a).”\nMinor comment:\n4. Abstract: “scalability nature” should be “scalable nature”\nThank you. We updated the text in the Abstract accordingly.\nReviewer #3 (Remarks to the Author):\nIn this manuscript, 3D printed hydrogel-based materials were developed and evaluated as novel tools for dual carbon sequestration. Photosynthetic living materials were engineered by embedding cyanobacteria within a bioinert hydrogel matrix, to achieve effective dual carbon sequestration through biomass accumulation and irreversible microbially-induced calcium carbonate precipitation. The authors investigated the platform performance not only in the short-term (1 month) but also conducted long-term studies to demonstrate the platform ability in forming biomass and producing mineral precipitation for over a year.\nThis original study brings interesting findings and useful knowledge to the field. In terms of novelty of the engineering research approach, similar photosynthetic 3D printed living materials have been recently reported in the literature (Datta, Weiss, Wangpraseurt, et al. Nat. Commun. 14, 4742, 2023). Regardless, the system novelty mainly relies in its application, as showed by the platform’s potential for concomitant biomass accumulation and irreversible microbially-induced calcium carbonate precipitation over a year timepoint. However, at the current manuscript state, several questions related to the system performance still need to be addressed.\nThe reviewer believes that, because of lack of quantitative evaluation supporting the system performance, the manuscript is not suitable for publication in Nature Materials at its current state. The manuscript may be considered for publication in other Nature family journals after addressing the following comments.\nWe thank the reviewer for acknowledging the novelty and the application oriented aspects of our work and its potential in the field. As stated by the Reviewer, additive manufacturing has been applied in the field of engineering with living materials and was our processing technique of choice specifically as it allows the encapsulation of microorganisms together with the fabrication of complex geometries needed for efficient growth of photosynthetic microorganisms. Successful CO sequestration requires not only suitable hydrogel material but also porous structures with 2 large light-exposed surface areas to overcome limitations posed by bulk materials.\nTechnical comments\n1. Hydrogel formulation – The authors identified a hybrid Pluronic F127 formulation composed by pristine F127 (13.2 wt %) and methacrylated F127 (7.3 wt %), which resulted in optimal cell viability and printability through direct ink writing and light-based additive manufacturing. However, little information is reported about how the formulation was optimized.\nWe appreciate the Reviewer’s feedback and prepared the explanation regarding the rationale behind our formulation. We expand on the specific rationale in the sub-points below and included additional data to support these points to the Supporting Information.\na. What is the rationale behind the selection of the final F127:F127-BUM ratio?\nThe specific overall polymer content was selected so that we had the lowest polymer content that maintained suitable printability. We now include a comparison of the printability of F127-based hydrogels of different polymer contents (Figure S5) in the Supporting Information. Hydrogels with polymer content >20 wt% could be printed via direct ink writing and the printing resolution 2 increased with polymer concentration. Therefore, we used a total polymer content of 20.5 wt% as it had the lowest polymer content while remaining printable.\nFigure S5. Printability of Pluronic F127 hydrogels of different polymer contents (22G nozzle, printing speed v = 5–15 mm s-1, printing pressure P = 16–45 kPa). Scale bar, 5 mm.\nIn addition, we compared different F127:F127-BUM ratios to rationalize the specific ratio of F127:F127-BUM used in this work (Figure S6). Formulations with low (<3 wt%) F127-BUM exhibited instability during photo-cross-linking, as observed by the drop in the storage modulus after the light was turned on at t = 60 s. By increasing the concentration of F127-BUM to 7 wt% and above the final modulus and rate of cross-linking increased, and there was no observable benefit of further increases in F127-BUM. Therefore, we selected a final formulation of F127 (13.2 wt%) and F127-BUM (7.3 wt%) to maintain a total polymer content above 20 wt% and to ensure rapid cross-linking and a stable final hydrogel.\nFigure S6. Photo-cross-linking of different ratios of F127 and F127-BUM studied as an increase in storage modulus during the reaction. The 405 nm light (I = 8 mW cm-2) was turned on 60 s after the beginning of the measurement.\nb. Similarly, how was the final photo initiator (LAP) concentration and cell density determined?\nWe chose photoinitiator concentration of 0.1 wt% as it is within a range of commonly used concentrations for biofabrication (0.05–0.2 wt%) and resulted in high cell viability and rapid reaction rate. For our material, only a small increase in cross-linking rate was observed for higher 2 LAP concentrations (0.2 wt%; Figure S31). Further, increasing LAP to 0.2 wt% compromised bacteria viability (Figure S32a). We decreased the concentration of LAP to 0.05 wt% to slow down the reaction for volumetric printing as the structures were overexposed with 0.1 wt% LAP. Both of the conditions showed similar cell growth in clusters on day 5 (Figure S32b).\nIn literature, starting OD values of 0.3–0.4 have been reported for similar work1. In our work, we used OD of 0.8 as this was the value measured in our liquid culture at the start of the exponential growth phase. The only different OD we used was in the case of volumetric printing where we decreased it to 0.3 to allow sufficient light transmittance for the fabrication of porous structures. Higher OD values are also possible and the cells show similar growth patterns in all cases (Figure S33), and further increasing the starting OD would increase the initial biomass required to prepare the bioink.\n1. Heveran, C. M., Williams, S. L., Qiu, J., Artier, J., Hubler, M. H., Cook, S. M., ... & Srubar, W. V. (2020). Biomineralization and successive regeneration of engineered living building materials. Matter, 2, 481-494.\nThe additional Figures that support these points were included in the Supporting Information and are appended below:\nFigure S31. a) Photo-cross-linking of the bioink with different photoinitiator LAP concentrations (λ = 405 nm; I = 8 mW cm-2; light turned on at t = 60 s). b) relative reaction rates of the same hydrogels, calculated based on the time derivative of the storage modulus.\nFigure S32. a) Differences in cell growth represented as green color of the living samples on day 5 after photo-cross-linking the bioink at with 0.05, 0.1, and 0.2 wt% LAP. The lowest cell growth was observed with 0.2 wt% LAP in the latter case. b) Cell viability on day 5 after photo-cross-linking the bioink at with 0.05 and 0.1 wt% LAP (SYTOX Blue dead cell staining, cyan; cell autofluorescence, green; maximum intensity Z-projection). Scale bar, 100 μm.\nFigure S33. Cell viability of samples with different starting cell densities (OD in the range from 0.4 to 1.6) on day 5 (SYTOX Blue dead cell staining, cyan; cell autofluorescence, green; maximum intensity Z-projection). Scale bar, 100 μm.\nFormulation optimization and related characterizations should be added to the Supplement.\nAs described above, we have now included substantial additional supporting data on cell viability and material characterization to the Supporting Information to show how we reached our reported formulation. We have also commented on these choices in the Methods section of the manuscript.\nText in the original Methods section Bioink preparation subsection (page 15):\nStock solutions of 18 wt% F127 in culturing medium, 30 wt% of F127-BUM (synthesis described in ESI, F127-bis-urethane methacrylate synthesis) in Milli-Q water, and 5 wt% LAP (synthesis described in ESI, LAP synthesis) in Milli-Q water were mixed at 3:1:0.082 ratio to reach final concentrations of 13.2 wt% F127, 7.3 wt% F127-BUM, and 0.1 wt% LAP. For the bioink preparation, cyanobacteria suspension was centrifuged at 3500 rcf for 5 mins, the supernatant was discarded, and the cell pellet was resuspended in the prepared bioink mixture to reach an equivalent optical density OD = 0.8. For volumetric printing, an equivalent optical density OD = 0.3 was 730nm 730nm used. The equivalent OD calculation is described in Equation S1. 730nm\nText in the revised Methods section Bioink preparation subsection (page 16):\nStock solutions of 18 wt% F127 in culturing medium, 30 wt% of F127-BUM (synthesis described in ESI, F127-bis-urethane methacrylate synthesis) in Milli-Q water, and 5 wt% LAP (synthesis described in ESI, LAP synthesis) in Milli-Q water were mixed at 3:1:0.082 ratio to reach final concentrations of 13.2 wt% F127, 7.3 wt% F127-BUM, and 0.1 wt% LAP. This formulation was optimized to have the lowest polymer content and stable hydrogel after photo-cross-linking (Figure S6 and S10). LAP concentration was chosen to have a high cross-linking rate without compromising cell viability (Figure S31 and S32). For the bioink preparation, cyanobacteria suspension from the early exponential phase was centrifuged at 3,500 rcf for 5 mins, the supernatant was discarded, and the cell pellet was resuspended in the prepared bioink mixture (liquefied on ice) via pipetting to reach a homogeneous solution with an equivalent optical density OD = 0.8, as this was the value measured in our liquid culture at the start of the 730nm exponential growth phase. For volumetric printing, an equivalent optical density OD = 0.3 was 730nm used to allow sufficient light transmittance for the fabrication. Choice of initial OD did not impact the cell growth (Figure S33). The equivalent OD 730nm calculation is described in Equation S1.\n2. PCC 7002 embedding – Little details are reported regarding the experimental procedures adopted to embed cyanobacteria in the hydrogel matrix. To ensure research reproducibility, please carefully describe the embedding procedure in the corresponding materials and methods section.\nWe thank the Reviewer for the feedback regarding the clarity of our experimental procedures and we apologize that this was not sufficiently clear in the initial version of the manuscript. We included additional information about the mixing of cyanobacteria and the hydrogel precursor solution. In brief, the bioink was prepared with the cyanobacteria prior to printing by liquefying the material on ice and mixing with a concentrated solution of bacteria.\nText in the original manuscript (page 15):\nFor the bioink preparation, cyanobacteria suspension was centrifuged at 3500 rcf for 5 mins, the supernatant was discarded, and the cell pellet was resuspended in the prepared bioink mixture to reach an equivalent optical density OD = 0.8. For volumetric printing, 730nm an equivalent optical density OD = 0.3 was used. The equivalent OD calculation 730nm 730nm is described in Equation S1.\nUpdated text in the revised version of the manuscript (page 16):\nFor the bioink preparation, cyanobacteria suspension from the early exponential phase was centrifuged at 3,500 rcf for 5 min, the supernatant was discarded, and the cell pellet was resuspended in the prepared bioink mixture (liquefied on ice) via pipetting to reach a homogeneous solution with an equivalent optical density OD = 0.8, as this was the 730nm value measured in our liquid culture at the start of the exponential growth phase. For volumetric printing, an equivalent optical density OD = 0.3 was used to allow sufficient 730nm light transmittance for high resolution fabrication. The choice of initial OD does not impact the cell growth (Figure S33). The equivalent OD calculation is described in Equation 730nm S1.\n3. PCC 7002 encapsulation efficiency and retention overtime – The presented F127 formulation is characterized by a high percentage of pristine F127 (i.e., 13.2 wt%). After crosslinking, this hybrid hydrogel formulation will be composed by a highly stable chemically crosslinked network (F127- BUM) mixed with a physical network constituted by F127. Because of the absence of chemical bonds, it is expected that the physical F127 phase will progressively dissolve overtime and leak out from the crosslinked network (or removed right after as reported for volumetric printing). As such, besides an initial loss of cyanobacteria (absorbed to the hydrogel surface only and not embedded in the inner structure), it is expected that there will be a progressive leakage of PCC 7002 embedded in the F127 phase.\na. What is the system PCC 7002 encapsulation efficiency overtime?\nWe thank the reviewer for raising this interesting point and we have now evaluated the encapsulation efficiency over time. Our system is not closed and cell leakage can occur. We calculated the ratio between cell count within the hydrogel and surrounding media and added it to the Supplementary Information (Figure S13). The encapsulation efficiency was above 90% over the first 25 days and decreased to 80% by day 30. The efficiency would potentially be higher in the samples that are only partially immersed in the medium.\nUpdated text in Results section (page 7):\n“The pH did not change in abiotic control samples throughout the incubation (Figure S11b) the majority of the encapsulated bacteria remained within the hydrogel throughout the incubation in biotic samples (Figure S13).”\nText added to the updated Methods section (page 17):\nBiomass quantification and cell viability characterization\n“<...> Chlorophyll extraction was done on randomly chosen disc samples (n = 5) every 5 days starting from day 10. The samples were fully immersed in 0.5–1 mL of isopropyl alcohol and homogenized (IKA, T25 Ultra Turrax with S25N-8G-ST inset). The slurry was then sonicated for 85 min followed by centrifugation at 19000 rcf for 10 min. Technical triplicates of 100 μL supernatant were transferred into a 96-well plate and the absorption at 663 nm was measured by UV-vis plate reader (HIDEX Sense). Suspension samples of different optical densities were used to calculate an equivalent cell concentration from the chlorophyll amount using exponential fitting. (Figure S25). A non-linear fitting was chosen as we observed an increase in encapsulated cell concentration with chlorophyll extraction due to the interference of other cyanobacterial pigments, such as phycobiliprotein68-70.\nFor suspension and medium samples from the well plate, samples were collected every 5 days and the cell number was measured on a Multisizer 4e (Beckman Coulter) equipped with a 30 µm aperture tube (gain: 4"}, "author_response": "RESPONSE TO REVIEWERS: All responses are written in blue font and all changes to the main\ntext or supplementary information have been highlighted in yellow.\nReviewer #1 (Remarks to the Author):\nThe manuscript of Dranseike and co-authors reports on a photosynthetic living material that\nsequesters CO2 via two routes: biomass accumulation and MICP. The living material is based on\ncyanobacteria encapsulated into a Pluronic F127-based hydrogel matrix. The use of Pluronic F127\nenables them to 3D-print their living material. Over the course of 30 days of growth, the dry mass\nof the material is shown to increase. Such increase is then associated biomass growth and MICP.\nMy most major concerns on this study relate to the novelty of this “dual sequestration” and to some\ntechnical aspects.\nWe appreciate the Reviewer’s consideration of our work. By addressing specifically the\ncomments below, we believe that a revised version of our manuscript will articulate more clearly\nthe ‘dual sequestration’ nature of our approach as well as clarify the technical aspects that were\nnot sufficiently clear in the original manuscript.\n1. From what I understand, the main novel aspect is this dual sequestration obtained in a living\nmaterial, i.e. combination of CO2 sequestration via biomass accumulation and photosynthetic\nMICP. This manuscript claims that such dual sequestration has not yet been explored in\nphotosynthetic living materials. I am not sure that I agree with this. For example, the work of\nReinhardt et al. Front. Bioeng. Biotechnol. 2023, which the authors also cite, reports on a living\nmaterial that is using CO2 sequestration for biomass accumulation (cell growth) and photosynthetic\nMICP.\nWe appreciate the Reviewer noting the main novel aspect of the work, which is to tailor a living\nmaterial specifically for dual carbon sequestration. We apologize that the original writing gave\nthe impression that systems with both biomass generation and precipitation have not yet been\nexplored. However, we maintain that, to our knowledge, no system has been engineered\nspecifically for efficient dual carbon sequestration using photosynthetic living materials. The work\nof Reinhardt et al., and others, explored photosynthetic cyanobacteria primarily to reinforce\nmaterials via microbial-induced carbonate precipitation (MICP). This process also inherently\ninvolves biomass accumulation, but the focus of the materials design has been on the use of\nprecipitation to augment the mechanical properties of the structure and the design was not\ntailored for efficient dual carbon sequestration.\nMore specifically, the manuscript by Reinhardt et al. encapsulated photosynthetic cyanobacteria\nwithin a printable mixture of alginate, methylcellulose, and sand. Their work focused on the\nchange in mechanical properties of these living building materials with the aim of fabricating an\nenvironmentally benign alternative to the carbon emission intensive cement sector. The authors\ncarried out a thorough analysis of the mechanical properties and cell viability of the living building\nmaterials and the materials were not engineered for efficient carbon sequestration specifically.\nIn the present work, we tailored the material design specifically to enable efficient dual carbon\nsequestration (via both biomass and precipitate generation) of the encapsulated cyanobacteria.\nThis required rational material selection and 3D design that were combined with rigorous analysis\nof the developed photosynthetic living materials. That said, the work of Reinhardt et al. and others\n1\nmotivated the feasibility of our approach, and we have reframed the Introduction and Discussion\nto articulate these points more clearly.\nText in the original version of the Introduction:\n“Recently, photosynthetic MICP was used to design living building materials that\nmineralized over time17.”\n“Drawing inspiration from natural systems, we engineered photosynthetic living materials\nfor dual CO sequestration by immobilizing MICP-capable photosynthetic microorganisms\n2\nwithin a printable polymeric network.”\nUpdated text in the revised version of the Introduction (page 3, 4):\n“Recently, photosynthetic MICP was used to design living building materials that were\nreinforced by mineralization over time17.”\n“Drawing inspiration from natural systems and recent advances in engineered living\nmaterials17, we engineered photosynthetic living materials for dual CO2 sequestration by\nimmobilizing MICP-capable photosynthetic microorganisms within a printable polymeric\nnetwork.”\nText in the original version of the Discussion:\n“The design of our photosynthetic living materials builds upon many recent advances in\nengineered living materials9,13,50,51. The diverse functionality of microorganisms has\nenabled the design and application of engineered living materials in various fields. For\ninstance, bacteria-based biosensors are used to detect target molecules, such as glucose\nor specific pathogens52,53. Living skin patches adopt skin microbiomes to control bacterial\noutgrowth on the wounded area54. Utilizing the bacterial-mineralization process,\nmicroorganism-based living building materials can self-heal and regenerate16,17,27. Living\nmaterials have also been used to replace conventionally pollution-intensive processes,\nsuch as applications in the energy industry, the production of textiles or structural\nmaterials, due to their inherent bioremediation abilities, especially through the carbon\nsequestration capabilities of certain microorganisms55–59.”\nUpdated text in the revised version of the Discussion (page 14):\n“The design of our photosynthetic living materials builds upon many recent advances in\nengineered living materials9,13,50,51. The diverse functionality of microorganisms has\nenabled the design and application of engineered living materials in various fields. For\ninstance, bacteria-based biosensors are used to detect target molecules, such as glucose\nor specific pathogens52,53. Living skin patches adopt skin microbiomes to control bacterial\noutgrowth on the wounded area54. Notably, bacterial mineralization processes have been\nexploited to design microorganism-based living building materials that can self-heal,\nstrengthen over time, and regenerate16,17,27, demonstrating the potential to combine\nbiomass accumulation and mineralization within a single living material. Living materials\nhave also been used to replace conventionally pollution-intensive processes, such as\napplications in the energy industry, the production of textiles or structural materials, due\nto their inherent bioremediation abilities, especially through the carbon sequestration\ncapabilities of certain microorganisms55–59.”\n2\n2. The authors write that, in their living materials MICP is using atmospheric CO2 as the sole carbon\nsource. I do not agree with this since they use the BG11 medium, which contains sodium carbonate.\nWe appreciate the Reviewer’s comment and agree that identifying the precise carbon source is\nimportant and tracking this would require further investigation, such as isotope labeling.\nTherefore, we revised the manuscript text to state that atmospheric carbon was the main source\nand include additional experimental data on the carbon balance and equilibrium in the culture\nmedium throughout the sample incubation to support the claim that atmospheric CO is the main\n2\nsource of carbon (see more details in the response to the next comment R1.3). We have also\nupdated the composition table to include the full list of ions from ASN II and BG11 media in the\nSupplementary Information.\nText in original version (page 13):\n“Our photosynthetic living materials function at ambient conditions with atmospheric CO\n2\nas the sole carbon source, which highlights the ability to use photosynthetic living\nmaterials as a complementary approach in a more distributed manner.”\nText in the revised version (page 14):\n“Our photosynthetic living materials function at ambient conditions with atmospheric CO\n2\nas the main carbon source, [...]”\n3. The amount of sodium carbonate in BG11 should be considered in their calculations of CO2\nsequestration, also considering that the medium is changed regularly, so more sodium carbonate\nin accessible to the living material.\nTo address the comment of the Reviewer, we further elaborated our analysis of the dissolved\ninorganic carbon equilibria in the Results section of the manuscript and clarified the information\nabout the carbon source to avoid confusion for the reader. In total, our findings demonstrate that\nthe dissolved inorganic carbon in the medium was always higher than the amount supplied by\nthe sodium carbonate in the medium. Therefore, the majority of the CO taken up by the system\n2\narose from the atmospheric carbon that equilibrates the system. As it is not possible to track each\nmolecule specifically, we have softened the text in the results to state that the CO sequestration\n2\nwas achieved “with atmospheric carbon as a primary carbon source” (page 7).\nWe analyzed the culture medium according to the established protocols for inorganic carbon\nequilibria used in seawater analysis, known as CO2SYS [ref. 1]. We also added the\ncorresponding data and an extended version of the CO2SYS calculations in the Supplementary\nInformation (page 7) and a brief version below. As can be seen in Figure S8, the dissolved\ninorganic carbon (both from the culture medium and atmosphere) is in equilibrium. The equilibria\ndepend on medium conditions such as salinity, alkalinity, pH, and temperature. At the end of\neach 5-day incubation period, we recollected and pooled the media, measured its pH, total\nalkalinity, and salinity. From that, we were able to calculate the total amount of dissolved\ninorganic carbon under equilibrium. We can show that in the case of biotic samples, the total\namount of dissolved inorganic carbon was greater than the total amount of CO 2- coming from\n3\nthe medium components despite:\n1. the simultaneous occurrence of MICP and;\n3\n2. the increase in pH as biomass grows, which decreases the solubility of carbon species\noverall.\nDue to the mentioned equilibria of carbon species, total dissolved inorganic carbon in abiotic\ncontrol samples was substantially higher than the CO 2- coming from BG11–ASNIII medium,\n3\nwhich did not decrease as much as in the biotic case because the pH is lower (due to the absence\nof biomass growth and bacterial metabolic activities). This highlights that substantial amounts of\natmospheric carbon dissolve in the medium in our open culture system even with the moderate\naddition of dissolved carbonates in the BG11–ASNIII medium.\nOverall, as the total amount of dissolved inorganic carbon in the culture medium throughout the\nentire incubation period was more than the amount of carbon we supplemented through the\naddition of sodium carbonate, we are confident that a substantial portion of the carbon in the final\nsystem came from atmospheric carbon. Based on our data, the medium components define the\ninitial amount of dissolved inorganic carbon in the medium and either remain or are replenished\nby atmospheric carbon. On day 25, the total dissolved inorganic carbon was slightly below (343\nµmol/kg) the amount of inorganic carbon added to the medium in the form of Na CO\n2 3\n(381µmol/kg), which we associated with the high pH of the media and continuous MICP.\n1. Lewis, E. R., & D. W. R. Wallace (1998). Program developed for CO2 system calculations.\nNo. cdiac: CDIAC-105. Environmental System Science Data Infrastructure for a Virtual\nEcosystem (ESS-DIVE)(United States). https://www.osti.gov/biblio/1464255.\nPrevious text in Results section (page 7):\n“During the incubation, BG11–ASNIII medium was changed every 5 days. From day\n5, the concentration of Ca2+ in the medium was set to 8.65 mM via CaCl addition to\n2\nsimulate natural seawater conditions37 and to promote MICP. To highlight the CO\n2\nsequestration abilities of the living material, no additional HCO - was dissolved in the\n3\nmedium. As such, all HCO - essential for cell metabolism came from the dissolution of\n3\ngaseous atmospheric CO .”\n“During the incubation, BG11–ASNIII medium was changed every 5 days and the\ndissolved inorganic carbon (DIC) was quantified in the collected samples (see\nSupplementary Information, Calculation of dissolved inorganic carbon in BG11-\nASNIII medium with CO2SYS program). From day 5, the concentration of Ca2+ in the\nmedium was set to 8.65 mM via CaCl addition to simulate natural seawater\n2\nconditions37 and to promote MICP. To highlight the CO sequestration abilities of the\n2\nliving material, no additional HCO - was dissolved in the medium. 40.4 mg L-1 (0.38\n3\nmM) of Na CO was dissolved in the medium to simulate natural seawater; however,\n2 3\nCO 2- ions are not directly taken up by cyanobacteria as source of inorganic carbon.\nshowed similar or even higher value than the amount of carbon initially dissolved in\nthe form of CO 2- (Figure S11a). The higher DIC capacity of the culture medium\n3\nconfirms the dissolution of atmospheric CO and, therefore, the CO sequestration\n2 2\nability of the living materials using atmospheric CO as a primary carbon source.”\n2\ncollected abiotic (black) and biotic (green) medium. Gray dashed line indicates the\namount of dissolved inorganic carbon in the form of Na CO that was used in simulated\n2 3\nseawater medium (ASNIII). b) pH changes of pooled biotic (green) and abiotic (black)\nmedium every 5 days.\n4. The pH change was measured in the culture medium around the living material. Have the authors\nchecked that the culture medium is free of cyanobacteria during the whole period of incubation?\nCyanobacteria are likely to also grow in this culture solution and to affect the pH. Hence what is\nmeasured is (at least in part) the CO2 sequestration capability of the cyanobacteria in the liquid and\nnot only in the hydrogel.\nWe appreciate the Reviewer’s comment and have added data on cell concentration in hydrogels\nand surrounding medium to the Supplementary Information. The ratio between encapsulated and\nescaped cells shows that less than 20% of the total number of the cells escaped. The retention\nefficiency, calculated based on 1 disc sample immersed in 2 mL of culture medium, was more\nthan 90% for the first 25 days and more than 80% on Day 30 (Figure S13). Increase in bacteria\nleakage/growth on day 30 (18%) might be attributed to the space limitation within the gel as we\nalso observed a slower increase in the number of encapsulated cells potentially reaching a\nsteady state. Additionally, we observed that while the number of escaped cells was relatively\nconstant, the increase in pH of the surrounding culture medium every 5 days was increasing.\nFurthermore, we exchanged the whole medium every 5 days to limit growth and accumulation of\ncyanobacteria outside of the hydrogel. Any contribution to the media’s pH change from escaped\ncyanobacteria would be marginal and we thus conclude that the major contribution to the\nincrease in pH came from the encapsulated cells. Therefore, we do not anticipate that this\ncontributes substantially to the overall analysis. The contribution would potentially be even lower\nin the samples that are only partially immersed in the medium.\n(green) and cell count in the surrounding culture medium (pink) measured via Coulter\nCounter. The right axis indicates the percentage of cells that leaked into the\nsurrounding culture medium (2 mL) normalized to the total number of cells in 1 printed\ndisc sample (gray), as well as the encapsulation efficiency calculated as 100% –\nleakage % (brown).\nUpdated text in Results section (page 7):\n“The pH did not change in abiotic control samples throughout the incubation (Figure\nS11b) and the majority of the encapsulated bacteria remained within the hydrogel\nthroughout the incubation in biotic samples (Figure S13).”\n5. Since there is a strong focus on carbon sequestration, I recommend a more rigorous analysis on\nCO2 sequestration values. For example, would it be possible to directly measure the evolution of\nCO2 used in the atmosphere during material growth? Given the previous point, it is preferable to do\nthis without liquid at the base of the material, or only when there is freshly added culture media\n(without cyanobacteria).\nWe thank the Reviewer for the recommendation, and we think this is a great idea. We performed\nCO sequestration measurements in a closed system. Over the measurement period, we\n2\nobserved a decrease in atmospheric CO level. In brief, a 0.25 mL sample with approximately 10\n2\nmL of culture medium was placed in a glass jar in an air-tight closed container (250 mL) with an\nembedded CO sensor (Sensirion, SEK-SCD41 Multiple Function Sensor) for 10 days. The\n2\ncontainer was opened every day and was allowed to equilibrate with atmospheric air to replenish\nthe CO within the closed system. The rate of change of CO concentrations were measured\n2 2\nduring the 12-hour day cycle (Figure S30a). Starting from measurement day 3, the rate of change\nin CO concentrations within the closed container was below zero, indicative of CO\n2 2\n6\nsequestration. The rate of change in CO concentration reached a value of approximately -30\n2\nppm per hour beyond day 5. We also collected a representative snapshot of the CO\n2\nconcentration within the closed container for one of the samples over the course of 1 hour (Figure\nS30b). Average atmospheric CO concentration decreased from approximately 947 ppm to 940\n2\nppm during the measurement time.\nThe new Figure S30 is appended below:\nFigure S30. a) Rate of change of CO concentration in a 250 mL closed container\n2\nover a measurement period of 10 days with a 0.25 mL sample and 10 mL culture\nmedium. CO sequestration was observed from day 3 and plateaued beyond day 5.\n2\nb) Representative snapshot of CO evolution (black) in a closed container and linear\n2\ninterpolation of the measurement (blue).\nIn our future research, we plan to build a continuous measurement system to obtain more\naccurate read-outs of the rate of CO sequestration of the living materials. We believe that further\n2\ndevelopment on the experimental design will be a valuable area of research for the future work.\nMore accurate analysis on direct CO measurement would require a tailored measurement setup\n2\nconsidering all the environmental parameters under dynamic equilibrium. In future work for\nupscaling the use of photosynthetic living materials, we plan to engineer controlled bioreactors\nthat quantitatively control and monitor CO and O input/output, temperature, humidity, and the\n2 2\nsupply of culture media. In this manner, we would be able to more quantitatively analyze the CO\n2\nsequestration directly during growth. We agree that this is an important area of research for future\nwork, especially for the scale-up and translation of these concepts. We would like to emphasize\nagain that in the present work, we relied on extensive post mortem analyses of our photosynthetic\nliving materials to ensure robust quantification of the CO uptake and mineralization of the\n2\nmaterial, which convincingly demonstrate the activity and importance of the present system.\nTo address the continued need for more robust quantification of CO uptake during the life cycle\n2\nof the photosynthetic living materials, we have added the following text to the Discussion.\nText in original version of the Discussion (page 13):\n“Implementing photosynthetic living materials in broad application still requires\nimproved usability and upscaling of the material production. This can be achieved by\ntaken advantage of recent advances in biofabrication for the creation of larger scale\nporous62 or granular63 structures.”\n“Implementing photosynthetic living materials in broad application still requires\nimproved usability and upscaling of the material production. This can be achieved by\ntaking advantage of recent advances in biofabrication for the creation of larger scale\nporous62 or granular63 structures. Upscaling would provide the means to do more\nrobust CO sequestration analysis using gas composition monitoring (Figure S30) as\n2\nwell as control over carbon sources via media supply.”\n6. The pH reaches values above 9 in the growth medium. Was there also calcium precipitation\nobserved in the dish containing the growth medium (outside the hydrogel)?\nWe observed calcium precipitates in the culture dish (24-well plates). However, the calcium\nprecipitation in the well plates did not affect our measurements of the quantification of the CO\n2\nsequestration ability of the living materials as all samples were transferred to fresh, unused wells\nprior to any analysis.\nWhile there were minimal cyanobacteria observed in the growth medium outside of the hydrogel\nduring the incubation period of 30 days (see R1.4), we did observe calcium precipitation in the\nwell plates of biotic samples as the well plate surface was negatively charged and could serve\nas additional nucleation sites. We also quantified the difference in the amount of precipitates on\nthe well plates (samples transferred every 5 days to new wells) between biotic and abiotic\nsamples throughout the incubation period (Figure S15). Again, it should be noted that these\nadditional precipitates were not counted in the quantification of the CO sequestration ability of\n2\nthe living materials.\nFigure S15. a) Calibration curve to calculate equivalent CaCO amount from the\n3\nabsorbance of Alizarin Red S staining. b) Equivalent CaCO amounts calculated as\n3\ndifference in Alizarin Red S staining of the precipitates in the biotic and abiotic wells\nnormalized per milliliter of hydrogel. The printed disc samples were transferred into\nnew well plates every 5 days and Alizarin Red S quantification was done on the\nrecovered well plates.\nTo quantify the CO sequestration, we carried out extensive post mortem analyses on all hydrogel\n2\nsamples and that the calcium carbonate precipitates within the well plate or the growth medium\n8\ndid not affect the analysis of carbon sequestration within the photosynthetic living materials, as\nour samples were analyzed only after transfer to a new well thorough washing to remove any\nresidual salts on the surface from the growth medium prior to burning the construct itself and\nmeasuring quantitatively the mineral phase within the material. The presence of additional\ncarbonate precipitates in the surrounding environment of biotic samples, that were not accounted\nfor in our calculations, suggests that the CO sequestration ability of the photosynthetic living\n2\nmaterials may even go beyond the amount of CO sequestered within the living material itself\n2\ndepending on the local growth environment.\nSentence added to the revised version of the Results section of the manuscript (page 8):\n“Biotic samples stained red by day 10, indicating Ca2+ accumulation in the whole\nvolume of the hydrogel and the dark red color was observed around growing bacteria\nclusters over time. From the analysis of Alizarin Red S staining, we observed that the\namount of precipitates was equivalent to > 1 mg of CaCO (mL-1 of hydrogel) already\n3\nafter 10 days (Figure 2e). It reached 4.8 ± 2 mg of CaCO (mL-1 of hydrogel) by the\n3\nend of the 30-day period. Precipitates were also observed in the bottom of the\nincubation wells, and all analyses were carried out in fresh wells to avoid any influence\non precipitation outside of the living material on our quantification (Figure S15).”\nUpdated Methods section about Alizarin Red S staining quantification (page 18):\n“Calcium-containing precipitate deposits in circular disc-shaped hydrogels were\nstained using 40 mM Alizarin Red S solution (pH = 4.2) in Milli-Q water. Each sample\nwas fully immersed in 2 mL of Alizarin red S solution for 20 s. The sample was then\nwashed in 200 mL of Milli-Q water 3 times for 5 min and imaged using an optical\nmicroscope. To quantify the extent of precipitation, we extracted the Alizarin Red S\nstain from the samples using 10% acetic acid (0.5 mL per sample, n = 3) with vortexing\nfor at least 1 hour. The solutions were then neutralized using 1 M NaOH and 100 μL\nof the neutralized extraction solution was used to measure the absorbance at 405 nm\nusing a UV-vis plate reader (HIDEX Sense). A calibration curve was generated using\npure CaCO following the same method to quantify the amount of CaCO in the\n3 3\nsamples based on the Alizarin Red S absorbance (Figure S15a). Alizarin Red S\nstaining was also performed on randomly chosen recovered wells (n = 3) of 24-well\nplate every 5 days. 0.5 mL of 40 mM Alizarin Red S solution was added to each well\nfor 20 s. Each well was then washed with 0.5 mL Milli-Q water 3 times to collect the\nprecipitates, which were then recovered by centrifugation at 12,300 rcf for 10 minutes.\nThe supernatant was then discarded and the remaining precipitate pellet was\ndissolved in 0.5 mL of 10% acetic acid by pipetting. The solution was neutralized and\nthe absorbance was measured under the same conditions as those of the disc-shaped\nhydrogel samples.”\n7. What is the viability of bacterial cells within gels after post-processing (e.g., printing, curing at 405\nnm)? The authors use a 22G conical nozzle, which could have high shear stress exerted on cells.\nAnd the cells are exposed to UV during the curing.\nWe agree that this is a valuable set of data to include in the manuscript to guide the design of\nphotosynthetic living materials, and therefore, we added data on cell viability after the printing\n9\nprocess to the Supplementary Information. To confirm that the bacterial cells survive the printing\nprocess (extrusion and photo-cross-linking), we include viability data for day 0 and day 1 (Figure\nS2). The cells were viable (> 50%) after printing and the population recovered to > 85% viability\nby day 1. Given this, we can tailor the initial concentration of cyanobacteria to ensure a robust\npopulation of the sample at day 1 that remains viable and grows for over 1 year.\nWe also agree that the photo-cross-linking step using 405 nm light is likely one factor that impacts\ncell viability during printing, and therefore added the information on the impact of different light\nintensities on the reaction rate and cell viability to the Supplementary Information. We observed\nsubstantially compromised cell viability when using light intensity I = 12 mW cm-2 (Figure S10a).\nTherefore, we chose to use a light intensity of I = 8 mW cm-2 for our experiments to minimize\nphoto-cross-linking time while achieving sufficient cell viability (Figure S10b).\nThe Supplementary Figures are appended below:\nFigure S2. a) Cell viability on day 0 and day 1 after bioprinting. b) Representative\nmaximum intensity Z-projection images of dead cell staining (cyan) and cell\nautofluorescence (green). Scale bar, 100 μm.\nFigure S10. a) Cell viability on day 5 after photo-cross-linking the bioink at different\nlight intensities (SYTOX Blue dead cell staining, cyan; cell autofluorescence, green;\nmaximum intensity Z-projection). Scale bar, 100 μm. b) Bioink photo-cross-linking at\ndifferent light intensities of 4, 8, and 12 mW cm-2 (λ = 405 nm; light turned on at t =\n60 s).\n“Biomass characterization\n5 of the freshly printed (day 0) abiotic and biotic disc samples were washed in Milli-Q\nwater for 10 min to remove salt deposited on the surface from the culturing medium.\nThe samples were then dried under a constant laminar flow in a 1.5 mL microcentrifuge\ntube of known mass to evaporate the water entrapped within the material. The dried\nsamples together with the microcentrifuge tube were weighed and the mass was used\nas the initial benchmark for characterization.\nOn day 5, 10, 20 and 30, five of the abiotic and biotic samples were randomly selected\nand washed with the same procedure. The samples were subsequently dried under\nthe same condition before weighing. The mass was normalized to the average mass\nof the abiotic or biotic discs on day 0.”\nUpdated Methods section (pages 17–18):\n“Biomass quantification and cell viability characterization\nTo measure the sample mass change, 5 of the freshly printed (day 0) abiotic and biotic\ndisc samples were washed in Milli-Q water for 10 min to remove salt deposited on the\nsurface from the culturing medium. The samples were then dried under a constant\nlaminar flow in a 1.5 mL microcentrifuge tube of known mass to evaporate the water\nentrapped within the material. The dried samples together with the microcentrifuge\ntube were weighed and the mass was used as the initial benchmark for\ncharacterization. On day 5, 10, 20 and 30, 5 of the abiotic and biotic samples were\nrandomly selected and washed with the same procedure. The samples were\nsubsequently dried under the same condition before weighing. The mass was\nnormalized to the average mass of the abiotic or biotic discs on day 0.\nCell viability and growth within the hydrogels were evaluated using confocal\nmicroscopy and chlorophyll extraction (Figure S34). Dead cells within the printed disc\nsamples were stained using SYTOX Blue dead cell staining. Disc samples were first\nwashed 3 times in 1 mL of MilliQ water then stained using 0.2 μM staining solution\ndiluted with 1x ASNIII medium. The samples were then incubated in the dark for 15\nmin, washed 3 times with 1x ASNIII medium for 5 min, and stored in the same medium\nuntil imaging. The images of SYTOX Blue dead cell staining and cell autofluorescence\nwere acquired with confocal microscopy (TCS SP8, Leica with a 506357 ∞/0.14 -\n0.18/OFN25/E 40x, water objective, 26 slices, 2 μm each). A 405 nm laser was used\nto excite SYTOX Blue and emission was collected at 450–510 nm (HyD detector).\nChlorophyll autoflorescence was excited with a 488 nm laser and was collected at 682–\n720 nm (HyD detector). Z-stacks of 50.4 µm were used for maximum intensity Z-\nprojections using LAS X (Leica). Separate images (n = 3–5) of dead cell and\nautofluorescence were imported in CellProfiler 4.2.5, (https://cellprofiler.org/).\nGaussian filter was applied to the images followed by primary object identification using\na built-in module (Adaptive Otsu threshold algorithm was applied with 3 classes where\n11\nmiddle intensity was considered as background), resulting in a binary image and object\ncount.\nChlorophyll extraction was done on randomly chosen disc samples (n = 5) every 5\ndays starting from day 10. The samples were fully immersed in 0.5–1 mL of isopropyl\nalcohol and homogenized (IKA T25 Ultra Turrax with S25N-8G-ST inset). The slurry\nwas then sonicated for 85 min followed by centrifugation at 19000 rcf for 10 min.\nTechnical triplicates of 100 μL supernatant were transferred into a 96-well plate and\nthe absorption at 663 nm was measured by UV-vis plate reader (HIDEX Sense).\nSuspension samples of different optical densities were used to calculate an equivalent\ncell concentration from the chlorophyll amount (Figure S25).”\nFigure S25. a) Calibration curve fitting to relate chlorophyll absorption at OD to cell\n663nm\nsuspension OD (black) and calibration curve fitting to relate cell suspension OD and\n730nm 730nm\ncell number measured via particle counting (black). b) Cell concentration in liquid suspension\n(particle counting, black) compared to average cell concentration in encapsulated samples\nobtained via confocal microscopy max projection (light green) and chlorophyll extraction (dark\ngreen). c) Comparison of equivalent OD of encapsulated cells (green) and actual OD\n730nm 730nm\nof liquid suspension (black) from day 10.\n8. The modulus and the toughness of the material increased after 30 days, but the values are\nconsiderably lower than wood-based materials for example (another photosynthetic living materials\nwith CO2 sequestration). It would be useful if the authors compare the values to other materials,\nincluding those based on ureolytic MICP for comparison.\nof these materials to wood-based alternatives or any other material would distract from the central\nline of the paper. Our aim was solely to design a photosynthetic living material for long-term CO\n2\nsequestration, and the increase in mechanical properties was a “by-product” of the calcium\ncarbonate precipitation pathway and further proof that the material was filling with a substantial\nmineral phase. This photosynthetic living material was not designed for load-bearing but\nspecifically for efficient (compared with chemical processes), permanent (compared with wood\nmaterials), and environmentally-friendly (compared with ureolytic bacteria) CO sequestration\n2\nmethod. The work of Reinhardt et al., described above, nicely demonstrated how mineralization\njoining sand particles can be used to enhance mechanical properties and we have referred to\nthis nice demonstration in several places. In order to focus our work on dual carbon sequestration\nand distinguish it from Reinhardt et al. and other studies in the field, we prefer to de-emphasize\nthe mechanical reinforcement aspects of our work and further highlight the carbon sequestration.\n9. A ratio of 70% CaCO3 and 30% MgCO3 is obtained from the shift of the main diffraction peak\nrepresenting the calcite (104) plane. To which level of accuracy can this approach measure ratios\nin carbonate precipitates?\nCa to Mg atom % ratios of EDX analysis (number of analyzed samples n = 10) was 78 ± 5 % to\n22 ± 5 %, which indicates lower Mg amount in the mixture. Therefore, the 70% CaCO and 30%\n3 3\ncompare the peak position of (104) plane to carbonates with increasing Mg content, the peak\nposition is also similar to the pattern of dolomite, which has 50% Ca and 50% Mg. We used a\nratio of 70% Ca and 30% Mg as a conservative estimate of the CaCO content given the EDX\n3\nquantification. We compared the difference in sequestered CO calculations between pure\n3\na calculated difference of 5 %, indicating that the precise ratio of Ca:Mg did not have a substantial\neffect on the quantification.\nIn this work, we aimed to produce crystalline precipitates that are less soluble than amorphous\nones to provide more stable carbon storage. We rephrased the Results section to emphasize\nthis aspect instead of identifying a particular crystalline structure. We also include an extended\ncomparison to potential components of the precipitates in Figure S18 also showing aragonite\nas another CaCO form that could be in the precipitates.\n3\n600 °C and reference XRD patterns of aragonite, magnesite, dolomite and calcite7.\nText in the Results section of the original manuscript (page 9):\n“Having demonstrated that photosynthetic living materials are capable of CO\n2\nsequestration via both biomass accumulation and carbonate formation, we further\ninvestigated the composition of the mineral phase produced during incubation. MICP\nresults in the formation of calcium carbonates that vary in stability and solubility with\ncalcite being the most stable polymorph and, therefore, desirable for carbon sinking\npurposes (Figure 3a)41. XRD analysis of precipitates in biotic samples revealed a\nhighly crystalline structure with the main peaks corresponding to calcite structure with\na noticeable shift of the main diffraction peak representing the (104) plane (Figure 3b).\nThe observed shift likely indicated compositional variation due to the incorporation of\nmagnesium in the carbonate structure as previously observed in marine environments\nand, in this case, due to the use of simulated seawater (ASNIII medium) 42. The XRD\nwas obtained after thermal decomposition (T = 600 °C, which is below the\ndecomposition temperature of CaCO 40) as the signal from the F127-based matrix was\n3\nmuch stronger than the one of the precipitates (Figure S10).\nPrecipitates distributed throughout the polymeric matrix (Figure 3c). Energy dispersive\nX-Ray analysis (EDS) indicated mainly carbon and oxygen in the abiotic matrix\nwhereas peaks of calcium and magnesium were prominent in the areas with\nprecipitates in biotic samples. Elemental mapping corroborated these data with\ncalcium, magnesium, carbon, and oxygen in overlapping regions near clusters of\ncyanobacteria, suggesting the pericellular formation of carbonates (Figure 3d).”\nText in the revised version of the Results section (page 10):\n“Having demonstrated that photosynthetic living materials are capable of CO\n2\nsequestration via both biomass accumulation and carbonate formation, we further\ninvestigated the composition of the mineral phase produced during incubation. MICP\nresults in the formation of crystalline carbonates that vary in stability and solubility with\ncalcite being the most stable polymorph and, therefore, desirable for carbon sinking\npurposes (Figure 3a)41. TGA analysis confirmed carbonate presence in the\nprecipitates (Figure 3b), which was additionally confirmed by FTIR (Figure S17). XRD\nanalysis of precipitates in biotic samples revealed a highly crystalline structure with the\nmain peaks corresponding to calcium–magnesium carbonates indicated by the\nposition of the main diffraction peak representing the (104) plane (Figure S18). The\nobserved shift likely indicated compositional variation due to the incorporation of\nmagnesium in the carbonate structure as previously observed in marine environments\nand, in this case, due to the use of simulated seawater (ASNIII medium)42. The XRD\nwas obtained after thermal decomposition (T = 600 °C, which is below the\ndecomposition temperature of CaCO 40) as the signal from the F127-based matrix was\n3\nmuch stronger than the one of the precipitates (Figure S19).\nPrecipitates distributed throughout the polymeric matrix (Figure 3c). Energy\ndispersive X-Ray analysis (EDS) indicated mainly carbon and oxygen in the abiotic\n14\nmatrix whereas peaks of calcium and magnesium were prominent in the areas with\nprecipitates in biotic samples. Elemental mapping corroborated these data with\ncalcium, magnesium, and oxygen enrichment in overlapping regions near clusters of\ncyanobacteria, suggesting the pericellular formation of carbonates (Figure 3d and\nS20). EDX measurements also indicated that Ca to Mg atom % ratio in the area of the\nprecipitates was 78 ± 5 % to 22 ± 5 % (number of analyzed samples n = 10), which\nconfirmed that both elements were embedded in the precipitates.”\nThis manuscript describes a system in which Synechococcus sp. strain PCC 7002 cells are\nencapsulated in a hydrogel matrix, where they evidently grow via photosynthetic uptake of CO2 and\nsimultaneously promote formation of insoluble carbonates (e.g. calcium carbonate). Although this\nis an interesting system, I am not convinced by the indirect measurements of CO2 sequestration\nvia measurement of extracellular pH.\nWe are grateful for the Reviewer's remarks about our living material system for CO uptake. We\n2\nnow provide more detailed information about the biomass growth as one of the carbon\nsequestration mechanisms based on imaging and cell viability data in the updated version of the\nmanuscript, to support the measurements based on extracellular pH.\nMajor comments:\n1. On p. 4, the authors state “carbonic anhydrase (CA) within the carboxysome converts the\nbicarbonate species into a hydroxyl ion (OH-) and CO2.” On p. 7, this is restated as “Using HCO3-\nas a substrate, carbonic anhydrase generates equimolar amounts of OH- ions, which are excreted\ninto the surrounding medium.” These statements are incorrect. CA catalyzes the conversion of\nbicarbonate (HCO3-) and H+ to CO2 and H2O. OH- is not a product of the reaction. Overall, there\nshould be an uptake of H+ through CA activity, which will increase the pH, but this is a very indirect\nway to measure photosynthetic CO2 assimilation, and there are many other microbial processes\nthat could affect the pH of the extracellular medium.\nWe thank the Reviewer for raising this point and we apologize for having not explicitly described\nthe function of carbonic anhydrase in the context of marine β-cyanobacteria PCC 7002 clearly in\nthe original version of the manuscript. The function of carbonic anhydrase (CA) within the\ncarboxysome of PCC 7002 is indeed to convert HCO - to CO 1,2. However, according to the body\n3 2\nof literature describing carbon concentration mechanisms of cyanobacteria1-7, the species that\nperform MICP do it in this way:\n● CO uptake as HCO - into the cell occurs via active transport where CA enzymes convert\n2 3\nCO into HCO - at a uniport and then HCO - diffuses into the carboxysome. The\n2 3 3\nequilibrium of this reaction is, as described by the Reviewer, CO + H O ⇆ H+ + HCO −.\n2 2 3\n● However in the carboxysome, CA converts HCO - into CO , liberating OH- ions that are\n3 2\nthen released from the cell.\nAs a result, the pH increases in the sheath, which shifts the dissolved carbon equilibrium towards\nHCO - conversion to CO 2-, favoring CaCO nucleation within the sheath.\n3 3 3\nDue to this mechanism in the carboxysome of PCC 7002—corroborated by the references\nbelow—we believe that it is reasonable to do a first order estimation of the amount of carbon\nintegrated into biomass during growth based on the amount of hydroxyl ions released. To\ncorroborate this approach, we conducted cell count with confocal microscopy as well as\nchlorophyll extraction to benchmark the increase in biomass throughout the 30 days. Cell count\nand chlorophyll extraction further supports the increase in biomass previously quantified by the\nincrease in the pH due to biomass growth, which was positively correlated with the amount of\nCO sequestered via biomass accumulation.\n2\n16\n1. Riding, R. (2006). Cyanobacterial calcification, carbon dioxide concentrating\nmechanisms, and Proterozoic-Cambrian changes in atmospheric composition. Geobiology 4(4),\n299-316.\n2. Kamennaya, N., Ajo-Franklin, C., Northen, T. & Jansson, C. (2012). Cyanobacteria as\nbiocatalysts for carbonate mineralization. Minerals 2, 338–364.\n3. Reinhardt, O., Ihmann, S., Ahlhelm, M., & Gelinsky, M. (2023). 3D bioprinting of\nmineralizing cyanobacteria as novel approach for the fabrication of living building materials.\nFrontiers in Bioengineering and Biotechnology, 11, 1145177.\n4. Garg, R., Garg, R., & Eddy, N. O. (2023). Microbial induced calcite precipitation for self-\nhealing of concrete: a review. Journal of Sustainable Cement-Based Materials, 12(3), 317-330.\n5. Achal, V., Mukherjee, A., Kumari, D., & Zhang, Q. (2015). Biomineralization for\nsustainable construction–A review of processes and applications. Earth-science reviews, 148, 1-\n17.\n6. Dittrich, M., & Obst, M. (2004). Are picoplankton responsible for calcite precipitation in\nlakes?. AMBIO: A Journal of the Human Environment, 33(8), 559-564.\n7. Merz, M. U. E. (1992)The biology of carbonate precipitation by cyanobacteria. Facies 26,\n81–101.\nWe adjusted the manuscript text to be more specific about the function of carbonic anhydrase in\ncyanobacteria PCC 7002 and release of OH- ions that contributes to the MICP process. In\naddition, we removed the illustrations of chemical reactions from Figure 2 to avoid confusion for\nthe reader.\nText in the original manuscript:\n“For carbon concentration to occur, CO from the surrounding environment dissolves in\n2\naqueous solutions and forms bicarbonate ions (HCO -) that are then transported to the\n3\nbacterial carboxysome33. Here, carbonic anhydrase (CA) within the carboxysome\nconverts the bicarbonate species into a hydroxyl ion (OH-) and CO .”\n“For carbon concentration to occur, CO from the surrounding environment dissolves in\n2\naqueous solutions and forms bicarbonate ions (HCO -) that are then transported to the\n3\nmarine β-cyanobacteria 7002 carboxysome33. Here, carbonic anhydrase (CA) within the\ncarboxysome converts the bicarbonate species into CO , releasing a hydroxyl ion (OH-)\n2\ninto the surrounding media7,33–35.”\nFigure 2. Dual carbon sequestration in photosynthetic living materials.\n2. One example of a microbial process that affects pH is the calcification reaction itself, which\nreleases 2 H+ for each CaCO3 formed.\nWe thank the reviewer for this comment. We agree that any release of H+ into the growth medium\nfrom precipitation or other biological processes would affect the measurements based on pH\nchange in the medium. However, in this context, release of H+ ions into the growth medium should\nbe compensated by the release of OH- ions during the bacterial metabolic activities and we\nobserved pH increase after each media change every 5 days. Therefore, the surrogate pH\nmeasurement conducted in the original manuscript is more likely to be an underestimation of the\nactual amount of biomass produced. To further support our findings on biomass growth, we have\ncorroborated the original biomass quantification via pH measurement with additional\nquantification of biomass accumulation based on cell count with confocal microscopy and\n18\nchlorophyll extraction in the revised manuscript (Figure S29). With these, we also observed an\nincrease in cell number and high viability, which positively correlated with the amount of biomass\nproduction and CO sequestration. We still include the data of cumulative CO sequestration as\n2 2\nbiomass calculated from pH measurements reported in the original Figure 2d as pH allowed us\nto estimate the order of magnitude of this process because it is not possible to quantify it from\nthe cell number directly.\n3. Cyanobacterial growth was also assessed qualitatively by microscopy. Increases in cell number\nshould be quantified by cell counting.\nWe thank the reviewer for this suggestion and agree that quantifying cyanobacteria growth would\nfurther support the conclusions around biomass accumulation, as described above. We\nperformed individual cell counting in the hydrogels during days 0 and 1 of the incubation using\nautofluorescence images taken via confocal microscopy. However, the cells later grew in clusters\n(Figure S12) and accurate evaluation of their number was no longer possible after the first days\nof incubation. Therefore, after day 5, we quantified the cell number using chlorophyll extraction.\nWe observed increasing cell number throughout the 30 day incubation (Figure S29b) with an\ninitial rapid growth rate up to day 15 followed by a slower growth rate from day 15 to day 30. We\nhave updated the Method section accordingly and we also added cell growth data to updated\nFigure 2d.\nSupplementary Figures S12 and S29 appended below:\nFigure S12. Representative sample maximum intensity Z-projection images at\ndifferent time points (cell autofluorescence, green). Scale bar, 100 μm.\n663nm\ncell suspension OD (black) and calibration curve fitting to relate cell suspension\n730nm\nOD and cell number measured via particle counting (black). b) Cell concentration\n730nm\nin liquid suspension (particle counting, black) compared to average cell concentration\nin encapsulated samples obtained via confocal microscopy max projection (light\ngreen) and chlorophyll extraction (dark green). c) Comparison of equivalent OD\n730nm\nof encapsulated cells (green) and actual OD of liquid suspension (black) from day\n730nm\n10.\nUpdated Figure 2 (panels d,e):\nFigure 2. d) An increase in cell concentration was observed throughout the 30-day\nincubation in the biotic samples, with an initial rapid growth rate up to day 15 followed\n20\nby a slower growth rate from day 15 to day 30 (green). Biotic samples continuously\nsequestered CO over the 30-day incubation in the form of biomass accumulation\n2\n(~0.31 mg CO per g of gel, black). e) Both biomass growth and CaCO precipitation\n2 3\ncontributed to the increase in dry mass of the biotic samples. The cumulative CO\n2\nsequestration by biotic samples was reflected by the difference in normalized dry mass\nbetween biotic and abiotic samples (black). Over the incubation period of 30 days,\nbiotic samples accumulated 36% more dry mass than the abiotic samples. Alizarin red\nS staining on the biotic samples showed a total of 4.8 mg CaCO precipitation per g of\n3\ngel (red).\nUpdated Methods section (Page 17):\nBiomass quantification and cell viability characterization\n“<...>\nChlorophyll extraction was done on randomly chosen disc samples (n = 5) every 5\ndays starting from day 10. The samples were fully immersed in 0.5–\n1 mL of isopropyl alcohol and homogenized (IKA, T25 Ultra Turrax with S25N-8G-ST\ninset). The slurry was then sonicated for 85 min followed by centrifugation at 19000 rcf\nfor 10 min. Technical triplicates of 100 μL supernatant were transferred into a 96-well\nplate and the absorption at 663 nm was measured by UV-vis plate reader (HIDEX\nSense). Suspension samples of different optical densities were used to calculate an\nequivalent cell concentration from the chlorophyll amount (Figure S29a).”\nMinor comment:\n4. Abstract: “scalability nature” should be “scalable nature”\nThank you. We updated the text in the Abstract accordingly.\nIn this manuscript, 3D printed hydrogel-based materials were developed and evaluated as novel\ntools for dual carbon sequestration. Photosynthetic living materials were engineered by embedding\ncyanobacteria within a bioinert hydrogel matrix, to achieve effective dual carbon sequestration\nthrough biomass accumulation and irreversible microbially-induced calcium carbonate precipitation.\nThe authors investigated the platform performance not only in the short-term (1 month) but also\nconducted long-term studies to demonstrate the platform ability in forming biomass and producing\nmineral precipitation for over a year.\nThis original study brings interesting findings and useful knowledge to the field. In terms of novelty\nof the engineering research approach, similar photosynthetic 3D printed living materials have been\nrecently reported in the literature (Datta, Weiss, Wangpraseurt, et al. Nat. Commun. 14, 4742,\n2023). Regardless, the system novelty mainly relies in its application, as showed by the platform’s\npotential for concomitant biomass accumulation and irreversible microbially-induced calcium\ncarbonate precipitation over a year timepoint. However, at the current manuscript state, several\nquestions related to the system performance still need to be addressed.\nThe reviewer believes that, because of lack of quantitative evaluation supporting the system\nperformance, the manuscript is not suitable for publication in Nature Materials at its current state.\nThe manuscript may be considered for publication in other Nature family journals after addressing\nthe following comments.\nWe thank the reviewer for acknowledging the novelty and the application oriented aspects of our\nwork and its potential in the field. As stated by the Reviewer, additive manufacturing has been\napplied in the field of engineering with living materials and was our processing technique of\nchoice specifically as it allows the encapsulation of microorganisms together with the fabrication\nof complex geometries needed for efficient growth of photosynthetic microorganisms. Successful\nCO sequestration requires not only suitable hydrogel material but also porous structures with\n2\nlarge light-exposed surface areas to overcome limitations posed by bulk materials.\nTechnical comments\n1. Hydrogel formulation – The authors identified a hybrid Pluronic F127 formulation composed by\npristine F127 (13.2 wt %) and methacrylated F127 (7.3 wt %), which resulted in optimal cell viability\nand printability through direct ink writing and light-based additive manufacturing. However, little\ninformation is reported about how the formulation was optimized.\nWe appreciate the Reviewer’s feedback and prepared the explanation regarding the rationale\nbehind our formulation. We expand on the specific rationale in the sub-points below and included\nadditional data to support these points to the Supporting Information.\na. What is the rationale behind the selection of the final F127:F127-BUM ratio?\nThe specific overall polymer content was selected so that we had the lowest polymer content that\nmaintained suitable printability. We now include a comparison of the printability of F127-based\nhydrogels of different polymer contents (Figure S5) in the Supporting Information. Hydrogels with\npolymer content >20 wt% could be printed via direct ink writing and the printing resolution\n22\nincreased with polymer concentration. Therefore, we used a total polymer content of 20.5 wt%\nas it had the lowest polymer content while remaining printable.\nFigure S5. Printability of Pluronic F127 hydrogels of different polymer contents (22G nozzle,\nprinting speed v = 5–15 mm s-1, printing pressure P = 16–45 kPa). Scale bar, 5 mm.\nIn addition, we compared different F127:F127-BUM ratios to rationalize the specific ratio of\nF127:F127-BUM used in this work (Figure S6). Formulations with low (<3 wt%) F127-BUM\nexhibited instability during photo-cross-linking, as observed by the drop in the storage modulus\nafter the light was turned on at t = 60 s. By increasing the concentration of F127-BUM to 7 wt%\nand above the final modulus and rate of cross-linking increased, and there was no observable\nbenefit of further increases in F127-BUM. Therefore, we selected a final formulation of F127 (13.2\nwt%) and F127-BUM (7.3 wt%) to maintain a total polymer content above 20 wt% and to ensure\nrapid cross-linking and a stable final hydrogel.\nFigure S6. Photo-cross-linking of different ratios of F127 and F127-BUM studied as an\nincrease in storage modulus during the reaction. The 405 nm light (I = 8 mW cm-2) was turned\non 60 s after the beginning of the measurement.\nb. Similarly, how was the final photo initiator (LAP) concentration and cell density determined?\nWe chose photoinitiator concentration of 0.1 wt% as it is within a range of commonly used\nconcentrations for biofabrication (0.05–0.2 wt%) and resulted in high cell viability and rapid\nreaction rate. For our material, only a small increase in cross-linking rate was observed for higher\nbacteria viability (Figure S32a). We decreased the concentration of LAP to 0.05 wt% to slow\ndown the reaction for volumetric printing as the structures were overexposed with 0.1 wt% LAP.\nBoth of the conditions showed similar cell growth in clusters on day 5 (Figure S32b).\nIn literature, starting OD values of 0.3–0.4 have been reported for similar work1. In our work, we\nused OD of 0.8 as this was the value measured in our liquid culture at the start of the exponential\ngrowth phase. The only different OD we used was in the case of volumetric printing where we\ndecreased it to 0.3 to allow sufficient light transmittance for the fabrication of porous structures.\nHigher OD values are also possible and the cells show similar growth patterns in all cases\n(Figure S33), and further increasing the starting OD would increase the initial biomass required\nto prepare the bioink.\n1. Heveran, C. M., Williams, S. L., Qiu, J., Artier, J., Hubler, M. H., Cook, S. M., ... & Srubar,\nW. V. (2020). Biomineralization and successive regeneration of engineered living building\nmaterials. Matter, 2, 481-494.\nThe additional Figures that support these points were included in the Supporting Information and\nare appended below:\nFigure S31. a) Photo-cross-linking of the bioink with different photoinitiator LAP\nconcentrations (λ = 405 nm; I = 8 mW cm-2; light turned on at t = 60 s). b) relative\nreaction rates of the same hydrogels, calculated based on the time derivative of the\nstorage modulus.\non day 5 after photo-cross-linking the bioink at with 0.05, 0.1, and 0.2 wt% LAP. The\nlowest cell growth was observed with 0.2 wt% LAP in the latter case. b) Cell viability on\nday 5 after photo-cross-linking the bioink at with 0.05 and 0.1 wt% LAP (SYTOX Blue\ndead cell staining, cyan; cell autofluorescence, green; maximum intensity Z-projection).\nScale bar, 100 μm.\nFigure S33. Cell viability of samples with different starting cell densities (OD in the range\nfrom 0.4 to 1.6) on day 5 (SYTOX Blue dead cell staining, cyan; cell autofluorescence,\ngreen; maximum intensity Z-projection). Scale bar, 100 μm.\nAs described above, we have now included substantial additional supporting data on cell viability\nand material characterization to the Supporting Information to show how we reached our reported\nformulation. We have also commented on these choices in the Methods section of the\nmanuscript.\nText in the original Methods section Bioink preparation subsection (page 15):\nStock solutions of 18 wt% F127 in culturing medium, 30 wt% of F127-BUM (synthesis\ndescribed in ESI, F127-bis-urethane methacrylate synthesis) in Milli-Q water, and\n5 wt% LAP (synthesis described in ESI, LAP synthesis) in Milli-Q water were mixed\nat 3:1:0.082 ratio to reach final concentrations of 13.2 wt% F127, 7.3 wt% F127-BUM,\nand 0.1 wt% LAP. For the bioink preparation, cyanobacteria suspension was\ncentrifuged at 3500 rcf for 5 mins, the supernatant was discarded, and the cell pellet\nwas resuspended in the prepared bioink mixture to reach an equivalent optical density\nOD = 0.8. For volumetric printing, an equivalent optical density OD = 0.3 was\n730nm 730nm\nused. The equivalent OD calculation is described in Equation S1.\n730nm\nText in the revised Methods section Bioink preparation subsection (page 16):\nStock solutions of 18 wt% F127 in culturing medium, 30 wt% of F127-BUM (synthesis\ndescribed in ESI, F127-bis-urethane methacrylate synthesis) in Milli-Q water, and\n5 wt% LAP (synthesis described in ESI, LAP synthesis) in Milli-Q water were mixed\nat 3:1:0.082 ratio to reach final concentrations of 13.2 wt% F127, 7.3 wt% F127-BUM,\nand 0.1 wt% LAP. This formulation was optimized to have the lowest polymer content\nand stable hydrogel after photo-cross-linking (Figure S6 and S10). LAP concentration\nwas chosen to have a high cross-linking rate without compromising cell viability\n(Figure S31 and S32). For the bioink preparation, cyanobacteria suspension from the\nearly exponential phase was centrifuged at 3,500 rcf for 5 mins, the supernatant was\ndiscarded, and the cell pellet was resuspended in the prepared bioink mixture (liquefied\non ice) via pipetting to reach a homogeneous solution with an equivalent optical density\nOD = 0.8, as this was the value measured in our liquid culture at the start of the\n730nm\nexponential growth phase. For volumetric printing, an equivalent optical density\nOD = 0.3 was used to allow sufficient light transmittance for the fabrication. Choice\n730nm\nof initial OD did not impact the cell growth (Figure S33). The equivalent OD\n730nm\ncalculation is described in Equation S1.\n.\n2. PCC 7002 embedding – Little details are reported regarding the experimental procedures\nadopted to embed cyanobacteria in the hydrogel matrix. To ensure research reproducibility, please\ncarefully describe the embedding procedure in the corresponding materials and methods section.\nWe thank the Reviewer for the feedback regarding the clarity of our experimental procedures and\nwe apologize that this was not sufficiently clear in the initial version of the manuscript. We\nincluded additional information about the mixing of cyanobacteria and the hydrogel precursor\nsolution. In brief, the bioink was prepared with the cyanobacteria prior to printing by liquefying\nthe material on ice and mixing with a concentrated solution of bacteria.\nFor the bioink preparation, cyanobacteria suspension was centrifuged at 3500 rcf for 5\nmins, the supernatant was discarded, and the cell pellet was resuspended in the prepared\nbioink mixture to reach an equivalent optical density OD = 0.8. For volumetric printing,\n730nm\nan equivalent optical density OD = 0.3 was used. The equivalent OD calculation\n730nm 730nm\nis described in Equation S1.\nUpdated text in the revised version of the manuscript (page 16):\nFor the bioink preparation, cyanobacteria suspension from the early exponential phase\nwas centrifuged at 3,500 rcf for 5 min, the supernatant was discarded, and the cell pellet\nwas resuspended in the prepared bioink mixture (liquefied on ice) via pipetting to reach a\nhomogeneous solution with an equivalent optical density OD = 0.8, as this was the\n730nm\nvalue measured in our liquid culture at the start of the exponential growth phase. For\nvolumetric printing, an equivalent optical density OD = 0.3 was used to allow sufficient\n730nm\nlight transmittance for high resolution fabrication. The choice of initial OD does not impact\nthe cell growth (Figure S33). The equivalent OD calculation is described in Equation\n730nm\nS1.\n3. PCC 7002 encapsulation efficiency and retention overtime – The presented F127 formulation is\ncharacterized by a high percentage of pristine F127 (i.e., 13.2 wt%). After crosslinking, this hybrid\nhydrogel formulation will be composed by a highly stable chemically crosslinked network (F127-\nBUM) mixed with a physical network constituted by F127. Because of the absence of chemical\nbonds, it is expected that the physical F127 phase will progressively dissolve overtime and leak out\nfrom the crosslinked network (or removed right after as reported for volumetric printing). As such,\nbesides an initial loss of cyanobacteria (absorbed to the hydrogel surface only and not embedded\nin the inner structure), it is expected that there will be a progressive leakage of PCC 7002 embedded\nin the F127 phase.\na. What is the system PCC 7002 encapsulation efficiency overtime?\nWe thank the reviewer for raising this interesting point and we have now evaluated the\nencapsulation efficiency over time. Our system is not closed and cell leakage can occur. We\ncalculated the ratio between cell count within the hydrogel and surrounding media and added it\nto the Supplementary Information (Figure S13). The encapsulation efficiency was above 90%\nover the first 25 days and decreased to 80% by day 30. The efficiency would potentially be higher\nin the samples that are only partially immersed in the medium.\nUpdated text in Results section (page 7):\n“The pH did not change in abiotic control samples throughout the incubation (Figure\nS11b) the majority of the encapsulated bacteria remained within the hydrogel\nthroughout the incubation in biotic samples (Figure S13).”\nText added to the updated Methods section (page 17):\n“<...>\nChlorophyll extraction was done on randomly chosen disc samples (n = 5) every 5\ndays starting from day 10. The samples were fully immersed in 0.5–1 mL of isopropyl\nalcohol and homogenized (IKA, T25 Ultra Turrax with S25N-8G-ST inset). The slurry\nwas then sonicated for 85 min followed by centrifugation at 19000 rcf for 10 min.\nTechnical triplicates of 100 μL supernatant were transferred into a 96-well plate and\nthe absorption at 663 nm was measured by UV-vis plate reader (HIDEX Sense).\nSuspension samples of different optical densities were used to calculate an equivalent\ncell concentration from the chlorophyll amount using exponential fitting. (Figure S25).\nA non-linear fitting was chosen as we observed an increase in encapsulated cell\nconcentration with chlorophyll extraction due to the interference of other cyanobacterial\npigments, such as phycobiliprotein68-70.\nFor suspension and medium samples from the well plate, samples were collected\nevery 5 days and the cell number was measured on a Multisizer 4e (Beckman Coulter)\nequipped with a 30 µm aperture tube (gain: 4, current: 600 µA). For each solution, 25\nµL of media were diluted in 10 mL of 0.2-µm-filtered Isoton II solution within a 25 mL\nAccuvette (Beckman Coulter).\nPCC 7002 encapsulation efficiency was quantified using the cell count and chlorophyll\nextraction results. The encapsulation efficiency was calculated as the percentage of\nencapsulated cell number to the total number of encapsulated cells and cells in the\nculture medium. Leakage was defined as the percentage of cells found in the culture\nmedium to the total cell number of encapsulated cells and cells in the culture medium.”\nNew references added:\n68. Lauceri, R., Bresciani, M., Lami, A., & Morabito, G. (2018). Chlorophyll a interference in\nphycocyanin and allophycocyanin spectrophotometric quantification. Journal of limnology, 77(1).\n69. Sobiechowska-Sasim, M., Stoń-Egiert, J., & Kosakowska, A. (2014). Quantitative analysis of\nextracted phycobilin pigments in cyanobacteria—an assessment of spectrophotometric and\nspectrofluorometric methods. Journal of Applied Phycology, 26, 2065-2074.\n70. Strieth, D., Stiefelmaier, J., Wrabl, B., Schwing, J., Schmeckebier, A., Di Nonno, S., ... &\nUlber, R. (2020). A new strategy for a combined isolation of EPS and pigments from\ncyanobacteria. Journal of Applied Phycology, 32, 1729-1740.\nb. Did the author evaluate cyanobacteria leakage into the surrounding media?\nThis should be evaluated as it can affect the platform reproducibility and overall performance.\nWe evaluated the bacteria leakage from the samples and found that less than 20% of the total\ncells in the system are present in the surrounding media, indicating that the majority of the\ncyanobacteria remain encapsulated in the gel volume (Figure S13). For the first 25 days,\ncyanobacteria leakage was less than 10%. Increase in bacteria leakage/growth on day 30 (18%)\nmight be attributed to the space limitation within the gel as we also observed a slower increase\n28\nin the growth rate of encapsulated cells, indicating that the encapsulated cell number may reach\na steady state. We addressed this point of the reviewer in the Results section.\nUpdated text in Results section (page 7):\n“The pH did not change in abiotic control samples throughout the incubation (Figure\nS11b) and the majority of the encapsulated bacteria remained within the hydrogel\nthroughout the incubation in biotic samples (Figure S13).”\nFigure S13. Cell count in the printed disc sample quantified with chlorophyll extraction\n(green) and cell count in the surrounding culture medium (pink) measured via Coulter\nCounter. The right axis indicates the percentage of cells that leaked into the\nsurrounding culture medium (2 mL) normalized to the total number of cells in 1 printed\ndisc sample (gray), as well as the encapsulation efficiency calculated as 100% –\nleakage % (brown).\n4. PCC 7002 proliferation and C02 sequestration (Fig 2) – The authors report increasing cellular\ngrowth overtime, as shown by the presence of individual cells and cluster within the hydrogel matrix.\nMoreover, an increasing CO2 sequestration was observed over 30 days. Do the authors expect\nphotosynthesis to peak?\nThis is an interesting point. Yes, we do assume that photosynthesis will likely peak at some point,\nthough we anticipate the system reaches a pseudo-steady state (with cells dying and growing at\nsimilar rates) while the amount of insoluble precipitates increases. In long-term studies, beyond\none year, we also observed that the system undergoes cycles of more or less rapid growth,\n29\nincluding periodic yellowing and growth to a robust green sample. We elaborate on these aspects\nin the Results that describe Figure 2 and the related Discussion.\nText in the original Results section (page 7):\n“Over the first 15 days the cumulative amount of CO sequestered per gram of living\n2\nmaterial was 3.3 μmol (0.15 mg) and reached a value of 7 μmol (0.31 mg) by the end\nof the 30-day experiment, an increase that reflects CO sequestration corresponding\n2\nto biomass accumulation (Figure 2d).”\nRevised text (Page 7):\n“Over the first 15 days the cumulative amount of CO sequestered per gram of living\n2\nmaterial was 3.3 μmol (0.15 mg) and reached a value of 7 μmol (0.31 mg) by the end\nof the 30-day experiment, an increase that reflects CO sequestration corresponding\n2\nto biomass accumulation (Figure 2d). Beyond 30 days, we expect the biomass\naccumulation to reach a pseudo-steady state (with similar cell death and growth rates)\nand the component of carbon sequestration related to biomass accumulation to\nequilibrate within the hydrogel.”\nAdded sentence in the Discussion section (Page 14):\n“With minimum requirements of sunlight and atmospheric CO2, our photosynthetic\nliving materials showed a consistent carbon sequestration efficiency throughout the\nentire incubation period, and demonstrated the potential of using natural carbon\nsequestration processes at scale60. We expect the system to reach its peak\nphotosynthesis rate early as the cells go through an initial exponential growth phase\nand later operate at a pseudo-steady state while maintaining high viability (Figure\nS29).”\n5. Dual carbon sequestration in photosynthetic living materials (Figure 2, d; Figure 2, e) –\na. It is not clear the number of biological samples and technical replicates that were used to perform\nthe assay, as standard deviations are missing in both graphs. This info should be included and\nclarified in the corresponding materials & methods section, as well as in each caption (the authors\nmention 5 samples in the biomass and inorganic precipitation experimental section, but it is not\nclear whether technical or biological).\nIn Figure 2d, the pH was measured by pooling media from 20 randomly selected samples and\nthe graph shows results of CO calculated from the pH values obtained. The experiment was\n2\nperformed twice. We have updated the Text in the Methods accordingly (page 17):\n“The collected medium pooled from individual samples (n = 20) was used for pH\nmeasurements to evaluate total sequestered CO as fully described in the\n2\nliving materials).”\nIn Figure 2e, all raw data of biological replicates (n = 5) and calculations are provided in the\nSupplementary Information. The results are presented as a difference of means with the standard\nerror of the difference of means. We have updated the Text in the Methods accordingly (page\n17):\n30\n“On day 5, 10, 20 and 30, abiotic and biotic samples (n=5; biological replicates) were\nrandomly selected and washed with the same procedure. The samples were\nsubsequently dried under the same condition before weighing. The mass was\nnormalized to the average mass of the abiotic or biotic discs on day 0.”\nThe updated Figure 2d,e and the updated figure caption are appended below:\nFigure 2. d) An increase in cell concentration was observed throughout the 30-day\nincubation in the biotic samples, with an initial rapid growth rate up to day 15 followed\nby a slower growth rate from day 15 to day 30 (green). Biotic samples continuously\nsequestered CO over the 30-day incubation in the form of biomass accumulation\n2\n(~0.31 mg CO per g of gel, black). e) Both biomass growth and CaCO precipitation\n2 3\ncontributed to the increase in dry mass of the biotic samples. The cumulative CO\n2\nsequestration by biotic samples was reflected by the difference in normalized dry mass\nbetween biotic and abiotic samples (black). Over the incubation period of 30 days,\nbiotic samples accumulated 36% more dry mass than the abiotic samples. Alizarin red\nS staining on the biotic samples showed a total of 4.8 mg CaCO precipitation per g of\n3\ngel (red).\nb. Biotic positive control groups (e.g., PCC 7002 cells in suspension) should also be added in\naddition to the negative abiotic ones, to evaluate the effect of the hydrogel matrix on cell viability\nand photosynthetic ability.\nTo address this comment we added the results comparing cell growth in hydrogel matrix and\nsuspension positive control (cell count) to the Supplementary Information (Figure S29). It shows\nan increase in cell number for both encapsulated samples and liquid positive control during the\nfirst 15 days of incubation. Due to the cell growth in clusters in the living material, cell\nconcentration was much higher compared to the liquid suspension positive control in terms of\nboth equivalent cell number and equivalent OD (Figure S29b and c, respectively).\nAs the cells grow in clusters starting from day 5, chlorophyll extraction was conducted to quantify\nthe encapsulated cell number. It is important to note that the available medium volume was\ndifferent for our encapsulated samples and suspension. Since each disc sample was 40 µL with\nconcentrated cell clusters grown in 2 mL of medium, we observed the cell number within an\n31\nencapsulated state is higher than that in the liquid positive control. These data demonstrate that\nthere was no inherent limitation to cell viability and photosynthetic activity upon encapsulation.\nFigure S29. a) Calibration curve fitting to relate chlorophyll absorption at OD to\n663nm\ncell suspension OD (black) and calibration curve fitting to relate cell suspension\n730nm\nOD and cell number measured via particle counting (black). b) Cell concentration\n730nm\nin liquid suspension (particle counting, black) compared to average cell concentration\nin encapsulated samples obtained via confocal microscopy max projection (light\ngreen) and chlorophyll extraction (dark green). c) Comparison of equivalent OD\n730nm\nof encapsulated cells (green) and actual OD of liquid suspension (black) from day\n730nm\n10.\nc. Additional quantitative cell viability evaluation overtime as well as quantifying Alizarin red S\nstaining would complement the qualitative information reported in Figure 2 b and c.\nWe thank the Reviewer for the comment regarding quantification of cell viability and Alizarin Red\nS Staining. We calculated cell viability using chlorophyll extraction as well as confocal imaging of\ndead cells in the material at different time points in the Supplementary Information (Figure S34).\nAfter reaching a steady state, the cells were >98% viable.\nThe results of quantitative analysis of Alizarin Red S staining were added to the updated Figure\n2e and mentioned in the revised Results section. By the end of the 30 day experiment, the extent\nof precipitation measured by this method was 4.8 ± 2 mg of equivalent CaCO per mL of gel,\n3\nwhich was higher than the total amount of precipitates (2.2 ± 0.9 mg per mL of gel) previously\nfound by thermal decomposition.\nFigure S34. Cell viability (blue) and comparison of live (green) and dead (black) cell\nconcentration in encapsulated disc samples obtained via confocal microscopy max\nprojection (day 0, day 1, and all dead cell data) or chlorophyll extraction (live cells day\n15–30).\nThe updated Figure 2d, e with a second axis displaying the CaCO equivalents as quantified by\nFigure 2. d) An increase in cell concentration was observed throughout the 30-day\nincubation in the biotic samples, with an initial rapid growth rate up to day 15 followed\nby a slower growth rate from day 15 to day 30 (green). Biotic samples continuously\nsequestered CO over the 30-day incubation in the form of biomass accumulation\n2\n(~0.31 mg CO per g of gel, black). e) Both biomass growth and CaCO precipitation\n2 3\ncontributed to the increase in dry mass of the biotic samples. The cumulative CO\n2\nsequestration by biotic samples was reflected by the difference in normalized dry mass\nbetween biotic and abiotic samples (black). Over the incubation period of 30 days,\nbiotic samples accumulated 36% more dry mass than the abiotic samples. Alizarin red\n3\ngel (red).\nSentence added to the revised version of the Results section of the manuscript (page 8):\n“Biotic samples stained red by day 10, indicating Ca2+ accumulation in the whole\nvolume of the hydrogel and the dark red color was observed around growing bacteria\nclusters over time. From the analysis of Alizarin Red staining, we observed that the\namount of precipitates was equivalent to > 1 mg of CaCO per mL of hydrogel already\n3\nafter 10 days (Figure 2e). It reached 4.8 ± 2 mg of CaCO per mL of hydrogel by the\n3\nend of the 30 day period. Precipitates were also observed in the bottom of the\nincubation wells, and all analyses were carried out in fresh wells to avoid any influence\non precipitation outside of the living material on our quantification (Figure S15).”\nUpdated Methods section about Alizarin Red S staining quantification (page 18):\n“Calcium carbonate precipitate deposits in circular disc-shaped hydrogels were\nstained using 40 mM Alizarin Red S solution (pH = 4.2) in Milli-Q water. Each sample\nwas fully immersed in 2 mL of Alizarin red S solution for 20 s. The sample was then\nwashed in 200 mL of Milli-Q water 3 times for 5 min and imaged using an optical\nmicroscope. To quantify the extent of precipitation we extracted the Alizarin Red S\nstain from the samples using 10% acetic acid (0.5 mL per sample, n = 3 ) using\nvortexing for at least 1 hour. The solutions were then neutralized using 1 M NaOH and\n100 μL of the neutralized extraction solution was used to measure the absorbance at\n405 nm using a UV-vis plate reader (HIDEX Sense). Pure CaCO calibration curve\n3\nwas prepared using the same method to calculate an equivalent amount (Figure\nS15a). Alizarin Red S staining was also performed on randomly chosen recovered\nwells (n = 3) of the 24-well plate every 5 days. 0.5 mL of 40 mM Alizarin Red S solution\nwas added to each well for 20 s. Each well was then washed with 0.5 mL Milli-Q water\n3 times to collect the precipitates which were then recovered by centrifugation at\n12,300 rcf for 10 min. The supernatant was then discarded and the remaining\nprecipitate pellet was dissolved in 0.5 mL of 10% acetic acid by pipetting. The solution\nwas neutralized and the absorbance was measured under the same conditions as\nthose of the disc-shaped hydrogel samples.”\nd. Statistical analysis should also be included as it would be beneficial to validate the system\nadvantages over control groups, and strength the authors’ findings.\nFor most of our measurements we used abiotic samples as a control and we now included the\nresults of statistical analysis where applicable for the comparison between biotic samples and\nabiotic control.\nWe performed two sample t-tests to find time points at which sample properties change\nsignificantly compared to day 0, 1, or 2 depending on the experiment (Figure 3e,f). We\nobserved a significant increase (p = 0.001) in biomass from day 10 of incubation. A significant\nchange in mechanical properties compared with day 1 was observed after 10 days (p =\n0.05, Figure S21). No significant change was observed in the mechanical properties of abiotic\nsamples throughout the time course.\nquantification of calcium precipitates (Figure S15), calibration curve fitting to relate chlorophyll\nabsorption at OD to cell suspension OD and calibration curve fitting to relate cell\n663nm 730nm\nsuspension OD and cell number measured via particle counting (Figure S29) as well as\n730nm\nthe linear regression of direct CO measurement within a closed container (Figure S30).\nOver the incubation period of 30 days, the biotic samples had approximately 36% more\ndry mass than control abiotic samples (Figure 2e) with significant increase (p = 0.001) in\nbiomass observed from day 10 of incubation.\nFigure 2. e) Both biomass growth and CaCO precipitation contributed to the increase\n3\nin dry mass of the biotic samples. The cumulative CO sequestration by biotic samples\n2\nwas reflected by the difference in normalized dry mass between biotic and abiotic\nsamples (black). Over the incubation period of 30 days, biotic samples accumulated\n36% more dry mass than the abiotic samples. Alizarin red S staining on the biotic\nsamples showed a total of 4.8 mg CaCO precipitation per g of gel (red).\nFigure 3e,f. Statistical analysis added to the graphs.\nand abiotic (gray) samples during the 30-day incubation period. A significant change\nin mechanical properties compared with day 1 was also observed after 10 days (p =\n0.05).\nFigure S15. a) Calibration curve to calculate equivalent CaCO amount from the\n3\nabsorbance of Alizarin Red S staining. b) Equivalent CaCO amounts calculated as\n3\ndifference in Alizarin Red S staining of the precipitates in the biotic and abiotic wells\nnormalized per milliliter of hydrogel. The printed disc samples were transferred into\nnew well plates every 5 days and Alizarin Red S quantification was done on the\nrecovered well plates.\n663nm\ncell suspension OD (black) and calibration curve fitting to relate cell suspension\n730nm\nOD and cell number measured via particle counting (black). b) Cell concentration\n730nm\nin liquid suspension (particle counting, black) compared to average cell concentration\nin encapsulated samples obtained via confocal microscopy max projection (light\ngreen) and chlorophyll extraction (dark green). c) Comparison of equivalent OD\n730nm\nof encapsulated cells (green) and actual OD of liquid suspension (black) from day\n730nm\n10.\nFigure S30. a) Rate of change of CO concentration in a 250 mL closed container\n2\nover a measurement period of 10 days with a 0.25 mL sample and 10 mL culture\n37\nmedium. CO sequestration was observed from day 3 and plateaued beyond day 5.\n2\nb) Representative snapshot of CO evolution (black) in a closed container and linear\n2\ninterpolation of the measurement (blue).\n6. Confirmation of mineral precipitates (Figure 3a; 3b). The XRD results are not convincing; if calcite\nwere present, sharp peaks would be present that match the fingerprint of calcite whether or not\nmagnesium or other trace elements were incorporated into the CaCO3 framework. If magnesium\nwere incorporated into the CaCO3 network, only marginal broadening would be evident in the\nprincipal calcite peaks. The other peaks may be MgCO3 or other minerals. The authors should label\nall peaks in the XRD regardless whether they are attributable to calcite or another mineral\nprecipitate.\nMy suspicion is that there are other crystallized salts (e.g., CaCl2, MgCl2, CaSO4) that are present\na present\nafter the heat treatment the authors conduct. The SEM images (Fig 3a) also give some concern that\nthese are salts not calcite precipitates, as the morphology is not consistent with calcite. Furthermore,\nthe elemental mapping does not show that carbon is collocated with the others, further indicating\nthat these may, in fact, be crystallized salts. The additional peaks in the EDS spectra should be\nlabeled to substantiate the authors’ conclusions regarding the identification of mineral phases. The\nauthors are encouraged to repeat the mineral precipitate characterization experiments using\nthermogravimetric analysis (TGA). TGA is a more appropriate method to confirm the presence\n(or absence) of CaCO3, as it would indicate the onset of thermal degradation of CaCO3, which can\nexist between ~450-700C.\nWe thank the Reviewer for the insightful suggestions on thermogravimetric analysis. We\napologize that the characterization of the precipitates in our material was not convincing from the\nprovided data and compiled additional information. To supplement the original analyses of the\nformed carbonates, we performed extensive additional experiments, including TGA on the\nmineral precipitates (Updated Figure 3b), FTIR analysis of the mineral precipitates (Figure S17),\nand further SEM–EDX analysis of the mineral precipitates (Figure S20). We have also included\nadditional reference patterns in the XRD analysis (Figure S18). All of these additions corroborate\nour conclusion that the formed mineral precipitates are Ca–Mg carbonates and do not contain\nsubstantial amounts of other crystallized salts. We elaborate on each of these additions below\nand believe that this abundance of data supports the conclusions drawn in this work.\nThermogravimetric analysis was performed on the mineral precipitates under a full nitrogen\nenvironment. The decomposition temperature agrees well with the values reported for\ncarbonates in literature and the range mentioned by the Reviewer, thus showing that we indeed\nhave a Ca-carbonate phase. In addition, we performed FTIR analysis of the precipitates which\nshow characteristic peaks (at 873 and 711 cm-1) of carbonate and added it to the Supplementary\nInformation1.\nunder a full nitrogen environment of the remaining precipitates, following removal of\nthe polymer content and bacterial biomass by thermal decomposition, indicated the\npresence of carbonates in the biotic samples42,50.\nFigure S17. FTIR spectra of pure CaCO (black) and the precipitates after one year of\n3\nincubation (green) showing indicative peaks of carbonates at 873 and 711 cm-1. For\ninorganic carbonates ν , represents asymmetrical stretching while ν and ν indicate\n3 2 4\nin-plane and out-of-plane bending, respectively6.\nIn our work, we aimed to obtain crystalline precipitates that are less soluble than amorphous\nones to provide more stable carbon storage than biomass alone. The XRD analysis of the\nprecipitates formed confirmed that they were not amorphous. We rephrased the Results section\nto emphasize the crystallinity aspect and added a reference pattern of aragonite which has a\nsimilar peak pattern for the remaining CaCO related peaks in our precipitate mixture. We added\n3\nadditional reference patterns of calcium carbonates with increasing Mg content for comparison\nof the peak position of (104) plane to XRD pattern and moved it to Supplementary Information\nFigure S18. Such a shift of the peak has been observed in biogenic precipitates1.\n600 °C and reference XRD pattern of aragonite, magnesite, dolomite and calcite7.\nFurther, we checked the presence of other elements at the precipitate site and could not observe\nan increased amount of chloride compared to the rest of the sample. We added SEM–EDS\nelemental mapping of our sample (for Na+ and Cl-) in the Supplementary Information (Figure\nS20a) and labeled additional chloride peaks EDX spectra (updated Figure 3c). We added the\ndata analysis of EDX measurements confirming that Ca to Mg atom % ratios of (number of\nanalyzed samples n = 10) was 78 ± 5 % to 22 ± 5 %, which indicates that both elements were\nembedded in the precipitates. We agree that the intensity of the carbon signal appears low in the\nsame location as oxygen and calcium in the SEM-EDS elemental mapping. However, the carbon\nis visible in the EDX graph of Figure 3c of the area with the precipitates. As carbon is part of the\npolymeric matrix, we expected that the carbon signal will be everywhere as shown, also as a\ndominant peak of the abiotic matrix in Figure 3c. We added the full atom percentage distribution\nfrom two areas of the sample in Figure S20b, which shows that carbon and oxygen are dominant\nelements found in both areas. We changed the text to state that the areas of precipitates are\nenriched with oxygen and calcium signals in the results section.\nRegarding the morphology of the precipitates in the SEM images, we now refer to them as\n‘crystalline carbonate precipitates’ instead of ‘calcite’ as we can confirm crystallinity from XRD\nand carbonates from FTIR and TGA.\nregion with the precipitates of the biotic samples. Scale bar, 20 μm. b) Atom\npercentage distribution in polymer- and precipitate- dominant areas of the sample.\nFigure 3c. EDX update with marked chloride peaks.\nText in the Results section of the original manuscript (page 9):\n“Having demonstrated that photosynthetic living materials are capable of CO\n2\nsequestration via both biomass accumulation and carbonate formation, we further\ninvestigated the composition of the mineral phase produced during incubation. MICP\nresults in the formation of calcium carbonates that vary in stability and solubility with\ncalcite being the most stable polymorph and, therefore, desirable for carbon sinking\npurposes (Figure 3a) 41. XRD analysis of precipitates in biotic samples revealed a\nhighly crystalline structure with the main peaks corresponding to calcite structure with\na noticeable shift of the main diffraction peak representing the (104) plane (Figure 3b).\nThe observed shift likely indicated compositional variation due to the incorporation of\nmagnesium in the carbonate structure as previously observed in marine environments\nand, in this case, due to the use of simulated seawater (ASNIII medium) 42. The XRD\nwas obtained after thermal decomposition (T = 600 °C, which is below the\ndecomposition temperature of CaCO 40) as the signal from the F127-based matrix was\n3\nmuch stronger than the one of the precipitates (Figure S9).\nPrecipitates distributed throughout the polymeric matrix (Figure 3c). Energy dispersive\nX-Ray analysis (EDX) indicated mainly carbon and oxygen in the abiotic matrix\nwhereas peaks of calcium and magnesium were prominent in the areas with\nprecipitates in biotic samples. Elemental mapping corroborated these data with\ncalcium, magnesium, carbon, and oxygen in overlapping regions near clusters of\ncyanobacteria, suggesting the pericellular formation of carbonates (Figure 3d).”\nText in the revised version of the Results section (page 10):\n“Having demonstrated that photosynthetic living materials are capable of CO\n2\nsequestration via both biomass accumulation and carbonate formation, we further\ninvestigated the composition of the mineral phase produced during incubation. MICP\nresults in the formation of crystalline carbonates that vary in stability and solubility with\ncalcite being the most stable polymorph and, therefore, desirable for carbon sinking\npurposes (Figure 3a)41. TGA analysis confirmed carbonate presence in the\nprecipitates (Figure 3b) which was additionally confirmed by FTIR (Figure S17). XRD\nanalysis of precipitates in biotic samples revealed a highly crystalline structure with the\nmain peaks corresponding to calcium–magnesium carbonates indicated by the\nposition of the main diffraction peak representing the (104) plane (Figure S18). The\nobserved shift likely indicated compositional variation due to the incorporation of\nmagnesium in the carbonate structure as previously observed in marine environments\nand, in this case, due to the use of simulated seawater (ASNIII medium) 42. The XRD\nwas obtained after thermal decomposition (T = 600 °C, which is below the\ndecomposition temperature of CaCO 40) as the signal from the F127-based matrix was\n3\nmuch stronger than the one of the precipitates (Figure S19).\nPrecipitates distributed throughout the polymeric matrix (Figure 3c). Energy dispersive\nX-Ray analysis (EDX) indicated mainly carbon and oxygen in the abiotic matrix\nwhereas peaks of calcium and magnesium were prominent in the areas with\nprecipitates in biotic samples. Elemental mapping corroborated these data with\ncalcium, magnesium, and oxygen enrichment in overlapping regions near clusters of\ncyanobacteria, suggesting the pericellular formation of carbonates (Figure 3d and\nS20). EDX measurements also indicated that the Ca to Mg atom % ratio was 78 ± 5 %\nto 22 ± 5 % (number of analyzed samples n = 10), which confirmed that both elements\nwere embedded in the precipitates.”\nText added to the Methods section (pages 18–19):\n“The presence of carbonates in the precipitates was confirmed by thermogravimetric\nanalysis using a TGA/DSC 3+ (Mettler Toledo). The sample was placed inside a 70 μL\nalumina crucible and heated from 25 to 900 °C in pure N under a constant flow rate\n2\nof 150 mL min-1. Buoyancy effects were accounted for by performing measurements\nwith an empty crucible. The sample was first heated to 400 °C and held at this\ntemperature for 30 min to remove any bound water, before continuing heating to 900\n43\n°C at a heating rate of 5 °C min-1. The same measurement conducted with an empty\ncrucible was taken as the baseline and was subtracted from the measured data.\nThe extent of carbonate precipitate formation within abiotic and biotic samples was\nfurther characterized by thermal decomposition at atmospheric pressure. 5 dry abiotic\nor biotic samples were loaded into a porcelain crucible and heated to 600 °C (ramp\nrate = 2 °C min-1) for 2 h. The samples were then cooled to room temperature and the\nremaining mass was measured. In addition, the remaining precipitates were collected\nfor attenuated total reflectance Fourier transform infrared spectroscopy (ATR-FTIR,\nSpectrum Two, PerkinElmer; 1 cm-1 resolution, 32 scans) as well as X-ray diffraction\n(XRD) analysis with a PANalytical Empyrean diffractometer (Cu Ka radiation, 45 kV\nand 40 mA) with an X'Celerator Scientific ultrafast line detector and Bragg-Brentano\nHD incident beam optics. XRD samples were measured over the 2θ range 10–90° for\n1 h with a step increment of 0.016°. Results were compared with reference patterns\nfrom the ICDD database.”\n7. Hydrogel degradation and mechanical properties – Because of progressive dissolution of pristine\nF127, the authors observed a significant mass loss overtime (approx. 45% of the original mass as\nreported for the abiotic samples in Supplementary). This should translate in a significant reduction\nof mechanical properties for the abiotic samples, as well as an initial decrease for the biotic ones\nfollowed by progressive increase due reinforcement because of MICP deposition overtime.\na. How do the authors explain that no differences were observed for the abiotic samples between\nday 2 and 30, despite the remarkable mass loss?\nWhile the formulation with pristine F127 is a viscoelastic solid, photo-cross-linking of the\nF127:F127-BUM hydrogel resulted in an increase of the storage modulus from 16 kPa to 48 kPa\nprior to swelling (Figure 1e and S9). For improved clarity, we have updated the axes in the\nupdated Figure 1. Therefore, we concluded that the main contributor to the storage modulus in\nthe samples tested (abiotic and biotic) was the chemically cross-linked F127-BUM network.\nTherefore, dissolution of the weak viscoelastic F127 that was not cross-linked would not be\nexpected to make a substantial change in the final material properties. In addition, the reported\nmechanical properties were measured on day 2 and day 30, as we first allowed the samples to\nequilibrate in the culture medium prior to the initial mechanical measurement. Partial dissolution\nof non-cross-linked F127 from the hydrogel matrix could have already occurred prior to the\nmeasurement on day 2. Measurements on day 0 were not included as sufficient time is needed\nfor the samples to equilibrate their swelling state for proper mechanical analysis. Given this, any\npotential dissolution of pristine F127 did not result in a significant drop in modulus between day\n2 and day 30 samples.\nb. Adding additional intermediate timepoints would better characterize when MICP deposition\nbecomes relevant for material reinforcement.\nWe thank the reviewer for the comment. We performed rheological measurements and\nquantitative Alizarin Red S staining at intermediate time points to further assess when the MICP\ndeposition became relevant for material reinforcement. We observed that precipitate\naccumulation was > 1 mg mL-1 of hydrogel already after 10 days (see answer to R3.5c). A\nsignificant change in mechanical properties compared with day 1 was also observed after 10\n44\ndays (p = 0.05, Figure S21). No significant change was observed in the mechanical properties\nof abiotic samples throughout the time course.\nFigure S21. The storage modulus (G′), measured by shear rheometry, of biotic (green)\nand abiotic (gray) samples during the 30-day incubation period. A significant change\nin mechanical properties compared with day 1 was also observed after 10 days (p =\n0.05).\nText in the Results section of the original manuscript (page 9):\n“The formation of precipitates in hydrogel matrices via MICP can reinforce the\nmechanical properties of the material17,43,44. The storage moduli (G′) of the printed\nbiotic samples on day 2 of the experiment were slightly lower compared with abiotic\nsamples (5.4 ± 2 kPa and 6.7 ± 2 kPa, respectively).”\nRevised text in the Results section of the manuscript (page 10):\n“The formation of precipitates in hydrogel matrices via MICP can reinforce the\nmechanical properties of the material17,43,44. The storage moduli (G′) of the printed\nbiotic samples on day 2 of the experiment were slightly lower compared with abiotic\nsamples (5.4 ± 2 kPa and 6.7 ± 2 kPa, respectively). A significant increase (p = 0.05)\nin storage moduli (G′) in the case of biotic samples was observed already after 10 days\n(Figure S21).”\nText added in the Methods section (page 19):\n“Viscoelastic properties of the printed hydrogels at different time points (n = 3;\nbiological replicates) during incubation were evaluated using the same rheometer as\nfor the bioink characterization equipped with a Peltier stage and 8 mm sandblasted\nprobe (gap size h = 0.15–0.35 mm, normal force F = 0.1 N) at 25 °C.”\nN\nc. Statistical analyses should be included to confirm significant differences in mechanical\nperformance, if any.\nWe performed a two-sample t-test and added the p values to the graphs of mechanical analysis\nof Figure 3 where significant differences were found between samples. In case of rheology\nresults, on day 2 as well as between day 2 and day 30 of abiotic samples there was no significant\n45\ndifference p > 0.05. There were also no significant differences between samples in the toughness\nmeasurements p > 0.05. We added the methods of statistical analysis to the Methods section.\nFigure 3e,f. Statistical analysis added to the graphs.\nText added in the Methods section (page 20):\n“Statistical analysis of the samples at different time points was performed using two-\nsample t-tests (OriginPro 2019 software). All values are shown as average with\nstandard deviation or root mean square of standard deviations in cumulative cases.”\nRevised text in the Results section of the manuscript (page 10):\n“Over the course of 30 days, the modulus increased significantly to 10.1 ± 1 kPa (p =\n0.01) in the biotic case while it did not change for abiotic samples (6.5 ± 1 kPa) (Figure\n3e).”\nIn case of tensile testing, we updated the number of biological replicates in the Methods section\n(page 19).\n“To obtain homogeneous samples for tensile testing, abiotic and biotic bioink were\nmolded into a rectangular-shaped mold (12 mm × 50 mm × 1 mm, width x length x\nthickness) and cross-linked for 2 min (λ = 405 nm light, I = 8 mW cm-2). Samples were\nthen washed and incubated under the same conditions as the disc-shaped samples.\nUniaxial tensile testing was performed on the samples (n = 5; biological replicates)\nincubated for 2 and 30 days using a tensile testing machine (Stable Micro Systems)\nequipped with two parallel metal screw-clamps.”\n8. Volumetric printing of lattice structures – Volumetric 3D printing was adopted to achieve high\nresolution and a defined internal porosity. However, this cannot be appreciated from brightfield\nimages reported in Fig 4,b.\na. SEM of the external surface and internal cross-sections of the final structure would validate that\nthe desired high resolution and internal defined porosity was achieved.\nWe added images of the external surface (updated Figure 4a) as well as cross sections of\nlyophilized sample (Figure S25) prepared by volumetric printing to confirm the porosity of the\nsample to the Supplementary Information.\nScale bar, 1 cm. b) SEM image of the same sample with lattice structure falsed colored\nin blue. Scale bar, 1 mm. c) SEM image of the cross-section of the sample prepared by\nvolumetric 3D printing. Scale bar, 1 mm.\nUpdated Figure 4a:\nFigure 4. Digital fabrication of photosynthetic living structures for dual carbon\nsequestration. a) Lattice structures were digitally designed to facilitate light exposure\nand medium transport via passive capillary wetting, enabling growth of the photosynthetic\nliving materials. The structures were fabricated using volumetric printing. The porosity of\nthe printed structure can be observed in the lyophilized sample.\nb. False color high resolution SEM images of seeded samples should also be provided to better\nunderstand localization of cells within the hydrogel network.\nWe added false color high resolution SEM images of cells embedded in the polymer (Figure\nS1a) to the Supplementary Information. We also added lower magnification images of the same\narea for context (Figure S1b–d).\nrepresentative cluster of embedded cyanobacteria Synechococcus sp. PCC 7002 are\nhighlighted in green false color to indicate the location within the supporting matrix. It\nshould be noted that the sample was homogeneously populated with embedded\ncyanobacteria that have not been false colored, as best seen in panel b). Scale bars,\na) 1 µm, b) 10 µm, c) & d) 100 µm.\n9. Long-term viability evaluation of 3D printed photosynthetic living materials – The authors report\nextended cells viability for over a year.\na. To better characterize cell proliferation within the lattice structure overtime, quantitative cell\nviability assays should be performed.\nTo assess cell viability, we performed dead cell staining at different time points and evaluated\noverall biomass in the sample by chlorophyll content (Figure S34). After reaching a steady state,\nthe encapsulated cells were > 98% viable throughout the time course of analysis.\nconcentration in encapsulated disc samples obtained via confocal microscopy max\nprojection (day 0, day 1, and all dead cell data) or chlorophyll extraction (live cells day\n15–30).\nAdded sentence in the Discussion section (page 14):\n“With minimum requirements of sunlight and atmospheric CO , our photosynthetic\n2\nliving materials showed a consistent carbon sequestration efficiency throughout the\nentire incubation period, and demonstrated the potential of using natural carbon\nsequestration processes at scale60. We expect the system to reach its peak\nphotosynthesis rate early as the cells go through an initial exponential growth phase\nand later operate at a pseudo-steady state while maintaining high viability (Figure\nS29).”\nb. The authors indicate that transparency of the hydrogel facilitates light penetration and cell growth\nwithin the structure. However, because of progressive mineral deposition, the structure loses its\ntransparency overtime. Do the authors expect the reduce light penetration to affect viability of the\ninner layers of the structure? Evaluation of cell distribution within the material thickness overtime\nshould also be investigated by providing representative images of material cross sections at\ndifferent timepoints (both brightfield images as well as fluorescently stained live and dead cells).\nWe agree that precipitates would decrease the transparency of the matrix. However, from our\nobservations, cell encapsulation and further growth played a larger role in the decrease in\ntransparency (Figure S3). We observed a 5 mm depth to be an optimal material thickness to\nmaintain PCC 7002 viability (Figure S27). This motivated the design parameters for the textured\nsurfaces shown in Figure 4d. We added a comment regarding light penetration to the Results of\nthe revised manuscript as well as images of cell growth in living material 5 x 5 x 5 mm cube cross\nsections (Figure S24) with cell growth viability differences at the edges and in the center of the\ncube (Figure S24). The cells grow in big clusters at the edges while small clusters and individual\ncells are more prominent in the center. We assume that the lower cell growth in the deeper\nregions was due to light shielding from cell clusters in the outer regions of the cube.\nScale bar, 5 mm. Cell viability of samples from b) edge and c) center of a cross section\nof a 5 x 5 x 5 mm printed cube (SYTOX Blue dead cell staining, cyan; cell\nautofluorescence, green). Scale bar, 100 μm.\nText added to the Results (page 12):\n“The transport of gases and liquid medium as well as access to light are essential for\nthe efficient functioning of biologic processes (including carbon sinking) in the\nphotosynthetic living materials9,31. The decrease in light penetration over the course of\nthe life cycle is expected due to increased light absorption and scattering with\nincreased biomass, resulting in reduced proliferation in the volume fraction further from\nthe surface (Figure S24).”\ntext or supplementary information have been highlighted in yellow.\nReviewer #1 (Remarks to the Author):\nThe authors have performed several additional experiments. My comments are mostly related to\nthe additional data:\n1. Figure S11a: The BG11-ASNIII medium was changed every 5 days, so additional carbon\nsource is made available (potentially) each time that the medium is refreshed. How is that taken\ninto consideration?\nWe agree that this is an important point and we apologize that the consideration of added carbon\nwas not explained clearly in the original manuscript. We included sodium carbonate (Na CO )\n2 3\nin the medium to simulate seawater; however, the amount of Na CO present in the medium\n2 3\nwas below the level of dissolved inorganic carbon (DIC) at equilibrium under our operating\nconditions. Therefore, additional carbon was not made available each time the medium was\nrefreshed. When quantifying the DIC content after each 5-day period, we re-collected all the\nmedium and analyzed the total DIC after the 5 days of culture. Using this methodology, there\nwas no way for sequential accumulation of DIC from the medium itself and we were able to\ncompare the DIC in the medium after each 5-day culture period in the basal medium. The\namount of DIC in the re-collected culture medium was at or above the amount of inorganic\ncarbon initially present from the Na CO in the medium for each culture period (former Figure\n2 3\nS11a now Figure S1a). This means that during each 5-day culture period, additional\natmospheric carbon dissolved in the culture medium to maintain the equilibrium amount of DIC,\nas illustrated in Figure 1a. As the initially available inorganic carbon from Na CO did not affect\n2 3\nthe DIC equilibrium, the change in medium did not make additional carbon available.\nTo further support that the DIC equilibrium under these conditions was at or above that of the\ninitially supplemented carbon, we conducted additional DIC analyses on Na CO -free BG11–\n2 3\nASNIII medium at pH = 7 and pH = 9 (Figure S1c). Again, the DIC content in the medium\nwithout Na CO was higher than 381 μmol/L, which is the amount of Na CO in the culture\n2 3 2 3\nmedium. This indicates that the DIC equilibrium is not dependent on the presence of Na CO in\n2 3\nthe simulated seawater medium and that atmospheric carbon did dissolve in the culture medium\nunder all conditions.\nWhile no additional carbon is made available to the system due to the Na CO in the medium,\n2 3\nsome of the carbon in the final construct comes from the inorganic carbon from the Na CO in\n2 3\nthe culture medium, which was added to simulate the mineral content of seawater. In the\nprevious revision, we had already softened all language about atmospheric CO being the sole\n2\ncarbon source for carbon sequestration throughout the manuscript for this reason.\nFigure S1. a) Total amount of dissolved inorganic carbon (DIC) under equilibrium in the re-\ncollected abiotic (black) and biotic (green) medium after every 5-day culture period (n = 2, with\npooling of 20 individual samples in each case). The gray dashed line indicates the amount of\nDIC in the form of Na CO that was used in simulated seawater medium (BG11–ASNIII). b)\n2 3\npH changes of pooled biotic (green) and abiotic (black) medium every 5 days (n = 3). c) DIC\nanalyses on BG11–ASNIII medium without the addition of inorganic carbon through sodium\ncarbonate (Na CO ) at pH = 6.7 and pH = 8.9 (n = 2, black). The gray dashed line indicates\n2 3\nthe amount of DIC in the form of Na CO that was used in simulated seawater medium (BG11–\n2 3\nASNIII).\n2\ninduced carbonate precipitation (MICP) during photosynthesis using dissolved CO as the major carbon\n2\nsource.\nUpdated text in the Results section (page 7)\n“40.4 mg L-1 (0.38 mM) of Na CO was dissolved in the medium to simulate natural seawater.\n2 3\nAnalysis of the total DIC under equilibrium in the culture medium indicated that the DIC content\nwas above or at the quantity of carbon initially dissolved in the form of CO 2- (Figure S1). This\n3\nincrease in the DIC confirmed the dissolution of atmospheric CO , which served as the primary\n2\ncarbon source for CO sequestration.”\n2\n2. Figure S11a: Why are the values for the DIC in the abiotic medium varying like this with a peak\nat 20 days? Please also include the number of replicates and error bars.\nThe observed variation in dissolved inorganic carbon (DIC) value at day 20 was likely due to\ncumulative variability in the measurements. The DIC in the medium was calculated based on\nseveral parameters, including the medium pH, medium conductivity, salinity, and alkalinity.\nEach measurement of these parameters incurred a certain degree of variability, and the error\nbar in the final plot for DIC reflects the combined uncertainty from all the factors. The abiotic\nsamples exhibited increased variability starting from day 20—this variability was also observed\nin the pH data (Figure 1b). In contrast, the variability was less pronounced in the biotic samples,\nespecially in the later stages of the experiments.\nThe DIC calculation consisted of two sets of experiments, and each data point was obtained by\npooling the medium from 20 individual samples, which was required for more accurate titration\nresults.\nThe updated former Figure S11 now Figure S1 and caption appended below:\ncollected abiotic (black) and biotic (green) medium after every 5-day culture period (n = 2, with\npooling of 20 individual samples in each case). The gray dashed line indicates the amount of\nDIC in the form of Na CO that was used in simulated seawater medium (BG11–ASNIII). b)\n2 3\npH changes of pooled biotic (green) and abiotic (black) medium every 5 days (n = 3). c) DIC\nanalyses on BG11–ASNIII medium without the addition of inorganic carbon through sodium\ncarbonate (Na CO ) at pH = 6.7 and pH = 8.9 (n = 2, black). The gray dashed line indicates\n2 3\nthe amount of DIC in the form of Na CO that was used in simulated seawater medium (BG11–\n2 3\nASNIII).\n3. The amount of CO2 sequestered by precipitation is less than 0.1% of the whole mass of the\nmaterials. Based on Figure S16 (biotic culture) and Supplementary text on “CO2 sequestration\nby biomass growth (pH change) in living materials”, the precipitates’ weight is fairly low, so the\ndual carbon sequestration does not appear really efficient.\nWe have removed references to this process as being “efficient” throughout the manuscript and\ninstead focused on reporting the quantitative amounts of CO sequestered per unit mass of wet\n2\nhydrogel. The main text has been adapted accordingly.\nText in the original version of the Abstract:\n4\n“Digital design and fabrication of the living material ensured sufficient access to light and\nnutrient transport of the encapsulated cyanobacteria, which were essential for long-term\nviability (more than one year) as well as efficient photosynthesis and carbon sequestration.”\nUpdated text in the revised version of the Abstract:\n“Digital design and fabrication of the living materials ensured sufficient access to light and\nnutrient transport of the encapsulated cyanobacteria, which were essential for long-term\nculture (more than one year), photosynthesis, and carbon sequestration. […] These findings\nhighlight the potential of photosynthetic living materials for scalable carbon sequestration with\npossible future use in carbon-neutral infrastructure and green building materials.”\nText in the original version of the Results (Page 8):\n“CO sequestration via MICP was approximately one order of magnitude more than that\n2\nachieved via biomass growth, making the dual CO sequestration a more efficient carbon sink,\n2\nand indicating that the majority of the sequestered CO had been stored in a more stable\n2\nmineral form.”\nUpdated text in the revised version of the Result (Page 8):\n“CO sequestered via MICP has been stored in a more stable mineral form.”\n2\nbelieve that presenting the mass of carbon sequestered per unit mass is a more meaningful\nand representative value—both for potential future applications of this approach and for\ncomparison with other technologies. That said, the amount of CO sequestered via carbonate\n2\nprecipitation was quantified as 2.2 ± 0.9 mg of CO per gram of wet hydrogel material over a\n2\nculture period of 30 days, increasing to 26 ± 7 mg over 400 days. This corresponds to a mass\npercentage of ~2.6% after 400 days and ~0.22% after 30 days, based on wet (whole) mass of\nthe materials. In addition, we prefer to report these values in quantitative terms per unit mass\nof wet hydrogel rather than as percentages given the continuous nature of the sequestration in\nour living materials.\n4. Concerning the carbon sequestration, is it possible to provide a quantitative analysis of the\ncarbon sequestration contribution from each factor? (e.g., biomass accumulation, carbonate\nprecipitation without the source of sodium bicarbonate in ASNIII-BG11).\nWe thank the Reviewer for the comment and we would first like to clarify the culture medium\ncomposition. We did not add sodium bicarbonate (NaHCO ) to BG11–ASNIII, only sodium\n3\ncarbonate (Na CO ), which was included in the culture medium to simulate the composition and\n2 3\nionic strength of seawater. As the samples and culture medium were exposed to the open\natmosphere, a carbon equilibrium was established according to Henry’s Law (Figure 1a). As\nseen in Figure S1a of the updated manuscript, that initial dissolved inorganic carbon (DIC)\nconcentration was double that of the carbon content from Na CO alone. Therefore, the carbon\n2 3\nintegrated into biomass and insoluble carbonates is likely coming from both the atmosphere\nand the inorganic carbon included in the medium. In our current experimental setup, the amount\nof carbon (CO ) sequestration via carbonate precipitation was 2.2 ± 0.9 mg of CO per gram of\n2 2\nwet material after 30 days. We also observed an increase in cell number from day 0 to day 30,\nfrom an initial cell concentration of 3 x 107 cells mL -1 to a final cell concentration of 5 x 109\ngel\n5\ncells mL -1. While biomass increased during the same culture period, we were not able to\ngel\nprovide a quantitative measure of the CO sequestered into this biomass. Approximations\n2\nbased on cell counts, chlorophyll extraction, and medium pH indicated that the CO\n2\nsequestration by biomass accumulation was likely to be a minor fraction of the total CO\n2\nsequestered by our photosynthetic living materials (approximately one order of magnitude less\nthat by carbonate precipitation). However, as the CO sequestered into this biomass were\n2\napproximated based on the cell counts, chlorophyll extraction and medium pH, we only report\nthe quantitative measure of CO sequestration via carbonate precipitation in the revised form of\n2\nthe manuscript.\nIn the future, we agree that more quantitative measures of the relative contributions to the\ncarbon sequestered should be conducted. Such experiments could be carried out in closed\nsystems, and preferably large-scale experiments. These experiments should control carbon\ninput into the system and more accurately quantify the rate of photosynthesis via the direct\nmeasurement of O evolution. In this manner, one may be able to quantitatively assess the\n2\nrelative contributions of biomass accumulation and carbonate precipitation as well as to provide\na more precise number of the amount of atmospheric carbon sequestered per unit mass of\nmaterial and per unit time. We have revised the Discussion to indicate the importance of such\nfuture studies in the field.\nRevised text in the Discussion (pages 14 and 15):\n“[...] Future work should include a more quantitative evaluation of CO sequestration through\n2\nbiomass accumulation to better understand the photosynthetic rate of the cyanobacteria.\nDirect measurements of O evolution using an oxygen electrode would provide a more\n2\naccurate quantification of photosynthesis. Additionally, established computational models\ncould be applied to further explore the relationship between cytosolic pH of the cyanobacteria\nand CO assimilation into biomass65.”\n2\n“Upscaling would provide the means to perform more robust CO sequestration analyses\n2\nusing gas composition monitoring (Figure S30) as well as control and tracking of carbon\nsources via media and atmospheric supply.”\n5. Figure S30: The CO2 consumption rate data is quite noisy, and a very low R2 value (0.15) is\nshown on panel b. Also, in panel b, is there a reason why the x-axis is from 0.5 to 1.4h? Please\nalso include the number of replicates and error bars.\nThe measurement was conducted by measuring the CO concentration within a closed\n2\ncontainer over 10 days. Upon closing the container, the setup was allowed to equilibrate for\napproximately 30 min, and therefore, the x-axis starts from 0.5 h. Each data point was obtained\nevery 4–5 s (Figure S30b). The low R2 value was due to the high sampling frequency of the\nCO concentration within the closed container, as any external fluctuation (in temperature,\n2\nvibration, or movement) caused the value to vary.\nWe added error bars in Figure S30a for the 10-day measurement (n = 2). We have also updated\nthe Methods section accordingly to better explain how these data were acquired.\nUprated Figure S30:\n2\nmeasurement period of 10 days with a 0.25 mL sample and 10 mL culture medium (n = 2).\nCO sequestration was observed from day 3 and plateaued beyond day 5. b) Representative\n2\nsnapshot of CO evolution in a closed container and exemplary linear interpolation of\n2\nthe measurement (blue) used to calculate the rate of change of CO concentration.\n“Direct CO measurement in a closed environment\n2 2\n(Sensirion I2C SCD4x Multiple Function Sensor), and an Arduino microcontroller board (Uno\nr4 minima). An SD card reader was connected to the Arduino board to record the data. Two\n8.2 kΩ resistors were added into the electric circle of the sensor to amplify the voltage signal\nof the two data cables. A 0.25 mL sample with approximately 10 mL of culture medium was\nplaced in each measurement container. The containers were opened every day and were\nallowed to equilibrate with atmospheric air to replenish the CO within the closed system. The\n- The authors use expressions like carbon-negative and carbon-neutral infrastructure in the\nmanuscript. To use such wording, it would require a quantitative cradle-to-grave CO2 emission\nanalysis (LCA).\nWe thank the reviewer for the comment and we agree that a quantitative life cycle analysis\nwould be needed for these materials. We have softened our statement and included a\ncomment related to the need for life cycle assessment in the revised Discussion.\nOriginal text in the manuscript (page 15):\n“Photosynthetic living materials also hold promise for future applications as surface\ncoatings for green building materials or bioreactors in commercial-scale sequestration\nplants, enabling bioremediation of CO emissions and supporting the realization of\n2\ncarbon-neutral to carbon negative infrastructure.”\nUpdated text in the manuscript (page 15):\n“Photosynthetic living materials also hold promise for future applications as surface\ncoatings for green building materials or bioreactors in commercial-scale sequestration\nplants, enabling bioremediation of CO emissions and could potentially support the\n2\nreduction of CO emission in the built environment. However, a comprehensive life cycle\n2\nassessment would be necessary to quantify the net carbon impact of such systems.”\n- Caption of Figure S11a: Shouldn’t that be BG11-ASNIII medium?\nWe thank the reviewer for pointing this out and we have corrected this accordingly.\n- Typo in Figure S11: mediuim (twice)\nWe thank the reviewer for noticing the typo and these have been corrected.\n- Typo in Figure 2a: medum\nWe thank the reviewer for noticing the typo and the word has been corrected.\nAfter reading the authors’ Response to Reviewers, I appreciate the addition of cell counts and\nchlorophyll measurements to support the conclusions regarding biomass growth. However, I\nremain unconvinced that measuring extracellular pH is a valid way to quantify uptake of inorganic\ncarbon into biomass in this system, which is open to the atmosphere. The standard way to quantify\nphotosynthesis in cyanobacteria is to measure O2 evolution using an oxygen electrode or by\nmembrane inlet mass spectrometry.\nI understand that there are several references (cited in the Response to Reviewers) that describe\nthe release of OH- ions by carbonic anhydrase located in the carboxysome as part of a model for\nthe cyanobacterial CCM and MICP, but this is an oversimplification that is not biochemically\ncorrect. A paper on quantitative modeling of the CCM in beta-cyanobacteria (Mangan et al., 2016.\nPNAS 113: E5354-E5362) notes that the combined action of CA and Rubisco in the carboxysome\nactually produces a net H+, not an OH- ion.\nThe statements on p. 4 (“Here, carbonic anhydrase (CA) within the carboxysome converts the\nbicarbonate species into CO2, releasing a hydroxyl ion (OH-) into the surrounding media7,33–35.\nThe OH- is excreted to the exterior of the bacterium....”) and p. 7 (“Using HCO3- as a substrate,\ncarbonic anhydrase generates equimolar amounts of OH- ions, which are excreted into the\nsurrounding medium, and CO2 molecules, which are integrated into bacterial metabolism.17,27”)\nand the depiction in Fig. 1 are incorrect.\nWe thank the Reviewer for the additional comments on this point and we agree that\ncomplementing the initial data in the original version of the manuscript with cell counts and\nchlorophyll measurements were important and helpful additions to corroborate the conclusions\nrelated to biomass growth. In order to provide an estimate of the amount of CO sequestered\n2\nvia biomass, we had initially used the increase in pH and these data were in agreement with\nthe cell counts and chlorophyll measurements. However, to avoid any incorrect statements and\nto focus on the central conclusions of the manuscript, we have removed the pH data from the\nmain text and no longer use pH changes as a metric to quantify biomass accumulation. In\naddition, we have softened the statements on this topic pointed out by the Reviewer to avoid\nconfusion or incorrect statements related to these points. Further, we would like to re-emphasize\nthat while biomass accumulation is a necessary feature of our system, our approximate values\nindicated that it was only a minor fraction of the CO sequestered by these photosynthetic living\n2\nmaterials.\nWe have now rephrased the text on both page 4 and page 7 to avoid any misstatements as\npointed out by the Reviewer:\nOriginal text on page 4:\n“We engineered photosynthetic living materials for dual carbon sequestration by exploiting the\ncarbon concentrating mechanism of cyanobacteria strain PCC 7002 (Figure 1a). Dual CO\n2\nsequestration proceeds via both reversible biomass accumulation and irreversible mineral\nprecipitation. For carbon concentration to occur, CO from the surrounding environment\n2\ndissolves in aqueous solutions and forms bicarbonate ions (HCO -) that are then transported\n3\nto the marine β-cyanobacteria 7002 carboxysome33. Here, carbonic anhydrase (CA) within\nthe carboxysome converts the bicarbonate species into CO , releasing a hydroxyl ion (OH-)\n2\ninto the surrounding media7,33–35. The OH- is excreted to the exterior of the bacterium and the\n2\nphotosynthesis into two molecules of 3-phosphoglycerate, which is enzymatically converted\ninto sugars that support cell growth and biomass production. Due to the metabolic production\nof OH-, the local pH value of the local medium surrounding the bacteria increases. This\nincrease in pH, combined with negatively charged extracellular polysaccharides on the\nbacterial membrane, creates a favorable environment for the nucleation and formation of\ninsoluble carbonates31. In the presence of divalent cations, such as Ca2+ and Mg2+, CO 2- in\n3\nthe medium is consumed and fixed irreversibly into calcium or magnesium carbonates.\nChemical equilibrium then favors the dissolution of additional atmospheric CO into the culture\n2\nmedium, continuously driving the dual carbon sink.”\nRevised text (Page 4):\n“We engineered photosynthetic living materials for dual carbon sequestration by exploiting the\ncarbon concentrating mechanism of cyanobacteria strain PCC 7002 (Figure 1a). Dual CO\n2\nsequestration proceeds via both reversible biomass accumulation and irreversible mineral\nprecipitation via MICP. For carbon concentration to occur, CO from the surrounding\n2\nenvironment dissolves in aqueous solutions and forms bicarbonate ions (HCO -) that are then\n3\ntransported to the marine β-cyanobacteria 7002 carboxysome33 Within the carboxysome,\ncarbonic anhydrase (CA) catalyzes the conversion of bicarbonate into CO , which is then fixed\n2\nby ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCo) during photosynthesis into\ntwo molecules of 3-phosphoglycerate. This product is enzymatically converted into sugars\nthat support cell growth and biomass production, representing the reversible part of CO\n2\nsequestration.\nIrreversible CO sequestration as mineral precipitation occurs outside the cells, near the\n2\ncyanobacteria. Negatively charged extracellular polysaccharides on the bacterial membrane\ncombined with a suitable extracellular environment (alkaline pH, presence of divalent cations),\nfacilitates the nucleation and formation of insoluble carbonates33–37. Thus, by culturing\nphotosynthetic living materials in simulated seawater that contains Ca2+ and Mg2+, an increase\nin medium pH, which occurred in cyanobacteria-containing samples (Figure S1), allows CO 2-\n3\nin the medium to be fixed irreversibly into calcium or magnesium carbonates. Chemical\nequilibrium then favors the dissolution of additional atmospheric CO into the culture medium,\n2\ncontinuously driving the dual carbon sink.”\nOriginal text on page 7:\n“PCC 7002 growth within the living material was observed directly by microscopy and\nindirectly by measured pH changes in the surrounding medium (Figure 2b). After 10 days of\nincubation samples exhibited visible clusters of PCC 7002 cells as well as many individual\ncells throughout the volume of the hydrogel matrix. The number of clusters and individual cells\nincreased with time (Figure S12). The change in pH of the culture medium was attributed to\ncarbonic anhydrase activity in the carboxysome of the bacterial cells. Using HCO - as a\n3\nsubstrate, carbonic anhydrase generates equimolar amounts of OH- ions, which are excreted\ninto the surrounding medium, and CO molecules, which are integrated into bacterial\n2\nmetabolism17,27. As the medium was changed every 5 days, the cumulative amount of OH-\nions was reflected in the pH increase of the medium from the initial pH value of 6.5. The\ncumulative OH- ion production served as a surrogate measure of carbonic anhydrase activity\nand CO integration into biomass 40. Between days 5 and 10, the pH of the biotic samples\n2\n10\nincreased to 9.5, corresponding to approximately three orders of magnitude increase in OH-\nconcentration. In each subsequent 5-day interval, the pH increased by 1.8–2.9 with an\naverage of more than two orders of magnitude increase in OH- concentration (Figure S11b).\nThe cumulative CO sequestered was calculated based on the pH change of the medium\n2\n(calculations in the Supplementary Information, CO sequestration by biomass growth (pH\n2\nchange) in living materials). Over the first 15 days the cumulative amount of CO\n2\nsequestered per gram of living material was 3.3 µmol (0.15 mg) and reached a value of 7\nµmol (0.31 mg) by the end of the 30-day experiment, an increase that reflects CO\n2\nsequestration corresponding to biomass accumulation (Figure 2d). Beyond 30 days, we\nexpect the biomass accumulation to reach a pseudo-steady state (with similar cell death and\ngrowth rates) and the component of carbon sequestration related to biomass accumulation to\nequilibrate within the hydrogel. The pH did not change in abiotic control samples throughout\nthe incubation (Figure S11b) and the majority of the encapsulated bacteria remained within\nthe hydrogel throughout the incubation in biotic samples (Figure S13).”\nRevised text (Page 7):\n“PCC 7002 biomass growth within the living material for CO capture was evaluated by\n2\nfluorescence microscopy (cell counts) and chlorophyll extraction (Figure 2b). After 10 days\nof incubation samples exhibited visible clusters of PCC 7002 cells as well as many individual\ncells throughout the volume of the hydrogel matrix. The number of clusters and individual cells\nincreased over time with cell concentrations reaching 5 x 109 cells mL -1 after 30 days (Figure\ngel\n2d, S12). Further, the choice of initial OD , within the range of 0.3–1.6, did not impact cell\n730nm\ngrowth (Figure S13a). Moreover, the majority of the encapsulated bacteria remained within\nthe hydrogel throughout the incubation in biotic samples (Figure S13b). Beyond 30 days, we\nexpect the biomass accumulation to reach a pseudo-steady state (with similar cell death and\ngrowth rates) based on the plateau observed in the cell concentration beyond day 25.”\nIn addition, we have removed the arrow indicating the secretion of OH- ions by the cyanobacteria\ninto the extracellular environment in Figure 1a to avoid confusion or incorrect statements and\nupdated Figure 2d to focus on the increase in cell concentration over time as measured by cell\ncounts and chlorophyll extraction. The corresponding sentence in the Abstract was also\nchanged accordingly.\nOriginal text in the Abstract:\n“Additionally, the metabolic production of OH- ions in the surrounding medium created an\nenvironment for the formation of insoluble carbonates via microbially-induced calcium\ncarbonate precipitation (MICP). Digital design and fabrication of the living material ensured\nsufficient access to light and nutrient transport of the encapsulated cyanobacteria, which were\nessential for long-term viability (more than one year) as well as efficient photosynthesis and\ncarbon sequestration. The photosynthetic living materials sequestered approximately 2.5 mg\nof CO per gram of hydrogel material over 30 days via dual carbon sequestration, with 2.2 ±\n2\n0.9 mg stored as insoluble carbonates.”\nRevised text in the Abstract:\n11\n“Additionally, the living materials created a favorable environment for the formation of\ninsoluble carbonates via microbially-induced calcium carbonate precipitation (MICP). Digital\ndesign and fabrication of the living materials ensured sufficient access to light and nutrient\ntransport of the encapsulated cyanobacteria, which were essential for long-term culture (more\nthan one year), photosynthesis, and carbon sequestration. The photosynthetic living materials\nsequestered 2.2 ± 0.9 mg of CO per gram of hydrogel material in the form of insoluble\n2\ncarbonates over 30 days, with continuous increase in cyanobacteria cell number, enabling\nfurther CO sequestration in the form of biomass.”\nFigure 1. a) PCC 7002 are capable of CO sequestration via biomass accumulation and microbially\n2\ninduced carbonate precipitation (MICP) during photosynthesis using dissolved CO as the major carbon\n2\nsource.\nin the encapsulated samples (n = 3), with an initial rapid growth rate up to day 15 followed by\na slower growth rate from day 15 to day 30 (green). Liquid suspension with the same starting\ncell concentration was used as a control of cell growth (black).\nFinally, we agree that more sophisticated methods can be used to quantify photosynthesis in\ncyanobacteria, such as direct measurements of O evolution using an oxygen electrode or using\n2\nmembrane inlet mass spectrometry. We have included a comment on this in the Discussion as\nan important focus for future work, from our group and others, that aim to quantitatively measure\nthe photosynthetic rate of encapsulated cyanobacteria in photosynthetic living materials.\nThe updated text in the Discussion now reads (page 13):\n“[...] We expect the system to reach its peak photosynthesis rate early as the cells go through\nan initial exponential growth phase and later operate at a pseudo-steady state while\nmaintaining high viability (Figure S29). Future work should include a more quantitative\nevaluation of CO sequestration through biomass accumulation to better understand the\n2\nphotosynthetic rate of the cyanobacteria. Direct measurements of O evolution using an\n2\noxygen electrode would provide a more accurate quantification of photosynthesis.\nAdditionally, established computational models could be applied to further explore the\nrelationship between cytosolic pH of the cyanobacteria and CO assimilation into biomass65.”\n2\nGeneral Comments:\nThe manuscript describes 3D-printed hydrogel-based materials as novel tools for dual carbon\nsequestration. Photosynthetic living materials were engineered by embedding cyanobacteria\nwithin a bioinert hydrogel matrix, to achieve effective dual carbon sequestration through biomass\naccumulation and irreversible microbially-induced calcium carbonate precipitation. The authors\ninvestigated the platform’s performance not only in the short term (1 month) but also conducted\nlong-term studies to demonstrate the platform’s ability in forming biomass and producing mineral\nprecipitation for over a year.\nWhile the authors have made commendable efforts to address the reviewer’s questions, some\nquestions were not adequately addressed. The reviewer acknowledges these efforts but believes\nthat several questions need further clarification. Therefore, the manuscript is not recommended\nfor publication in its current state.\nTechnical Comments:\n1. Hydrogel formulation\na. The authors have adequately addressed the printability of Pluronic F127 hydrogels of\ndifferent polymer contents. However, the images in Figure S5 are not consistent in lighting,\nresulting in difficulty to access the resolution of prints. Higher-quality images with proper and\nconsistent lighting are recommended.\nTo improve the quality of the transparent hydrogel images, we took additional pictures of the\nprinted samples using a camera (alpha 7R, Sony) equipped with a macro lens (FE 2.8/90\nMACRO G OSS, Sony) with consistent lighting.\nWe have updated the former Figure S5, current Figure S6 with these new images.\nFigure S6. Printability of Pluronic F127 hydrogels of different polymer contents (22G nozzle,\nprinting speed v = 5–15 mm s-1, printing pressure P = 16–45 kPa). Top and planar view. Scale\nbar, 10 mm.\nb. How was the stability in Figure S6 evaluated? What causes the instability?\nrheology during photo-cross-linking, as observed in the decrease in the storage modulus upon\nirradiation (λ = 405nm; I = 8 mW cm-2). We suppose that the low concentration led to phase\nseparation and micelle rearrangement rather than network formation, rendering these\nformulations unsuitable for use in printed structures.\nc. The authors mentioned that “The choice of initial OD does not impact the cell growth\n(Figure S33)”. This statement should be moved to Results and Discussion to enhance clarity.\nWe thank the Reviewer for the suggestion and we have moved the statement to Results section\naccordingly. Previous Figure S33 is named Figure 13a in the revised manuscript.\nRevised text in Results (page 7):\n“The number of clusters and individual cells increased over time with cell concentrations\nreaching 5 x 109 cells mL -1 after 30 days (Figure 2d, S12). Further, the choice of initial\ngel\nOD , within the range of 0.3–1.6, did not impact cell growth (Figure S13b).”\n730nm\n2. The authors have adequately addressed the comment.\n3. Cyanobacteria encapsulation\na. The authors are encouraged to expand the time points for biomass quantification and cell\nviability measurements to include day 0 and day 5, in addition to the current intervals starting from\nday 10, due to swelling and dissolution of the un-crosslinked F127 during this period. The newly\nadded Figure S21 suggests that most changes predominantly occur within the day 1-10\ntimeframe.\nWe thank the Reviewer for the suggestion and we added viability measurements on day 0 and\nday 5 to Figure 2d.\nThe updated Figure 2d is appended below:\nin the encapsulated samples (n = 3), with an initial rapid growth rate up to day 15 followed by\na slower growth rate from day 15 to day 30 (green). Liquid suspension with the same starting\ncell concentration was used as a control of cell growth (black).\nUpdated text in the Methods section:\n“Cell viability and growth within the hydrogels were evaluated using confocal microscopy for\nday 1 to 5 and chlorophyll extraction for day 10 to 30 (Figure S33).”\nb. The same comment applies to the evaluation of cyanobacteria leakage.\nWe thank the Reviewer for the suggestion and we have also included data for day 1 and day 5\nto the leakage data (Figure S13b).\nFigure S13. b) Cell count in the printed disc sample quantified with chlorophyll extraction\n(green) and cell count in the surrounding culture medium (pink) measured via Coulter Counter\n(n = 3). The right axis indicates the percentage of cells that leaked into the surrounding culture\nmedium normalized to the total number of cells in the printed disc and the culture medium\ncombined (gray), as well as the encapsulation efficiency calculated as 100% – leakage %\n(brown).\n4. The authors have adequately addressed the comment.\n5. Dual carbon sequestration\na. In Figure 2E, the variability in the abiotic samples on day 1, day 5, and day 30 is very high.\nPlease provide an explanation. In addition, why is there a drop from day 0 to day 1?\nDiscussion of the results should be added to the main text.\nWe would like to clarify the data presented in Figure 2e. The left axis of Figure 2e (black)\nrepresents the difference in dry mass between 5 randomly selected biotic samples and 5\nrandomly selected abiotic samples. The variability on days 1 and 5 was high as the error bars\nreflected the cumulative error of these 5 randomly selected biotic samples and the 5 randomly\n16\nselected abiotic samples (Figure S-I, see Equivalent CO sequestration calculation in living\n2\nmaterials in the Supplementary Information, page 11). Due to the differences in the diffusion\nrate of unfunctionalized F127 polymer from the printed discs, there was sample to sample\nvariation in dry mass on days 1 and 5 (Figure S-I). This resulted in a relatively high error when\ncalculating the mass difference between the 5 biotic and 5 abiotic samples. We also attribute\nthe drop in mass from day 0 to day 1 to the diffusion of unfunctionalized F127 out of the samples.\nWe have updated the caption of Figure SX to explain these points.\nThe right axis of Figure 2e (right axis, red) represents the CaCO concentration quantified by\nmeasured the absorbance of 3 randomly selected biotic samples. The average absorbance of\n3 randomly selected abiotic samples (measured at each time point) was subtracted as\nbackground to obtain the final values. The error bars, therefore, reflected the cumulative\nvariability between the biotic and abiotic samples. We then calculated the corresponding CaCO\n3\nconcentration based on these absorbance values using the calibration curve (Figure S15 a).\nText in the original Results section (page 8):\n“To assess the amount of biomass and precipitate formation, the mass of abiotic and biotic\nsamples was measured and compared. Over the incubation period of 30 days, the biotic\nsamples had approximately 36% more dry mass than control abiotic samples (Figure 2e) with\nsignificant increase (p = 0.001) in biomass observed from day 10 of incubation.”\nRevised text in the Results section (page 7):\n“To assess the amount of biomass and precipitate formation, the mass of abiotic and biotic\nsamples was measured and compared (Figure 2e). The initial decrease in sample mass was\nattributed to the diffusion of unfunctionalized F127 polymer out of the printed discs, which also\ncaused high variability in the data of day 1 and day 5. Over the incubation period of 30 days,\nthe biotic samples had approximately 36% more dry mass than control abiotic samples with\nsignificant increase (p = 0.001) in dry mass observed from day 10 of incubation.”\n3\ndry mass of the biotic samples. The cumulative CO sequestration by biotic samples was\n2\nreflected by the difference in normalized dry mass between biotic and abiotic samples\n(black). Over the incubation period of 30 days, biotic samples accumulated 36% more\ndry mass than the abiotic samples. Alizarin Red S staining on the biotic samples indicated a\ntotal of 4.8 mg CaCO precipitation per gram of gel (red).\nincubation period of 30 days. The dry mass of each sample was normalized to the average\ninitial mass of day 0. The non-cross-linked F127 diffused out over time and caused higher\nsample to sample variation in dry mass on day 0 to 5.\nb. The authors stated that “It is important to note that the available medium volume was different\nfor our encapsulated samples and suspension. Since each disc sample was 40 μL with\nconcentrated cell clusters grown in 2 mL of medium, we observed the cell number within an\nencapsulated state is higher than that in the liquid positive control.” In order to compare the two\nexperimental groups, the experiment should be carried out at same concentration or volume,\notherwise there is an overestimation of the performance of the encapsulated samples.\nWe apologize for any confusion regarding the control experiment setup in this case. We indeed\nstarted the encapsulated samples and the liquid suspension control at the same cell\nconcentration per volume. Both the liquid suspension control and the encapsulated samples\nwere prepared at the same initial concentration (equivalent optical density of OD = 0.8) to\n730nm\nensure that the cells are in the same growth phase in both cases. For the encapsulated sample,\nan equivalent starting OD = 0.8 was reached within a gel volume of 40 μL, which we\n730nm\nconsidered as the primary cell growth volume. The encapsulated samples were immersed in 2\nmL of culture medium, but cell growth occurred in the gel volume of 40 μL and very limited cell\nleakage was observed (Figure S13b). The cells grew in immobilized clusters causing their high\nfinal concentration (Figure S12). For the suspension sample, the starting cell concentration was\nalso OD = 0.8 with the individual cyanobacteria cells dispersed in the culture medium\n730nm\nwithout forming clusters. This led to a lower final concentration compared with that observed in\nthe encapsulated state after the 30-day culture period. We would like to note that the\n18\ncomparative value between encapsulated and liquid suspension culture is not a conclusion of\nthe manuscript. We include these data to show that the cyanobacteria grow efficiently in both\nthe liquid suspension control and within our photosynthetic living materials, indicated that\nencapsulation does not inhibit their growth.\nFigure S12. Representative sample maximum intensity Z-projection images at different\ntime points (cell autofluorescence, green). Scale bar, 100 μm.\nFigure S13. b) Cell count in the printed disc sample quantified with chlorophyll extraction\n(green) and cell count in the surrounding culture medium (pink) measured via Coulter Counter\n(n = 3). The right axis indicates the percentage of cells that leaked into the surrounding culture\nmedium normalized to the total number of cells in the printed disc and the culture medium\ncombined (gray), as well as the encapsulation efficiency calculated as 100% – leakage %\n(brown).\nc. Where are the standard deviations in Figure 2D?\nWe have added the standard deviation accordingly.\nin the encapsulated samples (n = 3), with an initial rapid growth rate up to day 15 followed by\na slower growth rate from day 15 to day 30 (green). Liquid suspension with the same starting\ncell concentration was used as a control of cell growth (black).\n6. The authors have adequately addressed the comment.\n7. The authors have adequately addressed the comment.\n8. Volumetric lattice structure\na. Please provide the information on time points for the SEM cross-section. Which crosssection\nof the sample was imaged?\nThe samples were printed and incubated for 5 days before lyophilization and mounting for SEM\nimaging. To showcase the porosity on the sample surface, a sample (Figure S25a, b) was\ndissected along the vertical axis and flattened on the SEM holder. Another sample (Figure\nS25c) was dissected horizontally (transverse cross-section) to showcase the internal structure\nradially.\nincubated for 5 days. Scale bar, 1 cm. b) SEM image of the sample surface with lattice structure\nfalse colored in blue. Scale bar, 1 mm. c) SEM image of the transverse cross-section\nof the sample prepared by volumetric 3D printing. Scale bar, 1 mm.\nb. The authors stated that the material “exhibited robust viability up to 365 days” in Figure 4.\nHowever, cell viability was evaluated for up to 30 days only. Further evaluation at long-\nterm time points (days 60, 120, and 365) should be carried out to corroborate the authors’\nstatement.\nWe thank the Reviewer for this comment and as we did not quantify cell viability using\nchlorophyll extraction and confocal microscopy beyond day 30, we have softened these\nstatements in the manuscript. Fortunately, cyanobacteria, owing to their continual production of\nchlorophyll when metabolic active, provide a visual representation of their active state. We\nobserved a vibrant green color in the samples, indicative of continuous chlorophyll production\nthroughout the lifecycle reported in this manuscript. When samples die, they yellow, in our\nhands, within 48 h. We elaborate on this point below and have included additional images of\nour samples that were cultured long-term as well as representative images of samples that died,\nshowing yellowing.\nThe updated text in the revised manuscript now reads (Figure 4 caption, page 12):\n“[...] maintained a vibrant green color up to 365 days, indicative of continuous chlorophyll\nproduction.”\nWe collected images throughout the life cycle of the photosynthetic living materials cultured\nover 1 year that were presented in Figure 4b. The green shade in the optical images comes\nfrom the chlorophyll a content of the encapsulated cyanobacteria1. When chlorophyll a in\ncyanobacteria degrades upon or after cell death, it will be converted to phaeopigments, which\ngives a brownish-yellow color. Throughout the sample incubation time of one year, there was\nno yellowing observed, which would be indicative of cyanobacteria death and associated\nchlorophyll oxidation. We included additional images in the series in the Supporting Information\n(Figure S26a), appended below. In addition, we included an image of a typical unhealthy\nsample where the embedded cyanobacteria show the phaeopigment-like brownish-yellow color2\nwithin 48 h of cell death (Figure S26b).\nThe updated text in the revised manuscript now reads (page 11):\n“The 3D-printed photosynthetic living structure was cultured for over a year, during which it\ncontinuously produced chlorophyll and performed dual carbon sequestration (Figure S26).”\nIn addition, we updated the text in the original manuscript (page 11, page 13):\n“This prolonged viability of the photosynthetic living materials showed that dual carbon\nsequestration can take place beyond 30 days, especially within rationally designed 3D\nstructures.”\nAnd\n“Tailoring construct geometry via additive manufacturing enabled our photosynthetic living\nstructures to survive beyond one year while continuously undergoing dual carbon sinking.”\n“This prolonged culture of the photosynthetic living materials showed that dual carbon\nsequestration can take place beyond 30 days, especially within rationally designed 3D\nstructures.”\nAnd\n“Tailoring construct geometry via additive manufacturing enabled our photosynthetic living\nstructures to continuously perform dual carbon sequestration beyond one year.”\nFigure S26. a) Optical images of 3D printed photosynthetic living material at different time\npoints up to 365 days. Scale bar, 1 cm. b) 3D-printed photosynthetic living material yellowed\nwithin 48 h upon nutrient depletion, indicative of cyanobacteria death and chlorophyll a\noxidation. Scale bar, 1 cm.\nAs stated above, the samples reached a pseudo-steady state of cell number prior to day 30\n(Figure 2d). Thus, given the visual indication of continuous chlorophyll production throughout\nthe life cycle of the photosynthetic living materials, we do not see added value in further\nmeasures of individual cell viability. In order to be precise, we have therefore softened all\nlanguage related to ‘robust viability’ or ‘viability’ in the long-term samples and rephrase this as\ncontinuous chlorophyll production.\nReferences:\n1. Alvey, R. M., Biswas, A., Schluchter, W. M. & Bryant, D. A. Effects of modified Phycobilin\nbiosynthesis in the Cyanobacterium Synechococcus sp. Strain PCC 7002. J. Bacteriol.\n193, 1663–1671 (2011).\n22\n2. Sanmartín, P., Villa, F., Silva, B., Cappitelli, F. & Prieto, B. Color measurements as a\nreliable method for estimating chlorophyll degradation to phaeopigments. Biodegradation\n22, 763–771 (2011).\nFigure 2. d) An increase in cell concentration was observed throughout the 30-day incubation\nin the encapsulated samples (n = 3), with an initial rapid growth rate up to day 15 followed by\na slower growth rate from day 15 to day 30 (green). Liquid suspension with the same starting\ncell concentration was used as a control of cell growth (black).\n9. The authors reported cell viability up to day 30. The viability test should be evaluated at the\nlatest time points (days 60, 120, and 365).\nWe refer to the response to comment 8b from the same Reviewer directly above.\n10. The authors claimed that “Beyond 30 days, we expect the biomass accumulation to reach a\npseudo steady state (with similar cell death and growth rates) and the component of carbon\nsequestration related to biomass accumulation to equilibrate within the hydrogel” However, further\nexplanation is needed to clarify why this outcome is expected. Could the authors provide the\nrationale or reference supporting this expected outcome?\nWe drew this conclusion as we observed that cell number reached a plateau beyond 25 days\nwithin the encapsulated samples, as shown in Figure 2d. This suggests that the cell growth\nand cell death rates likely balanced out, leading to a plateau in biomass accumulation. Even\nthough the cell concentration reached a plateau, we still observed increased pH (Figure S11b),\nindicating that the cyanobacteria within the encapsulated samples remain metabolically active.\nin the encapsulated samples (n = 3), with an initial rapid growth rate up to day 15 followed by\na slower growth rate from day 15 to day 30 (green). Liquid suspension with the same starting\ncell concentration was used as a control of cell growth (black).\nFigure S1. b) pH changes of pooled biotic (green) and abiotic (black) medium every 5 days\n(n = 3).\n11. The authors mentioned that “The collected medium pooled from individual samples (n = 20)\nwas used for pH measurements to evaluate total sequestered CO as fully described in the\nthan taken from individual samples, which would allow reporting of the averages and standard\ndeviations. Additionally, while the authors stated that the experiment was performed twice, no\ninformation was provided on the averages or standard deviations for these repeated trials.\nmedium. Titration on 2 mL of culture medium is challenging and, in our hands, resulted in larger\nerror. Therefore, we pooled the medium from 20 samples together and titrated on the pooled\nmedium for more accurate and reliable results. The error bars have now been added to Figure\nS11. The DIC calculation also consisted of two sets of independent experiments, and each data\npoint was obtained by sampling the pooled medium from 20 individual samples. \nFigure S1. a) Total amount of dissolved inorganic carbon (DIC) under equilibrium in the re-\ncollected abiotic (black) and biotic (green) medium after every 5 day culture period (n = 2, with\npooling of 20 individual samples in each case). The gray dashed line indicates the amount of\nDIC in the form of Na CO that was used in simulated seawater medium (BG11–ASNIII). b)\n2 3\npH changes of pooled biotic (green) and abiotic (black) medium every 5 days (n = 3).\nMinor Comments:\n12. Page 8, Line 9 – There is a spelling error in “asses”. This should be corrected.\nWe thank the reviewer for pointing out the typo and it has been corrected accordingly.\n13. The reference to “Biomass quantification and cell viability characterization” (currently referred\nto Figure S25) appears to be incorrect. Please verify and correct this figure reference.\nWe thank the reviewer for pointing this out, the reference has been corrected.\n25"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have made improvements in the manuscript. However, the authors have still not addressed two critical comments.\nMajor comments:\n1. It’s still questionable whether atmospheric CO₂ is truly the primary carbon source for sequestration (lines 183-184). It’s not sure whether we can regard these materials as meaningful sequestration system if the authors take initial input [CO3 2-] into consideration as “cost” carbon sources.\nMaybe I miss something, but when I look at Figure S1a, I find that the amount of carbonate refreshed every 5 days is not an insignificant amount. For example, let’s consider the decrease in DIC from day 5 to day 10: Right after the medium refreshment at day 5, [DIC] should be similar to the one at day 0 (i.e. ~900umol/L with ~380umol/L coming from Na₂CO₃ in the medium). At day 10, [DIC] have reduced to ~600umol/L, corresponding to a reduction of 300umol/L used for biomass and carbonate precipitates. Why could these 300umol/L not come (in significant part) from Na₂CO₃ in the medium? And then from day 15 to 20, it goes from ~900umol/L to ~400umol/L, and why could these 500umol/L “used” not come in large part from Na₂CO₃ in the medium?\nThe authors should make two things clear in the manuscript or supplementary information: 1] How much CO3 of carbonate precipitates or biomass originally come from the atmospheric CO2, at least with clear simulation or calculations. The [CO3 2-] supplied every 5 days (refreshed) can be used for both clearly. 2] The contents of Figure S1a should include more details to make it clearer. It would help a lot if this graph also included the datapoints for [DIC] right after the refreshing of the medium.\nThese are critical points, which are essential to convincingly demonstrate the claims of “dual sequestration”. So, once this has been demonstrated, I recommend the data to be included as a main Figure or Table rather than in the S.I.\n2. Figure S30: The rate of CO2 change in a closed chamber is indeed a way to directly demonstrate CO2 sequestration. However, the uncertainty on this measurement is very high, with quite low R2. The addition of 2 controls is needed to validate this result: A control in dark conditions where CO2 should increase because of respiration of the cells (this data should already be available to the authors since the materials were monitored for 10 days, so I guess also while in dark condition). A control without cells (but in the same chamber and with an initial CO2 concentration higher than atmospheric) to check whether the sensor reports a constant amount of CO2 in such conditions, without any leakage, etc. It would be nice to show these controls and calibration data together. The authors can refer to the following paper for further information:\nBanerjee, S., Siemianowski, O., Liu, M., Lind, K. R., Tian, X., Nettleton, D., & Cademartiri, L. (2019). Stress response to CO2 deprivation by Arabidopsis thaliana in plant cultures. PLoS One, 14(3), e0212462.\nAlso, the authors should display the full data from their CO2 measurements including the 30min equilibration time and also the data after 1.4h, and explain clearly why the period between 0.5h-1.4h was specifically used to calculate the rate in change of [CO2].\nIf these are not possible, I would recommend to just remove figure S30 from the Supplementary data.\nMinor comment:\n1. In the revised manuscript (Line 173), the authors mentioned “uniform circular samples (Vsample = 40 μL, d = 10 mm)”. Is the volume (40uL) correct here?", "reviewer_2": "In this revised manuscript, the authors have satisfactorily addressed the comments from my previous review.", "reviewer_3": "The authors have satisfactorily addressed all of my comments."}, "author_response": "Response to Reviewers\nReviewer #1 (Remarks to the Author):\nThe authors have made improvements in the manuscript. However, the authors have still not\naddressed two critical comments.\nMajor comments:\n1. It’s still questionable whether atmospheric CO₂ is truly the primary carbon source for\nsequestration (lines 183-184). It’s not sure whether we can regard these materials as\nmeaningful sequestration system if the authors take initial input [CO3 2-] into consideration as\n“cost” carbon sources.\nMaybe I miss something, but when I look at Figure S1a, I find that the amount of carbonate\nrefreshed every 5 days is not an insignificant amount. For example, let’s consider the decrease\nin DIC from day 5 to day 10: Right after the medium refreshment at day 5, [DIC] should be\nsimilar to the one at day 0 (i.e. ~900umol/L with ~380umol/L coming from Na₂CO₃ in the\nmedium). At day 10, [DIC] have reduced to ~600umol/L, corresponding to a reduction of\n300umol/L used for biomass and carbonate precipitates. Why could these 300umol/L not\ncome (in significant part) from Na₂CO₃ in the medium? And then from day 15 to 20, it goes\nfrom ~900umol/L to ~400umol/L, and why could these 500umol/L “used” not come in large\npart from Na₂CO₃ in the medium?\nThe authors should make two things clear in the manuscript or supplementary information: 1]\nHow much CO3 of carbonate precipitates or biomass originally come from the atmospheric\nCO2, at least with clear simulation or calculations. The [CO3 2-] supplied every 5 days\n(refreshed) can be used for both clearly. 2] The contents of Figure S1a should include more\ndetails to make it clearer. It would help a lot if this graph also included the datapoints for [DIC]\nright after the refreshing of the medium.\nThese are critical points, which are essential to convincingly demonstrate the claims of “dual\nsequestration”. So, once this has been demonstrated, I recommend the data to be included\nas a main Figure or Table rather than in the S.I.\nTo ensure there is no confusion in the manuscript about the initial dissolved inorganic carbon\n(DIC) present in the medium, we have modified the statement of the primary source of\ncarbon in the manuscript accordingly.\nUpdated text in the revised version of the Results (page 7):\n“This increase in the DIC confirmed the dissolution of atmospheric CO , which together\n2\nwith dissolved Na CO in the simulated seawater medium, served as the carbon sources\n2 3\nfor CO sequestration.”\n“Our photosynthetic living materials function at ambient conditions with atmospheric CO\n2\nand simulated seawater as the carbon sources, [...]”\nUpdated text in the revised version of the Conclusion (page 14):\n“The photosynthetic living materials performed dual carbon sequestration over an\nextended life of up to one year with light as its energy source; atmospheric carbon and\nsimulated seawater as its carbon sources.”\nHowever, we would like to note that the precise source of inorganic carbon—whether\noriginating from the initial sodium carbonate or dissolved from the atmosphere to maintain\nthe carbon equilibrium in the medium—has no bearing on the claim of “dual carbon\nsequestration” in the manuscript.\nIn the prior revision, we included the DIC calculations for the biotic and abiotic samples\ntaking measurements at the end of each 5-day cycle. These data demonstrated that the DIC\nremained above the level given by the addition of Na CO at all times, independent of\n2 3\nmedium pH, temperature, and bacterial consumption of CO 2- ions, all of which affect the\namount of DIC in the fresh medium at the start of each 5 day cycle (Figure S1a). As the\nliving materials undergo carbon sequestration, they decrease the amount of DIC in the\nculture medium, yet atmospheric carbon dissolves in the culture medium to restore the\ncarbon equilibrium (which depends on temperature and pH). Notably, Figure S1c shows\nthat even without Na CO , the amount of DIC exceeded 381 μmol L-1, confirming that\n2 3\natmospheric carbon dissolved in the culture medium in all conditions.\nIn this context, even if the initial Na CO is considered a distinct source of inorganic carbon,\n2 3\nthe maximum percentage of inorganic carbon from Na CO that is used by the living\n2 3\nmaterials is 381μmol L-1/892 μmol L-1 x 100% = 42.7%. Therefore, atmospheric carbon\nremains the primary source of carbon for the living materials (> 50%). Nonetheless, we have\nnow explicitly acknowledged in the manuscript that the simulated seawater contributed to\nthe overall carbon pool, as stated above.\nWe would like to emphasize that the use of sodium carbonate (Na CO ) in the medium was\n2 3\nto simulate natural seawater conditions in a controlled lab environment. In natural seawater,\ncarbonate ions result from the dissolution and equilibration of atmospheric CO , where the\n2\nequilibrium is dictated by pH, temperature, and biological activity. We used Na CO to\n2 3\nreplicate this carbonate buffering system, which is a standard approach in the field. The\nphotosynthetic living materials then performed dual carbon sequestration in this simulated\nseawater environment by consuming dissolved inorganic carbon (DIC), which was in\ndynamic equilibrium with atmospheric CO .\namount introduced in the medium by Na CO , independent of pH and temperature. In an\n2 3\nopen system, if a CO 2- ion is consumed it will be replaced with atmospheric carbon at\n3\nequilibrium, regardless of whether the CO 2- ion came from Na CO or the atmosphere. This\n3 2 3\nwas also observed experimentally in closed-container experiments, which indicated a\ndecrease in the concentration of atmospheric CO during the day cycle (Figure S30 b,c,d)\n2\nas elaborated in response to point 2.\nFigure S1. a) Total amount of dissolved inorganic carbon (DIC) under equilibrium in the re-\ncollected abiotic (black) and biotic (green) medium after every 5-day culture period (n = 2,\nwith pooling of 20 individual samples in each case). The gray dashed line indicates the amount\nof DIC in the form of Na CO that was used in simulated seawater medium (BG11–ASNIII).\n2 3\nb) pH changes of pooled biotic (green) and abiotic (black) medium every 5 days (n = 3). c)\nDIC analysis of BG11–ASNIII medium without the addition of inorganic carbon through sodium\ncarbonate (Na CO ) at pH = 6.7 and pH = 8.9 (n = 2, black). The gray dashed line indicates\n2 3\nthe amount of DIC in the form of Na CO that was used in simulated seawater medium (BG11–\n2 3\nASNIII).\n2. Figure S30: The rate of CO2 change in a closed chamber is indeed a way to directly\ndemonstrate CO2 sequestration. However, the uncertainty on this measurement is very high,\nwith quite low R2. The addition of 2 controls is needed to validate this result: A control in dark\nconditions where CO2 should increase because of respiration of the cells (this data should\nalready be available to the authors since the materials were monitored for 10 days, so I guess\nalso while in dark condition). A control without cells (but in the same chamber and with an\ninitial CO2 concentration higher than atmospheric) to check whether the sensor reports a\nconstant amount of CO2 in such conditions, without any leakage, etc. It would be nice to show\nthese controls and calibration data together. The authors can refer to the following paper for\nfurther information:t\nBanerjee, S., Siemianowski, O., Liu, M., Lind, K. R., Tian, X., Nettleton, D., & Cademartiri, L.\n(2019). Stress response to CO2 deprivation by Arabidopsis thaliana in plant cultures. PLoS\nOne, 14(3), e0212462.\nAlso, the authors should display the full data from their CO2 measurements including the\n30min equilibration time and also the data after 1.4h, and explain clearly why the period\nbetween 0.5h-1.4h was specifically used to calculate the rate in change of [CO2].\nIf these are not possible, I would recommend to just remove figure S30 from the\nSupplementary data.\nWe thank the Reviewer for the careful look at these additional data. In order to build\nconfidence in our setup to monitor CO concentration within a closed chamber containing\n2\nour photosynthetic living materials, we had carried out the 2 controls requested and we have\nnow included these data in the Supporting Information.\nFirst, we loaded the chamber with an excess concentration of CO (~40’000 ppm) and\n2\nmonitored the concentration over time. The CO concentration remained constant\n2\nthroughout the measurement period, after an initial stabilization duration of ~0.5 h in two\nindependent experiments, suggesting no leakage from our system (Figure S30 e,f). Based\non these leakage tests, we also identified 0.5 h (30 min) as the initial equilibration time for\nall subsequent measurements.\nFigure S30. e, f CO leakage tests in 250 mL closed containers without cells loaded with an\n2\nexcess concentration of CO .\n2\nexperiment period and Figure S30 b,c,d show representative snapshots of the CO evolution\n2\nwithin the closed chambers. In dark conditions (night; areas shaded in grey), an increase (or\nplateau) in CO concentration was observed and this can be attributed mainly to cell respiration,\n2\nas indicated by the Reviewer. During the light cycles (day; areas shaded in green), CO\n2\nconcentration decreased within the closed chamber as expected for active photosynthesis.\nFigure S30. a Rate of change of CO concentration in a 250 mL closed container over a\n2\nmeasurement period of 10 days with a 0.25 mL sample and 10 mL culture medium (n = 2).\nCO sequestration was observed from day 3 and plateaued beyond day 5. b,c,d\n2\n3 day-night cycles. Black scatters: CO concentration; green shading: light (day) conditions;\n2\ngrey shading: dark (night) conditions.\nWe would like to clarify that the quantity of dual CO sequestration and conclusions reported\n2\nin the main text of the manuscript were not based on any closed-container measurements.\nThese experiments were included to corroborate that the systems consume atmospheric\ncarbon via photosynthesis during their life cycle and are therefore included in the Supporting\nInformation. We extracted rate information by interpolating stable sections of our data from\nthe CO sensor, as previously shown in Figure S30b. Such interpolations were taken over\n2\nmany sections of the data and we agree that it is better to show whole sections of the sensor\ndata, which are now present in the light(day)/dark(night) cycles presented in Figure S30\nb,c,d. We have kept the extracted rate information as it nicely shows that the system takes\nsome days to reach a steady state of CO consumption during the day cycles. Again, we\n2\nthank the Reviewer for this comment and agree that with the additional controls and\nexplanation, the data more clearly demonstrate that our systems convincingly consume\natmospheric CO during their life cycle.\n1. In the revised manuscript (Line 173), the authors mentioned “uniform circular samples\n(Vsample = 40 μL, d = 10 mm)”. Is the volume (40uL) correct here?\nYes, the volume of the printed samples is correct here, V = 40 μL = 40 mm3, d = 10\nsample\nmm, thickness l = 0.5 mm. The samples were printed using 22G conical nozzles (Ø = 0.41\nmm). We clarified the information of the final printed sample height in the revised manuscript."}, {"round": 3, "reviewer_comments": {"reviewer_1": "All my comments are properly addressed. I recommend publication."}, "author_response": ""}]

Decision letter
Title: Dual carbon sequestration with photosynthetic living materials
General Comments:
The manuscript describes 3D-printed hydrogel-based materials as novel tools for dual carbon sequestration.
Photosynthetic living materials were engineered by embedding cyanobacteria within a bioinert hydrogel
matrix, to achieve effective dual carbon sequestration through biomass accumulation and irreversible
microbially-induced calcium carbonate precipitation. The authors investigated the platform’s performance
not only in the short term (1 month) but also conducted long-term studies to demonstrate the platform’s
ability in forming biomass and producing mineral precipitation for over a year.
While the authors have made commendable efforts to address the reviewer’s questions, some
questions were not adequately addressed. The reviewer acknowledges these efforts but believes
that several questions need further clarification. Therefore, the manuscript is not recommended
for publication in its current state.
Technical Comments:
1. Hydrogel formulation
a. The authors have adequately addressed the printability of Pluronic F127 hydrogels of
different polymer contents. However, the images in Figure S5 are not consistent in lighting,
resulting in difficulty to access the resolution of prints. Higher-quality images with proper and
consistent lighting are recommended.
b. How was the stability in Figure S6 evaluated? What causes the instability?
c. The authors mentioned that “The choice of initial OD does not impact the cell growth
(Figure S33)”. This statement should be moved to Results and Discussion to enhance
clarity.
2. The authors have adequately addressed the comment.
3. Cyanobacteria encapsulation
a. The authors are encouraged to expand the time points for biomass quantification and cell
viability measurements to include day 0 and day 5, in addition to the current intervals
starting from day 10, due to swelling and dissolution of the un-crosslinked F127 during
this period. The newly added Figure S21 suggests that most changes predominantly occur
within the day 1-10 timeframe.
b. The same comment applies to the evaluation of cyanobacteria leakage.
4. The authors have adequately addressed the comment.
5. Dual carbon sequestration
a. In Figure 2E, the variability in the abiotic samples on day 1, day 5, and day 30 is very high.
Please provide an explanation. In addition, why is there a drop from day 0 to day 1?
Discussion of the results should be added to the main text.
b. The authors stated that “It is important to note that the available medium volume was
different for our encapsulated samples and suspension. Since each disc sample was 40 µL
with concentrated cell clusters grown in 2 mL of medium, we observed the cell number
within an encapsulated state is higher than that in the liquid positive control.” In order to
compare the two experimental groups, the experiment should be carried out at same
concentration or volume, otherwise there is an overestimation of the performance of the
encapsulated samples.
c. Where are the standard deviations in Figure 2D?
6. The authors have adequately addressed the comment.
7. The authors have adequately addressed the comment.
8. Volumetric lattice structure
a. Please provide the information on time points for the SEM cross-section. Which cross-
section of the sample was imaged?
b. The authors stated that the material “exhibited robust viability up to 365 days” in Figure 4.
However, cell viability was evaluated for up to 30 days only. Further evaluation at long-
term time points (days 60, 120, and 365) should be carried out to corroborate the authors’
statement.
9. The authors reported cell viability up to day 30. The viability test should be evaluated at the latest
time points (days 60, 120, and 365).
10. The authors claimed that “Beyond 30 days, we expect the biomass accumulation to reach a pseudo-
steady state (with similar cell death and growth rates) and the component of carbon sequestration
related to biomass accumulation to equilibrate within the hydrogel” However, further explanation
is needed to clarify why this outcome is expected. Could the authors provide the rationale or
reference supporting this expected outcome?
11. The authors mentioned that “The collected medium pooled from individual samples (n = 20) was
used for pH measurements to evaluate total sequestered CO as fully described in the
than taken from individual samples, which would allow reporting of the averages and standard
deviations. Additionally, while the authors stated that the experiment was performed twice, no
information was provided on the averages or standard deviations for these repeated trials.
Minor Comments:
12. Page 8, Line 9 – There is a spelling error in “asses”. This should be corrected.
13. The reference to “Biomass quantification and cell viability characterization” (currently referred to
Figure S25) appears to be incorrect. Please verify and correct this figure reference.