Engineered microbial hydrogels with confined architecture and binary microbes for efficient hydrogen production

Domain: biology
Published: 2026-03-23
Editorial outcome: major_revision

Abstract
Renewable energy sources, particularly hydrogen, offer a promising solution to address global
energy crisis and carbon emissions. Microalgae-driven hydrogen production has attracted
immense interest in both scientific and industrial fields. However, challenges such as high oxygen
sensitivity, substantial water demand, and low hydrogen production efficiency limit their potential.
Here, we develope a core–shell symbiotic hydrogel system for enhanced hydrogen production via
leveraging coaxial 3D bioprinting to spatially separate microalgae (i.e., core component) and
bacteria (i.e., shell component). These networks optimize light and nutrient utilization while
providing a localized anaerobic microenvironment to facilitate hydrogen production from
microalgal photosynthesis. The symbiotic systems enable a high hydrogen yield (1763 ± 98 mL L-
1). The system not only provides a highly efficient, liquid-free strategy for biohydrogen generation,
but also advances the understanding of symbiotic relationships and microorganism-material
interactions for creating advanced living material systems.

Introduction
The increasing global energy demand and environmental concerns have catalyzed the development
of sustainable and clean energy solutions1,2. Hydrogen is considered a key element in the future of
energy transition due to its high energy density, zero carbon emissions, and renewable
characteristics3. Traditional hydrogen production methods, such as steam reforming of fossil fuels,
are often accompanied by high energy consumption and greenhouse gas emissions. Water
electrolysis is a promising approach for obtaining clean and renewable energy, but it requires
relatively high energy consumption at present, raising potential concerns about the viability of
hydrogen as a clean and renewable energy source4,5. Biological hydrogen production has been
widely studied as a sustainable technology including photo-fermentation production, microbial
electrolysis production, and synthetic biology-enabled production6-8. In particular, microalgae-
driven hydrogen production via photosynthetic activity has attracted much attention, as it directly
utilizes the abundant and clean solar energy. The microalgal [FeFe]-hydrogenase uses electrons
obtained from the photosynthetic electron transport chain to reduce protons into hydrogen under
anaerobic conditions9, representing an ideal method for sustainable hydrogen production. However,
another intrinsic product of photosynthesis, oxygen, is a strong inhibitor of the activity and
expression of [FeFe]-hydrogenase10. Therefore, depleting oxygen and creating an anaerobic
environment are essential for microalgae hydrogen production.

Several strategies have been developed to overcome this challenge11-13, including genetic
engineering, cell-material interface engineering, microalgae immobilization, and co-culture
method. For example, the lba gene from soybean was introduced into the green microalga
Chlamydomonas reinhardtii, resulting in a significant increase of hydrogen production due to
enhanced respiration rates and altered starch metabolism14. However, this approach involves
complex transgenic biotechnology and may lead to potential ecological and biosafety concerns. In
contrast, engineering the microalgae-material interface (e.g., modulating microbial metabolic
pathways through the intervention of materials15) eliminates the gene risk (e.g., potential
biosecurity concerns caused by transgenic technology) and provides an effective strategy for
creating anaerobic microenvironments. While enzyme-regulated anaerobic encapsulation
technique was introduced to encapsulate single Chlorella cell by using laccase and polydopamine
for sustained H production16, the application of such enzymes and materials increases the cost and
2
limits the large-scale application. Additionally, immobilized microalgae maintained high
photosynthetic activity while effectively reducing oxygen diffusion, thereby enhancing
hydrogenase activity. For example, gel immobilization increased hydrogen yield of
Chlamydomonas reinhardtii by approximately twofold compared with suspended counterparts.

Nevertheless, conventional immobilized systems exhibit limited light penetration and low mass
transfer efficiency, which constrain efficient hydrogen production17. Co-culturing algal-bacterial
symbiosis is able to generate hydrogen, where the anaerobic conditions are maintained by oxygen-
scavenging bacteria18. Such methods typically exhibit the maximum accumulation of hydrogen
around 120 mL L⁻¹, primarily due to nutrient competition, toxic metabolite accumulation19, and
particularly reduced light transmission efficiency20 (limited light availability to microalgae), thus
reducing hydrogen production efficiency. Given these limitations, optimizing the symbiotic system
is crucial to improving hydrogen yield while maintaining its eco-friendly and cost-effective
advantages.

Here, we show a core–shell binary hydrogel network system containing microalgae and bacteria
for efficient hydrogen production, denoted as microbial symbiotic networks (MSNs), which
enhances and prolongs the activity of hydrogenases by creating a localized anaerobic
microenvironment to boost the hydrogen production capacity of microalgae (Fig. 1). Specifically,
we prepare hydrogel networks composed of sodium alginate (SA), carrageenan, and poly (ethylene
glycol) diacrylate (PEGDA), with Chlamydomonas reinhardtii (C. reinhardtii) and Bacillus
subtilis (B. subtilis) spatially compartmentalized in the core and shell parts, respectively, through
coaxial 3D printing technology (Supplementary Fig. 1). C. reinhardtii is a model species for
photosynthetic hydrogen production due to its high hydrogenase activity, fast growth rate, and low
culture cost21,22, while B. subtilis is able to consume oxygen by respiration20. The hydrogel nature
reduces the need of liquid medium and its core–shell structure mitigates nutrient competition.
Moreover, the customized hydrogel networks effectively distribute light, allowing the microalgae
within the scaffold to adequately receive light for enhanced hydrogen production. The MSNs not
only enhance the hydrogen production efficiency of microalgae in hydrogel, but also shed light on
the regulated symbiotic relationship in living material systems.

Results
Preparation of microbial symbiotic networks (MSNs)
SA and PEGDA were the main components used in the coaxial bioprinting technology for
preparing core–shell hydrogels due to their mechanical properties and biocompatibility23,24. SA
and carrageenan were mixed to prepare the bioink for the core part (i.e., microalgae region), while
they were further combined with PEGDA for the shell part (i.e., bacteria region) to improve the
mechanical properties and structural stability of the final hydrogel networks25,26. Specifically,
extrusion-based 3D printing requires materials with shear-thinning properties, and hence
carrageenan was incorporated into bioink to modulate its viscosity. The addition of an appropriate
amount of carrageenan improved the quality of the printed filaments (Supplementary Fig. 2).
Moreover, such composite bioink exhibited effective shear thinning behavior and good elastic
recovery, and the addition of C. reinhardtii or B. subtilis did not change the shear thinning behavior
(Supplementary Fig. 3). Introducing the corresponding culture medium (TAP medium for C.
reinhardtii and LB medium for B. subtilis) to each bioink enables the precise core–shell spatial
manipulation of C. reinhardtii and B. subtilis with suitable nutrient distribution for their mutual
growth. The crosslinked MSNs with stable architecture were eventually obtained after calcium
chloride treatment and UV curing process (Supplementary Fig. 4 and Supplementary Table 1).

The printing properties of core–shell hydrogels were investigated by adjusting different parameters,
including filament diameter and structural patterns. We demonstrated that core–shell hydrogels
can be printed in different sizes and heights, with C. reinhardtii (emitting red fluorescence due to
chlorophyll) and B. subtilis (emitting green fluorescence due to green fluorescent protein (GFP)
expression) uniformly distributed in the core and shell regions of the hydrogels (Fig. 2a and
Supplementary Fig. 5). In addition, the MSNs can be fabricated into structures of various shapes
with excellent flexibility (Fig. 2b,c). The obtained MSNs can withstand more than 50%
compressive strain and 30% tensile strain, and the compressive strength can reach 94 kPa
(Supplementary Fig. 6), showing excellent mechanical strength for potential large-scale
applications.

Flow cytometry revealed that C. reinhardtii and B. subtilis maintained high viability throughout
the processing steps, including extrusion, cross-linking, and photocuring (Fig. 2d and
Supplementary Fig. 7). In addition, glutathione S-transferase activity was monitored, as it is the
critical indicator to evaluate the level of microbial oxidative stress. No significant difference was
observed in glutathione S-transferase activity of C. reinhardtii and B. subtilis before and after
photocuring (Supplementary Fig. 8a). Furthermore, when the UV-treated cells were inoculated at
the same concentration as the pristine ones, no difference in the number of colonies was observed
(Supplementary Fig. A8b,c), further confirming that the negligible effect of printing and
photopolymerization on microbial activity. MSNs was placed in a sealed vial and an agar substrate
was provided at the bottom to maintain the humidity for the MSN culture (Supplementary Fig. 9).
The growth of C. reinhardtii and B. subtilis were monitored during 14-day culture, during which
additional liquid medium was not required to suspend the MSNs. Compared to the scanning
electron microscope (SEM) images and optical microscope of native C. reinhardtii and B. subtilis
(Fig. 2e and Supplementary Fig. 10), this culture method had no adverse effects on the morphology
of cells. Moreover, the MSN hydrogel constructs exhibited the increased chlorophyll content and
transitioned from a light green color on day 0 to a dark green after 14 days of culture (Fig. 2f,g),
confirming the growth of microalgae. The steady-state fluorescence spectrum further revealed that
the chlorophyll (at 680 nm) corresponding to C. reinhardtii continued to increase over a period of
two weeks, while the peak intensity of the green fluorescent protein (in the range of 400–550 nm)
significantly enhanced in the first three days and then gradually decreased to a stable level, which
might be due to nutrient deficiency (Fig. 2h and Supplementary Fig. 11). C. reinhardtii and B.
subtilis were largely confined in their designated compartments after 14-day culture with clear
boundaries for each microbial community (Fig. 2i). We ascribed this superior confinement to the
physical properties of the hydrogel (e.g., porosity and microenvironment of the hydrogel), which
restricted their movement and made C. reinhardtii and B. subtilis more inclined to colonize within
their inoculated regions. Additionally, the composition of the core and shell media, which were
tailored for C. reinhardtii and B. subtilis respectively, further facilitated their partitioned growth.
Collectively, we created an engineered binary living materials with complex structures, providing
a favorable environment for the growth and interaction of C. reinhardtii and B. subtilis.

Photosynthetic hydrogen production in MSNs
We next investigated whether B. subtilis could deplete oxygen through respiration to create an
anaerobic environment, which thereby activated hydrogenase to achieve hydrogen production.
Tungsten oxide (WO ), which can react with hydrogen to produce tungsten bronze and therefore
3
changes color from pale yellow to blue, was used for perceivable hydrogen detection27,28. In the
typical experimental setup, the MSNs were sealed in a glass container with WO powder. After a
3
48-hour illumination of the MSNs, the powder transformed from yellow-green to blue-gray color,
indicating the presence of hydrogen. Whereas, the color change was negligible in the absence of
light (Fig. 3a), indicating that light is essential for microalgae-driven hydrogen production. Oxygen
availability was typically constrained by low dissolved oxygen levels in traditional liquid cultures,
while MSNs under liquid-free conditions were exposed to sealed headspace air which contains
~20% oxygen. Therefore, the effect of oxygen in the headspace of container on hydrogen
production efficiency was investigated. Initially, the overall oxygen level decreased, as the bacteria
within the shell consumed oxygen through respiration. After the first cycle of hydrogen production
(72 hours), the overall oxygen generated by MSNs system was higher, leading the system to
gradually shift toward an aerobic state (Supplementary Fig. 12). We referred that the depletion of
acetic acid in the medium may reduce microalgal respiratory oxygen consumption, as acetate
serves as a primary substrate for mitochondrial respiration29. Consequently, reduced oxygen uptake
led to the accumulation of photosynthetically generated oxygen, driving the system from an
anaerobic to an aerobic state and thereby inhibiting the oxygen-sensitive hydrogenase. In this
scenario, the system exhibited a hydrogen production rate of 905 ± 47 mL H L-1 (Supplementary
2
Fig. 13), representing merely 51.3% of the purged system. Therefore, after introducing the MSNs
into the setup, nitrogen purging is recommended to remove oxygen in the headspace and establish
an anaerobic environment for initiating hydrogen production.

Hydrogenase activity governs the efficiency of electron transport and hydrogen production, while
it is highly sensitive to oxygen and rapidly deactivated even at low concentrations of oxygen17. To
maintain the hydrogenase activity, a sustained anaerobic environment is required. During this
process, B. subtilis were used to continuously create the atmosphere of low oxygen level by
consuming oxygen via respiration (Supplementary Fig.14). We varied the proportions of C.
reinhardtii and B. subtilis to investigate the optimal ratio for hydrogen production (Fig. 3b). The
C. reinhardtii-containing hydrogel system (i.e., shell bioink without bacteria) produced hydrogen
with a yield of 933 ± 64 mL H L-1, which is superior to most conventional suspension-based
2
microalgal (SM) systems (Supplementary Table 2). We ascribed this enhanced hydrogen
production to the improvement of cell density and light utilization30,31. No hydrogen was detected
in the monoculture system with B. subtilis alone for 48 h, while the addition of B. subtilis into C.
reinhardtii increased hydrogen production (Supplementary Fig. 14 and 15), as B. subtilis helped
maintain the oxygen concentration at a low level, which is essential for regulating the activity of
hydrogenase. For example, when the C. reinhardtii and B. subtilis were employed at a cell number
ratio of 1:15, the yield of hydrogen was 1763 ± 98 mL H L-1, approximately 3985 ± 327 μmol H
2 2
(mg Chl)-1, which increased by 89.0% compared to the C. reinhardtii-containing hydrogel. In
contrast, excessive use of B. subtilis (i.e., the ratio of C. reinhardtii to B. subtilis was 1:20) led to
their obvious diffusion into the algal region in the MSNs (Supplementary Fig. 16), where it likely
competed with C. reinhardtii for nutrients and interfered with light transmission20. Moreover, the
MSN with B. subtilis had a longer hydrogen production time than the group without B. subtilis
(Fig. 3b). We refer that B. subtilis consumed oxygen through respiration to maintain a low-oxygen
environment, thereby improving the hydrogenase activity and prolonging the time of hydrogen
production at relative high levels. To investigate whether MSNs with C. reinhardtii : B. subtilis
ratio of 1:15 can establish and maintain a local anaerobic environment, time-dependent dissolved
oxygen (DO) concentration of C. reinhardtii-containing hydrogels, B. subtilis-containing
hydrogels, and MSNs were monitored (Fig. 3c). The results showed that the C. reinhardtii-
containing hydrogels elevated DO levels through continuous production of photosynthetic oxygen.
In contrast, B. subtilis-containing hydrogels caused a rapid decline in DO due to the bacterial
respiration. In the MSN system, the oxygen generated by C. reinhardtii in the core was consumed
by the excessive B. subtilis in the shell, leading to a decrease in DO to a relatively low level. This
confirmed that MSNs efficiently created an anaerobic environment conducive to hydrogen
production.

Under light conditions, algal cells decompose water through the oxygen-evolving complex (OEC)
in Photosystems II (PSII) to produce electrons, which are the main source of algal photosynthesis
hydrogen32. Therefore, the activity of PSII can significantly affect the hydrogen production
efficiency of microalgae. Pulse-amplitude-modulation chlorophyll fluorescence imaging
(Imaging-PAM) was used to evaluate the PSII activity of C. reinhardtii under three different
culture conditions, including SM system, homogenous algal-bacterial hydrogel (Homo system),
and core–shell MSNs. The results showed that the PSII activity of C. reinhardtii under the three
culture conditions was similar at the beginning, indicating that the PSII activity of microalgae
would not be restricted in hydrogel. The PSII activity in Homo system slightly decreased after 4
days of cultivation, whereas that in MSNs system increased (Fig. 3d, e). These results suggested
that the microalgae of MSNs maintained higher photosynthetic activity over time, indicating that
the core–shell structure provided a favorable microenvironment for supporting efficient
photosynthesis.

The influence of core–shell MSN architecture (e.g., filament diameter, projected area, filling
interval, and construct height) on hydrogen production was evaluated (Supplementary Fig. 17).
Varying filament diameter (0.8, 1.0, and 1.2 mm) or projected area (300, 600, and 900 mm²) at a
constant height of 4 mm had minimal impact on hydrogen yield per unit volume (Supplementary
Fig. 17b, c). In contrast, MSNs with no filling intervals produced less hydrogen than those with
2.5 or 5.0 mm intervals (Supplementary Fig. 17d). This reduction is attributed to restricted gas
diffusion and high light attenuation, which suppress microalgal growth within the 1–2 mm thin
layer beneath the cube periphery33. However, height significantly affected production, with 4 and
8 mm constructs showing similar efficiency (1742 ± 47 and 1745 ± 9 mL L-1, respectively), while
the height of 12 mm reduced efficiency (1419 ± 91 mL L-1) probably due to limited light
penetration (Supplementary Fig. 17e). Moreover, F
v
/F
m
measurements after 48 hours confirmed
that higher heights (e.g., 12 mm) led to relatively lower value (Supplementary Fig. 18).

To assess the advantage of spatial compartment of the two microorganisms for hydrogen
production, we compared the core–shell MSNs to a Homo system. The core–shell MSNs exhibited
robust growth for 8 days, while the Homo system turned yellow on the fourth day, indicating the
decreased chlorophyll content for photosynthetic activity (Fig. 3f, g, and Supplementary Fig. 19).
It is noteworthy that C. reinhardtii-containing hydrogel (i.e., non-core–shell structure) also
exhibited robust growth (Supplementary Fig. 19), suggesting that the death of algae in the Homo
system was related to the bacteria. This highlights the advantage of MSNs system, which spatially
separates microalgae and bacteria, effectively avoiding the potential inhibitory effect of bacteria
on microalgae growth. On day 8, the chlorophyll content of MSNs reached 29.8 ± 1.8 mg L-1,
which was 2.2 times higher than that of Homo system (Fig. 3g). Moreover, the core–shell MSNs
produced higher amounts of hydrogen (Fig. 3h). This enhancement is attributed to
compartmentalizing C. reinhardtii and B. subtilis in distinct spatial domains, thereby providing
tailored microenvironments for each microorganism, which are conducive to maintaining the ideal
growth and metabolic state of each microorganism for hydrogen production34, 35. During the
cultivation process of MSNs and Homo systems, the metabolic process of B. subtilis significantly
reduced the mechanical properties of the hydrogel over time by microbial degradation of hydrogel
matrix (Supplementary Fig. 20)36.

Particularly, the MSNs can not only conserve water resources and improve space utilization, but
also outperform SM system in hydrogen production (Fig. 3i). TAP medium was used for hydrogen
production in the SM system, as C. reinhardtii grew well in the TAP medium whereas B. subtilis
underwent limited proliferation due to insufficient necessary nutrients (Supplementary Fig. 21).
When the initial cell numbers of C. reinhardtii and B. subtilis were consistent with those in MSNs,
the yield of hydrogen in SM is 22.4 ± 3.2 mL H₂ L-1. In contrast, the hydrogen yield of MSNs
achieved 1763 ± 98 mL H
2
L-1, 78 times higher than that of conventional SM system. MSNs were
able to create a more tailored microenvironment for C. reinhardtii and B. subtilis, which facilitated
the accumulation of microbial population and in turn promoted the efficient generation of
hydrogen. Several reports showed that in the symbiotic system of microalgae and bacteria, the
hydrogen production commonly ceased after reaching a certain level due to nutrient deficiency
and anaerobic environment disruption37,38. In this MSN system (the ratio of C. reinhardtii to B.
subtilis was 1:15), hydrogen production reached its maximum accumulation after 48 hours.
Therefore, the resulting hydrogen was collected, and nitrogen was introduced into the reaction
system to re-establish anaerobic conditions for another production cycle. The results indicated that
MSNs were capable of producing hydrogen in consecutive experiments for 8 days, though
hydrogen production showed obvious decline with increased cycle numbers (Fig. 3j). We speculate
that nutrients in the medium may be gradually depleted, limiting the growth and metabolic activity
of the binary microbial systems. The hydrogel system containing only microalgae also showed a
decreasing production trend in successive rounds, with hydrogen production in each cycle
significantly lower than that of the MSNs system containing B. subtilis (Supplementary Fig. 22).
These results indicated that the initial nitrogen purging cannot maintain a continuous anaerobic
environment and therefore additional B. subtilis was necessary to ensure the activity of microalgal
hydrogenase for enhanced hydrogen production.

Current microalgae-based photobiological hydrogen production mainly include the use of oxygen-
resistant strains39, adoption of pulse illumination40, depletion of generated oxygen from algae41-43,
co-culturing microalgae and bacteria19,44, promoting algae cell aggregation27, material-modified
algae45, immobilization system46, and employing genetic engineered strains with high hydrogen
production capabilities (Fig. 3k and Supplementary Table 2)47,48. In contrast, the spatial separation
strategy of C. reinhardtii and BC in MSNs significantly improves hydrogen production efficiency
and extends operational duration by optimizing nutrient and light utilization, while also creating a
localized anaerobic environment. Additionally, the spatial arrangement facilitates the
establishment of localized anaerobic microenvironments essential for efficient biohydrogen
synthesis. Consequently, this strategy enables the development of a sustainable biohydrogen
production platform while reducing environmental footprint.

Variation of photosynthetic system and hydrogenase gene expression
During the photosynthesis of microalgae, the electron transport process involves Photosystems II
(PSII) and I (PSI), where PSII and PSI work in tandem to drive the electron transport chain. PSII
initiates the process by capturing light energy and generating high-energy electrons, which are then
conveyed through the electron transport chain to plastoquinone (PQ), cytochrome b6/f complex
(Cyt b6/f), and plastocyanin (PC) (Fig. 4a)49. Following light-induced charge separation, PSI
donates its light-generated electron to ferredoxin (Fd) and accepts the electron from PC. Fd acts as
the final electron carrier in the photosynthetic linear electron transport chain and is responsible for
distributing electrons to different metabolic pathways. Under anaerobic conditions, Fd translocates
a portion of its electrons to [FeFe]-hydrogenase, wherein the enzymatic catalysis facilitates the
combination of these electrons with protons for the biogenesis of hydrogen50,51. The transfer of
electrons from the primary quinone receptor (QA) to the secondary quinone receptor (QB) of PSII
is one of the key steps in the whole process. The compound 3-(3, 4-dichlorophenyl)-1,1-
dimethylurea (DCMU) is a specific inhibitor of PSII that effectively blocks the electron transfer
process from the QA to QB52,53. When DCMU was added to the core ink, hydrogen production
showed an obvious decrease with the increasing of DCMU concentration (Fig. 4b). This confirms
the critical coupling of photosynthetic pathways and hydrogenase in MSN-driven hydrogen
generation.

We further explored the mechanism of improved hydrogen production of MSNs by transcriptome
analysis of C. reinhardtii in different coculture systems (i.e., the SM system or the Homo system
served as the control group, while the MSNs system was the experimental group). The repeatability
and correlation of each sample in transcriptome sequencing were reasonable, and the differential
gene clustering effect was significant, which proved the reliability of transcriptome analysis data
(Supplementary Fig. 23a). The expression patterns of C. reinhardtii in MSN system were
significantly different from those in SM system or Homo system, as well as the data distribution
and density of C. reinhardtii in MSNs (Supplementary Fig. 23b). Compared to the SM system,
9313 differentially expressed genes (DEGs) were identified, including 5056 upregulated and 4257
downregulated genes. While compared to Homo system, there were 2553 up-regulated genes and
1985 down-regulated (Fig. 4c and Supplementary Fig. 23c). The MSN system showed more gene
expression differences compared to the SM system (Supplementary Fig. 23d). Compared to SM
system, a certain number of DEGs in MSN systems were enriched in the pathways related to the
expression of metabolic pathways (i.e., pyruvate metabolism, citrate cycle, etc.) according to the
Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment (Fig. 4d). Moreover, the gene
ontology (GO) enrichment analysis revealed that the pathways related to photosynthesis were
significantly enriched (Fig. 4e and Supplementary Fig. 24). Compared to Homo system, some
pathways related to metabolism and photosynthesis were also enriched (Supplementary Fig. 25).
The microalgae photosynthesis pathway (e.g., photosynthesis, light reaction, light harvesting) was
enriched, which was directly related to the improved light utilization efficiency. Enrichment of
pathways related to nitrogen metabolism and ABC transporters indicated the enhanced metabolic
activity and efficient nutrient utilization in the microalgae17. Collectively, the improved hydrogen
production capacity of the MSNs system is likely linked to the activation of photosynthesis and
energy metabolism.

Gene expression of the photosynthetic pathways in MSNs was significantly upregulated (Fig. 4f).
In PS II, the expression of extrinsic proteins (psbO, psbP, psbQ, psbR) and structural proteins (psbY,
psb27) is stimulated compared to SM system, and their expression levels significantly affected the
efficiency of PSII. Similarly, the upregulation of gene encoding PSI led to an increasd PSI activity.
Moreover, the genes encoding Fd, cyt b6f complex, and ATPase in the MSNs system were
generally upregulated compared to SM system. The optimization of the electron transfer pathway
enhanced the energy conversion efficiency, thereby activating the expression of hydrogenase.
Furthermore, the expression levels of [Fe-Fe] hydrogenase and photosystem genes in different
system were compared by heat maps (Supplementary Fig. 26). The expression of most genes in
the SM system were lower than those in the other two systems, indicating that the liquid-free
cultivation environment had a positive effect on the photosynthesis and hydrogen production
efficiency of microalgae. Moreover, the expression of C. reinhardtii hydrogenase-related genes in
three systems were compared (Fig. 4g). HydEF and HydG are related genes encoding hydrogenase
maturation proteins. These two genes in MSNs systems were up-regulated by 1.79 and 2.10 times
compared to the SM system, and by 2.00 and 2.69 times compared to the Homo system. Compared
to the SM system and Homo system, the HydA1 gene encoding the catalytic subunit of hydrogenase
in MSN system was up-regulated by 1.16 and 3.95 times, respectively. In contrast, the low
expression of these genes in the Homo system highlighted that its microenvironment suppressed
hydrogenase transcription, confirming that the microenvironment of core–shell structure was more
conducive to the growth of C. reinhardtii for improved hydrogen production.

In summary, this study demonstrates that the binary microbial hydrogels featuring a confined core–
shell architecture represent a breakthrough approach for enhancing biohydrogen production. The
spatial separation strategy achieved through coaxial 3D bioprinting creates optimal photosynthetic
microenvironments for microalgae while establishing a localized anaerobic condition that
facilitates hydrogen generation via bacteria respiration. The MSN system not only achieves high
hydrogen production efficiencies but also reduces reliance on aqueous environments and
maximizes space utilization, highlighting its great potential for sustainable and efficient green
energy production. We expect that this innovative strategy provides a promising solution for
microalgae-based biomanufacturing, laying a solid foundation for the future advancement of
sustainable technologies and synthetic biology.

Methods
Chemicals and materials
All chemical reagents were used without further purification. Alginic acid sodium salt (viscosity
500–1000 mPa·s), carrageenan (E407), calcium chloride anhydrous (CaCl
2
), sodium citrate
dihydrate (≥ 99%), sodium chloride (NaCl), acetone, glutaraldehyde (25% in water), tungsten
trioxide (WO ), (3-(3,4-dichlorophenyl)-1,1-dimethylurea) (DCMU), lithium phenyl (2,4,6-
3
trimethylbenzoyl) phosphinate (LAP), Luria-Bertani (LB) medium, and poly (ethylene glycol)
diacrylate (PEGDA, Mw = 1000) were purchased from Shanghai Aladdin Biochemical
Technology CO., Ltd. Agar was purchased from Beijing Coolaibo Technology Co., Ltd.. Tris-
acetate-phosphate (TAP) medium was purchased from Institute of Hydrobiology, Chinese
Academy of Sciences. Agar was purchased from Bei-jing Coolaber Technology Co., Ltd.
Propidium iodide (PI) was purchased from Shanghai Aladdin Biochemical Technology CO., Ltd.
SYTOX Green Nucleic Acid Stain was purchased from Thermo Fisher Scientific (Waltham, MA,
USA). Glutathione S-transferase (GST) activity assay kit (BC0355) was purchased from Solarbio
Science & Technology Co., Ltd (Beijing, China). High-purity water with a resistivity of 18.2
MΩ·cm was obtained from a Millipore water purification system.

Characterization
The rheological behavior of hydrogels were characterized on an Anton Paar MCR302 rotational
rheometer. UV-vis absorption measurements were recorded with a PerkinElmer LAMBDA1050+
UV/Vis/NIR spectrophotometer. Microscopic and fluorescence images were obtained using a
Nikon ECLIPSE Si biomicroscope and an Olympus IX73 inverted fluorescence microscope,
respectively. Z-stack fluorescence images were collected with a Leica STELLARIS 5 CLSM. The
viability of microalgal cells was assessed using Agilent Technologies' NovoCyte Advanteon flow
cytometry and steady-state transient fluorescence spectrometer (Fluorolog-3). Scanning electron
microscopy was performed on a Phenom XL system (Thermo Fisher Scientific) at an accelerating
voltage of 10 kV. The Photosynthetic activity was assessed with a pulse amplitude modulation
chlorophyll fluorescence imaging (Imaging- PAM). Hydrogen was measured using a calibrated gas
chromatograph (GC 7900) with a column packed with molecular sieve 13X using nitrogen as
carrier gas.

Cells
Chlamydomonas reinhardtii (CC5325, denoted as CC) were kindly provided by Shenzhen Key
Laboratory of Marine Microbiome Engineering at Shenzhen University. C. reinhardtii was
cultured at 23 ℃ with agitation at 150 rpm and a continuous illumination of 50 µmol photons m-2
s-1 in a shaker incubator. Green fluorescent protein (GFP)-labeled Bacillus subtilis (Bacillus
subtilis 168-GFP) was obtained from Beijing Biobw Co., Ltd (China).

Preparation of MSNs
Due to the different bioink composition and nozzle parameters during coaxial printing, the
carrageenan concentration was investigated for the core bioink (3% SA) and the shell bioink (3%
SA + 10% PEGDA). For visualization and structural characterization, microalgae or bacteria were
separately incorporated into the bioinks, and their fluorescence signals were used to examine the
printed filament structures under a fluorescence microscope. Filament diameters were further
quantified by image analysis to assess the relationship between carrageenan concentration and
nozzle dimeter.

Based on the obtained formulations, 10 mL C. reinhardtii cells (1×107 cells mL-1) at exponential
growth stage were centrifuged (4000 g, 5 min) and concentrated into 75 µL (4000 g, 6 min), and
then suspended in 10 mL of 30 mg mL-1 sodium alginate (SA) solution, where TAP medium was
used to dissolve SA. Then, 450 mg carrageenan was added to the mixture to obtain core bioink for
the printing process.

Next, different amounts of B. subtilis (5 mL, 10 mL, 15 mL, 20 mL, 25 mL) in exponential growth
period (1×108 cells mL-1) were centrifuged and concentrated to 75 µL (8000 g, 6 min) and
suspended in 5 mL of 60 mg mL-1 SA solution, where LB medium was used to dissolve SA. The
cell number ratio of C. reinhardtii to B. subtilis in MSNs was 1:0, 1:5, 1:10, 1:15, 1:20, and 1:25,
respectively. The MSN with C. reinhardtii : B. subtilis=1:0 refers to a hydrogel system containing
only C. reinhardtii (i.e., shell bioink without bacteria). Then 5 mL of 20% PEGDA and 5 mg of
photoinitiator LAP were added. Then, 900 mg of carrageenan was added to the mixture to obtain
shell bioink for the printing process.

The core and shell bioinks were transferred into 5 mL and 10 mL syringes matched with the 3D
printer (Bio-architect® SR, Regenovo, Hangzhou, China), respectively. The core and shell bioinks
were distributed pneumatically through nozzles with pressure of 0.3 MPa and 0.4 MPa,
respectively. Customized 3D structures with different sizes and shapes were designed using printer
software, and the diameter of the material was adjusted by using different nozzles. Finally, MSNs
were cross-linked in 100 mM CaCl solution for 10 min and cured under UV light for 30 s (UV
2
light with a wavelength of 365 nm and a power density of 3 mW/cm²).

Preparation of Homo system
To prepare Homo system, 10 mL C. reinhardtii cells (1×107 cells mL-1) at exponential growth stage
were centrifuged and concentrated into 75 µL (4000 g, 5 min), and then suspended in 10 mL of 30
mg mL-1 sodium alginate (SA) solution, where TAP medium was used to dissolve SA. Next, 15
mL B. subtilis at exponential growth period (1×108 cells mL-1) were centrifuged and concentrated
to 75 µL (8000 g, 6 min) and suspended in 5 mL of 60 mg mL-1 SA solution, where LB medium
was used to dissolve SA. Then 5 mL of 20% PEGDA and 5 mg of photoinitiator LAP were added.
The two solutions were combined and thoroughly mixed. Finally, 1800 mg carrageenan was added
to the mixture to obtain the bioink for printing process. The bioinks were distributed pneumatically
through nozzles with pressure of 0.4 MPa. The Homo system was cross-linked in 100 mM CaCl
2
solution for 10 min and cured under UV light for 30 s.

Determination of the chlorophyll content
The method established in reference was used to quantitatively determine the chlorophyll content
of microalgae in MSNs53,54. Specifically, the freeze-dried MSNs were ground into powder and
soaked in acetone in a thermostatic mixer (600 rpm, 37 °C, Eppendorf ThermoMixer C) until the
cells were colorless. The mixture was then centrifuged (4000 g, 5 min) and the chlorophyll
concentration in the supernatant were determined by the absorbance values at 665 nm (A ) and
665
649 nm (A ). The total concentration of chlorophyll (C, mg L-1) was calculated according to the
649
equations below:
Chl a = 13.70×A − 5.76×A （1）
665 649
Chl b = 25.80×A − 7.60×A （2）
649 665
Chl (total) = 6.10×A + 20.04×A （3）
665 649

Photosynthetic performance measurements
The photosynthetic performance was measured using a pulse amplitude modulation chlorophyll
fluorescence imaging (Imaging-PAM). Specifically, a certain amount of C. reinhardtii and B.
subtilis was inoculated in liquid medium or hydrogel in the same volume, and the samples were
then placed in darkness for 30 min. The steady-state chlorophyll fluorescence (F ) and maximum
0
chlorophyll fluorescence (F ) were obtained by using a modulated light and a brief pulse of
m
saturating light, respectively. The parameter F /F indicated the maximum quantum efficiency of
v m
PSII. This factor was determined by calculating the ratio of (F – F ) to F .
m 0 m

Determination of cell viability by flow cytometry
To avoid the interference of fluorescence signals among chlorophyll, GFP, and dyes (e.g.,
propidium iodide and SYTOX Green), monoculture hydrogels containing microalgae or bacteria
were prepared to evaluate the viability of each component. The assay of dead cells for C.
reinhardtii and B. subtilis was conducted by incubating with 10 μM SYTOX Green and 5 μM
propidium iodide in the dark at room temperature for 15 minutes, respectively. The FITC channel
is used to detect microalgal dead cells labeled with SYTOX Green dye, while the PE channel is
used to detect bacterial dead cells labeled with PI. To avoid the interference of the green
fluorescence of B. subtilis-GFP with other dyes, we used the native B. subtilis in the analysis of
dead cells.

Qualitative analysis of hydrogen using tungsten oxide
Tungsten oxide (WO ) can react with hydrogen to produce tungsten bronze (2xH + WO ↔
3 ad 3
WO ·xH O) and therefore changes color from pale yellow to blue. It was used for perceivable
3−x 2
hydrogen detection. Initially, N was used to remove oxygen from the headspace of the vial
2
containing MSN and tungsten oxide (WO₃) powder. One group was incubated in dark, while the
other group was exposed to continuous illumination (50 µmol photon m-2 s-1) for 48 hours.

Preparation of SM system and measurement of H production
2
For hydrogen production in SM system, C. reinhardtii and B. subtilis at exponential growth stage
were suspended in a conical vial containing 10 mL TAP medium at a cell number ratio of 1:15 (C.
reinhardtii : B. subtilis). The initial cell numbers of C. reinhardtii and B. subtilis per milliliter of
TAP were consistent with those of MSN and the headspace volume of the hydrogen production
experiments was same. The headspace was purged with nitrogen for three minutes to remove
oxygen to accelerate hydrogen production and culture conditions (23 °C, 50 µmol photon m-2 s-1)
were the same as that of liquid-free culture. The amount of hydrogen produced is expressed in mL
L-1, and the liters in the unit refer to the volume of the TAP medium. The collection and detection
methods of hydrogen are consistent with those of the MSNs system.

Collection of C. reinhardtii and B. subtilis from hydrogel systems
For cells from the hydrogel for imaging and transcriptome sequencing experiments, MSNs were
dissolved with 0.9% NaCl solution containing 100 mM sodium citrate. The undissociated hydrogel
(shell part) was removed by gravity sedimentation and the remaining suspension was centrifuged
(4000 g, 5 min) to collect the C. reinhardtii. Subsequently, the supernatant was centrifuged again
(8000 g, 5 min) to collect the B. subtilis. Note: the hydrogel samples for flow cytometry assay was
prepared without adding photoinitiator (i.e., only calcium chloride cross-linking), and the
dissociation of the samples was performed in accordance with the steps described above.

Rheological performance testing
The rheological properties of bioink were tested using a MCR 302 rheometer (Anton Parr,
Germany), with a gap height of 1 mm at 25 ℃. The variation of shear stress with shear strain (the
strain range was from 0 to 100%) was tested by oscillatory amplitude sweeps. The fixed frequency
of the dynamic strain sweep experiment is 10 Hz, and the range of deformation is from 0.01% to
100%.

Mechanical performance testing
The tensile, compressive and cyclic compression tests of hydrogels were measured using a
universal tensile testing machine (Instron 5967, Instron Electronics Instrument Co., Ltd, USA).
For the tensile test, a dumbbell-shaped specimen with a filling interval of 0, a thickness of 4 mm,
a length of 30 mm, and a width of 10 mm was printed. For the compression test, a cylindrical
specimen with a filling interval of 0, a diameter of 14 mm, and a height of 20 mm was printed. The
tensile and compression speeds were both set at 10 mm min-1. Three specimens were prepared for
each mechanical property experiment.

Determination of glutathione S-transferase activity
To assess the potential effects of photopolymerization on microalgae and bacteria, three types of
MSN hydrogels were prepared (i.e., CC:BS = 1:0; CCP:BS = 0:1; CC:BS = 1:1). The experimental
group was cross-linked in 100 mM CaCl solution for 10 min and cured under UV light for 30 s
2
(UV light with a wavelength of 365 nm and a power density of 3 mW/cm²). The control group was
cross-linked in the same concentration of CaCl
2
solution for 10 minutes. Then, MSNs were
dissolved with 0.9% NaCl solution containing 100 mM sodium citrate and centrifuged (4000 g, 5
min) to collect the cells. Subsequently, glutathione S-transferase activity assay kit (BC0355,
Solarbio) was used to analyze the glutathione S-transferase activity according to the
manufacturer’s instructions.

Measurement of dissolved oxygen
The dissolved oxygen (DO) was measured according to previous method with modifications29.
Specifically, the MSNs (CC:BS=1:15), C. reinhardtii-containing hydrogels, and B. subtilis-
containing hydrogels were respectively incubated in the water, and the oxygen level was monitored
using a dissolved oxygen sensor (PyroScience OXF1100).

Culture of MSNs and measurement of H and O
2 2
In liquid-free culture for hydrogen production, MSNs was placed in a sealed bottle containing an
agar film at the bottom, where the agar film supplied water and maintained the humidity for the
MSN culture. The headspace was purged with nitrogen for three minutes to remove oxygen to
accelerate hydrogen production. All the experiments were purged after introducing the MSNs into
the setup or at the beginning of each cycle, except for the experiment investigating the time-
dependent changes of hydrogen and oxygen concentrations without nitrogen purging. The culture
environment was maintained at 38% relative humidity and 23 °C, and 50 µmol photon m-2 s-1 was
continuously illuminated. For hydrogen production in SM system, C. reinhardtii and B. subtilis at
logarithmic growth phase were suspended in a conical bottle containing TAP medium at a cell
number ratio of 1:15. The initial cell numbers of C. reinhardtii and B. subtilis were consistent with
MSNs. The headspace was purged with nitrogen for three minutes to remove oxygen to accelerate
hydrogen production and culture conditions (23 °C, 50 µmol photon m-2 s-1) are the same as that
of liquid-free culture. At different time intervals, 1 mL of the headspace was extracted with an
airtight syringe and injected into a calibrated gas chromatograph (GC 7900) with a column packed
with molecular sieve 13X using nitrogen as carrier gas. The concentrations of H and O in the
2 2
headspace were both determined by reported methods27,42. Specifically, the packed column was
held at 50 °C and the thermal conductivity detector (TCD) was set to 120 °C for H
2
analysis. The
volumes of hydrogen and oxygen gas were calculated using the external standard method. The
amount of hydrogen produced is expressed in mL L-1, and the liters in the unit refer to the volume
of the MSN. For hydrogen production at a cell number ratio of 1:15, the chlorophyll content was
measured as 20 ± 3 mg Chl L⁻¹. Therefore, the hydrogen yield of 1763 ± 98 mL H₂ L⁻¹ can be
converted to 3985 ± 327 μmol H₂ (mg Chl) ⁻¹.

The effect of DCMU on MSNs for hydrogen production
The MSNs with DCMU were prepared in the same protocol as the forementioned MSNs, except
that DCMU was dissolved in TAP medium at different concentrations (8 μM and 16 μM) when the
core ink was formulated.

Continuous H production
2
H production was analyzed in repeated batches. When no obvious increase of hydrogen was
2
observed, the reaction bottle was purged with nitrogen again to replace the gas to re-establish
oxygen-free conditions. At different time interval, 1 mL of headspace was extracted with an airtight
syringe and injected into a gas chromatograph to analyze the hydrogen content.

Transcriptome sample preparation
In order to ensure the rationality and validity of the experiment, three groups of samples with
different culture methods (i.e., conventional SM system, Homo system, and MSN system) were
designed. The control group was the C. reinhardtii in SM and Homo system, while the
experimental group was the C. reinhardtii in MNSN system. Three duplicate samples were set in
each group to enhance the reliability of the experimental results. Microalgae (i.e., C. reinhardtii)
under different culture methods were collected and washed with PBS to remove impurities. The
samples were then quickly transferred to a freezer tube and centrifuged (4000 g, 5 min) to remove
the supernatant. To maintain the activity and sequencing performance of the samples, the sample
was quickly frozen in liquid nitrogen for 15 minutes, after which the samples were transferred to
a refrigerator at -80 ℃ for long-term storage.

RNA sequencing and transcriptome analysis
Total RNA from microalgal samples was extracted using TRIzol reagent (Invitrogen, USA)
according to the manufacturer’s protocol with minor modifications to accommodate the
characteristics of microalgal cells. Briefly, fresh microalgal samples were thoroughly ground in
liquid nitrogen, and 1 mL of TRIzol reagent was added per 50–100 mg of tissue to ensure complete
cell lysis. The mixture was incubated at room temperature for 5 min, followed by the addition of
0.2 mL chloroform. After vigorous shaking, the samples were centrifuged at 13,000 g for 10 min
at 4 °C. The aqueous phase was carefully transferred to a new RNase-free tube, and total RNA
was precipitated with an equal volume of isopropanol, washed with 75% ethanol, air-dried, and
dissolved in RNase-free water. The concentration, purity, and integrity of RNA were assessed
using a NanoDrop spectrophotometer and agarose gel electrophoresis, and the qualified RNA
samples were subsequently used for transcriptome sequencing. The final cDNA library was
obtained by PCR enrichment. Enrichment analysis was performed using the clusterProfiler
package, which applied a hypergeometric test to calculate P values. The sequences were annotated
using the Kyoto Encyclopedia of Genes and Genomes (KEGG) database to identify functional
genes and perform metabolic pathway analysis.

Normalization of gene expression data and heatmap generation
The original FPKM values were Z-score normalized using the formula:
𝑧 = (𝑥−𝜇)/𝜎 （4）
where x is the original expression value, and μ and σ are the mean and standard deviation of that
gene across all samples, respectively.

Statistics and reproducibility
All studies were performed with three independent replications. All samples used in the study were
randomly allocated into different experimental groups. The average and standard deviation were
calculated in Prism9.5 software (GraphPad Software, USA). Data are presented as mean ± standard
deviation from at least three biological replicates.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript presents the development of a network of microbial hydrogels with core-shell architecture fabricated by 3D coaxial bioprinting, where Chlamydomonas reinhardtii (microalgae) is confined in the core and Bacillus subtilis (aerobic bacteria) in the shell. The central hypothesis is that this spatial configuration allows the maintenance of a local anaerobic microenvironment by bacterial oxygen consumption, thus enhancing hydrogen production by the microalgae through the photosynthetic pathway. The system achieves a H₂ production of up to 1763 mL L-¹, which is claimed to be the highest reported to date for systems of this type.\nThe manuscript uses a large variety of methodologies including functional assays, hydrogen and oxygen measurements, confocal microscopy imaging, 3D printing, physical characterizations of the hydrogels, transcriptomics.\nThe work has high originality and an innovative approach: The use of 3D coaxial bioprinting to create symbiotic systems with controlled architecture is technically advanced and represents a remarkable methodological innovation in the field of living materials. Moreover, it represents a significant advance in H₂ production: the yield achieved (up to 1763 mL H₂ L-¹) is far superior to most other reports on hydrogen production using Chlamydomonas, demonstrating the practical impact of the approach.\nUnfortunately, there are several drawbacks that severely affect the quality of the manuscript:\n• The introduction lacks background on H2 production using immobilized microalgae. Moreover, some relevant references describing high H2 production rates are absent.\n• References used throughout the manuscript are often inaccurate, misplaced, outdated or irrelevant. The use of references appears somewhat indiscriminate. Inappropriate references are often used. At other times, different references are used to support previously established knowledge that is repeated in different places in the manuscript, rather than reusing the same references. This gives the impression that references are included somewhat gratuitously, or at least with unsound inclusion criteria. A thorough review of the bibliography cited in the introduction is highly recommended.\n• The manuscript has an outstanding number of figures. However, the choice, quality and size, order, and distribution of the figures should be improved. One suggestion is to remove from the main manuscript all those figures that provide too little information for understanding the results, and to focus on those figures that provide relevant data. Some figure panels are difficult to visualize.\n• There are serious flaws in the experimental designs of some experiments.\n• The discussion of some results is poor.\n• Some conclusions are not supported by experimental data.\n• The wording of the text is repetitive and not very concise on the most relevant facts.\n• The information about the methodology used is scarce in some cases (e.g. H2 measurements, flow cytometry, RNA isolation).\nOverall, the manuscript has potential for publication in Nature Communications. However, it contains serious drawbacks that should be thoroughly addressed before publication can be reconsidered again.\nA more detailed review is as follows:\nIntroduction\nLines 41-42. The references used are neither the most suitable nor the most up to date.\nLines 42-44. The reference used could be improved or at least complemented with more current and specific ones to what is stated in this sentence.\nLines 44-47. The statements in this sentence are not very precise. Water electrolysis is a methodology that is highly compatible with obtaining clean and renewable energy. The authors should be more careful with this statement or at least qualify their point of view.\nLines 47-50. The idea of biohydrogen production has been in development for several decades. The term \"recently\" is debatable. Many of the biohydrogen production strategies are associated with CO2 co-production, so care should be taken when selling this strategy as \"environmentally friendly\". Finally, some of the references used can be updated by more modern ones.\nLines 56-58. Are the references corresponding to this sentence misplaced?\nLines 59-60. The reference linked to this sentence does not cover co-culturing approaches\nLines 60-63. This sentence has no reference\nLines 63-65. The reference of this sentence is likely misplaced.\nLine 65-67. Authors may consider explaining in the introduction the concept of “microalgae-material interface” and include references. Also, the term “gene risk” is not well defined.\nLines 72-76. Why is self-shading effect the most relevant drawback in co-culturing? Can the author support this statement?\nLines 87-89. References for Chlamydomonas reinhardtii as model organism for H2 production can be improved.\nLine 89. Reference number 28 is not well supported here.\nThe use of Chlamydomonas (or C. reinhardtii) and B. subtilis instead of CC and BS, respectively, is advisable throughout the manuscript.\nResults\nThe use of Flow Cytometry methodology to evaluate cell viability is poorly explained. (e.g. dyes to differentiate dead and viable cells, controls, percentage meanings, etc).\nLines 127-130. Measuring fluorescence derived from chlorophyll or GFP does not necessarily reflect the effect of UV on cell viability during photocuring, especially if photocuring last only 30 seconds.\nHas the mutagenic potential of photocuring been evaluated?\nSEM images of the cells before (Fig. S9) and after MSN (Fig. 2E) should be in the same figure for better comparison.\nLine 139. Type with “howed”\nWhat relevant information is providing Fig. S10 (3 days) relative to Fig. 2I (14 days)? Can Fig. S10 be omitted from the manuscript?\nLines 155-157. Should the composition of core and shell media (TAP vs. LB) be taken into account?\nLines 164-166. The addition of references originally describing or optimizing this methodology is advised.\nLines 167-171. The quality of the image of Fig. 3A is poor.\nFor the tungsten oxide experiments there is almost no methodological information.\nThis reviewer does not understand why data obtained with tungsten oxide (Fig. 3A) are more relevant than those obtained with gas chromatography (Fig. S11), which is a much more accurate technique. In fact, the hydrogen production depicted in Fig. S11 is not even discussed.\nLines 172-173. The initial decrease in O2 level, followed by its increase, is poorly discussed.\nLines 171-172. The cited references are not very suitable.\nThe methodology used to obtain the data depicted in Fig. S12 is poorly explained. Why are the O2 levels at time 0 at 0%? Were the cultures purged? If so, please identify clearly in the text which experiments were purged and which were not (e.g. Fig S11).\nOverall, the information about the methodology used for hydrogen production experiments is scarce. What is the volume of the bottles used to introduce the hydrogels? How is it possible to express H2 production as mL/L in an experimental design that has no liquid phase? What do the litres in these units refer to? To the volume of the bottles? To the volume of the hydrogels? How are the volumes of the hydrogels calculated? Do all hydrogels used in this manuscript have the same volume? For H2 production experiments in liquid media (without hydrogels). What volumes of liquid phase and headspace have been used?\nThis kind of information is absent in the figure captions or in the main text.\nMoreover, for all H2 production experiments, controls with only bacteria cultures are absent. Without these controls the contribution of the bacteria to H2 production is unknown.\nFor the H2 experiment using different algae-bacteria ratios (Figure 3B y 3C), how was the 1:0 condition performed? Were hydrogels made only with core bioink, or was shell bioink also added but without bacteria? The detail of this control is important to understand the results.\nWhat is the difference between Fig. 3B and 3C? Are the same data represented in different ways? Different representations of the same data must be avoided. Otherwise, the differences must be explained.\nTable S1. What do the colored markings mean? The choice of references included in this table seems capricious. Why are there references that show little H2 production capacity along with other works that show a high production capacity? Relevant references are absent, especially those showing higher H2 production or those using immobilized systems.\nLines 184-186. No data supports this statement. Moreover, the cited reference is not very suitable. Same for lines 190-192. Many other reasons could explain the observed data.\nLines 192-193. “….significantly prolonged after the addition of BS…”. Author may consider rephrase this sentence since BS was used at time 0 in all cases.\nFig. S13. This figure shows the influence of core–shell MSN architecture (e.g., projected area and construct 219 height) on hydrogen production. This experiment would have been useful and relevant if the cell concentrations or the total volume of the hydrogels had been kept constant. However, the same numbers of cells were used for different volumes of hydrogels, so much of the observed effect is likely due to a simple cell concentration effect. For example, the bacterial concentrations surrounding the algae can seriously affect O2 levels and H2 production. A clear example is in Fig. S13A, where the smallest MSN (300 mm2) produces the highest amount of H2. The authors conclude that some shapes impede light penetration and reduce H2 production. However, this conclusion needs to be contrasted by altering the experimental design and using a constant number of cells/mm2. Moreover, it will be convenient to express H2 using the same units throughout the manuscript.\nFor the experiments comparing growth between MSN and Homo systems, there is no discussion covering the fact that the algae die in the Homo systems after 4 days. Are there any controls using only the alga (no bacterium)? This would help to know whether or not the algal death is related to the presence of the bacteria. It would also be useful to know the viability of the bacteria in the homo systems.\nFinally, according to the Methods section, there are different characteristics between the MSN and Homo systems that make them difficult to compare. For example, the algal cell number for MSN systems and Homo systems are not comparable (10 ml of 1·107 cells mL-1 diluted in 10 ml of sodium alginate vs 50 ml OD=0.6 diluted in 5 ml of sodium alginate).\nFig 3E and S14A are very similar.\nLines 239-242. Why are two references cited for this statement? Please justify the use of these references.\nLines 242-245. Which data supports the idea that the alga and the bacterium are degrading the hydrogels? Why is there a reference cited at the end of this sentence? Please justify the use of this reference. The results indicated in these lines are not discussed.\nLines 252-253. Are there measures comparing algal and bacterial growth in liquid media vs. MSN systems?\nLines 253-256. The references cited do not cover algae-bacteria co-cultures\nLine 256-257. “In this MSN system, hydrogen production reached its maximum accumulation after 48 hours” Please, clearly indicate which MSN system is referred here.\nLine 269-272. The idea that nutrient and light utilization is optimized in the MSN system, which in turn favors H2 production, is not well supported by experimental data in this manuscript.\n272-274. Why does this approach minimizes environmental impact?\nFig. 4A can be spared\nLines 285-287. The reference 57 is not very suitable for this sentence.\nLines 277 to 289. All this information is more suitable for the introduction section rather than the result section. Consider moving all this information to the introduction section.\nMany of the figures and panels related to transcriptional analysis do not have sufficient information to be interpreted correctly. For example, it is unclear whether the data presented in these figures relates to liquid or hydrogel cultures. Which system is acting as the reference? More information is needed in order to interpret the figures correctly. There are also numerous abbreviations and color codes with no explanation of their meaning.\nThe idea of presenting clusters of genes that are overexpressed in Figure 4C, while clusters of genes that are downregulated are shown in Figure S16 is odd and does not make the manuscript any easier to read.\nLines 327-328. There is a typo in “stugy”\nThe experimental design of comparing transcriptomes between MSN hydrogels and liquid cultures, and between MSN and Homo systems is interesting and could shed light on the key pathways or genes that benefit hydrogen production. However, the study is very descriptive, and the discussion of the results is very basic; it does not provide any relevant information, except for the expected results (hydrogenases, maturation factors, ferredoxins, etc). It is a pity that the authors have not attempted to dive into the data to provide novel information or to discuss the data obtained in greater depth. A three way comparison (liquid vs MSN vs Homo) may help in identifying key genes related to the enhanced hydrogen production in MNS systems.", "reviewer_2": "Review is uploaded as an attachment", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_4": "The manuscript “Engineered binary microbial hydrogels with confined architecture for exceptional hydrogen production” presents a novel way to leverage co-axial 3D printing with compartmentalized microalgae and bacterial cultures for enhanced hydrogen production. Building upon the existing technologies that the co-culture of microalgae and bacteria species for photobiohydrogen production, the authors introduced a core-shell hydrogel system to improve the hydrogen production performance. The authors claimed that the core-shell 3D printing methods created an anaerobic environment that aids microalgae hydrogenase activity, which led to improved hydrogen production compared to conventional methods.\nIn the manuscript, the authors explained the preparation of microbial symbiotic networks and investigated the hydrogel production compared to non-core-shell structures as well as conventional suspension-based culture methods. The authors also investigated the change in gene expression of the microalgae within the symbiotic network to explain the increase in hydrogen production. The manuscript was well elaborated with convincing characterizations. However, it would benefit from further explanations of several core concepts and methods as well as improved control experiments to strengthen the central claims of the manuscript.\nMajor points:\n1. Regarding the claim on “anaerobic conditions”.\n1a. In the Introduction, the authors claimed that “Notably, co-culturing native algal-bacterial symbiosis is able to generate hydrogen, where the anaerobic conditions are maintained by oxygen-scavenging bacteria.” And therefore, they introduced bacillus subtilis to “creat[e] a localized anaerobic microenvironment”. As bacillus subtilis is a facultative anaerobe with limited oxygen consumption rates, the claim on “anaerobic microenvironment” might be an overstatement, especially when there is no direct evidence to show that there is indeed no oxygen present in the hydrogel in the manuscript.\n1b. On the same point, as the alginate-based hydrogel used in the manuscript is highly porous, oxygen diffusion may happen from the surrounding environment into the gel. Thus, the claim on “anaerobic conditions” created by bacillus subtilis may not be valid and if they would like to keep this claim needs to be demonstrated rigorously.\n1c. In the later part of the manuscript, the authors explained “[…] nitrogen was introduced into the reaction system to re- establish anaerobic conditions for another production cycle.” Does this mean that the oxygen-limited environment created by b. subtilis was only relevant for the first cycle (~48h?). Why is b. subtilis essential if the system can reach an oxygen-limited environment with nitrogen purging? The authors should include a control experiment without b. subtilis but with the nitrogen purging to strengthen the claim?\n1d. As a minor point, under the section “photosynthetic hydrogen production in MSNs”, the author also wrote “ […] BS was able to deplete oxygen […]”,the use of “deplete” needs to be changed if there is no direct evidence.\n2. For the material selection\n2a. The rationale behind the selection of the hydrogel materials was not fully elaborated. For example, carrageenan was used as a rheological modifier, but this point was not explained in the manuscript.\n2b. For the selection of the shell material, why was PEGDA used? What are the advantages that made the authors decide on using PEGDA and not other photo-cross-linkable hydrogels such as alginate methacrylate (which is similar to the base material of alginate)? Additionally, what was the molecular weight of PEGDA used in the formulation was how was this chosen?\n2c. In the method section, under the “Preparation of MSNs”, the authors used 450mg of carrageenan in the 10 mL of core material and 900 mg in the 10 mL of shell material. However, in the comparison with the “Homo system”, 900 mg of carrageenan was used per 10 mL of the hydrogel. The alginate concentration used in the core (30 mg/mL)-shell (60 mg/mL) was also different from that in the homo system (30 mg/mL).\nThese would make the core-shell system and homo system different in terms of hydrogel compositions and, therefore, the homo system is not the perfect control for comparing the hydrogen production rate. Is there a reason why the experiment was designed this way? Why was the concentration in the homo system not the average of the core-shell system?\n3. For the culture of algae and bacteria\n3a. The author used GFP-labelled bacillus subtilis to visualize the bacteria growth and distribution within the hydrogel. Was GFP chromosomally integrated? If not, were there any antibiotics used to keep the selection pressure? If GFP is carried on the plasmid and no antibiotics was used, wouldn’t the loss in plasmids affect the fluorescence readout (especially for Fig. 2H)?\n3b. The authors claimed that “The viability of CC and BS showed the negligible change before and after processing including after extrusion, crosslinking and photocuring, as evaluated by flow cytometry (Fig. 2D and fig. S7).” How was flow cytometry conducted when algae and bacteria were embedded in the photo-cross-linked hydrogel?\n3c. The authors stated that “The growth of CC and BS were monitored during 14 days of liquid-free culture, during which additional liquid medium was not required to suspend the MSNs, thus eliminating the liquid phase.” Could the authors explain the importance of having a liquid-free system? Why is it important to keep the sample liquid free when the final product (hydrogen) is not highly water-soluble? Wouldn’t the system benefit more from being incubated in liquid medium to 1. avoid nutrient depletion of b. subtilis and 2. limit oxygen diffusion to Chlamydomonas reinhardtii?\nAdditionally, it would help the readers to understand the system better by stating that the 3D prints were incubated with agar to maintain humidity in the main text. Without specifying this culture condition and without showing the supporting agar in any of the optical images, the readers will likely question how the hydrogel was not drying out throughout the duration of 14 days.\n3d. It is understandable that during the light cycle, C. reinhardtii produces hydrogen in the absence of oxygen. How does the system look like during the night cycles when C. reinhardtii undergoes respiration? Would the depletion in oxygen during the day cycle impact their viability during the night cycles (and thus harming the following hydrogen production)?\n3e. In Method section, “Determination of chlorophyll content”, how was the equation obtained? Was there any calibration curve for the chlorophyll quantification?\n3f. It is also helpful for the readers to know the type of chlorophyll present in C. reinhardtii, and how the absorbance wavelengths were chosen accordingly.\n3g. For the control experiment in SM (liquid) system, C. reinhardtii and b. subtilis were cultured only in TAP medium. As TAP medium is purely inorganic and is meant to support the growth of C. reinhardtii alone, b. subtilis is not expected to thrive in this condition, and thus this control simulated more a mono-culture condition for hydrogen production. A better alternative control would be with a mix of both media to ensure the viability of both species.\n4. Regarding the photosynthetic hydrogen production in MSNs,\n4a. It is a bit hard to see but was the experiment with tungsten oxide conducted with MSNs in liquid (Fig. 3A)? If so, why was this experiment conducted in a liquid state?\n4b. While it is important to 3D print the structures to have the core-shell compartmentalization of the species, is the grid structure necessary? As the hydrogen production is directly proportional to the amount of algae cells in the structure, and the authors noted that only the height of the structure had an impact on the hydrogen production, would printing a solid slab increase the hydrogen production per unit area of footprint (and thus better for any future mass production)?\n5. In section “Variation of Photosynthetic System and Hydrogenase Gene Expression”\n5a. In Fig. 4F, is the data presented as the ratio between MSNs and SM?\nMinor points:\n1. The use of “exceptional” in the title might be an overstatement and only relevant for the time of the manuscript. This wording should be changed and exceptional should be removed.\n2. The use of “binary” in the title may lead to confusion, and it does not elaborate the nature of the system very well. Maybe consider using another word.\n3. The author stated that “We ascribed this excellent hydrogen production to the improvement of cell density and optimization of light utilization.” The use of “excellent” is very subjective and “optimization” was not really illustrated through the experimental design. These phrasings should be changed and softened to make the writing more precise and aligned with what is shown in the manuscript.\n4. In the Introduction, the authors write “Such methods typically exhibit the maximum accumulation of hydrogen around 120 mL L⁻¹”. Given the algae-bacteria system is a dynamic biological system, it might be good to specify the hydrogen production in terms of production rate (for e.g. X mL/L/day)\n5. In Methods, under “preparation of MSNs” and “preparation of Homo system”, the cell concentration was reported in different ways (cells/mL vs. OD). The PEGDA concentration was also reported in either % or gram. It’s recommended to standardize these two.\n6. For Fig. S2A, the stress-strain curve. Stress is plotted on the x-axis and strain on the y axis. Was shear stress the independent variable?\n7. In Fig. S4 & S5, the duration of incubation was not specified.\n8. The author stated that “We ascribed this superior confinement to the physical properties of the hydrogel (e.g., porosity and microenvironment of the hydrogel), which restricted their movement and made CC and BS more inclined to colonize within their inoculated regions.” The use of “superior confinement” might be an overstatement especially in Fig. S10, mild mixing can be observed at the layer boundary and certain areas appeared to be yellow (which indicates mixing between the two species). Given the theoretical pore size of alginate networks to be much bigger than either Chlamydomonas reinhardtii or bacillus subtilis, and both species are motile, it is not very convincing to state that hydrogel network confined their movement. Also, this is not a central claim of the manuscript and toning the statement down would make the manuscript more scientifically sound.\n9. The hydrogen production rate reported in the manuscript was not consistent with the units. For example, in Fig. 3G & H, the unit was normalized to volume (mL/L), but in Fig. S13, it’s reported in micromole H2/mg Chl. Standardization across these plots can give the readers better comparisons.\n10. Certain experimental procedures are missing from the method section, such as the cell count with flow cytometry; UV cross-linking intensity and the calibration of the light; the hydrogen production testing with tungsten oxide; time dependent change of H2 and O2 concentration in the top space of a sealed bottle; rheological measurements on the hydrogel and the characterization on mechanical properties of MSNs.\n11. In the caption of Figure 2E, “Optical microscope and SEM images of CC and BS extracted from the MSN hydrogel constructs after 14 days of culture.” How were the algae and bacteria extracted from the hydrogel for imaging especially when the shell layer was photo-cross-linked? The extraction method should also be specified.\n12. In Fig. 3J, the unit is also missing from the x-axis."}, "author_response": ""}, {"round": 2, "reviewer_comments": {"reviewer_1": "The manuscript continues to provide a very interesting perspective on the potential to optimize H₂ production using microalgae. The alga–bacterium immobilization technology based on 3D printing is novel, and the H₂ production results (in absolute terms) are highly remarkable. In addition, the manuscript presents a valuable analysis of how to optimize 3D printing to achieve robust growth and production. The authors have made numerous changes that have improved some aspects of the previous manuscript. However, the essence of the criticisms raised in my first review still remains inadequately addressed. I regret that I cannot recommend this article for Nature Communications in its current form.\nThese were some of my initial criticisms:\nA. The introduction lacks background on H2 production using immobilized microalgae. Moreover, some relevant references describing high H2 production rates are absent.\nThe new version is still lacking background on this aspect, specifically in relation to Chlamydomonas.\nB. References used throughout the manuscript are often inaccurate, misplaced, outdated or irrelevant. The use of references appears somewhat indiscriminate. Inappropriate references are often used. At other times, different references are used to support previously established knowledge that is repeated in different places in the manuscript, rather than reusing the same references. This gives the impression that references are included somewhat gratuitously, or at least with unsound inclusion criteria. A thorough review of the bibliography cited in the introduction is highly recommended.\nMany references have been updated. However, the current version of the manuscript still contains many references which suitability could be questioned. In addition, the inclusion of references that describe the state of the art in a general and comprehensive manner is lacking.\nC. The manuscript has an outstanding number of figures. However, the choice, quality and size, order, and distribution of the figures should be improved. One suggestion is to remove from the main manuscript all those figures that provide too little information for understanding the results, and to focus on those figures that provide relevant data. Some figure panels are difficult to visualize.\nThe current version of the manuscript has not reduced the number of figures or panels within the figures; on the contrary, their number has increased (from 21 supplemental Figures to 26). Many of these figures or panels contain data of limited relevance, or even redundant information. Moreover, the size of many figures is so small that it is very difficult to clearly discern the results presented in them\nD. There are serious flaws in the experimental designs of some experiments.\nSome aspects have been corrected, such as the inclusion of a monoculture control of B. subtilis in the H₂ production experiments. However, the new data do not appear to be included in the manuscript, nor are they even mentioned in it.\nE. The discussion of some results is poor.\nF. Some conclusions are not supported by experimental data.\nG. The wording of the text is repetitive and not very concise on the most relevant facts.\nFurthermore, the coherence of the manuscript is affected by frequent digressions. For instance, the narrative on H₂ production is repeatedly interrupted by sections on hydrogel stability or photosynthetic activity, which hinders the overall flow of the text.\nH. The information about the methodology used is scarce in some cases (e.g. H2 measurements, flow cytometry, RNA isolation).\nSome new examples of aspects that should be improved are as follows:\n1. Lines 42-43: I am still not sure about the suitability of some references. Particularly, reference number 2.\n2. Lines 43-45. Reference number 4 focuses on hydrogen from biomass. This sentence should include more general references on the use of H₂ as an energy vector\n3. Lines 47-53. Similarly, most of the references (5–16) are too specific, addressing very concrete and discrete aspects of the different methodologies or topics mentioned. It would be advisable to substitute them with review articles that comprehensively cover the state of the art of the subjects referred to in these sentences.\n4. Line 51. I do not think the term 'facile' is appropriate.\n5. Relevant literature about Chlamydomonas immobilization/encapsulation for H2 production is missing.\n6. Lines 58-59. Again, references 18-19 are not the most suitable matches\n7. Line 79. Reference 28 does not cover algae-bacteria cocultivation.\n8. Lines 91-93. Again, references 29-31 are not the most suitable matches. References 29 and 31seems to be very specific references, selected rather randomly, whose only common point is that they use Chlamydomonas.\n9. Previous comment #13. The use of Chlamydomonas (or C. reinhardtii) and B. subtilis instead of CC and BS, respectively, is advisable throughout the manuscript.", "reviewer_2": "X. Li and co-authors have thoughtfully and thoroughly addressed my recommendations for the manuscript. Results from additional experiments support their findings that the binary living environment facilitates biohydrogen production by the decrease in oxygen present. This work advances creative microenvironmental approaches for solar fuels research. I support publication in Nature Communications.\nMinor comments/suggestions:\n1. Report H2 production in both micromole H2/mg Chl as well as the normalized mL/L. Many biohydrogen systems are reported as micromole H2/mg Chl. This could be done once (abstract) or in the paper once with the conversion factor explained. This will make their results more impactful to the wider scientific community.\n2.SI Table 2: \"CC\" is used two times. I do not think \"cellular component\" is used in the paper. Perhaps this could be deleted.\n3. SI Fig 20: please explain Mn and MW. Why are there 2 values listed in some cases?\n4. SI Fig 9: The abbreviations \"FITC and PE\" are not explained\n5. Fig 4A: both PSII and PSI are excited by light. Thus each should have a lightning bolt!\n6. line 357. \"Following light-induced charge separation, PSI donates its light-generated electron to ferredoxin (Fd) and accepts and electron from PC.\"\n7. In my opinion, I feel that the scientific findings reported in this paper are substantial without section 2.3. Perhaps think about this in your next papers.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_4": "The authors made substantial efforts to improve the manuscript “Engineered microbial hydrogels with confined architecture and binary microbes for efficient hydrogen production”, and the revised manuscript is more comprehensive than the original with clearer experimental designs and methods. However, in order to make the manuscript suitable for publication, the following points should still be addressed.\n1. a. What were the materials used for testing the impact of carrageenan concentration on filament diameter (Suppl. Fig. 2)? Was it carrageenan with alginate? And is this only applicable to the homogenous algal–bacterial hydrogel system? These information should be clarified in the figure caption. Additionally, the imaging method and the source of red color should also be indicated in the caption.\nb. For the selection of the carrageenan concentration, the Figure showed that 6 or 9 wt% of carrageenan had the best shape fidelity (Suppl. Fig. 2). And the authors stated in the response letter that “In contrast, 9% carrageenan resulted in poor extrusion performance during coaxial printing due to the long nozzle pathway for the core part. Therefore, a low carrageenan concentration (i.e., 4.5%) was used for the core material due to its relatively low viscosity.” It will help the reader to clarify the choice of different concentrations for different fabrication conditions as this is rather confusing in the current format.\n2. For the mechanical properties of the microbial symbiotic networks, the authors claimed that “The obtained MSNs can withstand more than 50% compressive strain and 30% tensile strain, and the compressive strength can reach 94 kPa (Supplementary Fig. 7)”. What were the microbial symbiotic networks compared with?\n3. The authors claimed that “Steady-state fluorescence further showed that the fluorescence of CC continued to increase over the two weeks, [...]”. While the growth of Chlamydomonas reinhardtii normally reaches the stagnation phase after 100 hours (~4 days). Was this due to the fact that the encapsulated C. reinhardtii had a longer doubling time? And why did the system reach the highest production on days 0–2 (Figure 3i and Figure RI.2) at the same time when the cell concentration was the lowest with C. reinhardtii?\n4. The authors compared the mechanical properties of the homogenous algal–bacterial hydrogel and microbial symbiotic networks and concluded that “compressive strength of MSNs was 1.7 times higher than that of the Homo system after 8 days of culture, indicating that the core–shell structure delayed the degradation of the hydrogel, demonstrating the robust structural integrity of core–shell hydrogel”. However, the final PEGDA concentrations in microbial symbiotic networks and homogenous algal–bacterial hydrogel systems were different. As the outer shell of the microbial symbiotic networks had a higher PEGDA and LAP concentration, it should naturally be a more robust system and the “delayed degradation of hydrogel” should be a result of higher PEGDA concentration and not the core-shell structure. This should be commented on in the manuscript text or a different comparison should be included with different architectures but the same PEGDA and LAP concentrations throughout the two constructs.\nMinor points:\n1. The correct use of “crosslinking” should be “cross-linking”.\n2. Figure RL2 shows the overview of hydrogen production over the 14 days of culture period, it might be helpful to replace Fig 3i with this.\n3. As a general comment, I agree with Reviewer #1 that reducing the use of non-standard acronyms (such as, SA for sodium alginate, BS for Bacillus subtilis, MSN for microbial symbiotic network, etc.) would enhance the readability of the manuscript. If re-formatting of the figures is truly not possible, the use of acronyms should at a minimum be reduced in the main text, and the acronyms used in the Figures can be defined explicitly in the caption."}, "author_response": "Thanks for your constructive suggestions. We understand that your suggestion aims to make the presentation more accessible. These two abbreviations (CC and BS) are frequently used in the figures. If they are replaced with “C. reinhardtii” and “B. subtilis”, it could cause some layout issues, which would affect the readability of the figures. Therefore, we would like to keep the abbreviated form “CC” and “BS”. Meanwhile, we have now added a table that includes all the abbreviations to facilitate readers' understanding (Table S2). The corresponding table has now been added as follows.\n2nd Reviewer response: As a suggestion, CC and BS may be kept in the figures if they are properly indicated in the figure legends, and CC and BS could be replaced by C. reinhardtii and B. subtilis, respectively, throughout the manuscript, as these are more conventional ways of referring to these microorganisms"}, {"round": 3, "reviewer_comments": {"reviewer_1": "Most of the issues have been properly addressed, and the manuscript has improved significantly.", "reviewer_2": "The authors have sufficiently addressed my recommendations for the paper. I fully support publication of this manuscript in Nature Communications.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts.", "reviewer_4": "The additionally revised version of the manuscript has improved substantially from the previous rounds. However, there remain a few points that should be addressed to support the central conclusions and improve its clarity.\n1. Regarding Point 4 in the previous round: although a Homo and a MSN system both have an average of 5 wt% PEGDA, the shell layer in the MSN itself contains 10 wt% PEGDA, as well as a higher LAP concentration. This local high concentration of polymer and photoinitiator results in a more highly cross-linked material. Therefore, the shell layer will have higher modulus and can retain its shape longer. Therefore, these observations were not purely the result of the core-shell structure but rather differences in the local PEGDA and LAP concentrations. While the core-shell structure had many other advantages in terms of hydrogen production, the increase in compressive strength can not also be attributed to the structure with the current data and conclusions in this direction should be removed.\n2. In order to claim the composite bioink to be shear-thinning, a shear rate ramp test is needed to see the change in viscosity with increasing shear rate. While it is likely that the material does exhibit shear-thinning, the current Suppl. Fig 3 (strain sweep) does not directly support this claim and similar curves can come from other material features (fracture, slippage, etc.). In addition, a suitable bioink should also demonstrate elastic recovery/self-healing properties, and a step strain measurement would support this property. These authors should refine the rheological characterization here to improve the characterization of the bioink and to be able to support the conclusions related to these points.\n3. The authors added “In contrast, excessive B. subtilis diffused into the algal region in the MSNs (Supplementary Fig. 16), where it likely competed with C. reinhardtii for nutrients and interfered with light transmission.” It was a bit confusing to understand the comparison here. What is this sentence in contrast with? Additionally, does B. subtilis really compete with C. reinhardtii for nutrients? This contradicts the claim later on (Line 325-326).\nMinor points\n1. As the field is still moving, the authors should avoid using the word “optimized”.\n2. Line 62: gene engineering should be genetic engineering\n3. Line 81: the use of “native algal-bacterial symbiosis pair” might be confusing. I understand the author meant “non- engineered species”, but it could also be understood as “naturally occurring symbiosis”, which is not the case.\n4. Line 100: The authors claimed “The hydrogel nature and core–shell structure of MSNs significantly eliminate the need for liquid medium and mitigate nutrient competition, improving both water and space utilization.” This one sentence is claiming two separate and non-interchangeable things. It would be better to separate them.\n5. Suppl. Fig 14: The figure caption was not written in complete sentences or correct grammar.\n6. Line 171: The sentence “The growth of C. reinhardtii and B. subtilis were monitored during 14 days of liquid-free culture, during which additional liquid medium was not required to suspend the MSNs, thus eliminating the liquid phase.” has many redundancies.\n7. Line 254: the use of “immediately consumed” is not supported by the data.\n8. Similarly, in line 255, the dissolved oxygen level was not stabilized but continued to decline according to the data shown in Figure 3c. More appropriate wordings should be used to describe the data.\n9. Line 298 and 299, extra quotation mark and line spacing.\n10. Line 304: the authors wrote “It is noteworthy that C. reinhardtii- containing hydrogel also exhibited robust growth [...].” This sentence needs clarification to indicate it is a hydrogel with only C. reinhardtii and not an MSN.\n11. Line 312, the abbreviation “BC” should be spelt out.\n12. Section numbers should be standardized."}, "author_response": ""}, {"round": 4, "reviewer_comments": {"reviewer_4": "The authors have adequately addressed the remaining comments and the manuscript is now suitable for publication."}, "author_response": ""}]

Decision letter
Peer Review File
Engineered microbial hydrogels with confined architecture
and binary microbes for efficient hydrogen production
Corresponding Author: Professor Jiajing Zhou
This file contains all reviewer reports in order by version, followed by all author rebuttals in order by version.
Attachments originally included by the reviewers as part of their assessment can be found at the end of this file.