Fully Reshapeable and Recyclable Protein Hydrogels

Abstract

Hydrogels have emerged as a new generation of functional materials with a broad range of applications in diverse fields. They usually cannot be reshaped or recycled, restricting their ability to respond to evolving demands and emerging applications, and contributing to the accumulation of synthetic polymer waste and associated environmental concerns. Here, a robust and general strategy is reported to engineer fully reshapeable and recyclable protein hydrogels by integrating protein folding‐unfolding with reversible disulfide crosslinking. Taking advantage of the substantial stiffness contrast between protein hydrogels in their folded and unfolded protein states, protein hydrogels are reversibly reshaped across one‐, two‐, and three‐dimensional geometries. Due to the denaturant concentration dependency and reversibility of the protein folding‐unfolding process, the resulting reshaping is highly tunable, reproducible, and chemically erasable, enabling consecutive shape transformations from a single hydrogel precursor. To impart full recyclability, irreversible crosslinking chemistry is replaced with a dynamic disulfide‐based one. The resulting hydrogels can be fully recycled, and the recovered proteins can be reused to form new hydrogels, allowing for complete recycling and repeated remolding of hydrogels into new shapes without compromising their mechanical properties. This approach establishes a robust platform for developing next‐generation protein‐based materials with dynamic formability and true material circularity.

By taking advantage of the substantial stiffness contrast between protein hydrogels in their folded and unfolded protein states as well as reversible disulfide crosslinking, a robust and general strategy is reported to engineer fully reshapeable and recyclable protein hydrogels. These protein hydrogels can be reversibly reshaped across one‐, two‐, and three‐dimensional geometries, and be fully recycled for material circularity.

1. Introduction

Hydrogels are hydrophilic polymeric networks capable of absorbing large amounts of water while maintaining structural integrity. Thanks to their versatile properties, such as high water content, soft nature, porosity, and tunable mechanical and physical characteristics, hydrogels have garnered significant attention across diverse fields, including drug delivery,^{[ 1} , ² , ^{3 ]} tissue engineering,^{[ 4} , ⁵ , ⁶ , ⁷ , ^{8 ]} cosmetics,^{[ 9 ]} environmental engineering,^{[ 10 ]} and soft robotics.^{[ 11} , ¹² , ^{13 ]} Conventionally, polymer hydrogels are fabricated using molds to attain specific shapes tailored to predefined applications. However, once manufactured, these hydrogels often exhibit limited adaptability, being able only to swell or contract without the capacity for substantial reshaping or recycling. Such inherent rigidity severely restricts the materials’ ability to respond to evolving demands and emerging applications. Additionally, the lack of recyclability significantly contributes to the accumulation of synthetic polymer waste and its associated environmental concerns.^{[ 14} , ¹⁵ , ^{16 ]} Therefore, developing hydrogels that can be dynamically reshaped and recycled for multiple uses is critical, not only for reducing environmental impact but also for transforming the traditional paradigm of hydrogel production and end‐use customization. Although several approaches, such as those based on supramolecular chemistry,^{[ 17} , ¹⁸ , ^{19 ]} have been explored to create recyclable hydrogels, efforts to develop hydrogels that are both recyclable and reshapeable remain limited. In this study, we report a novel strategy for engineering a protein hydrogel that seamlessly integrates both reshaping and recyclability into a single protein hydrogel system.

Reshaping is a common practice in industrial manufacturing techniques, such as die stamping and forging.^{[ 20} , ²¹ , ²² , ^{23 ]} These methods, which rely on the inherent or enhanced malleability of materials at elevated temperatures, enable the transformation of metal and polymeric materials into various geometric forms. Such reshaping capability substantially improves cost efficiency, sustainability, on‐demand modification, and material flexibility. While these processes are widely used in synthetic polymers, their direct application to hydrogel materials remains rare. Instead, hydrogels often rely on shape‐memory and stimuli‐responsive shape‐morphing technologies,^{[ 24} , ²⁵ , ²⁶ , ²⁷ , ²⁸ , ^{29 ]} which program a material's physical and mechanical properties to achieve predefined shape changes. Typically, these technologies involve programming temporary or intermediate shapes to facilitate efficient application processes, such as minimally invasive implantation^{[ 30} , ^{31 ]} or delayed deployment;^{[ 32 ]} whereas the permanent shape, which corresponds directly to the material's final functioning form, is predetermined during manufacturing.

To develop more adaptable and sustainable hydrogels, protein‐based hydrogels have attracted increasing interest due to their intrinsic biocompatibility, biodegradability, and tunable mechanical properties. Numerous strategies have been employed to achieve shape‐memory and shape‐morphing properties, including using thermal‐responsive proteins^{[ 33} , ³⁴ , ^{35 ]} or peptides^{[ 36 ]} to program shape recovery, leveraging the Hofmeister effect to induce conformational transitions in gelatin,^{[ 35} , ^{37 ]} and utilizing polyelectrolytes^{[ 38 ]} or metal ions^{[ 39 ]} to stiffen bovine serum albumin (BSA)‐based networks. In this work, we present a robust and general approach that enables protein hydrogels to be reshaped repeatedly and fully recycled for reuse. On the one hand, the reshaping capability allows the protein hydrogels to adapt to various application requirements even after manufacturing, offering enhanced flexibility in practical uses. On the other hand, full recyclability allows for the recovery of the fully functional proteins that can be reused for constructing new protein hydrogels. This strategy marks a significant step toward truly sustainable, efficient and versatile protein‐based biomaterial technologies.

2. Results and Discussion

2.1. Reshaping Protein Hydrogels via Protein Folding and Unfolding

To enable the reshaping of protein hydrogels, we propose to adapt the malleability‐based reshaping approach. Rather than relying on malleability, we instead exploit the substantial stiffness difference of protein hydrogels between folded and unfolded protein states as a means of reshaping. Our previous work demonstrated that the stiffness (Young's modulus) of protein hydrogels can be dynamically modulated by controlling the folding/unfolding state of the constituent protein blocks,^{[ 6} , ^{27 ]} with protein hydrogels in their folded state significantly stiffer than those in the unfolded state. This significant stiffness difference renders unfolded protein hydrogels pliable and easily deformable by physical manipulation, such as bending and rolling, whereas folded protein hydrogels are not. We hypothesized that if, after being physically deformed, the unfolded protein hydrogel is allowed to renature while being physically constrained in its deformed configuration, the folded protein hydrogel can be “locked” into the desired shape, as protein refolding stiffens the hydrogel network and thereby stabilizes the new configuration.

This shaping process would allow the protein hydrogels to be shaped into different forms from the same initial unfolded protein hydrogel. Moreover, the shaped folded protein hydrogels can then be reshaped by going through the unfolded protein hydrogel state again. Figure 1 schematically illustrates this reshaping strategy based on protein folding and unfolding: In step 1, the protein hydrogel prepared in the denatured state is manually deformed into the desired shape in the presence of chemical denaturant guanidine hydrochloride (GdmCl). In step 2, proteins are allowed to refold while the deformed hydrogel remains constrained, resulting in a stiffer and deswelled hydrogel that is reshaped into the desired new configuration. When necessary, the shaped hydrogel can be further reshaped by going through step 3, in which the hydrogel is brought back to the denatured state by incubating it in GdmCl. In so doing, the shape of the stiffer hydrogel is erased, allowing it to be subsequently reshaped into an entirely new configuration.

Figure 1.

Schematic illustration of the reshaping process for protein‐based hydrogels via protein unfolding and refolding dynamics. The starting hydrogel is crosslinked via dityrosine bonds in the denatured state. It is first deformed by mechanical forces into a desired shape. The following renaturation step in PBS enables the proteins to refold and stabilize the programmed configuration. In the last step, the proteins are unfolded again using the chemical denaturant GdmCl to restore the original shape of the hydrogel. This cyclic process allows protein hydrogels to be reshaped into multiple configurations as needed.

To test the feasibility of this proposed strategy, we used the (FL)₈ hydrogel as a model system. (FL)₈ consists of eight tandem repeats of FL, a de novo designed protein adopting the characteristic ferredoxin‐like α/β fold^{[ 40 ]} (Figure 2a), and has been used as a building block to construct protein hydrogels.^{[ 6} , ²⁷ , ^{41 ]} Here, we employed our recently developed denature crosslinking (DC) strategy to engineer (FL)₈ hydrogels.^{[ 6 ]} In the DC strategy, protein hydrogels are prepared from a concentrated solution of the unfolded protein, where the unfolded polypeptide chains are entangled, by using the well‐developed Ru²⁺‐mediated photochemical crosslinking method, which crosslinks two tyrosine residues that are in close proximity into a dityrosine adduct. The resultant denatured DC (FL)₈ hydrogel is referred to as the D‐DC hydrogel, in which chain entanglements are retained. The D‐DC (FL)₈ hydrogel is transparent and self‐standing, exhibiting a Young's modulus of ≈56 kPa in tensile tests (Figure 2b). Renaturing the (FL)₈ hydrogel in phosphate buffered saline (PBS) resulted in the renatured DC (FL)₈ hydrogel (named N‐DC), in which ≈50% of the FL domains refolded.^{[ 6 ]} Due to the refolding of FL domains and the entanglements of polypeptide chains, the N‐DC hydrogel significantly deswelled and became much stiffer, showing a swelling ratio (SR) of −72% and Young's modulus of ≈700 kPa. In addition, a large hysteresis is observed between stretching and relaxation curves of the N‐DC (FL)₈ hydrogel (Figure 2b), which is due to the force‐induced unfolding of FL domains at high strains. Moreover, the D‐DC and N‐DC hydrogels can be reversibly interconverted by repeated cycles of renaturation and denaturation of FL domains, leading to reversibly modulated physical and mechanical properties.

Figure 2.

a) Modeled structure of the FL protein generated using SWISS MODEL. b) Tensile stress–strain curves of 15% DC (FL)₈ hydrogel in native (red) and denatured (black) states. Inset: photographs of N‐DC (FL)₈ hydrogel (left) and D‐DC (FL)₈ hydrogel (right). The N‐DC hydrogel was translucent, significantly deswelled, and exhibited a high Young's modulus of ≈700 kPa. The D‐DC hydrogel showed an increased stiffness relative to a D‐NC hydrogel, with a Young's modulus of ≈50 kPa. The Young's modulus of the hydrogel was calculated as the slope of the stress–strain curve at 15% strain. c) A scheme illustrating the reshaping of a 15% D‐DC (FL)₈ hydrogel strip into a bent configuration and its restoration to the original form through protein unfolding in GdmCl. Scale bar: 5 mm. d) Photos demonstrating that a straight D‐DC (FL)₈ hydrogel strip could be reshaped into different 1D shapes via the protein unfolding and refolding strategy. Scale bar: 5 mm.

The D‐DC (FL)₈ hydrogel is relatively soft and flexible, making it possible to deform the hydrogel into different shapes by physical means. For example, a strip of 15% D‐DC (FL)₈ hydrogel can be easily bent by 180° without fracturing (Figure 2c). We therefore used this D‐DC (FL)₈ hydrogel as the starting form to evaluate the feasibility of our reshaping strategy. After bending by 180°, the D‐DC (FL)₈ hydrogel was incubated in PBS while under constraints to allow the FL domains to refold. After renaturation and removal of the physical constraints, the N‐DC hydrogel strip retained its bent shape, i.e., the straight D‐DC hydrogel strip was successfully reshaped into a stable bent N‐DC form. Moreover, once shaped, the N‐DC hydrogel can be reshaped by bringing the hydrogel back to the softer D‐DC form. As shown in Figure 2c, after the bent N‐DC hydrogel was incubated in 7 m GdmCl, the resulting D‐DC hydrogel restored back to being largely straight, closely matching the initial starting shape. From the initial state, the D‐DC hydrogel strip can be reshaped into different 1D configurations of N‐DC hydrogels (Figure 2d). This result demonstrates the feasibility of our stiffness‐based reshaping strategy. In a sense, this reshaping process is analogous to reshaping a steel rod at high temperatures.

It is of note that the reshaping and restoration depend on the folding and unfolding of proteins in the hydrogel and are relatively slow. The kinetics of the reshaping process depend on the time required for GdmCl to completely diffuse out of the hydrogel, and typically, 3 h is sufficient to achieve stable reshaping (i.e., shape fixation). In contrast, shape restoration in 7 M GdmCl is much faster. The hydrogel begins to lose its fixed shape almost immediately upon immersion, and typically restores its original shape within 20 min (Figure S1, Supporting Information).

Technically, the reshaping strategy we demonstrated here is similar to the conventional shape‐memory processes, where the reshaped configuration of the N‐DC hydrogel can be viewed as a programmed temporary state, while the D‐DC hydrogel represents the permanent shape. However, one major distinction is that the permanent shape in traditional shape‐memory/shape‐morphing typically serves as the intended functional configuration, whereas in our approach, the “temporary” shape is the intended form for applications. In a nutshell, our reshaping strategy can be regarded as a versatile manufacturing method to produce protein hydrogels with diverse functional shapes starting from a single initial hydrogel precursor.

2.2. Reshaping Performance can be Tuned by Varying GdmCl Concentrations

To characterize the reshaping efficacy of DC‐(FL)₈ hydrogels, we measured the fixity ratio (R_f), which is commonly used in characterizing shape‐memory materials, during the bending reshaping process (Figure S2, Supporting Information). A 15% D‐DC (FL)₈ hydrogel strip was bent by 180° and subsequently “fixed” in PBS buffer to convert the hydrogel to the N‐DC state. The significant stiffening and deswelling of the hydrogel network following FL refolding effectively locked the imposed deformation, yielding an R_f of 80% ± 5% (n = 9), demonstrating the high efficacy of the reshaping strategy. To assess the efficacy of “erasing” the shape of the N‐DC (FL)₈ hydrogel, the bent N‐DC hydrogel strip was brought back to 7 m GdmCl. The bent configuration was largely lost in the resultant D‐DC hydrogel, achieving a high recovery ratio (R_r) of 88% ± 7% (n = 9). This result supported our hypothesis that the significant stiffness difference between the N‐DC and D‐DC hydrogels allows for reshaping while avoiding permanent deformation. Moreover, when subjected to consecutive bending reshaping cycles, the DC (FL)₈ hydrogel demonstrated excellent reproducible reshaping performance for at least 10 cycles (Figure 3a). This notable reproducibility arises from the reversible nature of FL folding and unfolding, which underscores our reshaping strategy. In addition, during these repeated bending reshaping‐restoration cycles, no visible signs of cracking, tearing, or permanent deformation were observed in the hydrogel sample for up to 20 cycles, indicative of low fatigue in the hydrogel. Since this study is proof‐of‐concept in nature, we did not attempt to determine the maximum number of reshaping cycles the hydrogel can withstand before significant fatigue occurs. However, material fatigue is a critical factor in reshaping processes. Therefore, it is essential to determine the maximum number of reshaping cycles in applications involving repeated deformation.

Figure 3.

Reshaping can be modulated by denaturant concentration. a) R_f (gray) and R_r (red) values of the 15% DC (FL)₈ hydrogel over 10 consecutive programming cycles. The hydrogel was alternated between bent and straight configurations, with both R_f and R_r demonstrating excellent reproducibility across the tested cycles. b,c) The Young's modulus (b) and swelling ratio (c) of 15% DC (FL)₈ hydrogels depend on [GdmCl]. Error bars indicate standard deviation (n = 6). d) R_f values for DC (FL)₈ hydrogels reshaped at different GdmCl concentrations when using hydrogels in 7 m GdmCl as the starting material. Error bars indicate standard deviation (n = 3). R_f decreases with increasing [GdmCl] and reaches a plateau at 2 m and above. For comparison, the chemical denaturation curve of FL is included in panels b‐d).

The reshaping of the (FL)₈ hydrogel is driven by the folding and unfolding of FL domains and the resultant difference in the hydrogel's stiffness. Since protein folding/unfolding is dependent upon denaturant concentration, the stiffness of the hydrogels should depend on the denaturant concentration, which should in turn affect the reshaping efficacy. To examine this effect, we first determined the dependence of the (FL)₈ hydrogel's stiffness and swelling ratio on the GdmCl concentration ([GdmCl]). As shown in Figure 3b,c, the stiffness of DC‐(FL)₈ hydrogel decreased while the swelling ratio increased with increasing [GdmCl]. Both dependencies largely mirror the chemical denaturation curve of FL. We then determined the reshaping efficacy of DC‐(FL)₈ hydrogels (R_f), by using the D‐DC hydrogel (7 m GdmCl) as the starting form and fixing the hydrogel at different [GdmCl]. As shown in Figure 3d, as [GdmCl] increased from 0 to 2.0 m, R_f decreased from ≈84% to only ≈22%, which then plateaued at higher denaturant concentrations. This trend again largely mirrors the chemical denaturation curve of (FL)₈. This result indicates that the reshaping is effective when the fixing is done in solutions with [GdmCl] lower than 1.5 m, suggesting that a lower [GdmCl] (≤1.5 m) is necessary in order to reshape DC‐(FL)₈ hydrogel effectively when D‐DC‐(FL)₈ hydrogel in 7 m GdmCl is used as the starting material.

2.3. Reshaping Can Be Achieved in 2D and 3D Configurations

The reshaping of DC (FL)₈ hydrogels is not limited to 1D samples but also extends to 2D and 3D configurations. Figure 4a illustrates an example of 2D reshaping of DC (FL)₈ hydrogels through iterative reshaping cycles. In this experiment, a ring‐shaped hydrogel, which was first prepared in 7 m GdmCl, was reshaped into various multigon configurations consecutively starting from the same D‐DC hydrogel ring. Initially, a 15% ring‐shaped D‐DC (FL)₈ hydrogel was fabricated and then deformed into a triangular configuration by stretching. After renaturation and subsequent removal of the applied force, the N‐DC (FL)₈ hydrogel stably retained the triangular configuration. After incubating in 7 m GdmCl, the triangular configuration was erased, and the reinstated D‐DC (FL)₈ hydrogel ring was then reshaped into different configurations (rectangular and pentagonal). The reproducible fixity performance enabled by protein refolding allowed the N‐DC (FL)₈ hydrogel to maintain the new configuration effectively. These sequential reshaping cycles, transitioning between distinct geometrical configurations, illustrate a high degree of reshaping capability and the remarkable adaptability of our approach based on protein folding and unfolding.

Figure 4.

a) Photographs showing the reconfiguration of a ring‐shaped 15% DC (FL)₈ hydrogel into a series of 2D geometries. Scale bar: 5 mm. The hydrogel was sequentially transformed into a triangle, rectangle, and pentagon. The original ring shape was effectively recovered using GdmCl between each reshaping cycle. b) Photographs depicting the reshaping of a flat 15% DC (FL)₈ hydrogel sheet into various 3D configurations. Double‐headed arrows indicate the reversible deformation achieved through the protein folding and unfolding mechanism. Scale bar: 5 mm. c) Photographs illustrating that a straight 15% DC (FL)₈ tubing gel could be reconfigured into opposing 3D helical configurations.

Building upon the success of 2D reshaping, we continued to explore our strategy for 3D reshaping. Owing to the inherent high stiffness of the N‐DC (FL)₈ hydrogel, which is sufficient to sustain its structural form under its own weight, we were able to reshape a flat DC (FL)₈ hydrogel sheet into intricate 3D configurations. Figure 4b shows that a flat DC (FL)₈ hydrogel sheet was sequentially reshaped into a tubular form, a wave pattern, and finally a bowl conforming to the contours of a spherical mold. Additionally, a straight tubing‐shaped hydrogel was reshaped into a right‐handed helix (Figure 4c), as well as a left‐handed helix, demonstrating the capacity of the DC (FL)₈ hydrogel to adopt opposing 3D helical configurations. This series of reshaping processes highlights the hydrogel's exceptional adaptability and structural versatility, akin to the malleability of aluminum foil in taking on diverse customized forms while preserving its overall structural integrity and functionality.

The configurations obtained through reshaping exhibited remarkable stability in PBS, which was largely retained even after 10 days (Figure S3, Supporting Information). Moreover, the universal nature of protein folding and unfolding enables this reshaping approach to be applied across a wide range of proteins. For instance, hydrogels fabricated from (NuG2)₈ polyprotein^{[ 42 ]} can be readily reshaped, transforming a straight strip to a lightning‐shaped figure and then into a half‐circle form (Figure S4, Supporting Information). This combination of reversibility, tunability, stability, and versatility establishes a robust reshaping strategy to process hydrogels into different forms from the same starting precursor hydrogel.

2.4. Engineering Fully Recyclable and Reshapeable Protein Hydrogels

Having demonstrated the feasibility of reshaping protein hydrogels through protein unfolding and refolding, we then sought to impart recyclability to these reshapeable hydrogels to achieve full‐circle recycling. In practical applications, metal‐based materials are commonly recycled from end‐of‐life products, with metals being melted and recast into new forms. This process significantly reduces manufacturing costs and greenhouse gas emissions. However, similar recyclability is currently absent in hydrogels. Achieving full recyclability would not only minimize environmental waste associated with hydrogels but also greatly enhance sustainability by allowing these materials to be reused in new applications. Here, we aim to develop fully recyclable and reshapeable DC (FL)₈ hydrogels.

The reshapeable DC (FL)₈ hydrogels are covalently crosslinked via dityrosine adducts, rendering them non‐recyclable, although they can be degraded. To make the protein hydrogels fully recyclable, it is essential that the crosslinking points can be undone under mild conditions, so that the starting protein materials can be recovered and reused. Toward this goal, we explored the use of dynamic disulfide bonds as reversible crosslinks, which can be disrupted under mild reducing conditions (Figure 5a). To achieve this, we mutated the buried Met23 in FL to Cys to obtain FL‐M23C.^{[ 6} , ^{41 ]} To prepare DC (FL‐M23C)₈ hydrogels, denatured (FL‐M23C)₈ protein solutions were oxidized with air or H₂O₂ to form disulfide‐linked networks. The resulting N‐DC (FL‐M23C)₈ hydrogels displayed swelling and mechanical properties similar to the N‐DC (FL)₈ hydrogels, with a Young's modulus of 671 ± 83 kPa and a swelling ratio of −71 ± 3% (n = 6). Since disulfide bonds can reversibly break and reform in response to redox stimuli, we hypothesized that the DC (FL‐M23C)₈ hydrogel network could be broken down using reducing agents, allowing (FL‐M23C)₈ proteins to be recycled to produce new hydrogels.

Figure 5.

a) Schematic diagram of the formation and recycling process of the (FL‐M23C)₈ hydrogel through reversible redox reactions of disulfide crosslinks. Buried cysteine residues are exposed using GdmCl and can be crosslinked via H₂O₂ (or air) oxidation. Renaturation in PBS results in a stable N‐DC (FL‐M23C)₈ hydrogel. Recycling is achieved by denaturing the disulfide‐crosslinked hydrogel and then reducing it with TCEP. The recovered un‐crosslinked (FL‐M23C)₈ proteins can be further remodeled and re‐oxidized into a new hydrogel. b) Photos demonstrating the recycling of a column‐shaped DC (FL‐M23C)₈ hydrogel and its subsequent remodeling into a cone shape via sequential reduction and oxidation. c) 10% Coomassie blue G250 stained SDS‐PAGE gel of pristine (right lane) and recycled (FL‐M23C)₈ proteins (left lane). A prominent band corresponding to the fulllength (FL‐M23C)₈ was observed, while bands at higher molecular weights, which correspond to partially reduced multimers of (FL‐M23C)₈, were very faint, indicating most (FL‐M23C)₈ protein polymers were recovered by reducing the disulfide‐crosslinked hydrogel. d) Tensile stress–strain curves of 20% N‐DC (FL‐M23C)₈ hydrogels across 5 consecutive recycling and reformation cycles. The original hydrogel (cycle 1) displayed a notably higher Young's modulus, while the reformed hydrogels (cycles 2–5) showed comparable stiffness among themselves. e) Summary of the Young's modulus (gray) and toughness (blue) for 20% N‐DC (FL‐M23C)₈ hydrogels over 5 recycling and reformation cycles. Error bars represent standard deviation (n = 6). In general, both the original and reformed hydrogels exhibited consistent Young's modulus and toughness across all cycles. f) Swelling ratio of 20% DC (FL‐M23C)₈ hydrogels in PBS and 7 m GdmCl over 5 recycling and reformation cycles. Error bars represent standard deviation (n = 6). Highly consistent swelling behavior was observed between the original and reformed hydrogels.

To test this hypothesis, a column‐shaped N‐DC (FL‐M23C)₈ hydrogel was incubated in 7 m GdmCl containing 10 mm TCEP for 8 h at room temperature. As shown in Figure 5b, the hydrogel underwent a gel‐to‐sol transition, resulting in complete dissolution. SDS‐PAGE analysis of the recovered protein (Figure 5c) confirmed successful recovery of the starting protein building block. The recovered protein was then re‐oxidized using H₂O₂ to form a new transparent cone‐shaped (FL‐M23C)₈ hydrogel (Figure 5b). These results demonstrated that the redox‐driven gel‐sol transition and subsequent re‐gelation enabled the effective recycling of (FL‐M23C)₈ protein from its hydrogel. Moreover, the regenerated N‐DC (FL‐M23C)₈ hydrogels showed a Young's modulus (538 ± 76 kPa) and swelling ratio (−71 ± 3%) that are comparable to those of the pristine hydrogels, and the mechanical properties (Young's modulus, toughness and swelling ratios) of the recycled hydrogels remained consistent across multiple recycling cycles (Figure 5d–f). These highly reproducible mechanical and swelling characteristics demonstrate the robustness and reliability of the recycled protein hydrogels. The incorporation of dynamic disulfide bonds ensures both reversibility and the retention of network strength in N‐DC (FL‐M23C)₈ hydrogels, offering outstanding mechanical stability alongside recyclability.

In addition to recyclability, the DC (FL‐M23C)₈ hydrogel maintained its reshaping ability through the protein folding and unfolding, owing to the significant difference in stiffness between the recycled N‐DC and D‐DC (FL‐M23C)₈ hydrogel. As depicted in Figure 6 , a ring‐shaped N‐DC (FL‐M23C)₈ hydrogel was reshaped into a triangular shape. The triangular hydrogel was then recycled under reducing conditions to recover the protein building block (FL‐M23C)₈. The recycled protein solution was then cast into a square mold and oxidized using H₂O₂ to generate a square‐shaped D‐DC (FL‐M23C)₈ hydrogel sheet, which was further reshaped into a rolled form of the N‐DC hydrogel. Evidently, the DC (FL‐M23C)₈ hydrogel represents a fully recyclable protein hydrogel, and the recovered (FL‐M23C)₈ protein can be used to engineer new reshapeable DC (FL‐M23C)₈ hydrogels in a way identical to that used for the pristine protein building block. It is of note that although the recyclability and reshapeability were demonstrated on 20% DC (FL‐M23C)₈ hydrogels, (FL‐M23C)₈ hydrogels with different protein concentrations (>10%) are also fully recyclable and reshapeable.

Figure 6.

Photographs showing the consecutive reshaping and remolding of DC (FL‐M23C)₈ hydrogels. Scale bar: 5 mm. The original ring‐shaped hydrogel was first programmed into a triangular geometry and subsequently remolded into a flat sheet. The flat sheet was then reconfigured to a rolled form before being further recycled and remolded into a sea star shape. The arms of the sea star could be bent and later restored. The recycling and reformation process of the DC (FL‐M23C)₈ hydrogel via redox reactions enables virtually unlimited remolding possibilities.

Moreover, due to the reversible nature of the redox reaction of disulfide bonds, DC (FL‐M23C)₈ hydrogels can be recycled for many cycles and remolded into completely new shapes. For example, through the first recycling cycle, a (FL‐M23C)₈ hydrogel sheet was fabricated. This regenerated hydrogel sheet was recycled again and used to produce a flat star‐shaped D‐DC (FL‐M23C)₈ sheet, which was subsequently reshaped into a stellate 3D shape. These results demonstrated the seamless integration of recyclability and reshapeability into the protein hydrogel, significantly extending the service life of protein hydrogels and enhancing their adaptability. This strategy for engineering recyclable and reshapeable protein hydrogels can be easily generalized to cysteine‐rich proteins, either naturally occurring or engineered ones. For example, hydrogels constructed using cysteine‐rich BSA protein can be fully recyclable using this strategy (Figure S5, Supporting Information).

3. Conclusion

We have developed a simple yet effective and robust approach for engineering fully reshapeable and recyclable protein hydrogels by combining protein folding‐unfolding dynamics with dynamic disulfide crosslinks. Benefiting from the substantial stiffness difference between the native and denatured states of DC (FL)₈ hydrogels, we successfully achieved 2D and 3D reshaping of protein hydrogels with excellent stability. The reversible and GdmCl concentration‐dependent nature of protein folding and unfolding endowed the hydrogels with highly reproducible and tunable reshaping behaviors.

Compared to systems that require reinforcement with polyelectrolytes or metal ions, the diverse configurations achieved in our hydrogels are stabilized solely by the protein's folded structures, thereby leaving the biochemical functionality of the protein network unaffected. This unique feature ensures the compatibility of the hydrogels with a wide range of applications. Moreover, by replacing static dityrosine bonds with dynamic disulfide bonds, we created fully recyclable protein hydrogels that can be repeatedly remolded into entirely different shapes and further reconfigured, offering exceptional adaptability to customized needs.

The recyclability and reprocessability of these protein hydrogels not only extend their service life but also improve resource efficiency and reduce costs. We envision that these combined attributes will transform the application paradigm of protein hydrogels, where they can be manufactured and delivered in a precursor form, modified by end‐users to meet specific needs, and eventually recycled for alternative uses. Our approach paves the way for next‐generation biomaterials that combine sustainability and dynamic functionality, broadening the practical applications of protein‐based hydrogels in fields such as soft robotics, data encryption, sustainable packaging, and biomedical engineering.

4. Experimental Section

Protein Engineering

Polyproteins (FL)₈, (NuG2)_8, and (FL‐M23C)₈ were previously constructed in the lab using the identical sticky ends generated by BamHI and BglII restriction enzymes.^{[ 6} , ²⁷ , ⁴¹ , ^{42 ]} The amino acid sequence of the FL domain was MGEFDIRFRT DDDEQFEKVL KEMNRRARKD AGTVTYTRDG NDFEIRITGI SEQNRKELAK EVERLAKEQN ITVTYTERGS LE. The FL‐M23C domain was constructed based on the predicted structure of FL using standard site directed mutagenesis.^{[ 6 ]} NuG2 was a computationally designed fast‐folding mutant of protein GB1,^{[ 42 ]} with the sequence MDTYKLVIVL NGTTFTYTTE AVDAATAEKV FKQYANDNGV DGEWTYDDAT KTFTVTE. For this work, (FL)₈, (NuG2)₈ and (FL‐M23C)₈ were newly expressed in E. coli strain DH5α. All polyproteins were purified via Co²⁺‐affinity chromatography using previously reported methods.^{[ 27} , ⁴¹ , ^{43 ]} The purified proteins were dialyzed in deionized water at 4 °C for 24 h and then lyophilized.

Hydrogel Preparation

(FL)₈ and (NuG2)₈ hydrogels were synthesized based on the [Ru(bpy)₃]²⁺‐mediated photochemical crosslinking strategy, following previously reported protocols.^{[ 6} , ⁴³ , ⁴⁴ , ⁴⁵ , ^{46 ]} To prepare 15% DC (FL)₈ hydrogels, pre‐gel solutions containing 150 mg mL⁻¹ polyprotein, 50 mm ammonium persulfate, and 0.26 mm [Ru(II)(bpy)₃]Cl₂ in 7 m GdmCl were centrifuged at 12,000 rpm for 3 min to remove air bubbles. The solutions were then cast into molds and irradiated with a 200 W fiber‐optical white light source from a distance of 10 cm for 10 min. To tune the R_f and R_r values of 15% DC (FL)₈ hydrogels by varying their extent of chain entanglement, pre‐gel solutions were prepared in GdmCl solutions of 0, 0.5, 1, 1.5, 2, 2.5, 3, 4, 5, 6, and 7 m, followed by irradiation under the same conditions.

For DC (FL‐M23C)₈ hydrogels, a 20% (FL‐M23C)₈ solution in 7 m GdmCl was centrifuged at 12 000 rpm for 3 min to eliminate air bubbles. After being transferred into molds, the protein solution was left in a moist Petri dish at room temperature overnight. Oxidation was induced by dissolved O₂ in the solution. 10% DC BSA hydrogel was prepared by first dissolving BSA in 7 m GdmCl at 100 mg mL⁻¹ protein concentration. The pre‐gel solution was reduced by 50 mM TCEP and then crosslinked by adding 100 mm H₂O₂.

Ring‐shaped hydrogels were cast in molds with an inner diameter 8 mm, an outer diameter 10 mm, and a depth 3 mm. Flat sheet hydrogels were prepared in a 20 mm × 20 mm square mold with a depth of 1 mm. Tubing gels were fabricated using PTFE #18 AWG tubing.

Swelling Test

To determine the swelling ratio, the initial mass (m_i ) of the hydrogel samples was measured immediately after demolding. For the denatured condition, hydrogels were soaked in 7 m GdmCl solution for a minimum of 3 h to achieve swelling equilibrium. After soaking, the hydrogels were gently blotted and weighed to obtain the swollen mass (m_s ). For the renatured condition, hydrogels were washed and renatured in PBS before measuring m_s . The swelling ratio (SR) was calculated as

$$SR=\frac{m_s-m_i}{m_i}\times 100\%$$ (1)

Tensile Experiment

For tensile tests, hydrogels were prepared using a ring‐shaped Plexiglass mold (d _in = 8 mm, d _out = 10 mm, h = 3 mm).^{[ 47 ]} The tests were conducted using an Instron‐5500R tensometer equipped with a customized force gauge. Prior to testing, hydrogel samples were equilibrated in PBS or 7 m GdmCl solution. During the tensile test, samples were immersed in the corresponding buffer at 25 °C, matching the conditions used in swelling tests to ensure swelling equilibrium. The hydrogel sample was affixed at one end and stretched by a hook connected to the force gauge at a rate of 10 mm min⁻¹. Tensile stress was calculated as σ = F/A, where F was the measured loading force and A was the cross‐sectional area under tension. Tensile strain was calculated as ε = (L − L_init)/L_init  × 100%, where L was the recorded extension, and L_init was the initial length of the sample at zero tension. Young's modulus was determined as the slope of the stress–strain curve at 15% strain.^{[ 47 ]}

Shape Programming and Recovery Characterization

After demolding, the D‐DC (FL)₈ hydrogel strips were first equilibrated in 7 m GdmCl without constraint to establish their initial configuration (180° in this study). The hydrogels were then bent to the maximum angle θ _m . Denaturation was carried out in PBS while maintaining the bending constraint. Upon removing the force, the fixed bending angle θ _f of the hydrogel was immediately measured. Next, the hydrogels were immersed in 7 m GdmCl again to erase the fixed bent form. Following final renaturation, the residual bending angle θ _r was measured.

To study the effect of programming GdmCl concentration on R_f, hydrogels were equilibrated in 7 m GdmCl solution, deformed, and then immersed in 0, 0.5, 1, 1.5, 2, 2.5, 3, 4, 5, 6, and 7 m GdmCl solutions for fixation.

All angles were recorded on camera and quantified using ImageJ, as illustrated in Figure S2 (Supporting Information). The fixity ratio (R_f) and recovery ratio (R_r) were calculated as

$$R_f=\frac{{\theta }_f}{{180}^{\circ }-{\theta }_m}\times 100\%$$ (2)

$$R_r=\frac{{\theta }_f-{\theta }_r}{{\theta }_f}\times 100\%$$ (3)

Hydrogel Reshaping

To reshape a ring‐shaped DC (FL)₈ hydrogel into various 2D geometries, the hydrogel was first denatured in 7 m GdmCl and then programmed into a triangular form using a custom‐made fixer. The new geometry was stabilized in PBS through protein refolding. Before initiating the second reshaping cycle, the triangular hydrogel was treated with 7 m GdmCl to restore its original ring shape. It was then deformed into a rectangle and renatured to fix the configuration. Lastly, the hydrogel was reshaped into a pentagon following the same procedure. 3D reshaping of a flat sheet and strip‐shaped DC (FL)₈ hydrogel was performed using a similar approach.

Hydrogel Remolding

Remolding of a column‐shaped N‐DC (FL‐M23C)₈ hydrogel was conducted in two steps: recycling and reformation. For recycling, the hydrogel was first denatured with 7 m GdmCl to expose the buried cysteine crosslinks. Reduction was then initiated with 2 mm TCEP and allowed to proceed for 8 h at room temperature to un‐crosslink the hydrogel network. Next, the recycled (FL‐M23C)₈ polyproteins, now in a denatured state, were transferred to an Eppendorf tube and centrifuged at 12000 rpm for 3 min to remove air bubbles. To reform the hydrogel, an excess of H₂O₂ was added to the protein solution to induce the oxidation of exposed cysteine residues, resulting in a D‐DC (FL‐M23C)₈ hydrogel in a cone shape. The D‐DC hydrogel was then washed with PBS to refold the FL‐M23C domains.

The alternating reshaping and remolding scheme of a ring‐shaped DC (FL‐M23C)8 hydrogel was performed by combining the approaches outlined here and in the Hydrogel Reshaping section.

Statistical Analysis

The experimental data were analyzed using Igor Pro 8.0 (Wavemetrics), and the data are presented as the mean ± standard deviation.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Supporting Information

Bian Q. and Li H., “Fully Reshapeable and Recyclable Protein Hydrogels.” Adv. Mater. 37, no. 42 (2025): e09812. 10.1002/adma.202509812

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.