Sub-400 nanometer-thick skin and environment adaptable organohydrogel nanofilm epidermal electrode

Wang, Zonglei; Wang, Yuli; Yang, Jiawei; Zhou, Pengcheng; Yan, Wenqing; Lin, Shihong; Zhang, Yujie; Sun, Qingyuan; Xu, Yumiao; Ji, Zichong; Wang, Mingzhe; Zhang, Zongman; Yi, Junhong; Zheng, Meiqiong; He, Xuezhong; Sun, Lulu; Lee, Sunghoon; Yokota, Tomoyuki; Haick, Hossam; Someya, Takao; Wang, Yan

doi:10.1038/s41467-025-65089-0

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Published: 18 November 2025

Sub-400 nanometer-thick skin and environment adaptable organohydrogel nanofilm epidermal electrode

Nature Communications volume 16, Article number: 10103 (2025) Cite this article

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Abstract

Hydrogels are crucial for soft bioelectronics in long-term health monitoring; however, reconciling skin comfort with environmental resilience remains a major challenge. We present a 392 nm-thick organohydrogel nanofilm electrode that mimics skin deformation, offers high gas/water vapor/sweat permeability and heat transfer, and remains functional under various extreme conditions. The electrode comprises a genipin-crosslinked gelatin matrix, reinforced by polyurethane nanomeshes and plasticized with a glycerol/sodium chloride/tannic acid electrolyte. It achieves ultralow flexural rigidity (8.7 × 10⁻¹¹ nN·m), high stretchability (166.3% strain), toughness (3.0 MJ m⁻³), adhesion (365.8 µJ cm⁻²), and durability (1000 cycles at 100% strain). Solvent replacement strategies suppress ice formation and evaporation, preserving its physical and electrical performance under extreme conditions (−80–150 °C, 2% relative humidity, vacuum) and 200-day ambient storage. The organohydrogel nanofilm electrodes record stable electrocardiograms for 9 consecutive days with superior resistance to motion and sweat artifacts, offering a resilient platform for skin-integrated bioelectronics.

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Introduction

Hydrogels, whose softness, water content, and tunable chemistry echo those of living tissue, are foundational to the next generation of skin-conformal bioelectronics for continuous health monitoring and early disease intervention, even in extreme settings^1,2,3. To function reliably, hydrogel-enabled epidermal electronics must satisfy two demanding but often conflicting requirements: skin adaptability, the ability to conform, breathe, adhere, and move with epidermis during daily life, and environmental adaptability, the capacity to retain mechanical integrity, ionic conductivity, and biocompatibility under freezing, arid, or vacuum conditions^4,5,6,7,8.

Skin adaptability is a pivotal requirement for continuous on-body health monitoring, necessitating that hydrogel-based epidermal electronics conform seamlessly to the skin without disrupting daily activities^9,10,11. Skin adaptability encompasses mechanical compatibility, dynamic skin compliance with high skin adhesion and robust hydrogel-skin interfaces, and wearing comfort, characterized by favorable air and water vapor permeability, efficient sweat transport, and effective heat dissipation^5,12. Incorporating catechol groups is a widely used strategy to enhance hydrogel adhesion, while reinforcing fillers significantly improve mechanical strength without sacrificing softness^13,14,15. Crucially, reducing film thickness lowers flexural rigidity and shortens permeation paths for gases and heat^16,17,18. For example, a 5 μm-thick microfiber-reinforced poly(vinyl alcohol) (PVA)/glycerol/sodium chloride (NaCl) hydrogel sensor conforms to human skin and reliably captures high-quality electromyography (EMG) signals for up to 48 h. It exhibits a tensile strength of approximately 6 MPa, along with good resistance to tearing and dehydration¹⁹. Similarly, a 10 μm-thick nanomesh-reinforced, gas-permeable gelatin/borax/glycerol/sodium sulfate hydrogel sensor enables continuous daily electrocardiogram (ECG) monitoring for up to 24 h²⁰. Further reducing thickness into the sub-micrometer regime could dramatically enhance gas permeability and skin compliance^17,21, however, achieving ultrathin dimensions without compromising mechanical robustness and anti-dehydration performance remains a major challenge.

Environment adaptability is equally essential for hydrogel-based devices intended for long-term use, since they need to preserve physical and electrical functionality when exposed to extreme temperatures, low humidity, vacuum, and bacteria^3,22. Conventional hydrogels, due to their high water content, are highly susceptible to freezing, rapid dehydration, and mechanical failure in such environments. Approaches such as incorporating inorganic salts, binary solvents, and employing solvent replacement can help maintain conductivity and mechanical flexibility across wide temperature ranges^23,24,25, while robust covalent crosslinks bolster the thermal resistance of these materials^26,27,28. Nonetheless, as summarized in Supplementary Tables 1^{19,20,29,30,31,32,33} and 2^{34,35,36,37,38,39}, achieving both high skin adaptability and environmental compatibility in a single hydrogel platform for sustained health monitoring remains a considerable challenge.

Here, these challenges are addressed by developing a sub-400 nm-thick organohydrogel nanofilm epidermal electrode for high-fidelity, long-term health monitoring, even under extreme environments. The fabrication involves stepwise dip-coating a polyurethane (PU) nanomesh with a genipin and a gelatin hydrogel dilute solution, followed by immersion in a ternary electrolyte solution of glycerol, NaCl, and tannic acid (TA). The resulting organohydrogel nanofilm achieves a thickness of 392 nm, ultralow flexural rigidity (8.7 × 10⁻¹¹ nN·m), strong areal adhesion (365.8 μJ cm⁻²), high stretchability (166.3% strain), and robust mechanical stability (over 1000 cycles at 100% strain). It also exhibits effective air/water vapor/ sweat permeability, and heat transfer, ensuring superior wearability. This organohydrogel nanofilm maintains mechanical and electrical integrity after exposure to bacteria, extreme temperatures (−80 to 150 °C), low humidity (2% relative humidity (RH)), vacuum (−0.1 MPa), and prolonged storage (200 days at room temperature). By reliably maintaining high-quality ECG signals over 9 consecutive days of continuous use, even under challenging environmental conditions, this approach establishes a promising pathway for enduring, versatile, and accurate health monitoring across diverse scenarios, spanning everyday activities to extreme polar and aerospace environments.

Results

A sub-400 nm-thick, adaptable organohydrogel film

Previous studies demonstrated that PU nanomesh reinforcement enables nano/microfilms to simultaneously achieve high mechanical strength and ultrathin dimensions^17,20. Building on this concept, a stepwise dip-coating strategy using electrospun PU nanomeshes was employed (Supplementary Fig. 1). The nanomeshes were first dip-coated in a genipin solution⁴⁰, then in a high-temperature (80 °C) dilute gelatin sol containing glycerol to enhance wetting and coating uniformity. Each coating step was followed by approximately 5 s of air-drying. This stepwise dip‑coating limits genipin-gelatin contact at elevated temperature, preventing premature gelation while ensuring a homogeneous, ultrathin coating. After gelation at ambient or physiological temperature, the composite film was soaked for 24 h in a ternary electrolyte (glycerol, NaCl, TA) and finally air‑dried under ambient conditions (Supplementary Note 1, Supplementary Figs. 2 and 3).

Under optimal conditions, namely, a PU nanomesh density of 0.1 mg cm⁻², 2 g of gelatin, 60 mg of genipin, and a ternary electrolyte solution with glycerol:NaCl:TA mass ratio of 92:8:2, the organohydrogel nanofilm achieves a thickness of only 392 nm (Fig. 1A, Supplementary Fig. 4). For clarity, the formulation is designated as PU_0.1-GHGNT, where: PU_0.1 denotes a PU nanomesh with a density of 0.1 mg cm⁻²; H denotes a physically cross-linked hydrogel composed of gelatin, water, and glycerol; GH represents a genipin-crosslinked gelatin hydrogel (genipin, gelatin, water, glycerol); GNT indicates the electrolyte solution (glycerol, NaCl, and TA); GHGNT refers to the organohydrogel formed by soaking the GH in the GNT; PU_0.1-GHGNT thus denotes the fully reinforced nanofilm combining PU_0.1 with the GHGNT layer. For comparison, PU_0.1-HGNT is also defined, in which a non-crosslinked gelatin hydrogel (gelatin, glycerol, water) is soaked in GNT and reinforced with the same PU_0.1 nanomesh. Subscripts indicate specific component quantities (Supplementary Table 3).

**Fig. 1: A sub-400 nm-thick, adaptable organohydrogel film.**

As illustrated in Fig. 1B, C, the resulting organohydrogel nanofilm conforms readily to the curvature of a human wrist, demonstrating its sub-micron thickness, high transparency (90.5% transmittance at 550 nm). Supplementary Fig. 5 further confirms its suitability for uniform, large-scale fabrication. Figure 1D presents a schematic depiction of the internal structure of the organohydrogel nanofilm. The scanning electron microscopy (SEM) image shows the random distribution of PU nanomeshes embedded in the organohydrogel matrix, and the organohydrogel nanofilm can seamlessly conform to the surface of the rough fingerprint replica (Fig. 1E). Although the average diameter of the PU nanomesh is approximately 410 nm, the average organohydrogel nanofilm thickness is even smaller. Junction points between these nanomeshes contribute to subtle surface undulations.

The organohydrogel nanofilm demonstrates skin adaptability, characterized by permeability to gases, vapors, and sweat, favorable thermal conductivity, and dynamic mechanical compliance with human skin. Additionally, the organohydrogel nanofilm offers robust environmental resilience, exhibiting properties such as anti-freezing capability, heat and dehydration resistance, vacuum stability, and antibacterial performance. These multifunctional advantages are attributed to the ultrathin geometry, reinforced nanomesh structure, moisture retention provided by glycerol, and robust covalent crosslinking (Fig. 1F).

Mechanical and adhesive properties of the PU nanomesh reinforced organohydrogels

Figure 2A highlights the remarkable mechanical robustness of the organohydrogel nanofilm (7.2 mg), which can support a weight of 7.9 g, 1100 times its own weight. The mechanical properties are influenced by several parameters, including the gelatin content, genipin concentration, PU nanomesh density, and electrolyte composition.

To obtain thinner yet robust organohydrogel films, the gelatin content was first optimized while fixing a PU nanomesh density of 0.1 mg cm⁻² for dip-coating. Reducing gelatin from 8.0 to 2.0 g lowered film thickness from 2400 to 373 nm (Supplementary Fig. 6A). Below 2.0 g, a continuous film no longer formed, disrupting stress transfer and causing stress concentration, which results in a sharp drop in tensile strength and toughness (Supplementary Fig. 6B). Consequently, for practicality and balancing minimal thickness with mechanical integrity, 2.0 g of gelatin was selected for all subsequent experiments.

Genipin covalently cross‑links gelatin (Supplementary Note 2, Supplementary Fig. 7). As genipin content rises, the tensile strength increases (Fig. 2B). At 60 mg genipin (PU_0.1‑G₆₀HGNT) the tensile strength reaches 2.2 MPa with an elongation‑at‑break of 166.3%, 4.5 times higher than the physically cross‑linked control (PU_0.1‑HGNT). Beyond 60 mg, excessive cross‑link density suppresses chain mobility, reducing the elongation and fracture toughness (Supplementary Fig. 8).

Reinforcement with the PU nanomesh is pivotal. Relative to the unreinforced hydrogel (GHGNT), the PU‑reinforced nanofilm (PU_0.1‑GHGNT) shows a 22‑fold gain in tensile strength and a 163‑fold gain in fracture toughness, reaching 2.2 MPa and 3.0 MJ m⁻³, respectively (Fig. 2C, Supplementary Fig. 9). The mechanical enhancement arises from a “reinforced concrete” architecture, where the nanomesh serves to redistribute stress and dissipates energy across the organohydrogel-mesh interface. Increasing PU loading above 0.1 mg cm⁻² slightly lowers strength because of stress concentration within densely packed nanomeshes, but further elevates the elongation‑at‑break (Fig. 2C).

The soaking electrolyte acts as a conductor, plasticizer and secondary cross‑linker. Glycerol softens the network, while TA strengthens it through hydrogen‑bonding (Supplementary Note 3, Supplementary Figs. 10 and 11). As a result, soaking PU_0.1‑GH in the glycerol/NaCl/TA solution (forming PU_0.1‑GHGNT) reduces Young’s modulus from 3.4 ± 0.2 to 1.3 ± 0.1 MPa and significantly increases the elongation‑at‑break (Fig. 2D, Supplementary Fig. 12). The modulus of 1.3 MPa closely matches that of human epidermis⁴¹. The organohydrogel nanofilm endures 1000 loading-unloading cycles at 100% strain with negligible hysteresis (Supplementary Fig. 13).

The flexural rigidity (D) of the two-dimensional film is critically related to its compliance on three-dimensional (3D) surface and is defined by the equation

$$D=\frac{E{t}^{3}}{12\left(1-{v}^{2}\right)}$$

(1)

Where E, t and ν represent the Young’s modulus, thickness and Poisson’s ratio of the film, respectively. By reducing the film thickness, D is effectively decreased. The flexural rigidity of the organohydrogel nanofilm is calculated as 8.7 × 10⁻¹¹ nN·m, well below those of the stratum corneum (1.1 × 10⁻¹⁰ nN·m), basal epidermis (4.2 × 10⁻⁹ nN·m), and dermis (6.7 × 10⁻⁵ nN·m)⁴², ensuring reliable skin compliance. Compared to representative epidermal electronics and skin tissues, the organohydrogel nanofilm demonstrates the lowest flexural rigidity, enhancing its ability to mimic natural skin topography (Fig. 2E)^{17,21,42,43,44,45}.

When applied to artificial skin, the organohydrogel nanofilm seamlessly conforms without visible transition at the interface (Supplementary Fig. 14). Surface contour analysis shows close topographic matching between covered and uncovered regions, unlike polydimethylsiloxane (PDMS) films, which significantly alter surface roughness. Microscopy confirms the preservation of fine skin textures beneath the organohydrogel nanofilm, in stark contrast to the blurring seen with PDMS films (Supplementary Fig. 15A−D).

Thanks to abundant polar groups (hydroxyl, carboxyl, amino, phenolic hydroxyl), the organohydrogel nanofilm adheres effectively to human skin via hydrogen bonding. Adhesion was quantified using tack-separation tests (Fig. 2F). PU_0.1-GHGNT exhibited an adhesion force of 44 mN and a separation distance of 13 mm (Fig. 2G), corresponding to an areal adhesive energy of 365.8 μJ cm⁻², 3.9 and 2.2 times higher than PU_0.1-GH and PU_0.1-GHGN, respectively (Fig. 2H). These robust adhesive properties arise from: (1) high compliance due to ultralow flexural rigidity and (2) strong molecular interactions. High compliance enlarges the contact area between the human skin and the organohydrogel nanofilm, strengthening van der Waals forces and further boosting adhesion.

Cyclic adhesion tests confirm durable adhesion across 200 attachment-detachment cycles on artificial skin (Supplementary Fig. 16). The organohydrogel nanofilm dynamically deforms with skin stretching, aided by its self-adhesive and stretchable nature (Supplementary Fig. 15E, F). On human skin, the organohydrogel nanofilm adapts effortlessly to wrinkles and folds during daily movements (Supplementary Fig. 17).

The organohydrogel nanofilm also adheres robustly to various materials, including stainless steel (SS), polytetrafluoroethylene (PTFE), glass, kraft paper (KP), and rubber (Supplementary Fig. 18). The organohydrogel nanofilm exhibited strong interfacial adhesion to Ag/AgCl snap electrodes, with a measured adhesion force of 55.7 mN and area adhesion energy of 450.8 μJ cm⁻², achieved through TA-metal coordination bonding⁴⁶. Even on inert, non-polar substrates like PTFE, the organohydrogel nanofilm achieves areal adhesion energy of up to 152.2 μJ cm⁻², underscoring its versatility.

Skin adaptability of organohydrogel nanofilm

Skin adaptability involves not only mechanical compliance but also wearing comfort, primarily sweat and gas/vapor permeability and heat transfer¹². These factors directly influence both wearing comfort and signal quality⁹. Figure 3 A and B summarize transport at the organohydrogel film/human skin/ambient interface. ${\theta }_{C}$ represents the contact angle between sweat and the film, and ${\gamma }_{{LG}}$, ${\gamma }_{{SL}}$, and ${\gamma }_{{SG}}$ denote the liquid‑gas, solid‑liquid, and solid‑gas interfacial tensions, respectively. According to Young’s equation^47,48, wetting and subsequent permeation occur when the contact angle ${\theta }_{C}$ < 90°, a condition governed by the balance of these interfacial tensions⁴⁹. Once the film adheres to skin, two contrasting micro‑environments arise: the sweat‑saturated (37 °C, 100% RH) skin side and the cooler, drier ambient side (25 °C, 30% RH). Driven by the second law of thermodynamics, heat, liquid, and vapor flow from the high‑ to the low‑concentration side until equilibrium is reached (Fig. 3B).

Sweat permeability depends mainly on film thickness and surface hydrophilicity. To visualize sweat transport, the organohydrogel nanofilm was clamped over a 4 mm hole in a glass slide, and the contact-angle change of an artificial-sweat droplet was tracked (Fig. 3C). Commercial gel, PDMS film, and organohydrogel films of various thicknesses showed markedly different behaviors (Fig. 3D, E). All organohydrogel films allowed sweat to pass; thinner films exhibited faster permeation (Supplementary Figs. 19 and 20, Supplementary Movie 1).

Increasing hydrophilicity accelerates wetting. After soaking in the electrolyte, the PU_0.1-GHGNT compared to the PU_0.1-GH exhibited a lower contact angle and a higher permeation rate (Supplementary Fig. 21). In contrast, hydrophobic PDMS film and PU_0.1 nanomesh sheet remained impermeable for ≥ 10 min (Supplementary Fig. 22). This discovery reveals that hydrophobic films exclusively allow vapor-phase sweat transport. A fluorescein sodium-labeled two‑liquid permeation test corroborated this: the organohydrogel nanofilm promoted rapid sweat permeation, whereas PDMS film acted as a barrier (Supplementary Fig. 23, Supplementary Movie 2).

The trend is general. For PU/PVA nanomeshes, raising the PVA (hydrophilic) fraction progressively lowered the contact angle and enhanced sweat flow (Supplementary Fig. 24). Effective sweat release stabilizes the organohydrogel nanofilm electrode-skin interface even during vigorous exercise, whereas significant sweat pooling was observed under the PDMS film (Supplementary Fig. 25).

Poor heat transfer of epidermal electronics can cause thermal discomfort to users^9,12. Thermal comfort was assessed by applying an organohydrogel nanofilm, a 2 μm-thick PDMS film, and a commercial gel to a volunteer’s chest (healthy male, 31 years old) and recording the central patch temperature (T_c) and adjacent bare skin temperature (T_s) with infrared imaging (Supplementary Fig. 25A). After 10 minutes running, $\triangle T={T}_{C}-{T}_{S}$ was negligible for the organohydrogel nanofilm but rose by around 1.8 °C for the commercial gel and 0.8 °C for PDMS film (Fig. 3F, G). After 20 min, the commercial gel lost adhesion, $\triangle T$ rose to approximately 2.2 °C, and the PDMS film showed an increase of approximately 0.4 °C, whereas the organohydrogel nanofilm remained thermally indistinguishable from bare skin. Heating tests confirmed thickness-dependent heat dissipation: reducing the organohydrogel films’ thickness from 200 µm to 392 nm shortened the time to reach 37 °C from room temperature from 172 to 7 s (Supplementary Fig. 26).

Air permeance, measured by the constant volume method, increased with temperature and decreased thickness, reaching 0.2 cm³cm⁻² s⁻¹ cmHg⁻¹ at 37 °C for 392 nm-thick organohydrogel nanofilm (Fig. 3H). Even at 25 °C, the organohydrogel nanofilm (0.1 cm³cm⁻² s⁻¹ cmHg⁻¹) outperformed a 3 μm-thick parylene film by about 1700 times. The water vapor transmission rate (WVTR) of the organohydrogel nanofilm was similarly high, reaching 1318.1 ± 3.5 g m⁻² d⁻¹ at 25 °C, which is 99.3 % of the open container rate. The value increased to 3069.7 ± 27.6 g m⁻² day⁻¹ at 37 °C (Fig. 3I, Supplementary Fig. 27).

Impedance measurements at 100 Hz during 20 min of running followed by 100 min rest revealed that sweat initially lowered contact impedance for all conditions. Bare human skin fell from 51.5 to 37.3 kΩ, organohydrogel nanofilm-covered skin from 45.7 to 35.2 kΩ, and the commercial gel sharply dropped to 22.4 kΩ. Crucially, impedance under the organohydrogel nanofilm returned to its baseline after rest, mirroring bare skin, whereas the commercial gel detached after 20 min and could not recover (Fig. 3J). The organohydrogel nanofilm combines rapid sweat/gas transport and efficient heat dissipation, enabling a stable, low‑impedance skin interface even under strenuous activity.

Environment adaptability of organohydrogel nanofilm

The environmental adaptability of the organohydrogel nanofilm was furthermore evaluated under four extreme conditions: very low or high temperature, low RH, and high vacuum. Adaptability depends on both the polymer network and the solvent composition. The H hydrogel is held together only by hydrogen bonds. After genipin cross‑linking, the GH hydrogel gains covalent junctions, producing an irreversible network. Subsequent immersion in an electrolyte replaces most free water with glycerol, TA, and NaCl, yielding the GHGNT organohydrogel (Fig. 4A). Infrared images show that the organohydrogel remains flexible between −20 to 120 °C, even at the limits it can still be stretched and twisted (Supplementary Fig. 28).

At sub‑zero temperatures the strong hydrogen bonding between glycerol and water disrupts the ice crystal lattice, lowering the freezing point (Supplementary Fig. 29). Differential scanning calorimetry (DSC) curves reveal exothermic peaks at −7 °C for H and −23 °C for GH, confirming that the denser GH network suppresses freezing. In contrast, GHGNT shows no exothermic from −80 to 20 °C, indicating superior anti‑freezing performance (Fig. 4B). Correspondingly, the organohydrogel nanofilm remains transparent and elastic at −80 °C, whereas PU_0.1‑GH is already glassy (Supplementary Fig. 30). At high temperature the free water in H and GH evaporates completely near 100 °C. Replacing water with glycerol lowers the matrix vapor pressure and raises the onset of thermal decomposition, raising it to 187.2 °C for GHGNT compared to 89.3 °C for GHGN (Fig. 4C, Supplementary Fig. 31). Extra hydrogen bonds from TA further reinforce the network and enhance thermal stability.

Long‑term anti-dehydration is another key metric. Owing to the low vapor pressure, increased osmotic pressure, and hydration effect of electrolyte solution, the organohydrogel nanofilm still retained ≥ 92.7% of its initial weight after 30 days under extreme conditions (−80 °C, 2% RH, or high vacuum). Even at 150 °C, the loss was slight (Fig. 4D, Supplementary Fig. 32). By contrast, PU_0.1‑GH lost 73.5% of its mass in 24 h at room temperature, whereas the organohydrogel nanofilm retained 96.3% after 200 days. Mechanical strength and self‑adhesion also remained unchanged (Supplementary Fig. 33). Compared with recently reported wide‑temperature organohydrogels, the organohydrogel nanofilm covers a broader range (Fig. 4E)^{34,35,36,37,38,39}. To further probe environmental adaptability, the organohydrogel nanofilms were stored under exceptionally harsh conditions, including −80 and 150 °C, 2% RH, and high vacuum for 30 days, and at room temperature for 200 days. Throughout these tests, the electrodes preserved an almost unchanged electrical impedance (Supplementary Fig. 34), highlighting their reliable performance across an ultrawide range of temperature, humidity, and pressure.

Antibacterial activity of the organohydrogel nanofilms was tested against Gram‑negative E. coli and Gram‑positive S. aureus. Clear inhibition zones of 1.7 and 2.1 cm, respectively, were observed (Fig. 4F, Supplementary Fig. 35). Colony counts showed ≥ 99.3% killing efficiency, which is attributed to the polyphenolic tannins that disrupt bacterial membranes⁵⁰.

Finally, benchmarking against state‑of‑the‑art hydrogel epidermal electronics highlights the organohydrogel nanofilm’s balanced performance in thickness, skin adhesion, weight retention, mechanical durability, antibacterial properties, and air permeance (Fig. 4G, Supplementary Table 1)^19,20,29,32.

Electrophysiological signals monitoring under different conditions

Organohydrogel nanofilm electrodes couple the skin‑adaptability discussed above with reliable environmental stability, making them well-suited for long‑term, high‑fidelity electrophysiological recording. Reliable acquisition hinges on (i) intimate, conformal skin contact, (ii) a persistently low contact impedance, and (iii) minimal resistance drift under strain. Compared with commercial gel electrodes, organohydrogel nanofilm electrodes show a lower initial skin impedance due to the superior conformal and permeable properties and remain almost unaffected by sweat and sebum accumulation over 9 consecutive days of wear^51,52 (Supplementary Fig. 36). Their relative resistance rises only 8.4% at 100% uniaxial strain (Supplementary Fig. 37A) and remains stable through 2500 stretch-release cycles at the same strain (Supplementary Fig. 37B), confirming stable electromechanical durability.

For ECG monitoring, a pair of organohydrogel nanofilm electrodes was placed on the subject’s chest (Supplementary Fig. 38). At rest, the organohydrogel nanofilm electrodes delivered waveforms and signal‑to‑noise ratios (SNR) comparable to those of commercial gel electrodes (Supplementary Fig. 39). During treadmill running at 7.5 km h⁻¹, sweat rapidly accumulated beneath the commercial gel electrodes, blurring the P and T waves and causing the electrodes to detach after around 16 min. In contrast, the organohydrogel nanofilm electrodes wicked sweat away, remained securely adhered, and delivered clear, undistorted ECG waveforms for the full duration of the test⁵³ (Fig. 5A, B; Supplementary Fig. 40, Supplementary Movie 3).

**Fig. 5: Sweat and motion artifacts and long-term health monitoring using organohydrogel nanofilm electrodes.**

Ambulatory ECG monitoring was then carried out for 9 consecutive days covering desk work, laboratory experiment, wash up, sleep, meal, rest, and walk (Fig. 5C). Continuous traces (Fig. 5D) and enlarged excerpts (insets) show no morphological drift. Over the first 24 h, heart‑rate profiles derived from the R-R intervals (upper, Fig. 5E) tracked activity levels: 57.6 ± 3.1 bpm during sleep, 92.5 ± 4.7 bpm while walking. Transient spikes linked to turning over or increased receiver distance. The average ECG SNR remained above 34.1 dB for all activities on day 1 (Supplementary Fig. 49A) and above 32.2 dB throughout days 2 to 9 (Supplementary Figs. 41−48, Supplementary Fig. 49B).

For alpha (α) wave EEG monitoring, organohydrogel nanofilm electrodes were applied to the right mastoid adjacent to a commercial gel electrode reference (Fig. 5F). Resting‑state recordings with eyes open or closed exhibited matching amplitudes and spectral content (Supplementary Fig. 50); eye closure increased α power as expected. To probe environmental robustness, organohydrogel nanofilm electrodes were stored for 30 days at −80 °C, 150 °C, 2% RH, or high vacuum (−0.1 MPa), and for 200 days at room temperature. Impedance spectra showed negligible change (Supplementary Fig. 34). Post‑storage α wave EEG still displayed a pronounced 10 Hz α peak in the power spectral density (PSD) (Fig. 5G−J) and retained stable periodic α rhythm during 10 min recording after 200 days (Supplementary Fig. 51).

The organohydrogel nanofilm electrodes proved adept at capturing a diverse array of electrophysiological signals, EMG, motor conduction velocity (MCV), electrooculography (EOG), and the event‑related potential (ERP) P300, matching or outperforming commercial gel electrodes across every test (Supplementary Fig. 52). When positioned over the flexor carpi radialis, they generated EMG waveforms identical in shape to those from gel electrodes but with a higher SNR. In MCV measurements, stimulation of the median nerve and recording from the abductor pollicis brevis elicited a robust 14.3 mV M‑wave that rose and decayed within roughly 4 ms, demonstrating precise temporal fidelity. During horizontal eye movements, the organohydrogel nanofilm electrodes produced clean, periodic EOG traces, underscoring their suitability for ocular‑signal acquisition. Finally, in a visual oddball paradigm, they yielded a clear P300 response whose target‑versus‑non‑target difference closely mirrored that obtained with conventional commercial gel electrodes. Collectively, these results highlight the electrodes’ versatility and high‑fidelity performance across multiple electrophysiological modalities.

Discussion

This work establishes that sub‑400 nm organohydrogel nanofilms can marry ultrathin form with notable compliance to human skin, remaining reliable even under harsh mechanical and environmental stress. Until now, sub‑micrometer thickness was largely the province of “dry” epidermal electronics; here we show that “wet” hydrogel‑based devices can achieve the same nanoscale profile without sacrificing performance^18,54. Critically, the films combine mechanical toughness, permeability to air, vapor, and sweat, superior skin adhesion, and resistance to temperature swings and dehydration, traits essential for comfortable, long‑term skin applicability. These properties enable reliable function not only during skin contact, but also across storage, transport, and deployment in extreme or mission-critical environments where conventional hydrogels typically fail, such as in arctic medical monitoring, space-suit biosensing, cold-chain vaccination, and high-temperature industrial settings. The fabrication technique is efficacious, universally applicable, and amenable to upscaling. Collectively, the resilience and versatility of organohydrogel nanofilms promise to reshape materials and design strategies for next‑generation soft bioelectronics in personalized healthcare.

Methods

Materials

Type A pigskin-derived gelatin powder (Bloom value ~300 g) was obtained from Sigma-Aldrich. Genipin, NaCl, glycerol, TA, N, N-dimethylformamide (DMF), and tetrahydrofuran (THF) were purchased from Aladdin Industrial Corporation. Artificial sweat (pH 6.5), containing NaCl, KH₂PO₄, Na₂HPO₄, urea, and lactic acid, was purchased from Codow Chemical Co., Ltd. Fluorescein sodium was purchased from Macklin. PU (1185 A) was provided by BASF GmbH. Artificial skin (BIOSKIN plate) was purchased from Beaulax. The commercial gels (Vitrode L, primarily composed of polymethyl acrylate) were purchased from NIHON KOHDEN. Porous alumina membranes (Anodisc, diameter: 25 mm, thickness: 60 μm) were supplied by Whatman International. The thin glass slide with a 4 mm hole was fabricated in-house using a laser cutting machine. All chemicals were used as received unless otherwise specified.

Fabrication of organohydrogel nanofilm

A 13 wt% PU electrospinning solution was prepared by dissolving PU in a 1:1 (w/w) mixture of DMF and THF. The PU nanomesh was fabricated using an electrospinning process as described in previous studies^10,55. Briefly, the prepared solution was loaded into a 5 mL syringe with a 27-G needle and placed in an electrospinning apparatus (Yunfan YFSP-T, Tianjin, China). Electrospinning was performed under the following conditions: 11 kV voltage, 0.001 mm s⁻¹ injection rate, and 500 rpm drum speed. The needle tip was positioned 12.5 cm from the collection drum, which was lined with silicon-coated paper to collect the PU nanomesh. The final nanomesh was transferred onto polyimide (PI) or polyethylene terephthalate (PET) frames of various sizes.

The genipin solution was prepared by dissolving genipin in a glycerol-deionized water mixture solution (20:80, w/w), and the hydrogel solution was obtained by dissolving gelatin in the same mixture solution. During the preparation of the hydrogel films, both the genipin solution and the hydrogel solution were maintained at 80 °C. The hydrogel films were fabricated by a stepwise dip-coating process using PU nanomesh. First, the PU nanomesh was soaked in the genipin solution, then in the gelatin solution, and finally suspended at room temperature for 24 h to form the hydrogel films.

To prepare the organohydrogel nanofilms, the hydrogel films were immersed in the ternary electrolyte solution (glycerol/NaCl/TA = 92:8:2) to replace the free water in the hydrogel. TA was added to the electrolyte solution to enhance the adhesion of organohydrogel film. To facilitate the dissolution of TA in electrolyte solution, 10% deionized water (by weight of glycerol) was used as a co-solvent. After complete dissolution of TA, the electrolyte solution was heated at 100 °C and stirred for 12 h to remove free water.

Characterizations

Fourier transform infrared (FTIR) spectra were recorded using a FTIR spectrometer (Nicolet iN10, Thermo Scientific, USA) in ATR mode. Each spectrum was acquired with 32 scans at a resolution of 4 cm⁻¹ in the range of 4000−500 cm⁻¹. Transparency was measured using UV-visible spectrometer (TU 1901, Beijing PERSEE, China) in the wavelength range of 400−800 nm. The cross-sectional thickness and surface morphology of the organohydrogel nanofilms were observed using a SEM (TM4000, HITACHI, Japan). The organohydrogel films were adhered to an anodized membrane and treated with liquid nitrogen to induce brittle fracture. Thickness measurements were taken using a stylus profilometer (Dektak XT, Bruker, Germany). Optical images and profile features were obtained using a 3D laser microscope (OLS5100-SAF, Olympus, Japan). The freezing point of the organohyrogel was determined using a DSC (TA25, TA Instruments, USA). Thermogravimetric analysis (TGA) was performed by heating the samples in a nitrogen atmosphere using a thermogravimetric analyzer (TGA55, TA Instruments, USA). Contact angle of the samples was detected with a goniometer at 25 °C (DSA25, Kruss, Germany).

Antibacterial properties evaluation

The antibacterial activity of organohydrogel against E. coli (Gram-negative) and S. aureus (Gram-positive) was evaluated using the inhibition zone and colony counting methods. E. coli (ATCC 25922) and S. aureus (ATCC 29213) were sourced from the China Center of Industrial Culture Collection.

For the inhibition zone test, bacterial suspensions of E. coli or S. aureus were diluted in PBS to a concentration of 10⁵ CFU mL⁻¹. A 200 µL aliquot of the suspension was spread evenly on a 9 cm nutrient agar plate. A 1 cm diameter organohydrogel was placed on the agar surface, and the plate was incubated at 37 °C for 24 h. Antibacterial activity was assessed by measuring the inhibition zone diameter with a vernier caliper. The inhibition zone diameter was calculated using the formula (2):

$${Inhibition\; zone\; diameter}={{{{\rm{d}}}}}_{t}-{{{{\rm{d}}}}}_{n}$$

(2)

where ${d}_{t}$ is the total inhibition zone diameter, and ${d}_{h}$ is the sample diameter.

In the colony counting method, 0.2 g of organohydrogel was wrapped in sterile filter paper and immersed in a 10⁷ CFU mL⁻¹ bacterial suspension. After incubation at 37 °C for 12 h, 50 µL of the suspension was plated on a 9 cm nutrient agar plate and incubated at 37 °C for 72 h. The number of colonies was counted, and the antibacterial rate was calculated by comparing the colony counts of the experimental and control groups. The antibacterial rate (%) was calculated using the formula (3):

$${Antibacterial\; rate}=\left(\frac{{N}_{c}-{N}_{h}}{{N}_{c}}\right)\times 100\%$$

(3)

where ${N}_{c}$ is the number of viable bacterial colonies in the blank control group, ${N}_{h}$ is the number of viable bacterial colonies in the organohydrogel group.

Tensile tress test

The organohydrogel films were vertically mounted on a fixture of a universal tensile tester with a 10 N sensor (ESM303, Mark-10, USA). The specimens were transferred onto a 10 mm × 20 mm PI frame. The left and right borders of the PI window were carefully separated using a sharp scalpel. The sides of the PI frame were then excised before measuring the tensile stress. The gauge length was 5 mm, and testing was conducted at a strain rate of 20 mm min⁻¹.

Tack separation test

A 3D-printed holder with a 2 cm × 2 cm window was used to support the PI frame. All samples were transferred to the PI frame (2 cm × 2 cm) and fixed to the holder. The initial adhesion area between the organohydrogel nanofilm and the target substrate was 1 cm × 1 cm. The holder was gradually lifted at a rate of 20 mm min⁻¹ until complete separation from the substrate, with the adhesion force recorded using a universal tensile tester (ESM303, Mark-10, USA). The areal adhesion energy was calculated from the adhesion force versus displacement curve by following Eq. (4):

$$E={\int }_{o}^{x}y \; {{{\rm{d}}}}L$$

(4)

where $E$, $x$, ${y}$, and $L$ are areal adhesion energy, the separation distance when sample was detached, adhesion force between the sample and substrate, and separation distance, respectively.

Electrical properties characterization

The electrical impedance of the samples was measured using an inductance-capacitance-resistance (LCR) meter (E4980AL, Keysight, USA) with an amplitude of 100 mV. Skin contact impedance was assessed using an electrochemical workstation (VersaSTAT3F-500, Princeton Applied Research, USA) over a frequency range of 10⁶ to 10⁻¹ Hz, with organohydrogel nanofilm electrodes attached to the human skin. Measurements were conducted using a pair of plate electrodes (1.25 cm in diameter), adhered to the forearm skin with a 2.5 cm center-to-center distance, and compared with commercial gel electrodes of the same size.

Gas permeance

Gas permeance was measured using a constant permeation volume method with a low-pressure membrane permeation system (HR-21, Gaorui, China), at a feed pressure of 0.5 bar and a temperature of 25 °C. The permeance of gas (${P}_{P}$) is calculated by Eq. (5):

$${P}_{P}=\frac{273}{76}\frac{V}{{AT}{P}_{0}}\frac{{{{{\rm{d}}}}}_{P}}{{d}_{t}}$$

(5)

where ${P}_{P}$ (cm³ cm⁻² s⁻¹ cmHg⁻¹) is the single gas permeance, $V$ (cm³) is the gas permeance, $T$ (K) is the temperature, $A$ (cm²) is the permeation area, ${P}_{0}$ and are the feed-side and permeate-side pressures, and $\frac{{d}_{P}}{{d}_{t}}$ (cmHg⁻¹ s⁻¹) is the rate of pressure increase on the permeate side.

Water vapor transmission rate

A modified WVTR measurement method was employed due to the sensitivity of hydrogel films to ambient humidity. The WVTR was determined by recording the mass change of water-filled glass bottles sealed with the organohydrogel nanofilm. Each glass bottle (with an opening diameter of 11.6 mm) was filled with 10 g of deionized water and then covered with the test nanofilm. The bottles were placed in a temperature- and humidity-controlled chamber maintained at 25 °C and 30% RH for a predetermined duration of 24 h for 7 days, after which the mass of each bottle was measured.

To distinguish water loss through the test hydrogel film from water exchange of the hydrogel itself with the environment, a reference (calibration) hydrogel sample with identical initial water content, thickness, and exposed area was tested under the same conditions. By comparing the test sample’s mass change to that of the reference sample, any mass change attributed to the organohydrogel’s inherent water exchange could be corrected, thereby isolating the true water vapor transmission.

The corrected WVTR was calculated according to Eq. (6):

$${WVTR}=\frac{{W}_{m}-{W}_{{ref}}}{A\,S}$$

(6)

where ${W}_{m}$, ${W}_{{ref}}$, $A$, and $S$ are the water loss weight, weight change of the reference sample, test duration, and area of the bottle’s opening, respectively.

Weight retention property

The initial weight of different samples was measured. The samples were then placed in a thermostatic chamber at 25 °C and 30% RH, and the weight change was recorded over time. The weight retention was calculated using the following formula (7):

$${Weight\; retention}=\frac{{m}_{t}}{{m}_{0}}\times 100\%$$

(7)

where ${m}_{0}$ is the weight of the sample at the initial time (day 0), and ${m}_{t}$ is the weight of samples at different time.

Electrophysiological monitoring

For continuous long-term ECG monitoring during daily activities, two organohydrogel nanofilm electrodes were placed on the left chest and connected to a commercial wireless module (GM3 Co., Japan). The setup and electrode placement are shown in Supplementary Fig. 38.

α wave EEG were recorded following the international 10−20 system. Commercial gel electrodes were applied to the subject’s ears as reference electrodes and to the forehead as the ground electrode. Organohydrogel nanofilm electrodes were placed on the right mastoid process for recording long-duration signals. ERP P300 signals were recorded at the same site. In the P300 test, high-frequency signals (“do”) accounted for 80% at 70 dB, and low-frequency signals (“di”) for 20% at 70 dB. All electrodes were connected to the signal acquisition system with a bandpass filter (0.02 to 50 Hz) and a sampling rate was 500 Hz.

EMG signals were recorded by a pair of organohydrogel nanofilm electrodes placed on the flexor carpi radialis muscles as the recording electrodes, with a commercial gel electrode on the back of the hand as the ground electrode. All electrodes were connected to the signal acquisition system through a band-pass filter (10 to 10⁴Hz). The EMG signals were generated by muscle contractions induced by a fixed grip force.

For MCV monitoring, a pair of organohydrogel nanofilm electrodes were placed on the abductor pollicis brevis muscle as recording electrodes. The electrodes were connected to the signal acquisition system through a band-pass filter (10 to 10³Hz). Signals were elicited by stimulating the median nerve with an amplitude of 6 mA and a frequency of 2 Hz.

For EOG monitoring, two organohydrogel nanofilm electrodes were placed horizontally at the outer eyebrow canthi, with two more positioned above and below the eyebrows, perpendicular to the eyeball. A ground electrode at the forehead completed the setup. Signals were filtered (1 to 200 Hz) and transmitted to the acquisition system.

Electrophysiological signal analysis

Resting state α wave EEG data were bandpass filtered (0.5 to 50 Hz) and plotted against time. The filtered α wave EEG data were then analyzed using the short-time Fourier transform (STFT) in MATLAB R2021b for PSD analysis. For ECG data, the SNR was calculated as the ratio of peak voltage to the standard deviation of the noise voltage.

Experiments on human subjects

All experiments involving electrophysiological signal recordings were thoroughly reviewed and approved by the ethical committee of Guangdong Technion-Israel Institute of Technology (approval number E20240902001). Informed consent was obtained from the subject participating in the experiment.

Statistics and reproducibility

All experiments were repeated independently with similar results for at least 3 times.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The data supporting the findings of this study are available within the article and its Supplementary Information/Source Data. Source data are provided with this paper.

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Acknowledgements

The authors sincerely acknowledge the support from the National Natural Science Foundation of China (grant Nos.: 52303371, Yan Wang; W2521021, Yan Wang), JSPS Bilateral Program (grant No.: JPJSBP120257419, Sunghoon Lee), JSPS Kakenhi (grant No.: JP22K21343, Sunghoon Lee), Guangdong Science and Technology Department (grant Nos.: STKJ2023075, Yan Wang; 2022A1515110209, Yan Wang; 2021B0301030005, Yan Wang), Guangdong Education Department (grant No.: 2022KQNCX112, Yan Wang), seed fund (grant No.: GCII-Seed-202406, Yan Wang) from GTIIT Changzhou Innovation Institute, and the Key Discipline (KD) Fund, the Technion, and the Start-Up Fund from Guangdong Technion.

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Department of Chemical Engineering, Guangdong Technion-Israel Institute of Technology, 241 Daxue Road, Shantou, Guangdong, China
Zonglei Wang, Yuli Wang, Jiawei Yang, Pengcheng Zhou, Wenqing Yan, Shihong Lin, Yujie Zhang, Qingyuan Sun, Zichong Ji, Mingzhe Wang, Zongman Zhang, Junhong Yi, Meiqiong Zheng, Xuezhong He & Yan Wang
The Wolfson Department of Chemical Engineering, Technion-Israel Institute of Technology, Haifa, Israel
Zonglei Wang, Yuli Wang, Jiawei Yang, Pengcheng Zhou, Wenqing Yan, Shihong Lin, Yujie Zhang, Qingyuan Sun, Zichong Ji, Zongman Zhang, Junhong Yi, Meiqiong Zheng, Xuezhong He, Hossam Haick & Yan Wang
Department of Materials Science and Engineering, Guangdong Technion-Israel Institute of Technology, 241 Daxue Road, Shantou, Guangdong, China
Yumiao Xu
Department of Materials Science and Engineering, Technion-Israel Institute of Technology, Haifa, Israel
Yumiao Xu
Guangdong Provincial Key Laboratory of Materials and Technologies for Energy Conversion, Guangdong Technion-Israel Institute of Technology, 241 Daxue Road, Shantou, Guangdong, China
Xuezhong He & Yan Wang
RIKEN Center for Emergent Matter Science (CEMS), 2-1 Hirosawa, Wako, Saitama, Japan
Lulu Sun, Sunghoon Lee & Takao Someya
Department of Electrical Engineering and Information Systems, The University of Tokyo, 7-3-1 Bunkyo-ku, Tokyo, Japan
Sunghoon Lee, Tomoyuki Yokota & Takao Someya

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Contributions

Yan Wang conceived and designed the experiments. Z.W., Y.W., J.Y., P.Z., Q.S., Y.X., L.S., S.L., and T.Y. fabricated the materials. Z.W., Y.W., Z.J., W.Y., S.L., Y.Z., M.W., and X.H. performed the experiments for material characterization. Z.W., Yuli Wang, Z.Z., Junhong Yi, and M.Z. performed the biosignal recording and data processing. L.S., Sunghoon Lee, T.Y., H.H., T.S., and Y.W. analysed the data and/or discussed the results. Z.W. and Yuli Wang wrote the manuscript. Yan Wang supervised the project. All authors discussed the results and commented on the manuscript.

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Correspondence to Yan Wang.

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Wang, Z., Wang, Y., Yang, J. et al. Sub-400 nanometer-thick skin and environment adaptable organohydrogel nanofilm epidermal electrode. Nat Commun 16, 10103 (2025). https://doi.org/10.1038/s41467-025-65089-0

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Received: 14 May 2025
Accepted: 06 October 2025
Published: 18 November 2025
Version of record: 18 November 2025
DOI: https://doi.org/10.1038/s41467-025-65089-0