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Frontiers in Biomaterials Science ·

Volume 5 - 2026 | https://doi.org/10.3389/fbiom.2026.1924191

Hemocompatible polymeric hydrogels with tunable photoresponsive volumetric properties

  • Akari J. Seiner1
  • Victor Mishin1
  • Shreya S. Soni1
  • Elisabeth Posthill1
  • Biplab Sarkar1
  • Kara L. Spiller1
  • Amy L. Throckmorton1,2
  • Christopher B. Rodell1,3*corresponding author
  1. 1School of Biomedical Engineering, Science and Health Systems, Drexel University, Philadelphia, PA, United States
  2. 2Department of Pediatrics, College of Medicine, Drexel University, Philadelphia, PA, United States
  3. 3Department of Microbiology and Immunology, College of Medicine, Drexel University, Philadelphia, PA, United States
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Abstract

Hydrogels exhibit crosslink density-dependent properties that can be tuned to achieve spatiotemporal control, making them well-suited for a wide range of biomedical applications. In particular, light reactive hydrogels have requisite features for use in size-changing biomaterial devices such as pediatric blood shunts, that can uniquely meet evolving patient-specific needs after in vivo implantation. Here, we report on the design of a tunable photoresponsive and hemocompatible methacrylated dextran hydrogel suitable for use in such devices. The formation and subsequent deswelling of hydrogels proceeded through discrete sequential photopolymerization steps. The range of geometric tunability (up to 20% volume change) was a function of methacrylate density (e.g., polymer modification and concentration) and the extent of initial polymerization that was controlled via exposure time, intensity, and photoinitiator concentration. Furthermore, the hydrogels exhibited long-term (>3 months) stability and hemocompatibility, including with blood-borne cell types (e.g., erythrocytes and leukocytes). In vitro verification demonstrated the feasibility of this concept; our elucidation of the parameter space governing hydrogel properties will facilitate the use of the platform in various blood-interface constructs, specifically pediatric blood shunts.

Keywords

1 Introduction

Hydrogels are water-swollen networks, well suited to a wide range of biomedical applications owing to their adjustable chemical and biophysical properties (Kim and Chu, 2000; Majee, 2016; Li et al., 2019; Xia et al., 2025). Hydrogels formed through conventional covalent crosslinking of hydrophilic polymer chains usually render polymeric networks with static properties (Park et al., 1993; Burdick and Murphy, 2012). In contrast, responsive hydrogel systems have transformed the field of soft materials from static to dynamic complexity (Burdick and Murphy, 2012; Li et al., 2019), as the attributes of these materials can be adjusted through various external triggers, such as light, temperature, pH, magnetic or electric fields, mechanical strain, enzymatic activity, or changes in ion concentrations (Kirschner and Anseth, 2013; Lim et al., 2014; Duan and Zhang, 2017; Ferreira et al., 2018; Liu et al., 2018; Aguilar and San Román, 2019; Matsuda et al., 2019). These stimuli-responsive hydrogels have been applied as controlled drug delivery vehicles (Hajebi et al., 2019; Abasalizadeh et al., 2020; Vigata et al., 2020; D'Elia et al., 2024), dynamic 3D cell culture matrices (Tibbitt and Anseth, 2009; Baruffaldi et al., 2021; Zhang et al., 2023), soft actuators (Yang et al., 2017; Han et al., 2018; Zhao et al., 2019), and chemical sensors (Brouns and Dankers, 2020; Gao et al., 2024).

Among responsive hydrogels, photoresponsive constructs are particularly attractive for biomedical applications (Tomatsu et al., 2011; Lee et al., 2018; Li et al., 2019; Chen et al., 2020; Li et al., 2020; Zhu et al., 2020; Pearson et al., 2021; Lin et al., 2022; Guo et al., 2025). Light stimulation allows for precise, user-defined spatial and temporal control with minimal additional reagents (Ge et al., 2021; Pearson et al., 2021; Günay et al., 2023; Kopyeva et al., 2025). Irradiation dosage, governing final material properties (Huang et al., 2020), may be altered via light intensity, exposure time, and wavelength (Truong et al., 2017; Peng et al., 2022). These parameters influence hydrogel formation, subsequent changes in functional group presentation or crosslink density, and controlled photodegradation—with corresponding alterations in properties (Vining et al., 2019; Raman et al., 2020). While these methodologies have mainly been used for the triggered release of drugs (Xing et al., 2022) or biomolecules (Wang et al., 2017; Jiang et al., 2020) as well as dynamic switching of cell-culture environments (Lee et al., 2018; Ruskowitz and DeForest, 2018), they may also be used to achieve physical changes (e.g., volumetric swelling or deswelling) associated with the polymer network’s crosslink density (Willner, 2017; Shi et al., 2021; Gao et al., 2024). The extent of hydrogel swelling depends upon both polymer hydrophilicity and crosslinking density, where increased crosslink density is typically associated with deswelling (i.e., exclusion of water).

Interestingly, the systematic investigation of volumetric responsiveness of photodynamic hydrogels has unexplored potential in medical device development. For example, the distinctive features of these materials open new avenues for individualized therapy—particularly in the emergent field of pediatric cardiovascular engineering—potentially enabling precise modulation of the size of implanted medical devices consistent with the growth of organs, addressing a critical challenge in this population (Feins et al., 2017; Hirschhorn et al., 2021). Recently, we have demonstrated a geometrically tunable blood shunt for use in pediatric heart-reconstruction surgeries as a proof-of-concept, wherein tunable properties are attributed to an internal photoresponsive hydrogel lining (Figure 1). For such shunts, computational fluid dynamic models revealed that a 15%–18% increase in lumen diameter, achievable by a 20% change in the volume of hydrogel coating the luminal surface, can accommodate the first 2 years of infant growth (Garven et al., 2022). These values are corroborated by a case study, where a dilatable shunt’s lumen diameter was expanded from 2.7 to 3.3 mm (18.2% expansion) to correct for hypoperfusion (Chikada et al., 2002). By enabling implantation of shunts with controllable lumen diameters, photoresponsive hydrogel linings may alleviate the need for repetitive open thoracotomy in these children, significantly improving clinical outcomes for patients with congenital heart defects, such as atrial/ventricular septal defects, hypoplastic left heart syndrome, and pulmonary atresia (Rao, 2019). In vivo, inner lumen diameter increase could feasibly be achieved via a blue light emitting catheter, providing a minimally invasive method to correct for hypoperfusion (Figure 1). To enable these applications, systematic characterization of the photoresponsive system is essential.

Schematic Overview. A geometrically tunable blood shunt consisting of a semi-rigid outer material (gray) to provide mechanical support and a blue light-responsive methacrylated dextran (DexMA) hydrogel lining (blue) to allow for inner lumen dilation.
Figure 1. Schematic Overview. A geometrically tunable blood shunt consisting of a semi-rigid outer material (gray) to provide mechanical support and a blue light-responsive methacrylated dextran (DexMA) hydrogel lining (blue) to allow for inner lumen dilation.

Elucidating how hydrogel composition and formation affect geometric tunability is crucial for designing size-changing biomedical implants. Moreover, the biological response to these materials must be comprehensively investigated, especially at tissue or blood interfaces (Weber et al., 2018). We have previously developed methacrylated dextran (DexMA) hydrogels with photoresponsive volumetric properties as a proof-of-concept (Garven et al., 2022). Building upon our initial proof-of-concept, the current work provides a comprehensive, systematic investigation of the formulation and processing parameters of photoresponsive DexMA hydrogels. Herein, we further evaluated feasibility of using DexMA hydrogels by investigating their physical, chemical, and biological responses. Hydrogels were formed and modified through two independent and sequential photopolymerization steps, allowing for initial hydrogel casting while preserving unreacted methacrylate groups, followed by a secondary cure to induce volumetric changes. We investigated the effects of polymer concentration, extent of methacrylate modification, and the extent of initial polymerization on changes in hydrogel volume. Furthermore, multiple cell types relevant to vascular applications were exposed to DexMA hydrogels to evaluate their cytocompatibility, hemocompatibility, and propensity to induce a deleterious innate immune response. In the current study, we systematically characterized photoresponsive DexMA hydrogels that may be used as blood-interface biomaterials, offering insights into the parameter space for geometric tunability and establishing their hemocompatibility. This study could pave the way for utilizing these materials in personalized and dynamically adaptable medical devices for children with complex congenital heart defects.

2 Experimental section

2.1 Materials

Unless otherwise stated, reagents and solvents were of a minimum ACS grade and purchased from Thermo Fisher Scientific (Waltham, Massachusetts), TCI America (Portland, Oregon), or Sigma-Aldrich (Burlington, Massachusetts).

2.2 Preparation of methacrylated dextran

DexMA was synthesized via the modification of dextran by glycidyl methacrylate (GMA; TCI America, M0590) through base-driven esterification (Figures 2A,B). A round bottom flask was charged with a stir bar, 1.0 g of dextran (Fisher; 75 kDa, AAJ6098918; or 500 kDa, AAJ6370218), and 100 mg of 4-dimethylaminopyridine (DMAP; TCI America, D145025G). The reaction vessel was purged by dry nitrogen prior to the addition of 30 mL anhydrous dimethyl sulfoxide (DMSO; Thomas Scientific, C990D20) and GMA. The molar ratio of GMA to repeat glucose units comprising dextran in the reaction was adjusted to control the degree of substitution (DS = 5%, 0.25 eq.; 25%, 0.5eq.; 50%, 1 eq.; 75%, 1.5 eq.; 95%, 2 eq.). The DS, or methacrylate modification, directly impacts the properties of formed hydrogel, including light-responsive behavior and moduli (Reis et al., 2009). The reaction was allowed to proceed for 24 h at 45 °C with subsequent dialysis (6–8 kDa MWCO; Fisher, 08670D) against deionized (DI) water for 1 week with water changes twice daily. Products were frozen at −80 °C and lyophilized. The DS was determined using 500 MHz 1H NMR spectra of polymer samples in D2O (Supplementary Figure S1), by integrating the vinyl proton (δ = 5.55–5.85 ppm, 1H) relative to the anomeric proton of dextran (δ = 4.8–5.05 ppm, 1H); the DS was further confirmed by the integral of the methyl singlet (δ = 1.6–2.0 ppm, 3H). To assess integration error, results were blindly analyzed by three individuals and data are reported as mean ± s.d.

Synthesis of methacrylated dextran (DexMA). **(A,B)** DexMA was prepared by the base-catalyzed esterification of dextran with glycidyl methacrylate (GMA). **(C)** The degree of substitution was linearly dependent (R<sup>2</sup> = 0.97) on the molar feed ratio of GMA to glucose residues comprising dextran.
Figure 2. Synthesis of methacrylated dextran (DexMA). (A,B) DexMA was prepared by the base-catalyzed esterification of dextran with glycidyl methacrylate (GMA). (C) The degree of substitution was linearly dependent (R2 = 0.97) on the molar feed ratio of GMA to glucose residues comprising dextran.

2.3 Hydrogel fabrication and mechanical characterization

DexMA was dissolved in phosphate buffered saline (PBS; VWR, VWRL0119-0500) containing the photoinitiator lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP; Sigma-Aldrich, 900,889-1G). Rheological assessment was performed using a Discovery HR20 rheometer (TA Instruments, New Castle, Delaware) fitted with a sand-blasted 20 mm parallel-plate geometry at a 200 µm gap to investigate the effect of LAP concentration, light intensity, and light exposure time on photopolymerization kinetics. Real-time photopolymerization was monitored using a light curing stage (Omnicure S2000, λ = 400–500 nm, 1–10 mW/cm2) throughout oscillatory time sweeps (1% strain, 5 Hz). Oscillatory frequency (0.1–100Hz, 1% strain) and strain (1.0 Hz, 0.01%–500% strain) sweeps were then performed to ensure measurements were conducted within the plateau region.

A dual-stage photopolymerization scheme consisting of two independent light exposures was developed to i) cast a solid hydrogel while preserving unpolymerized methacrylate groups and ii) subsequently cure the hydrogel via secondary light exposure after overnight equilibrium swelling to achieve volumetric changes (Figure 3A). Samples of varying polymer modifications (5%, 25%, 50%, 75%, and 95%) and concentrations (2.5%, 5%, 10%, and 20%w/v) were pipetted into cylindrical molds and sealed to minimize oxygen quenching during photopolymerization. Hydrogel casting (10–60 s) and secondary curing (60–300 s) steps were performed (Omnicure S2000, λ = 400–500 nm) with calibrated light intensity (1 mW/cm2; Sper Scientific, 850,009) while incubated with fresh LAP to ensure standardized photoinitator concentration within the network during light exposure. Compressive moduli were assessed after equilibrium swelling through unconfined compression (5 μm/s) and quantified as the slope of the stress–strain curve at 20% strain for all samples (n = 3 per group).

Photopolymerization of DexMA. **(A)** Schematic of hydrogel formation via 1) initial casting to form a solid, steady state hydrogel and 2) subsequent secondary curing to enable tuning of hydrogel properties as a result of deswelling due to crosslinking and the formation of a kinetic chain. **(B)** Rheological assessment of the dual-stage crosslinking scheme, characterized by storage and loss modulus. Oscillatory time sweeps (5 Hz, 1% strain) throughout dual stage photopolymerization (shaded blue: 20 s casting (left) and 300 s curing (right) at 1 mW/cm<sup>2</sup>) for hydrogel formation (50% DexMA, 10%<sub>w/v</sub>, 1 mM LAP).
Figure 3. Photopolymerization of DexMA. (A) Schematic of hydrogel formation via 1) initial casting to form a solid, steady state hydrogel and 2) subsequent secondary curing to enable tuning of hydrogel properties as a result of deswelling due to crosslinking and the formation of a kinetic chain. (B) Rheological assessment of the dual-stage crosslinking scheme, characterized by storage and loss modulus. Oscillatory time sweeps (5 Hz, 1% strain) throughout dual stage photopolymerization (shaded blue: 20 s casting (left) and 300 s curing (right) at 1 mW/cm2) for hydrogel formation (50% DexMA, 10%w/v, 1 mM LAP).

The impact of hydrogel composition and photopolymerization conditions on the volume of the hydrogel was assessed in cylindrical hydrogels (n > 3 per group). Samples with different polymer concentrations (2.5%, 5%, 10%, and 20%w/v) and methacrylate modifications (5%, 25%, 50%, 75% and 95%) were prepared as previously described. Photoinitiator concentration (1–10 mM LAP) was altered in either the initial casting (20 s) or curing (60, 300, and 600 s) process after overnight swelling. Hydrogel contraction occurred as a result of curing, corresponding to a desired increase in inner lumen diameter, the absolute change in volume from post-cast to post-cure condition was measured using a digital caliper (V = πr2h) and calculated as Δ V = C u r e − C a s t C u r e * 100.

2.4 DexMA hydrogel stability under physiological conditions

The stability of DexMA hydrogels was evaluated through the incorporation of fluorescein O-methacrylate (Sigma-Aldrich, 568864) at a concentration of 100 ng/mL during casting. Hydrogels were prepared using varying polymer concentrations (2.5%, 5%, and 10%w/v), modifications (25%, 50%, and 95%), and molecular weights (75 or 500 kDa) of the polymer (n = 4 hydrogels per group). Once cast, hydrogels were washed by excess PBS (3 washes, 2 h each) to remove unbound fluorophore. Hydrogels were subsequently incubated at 37 °C in 1 mL PBS that was collected at set timepoints (with replacement) over 3 months. To assess hydrogel swelling, mass and volume were measured weekly in parallel. Polymer degradation was assessed by sample fluorescence (BioTek Synergy H1 plate reader; λex/em = 490/518 nm) relative to standard curves. Degradation data are presented following normalization to cumulative degradation after enzymatic digestion of the remaining hydrogel by 0.2 mg/mL dextranase (Sigma, D5884-5KU); swelling data were normalized to baseline values for each sample.

2.5 In vitro evaluation of hydrogel cytocompatibility

Cells were maintained under normal culture conditions (37 °C, 5% CO2) for all studies and conducted in a minimum of triplicate. NIH 3T3 fibroblasts (Fisher) were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM, VWR, VWRL0101-500) supplemented with 10% fetal bovine serum (FBS, VWR, 97068-085) and 1% penicillin-streptomycin (100X, VWR, 970063-708). Cells were seeded at 5 × 103 cells/well in a 96-well plate on hydrogel surface (direct contact) or in conditioned media following incubation with hydrogels (hydrogel extracts). Cell proliferation was monitored over 3 days in culture using PrestoBlue assay (Fisher, A13261) following the manufacturer’s protocols, and data were normalized relative to untreated cells at baseline. Cell proliferation was evaluated in the presence of cast hydrogels of various polymer modifications (25%, 50%, and 95%) at a concentration of 10%w/v after equilibrium swelling, as well as in media conditioned by incubation with cured DexMA hydrogels for 24 h (extracts). To investigate the effect of soluble hydrogel components, cells were cultured in media containing DexMA at the prescribed concentration and modification.

RAW-Blue™ macrophages (ATCC, TIB-71) were cultured in DMEM supplemented with 10% heat-inactivated fetal bovine serum (FBS, Fisher, Corning™ 35010CV), 1% penicillin-streptomycin (VWR, 970063-708), and 0.2% Normocin™ (50 mg/mL, InvivoGen, ant-nr-1) to maintain selection pressure. To investigate the innate immune response towards various hydrogel formulations (25%, 50%, and 95% polymer modifications; 2.5%, 5%, and 10%w/v; n = 3 per group) after overnight equilibrium swelling, cells were seeded in unmodified polystyrene wells, directly on the surface of hydrogels, or with conditioned media from hydrogels, at 25 × 103 cells/well and incubated for 24 h at 37 °C. Cell activation was quantified by QUANTI-Blue™ assay (QB; InvivoGen, rep-qbs) according to the manufacturer’s protocol. Briefly, 1 mL of QB reagent and 1 mL of QB buffer were diluted in 98 mL MilliQ water. For each sample, 180 µL of the QB solution was mixed with 20 µL media, incubated for 3 h at 37 °C, and the absorbance value was measured at 620 nm. Data were normalized relative to positive inflammatory controls (100 ng/mL liposaccharides, LPS; Fisher, 501657334). To quantify adhesion, cells were fixed with 4% paraformaldehyde (Electron Microscopy Sciences, 15710-S) for 15 min and washed with PBS. Cell nuclei were stained with DAPI (NucBlue™, Fisher, R37606). The nuclei of adhered cells in each image were counted using fluorescent images (×10 magnification).

2.6 ASTM-compliant hemolysis assessment

Procedures followed ASTM standards F756-13: Standard Practice for Assessment of Hemolytic Properties of Materials and F619-03: Standard Practice for Extraction of Medical Plastics. Hemolysis of citrated whole rabbit blood (Thomas Scientific, R209-0050) in response to hydrogels (25%, 50%, and 95% polymer modifications, each at 10%w/v, direct contact) and their extracts were assessed after initial equilibrium swelling (hydrogel extracts). Plasma free hemoglobin was assessed prior to blood use, and the analysis is presented relative to recommended negative (low-density polyethylene, LDPE; McMaster-Carr, 8657K111) and positive (Buna-N Rubber; McMaster-Carr, 8635K162) controls. Sample absorbance values were measured (Thermo BioMate 3 spectrophotometer, λ = 540 nm) with appropriate background subtraction to corresponding extracts and whole blood dilutions to correct for background absorbance of the material and free hemoglobin, respectively. The percentage of hemolysis was then determined as the supernatant hemoglobin concentration divided by the total hemoglobin concentration, accounting for dilutions. Studies were conducted in triplicate, with two independent biological repeats.

2.7 Functionalization of quartz tubing for hydrogel integration

Surface modification of 6 mm ID quartz glass tubing (McMaster-Carr, 4100N422) was achieved using 3-(trimethoxysilyl)propyl methacrylate 98% (TMS; Millipore Sigma, C974V12). Briefly, the tubing was immersed in 10 M NaOH for 20 min, rinsed three times with Milli-Q water, and allowed to air dry. TMS was applied dropwise to tubing for coating, and the tubing was subsequently heated (100 °C, 1 h followed by 120 °C, 10 min). Samples were washed with ethanol and Milli-Q water, and subsequently dried. Hydrogels (n = 4) were loaded into the modified glass tubing and cured (1 mW/cm2, 20 s). After overnight equilibrium swelling, a 3.5 mm biopsy punch was used to create an inner lumen, measured using digital calipers upon 10 s incremental exposure via a cylindrical light diffuser (Medlight, ML30-2292) while incubated with LAP.

2.8 Presentation of data and statistical analysis

Data are presented as mean ± standard deviation (s.d.), unless otherwise noted. All statistical analyses were performed using GraphPad Prism v9.5.0. Analysis of variance (ANOVA) with post-hoc Tukey Honestly Significant Difference (HSD) test was used to determine statistical significance (p < 0.05) for multiple comparisons.

3 Results and discussion

3.1 Synthesis and characterization of methacrylated dextran (DexMA)

Physical or chemical properties of photoresponsive hydrogels change in response to irradiation (Lee et al., 2018; Chen et al., 2020; Li et al., 2020; Lin et al., 2022), and such materials are potentially useful for biomedical applications (Tomatsu et al., 2011). Specifically, dextran is a biocompatible polysaccharide comprised primarily of α-1,6-linked D-glucose residues, each containing three secondary hydroxyl groups that may be leveraged for modification (Kim and Chu, 2000). The polymer has been applied as a drug carrier or plasma expander because of its biocompatibility and high water solubility, and it also has well documented anti-thrombogenic properties, making it well suited for blood-contacting applications (Won and Chu, 1998).

Direct attachment of pendant functional groups to biopolymers yields polymers with controlled extent of modification. The modification of dextran by pendant methacrylate groups has previously been accomplished by esterification with methacrylic anhydride, either in anhydrous or aqueous conditions (Kim and Chu, 2000; Khetan et al., 2013). Here, we utilize esterification with glycidyl methacrylate (GMA (van Dijk-Wolthuis et al., 1995; Trappmann et al., 2017)), as the reaction proceeds under homogenous conditions and may be easily adjusted to control the extent of polymer modification (Figures 2A,B). Based on the hypothesis that the volumetric tunability of the material depends on the number of available methacrylate groups, polymer samples with various extents of modification were prepared by altering the initial GMA concentration. The chemical structure of DexMA was confirmed using 1H NMR (Supplementary Figure S1); the degree of substitution (DS) ranged from 5% to 95%, with linear dependence on GMA concentration (Figure 2C).

3.2 Development of a two-stage photocrosslinking strategy for tunable hydrogels

Multi-step hydrogel crosslinking approaches to modulate material properties have been previously investigated by several groups (Rizwan et al., 2017a; Zhu et al., 2017; González-Henríquez et al., 2019; Rebers et al., 2021). Most frequently, increases in crosslink density are used to achieve changes in mechanical properties. Examples include the creation of injectable hydrogels with subsequent stiffening (Dong et al., 2015; Rodell et al., 2015; Zhu et al., 2017), development of tunable cell-culture environments (Rodell et al., 2016; Vining et al., 2019), and increasing DexMA fiber stiffness (Davidson et al., 2019). While mechanical tuning has been extensively studied, the potential for volumetric responsiveness–particularly in geometrically dynamic environments–remains unexplored. Here, both initial hydrogel formation and subsequent tuning of hydrogel properties are accomplished via photopolymerization using the photoinitiator LAP within a concentration range of 0.1–10 mM, which exhibits cytocompatibility and water solubility (Fairbanks et al., 2009; Caliari et al., 2016). We developed a two-step photocrosslinking method that includes 1) initial casting to form a solid hydrogel, while intentionally retaining unreacted methacrylate groups for 2) subsequent curing to refine hydrogel properties and allow for hydrogel deswelling and contraction, thereby corresponding to an increase in inner lumen diameter of hydrogel-lined vessels in vivo (Figures 1, 3A).

To determine appropriate conditions for the initial casting step that preserve methacrylate groups, the photopolymerization reaction was monitored in real time by rheology (Supplementary Figures S2A,B). Photoinitiator concentration, light intensity, and the duration of light exposure each contributed to the rate of hydrogel formation. Dissolved DexMA was initially a moderately viscous solution (G′ < 1 kPa, tan(δ) > 1). Upon light exposure, a sharp increase in G′ and G″ was observed, with an eventual plateau occurring within 5 min of light exposure for all conditions, indicative of complete polymerization through the consumption of available methacrylate groups (De Smedt et al., 1995). Both high photoinitiator concentrations (3, 5 mM) and light intensities (5, 10 mW/cm2) produced higher final storage moduli; however, photopolymerization was also completed more rapidly (Supplementary Figures S2A,B). Under such conditions, hydrogels would have a limited capacity to undergo further crosslinking during the secondary curing phase. Therefore, the lowest photoinitiator concentration (1 mM) and light intensity (1 mW/cm2) were used throughout subsequent experiments.

Oscillatory rheology was used to investigate hydrogel response to a secondary crosslinking event. The two-step method consisted of an initial exposure of 20 s to form a solid hydrogel (G′ = 1 kPa). Notably, a mild degree of crosslinking persisted following the end of the irradiation period; this is likely attributable to radical diffusion and resulting kinetic chain elongation. Following the stabilization of hydrogel moduli, a secondary light exposure of 300 s induced further crosslinking and resulted in hydrogel deswelling (Kim and Chu, 2000; Lo and Jiang, 2010), needed for the expansion of the inner lumen diameter of hydrogel-lined vessels in vivo. Final hydrogel moduli plateaued at approximately 100 kPa (Figure 3B). The method developed here intentionally prevents complete consumption of the available vinyl groups during the initial casting phase, perpetuating drastic changes in hydrogel properties during subsequent curing events triggered by light exposure.

To further explore the range of hydrogel tunability achievable through this approach and its dependence on hydrogel formulation, the compressive modulus of hydrogels was assessed both after casting and curing (Supplementary Figures S3A,B). Initial hydrogel moduli post-casting were considerably dependent on the polymer concentration (E = 3.9 and 374 kPa for 5% and 20%w/v, respectively, Supplementary Figure S3A) but were relatively independent of polymer modification at ≥ 25% functionalization (E = 45–50.7 kPa, Supplementary Figure S3B). Across all groups examined, curing led to a drastic increase in modulus, indicative of secondary polymerization. An increase in polymer concentration led to a more drastic change in moduli, most likely due to the availability of more reactive methacrylate groups that can form kinetic chains (Supplementary Figure S3A). Interestingly, final moduli of the cured hydrogels decreased for polymer modifications exceeding 50% (Supplementary Figure S3B), despite the increased concentration of vinyl groups, at a fixed polymer concentration of 10%w/v. Given the hydrophobicity of highly methacrylated dextran, this finding is likely attributable to a decrease in the polymer’s effective hydrodynamic radius and related polymer entanglement (Miquelard-Garnier et al., 2006; Trappmann et al., 2017; Zhan et al., 2021).

3.3 Geometric tunability and volumetric responsiveness

In addition to the modulation of mechanical properties, responsive hydrogels can undergo volumetric changes in response to external stimuli. Hydrogels can exhibit this change in volume by swelling or deswelling (Holback et al., 2011), enabling applications such as the spatiotemporal control of hydrogel topography in tissue culture platforms (Guvendiren et al., 2009; Zawko et al., 2009; Ramanan et al., 2010), and controlled drug release (Park, 1988; Kim et al., 1992; Rizwan et al., 2017b). Here, geometric tunability, representing hydrogel deswelling, was expected to depend heavily on the number of methacrylate groups remaining at the start of the secondary photopolymerization step. The extent of geometric tunability is expected to be directly dependent on hydrogel formulation, as the initial concentration of methacrylate groups is controlled by the polymer concentration and modification. Furthermore, the extent of photopolymerization during casting impacts the availability of methacrylate groups remaining for the secondary reaction. The polymer concentration, modification, and photopolymerization conditions (LAP concentration and exposure time) were therefore systematically investigated to quantify the decrease in DexMA hydrogel volume upon blue light exposure. For the initial casting, increasing LAP concentrations impeded later volume change (reported as absolute values), with 1 mM LAP achieving significantly greater secondary volume change than either 3 or 5 mM LAP (Figure 4A, p < 0.0001), with no significant differences reported between 3 and 5 mM LAP (p = 0.7623). These results overall agree with the rationale of our two-stage polymerization approach: retention of unreacted vinyl groups is essential to subsequent geometric tunability; however, further work would be needed to characterize this hypothesized methacrylate conversion. Furthermore, an increase in photoinitiator concentration for the secondary light exposure contributed to moderately greater volume changes (Figure 4B, not statistically significant, p > 0.2287).

Effect of polymerization conditions on geometric tuning. **(A,B)** Absolute volume change of hydrogels (50% DexMA, 10%<sub>w/v</sub>) between initial casting (20 s) and curing (300 s), each at 1 mW/cm<sup>2</sup>. LAP concentration was altered during casting (A: 1–5 mM) or curing (B: 1–10 mM). **(C)** Absolute volume change of hydrogels (50% DexMA, 10%<sub>w/v</sub>) to varying cure times (60–600s, 3 mM LAP, 1 mW/cm<sup>2</sup>). Absolute volumetric changes at varying **(D)** polymer concentrations (50% DexMA, 1 mW/cm<sup>2</sup>, 20 + 300 s exposure) and **(E)** modifications (10%<sub>w/v</sub> DexMA, 1 mW/cm<sup>2</sup>, 20 + 300 s exposure time). Data represent mean ± s.d.; n ≥ 3 independent samples. **P* < 0.05, ***P* < 0.01, *****P* < 0.0001 (ANOVA, Tukey HSD). No comparison is shown if statistically insignificant.
Figure 4. Effect of polymerization conditions on geometric tuning. (A,B) Absolute volume change of hydrogels (50% DexMA, 10%w/v) between initial casting (20 s) and curing (300 s), each at 1 mW/cm2. LAP concentration was altered during casting (A: 1–5 mM) or curing (B: 1–10 mM). (C) Absolute volume change of hydrogels (50% DexMA, 10%w/v) to varying cure times (60–600s, 3 mM LAP, 1 mW/cm2). Absolute volumetric changes at varying (D) polymer concentrations (50% DexMA, 1 mW/cm2, 20 + 300 s exposure) and (E) modifications (10%w/v DexMA, 1 mW/cm2, 20 + 300 s exposure time). Data represent mean ± s.d.; n ≥ 3 independent samples. *P < 0.05, **P < 0.01, ****P < 0.0001 (ANOVA, Tukey HSD). No comparison is shown if statistically insignificant.

Taken together, these results suggest that lower initiator concentrations during casting and higher initiator concentrations during secondary curing may be combined to achieve the greatest range of volumetric change. As observed rheologically for hydrogel moduli, geometric tunability should also be directly dependent on the duration of light exposure during curing. Direct and significant differences were observed between 60 s, 300 s, and 600 s cure times (Figure 4C, p < 0.0198), with no statistically significant difference between 300 s and 600 s curing (p = 0.7782). This result demonstrates that curing is more highly dependent on the time of light exposure than initiator concentration. Hence, not only can the process be performed at cytocompatible initiator concentrations, the degree of geometric tunability can also be temporally controlled.

To further examine the impact of hydrogel formulation, which affects initial methacrylate availability, a range of DexMA concentrations (2.5–20%w/v) and modifications (5%–95%) were tested in combination. For all polymer concentrations, an increase in absolute volumetric change was associated with an increase in polymer concentration (Figure 4D; Supplementary Figures S4A,B), with some comparisons showing statistical significance (p < 0.0498). However, the behavior was more complex for varying DexMA modifications. Volume changes exhibited a maximum at an intermediate DS of 25%–50% (Figure 4E; Supplementary Figures S2C,D), dependent on polymer concentration. These results correspond directly to those of compressive moduli (Supplementary Figure S3). As discussed, these observations could be attributable to elevated DexMA hydrophobicity at high levels of methacrylation.

Full characterization of hydrogel properties through mechanical and geometric assessment revealed that a combination of high DexMA concentration and intermediate modification is the best candidate for obtaining geometrically tunable hydrogels. An initial short casting of 20 s at low initiator concentrations (1 mM) allowed for the preservation of functional groups to achieve the greatest volume change during the longer 300 s curing at 5 mM LAP, each at 1 mW/cm2. Under these conditions, volume changes of 20.4% ± 0.79% were observed, meeting the needs for geometrically tunable cardiovascular shunts (Garven et al., 2022), and perpetuating their use in alternative applications such as cell culture platforms or size-changing implants.

3.4 Long-term hydrolytic stability of DexMA hydrogels

Dextran itself is moderately resistant to degradation, owing to the hydrolytic stability of glycosidic linkages and a lack of specific enzymes to degrade it in vivo (van Dijk-Wolthuis et al., 1997). The swelling and degradation behavior of DexMA hydrogels is therefore highly dependent on the crosslink density and stability (van Dijk-Wolthuis et al., 1997; Annu and Ahmed, 2021). For this study, the long-term stability of DexMA hydrogels of various polymer concentrations, modifications, and molecular weights was investigated over the course of 3 months. All hydrogels showed exceptional stability with respect to hydrolysis. Across all groups, less than 10% cumulative degradation was observed throughout the course of the study (Figure 5A; Supplementary Figures S5A–C). This finding was further confirmed by assessing hydrogel swelling. Hydrogels exhibited less than 5% volume change over the course of study (Figure 5B; Supplementary Figures S5D–F). These results are corroborated by mass losses, with 10%w/v hydrogels exhibiting less than 5% loss in 3 months and moderate mass losses (nearing 25%) noted only in the 2.5%w/v groups (Figure 5C; Supplementary Figures S5G–I). Altering the molecular weight of dextran had no impact on these outcomes (Supplementary Figures S6A–C). In summary, the hydrogel properties (structural and volumetric integrity) outlined above are not compromised by hydrogel degradation, even over the long term.

Hydrogel degradation and swelling. Hydrogel (10%<sub>w/v</sub>) stability was assessed over 90 days, including **(A)** cumulative degradation, **(B)** hydrogel volume, **(C)** and mass. Data represent mean ± s.d.; n = 4 independent samples. Shaded regions represent error bars.
Figure 5. Hydrogel degradation and swelling. Hydrogel (10%w/v) stability was assessed over 90 days, including (A) cumulative degradation, (B) hydrogel volume, (C) and mass. Data represent mean ± s.d.; n = 4 independent samples. Shaded regions represent error bars.

3.5 Cytocompatibility and immunogenic assessment

Following the implantation of vascular prosthetics or other biomedical implants, direct material interaction with a number of cell types can be expected. These include, for example, fibroblasts and endothelial cells that are abundant in vascular tissues, as well as erythrocytes and circulating immune cells (Weber et al., 2018). We therefore evaluated the cytocompatibility of formed hydrogels and their components. Throughout the course of 3 days of cell culture, lower DexMA modifications (25% and 50%) did not hinder cell proliferation either in direct hydrogel contact or in hydrogel extracts (Figures 6A,B).

Cytocompatibility profile of DexMA hydrogels. Proliferation of 3T3 fibroblasts cultured in **(A)** direct contact with cast DexMA hydrogels or with **(B)** hydrogel extracts. **(C)** Macrophage adhesion to DexMA hydrogels. Untreated control wells (tissue culture treated polystyrene) are included for reference, shown in red. **(D)** Inflammatory activation of RAW264.7 cells cultured on DexMA hydrogels after initial cast only at varying polymer modifications and concentrations. Inflammatory (LPS treated, red dashed line) and untreated controls (green dashed line) are included for reference. Data represent mean ± s.d.; n = 3 independent samples. Hemolysis was evaluated for **(E)** direct material contact and **(F)** extracts for hydrogels of polymer modifications (25%, 50%, and 95%) and light exposure conditions (cast, cast + cure). Data represents mean ± s.d. of two biological repeats; n = 3 per study. All groups were significantly different than hemolytic controls (Buna-N, P < 0.0041) but did not differ from non-hemolytic controls (LDPE, P > 0.9905). **P* < 0.05, ***P* < 0.01, *****P* < 0.0001 (ANOVA, Tukey HSD). No comparison is shown if statistically insignificant.
Figure 6. Cytocompatibility profile of DexMA hydrogels. Proliferation of 3T3 fibroblasts cultured in (A) direct contact with cast DexMA hydrogels or with (B) hydrogel extracts. (C) Macrophage adhesion to DexMA hydrogels. Untreated control wells (tissue culture treated polystyrene) are included for reference, shown in red. (D) Inflammatory activation of RAW264.7 cells cultured on DexMA hydrogels after initial cast only at varying polymer modifications and concentrations. Inflammatory (LPS treated, red dashed line) and untreated controls (green dashed line) are included for reference. Data represent mean ± s.d.; n = 3 independent samples. Hemolysis was evaluated for (E) direct material contact and (F) extracts for hydrogels of polymer modifications (25%, 50%, and 95%) and light exposure conditions (cast, cast + cure). Data represents mean ± s.d. of two biological repeats; n = 3 per study. All groups were significantly different than hemolytic controls (Buna-N, P < 0.0041) but did not differ from non-hemolytic controls (LDPE, P > 0.9905). *P < 0.05, **P < 0.01, ****P < 0.0001 (ANOVA, Tukey HSD). No comparison is shown if statistically insignificant.

Population doubling times were in agreement with those typically reported (ca. 20 h) (Horbett et al., 1985). At the highest modification (95%), substantial declines in metabolic activity were noted (Figures 6A,B; Supplementary Figure S7A). While hydrophobic surfaces may negatively impact cell adhesion, proliferation, and viability (Chen et al., 2018), similar effects were observed for both direct contact and hydrogel extracts. We therefore expected that soluble polymer may impact cytocompatibility, particularly for highly modified and hydrophobic DexMA samples. At high polymer concentrations (5%w/v) in solution, a trend toward reduced proliferation rates was observed to correlate with the DS (Supplementary Figure S7A). Thus, polymer release from formed hydrogels may contribute to reduced cell proliferation at higher modification. However, it is important to consider that the concentration of free soluble polymer investigated here far exceeds what can be released from the pre-cast photopolymerized hydrogel. Further scrutiny of concentration dependence showed that cell proliferation proceeded at reduced polymer concentrations, even for 95% DexMA (Supplementary Figure S7B). Although highly modified soluble DexMA may hinder cell proliferation at high concentrations, cell growth proceeds normally in the presence of formed hydrogels at low and intermediate modifications, even in the absence of extensive washing to remove unbound DexMA. Hence, it may be judicious to avoid highly modified dextran-based hydrogels both in cell culture systems and implantable devices.

The provocation of an innate immune response is a risk factor for implantable biomedical devices. Because of the blood-contacting environment of vascular prosthetics, it is critical to investigate the potential inflammatory response of immune cells, including abundant monocytes and macrophages. Here, we directly evaluated macrophage adhesion and activation that mediate the foreign body response and fibrous encapsulation of implanted devices (Mariani et al., 2019). The effect of substrate stiffness on macrophage adhesion and phenotype may be context dependent (McWhorter et al., 2015). According to some studies, RAW264.7 cells adhere to stiffer hydrogels more robustly; however, others have found no difference (Irwin et al., 2008; Nemir et al., 2010). We therefore investigated both cast and cured DexMA hydrogels. Nuclear staining of adhered cells indicated that DexMA was relatively resistant to cell adhesion when compared to a polystyrene control (Figure 6C; Supplementary Figure S8A). We noted a tendency for a reduction in cell adhesion for increased polymer modifications, concentrations, and after curing.

Inflammatory pathway activation was further assessed using the RAW-Blue™ reporter cell line that incorporates a SEAP reporter construct downstream of AP-1 and NF-κB promoter regions. We assessed pro-inflammatory activation relative to positive (LPS treated) and negative (untreated) controls. Macrophage activation following 24 h of culture in direct contact with the material was minimal across all groups (Figure 6D; Supplementary Figure S8B), with a moderate tendency for secondary curing to increase activation. Taken together, the results indicate that DexMA hydrogels are resistant to cell adhesion and do not provoke RAW 264.7 macrophage inflammation under static culture conditions, regardless of hydrogel formulation or the extent of crosslinking.

3.6 DexMA hydrogel hemocompatibility

The hemolytic potential of blood-contacting biomaterials is an important factor to consider, particularly for the development of cardiovascular devices. Our previous study on 95% modified DexMA (using ASTM standards) demonstrated that DexMA is non-hemolytic (Sovadinova et al., 2011; Garven et al., 2022). However, it is critical to ensure this property across all polymer concentrations, modifications, and crosslinking conditions used in this study, as factors including surface hydrophobicity are known to affect hemolysis (Kuroda et al., 2009; Sovadinova et al., 2011; Tong et al., 2020). Here, extracts and direct material contact of 25%, 50%, and 95% DexMA at: 1) 20 s of initial casting light exposure and 2) 20 s and 300 s of secondary curing light exposure were investigated (Figures 6E,F). All hydrogels tested exhibited an average hemolytic index of less than 3%-significantly less than the positive control (Buna-N rubber), comparable to the negative control (LDPE), and within the standard’s recommended hemolytic threshold of 5%. It is important to note that given dextran’s documented hemocompatibility, thrombogenicity was not evaluated as part of this study. Ultimately, regardless of polymer modification, DexMA hydrogels are suitable for blood-contacting applications according to prevailing industry benchmarks.

3.7 In vitro validation of geometric tunable shunt prototype

The ability to precisely modulate shunt lumen diameter over time may significantly enhance clinical outcomes in pediatric patients by accommodating physiological growth. Previous computational modeling has indicated that a 15%–18% increase in inner lumen diameter may accommodate infant growth over 2 years (Garven et al., 2022). It is critical to ensure that this range of volumetric tunability is achievable using an in vitro model of photocrosslinked hydrogel-lined shunt prototype. Here, a TMS-coated glass tubing was loaded with 50%-modified hydrogels with an initial inner lumen diameter of 3.5 mm and incrementally exposed to 10 s radiation pulses, with corresponding inner lumen diameter measurements (Figure 7). The requisite 15%–18% inner lumen diameter increase was achieved within 50–70 s, indicating promising suitability for future studies.

Cardiovascular shunt proof of concept investigating inner lumen diameter response to incremental blue light exposure. TMS coated glass tubing (8 mm OD, 6 mm ID, n = 4) lined with DexMA hydrogels underwent 10 s incremental exposure to blue light via catheterization starting at an initial and final inner lumen diameter of 3.5 and 4.1 mm, respectively. The required inner lumen diameter change of 15%–18% needed to accommodate for infant growth was achieved within 50–70 s.
Figure 7. Cardiovascular shunt proof of concept investigating inner lumen diameter response to incremental blue light exposure. TMS coated glass tubing (8 mm OD, 6 mm ID, n = 4) lined with DexMA hydrogels underwent 10 s incremental exposure to blue light via catheterization starting at an initial and final inner lumen diameter of 3.5 and 4.1 mm, respectively. The required inner lumen diameter change of 15%–18% needed to accommodate for infant growth was achieved within 50–70 s.

A key consideration for the clinical translation of delayed, post-implantation photoactivation is photoinitator retention. Because LAP is a small, hydrophilic molecule, passive retention within the hydrogel matrix under physiological blood flow conditions may not be assumed over extended durations. While the present study demonstrates the parametric evaluation of secondary volumetric contraction in relation to geometric tunability, clinical implementation will necessitate local, catheter-based delivery of photoinitiator prior to or during irradiation.

4 Conclusion

Here, we report a stepwise strategy to effectively tune hydrogel volume through sequential photopolymerization reactions, where increasing crosslink density promotes deswelling. The initial photopolymerization step preserves unreacted methacrylate groups, enabling volumetric changes during the secondary polymerization step. Both geometry and associated material properties of the hydrogel network were dependent on the extent of biopolymer modification by methacrylate groups and the initial polymer concentration. However, these properties were not dependent on the concentration of functional groups alone, as intermediate polymer modifications displayed the widest range of both geometric and mechanical tunability. Moreover, the range of tunability was fundamentally limited by the initial polymerization reaction used for hydrogel casting. The use of high initial photoinitiator concentrations, light intensity, and exposure times during casting all contributed to terminal consumption of reactive groups, thereby limiting subsequent physiochemical changes. Conversely, geometric tunability during latter curing was moderately improved by increasing photoinitiator concentrations (keeping it within the established range of cytocompatibility). Volume changes were temporally dependent; therefore, hydrogel geometry could be precisely tuned as a function of exposure time. Regardless of formulation, hydrogels were resilient towards hydrolytic degradation and maintained their initial polymeric content and geometry for months under physiological temperature and pH. DexMA hydrogels formed through these procedures were generally cytocompatible, non-inflammatory, and non-hemolytic under static cell culture conditions, demonstrating overall hemocompatibility. Our results demonstrated that effective crosslink formation via kinetic chain growth is not solely a function of polymerizable group density, but it also exhibits complex dependence on the polymer’s degree of substitution. These findings reveal the parameter space for tuning hydrogel properties and may facilitate the development of photo-responsive hydrogels for biomedical applications. Our design approach may be particularly relevant to designing blood-interface materials and devices with phototunable volume, where long-term in vivo persistence and hemocompatibility are crucial features.

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Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fbiom.2026.1924191/full#supplementary-material

Supplementary material Supplementary material at the source

Footnotes

Edited by: Mohamed Sharafeldin, University College Cork, Ireland

Reviewed by: Ranjith Kankala, Huaqiao University, China

Luca Guida, Polytechnic University of Milan, Italy

Statements

Conflict of interest

The author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Funding

The author(s) declared that financial support was received for this work and/or its publication. This work was supported by Startup Funds from the School of Biomedical Engineering, Science and Health Systems at Drexel University (CR) and a 2020 Individual Biomedical Research Award from The Hartwell Foundation (CR).

Data availability

The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding author.

Author contributions

AS: Conceptualization, Investigation, Methodology, Supervision, Validation, Writing – original draft, Writing – review and editing, Data curation, Formal Analysis, Visualization. VM: Conceptualization, Data curation, Formal Analysis, Investigation, Methodology, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing. SS: Conceptualization, Data curation, Formal Analysis, Investigation, Methodology, Visualization, Writing – original draft, Writing – review and editing. EP: Conceptualization, Data curation, Investigation, Methodology, Visualization, Writing – original draft, Writing – review and editing. BS: Conceptualization, Supervision, Writing – review and editing. KS: Conceptualization, Funding acquisition, Methodology, Resources, Supervision, Validation, Writing – review and editing. AT: Conceptualization, Funding acquisition, Methodology, Resources, Supervision, Validation, Writing – review and editing. CR: Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Writing – original draft, Writing – review and editing.

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Citation

Seiner, A. J., et al. (2026). Hemocompatible polymeric hydrogels with tunable photoresponsive volumetric properties. Frontiers in Biomaterials Science, Volume 5. https://doi.org/
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5 - 2026

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Copyright © 2026 Seiner, Mishin, Soni, Posthill, Sarkar, Spiller, Throckmorton and Rodell.

License: https://creativecommons.org/licenses/by/4.0/

*Correspondence: Christopher B. Rodell

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