Surface Engineering for Endothelium-Mimicking Functions to Combat Infection and Thrombosis in Extracorporeal Life Support Technologies

Abstract

Blood-contacting medical devices routinely fail from the cascading effects of biofouling towards infection and thrombosis. Nitric oxide (NO) is an integral part of endothelial homeostasis, maintaining platelet quiescence and facilitating oxidative/nitrosative stress against pathogens. Recently, it has been shown that the surface evolution of NO can mediate cell-cell interactions. However, this technique alone cannot prevent the biofouling inherent in device failure with dynamic blood-contacting applications. In this work, we propose an endothelium-mimicking surface design pairing controlled NO release with an inherently antifouling polyethylene glycol interface (NO+PEG). This simple, robust, and scalable platform develops surface-localized NO availability with surface hydration, leading to a significant reduction in protein adsorption as well as bacteria/platelet adhesion. Further in vivo thrombogenicity studies show a decrease in thrombus formation on NO+PEG interfaces, with preservation of circulating platelet and white blood cell counts, maintenance of activated clotting time, and reduced coagulation cascade activation. It is anticipated that this bio-inspired surface design will enable a facile alternative to existing surface technologies to address clinical manifestations of infection and thrombosis in dynamic blood-contacting environments.

Graphical Abstract

Vascular devices routinely fail due to surface-induced thrombosis and infection. Nitric oxide (NO) is a promising agent to prevent platelet activation and microbial growth but cannot prevent any underlying protein surface fouling. A hierarchical strategy of controlled NO release with hydrophilic polyethylene glycol polymer interfaces is described for improved hemocompatibility and device patency in dynamic blood-contacting applications.

1. INTRODUCTION

Extracorporeal life support (ECLS) technologies are a staple of critical care for organ and system failure. Since their debut in 1953, great strides have been made in increasing the safety and therapeutic efficacy of ECLS technologies with significant effort placed towards the development of materials that deter thrombosis.^[1] However, even with modern improvements and practices, thrombotic complications are estimated to occur in 8–31% of patients undergoing ECLS therapies.^[2]

Upon blood contact, plasma proteins rapidly adsorb to foreign materials, such as ECLS circuit components. Plasma proteins initiate and largely facilitate the contact activation pathway of coagulation and the interconnected complement activation response of the immune system. These are universal concerns for blood-contacting medical devices (BCMDs); however, these undesirable responses are particularly prevalent in ECLS-related technologies including extracorporeal circuits (ECCs) and oxygenators with substantial surface areas subjected to dynamic blood flow and gas exchange. In addition to direct complications, the consumption of blood and immune cells in these processes can lead to hypocoagulability post-treatment.^[1] ECC procedures including extracorporeal membrane oxygenation (ECMO) systems routinely fail within 6–12 d, even with systemic heparinization, due to occlusion associated with developed coagulation disorder, bleeding complication, and worsened gas exchange.^[3] This often results in a changeover of the oxygenator before treatment termination (i.e., 1–2 weeks), with a more frequent turnover of circuit loops and other components.^[4]

The lack of materials capable of avoiding innate thrombo-inflammatory response activation has led to the use of systemic anticoagulants as a necessary step to prevent clotting and subsequent BCMD failure. While instrumental to the use of ECLS, anticoagulants come with inherent risks and complications. For example, heparin – the most commonly used anticoagulant for ECMO and hemodialysis, has been associated with heparin-induced thrombocytopenia and increased bleeding risk.^[5] When necessary, anticoagulation can be reversed, but this may lead to additional thrombotic complications.

Another clinical challenge with ECLS technologies is infection. Bloodstream infections in ECMO patients have an estimated prevalence of 3 – 18%,^[6] and there is a greater risk of infection with long-term usage.^[7] In addition to their contributions to morbidity and mortality, bloodstream infections are considered among the most expensive nosocomial complications. Thrombosis and infection are often connected as thrombi can act as focal points for bacterial adhesion, while some bacteria also contribute to thrombi formation (e.g., coagulase-positive Staphylococcus).^[8]

Hemocompatible surface design strategies have been developed with the goal of eliminating the need for systemic anticoagulation and reducing surface biofouling.^[9] Coating circuits with anticoagulants (e.g., heparin) provides anticoagulant effects with reduced risks compared to systemic administration of the same agents. However, these devices maintain the requirement for supplemental anticoagulation therapies.^[10] Most strategies attempt to mimic the native endothelium, as it remains the only truly hemocompatible material. These strategies include surface passivation (e.g., wettability, charge neutrality, etc.) and bioactive approaches (e.g., controlled release of antithrombotic agents).^{[1, 11]} Many of these bioinspired materials have been shown to increase hemocompatibility alone, but there is growing evidence that complementing strategies (representative of the native endothelium) can improve device performance under both dynamic and long-term applications.^[12]

Nitric oxide (NO) is an endogenous gasotransmitter that regulates diverse physiological processes.^[13] Within the immune system, NO production is triggered to facilitate oxidative and nitrosative stresses for antimicrobial effects.^[13] In the cardiovascular system, endothelial cells secrete NO at an estimated rate of 0.5 to 4.0 × 10⁻¹⁰ mol cm⁻² min⁻¹ to regulate vasodilation and mitigate thrombosis through localized inhibition of platelet activation.^[14–15] NO confers surface-localized platelet quiescence effects via downregulation of P-selectin and upregulation of cyclic guanosine monophosphate, leading to phosphorylation of vasodilator-stimulated phosphoprotein and platelet inhibition through cytoskeletal confinement by inactivation of glycoprotein IIb/IIIa.^[16–17]

As a free radical, NO has a short biological half-life (~2 s) that necessitates therapeutic applications with NO donor compounds to achieve prolonged effects.^[18] NO-based therapies and platforms have therefore developed NO donors into solutions, creams, gels, polymers, fibers, and coatings for various materials.^[18–19] S-nitroso-N-acetyl-DL-penicillamine (SNAP) is one well-characterized NO donor known for its relative stability compared to alternative compounds.^[20] In addition, SNAP has the capability of being integrated into prefabricated, medical-grade polymers.^[21] SNAP impregnated into silicone rubber (SR)-based BCMDs has been demonstrated to release NO at physiologically relevant levels while improving hemocompatibility.^[21–23]

Although surface-localized NO release has been shown to reduce thrombosis via platelet inhibition,^[9] it cannot prevent plasma proteins from adhering to surfaces. Prior work has shown that the combination of NO release with high surface wettability has complementary anti-biofouling properties.^[24–25] In this work, a combination strategy using SNAP-impregnated SR with a polyethylene glycol (PEG) interface is described for the improved blood compatibility and infection resistance of BCMDs, especially those subjected to dynamic flow conditions (e.g., ECLS technologies).

Hydrophilic biointerphases have previously been developed by grafting silanated PEG onto hydrophobic polymer surfaces to impart enhanced wettability and resistance to biofouling.^[9] Surface PEGylation has been shown to prevent binding between plasma proteins and the underlying hydrophobic material, leading to the formation of a water hydration layer that interferes with hydrophobic and electrostatic interactions.^[26] Previously, SR surfaces have been modified with silanated PEG to enhance the long-term hemocompatibility of microfluidic devices, while PEG has also acted as a basis for further surface functionalization with zwitterions to achieve charge-neutral interfaces for BCMDs.^[27–28] High grafting densities of PEG are known to repel plasma proteins (e.g., albumin, fibrinogen, etc.), leading to good resistance to platelet adhesion.^[29–30] However, long-term performance is known to be heavily dependent on spacer link and stability under flow environments.^[29–30] Higher molecular weight PEGs have been associated with complement activation, supporting the use of lower molecular weight silanated PEGs (< 1,000 kDa) for the development of stable hemocompatible coatings as well as consideration of other hydrophilic polymer alternatives such as polyglycerol, polyvinylpyrrolidone, zwitterions, and polysaccharides.^{[28, 31–32]}

Herein, a dual-action surface technology is developed with NO release and surface grafting of PEG towards the evaluation of a hydrophilic, NO-releasing platform in vivo for ECLS applications. The study hypothesis is that controlled NO release from the device surface in combination with a passive, hydrophilic polymer interface will attenuate thrombosis and infection better than either individual component owing to complementary bioactive and biopassive modalities. The NO+PEG surface technology is developed in a multi-step process involving the integration of the NO donor SNAP within SR substrates via solvent swelling, surface activation via O₂ plasma treatment, and covalent immobilization of silanated PEG chains. The resulting NO+PEG technology is demonstrated for multifunctionality via materials characterizations including attenuated total reflectance-Fourier transform infrared (ATR-FTIR) spectroscopy, surface contact angle goniometry, and NO release studies, confirming each step of the fabrication. Protein adsorption studies demonstrate enhanced resistance against the deposition of plasma proteins. In vitro biological studies showcase the NO+PEG surface technology’s excellent resistance to bacterial and platelet adhesion while maintaining biocompatibility. Finally, the efficacy of the NO+PEG formulation is assessed via the construction of NO+PEG-modified ECCs and their evaluation in a 4-h rabbit thrombogenicity model, determining the effects on thrombus formation, platelet preservation, and key relationships in coagulation pathway activation.

2. MATERIALS AND METHODS

2.1. Materials

Drabkin’s reagent and Saint-Gobain Tygon^™ Formula 3350 silicone tubing were purchased from VWR (Atlanta, GA). S-nitroso-N-acetyl-DL-penicillamine (SNAP) was purchased from PharmaBlock USA, Inc. (Hatfield, PA). 2-Methoxy polyethyleneoxy (6–9) propyl trimethoxysilane (PEG-silane, Product Code: SIM6492.7, molecular weight range of 459–591 g mol⁻¹) was purchased from Gelest, Inc. (Morrisville, PA). CarboSil^® Thermoplastic Silicone Carbonate Polyurethane was obtained from DSM Biomedical (Exton, PA). Acetone, ethanol, ethylenediaminetetraacetic acid (EDTA), human fibrinogen (Fg), porcine hemoglobin (Hb), methanol, tetrahydrofuran (THF), and Triton-X-100 were acquired from Sigma-Aldrich (St. Louis, MO). Fluorescein isocyanate (FITC)-labeled human Fg was acquired from Innovative Research (Novi, MI). Phosphate-buffered saline (1x PBS) without calcium or magnesium was purchased from Corning (Manassas, VA). All other chemical supplies were reagent-grade and used without further purification.

The Staphylococcus epidermidis (ATCC 35984) and Escherichia coli (ATCC 25922) strains were grown from stocks originally acquired from American Type Culture Collection (Manassas, VA). Luria-Bertani (LB) and Mueller-Hinton (MH) broth and agar were purchased from Sigma-Aldrich (St. Louis, MO). All other bacteria-related supplies were purchased from VWR (Atlanta, GA).

BJ fibroblast (ATCC CRL-2522) cells were subcultured from cryopreserved stocks that were originally purchased from American Type Culture Collection (Manassas, VA). Human vascular endothelial cells (HUVECs) were acquired from Thermo Fisher Scientific (Waltham, MA). Fetal bovine serum (FBS), minimum essential medium (MEM), and penicillin-streptomycin (5,000 units/mL) were purchased from VWR (Atlanta, GA). EGM-2 endothelial cell growth basal media as well as EMG-2 growth supplements were purchased from Lonza (Greenwood, SC).

2.2. Fabrication of NO+PEG Surfaces

NO-releasing substrates were prepared following a previously reported method with minor deviation.^[21] In brief, SR materials were placed into an anhydrous THF swelling solution with SNAP (25 mg mL⁻¹) and mechanically agitated for 24 h at room temperature with protection from light. Substrates were then removed from the swelling solution and dried on glass Petri dishes for an additional 24 h in a fume hood with protection from light. Finally, samples were briefly rinsed in methanol and dried under vacuum for an additional 24 h to afford NO-releasing SR (NO control, see Scheme 1). Samples were stored at −20 °C under desiccant and with protection from light until use.

Scheme 1.

Fabrication of hydrophilic NO-releasing surfaces (NO+PEG) with silanated polyethylene glycol (PEG) chains. SR is modified with the synthetic NO donor, SNAP, using a solvent impregnation technique for 24 h. Afterward, the NO-releasing materials are subjected to oxygen plasma treatment to form hydroxyl groups on the surfaces. The treated materials are then incubated in a solution of silanated PEG at different volumetric ratios to anhydrous acetone to form the hydrophilic NO-releasing (NO+PEG) surfaces.

PEG-grafted surfaces were prepared following a previously reported method with minor deviation.^[27] Substrates were first treated with low-pressure oxygen plasma at 30 W for 15 min to facilitate hydroxyl group coverage on surfaces. Immediately afterward, treated surfaces were submerged in PEG-silane at [1:2, 1:1, or 2:1] volumetric ratios to anhydrous acetone for 1 h. Samples were then briefly rinsed with deionized water and dried under a vacuum overnight. Both hydrophilic controls (PEG control) and hydrophilic NO-releasing surfaces (NO+PEG) were prepared (Scheme 1). Samples were stored at −20 °C under desiccant and with protection from light until use.

2.3. Characterization of NO+PEG Surfaces

Attenuated Total Reflectance-Fourier Transform Infrared (ATR-FTIR) Spectroscopy.

ATR-FTIR scanning of surfaces was performed to confirm the surface conjugation of the hydrophilic silanated PEG. Infrared spectra were collected from 4000 – 650 cm⁻¹ using a Spectrum 3 Spectrometer with a PIKE Technologies VeeMAX III accessory equipped with a flat Ge 60° crystal plate (Perkin-Elmer; Waltham, MA). A total of 128 scans with a resolution of 4 cm⁻¹ were collected for each sample.

Surface Contact Angle Measurement.

Static contact angle measurements of the SR substrates were performed using a contact angle goniometer (Ossila; Sheffield, UK) on treated SR samples (~1 cm²) secured on glass slides. Droplets of deionized water (5 µL) were dispensed on surfaces. Using video frame captures, static contact angles were calculated using the sessile drop approximation. Final values are reported as the mean contact angle for each formulation ± standard deviation (SD) (n = 3 samples from independent preparations).

2.4. Protein Adhesion Studies

The hydrophilic NO-releasing surfaces were tested for resistance towards plasma protein adsorption using human Fg as a test model by adapting previously reported methods.^[33–34] Circular coupons (diameter of 6 mm) were prepared with the surface modifications and fitted into 96-well plates. Samples were preincubated in 1x PBS (100 µL well⁻¹) for 1 h at 37 °C. FITC-Fg was diluted with non-labeled Fg (1:10 by mass) in 1x PBS to achieve a final working solution of 4 mg-Fg mL^-1. Following incubation, plates were treated with the Fg working solution (100 µL well⁻¹) and incubated for an additional 90 min with protection from light. Afterward, plates were washed in an excess of 1x PBS (100 µL well⁻¹ per step) for a total of 10x washing steps. Plates were finally resuspended with 1x PBS (200 µL well⁻¹) and read for fluorescence (Ex: 490 nm, Em: 525 nm) using a Cytation 5 plate reader (BioTek; Vinooski, VT). Standard curves were prepared from the Fg working solution to normalize readings to the mass of Fg adsorbed per unit surface area (µg-Fg cm⁻²). Final results are reported as the mean adsorbed Fg ± SD (n = 5 repeats).

2.5. SNAP Loading

SNAP loading studies were carried out for 24 h with samples incubated in anhydrous THF at a standard surface area-to-volume (SA:V) ratio of 3 cm² mL⁻¹ at 25 °C with protection from light. A standard curve was prepared from SNAP in THF at known concentrations, with absorbance recorded (λ_max = 340 nm) using an Agilent Cary 60 UV-Vis spectrophotometer (see Supporting Information, Fig. S1). Control SR samples were also processed to account for any secondary absorbance from polymer leachates. Absorbances were used to calculate the concentration of SNAP in THF (mg mL⁻¹) to determine the total mass of SNAP (mg) loaded into substrates and the corresponding loading ratios. Final values are reported as the mean wt% for each formulation (mg SNAP per mg total ×100%) ± SD (n = 5 samples from independently prepared substrates).

2.6. SNAP Leaching

SNAP leaching studies were conducted over 24 h under physiological conditions at 37 °C with protection from light. Samples were incubated in 1x PBS with EDTA (100 µM) at SA:V of 1 cm² mL^-1. SNAP leaching was quantified by absorbance (λ_max = 340 nm) with an additional standard curve (see Supporting Information, Fig. S2) prepared from SNAP in 1x PBS with EDTA (100 µM). Absorbances were measured to determine leaching at 30 min, 1 h, 4 h, and 24 h. The final mass of SNAP leached was then calculated and normalized against the initial amount of SNAP loaded into substrates to determine the weight percent of SNAP leached. Final values are reported as the mean weight percent of SNAP leached (defined as mean mg SNAP per mg total of each substrate type × 100%) ± SD (n = 3 samples from independently prepared substrates).

2.7. NO Release Studies

Realtime NO evolution from the SNAP-functionalized materials was determined using a Nitric Oxide Analyzer (NOA) 280i (Zysense; Boulder, CO). The NOA uses a chemiluminescence-based detection method with a purged sample vessel to continuously sweep NO from the solution via a nitrogen carrier stream into an internal reaction chamber. The NOA simultaneously passes an oxygen stream into an ozone generator, and the resulting ozone is fed into the reaction chamber. Ozone and NO react to form excited-state nitrogen dioxide, which upon relaxation emits a photon registered by an internally cooled photodetector (see Eqns. 1 and 2). This high-sensitivity analytical method allows for the determination of NO gas concentration (PPB scale) using an internal calibration constant (mol-NO PPB⁻¹ min⁻¹). This enables the determination of NO surface flux from polymeric materials (mol-NO cm⁻² min⁻¹). All samples were incubated in 1x PBS with 100 µM EDTA at 37 °C during and between measurements to mimic physiologically relevant conditions. Final results are reported as the mean surface flux of NO ± SD (n = 3 runs for each time point using independently prepared samples of each substrate type).

$$\text{NO}+{\text{O}}_3\to {\text{NO}}_2^{*}+{\text{O}}_2$$ (1)

$${\text{NO}}_2^{*}\to {\text{NO}}_2+\text{hν}$$ (2)

2.8. In Vitro Cytotoxicity Studies

Preparation of Mammalian Cell Culture.

BJ cells and HUVECs were revived from cryopreserved stocks and subcultured under a 5% CO₂-humidified atmosphere at 37 °C. BJ cells were cultured in MEM media supplemented with 10% FBS and 1% penicillin-streptomycin. HUVECs were cultured in EBM-2 with the manufacturer-specified growth supplements from the EGM-2 kit. Both cell lines were subcultured for up to ten passages between experiments. When monolayers showed >70% subconfluency, cells were detached with 0.25% trypsin supplemented with 5 mM EDTA. Detached cells were collected via centrifugation (200 RCF, 5 min), counted, and seeded onto 96-well plates (5,000 cells well⁻¹) for experiments.

Indirect Contact (Polymer Leachate) Cytotoxicity Screening (24 h).

Evaluation of cytocompatibility of substrates followed the International Organization for Standardization (ISO) 10993–5 standards for the biological evaluation of medical devices, with some minor modifications.^[35] Circular coupons of the NO+PEG and control materials were sterilized with ultraviolet light for 15 min on both sides. Samples were then individually soaked in supplemented media at a SA:V of 1 cm² mL⁻¹ at 37 °C for 24 h to collect any extracts. Concurrently, seeded 96-well plates were grown for 24 h to achieve >70% subconfluency. The media on the plates was then aspirated and replaced with extract-containing media or control (blank) media (100 µL well⁻¹), plated across several replicate wells for each sample. After an additional 24 h incubation, each plate was suspended with clean media supplemented with 10% CCK-8 reagent (100 µL well⁻¹). Plates were then incubated for an additional 1 h and absorbance was quantified (λ_max = 450 nm with λ_ref = 600 nm). Relative viabilities of wells treated with material extracts compared to control wells treated with blank media were calculated according to Eqn. 3. Final data are presented as mean percent cellular viability ± SD (n = 3 treatment sets developed across independent passages).

$$\text{Cellular\hspace{0.5em}Viability}\left(\text{\%}\right)=\frac{{\left({\text{OD}}_{450}-{\text{OD}}_{650}\right)}_{\text{treatment}}}{{\left({\text{OD}}_{450}-{\text{OD}}_{650}\right)}_{\text{control}}}\times 100\%$$ (3)

2.9. In Vitro Bacteria Studies

Preparation of Bacteria Cultures.

Bacteria cultures were grown from stocks in MH broth until reaching the log phase of growth as determined by optical density measurements (OD₆₀₀ ~0.8 – 1.0). The culture was then pelleted (4,400 RCF, 7 min), washed, pelleted again, and resuspended in 1x PBS. The bacterial suspension was then diluted in 1x PBS to an absorbance (OD₆₀₀) of 0.1 for use in adhesion experiments.

Bacteria Adhesion Studies (24 h).

Substrates were sterilized via ultraviolet irradiation for 15 min on each side before bacterial exposure. Individual samples were then placed in bacterial solutions (OD₆₀₀ = 0.1 at SA:V of 1 cm² mL⁻¹) and shaken for 24 h at 37 °C with protection from light. After incubation, the samples were rinsed with sterile PBS to remove any non-adherent bacteria and homogenized in 1x PBS (1 mL) to detach any adherent bacteria. The homogenates were diluted 10x times and plated with the assistance of an Eddy Jet W2 spiral plater (IUL Instruments; Farmingdale, NY) on MH agar. After overnight incubation at 37 °C, colony-forming units (CFUs) were counted using a SphereFlash automated colony counter (IUL Instruments; Farmingdale, NY). Final results are presented as the mean CFU cm⁻² ± SD (n = 4 per sample type). Percent bacterial reductions are calculated using Eqn. 4.

$$\text{Bacteria\hspace{0.5em}Reduction}\left(\text{\%}\right)=\frac{{\text{Count}}_{\text{control}}\left(\text{CFU\hspace{0.5em}}{\text{cm}}^{-2}\right)-{\text{Count}}_{\text{treatment}}\left(\text{CFU\hspace{0.5em}}{\text{cm}}^{-2}\right)}{{\text{Count}}_{\text{control}}\left(\text{CFU\hspace{0.5em}}{\text{cm}}^{-2}\right)}\times 100\%$$ (4)

2.10. In Vitro Blood Compatibility Screening

Collection and Processing of Porcine Whole Blood.

All procedures concerning the use of porcine whole blood (WB) and its components were approved by the Institutional Animal Care and Use Committee of the University of Georgia. Porcine WB was drawn using 3.2% sodium citrate solution at a 1:10 dilution through blind draws from a donor population. Collected blood was assessed for quality compliance (i.e., nominal platelet counts, white blood cell counts, and hemoglobin levels) and then fractionated into platelet-rich plasma (PRP) via centrifugation (280 RCF, 12 min).^[36] The PRP was extracted and analyzed for hematological parameters, with the remaining fractions further spun down (3,000 RCF, 20 min) to obtain platelet-poor plasma (PPP). All blood samples were processed and tested within 4 h of drawing.

In Vitro Platelet Adhesion Assessment (90 min).

Platelet adhesion to surfaces was assessed following previously reported methods with minor deviation.^[34] PRP was diluted with PPP to obtain a final count of 2 × 10⁸ platelets mL⁻¹ with anticoagulation reversed using calcium chloride (2.5 mM). Circular coupons prepared from the modified surfaces were then incubated with diluted PRP (SA:V of 1 cm² mL⁻¹) for 90 min at 37 °C with gentle rocking. After incubation, samples were removed and gently washed with 1x PBS (Ca²⁺/Mg²⁺-free) to remove nonadherent platelets. Adherent platelets were then lysed with 2 % v/v Triton-X-100 in 1x PBS (Ca²⁺/Mg²⁺-free) and quantified using the Roche^® Cytotoxicity Detection Kit based on the colorimetric detection of lactate dehydrogenase (LDH) from lysed cells (λ_max = 492 nm with λ_ref = 620 nm). These readings were calculated against a standard curve of lysed platelets to determine the total number of adherent platelets normalized to the sample surface area. Final data are reported as the mean number of platelets per cm² ± SD (n = 3 per treatment).

Hemolysis Screening (3 h).

Hemolysis screenings of the NO+PEG and other surface modifications were performed following the ISO 10993–4 standards for the biological evaluation of medical device interactions with blood with some minor deviation.^[37–38] Freshly drawn porcine WB was diluted in 1x PBS (Ca²⁺/Mg²⁺-free) to a final Hb concentration of 10.0 mg mL^-1. Materials were then placed in the diluted WB (SA:V of 3 cm² mL⁻¹). Reference samples included a diluted WB blank in 1x PBS (Ca²⁺/Mg²⁺-free), unmodified SR negative control (SA:V of 3 cm² mL⁻¹) in diluted WB, and diluted WB positive control in deionized water. Samples were incubated at 37 °C for 3 h with manual rocking every 30 min. Afterward, the WB and extract solutions were centrifuged (800 RCF, 15 min). The supernatants were mixed 1:1 with Drabkin’s reagent and incubated for 15 min with protection from light. Absorbance was then quantified (λ_max = 540 nm) and compared against a standard curve developed with Hb. Hb concentrations (${\text{C}}_{\text{Hb}}$) were determined and used to calculate percent hemolysis per Eqn. 5. Final data are reported as the mean hemolytic index (normalized against SR control) ± SD (n = 3 per treatment).

$$\text{Hemolysis}\left(\text{\%}\right)=\frac{{\text{C}}_{\text{Hb},\text{Sample}}-{\text{C}}_{\text{Hb},\text{Diluted\hspace{0.5em}WB\hspace{0.5em}Blank}}}{{\text{C}}_{\text{Hb},\text{Positive\hspace{0.5em}Control}}-{\text{C}}_{\text{Hb},\text{Diluted\hspace{0.5em}WB\hspace{0.5em}Blank}}}\times 100\%$$ (5)

2.11. In Vivo Extracorporeal Circulation Rabbit Model

Extracorporeal Circuit Fabrication.

The ECC loops used for in vivo evaluation were prepared following a previously reported method with slight modification.^[21] SR tubing was functionalized for NO release via swelling with SNAP in THF (see Section 2.2). The loops consisted of two segments of 1/4” inner diameter SR tubing (16 cm in length) on either side of one segment of 3/8” inner diameter SR tubing (8 cm in length), which served as the thrombogenicity chamber. The pieces were assembled using a silicone sealant and allowed to cure for 24 h before further modification. Unmodified SR tubing was likewise assembled and used as controls. The constructed, SNAP-modified loops were plasma treated for 15 min before being filled with 1:1 PEG-silane in anhydrous acetone (see Section 2.2). Both loop ends were clamped to prevent acetone evaporation. After 1 h of incubation, the loops were thoroughly rinsed with deionized water and dried. Angiocatheters (16- and 14-gauge) were coated with CarboSil^® Thermoplastic Silicone Polycarbonate Polyurethane dissolved in THF (control circuits) or silicone rubber containing 10 wt% SNAP in THF (NO+PEG circuits). All ECC loops were soaked in normal saline for 60 min before use.

In Vivo Thrombogenicity Model.

New Zealand white rabbits (Robinson Services Inc, MI) were used in these studies, weighing between 2.2 – 3.0 kg. An established 4-h arteriovenous (AV) shunt model was used to evaluate the thrombogenicity of the fabricated ECCs.^{[21, 39]} Briefly, the animals were pre-anesthetized with intramuscular injections of 5 mg kg⁻¹ xylazine injectable (AnaSed^®, Lloyd Laboratories; Shenandoah, IA) and 30 mg kg⁻¹ ketamine hydrochloride (Hospira Inc.; Lake Forest, IL), and the anesthesia was maintained via isoflurane gas inhalation at a rate of 2 – 4%. IV fluids of Lactated Ringers were given at a rate of 10 mL kg⁻¹ h⁻¹ to maintain blood pressure stability. The custom-built circuit (with a thrombogenicity chamber in the middle) was placed in the animal’s left carotid artery and right external jugular vein. The right carotid artery was cannulated using a 16-gauge IV angiocatheter (Jelco^®, Johnson & Johnson; Cincinnati, OH) for blood sample collection as well as for blood pressure and heart rate monitoring. Body temperature was maintained at 37 °C using a water-jacketed heating blanket and was monitored with a rectal probe. Animals were not systemically anticoagulated during the experiments. Blood flow through the ECCs was initiated by unclamping both the arterial and venous sides of the circuit and was monitored throughout the studies using an ultrasonic flow probe and meter (Transonic HT207; Ithaca, NY). The animal handling and surgical procedures were approved by the University of Michigan, University Committee on the Use and Care of Animals following university and federal regulations (UM protocol# PRO00010710).

After 4 h on ECC, animals were treated with sodium heparin (400 U/kg) to prevent necrotic thrombosis and euthanized using Fatal Plus (130 mg/kg sodium pentobarbital) (Vortech Pharmaceuticals; Dearborn, MI). Circuits were clamped and removed from the animal. Circuits were rinsed with saline (60 mL) and drained. The thrombogenicity chamber on each circuit (see Supporting Information, Fig. S8) was isolated and cut longitudinally, with any residual thrombus on the surfaces photographed. The degree of thrombus surface coverage was determined using ImageJ software from the National Institutes of Health (Bethesda, MD). The remaining thrombi were then collected and weighed. Circular punchouts of the thrombogenicity chamber were further collected and analyzed for NO release following 4 h flow using the chemiluminescence-based NO detection approach described in Section 2.7. Endpoint NO release measurements were compared against measurements of pristine segments prepared during loop construction prior to cutting the circuit to size.

Blood Sample Collection and Analysis.

Blood samples were collected at baseline and every hour after the initiation of the blood flow in the ECC for a total of 4 h of measurements. Hemodynamic parameters, as well as arterial blood pH, pCO₂, pO_2, and total hemoglobin, were measured using a blood-gas analyzer (ABL 825 Radiometer; Copenhagen, DK). Platelet and white blood cell counts (IDEXX ProCyte Dx; Westbrook, ME), plasma Fg levels (Dade Behring BCS XP, Siemens; Deerfield, IL), and activated clotting times (ACT, Hemochron Blood Coagulation System Model 801, International Technidyne Corp.; Edison, NJ) were also determined and monitored.

2.12. Statistical Analysis

All statistical comparisons were performed with GraphPad Prism 9 (GraphPad Software; San Diego, CA). All materials characterization and in vitro data are reported as the mean ± standard deviation (SD) unless explicitly stated otherwise. Comparisons between treated and control groups in these studies were made using ordinary one-way analysis of variance (ANOVA), implementing Tukey’s method for correction of multiple comparisons. Time-dependent comparisons were made using ordinary two-way ANOVA. Values of p ≤ 0.05 were deemed significant.

All in vivo data is reported as the mean ± standard error of the mean (SEM) for each animal group following outlier analysis (n = 4 screened rabbits per group) unless explicitly stated otherwise. All intragroup time-course-dependent in vivo comparisons were made using one-way ANOVA with Tukey’s method for multiple comparisons. Values of p ≤ 0.05 were deemed significant. All intergroup comparisons were made using two-way ANOVA. To account for multiple hypothesis testing, the false discovery rate (FDR) was controlled using the two-stage step-up method of Benjamini, Krieger, and Yekutieli. A false discovery rate q < 0.05 was deemed significant. The corresponding p-values are indicated for comparisons wherein p ≤ 0.05. Endpoint analyses of clot mass and chamber thrombus surface coverage were performed using unpaired t-tests assuming equal variance. In all analyses, p ≤ 0.05 was deemed statistically significant.

3. RESULTS AND DISCUSSION

3.1. Surface Characterization

The NO donor compound SNAP was integrated into SR (Scheme 1) following previously reported methods to develop a NO-releasing polymeric platform.^[21] SNAP is a well-characterized NO donor that has been incorporated into polyurethanes, polyvinyl chloride, silicone rubber, and other medical-grade plastics via blending, solvent impregnation, and covalent immobilization.^{[12, 19]} While SNAP-integrated materials are antithrombotic and antimicrobial,^[1] they lack mechanisms for preventing surface fouling that often compromises BCMDs. Hydrophilic surface modifications were therefore performed by grafting silanated PEG (Scheme 1) to confer exceptional surface wettability, improving the resistance of the blood-contacting interface against biofouling via complementation with NO (Fig. 1a).

Fig. 1.

Surface wettabilities and antifouling performances of (a) hydrophilic NO-releasing surfaces. Static contact angle measurements of the pristine surfaces (b) showed surface-dependent changes in wettability based on the volumetric ratio of silanated PEG to anhydrous acetone in the grafting step. Measurements of fibrinogen adsorption to the surfaces (c) demonstrated significant improvement in antifouling performance with a PEG ratio of (1:1) or higher. Final data are reported as the mean ± SD (n ≥ 3, see Sections 2.3 and 2.4). Comparisons in (b) and (c) are calculated from one-way ANOVA tests using Tukey’s method. *Indicates a significant difference against the SR control group. ^†Indicates a significant difference against the NO control group. ^‡Indicates a significant difference against the PEG (1:2) group. ^§Indicates a significant difference against the NO+PEG (1:1) group. ^‖Indicates a significant difference against the NO+PEG (1:2) group. Statistical significance is defined as p ≤ 0.05.

FTIR Analysis.

PEG (~0.46 – 0.59 kDa) grafting onto the NO+PEG surfaces was first characterized by FTIR (see Supporting Information, Fig. S3). Characteristic peaks for the condensed silanated PEG were observed through grazing angle reflection techniques, including the formation of siloxane (-Si-O-Si-) bonds (1095 cm⁻¹) on both PEG and NO+PEG surfaces. Hydrolysis of the organomethoxysilane leads to the formation of silanol groups, which facilitates condensation for subsequent monolayer and aggregate layers formation of the PEG interphase. Further evidence of PEG grafting was observed by hydroxyl group (-OH) stretching (3620–3160 cm⁻¹), alkylsilane (Si-C) stretching (~1280 cm⁻¹), as well as symmetric and asymmetric aliphatic (C-H) stretching (2924/2872 cm⁻¹) (Fig. S3). These findings are consistent with previous reports for silane-based immobilization of PEG chains.^[40–41]

Surface Wettability.

Static contact angle measurements were performed to assess changes in surface wettability during the fabrication processes. Solvent impregnation of SR with SNAP led to no appreciable difference in surface wettability on NO control surfaces compared to untreated SR control (p > 0.05, Fig. 1b). In contrast, all PEG-modified groups showed a significant reduction in static water contact angle compared to both the SR and NO groups (p ≤ 0.05, Fig. 1b). Among the PEG controls and NO+PEG, significant differences were observed with the PEG and NO+PEG (2:1) formulations against the respective (1:2) and (1:1) formulations (p ≤ 0.05, Fig. 1b). The increased PEG-silane volumetric ratio justifies the improved wettabilities of the (2:1) formulations, while SNAP has also been shown to increase surface retention of water which can facilitate silane condensation.^[42] Surface wetting is a good indicator of a material’s resistance to biofouling, as the formation of water hydration layers on wetting materials assists in the repulsion of hydrophobic plasma proteins and improves resistance towards bacterial colonization and platelet adhesion.^[24]

3.2. Resistance to Protein Adsorption

Surface passivation is a well-known strategy for the prevention of surface-induced thrombosis and bacteria contamination but does not fully protect BCMDs, especially under flow environments and longer-term applications.^[43] For this reason, the proposed hydrophilic NO-releasing surface technology may help avoid an untoward effect towards thrombosis and infection by facilitating a direct biocidal mechanism against bacteria and inducing localized platelet quiescence.^{[1, 19]}

Fibrinogen (Fg) is a hydrophobic plasma protein that undergoes enzymatic conversion in the presence of thrombin, leading to fibrin formation that is key to both platelet adhesion and bacterial colonization of BCMDs.^[1] A general antifouling trend was observed (Fig. 1c) following exposure of PEG-modified surfaces to physiological concentrations of human Fg (2 mg mL⁻¹). NO-releasing control materials showed a ~32% increase in Fg adsorption compared to unmodified SR (p ≤ 0.05, see Fig. 1c), which is consistent with prior observation.^[44] The addition of the silanated PEG on the NO+PEG surfaces led to ~74 and 80% reductions in adhered Fg compared to unmodified SR and NO controls, respectively (Fig. 1c). Interestingly, the NO+PEG (1:2) formulation exhibited poorer resistance to protein fouling compared to the NO+PEG (1:1) and (2:1) formulations. This is attributed to the possibility that the lower volumetric ratio of PEG in the deposition solution led to reduced surface functionalization. Similar reductions in Fg adhesion between the NO+PEG (1:1) and (2:1) formulations suggest that an antifouling threshold was reached with the surfaces, after which the higher volumetric ratio of PEG did not confer circumstantial benefit. Previously, others have shown that PEG coatings may reach saturation in both surface coverage and density, thereby limiting the added benefit of larger chain lengths, higher concentration grafting solutions, and incubation times.^{[40, 45]} Based on these findings, the hydrophilic NO-releasing surfaces were further evaluated for the influence of the grafting process on NO donor retention and any other effects on NO release kinetics.

3.3. Nitric Oxide Release Kinetics

The synthetic NO donor SNAP forms NO in the presence of heat, light, or metal ions via homolytic cleavage of its S-nitrosothiol bond (Fig. 2a). SNAP decomposition in NO-releasing polymers follows pseudo-first-order reaction kinetics, enabling the development of sustained surface release of NO when exposed to physiological conditions.^[46–47] Solvent impregnation of SNAP into SR implicates that a finite reservoir of the donor is loaded, therefore it is essential to evaluate how further processing of the SR substrates affects SNAP retention.^[21] As shown in Fig. 2b, PEG grafting at the (1:2) ratio led to a ~25% drop in SNAP loading (p ≤ 0.05), while both the (1:1) and the (2:1) formulations showed no significant differences in SNAP loading. SR exhibits moderate swelling capacity in anhydrous acetone (swelling ratio of 1.06 based on the change in length), while SNAP is moderately soluble in acetone (< 70 mg mL⁻¹).^[48–49] Therefore, it is reasonable to expect that the higher volumetric ratio of acetone in the (1:2) formulation leads to a proportionately higher loss in SNAP loading. This phenomenon coupled with other findings on surface wettability and protein adhesion implies that the (1:2) formulation is unacceptable for the hydrophilic NO-releasing surface application, as too many tradeoffs are made in performance. Concurrently, while the higher PEG ratio of (2:1) exhibited excellent material properties, no added advantage in anti-biofouling performance or SNAP loading is evident compared to the (1:1) formulation. Therefore, further studies of NO release kinetics and biological evaluation reported herein consider the NO+PEG (1:1) formulation.

Fig. 2.

NO release studies from hydrophilic NO-releasing materials prepared via solvent impregnation with (a) the NO donor SNAP that produces NO as a result of environmental stimuli. Hydrophilic NO-releasing materials showed (b) dissimilar retention of SNAP based on grafting conditions. A study of SNAP leaching (c) from the NO+PEG (1:1) formulation showed decreased diffusion of the donor compared to the NO control. Final data for both loading and leaching studies are reported as the mean ± SD (n ≥ 3 replicates per formulation, see Sections 2.5 and 2.6). Further study of the NO+PEG (1:1) formulation showed that the (d) instantaneous NO surface flux profiles, (e) average NO surface fluxes at several time points, and (f) total NO release thresholds after 4 h were comparable to NO controls under physiological conditions (37 °C in 1x PBS with 100 mM EDTA). Final data are reported as the mean ± SD (n = 3 replicates per formulation, see Section 2.7). Comparisons in (b) and (f) are calculated from one-way ANOVA tests using Tukey’s method. Time-dependent intergroup comparisons in (c) and (e) are calculated from two-way ANOVA tests using FDR procedure (see Section 2.12). *Indicates a significant difference against the NO control group in (b) and at each corresponding time point in (c) against NO+PEG (1:1). ^†Indicates a significant difference against the NO+PEG (1:2) group. Statistical significance is defined as p ≤ 0.05.

The extent of NO donor leaching from the polymer is a key consideration for the translation of the surface technology. Ideally, the NO-releasing surface will evolve NO via intrapolymeric degradation of SNAP without significant leaching of the donor into the surrounding physiological environment which could trigger secondary biocidal effects.^[50] Therefore, the NO-releasing materials were further evaluated for leaching over 24 h under physiological conditions (1x PBS with 100 mM EDTA, 37°C), showing remarkably decreased SNAP leaching with NO+PEG (1:1) compared to the NO control (Fig. 2c). One explanation for this trend develops from the emergence of both amorphous and crystalline states of SNAP in SR as a result of SNAP being unable to reach solubility equilibrium within the polymer matrix during the impregnation step (24 h), as was previously reported for other SR-based materials.^{[21, 49]} This is a key tradeoff of the solvent impregnation process, wherein SNAP may also become degraded over time due to trace water, metal contaminants, or other process parameters. Therefore, decreased leaching could result from some loss of SNAP during the PEG grafting step. The combination of acetone solvent and H₂O/EtOH formation from silane condensation may contribute to some solubilization and extraction of SNAP, especially from the amorphous (dissolved) state, which has been previously associated with the elevated NO release kinetics from SR substrates when first placed under physiological conditions.^{[22, 51]}

To further understand the effects of reduced SNAP leaching, NO release kinetics of the hydrophilic NO-releasing materials were evaluated during an initial 24-h timeframe via a chemiluminescence-based detection method using the NOA 280i instrumentation. Representative NO flux profiles from both the NO and NO+PEG (1:1) materials demonstrated that the NO control surfaces could exhibit higher initial NO release rates (Fig. 2d). However, this difference was insignificant by 1 h, with the surface flux from both NO and NO+PEG (1:1) surfaces showing no significant differences through 24 h (Fig. 2e). Similarly, the NO and NO+PEG (1:1) materials showed no appreciable difference in the total NO released after 4 h (Fig. 2f). Prior reports of SNAP-impregnated SR materials have demonstrated similar surface fluxes, with the NO fluxes being heavily dependent on the thicknesses of the substrates, SNAP concentrations in the swelling solutions, and durations of swelling.^{[22, 52]} Based on prior work, it is reasonable that the NO surface flux can be further tailored by manipulation of these parameters, such as to develop higher surface fluxes or longer-term thresholds in an application-dependent context.^{[50, 52]} Careful manipulation of the NO surface flux within the endothelial range (i.e., <0.5 – 4.0 × 10⁻¹⁰ mol cm⁻² min⁻¹) is essential, as higher fluxes and NO donor leaching have been reported to induce hemolytic responses.^[53]

Although the surface flux was investigated only during the initial 24 h (corresponding to in vitro and in vivo studies described herein), prior reports of SNAP-impregnated SR have demonstrated physiologically actionable NO release for over three weeks.^[22] Prior work with other hydrophilic NO-releasing materials has considered the incorporation of high water-uptake polyurethanes^[24] or post-swelling processing by further surface functionalization with polyphosphorylcholine zwitterion brushes.^{[25, 54]} In both cases, polyurethanes with either polyether or polydimethylsiloxane soft segments were used to coat existing silicone-based medical devices for infection-resistant applications. In contrast, the approach reported herein offers several unique advantages by ready modification of existing silicone-based devices. These advantages include fewer fabrication steps, a more robust sense of scalability (e.g., solvent impregnation of preformed devices versus solvent blending and dip-coating), and broad substrate compatibility using plasma treatment with silane chemistry.^{[25, 54]} Based on these findings, the hydrophilic NO-releasing materials were further evaluated for translational feasibility in several in vitro and in vivo studies.

3.4. Stability Studies

The clinical relevancy of the hydrophilic NO-releasing surface technology is predicated on its stability under storage and after sterilization. Typical blood-contacting devices such as oxygenators, catheters, and coated silicone-based technologies have shelf lives on the order of two years and are compatible with an appropriate, clinically available sterilization technique.^[55] Previously, SNAP-based NO-releasing polymer composites have been shown to exhibit six months of storage stability, with ~90% retention of SNAP loading following ethylene oxide or vaporized hydrogen peroxide (vH₂O₂) sterilization.^[56–57] While silanated PEGs are an efficacious surface strategy, there is the risk of oxidative degradation over time compromising antifouling features.^[32] This phenomenon has been observed with PEGs as low as 1 kDa via metal ion catalysis and can lead to further complement system activation in implanted devices.^[58]

The stability of the NO-releasing PEGylated surfaces was assessed after two years of cold storage (−20 °C), showing retention of SNAP loading, NO release rates, and surface wettability compared to freshly prepared materials (see Supporting Information, Figs. S4-S5). In similar studies with the NO+PEG surfaces sterilized via a clinically available vH₂O₂ process, > 83% retention of SNAP was observed in NO samples following sterilization, while NO+PEG samples displayed > 90% retention of SNAP (Fig. S6). The addition of the silanated PEG interface on NO+PEG surfaces may confer added protection towards the embedded NO donor while exhibiting no significant change in surface wettability after sterilization (Fig. S7). A minor increase in NO release rate is observed post-sterilization with NO+PEG, though this difference borders insignificant (Fig. S6a). Overall, storage and sterilization stability studies support the combination NO+PEG design pathway as efficacious towards long-term cold storage and is compatible with an existing sterilization technique appropriate for heat-sensitive devices.

3.5. In Vitro Evaluation of Cytocompatibility

Ensuring that the hydrophilic NO-releasing surfaces do not induce a cytotoxic effect is a critical checkpoint to pass before any further evaluation in an in vivo model as a medical device surface technology. Therefore, the dual-action NO+PEG materials were tested in 24-h indirect contact (leachate) cytotoxicity screenings following ISO 10993–5 standards concerning the biological evaluation of medical devices, with minor deviation.^[35] Human-derived fibroblasts (BJ) and endothelial cells (HUVECs) were used as model cell lines. As shown in Fig. 3a, leachates from the NO and PEG control materials caused no significant changes in BJ fibroblast viability, while the combination NO+PEG (1:1) maintained >90% viability. This slight decrease in viability is attributed to the possibility of reactive oxygen species (ROSs) such as peroxynitrite (${\text{ONOO}}^{–}$) and hydroxyl radicals (${\square }^{•}\text{OH}$) to form in the presence of NO and superoxide (${\text{O}}_2^{-}$).^[59] These reactive species can facilitate oxidative degradation of the PEG chains at terminal hydroxyl groups, resulting in a minor decrease in viability.^[60–61] The NO+PEG (1:1) material was previously shown to have decreased SNAP leaching compared to the NO control, with comparable total NO release rates (Figs. 2c-f).

Fig. 3.

Cytocompatibility screening of the NO+PEG (1:1) surfaces against (a) human fibroblasts (BJ) and (b) human umbilical vascular endothelial cells (HUVECs) in 24-h indirect contact leachate studies. All materials were found to retain greater than 80% cellular viability compared to the SR controls. Final data are reported as the mean percent viability of cell cultures treated with leachates from each formulation ± SD (n = 9 independent repeats, see Section 2.8). Comparisons are calculated from one-way ANOVA tests using Tukey’s method. *Indicates a significant difference against the SR control group. ^†Indicates a significant difference against the NO control group. ^‡Indicates a significant difference against the NO+PEG (1:1) group. Statistical significance is defined as p ≤ 0.05.

Further evaluation of the hydrophilic NO-releasing materials with HUVECs demonstrated similar findings, with the PEG (1:1) control causing no significant difference in the cellular viability compared to the SR control (Fig. 3b). On the other hand, the active NO release components of both the NO control and NO+PEG (1:1) surfaces enhanced cellular proliferation (p ≤ 0.05, see Fig. 3b), consistent with prior findings of increased endothelial cell proliferation in the presence of NO.^{[38, 50]} In this sense, any cytotoxic effect from the PEG component or ROSs with the NO+PEG (1:1) formulation is countered by the enhanced proliferative effect of NO with HUVECs. With clear trends of cytocompatibility during the initial 24-h period, the materials were further analyzed for their ability to resist bacterial adherence and reduce infection.

3.6. In Vitro Evaluation of Antimicrobial Performance

Another key consideration for the construction of blood-contacting materials is evaluating microbial surface biofouling, which can trigger long-term risks for infection and device failure. NO-releasing materials are known to be bactericidal and reduce the viability of adhered bacteria on substrates. The addition of silanated PEG chains further creates an antifouling interface that can prevent initial microbial attachment, granting multifunctionality to the hydrophilic NO-releasing surfaces.

The antibacterial efficacy of the hydrophilic NO-releasing surfaces was evaluated through 24-h adhesion studies with the representative Gram-positive S. epidermidis strain and the Gram-negative E. coli strain. Upon exposure to S. epidermis, PEG and NO controls individually decreased viable bacterial adhesion by 44.84% and 93.85%, respectively, compared to the unmodified SR controls (p ≤ 0.05, see Fig. 4a and Table S1). However, the combination of the surface-immobilized PEG and NO release with the NO+PEG materials reduced adhesion by 99.02% compared to unmodified SR controls (p ≤ 0.05), showcasing the complementary nature of the dual active-passive mechanism toward anti-biofouling performance.

Fig. 4.

Antimicrobial studies of the NO+PEG (1:1) surfaces against clinically relevant (a) S. epidermidis and (b) E. coli demonstrated an enhanced reduction in viable, adherent bacteria to surfaces with the combination strategy. The NO+PEG (1:1) strategy resulted in 2.01 and 1.55-log reductions of S. epidermidis and E. coli after 24 h, respectively. Final data are reported as the mean colony-forming unit counts adjusted to surface area (CFU cm⁻²) ± SD (n = 4, see Section 2.9). Comparisons are calculated from one-way ANOVA tests using Tukey’s method. *Indicates a significant difference against the SR control group. ^†Indicates a significant difference against the NO control group. ^‡Indicates a significant difference against the PEG (1:1) group. Statistical significance is defined as p ≤ 0.05.

In separate studies with E. coli, PEG and NO control materials decreased counts of viable adhered bacteria by 92.20% and 77.62%, respectively, compared to SR controls (p ≤ 0.05, see Fig. 4b and Table S2). The combination of the two strategies with the NO+PEG surfaces again resulted in a superior reduction of 97.16% (p ≤ 0.05, see Fig. 4b and Table S2). These results are consistent with prior findings, wherein several groups have repeatedly shown an enhanced susceptibility of many Gram-positive bacteria towards NO compared to Gram-negative counterparts.^{[50, 62]} Prior studies have also demonstrated the decreased viability of Gram-negative bacteria on PEG-coated materials compared to Gram-positive counterparts.^[63] These trends may be attributed to differences in bacterial membranes and the influence of the underline surface chemistries. Overall, the combination NO+PEG surfaces exhibit complementary antibacterial surface effects through both a contact-active mechanism with NO and surface passivation with PEG, leading to decreased viability of adhered bacteria for both Gram-positive and Gram-negative strains (2.01 and 1.55-log reductions respectively).

3.7. In Vitro Screening for Blood Compatibility

In vitro hemocompatibility screenings of the hydrophilic NO-releasing materials and controls were conducted against diluted porcine PRP and WB for platelet adhesion and hemolysis testing, respectively. These screenings are critical before in vivo testing to verify the performance of the materials in a blood-contacting environment, especially in view of the effects on platelets and red blood cells. Platelet adhesion testing in fresh, diluted porcine PRP (~2 × 10⁸ platelets mL⁻¹) for 90 min showed significant reductions in platelet adhesion with all test groups compared to SR controls (p ≤ 0.05, see Fig. 5a). Notably, the NO+PEG (1:1) group showed the highest, statistically significant reduction in platelet adhesion compared to all other groups, with >96% reduction compared to SR controls (p ≤ 0.05, see Fig. 5a). Interestingly, even without an active mechanism for platelet quiescence from NO release, the PEG (1:1) control led to a nearly 85% reduction in platelet adhesion relative to the SR control (p ≤ 0.05, see Fig. 5a). These findings were consistent with previous reports with other NO-releasing control materials and confirm the complementary effect of PEG with NO to improve resistance to platelet adhesion and activation.^{[23, 64]}

Fig. 5.

In vitro blood compatibility screening of NO+PEG (1:1) and related surface modifications. Studies of platelet adhesion (a) with diluted porcine PRP showed a greater than 96% reduction in platelet adhesion to surfaces relative to the SR control with (b) maintenance of blood compatibility by not inducing a hemolytic response. Final data are reported as the mean ± SD (n = 3, see Section 2.10). Comparisons are calculated from one-way ANOVA tests using Tukey’s method. *Indicates a significant difference against the SR control group. ^†Indicates a significant difference against the NO control group. ^‡Indicates a significant difference against the PEG (1:1) control group. Statistical significance is defined as p ≤ 0.05.

While the hydrophilic NO-releasing materials were able to confer a platelet quiescence effect as a pathway to adhesion resistance, it was necessary to further evaluate the blood compatibility of the materials via benchmarking any tendency to cause red blood cell lysis. Following ISO 10993–5 standards for hemolysis testing, none of the NO control, PEG control, or NO+PEG materials were found to induce a hemolytic response (Fig. 5b). This trend is consistent with prior findings with other NO-releasing materials^{[38, 65]} and supports the general hemocompatibility of the NO+PEG surfaces for further performance evaluation in a rabbit thrombogenicity model.

3.8. In Vivo Rabbit Model of Extracorporeal Circulation

Based on promising in vitro data for biocompatibility, protein adsorption resistance, antibacterial action, and antiplatelet effects, the hydrophilic NO-releasing materials (NO+PEG (1:1)) were further assessed in an in vivo extracorporeal rabbit thrombogenicity model (see Supporting Information, Figure S8). The NO+PEG (1:1) formulation was used in the construction of extracorporeal circuits (ECCs) which were applied in 4-h arteriovenous shunt studies through a well-established thrombogenicity model without systemic anticoagulation in New Zealand white hares.^{[21, 39, 66]} Both NO+PEG (1:1) and SR control circuits were evaluated, with circuits primed for 1 h with normal saline before implantation to deter any systemic effects from burst release of NO that may be induced by initial SNAP leaching.^{[21, 66]} The NO release from the NO+PEG (1:1) circuits was measured following the 1-h saline flush and 4-h implantation using chemiluminescence-based NOA detection (see Section 2.7), with nominal NO release levels of > 0.50 × 10⁻¹⁰ mol cm⁻² min⁻¹ maintained by the circuit throughout the study. This finding was consistent with prior studies of the NO+PEG (1:1) formulation in PBS (Fig. 2e) and other NO-releasing materials, implying maintenance of NO surface flux within endothelial levels without significant change in the overall release kinetics as a result of blood exposure.^{[21, 62]}

Key hemodynamic parameters were monitored throughout the 4 h experiment, including mean arterial pressure (MAP), heart rate (HR), and circuit blood flow (ECC BF) (Table 1). No key changes were observed in MAP and HR vital parameters with control circuits, while a slight drop in MAP was recorded for NO+PEG (1:1) circuits by 4 h. This less than 25% drop in MAP relative to baseline for NO+PEG (1:1) was unexpected, compared to prior studies of NO-releasing circuits showing >50% drop in MAP.^[21] The greater preservation in MAP may be correlated with the reduced SNAP leaching of the NO+PEG circuits, as NO is known to lower pressure by enhancement of vasodilation.^[67] Reduced SNAP leaching observed with the NO+PEG (1:1) formulation (Fig. 2c) may therefore reduce systemic effects from nonlocalized NO release, which is ideal for pinpointing the effects of the circuits proximal to the blood-contacting, polymer-solution interface. Blood flow through the ECCs is highly dependent on MAP and surface occlusion from thrombi. ECC blood flow proportionately increased for the NO+PEG (1:1) group (Table 1), which can be attributed to hemodilution effects from IV fluids administration, NO release mediating vasodilation (supported by significant decrease in MAP by 4 h), as well as possible lubrication effects of the hydrophilic NO-releasing surfaces.^{[21, 68–69]}

Table 1.

Evaluation of rabbit hemodynamic parameters as a result of the NO+PEG (1:1) and SR control ECC circuits during 4 h of blood flow. Mean arterial pressure (MAP), heart rate (HR), and extracorporeal circuit blood flow (ECC BF).

Hemodynamics
ECC	Parameter	Time on ECC (h)
		Baseline	1	2	3	4
SR	MAP (mmHg)	46 ± 5	35 ± 4	38 ± 3	39 ± 2	40 ± 3
	HR (beats min−1)	201 ± 8	179 ± 10	191 ± 10	206 ±4	221 ± 5
	ECC BF (mL min−1)	77 ± 2	72 ± 9	81 ± 9	88 ± 14	93 ± 12
NO+PEG (1:1)	MAP (mmHg)	49 ± 2	39 ± 2	40 ± 5	35 ± 3*	37 ± 1*
	HR (beats min−1)	191 ± 14	195 ± 2	195 ± 3	207 ± 12	213 ± 16
	ECC BF (mL min−1)	62 ± 4	83 ± 3*	84 ± 4*	80 ± 3*	83 ± 4*

Final values are reported as mean ± SEM (n = 4). Intragroup time-course comparisons against BL values are calculated using one-way ANOVA with Tukey’s method.

^*Indicates a significant difference against baseline for the group.

Statistical significance is defined as p ≤ 0.05.

Throughout the 4 h experiment, platelet activation and functionality were also monitored by assessing platelet count, white blood cell count, plasma Fg levels, and pathway activation. The NO+PEG (1:1) circuits led to superior platelet preservation (adjusted for hemodilution and normalized to baseline) at all time points (Fig. 6a). Platelet counts were maintained at near baseline levels for NO+PEG (1:1) circuits after 4 h of blood flow. In contrast, SR circuits exhibited a significant time-dependent loss in counts, dropping to less than < 75% of baseline by 4 h. While both loop configurations appeared to exhibit a modest upward trend in WBC counts (including lymphocytes, monocytes, and granulocytes) with leveling off by 4 h, no significant differences were found (Fig. 6b). This finding was consistent with prior studies of NO-releasing and catalyzing materials.^{[21, 68]}

Fig. 6.

Hematological assessment of circuits was conducted every hour of the ECC flow studies. Circulating (a) platelets and (b) white blood cells (WBC), adjusted for hemodilution, and normalized against baseline showed greater platelet preservation with the NO+PEG (1:1) group with no significant difference in WBC counts. Plasma fibrinogen levels (c) showed a general decrease over time compared to baseline, likely due to hemodilution effects and adsorption to circuits. Clotting assessment of WB showed a significant increase in (d) activated clotting time (ACT) in the SR group compared to baseline (BL). Further pathway activation assessment of isolated plasma showed (e) elevated prothrombin time for both the SR and NO+PEG (1:1) groups as well as (f) elevated activated partial thromboplastin time (aPTT) with the SR group compared to BL values, suggesting pathway dysregulation. Final data reported as mean ± SEM (n = 4 circuits per group, see Section 2.11). Intragroup time-course comparisons against BL values are calculated using one-way ANOVA with Tukey’s method. Time-dependent intergroup comparisons are calculated from two-way ANOVA tests using FDR procedure (see Section 2.12). *Indicates a significant difference between the SR and NO+PEG (1:1) groups at the specified timepoint. ^†Indicates a significant difference against SR control at 1 h. ^‡Indicates a significant difference against NO+PEG (1:1) at 1 h. ^§Indicates a significant difference against SR control at 2 h. ^‖Indicates a significant difference against the SR baseline. ^¶Indicates a significant difference against the NO+PEG (1:1) baseline. Statistical significance is defined as p ≤ 0.05.

Initial plasma Fg levels were 246.6 ± 14.9 and 230.0 ± 14.7 mg dL⁻¹ for SR and NO+PEG (1:1), respectively, and corrected for hemodilution. During the time course of flow studies, the SR group exhibited a significant ~40% drop in plasma Fg levels, whereas the NO+PEG (1:1) group exhibited a ~30% drop by 4 h (Fig. 6c). While these drops are significant within treatment groups, nominal levels are consistent with previous reports for the animal breed.^{[36, 70]} The loss in plasma Fg is associated with its adsorption to foreign surfaces and further role in thrombus formation.^[1] Previously, NO-releasing materials have been associated with increased Fg adsorption and d-dimer formation at the blood-polymer interface as a result of hydrophobic surface interactions and NO evolution.^{[44, 53]} The previously observed reduction in Fg on the NO+PEG (1:1) surfaces (Fig. 1c) is thereby able to act complementary to NO release and facilitate significantly improved platelet preservation and reduction in clot formation. Increased Fg adsorption has been observed on NO-releasing materials proportional to the magnitude of the NO surface flux,^[44] which can attract platelets and facilitate blood contact activation.^{[9, 44]} By reducing the amount of Fg lost from circulation, NO+PEG confers an additional protective effect through surface passivation that prevents Fg attachment and thereby reduces platelet binding. This ensures a dual-action strategy with NO for mediating platelet quiescence and mimicking endothelial homeostasis at the blood-contacting interface.

Further assessment of platelet function first investigated activated clotting time (ACT) for change in hemodynamics as a result of WB stimulation with factor XII activators at each hour mark in the ECC studies. Elevated ACTs were observed for the SR group by study termination (p ≤ 0.05), with a general trend towards elevation in the NO+PEG (1:1) group (p = 0.0513) (Fig. 6d). Elevated ACT is indicative of platelet dysfunction and thrombocytopenia but is typical of the ECC model due to hemodilution effects and concurrent plasma Fg adsorption onto the circuit surfaces.^[71] Further analysis of plasma at each time point showed elevated prothrombin time (PT) by study termination for both groups, indicative of extrinsic and common pathway activation (Fig. 6e). This finding is typical of the ECC surgery, which has the potential to cause injury to the vessel wall and disrupt blood homeostasis, resulting in some inherent platelet activation and other conformational changes.^{[1, 19]}

Testing for activated partial thromboplastin time (aPTT) showed elevation in the SR group but no statistically significant increase in the NO+PEG (1:1) group (Fig. 6f), implying intrinsic pathway activation in the SR control group. This finding idealizes the role of the NO+PEG (1:1) formulation in minimizing blood interaction with the circuit surfaces, thereby reducing the stimuli that induce contact activation of the intrinsic pathway and activation of circulating platelets. NO is an antiplatelet agent known to regulate function through the activation of guanylyl cyclase receptors and prevention of platelet morphological changes via spatial confinement.^[72–74] Concurrently, PEG surfaces are known not to inhibit coagulation processes in native blood but can reduce the extent of activation.^[75]

Gross analysis of the ECC circuits following 4 h implantation demonstrated a significant 79% reduction in thrombus mass on the thrombogenicity chambers with the NO+PEG (1:1) technology compared to the SR controls (p ≤ 0.05, see Fig. 7a). This finding was further corroborated by a 134% increase in chamber patency with NO+PEG (1:1) compared to SR controls (p ≤ 0.05, see Fig. 7b) based on ImageJ analysis of cross-sectional chamber occlusion. Visual inspection of the thrombogenicity chambers showed the emergence of dense, mature clots in the SR controls (Fig. 7c) leading to some degree of occlusion, whereas only thin blood films with some nascent clot coverage were observed with the NO+PEG (1:1) loops after 4 h (Fig. 7d). Taken together, the in vivo findings suggest that delayed thrombus formation on the NO+PEG (1:1) circuits is related to the downregulation of surface contact activation via the promotion of fewer interactions with the circuit walls as well as the facilitation of platelet quiescence via localized NO elution. Study termination at 4 h was consistent with prior work using the rabbit thrombogenicity model, as the heparin-free model often leads to circuit occlusion in controls (Fig. 7c) by study conclusion or requires early termination due to complications from thromboembolism or other systemic effects. While restricted to 4 h circulation, the study provides key insights into the hemocompatibility and antithrombotic potential of the NO+PEG combination strategy, supporting further investigation in longer-term, scaled animal studies with a porcine or ovine model for further evaluation of potential in ECLS technologies.^[1]

Fig. 7.

Analysis of extracorporeal circuits following termination of the study (4 h). Measurements of (a) thrombus mass and further calculations of (b) chamber patency via imaging software demonstrate suppression of aggregate clot formation on the NO+PEG circuits. Representative images of (c) unmodified SR circuits and (d) the NO+PEG (1:1) circuits are provided. Final data are reported as the mean ± SEM (n = 4, see Section 2.11). Endpoint comparisons are calculated using unpaired t-tests assuming equal variance. *Indicates a significant difference against the SR control group. Statistical significance is defined as p ≤ 0.05.

4. CONCLUSION

In summary, this work has investigated the surface properties and translational feasibility of a hydrophilic NO-releasing material strategy to confer enhanced resistance to infection and thrombosis, suitable for dynamic blood flow applications in extracorporeal life support technologies. In this work, the synthetic NO donor SNAP was impregnated into prefabricated SR tubing and surface treated with silanated PEG to confer potent antifouling, antimicrobial, and antiplatelet capabilities. This novel strategy exhibited low water contact angles while retaining NO release kinetics viable for physiological action. This contributed to over a 74% reduction in adsorbed Fg protein and >2-log reduction in adherent bacteria while maintaining the biocompatibility of the base silicone material. Blood compatibility screening of the materials demonstrated a greater than 96% reduction in platelet adhesion with negligible hemolytic potential. Evaluation in a 4-h ECC rabbit model of thrombogenicity demonstrated a 79% reduction in thrombus mass on the test chambers, a 134% increase in chamber patency with the combination NO+PEG strategy, and further highlighted the technology’s ability for superior platelet preservation, maintenance of plasma Fg levels, and reduced perturbations of the coagulation cascade.

The results of this study suggest the potential of improving both infection resistance and hemocompatibility of existing medical devices via a simple two-step fabrication technique suitable for dynamic blood flow applications. Findings support the further investigation and preclinical translation of the NO+PEG technology for extracorporeal applications towards clinical use on the order of hours to days, with strong support for sterilization and storage stability. Generally, the NO+PEG technology is well-suited as a hemocompatible, antifouling, and infection-resistant coating paradigm, but further application-specific requirements should be noted for longer-term and implanted medical device applications such as vascular stents and grafts. In view of these design spaces, caution is warranted to balance the potential immunogenicity of PEG coatings as well as their transient nature. Alternative hydrophilic coatings (e.g., polyglycerol, polysaccharides, polyvinylpyrrolidone, and other zwitterionic polymers) are of strong interest for overcoming these issues of complement activation and transience.^[32] As a facile route towards hemocompatible surface modification, the herein-described NO+PEG strategy offers the unique advantage of a dual bioactive/biopassive approach with acceptable stability over time suitable for extracorporeal application and potential for further adaptation to other membrane oxygenation and blood-contacting technologies.

ABBREVIATIONS

ANOVA

analysis of variance

BCMD

blood-contacting medical device

CCK-8

cell counting kit-8

EDTA

ethylenediaminetetraacetic acid

ECC

extracorporeal circuit

ECLS

extracorporeal life support

ECMO

extracorporeal membrane oxygenation

ISO

International Organization for Standardization

NO

nitric oxide

OD

optical density

PBS

phosphate-buffered saline

PEG

polyethylene glycol

PPP

platelet-poor plasma

PRP

platelet-rich plasma

SNAP

S-nitroso-N-acetylpenicillamine

SR

silicone rubber

THF

tetrahydrofuran

WB

whole blood