Smartphone compatible nitric oxide releasing insert to prevent catheter-associated infections

Abstract

A large fraction of nosocomial infections is associated with medical devices that are deemed life-threatening in immunocompromised patients. Medical device-related infections are a result of bacterial colonization and biofilm formation on the device surface that affects >1 million people annually in the US alone. Over the past few years, light-based antimicrobial therapy has made substantial advances in tackling microbial colonization. Taking the advantage of light and antibacterial properties of nitric oxide (NO), for the first time, a robust, biocompatible, anti-infective approach to design a universal disposable catheter disinfection insert (DCDI) that can both prevent bacterial adhesion and disinfect indwelling catheters in situ is reported. The DCDI is engineered using a photo-initiated NO donor molecule, incorporated in polymer tubing that is mounted on a side glow fiber optic connected to an LED light source. Using a smartphone application, the NO release from DCDI is photo-activated via white light resulting in tunable physiological levels of NO for up to 24 h. When challenged with microorganisms S. aureus and E. coli, the NO-releasing DCDI statistically reduced microbial attachment by >99% versus the controls with just 4 h of exposure. The DCDI also eradicated ~97% of pre-colonized bacteria on the CVC catheter model demonstrating the ability to exterminate an established catheter infection. The smart, mobile-operated novel universal antibacterial device can be used to both prevent catheter infections or can be inserted within an infected catheter to eradicate the bacteria without complex surgical interventions. The therapeutic levels of NO generated via illuminating fiber optics can be the next-generation biocompatible solution for catheter-related bloodstream infections.

1. Introduction

Intravascular (IV) catheters are fundamental to contemporary hospital practices and are frequently implanted in critically ill patients for the administration of drugs, fluids, blood transfusion, dietary solutions, and for hemodynamic monitoring. Approximately, 90% of all the patients admitted to the hospital encounter some sort of intravenous therapy during their hospital stay [1]. The typical duration of a catheter used in clinical settings like emergency rooms, operating theaters, and intensive care units (ICUs) ranges from minutes to months. While the acute catheters are used for shorter periods (weeks), longer insertion periods of a catheter, like those used in hemodialysis, may be used from several months to years [2]. Among all the medical devices used in a hospital setting, IV catheters account for an increased risk of device-related infections compared to any other medical device categories. The infection-causing bacteria can adhere to the catheter surface and colonize to develop biofilms. The primary contact of the bacterial cells on the surface of the catheter can emanate from the patient’s own skin flora which can colonize the catheter lumen triggering the bacteria to travel from the catheter insertion site into the vascular [3,4]. Furthermore, hematogenous seeding from a different contaminated site can become a source of infection in catheters. In rare occasions, catheter lumen contamination can occur when the infusate is contaminated [5]. These are the predominant sources of morbidity and mortality in patients contributing to conditions like catheter-related bloodstream infections (CRBSIs) (e.g., bacteremia and sepsis). Each year the occurrence of CRBSIs in the United States alone is estimated to be >250,000 incidences with a mortality rate of approximately 35%, where the cost of each catheter infection is estimated to be approximately $34,000–56,000 and results in an annual medical care outlay of approximately $2.3 billion [6–9].

Catheter-related infections have escalated especially due to the formation of biofilms [10]. As per the reports, over 1 million cases of hospital-acquired infections are reported every year [11]. About 60–70% of these infections are identified to arise from bacterial contamination and biofilm formation on the surface of the medical device which severely compromises the durability of the medical devices [12]. The steps to address a catheter infection typically include irrigation of the infection site with antibiotics, removal of the catheter, and initiation of antibiotic lock solution therapy [13]. Bacteria protected within biofilms require up to a 1000 times higher dosage of antibiotics than their free-floating (planktonic) counterparts [14]. This high dosage can increase the possibility of antibiotic resistance across the bacterial species, engender a great deal of economic burden, and is a threat to native beneficial bacteria and other healthy organs of the body [15,16]. Therefore, there is an urgent need for efficacious and harmless approaches that can not only handle the emergence, but also the propagation of pathogenic microorganisms. To overcome this issue and enhance the bactericidal effect of medical devices, several antimicrobial strategies including silver-doped catheters, incorporation of antibiotics, or antimicrobial peptide coatings are employed to thwart the replication of bacteria or increase the susceptibility of antibiotics [17–21]. Likewise, IV line devices with potent antibacterial activity against various bacterial and fungal strains via alcohol impregnation have been reported in the literature [22–25]. Many of these approaches have been extensively tested, evaluated in vivo, have reached clinical stages, and are commercially available. However, while many of these approaches are effective against planktonic bacteria, they remain ineffective in reducing biofilms [26]. Similarly, even the most robust antibiotics are growing ineffective against biofilm-forming microorganisms. Current clinical guidelines oppose the idea of prolonged use of antibiotic-impregnated catheters mainly due to the problem of uncontrolled leaching of antibiotic species from implanted devices, related toxicity, and emergence of antibiotic resistance [27]. Despite all these efforts, CRBSIs still remain one of the most significant concerns pertaining to biomedical devices as there is no definite solution for resisting biofilm that meets the prerequisite of clinically suitable catheter sizes.

Nitric oxide (NO) based therapy is emerging as a potential antibacterial treatment due to its bacteria-killing and biofilm dispersing abilities [28,29]. Nitric oxide is an innate signaling diatomic molecule utilized by the body’s defense system for fighting infection-causing microorganisms, preventing platelet activation, reducing localized and chronic inflammation, and enhancing wound healing [30]. The endogenous synthesis of NO in the body happens via NO synthase (NOS) enzymes which convert the amino acid L-arginine into citrulline and NO. [31,32] Macrophages and neutrophils utilize NO synthesized via the inducible nitric oxide synthase (iNOS) enzyme to eradicate the invading pathogens in the body by promoting biofilm dispersal and preventing the adherence of planktonic bacteria. Nitric oxide donor molecules, like S-nitrosothiols (RSNO), incorporated into a polymer substrate can mimic endogenous NO release levels, such as endothelial cells that release NO at a surface flux of 0.5–4 × 10⁻¹⁰ mol cm⁻² min⁻¹ to prevent platelet activation and adhesion and exhibit broad-spectrum antimicrobial properties [33,34]. Considering the potential benefits of endogenous NO, various studies have been designed that can utilize these benefits synthetically by either incorporating or impregnating the NO donors in the polymer matrix that will release their NO payload [35–37] or using a generation mechanism to stimulate the release of endogenous NO in blood [38].

The release of NO from polymeric substrates has been tremendously explored in the past two decades demonstrating its wide range of tunable properties for achieving controlled NO release depending on the trigger mechanism. Therefore, to enhance the NO payload and extend the lifetime of NO release, several distinct frameworks have been designed with NO-releasing mechanisms at the polymer interface. Such engineered polymer surfaces that can release NO have been comprised of physical dispersal of NO donors into the polymer substrate or covalent conjugation of NO donors to the polymer backbone [39–42]. The RSNO donors, like S-nitroso-N-acetylpenicillamine (SNAP), have been recognized to have extended storage stability in the crystallized form and can emit NO either photochemically, thermally by heat, or metal ions (Cu²⁺, Se, Zn, etc.). Photocatalytic release of NO from RSNOs and RSNO-based polymers have been explored [43,44]. The characteristic absorption maxima for the RSNOs occurs at wavelengths 340 nm and 520–590 nm corresponding to the π → π* and n → π* electronic transitions of the SNO functional group that has been primarily associated with their decomposition [45].

Encouraged by the promising capabilities of NO and light-mediated microbe killing, a novel smartphone-based Disposable Catheter Disinfection Insert (DCDI) is engineered in this study which, for the first time, combines a NO-releasing polymer and side glowing fiber optics. The NO-release polymer tubing was prepared by impregnating commercial silicone tubing with a light-activated NO donor molecule [2]. The NO-releasing polymer tubing is mounted on a side glowing fiber optic to illuminate the full length of the NO-releasing tubing and is connected to a light source (Fig. 1A). To activate the NO release and eradicate the viable pathogenic bacteria, the antimicrobial DCDI can be inserted within the lumen of IV catheters (Fig. 1B) and can be controlled via Bluetooth using a smartphone application. When the DCDI is powered by Bluetooth connected light source, the side glow fiber optic will illuminate, and the decomposition of SNAP can be triggered under irradiation by LED with a wavelength of 450–650 nm to generate NO (Fig. 1C). Taking this into consideration, NO release kinetics from the DCDI using an LED source in dark and with different nominal lights and intensities was evaluated using the chemiluminescence detection method. Using the UV–vis spectroscopy method, the amount of NO donor impregnated in the SR polymer, leaching from the device with and without light, and stability with various sterilization methods were investigated. The antibacterial efficacy of DCDI was studied using a 4 h bacterial adhesion assay against two prominent bacterial strains associated with CRBSIs, Gram-positive (S. aureus) and Gram-negative (E. coli). The DCDI was also evaluated in a more challenging in vitro infection model to evaluate its ability to disinfect an S. aureus infected catheter, which closely mimics the end-use clinical application of the DCDI device. The small DCDI is envisioned to be a part of the catheter lumen cap and inserted within the lumen of IV catheters between clinical use of the IV catheters (when not in use by clinicians for blood draws, infusion, and other fluid administration). Additionally, in clinical applications, the use of personalized smartphone-based devices can reduce the risk of disease transmission among patients in a hospital environment. Smartphone-based devices help in simplifying the healthcare system since these applications are widely used and offer effective ease of use in patients. In some cases, patients, such as those with a hemodialysis catheter, could control the DCDI at home using their smartphone application. Once the NO release is depleted, a new DCDI can easily be replaced which extends the NO release capability at the catheter interface to the entire indwelling lifetime of the IV catheter. It is expected that light as a catalyst from the fiber optic combined with the NO-releasing SR will enhance the antimicrobial activity of NO released from the catheter by both disinfecting the catheter in situ and preventing impending infections (Fig. 1D). Further, this DCDI device is immediately applicable to a wide range of catheters currently used in clinical applications (e.g., vascular, hemodialysis, urinary catheters).

Fig. 1.

Design and functional use of the NO-releasing Disposable Catheter Disinfection Insert (DCDI). (A) NO-releasing SR is mounted on a side glow fiber optic and connected to a LED light source to develop the DCDI device. (B) The DCDI device can then easily be integrated with an indwelling catheter (intravascular, urinary, etc.) for continuous decontamination in between clinical uses. (C) The DCDI is engineered using a light-sensitive NO donor molecule, S-nitroso-N-acetylpenicillamine (SNAP), incorporated in polymer tubing which is mounted on a side glow fiber optic. The photosensitivity of SNAP can be exploited using an LED light source that can illuminate the side glow fiber optic using a simple mobile phone application that activates and enables real-time control of the NO release levels. (D) The DCDI can both eradicate catheter infections and prevent Catheter-related bloodstream infections (CRBSIs) thereby extending the usage lifetime of medical devices and drastically reducing associated treatment costs.

2. Experimental section

2.1. Materials

N-Acetyl-D-penicillamine (NAP), sodium nitrite, L-cysteine, sodium chloride, potassium chloride, sodium phosphate dibasic, potassium phosphate monobasic, copper (II) chloride, ethylenediaminetetraacetic acid (EDTA), tetrahydrofuran (THF), and sterile phosphate buffer saline powder with 0.01 M, pH 7.4, containing 0.138 M NaCl, 2.7 mM KCl, were purchased from Sigma Aldrich (St. Louis, MO). Methanol, hydro-chloric acid, and sulfuric acid were obtained from Fisher Scientific (Hampton, NH). Helixmark^® silicone tubing (60–011-06 and 60—011-09) was purchased from VWR (Radnor, PA). PMMA side glow optical fiber (Huaxi, Amazon.com), 12 V 1.5 W LED light source (Rayauto, Amazon.com), and LED BLE Bluetooth 4.0 software were used for the light studies. All aqueous solutions were prepared using deionized water. Phosphate buffer saline (PBS) 0.01 M with 100 μM EDTA was used for all material characterization and NO analyzer studies. Dulbecco’s modified Eagle’s medium (DMEM) and trypsin-EDTA were purchased from Corning (Manassas, VA20109). The Cell Counting Kit-8 (CCK-8) was purchased from Sigma-Aldrich (St. Louis, MO). Penicillin-Streptomycin (Pen-Strep) and fetal bovine serum (FBS) were obtained from Gibco-Life Technologies (Grand Island, NY). The bacterial strain S. aureus (ATCC 6538), E. coli (ATCC 25922), and 3T3 mouse fibroblast cells (ATCC 1658) for cell compatibility experiments were obtained from American Type Culture Collection (ATCC). All the buffers and media were sterilized in an autoclave at 121 °C, 100 kPa (15 psi) above atmospheric pressure for 30 min prior to bacteria and biocompatibility experiments.

2.2. S-nitroso-N-acetylpenicillamine synthesis

S-nitroso-N-acetylpenicillamine (SNAP) synthesis procedure was adapted from a previously published report with a slight modification [42]. Briefly, NAP was dissolved in a 2:3 ratio of water and methanol. To this mixture, 0.7 and 1.6 M of H₂SO₄ and HCl were added, respectively followed by dropwise addition of sodium nitrite dissolved in water and stirring for 10 min at room temperature. The beaker was shielded from ambient light and incubated in an ice bucket with continuous nitrogen purging for 8 h. After incubation, the filter paper was used to collect the SNAP crystals into a Buchner funnel using vacuum filtration. The SNAP crystals were then rinsed with ice-cold DI water and placed in a vacuum desiccator overnight allowing the product to dry. Care was taken throughout the process to shield the samples from light. Each batch of synthesized SNAP was tested for its purity (quantifying the conversion of the parent thiol RSH moiety to the NO-rich RSNO moiety) using the chemiluminescence NOA and UV–vis calibration curve with a characteristic maximum absorbance peak of the SNAP molecule at 340 nm using the previously reported methods [46,47]. All batches of SNAP used in the study had recorded purity levels of >90%.

2.3. Preparation of NO-releasing polymer

The NO-releasing polymer was prepared by impregnating the silicone rubber (SR) tubing with SNAP,.A stock solution of SNAP and THF (125 mg mL⁻¹) was prepared based on previously reported methods [2]. A 3 cm long Helix Silastic SR tubing with an inner diameter of 0.058″ was incubated in SNAP-THF solution for 24 h in dark at room temperature. After 24 h, the SNAP-impregnated NO-releasing SR (SR-NO) was removed from the solution and dried overnight in a vacuum desiccator protected from light. All samples were rinsed with PBS to remove excess SNAP crystals from the outer surface and lumen of the impregnated tubing before conducting any further experiments.

2.4. Determining wt% SNAP using UV–vis spectroscopy

The amount of SNAP impregnated in the SR-NO tubing was quantified using a UV–vis spectrophotometer (Cary 60, Agilent Technologies). For this, first, the mass of each SR-NO sample was recorded using an analytical balance (Mettler Toledo^™ XS105DU, Columbus, OH). Each SR-NO sample was soaked in THF for 30 min to extract all the impregnated SNAP from the SR tubing. The tubing appeared clear after incubation in the THF, indicating all SNAP had been extracted into the THF. The SNAP extracted solution was evaluated by UV–vis spectroscopy at 340 nm wavelength. The molar absorptivity of SNAP in THF at 340 nm was determined to be 909 M⁻¹ cm⁻¹. The weight percentage (wt%) of SNAP loaded is reported as milligrams of SNAP loaded per milligram of tubing.

2.5. Fabrication of disposable catheter disinfection insert (DCDI)

The Disposable Catheter Disinfection Insert (DCDI) was produced using a 2.9 cm SR-NO tubing mounted on a 7 cm segment of 1.5 mm diameter PMMA side glow optical fiber (Huaxi, Amazon.com). The SR-NO-Light DCDI samples were connected to a 12 V 1.5 W LED light source and controlled via Bluetooth using a smartphone application. For controls, either unmodified SR or SR-NO were mounted on the fiber optic and operated with or without light, resulting in the SR, SR-Light, and SR-NO control groups.

2.6. Light emission spectroscopy measurements

To determine the wavelength of light emitted by the LED light source, a wireless spectrophotometer (PS-2600, PASCO Scientific) with a detection range of 380–950 nm was utilized. The light detecting fiber optic cable was held in place with a clamp, the DCDI samples were exposed to the detector, and the wavelength of light was recorded with four different colors (red, blue, green, and white) in addition to testing in the absence of light (0% intensity). Studies with light were done in the absence of ambient light to ensure only desired lights were being characterized.

2.7. Photoinitiated NO release

The NO release from samples with light (SR-NO-Light) and without light (SR-NO) was quantified using the gold standard Zysense chemiluminescence Nitric Oxide Analyzer (NOA) 280i (Frederick, CO) under a nitrogen atmosphere under physiological conditions. The NO release from the samples is normalized to the surface area of the DCDI and is presented as moles min⁻¹ cm⁻².

2.7.1. NO release vs. light color

In order to optimize the light from DCDI, samples were placed in an NOA sample cell at 37 °C connected to the LED light source (SR-NO-Light). To avoid the interference of leached SNAP in buffer conditions, a dry state under physiological conditions was used to study the influence of white light on DCDI at different light intensities. Using the smartphone application, the LED light source was set to emit light at 100% intensity, and the NO flux was recorded as the light color was adjusted. Four different light colors (white, blue, green, and red) were tested for their NO release.

2.7.2. Tunable photoinitiated NO release

The SR-NO-Light samples were placed in an NOA sample cell under a dry state at 37 °C protected from ambient light. The LED light source was set to emit white light from the connected fiber optic and the NO flux from samples was recorded as the light intensity was adjusted using the smartphone application. Starting without light (in dark), the intensity of light was increased in 25% increments until 100% intensity was reached. The light intensity was then decreased by 25% increments to 0% to demonstrate control over NO levels.

2.7.3. 24 hour NO release

The DCDI samples were immersed in PBS with EDTA (7.4 pH) at the physiological temperature of 37 °C in an amber NOA sample cell. For the SR-NO-Light samples, the LED light source was set to emit white light at 100% intensity. The 24 h NO release profiles for the DCDI samples were recorded using 0% (light off) and 100% white light intensity from SR-NO and SR-NO-Light samples, respectively.

2.8. Determination of SNAP diffusion

The amount of SNAP leached from samples in the presence (100% light intensity) and absence of light (0% light intensity) was determined by a UV–vis spectrophotometric method. Each sample SR-NO and SR-NO-Light was incubated in 10 mM PBS, pH 7.4, with 100 μM EDTA at 37 °C for 24 h. The soaking buffer was evaluated for SNAP concentration at 2, 4, 6, 8, and 24 h time points. The molar absorptivity of SNAP in 10 mM PBS, pH 7.4, with 100 μM EDTA at 37 °C was determined to be 1072 M⁻¹ cm⁻¹ at 340 nm. Samples were incubated at 37 °C in PBS throughout the duration of the experiment. Results were analyzed by calculating SNAP concentration from each sample, normalized by the surface area of DCDI.

2.9. Sterilization of NO-releasing tubing

Sterilization of medical devices is an important process for decontaminating medical device surfaces before in vivo application. Therefore, it is very critical for medical devices to withstand the sterilization process without compromising the desired properties. In this study, two common sterilization methods, ethylene oxide (EO) and ultraviolet (UV) light, were compared to identify the most compatible method with the NO-releasing polymer utilized in the DCDI device. For EO sterilization, the NO-releasing DCDI inserts were packaged into the sterilization pouch and exposed to EO under AN 74i Anprolene EO gas sterilizer (Anderson Sterilizers). The sterilizer was operated at room temperature (between 68 and 91 °F) with a Humidichip to ensure at least 35% humidity was achieved. Samples were sterilized for 12 h with 2 h of purging with ambient air. Similarly, for UV sterilization, SR-NO samples were sterilized with UV light under a biosafety cabinet administered by REDISHIP Purifier^® Logic^® + Class II A2 Biosafety Cabinets, Labconco^® for 30 min. Upon sterilization, all the samples (n = 4) were weighed and then suspended in THF (30 min) to fully extract the SNAP from the polymer. The amount of SNAP remaining in the SR-NO tubing after the sterilization process was compared to fresh samples using UV–vis spectroscopy by measuring the absorbance of SNAP extracted in THF at 340 nm wavelength. The results of the study are reported as normalized values compared to the initial wt% of SNAP (weight of SNAP/weight of polymer). To confirm the activity of SNAP in the SR-NO samples, the samples were also tested for their NO release levels using the same methodology as Section 2.7.2 using fresh samples and samples after sterilization.

2.10. Shelf-life storage of DCDI

To evaluate the long-term stability of DCDI at various storage conditions, SNAP-impregnated SR were stored at −20 °C, 4 °C, and RT for 30 d in a tightly closed vial, in the dark, and with desiccant to protect the DCDI from moisture. After 30 d, the SR-NO tubing was mounted on a fiber optic, connected to a light source, and the NO release from the samples before and after storage was evaluated using chemiluminescence NOA at 37 °C. The NO release from samples was recorded in the absence (0%) and presence of light (100% white light) under physiological conditions.

2.11. In vitro antibacterial evaluation

2.11.1. Antibacterial activity of DCDI using 4 h bacterial adhesion assay

Studies were conducted to determine the efficacy of the DCDI to prevent bacterial adhesion on the surface. The antibacterial activity of DCDI was examined against S. aureus (ATCC 6538) and E. coli (ATCC 25922) in terms of viable bacterial adhesion on the catheter surface. Individual colonies of S. aureus and E. coli were inoculated in LB media and grown until mid-log phase on a shaking incubator at 120 rpm, at 37 °C for 5 h. All DCDI samples (SR, SR-Light, SR-NO, and SR-NO-Light; n = 3 each) were exposed to the bacterial solution with the final OD₆₀₀ of bacteria ranging between 10⁶ and 10⁸ CFU mL⁻¹ for 4 h at 150 rpm at 37 °C to maintain the chronic infection conditions. After 4 h incubation, bacteria adhered to the DCDI surface were homogenized to detach, diluted, and plated using a spiral plater (Eddy Jet 2, IUL Instruments). Viable colony forming units (CFU) were determined after 24 h of incubation at 37 °C using an automated bacteria colony counter (Sphere Flash, IUL Instruments). The CFUs on the samples and individual controls were normalized by the surface area of the samples and the percentage of reduction in bacterial viability was determined by the following equation (with respect to SR control), where C represents the concentration of viable bacteria in CFU cm ⁻²:

$$\%\mathit{\text{bacterial}}\mathit{\text{reduction}}=\frac{\left(C_{SR}\right)-\left(C_{\mathit{\text{Test}}}\right)\times 100}{\left(C_{SR}\right)}$$

2.11.2. In situ catheter disinfection using catheter infection model

To evaluate the efficacy of DCDI in disinfecting an infected model medical catheter (Helixmark^® silicone tubing 60–011–09) with an ID of 0.078″ and OD of 0.125″ was used to develop an in vitro assay using S. aureus bacteria (ATCC 6538). S. aureus was grown overnight in an LB media following the same procedure as in Section 2.11.1. Then, 3 mL of bacterial suspension (0.1 optical density) was added to the model CVC catheter (Helix-09 tubing; 3 cm long) such that both intraluminal and extraluminal surfaces were exposed to bacteria. Samples were incubated for 24 h in LB medium under stagnant conditions at 37 °C. Nutrient media in the tubes was changed periodically to keep the supply of nutrients stable. After 24 h, catheter samples were taken out from media containing bacteria and briefly rinsed to remove any unadhered cells on the catheter tubing. Then the DCDI (SR-NO-Light) and control (SR) devices were inserted into the infected catheter tubing and incubated in PBS for 4 h at 37 °C under static conditions. The control samples contained no NO-releasing DCDI, just an unmodified SR tubing mounted on the fiber optic was inserted into the bacteria-infected catheter. Samples were removed after incubation and the bacteria remaining on the catheter tubing (both inside and outside surfaces) were enumerated using the same CFU counting protocol described in Section 2.11.1.

2.12. In vitro cytocompatibility study

2.12.1. Leachate preparation

All test and control groups (SR, SR-NO, SR-Light, and SR-NO-Light samples; n = 3) were first cleaned with ethanol and UV-sterilized for 30 mins. Next, DCDI samples were incubated in complete DMEM media (1 mL) to collect the leachates in the solution by following the ISO standards (ISO10993–5:2009 Test for in vitro cytotoxicity assessment of biomedical devices). All the vials were covered in aluminum foil to prevent the effect of ambient light and incubated for 24 h at 37 °C. After 24 h, the samples were removed, and the leachates were used for further analysis.

2.12.2. Cytocompatibility of DCDI

A cell culture treated 96-well plate was used to seed 3T3 mouse fibroblast cells (10,000 cells/mL) in each well and incubated in a humidified incubator at 37 °C, 5% CO₂ for 24 h. Later, leachate samples were exposed to the cells (100 μL) and incubated for another 24 h to let the leachates act on the cells. The cytocompatibility of samples was analyzed using a CCK-8 cell viability kit following the manufacturer’s instructions (Sigma Aldrich). To each of the wells, CCK-8 solution was added (10 μL) and incubated for 1 h. The absorbance (A) of the samples was recorded at 450 nm wavelength using a microplate reader (Cytation 5 imaging multi-mode reader, BioTek). Results from the experiment are reported as relative cell viability of the test group normalized to control (cells in media) using the following equation:

$$\mathit{\text{Relative}}\mathit{\text{cell}}\mathit{\text{viability}}=\frac{A_{\mathit{\text{test}}\mathit{\text{group}}}}{A_{\mathit{\text{cells}}\mathit{\text{control}}}}$$

2.13. Statistical analysis

All results in the study are presented with a sample size n ≥ 3. Data are all reported as mean ± standard error of the mean (SEM). Statistical significance between the sample types was determined using a student’s t-test. To ascertain the significance of the results, a value of p < 0.05 was used to evaluate statistical differences between the test (SR-Light, SR-NO, SR-NO-Light) and control groups (SR).

3. Results and discussion

3.1. Fabrication of the DCDI

To generate the NO-releasing DCDI insert, segments of SR tubing were soaked in the SNAP-THF impregnation solution for 24 h (125 mg mL⁻¹) (Fig. 2A). The solvent impregnation process is one of the simple and effective ways to incorporate NO donors in the polymer matrix. Incorporation and impregnation of SNAP in various polymers, such as polyurethanes and silicone elastomers, have been reported and studied for their clinical applications for devices including intravascular catheters, urinary catheters, insulin cannulas, and extracorporeal circuit tubing [36,48–50]. These polymers have exhibited uniform SNAP impregnation [47,49] long-lasting and controlled NO release [46,47,51,52], with enhanced shelf-life [50,53] and sterilization [34,46] stability, ability to withstand low rates of SNAP leaching, and photosensitivity [54]. This impregnation approach helps conquer challenges with the typical style of polymer manufacturing procedures that use elevated temperature possessing (i.e., polymer extrusion), which could negatively influence the thermally sensitive NO donors, like SNAP. In this study, the amount of SNAP impregnated into SR was confirmed by the UV–vis spectroscopy method which revealed 4.66 ± 0.16 wt% of SNAP impregnated into the SR-NO samples (Fig. 2B). The values obtained in this study are consistent with the previously reported values for SNAP impregnation of SR that accomplished ca. 5 wt% with the same concentration of SNAP-impregnation-solvent (125 mg mL⁻¹) [49,50].

Fig. 2.

(A) NO-releasing tubing is fabricated by soaking the SR tubing in SNAP-THF solution (125 mg mL⁻¹) for 24 h followed by drying in a vacuum desiccator for 24 h. (B) Quantification of SNAP impregnation in SR-NO samples is determined by UV–vis spectroscopy. Data represents mean ± SEM for n ≥ 3. (C) To develop the NO-releasing Disposable Catheter Disinfection Insert (DCDI), SR-NO tubing is mounted on a side glow fiber optic and connected to a LED Light source. Examples of the DCDI illuminated with different nominal lights, from left to right: light off, red, green, blue, and white light. Verification of wavelength of light emitted by LED light source (D) red (621 nm) (E) green (512 nm), (F) blue (447 nm), and (G) white (mixture of red, green, and blue) set at 100% light intensity.

3.2. Quantification and control of real-time NO release

3.2.1. NO release vs. light wavelength

To confirm the wavelength of light emitted by the LED source, a PASCO wireless spectrometer optical probe was utilized. The wavelength of light was determined at four different colors of light (Fig. 2C). Results confirmed that the red, green, and blue light had emissions ranging from 570 to 650 nm, 475–575 nm, 450–500 nm, respectively (Fig. 2 D–F). Furthermore, the study confirmed the white light provided by the light source is comprised of red, blue, and green wavelengths of light (Fig. 2G).

The NO donor SNAP can catalytically release NO upon exposure to heat, light, or metal ions (Fig. 3A). The small side glow fiber optic used in this study enables compatible application with the IV catheter. Fiber optics are used in clinical applications such as sterilization of medical device surfaces. The illuminating light has been shown to be compatible with endoscopes, respiratory devices, catheters (endotracheal tube, urinary catheters), and bandages for wound healing [55]. The thin, flexible nature, and tubular scattering factor make the light from fiber optic cable of uniform light dissemination. Therefore, a side glow fiber optic is utilized in the DCDI due to its significant advantages over end glow fiber optics in order to illuminate the full length of SNAP tubing for catheter applications. The DCDI can be inserted within the vascular catheters to disinfect the entire length of the catheter tubing (Fig. 3B).

Fig. 3.

(A) Chemical structure of the NO donor S-nitroso-N-acetylpenicillamine (SNAP). RSNOs like SNAP can be triggered by the stimulus of heat, light, or metal ions to cleave the S—N bond and release NO. (B) Cross-section of the Disposable Catheter Disinfection Insert (DCDI) device comprised of SNAP-impregnated SR tubing mounted on a side glow fiber optic. (C) Comparison of steady-state NO release from DCDI at physiological temperature (37 °C) in dark and photoinitiated at 100% light intensity of red (620 nm), green (530 nm), blue (450 nm), and white (mixture of red, green, and blue) light. (D) Representative example of tunable NO release via increasing and decreasing intensities of light (between 0% and 100%). (E) Quantification of NO release using chemiluminescence measured with the trigger of different light intensities at 37 °C (n ≥ 3). (F) Determination of real-time NO release from NO (dark) and SR-NO-Light (100% light intensity) using a chemiluminescence NO analyzer. The NO flux levels were measured at physiological conditions in PBS with 100 μM EDTA up to 24 h. (G) Quantitation of amount of SNAP present in the PBS (soaking buffer) from SR-NO and SR-NO-Light samples at 37 °C in dark and 100% white light intensity conditions, respectively. Data normalized to the surface area of the polymer. All data are reported as mean ± standard error of mean (n ≥ 3). (H) DCDI soaked in PBS-EDTA from left to right: DCDI without light on day 0, DCDI with 100% white light on day 0 and day 1 (24 h).

To optimize the light color for the study, NO release from the DCDI was tested against various colors of light red (620 nm), green (530 nm), blue (450 nm), and white (a mixture of red, green, and blue light) at 100% light intensity (n ≥ 3). DCDI samples were inserted in the amber NOA sample cell to protect the samples from ambient light. First, the NO release from the samples was recorded in the absence of light (dark) and then the color of the light was changed, and the intensity was adjusted using the mobile phone application. The NO release in dark was 0.09 × 10⁻¹⁰ mol cm⁻² min⁻¹ flux, and the red, green, blue, and white light at 100% light intensity triggered 0.14, 0.17, 1.23, and 1.69 × 10⁻¹⁰ mol cm⁻² min⁻¹ of NO from DCDI, respectively (Fig. 3C). The hemolytic cleavage of S—N bond of S-nitrosothiols has been well-established in the visible light spectrum. Once the results were confirmed with white light resulting in the maximum levels of NO, all further studies were conducted with white light. These results are consistent with previously published reports that utilized white light to maximize the NO release from the NO donating compound [56,57].

3.2.2. Real-time control of NO release

NO-releasing materials hold a huge potential in clinical applications. However, it is crucial to gain the ability to dynamically control the NO levels depending on the biomedical application. For instance, at the time of implantation, the catheter may require a higher level of NO to prevent bacterial colonization on the device surface. However, over time the same device may need lower levels of NO to maintain the biofilm-free state of the device. Similarly, a significantly contaminated device surface may require very high levels of NO to disinfect and eradicate the pre-established biofilm on the catheter surface. This modulatory capacity of NO release can be achieved by exploiting the photosensitivity of NO donating compounds that can tune the NO release in response to the intensity and wavelength of light. Based on this, other RSNOs like GSNO, N-nitrosoamine-based NO donors with a combination of antibiotics, or even modified RSNOs have been studied for their light-sensitive properties [58–61].

The ability to control the release of NO by changing the light intensity was assessed by increasing and decreasing the white light intensity in 25% intervals (Fig. 3D). These results demonstrate that each degree of light intensity can modulate the NO release levels from the DCDI in real-time (Fig. 3E). The NO release levels at each light intensity are tabulated in Table 1. The advantage of the DCDI design is that the NO release levels can be controlled via adjusting the intensity of the light source connected to the fiber optic. This data demonstrates that using light as a trigger to potentiate the NO release enables accurate modulation of the NO levels as required to achieve antibacterial properties.

Table 1.

NO release levels measured from SR-NO-Light DCDI at different intensities of white light at 37 °C using chemiluminescence nitric oxide analyzer. Data represents mean ± SEM (n ≥ 3).

Light intensity	NO flux (× 10−10 mol min−1 cm−2)
0%	0.06 ± 0.01
25%	0.46 ± 0.03
50%	0.85 ± 0.05
75%	1.25 ± 0.08
100%	1.66 ± 0.14
75%	1.31 ± 0.13
50%	0.92 ± 0.09
25%	0.50 ± 0.05
0%	0.08 ± 0.02

3.2.3. Real-time NO release

The NO release from the DCDI was measured under physiological conditions (37 °C in PBS buffer) using chemiluminescence for 24 h at 0% (dark), and 100% white light intensity. NO release from the samples was investigated over 24 h with the idea that the DCDI device could be replaced daily during clinical catheter applications. The average initial NO flux for 0% and 100% light intensity was found to be 3.92 × 10⁻¹⁰ mol cm⁻² min⁻¹ and 9.01 × 10⁻¹⁰ mol cm⁻² min⁻¹, respectively, demonstrating that the light intensity is directly proportional to the level of NO released from DCDI on day 0 (Fig. 3F). However, all samples ultimately reach an equivalent NO release level at 24 h timepoint as the SNAP payload becomes depleted. The SR-NO-Light DCDI samples have a fixed amount of NO and will exhaust over time as NO is constantly discharged. The 100% light intensity significantly increases the rate at which NO is depleted from the samples early in the experiment which explains the decrease in the NO flux level at the 24 h time point. Overall, the DCDI with and without light are able to closely mimic the levels of NO released by endothelium (0.5–4 × 10⁻¹⁰ mol cm⁻² min⁻¹) even after 24 h. These levels of NO are known to exhibit important biological functions such as reducing inflammation and fibrosis, killing various microbial species (bacteria, fungus, viruses), inhibiting, disrupting, and dispersing microbial biofilm formation, preventing platelet activation, reducing clotting and thrombosis [33]. Once the NO flux is below physiological levels, a new DCDI could be easily replaced within the catheter periodically (e.g., daily).

3.3. Quantification of SNAP in soaking buffer

To quantify the amount of SNAP leached from NO-releasing samples, both SR-NO (0% light) and SR-NO-Light (100% white light) DCDI samples were incubated in PBS-EDTA at 37 °C for 24 h. The soaking buffer was collected at various timepoints, and the absorbance was recorded using UV–vis spectroscopy. After 24 h, 84.61 ± 5.32 and 35.83 ± 2.04 μg cm⁻² of SNAP were detected from SR-NO and SR-NO-Light samples, respectively (Fig. 3G). The introduction of light to the SR-NO-Light significantly enhanced the catalysis rate to release NO, resulting in a lower concentration of SNAP in the soaking buffer as compared to the SR-NO samples. Overall, the low SNAP leaching results observed in this experiment demonstrate that the NO released is localized at the DCDI polymer interface. The fiber optic also showed the ability to be illuminated in buffer even after a day of soaking (Fig. 3H).

Impregnating the NO donor SNAP into hydrophobic polymers like SR has been reported to significantly reduce leaching which consequently prolongs the NO release from the polymer [47,62]. Due to the intra-molecular hydrogen bonding between SNAP molecules and the low water uptake of hydrophobic polymers, SNAP dissolution and dissemination out of the polymer is significantly contained [2,53]. The SNAP leaches out of the polymer and eventually degrades into N-acetyl-D-penicillamine (NAP) and NAP-dimers. It is anticipated that the levels of SNAP detected in the buffer are observed to be lower when the fiber optic is illuminated due to the relatively faster light-mediated conversion of SNAP to NAP disulfide [36]. These degradation products of SNAP are non-toxic and an FDA approved drug utilized to treat heavy metal poisoning [63,64].

3.4. Sterilization of SNAP impregnated SR

Infection-causing pathogens can easily adhere to the medical device surfaces in the healthcare setting. If left untreated, these pathogens can migrate into the patient’s body and can lead to bloodstream infections and, in severe scenarios, increase healthcare costs and prolong the patient’s treatment time. Therefore, sterilization of medical devices is an important process for decontaminating medical device surfaces before clinical application [65]. It is extremely critical for medical devices to withstand the sterilization process without compromising the desired properties. NO donating compounds, such as RSNOs, are known to degrade due to their sensitivity to temperature and thermal degradation [66]. To investigate the stability during the sterilization process, the DCDI samples were exposed to clinically relevant UV-light and EO sterilization methods commonly used with thermally sensitive medical devices [67,68]. The SR-NO DCDI retained 99.06 ± 2.26% and 99.33 ± 1.08% SNAP after UV and EO sterilization, respectively, as compared to the initial wt% of SNAP in freshly prepared DCDI samples (Fig. 4A). In order to evaluate and confirm the activity of DCDI insert in terms of NO-releasing characteristics, the NO release from the fresh and sterilized DCDI samples was measured in the absence (0% light) and presence of light (100%). The results from the study revealed no significant difference between NO-release kinetics from freshly prepared vs. sterilized DCDI samples. The data demonstrates that the freshly prepared samples had 1.740 ± 0.092 × 10⁻¹⁰ mol min⁻¹ cm⁻² of NO flux, while the EO and UV sterilized samples released 1.707 ± 0.083 × 10⁻¹⁰ mol min⁻¹ cm⁻² and 1.689 ± 0.078 × 10⁻¹⁰ mol min⁻¹ cm⁻², respectively with 100% white light stimulation (Fig. 4B). These results indicate that both sterilization methods are compatible with the SR-NO polymeric DCDI device without significant losses in the amount of NO payload in the device, critical for clinical translation. Similar results have been reported regarding the compatibility of SNAP-based polymers with various sterilization methods [46,53]. These prior studies also confirmed the ability of SNAP-based polymers in preserving >90% of the initial wt% of SNAP incorporated into Elast-eon E2As (a copolymer of combined soft portions of polydimethylsiloxane and poly(hexamethylene oxide)), silicone rubber, E5–325, and CarboSil polymers after EO sterilization and UV-light sterilization [46,53].

Fig. 4.

The impact of storage and sterilization on NO-releasing DCDI device. (A) The retention of SNAP in the polymer after the sterilization process was analyzed by extracting the SNAP remaining in the polymer in THF solvent and measuring the absorbance of SNAP at 340 nm using UV–vis. Data represents mean ± SEM normalized to initial wt% of SNAP in freshly prepared samples (n ≥ 3). (B) Measurement of NO release from SR-NO DCDI samples fresh and after sterilization with ethylene oxide and UV-light. (C) Storage stability of the DCDI device after 30 d at room temperature (RT), −20 °C, and 4 °C. Data represents mean ± SEM normalized to initial wt% of SNAP in freshly prepared samples (n ≥ 3).

3.5. Shelf-life storage of DCDI

The success of biomedical devices is highly dependent on their ability to retain their function for a prolonged period. To evaluate the potential of the DCDI device to be stored at various storage conditions, SR-NO DCDI samples were stored for 30 d at clinically relevant storage conditions (i.e., −20 °C, 4 °C, and RT) for 30 d in a tightly closed vial, in the dark, and with desiccant to protect the insert from moisture. After 30 d, the SR-NO tubing was mounted on the side glow fiber optic, connected to the light source, and the NO release from the samples was recorded with and without light using chemiluminescence NOA at 37 °C for both fresh DCDI samples and DCDI samples that had been stored. Results from the storage stability experiment confirmed the stability of samples at different storage conditions for at least 30 d with no significant difference in the NO release levels (Fig. 4C). These results are in agreement with previously reported studies that confirm the stability of SNAP-based polymer for up to 8 months [46,53].

3.6. Antibacterial activity of DCDI using 4 h bacterial adhesion assay

3.6.1. Evaluating the antibacterial efficacy of DCDI

Various nosocomial infections in hospitalized patients are frequently linked with biomedical devices. While these devices are expected to offer life-saving protection, they are reported to be a primary source of device-associated infections in many instances [69]. A recent study reported an upsurge in the rate of hospital-acquired infections in COVID-19 patients due to longer lengths of hospital stay [70]. In times like these when conventional antimicrobial therapies are failing to contain the infection rates, alternative strategies are critically needed. For this reason, novel antimicrobial approaches have been proposed as an alternative to traditional methods. Light-based antimicrobial therapy is one such strategy heavily being employed for combating biofilms on medical implants [71]. Clinical pathogens such as S. aureus and E. coli have been proven to be vulnerable to photodynamic inactivation by the wavelengths of the visible spectrum of light (400–800 nm) [72–74]. Similarly, photoactivation of silver and gold nanoparticles has also been explored for bactericidal efficacy; however, the metal-dependent killing is distinct for Gram-positive than for Gram-negative bacteria and often exhibits cytotoxicity towards mammalian cells [75,76]. For this reason, a broad-spectrum, biocompatible disinfecting device was developed in this study for eradicating both Gram-positive and Gram-negative bacteria.

The bactericidal efficiency of the DCDI device was evaluated against S. aureus (ATCC 6538) and E. coli (ATCC 25922) bacteria using a 4 h bacterial adhesion assay. The bacterial cells that adhered to the DCDI were enumerated and normalized to the surface area of the DCDI to obtain viable CFU cm⁻². Results from S. aureus adhesion on the synergy of SR-NO-Light unveiled a 99.45% reduction compared to the SR control (p < 0.05). The SR-Light and SR-NO samples also had a significant reduction in viable adhered cells as compared to SR control (p < 0.05) due to the action of the NO release (93.05% reduction) and light-mediated interface (41.30% reduction), respectively (Fig. 5A). A similar trend was observed after exposure to E. coli, where the SR-Light and SR-NO controls resulted in an 85.01% and 92.89% reduction in viable cells, respectively. The highest reduction in viable E. coli adhesion on the surface occurred with the SR-NO-Light DCDI resulting in a 99.36% reduction compared to SR control (p < 0.05) (Fig. 5B). These results align with the NO release levels obtained from SR-NO and SR-NO-Light samples, where both samples exhibit physiological levels of NO that are increased via the photoinitiated catalysis with the white light (Fig. 3F). Overall, the synergistic effect of NO and light resulted in broad-spectrum antimicrobial activity on the DCDI surface and significant inhibition of both S. aureus and E. coli bacteria. Mutations in the DNA sequence via reactive nitrogen species (RNS) is one of the major mechanism by which NO exhibits antimicrobial activity [31]. NO’s reaction with oxygen and peroxides leads to the formation of a range of free radical antimicrobial species such as peroxynitrite and nitrogen dioxide that can modify and destroy the DNA base pairs using oxidative and nitrosative mechanisms. The damage of DNA strands facilitates lipid peroxidation, constrains enzyme functions, and results in eventual membrane loss in microorganisms [31,77–79].

Fig. 5.

Antibacterial activity of the DCDI device calculated as a log of the colony forming units (CFU) cm⁻² per surface area against: (A) S. aureus; ** represents p ≤ 0.01, calculated for SR-NO, SR-NO-Light vs. SR, ## represents p ≤ 0.01, calculated for SR-NO vs. SR-Light, ### represents p ≤ 0.001, calculated for SR-NO-Light vs. Light, $ represents p ≤ 0.05 calculated for SR-NO-Light vs. SR-NO; and (B) E. coli; * represents p ≤ 0.05, calculated for SR-NO, SR-NO-Light vs. SR, # represents p ≤ 0.05, calculated for SR-NO-Light vs. light, $ $ represents p ≤ 0.01 calculated for SR-NO-Light vs. SR-NO. (C) Design of in situ catheter disinfection experimental model. A model catheter is exposed to S. aureus bacteria for 24 h allowing the bacteria to adhere and proliferate, creating the pre-infected catheter surface. The DCDI is inserted within the catheter lumen and the adhered bacteria are dispersed via the photoinitiated NO release. (D) Representative images of LB agar plates with viable S. aureus bacteria CFU after 4 h of exposure to SR control and SR-NO-Light DCDI in the in situ catheter disinfection model. (E) In situ disinfection of a contaminated model catheter with DCDI calculated as a log of CFU cm⁻¹ of catheter length against S. aureus. (F) Cytocompatibility of DCDI evaluated against 3T3 mouse fibroblast cell line relative to cell control in a 24 h cell viability assay using CCK-8 cell viability kit. All data are represented as mean ± SEM (n ≥ 3).

3.6.2. In situ surface disinfection using catheter infection model

The ability of DCDI to disinfect the catheter surface was characterized against S. aureus, one of the most common bacteria associated with biofilm-related infections. The S. aureus bacteria were grown in LB media until the mid-log phase and exposed to model CVC catheters for 24 h at 37 °C in order to develop a pre-established infection/biofilm on the CVC catheter surface. After 24 h, the DCDI and the corresponding SR control were then inserted into the infected catheter to investigate the disinfecting ability (Fig. 5C). After 4 h exposure to the DCDI device, the viable bacteria remaining on the surface of the catheter were enumerated using the plate counting method (Fig. 5D). The study demonstrates the ability of the SR-NO-Light DCDI to eradicate a pre-established S. aureus infection on the CVC catheter surfaces by ca. 96.99% compared to the unmodified SR control (p < 0.05) (Fig. 5E).

Nitric oxide is known to induce biofilm dispersal across many bacterial strains, which led to its importance in emerging as a therapeutic for biofilm-related infections. NO is a reactive gas with a very short half-life with the ability to diffuse through the cell membranes spontaneously. Previous reports have shown that NO at lower concentrations can trigger the switch of sessile cells to free-floating planktonic phenotype in bacterial cells enclosed within the biofilm [80]. It is understood that the control of intracellular secondary messenger such as cyclic di-GMP by NO imitates the effectors which can hamper the biofilm buildup and disperse the mature biofilm. [81] The reactive nitrogen species from NO and the superoxide ions lower the extracellular polysaccharide production which is an important intermediary component for bacterial attachment on a substratum. The role of NO in facilitating biofilm dispersion is maintained across a wide range of bacterial species. [82]

Silicone rubber is one of the most common polymer materials utilized for a wide range of biomedical devices. Due to its inherent biocompatibility, mechanical properties, and higher in vivo stability, previous studies have shown the compatibility of SR with NO-releasing chemistry [47,54,83]. These polymers incorporated with NO donating compounds have demonstrated excellent biocompatibility, stability, and antibacterial activity with prolonged physiological levels of NO release [84]. Meyerhoff and co-workers examined the transport of NO in various medical-grade polymers and concluded that NO has the ability to diffuse through a wide range of biomedical polymers, in particular silicones and polyurethanes that are commonly used in catheter devices [85,86]. Given the nature of gaseous NO and its high diffusion rate through such biomedical polymers, the activity of NO is retained both on the intraluminal and extraluminal catheter surfaces as demonstrated in this work and in prior literature reports [47,87]. Microbial contamination can occur both on the extraluminal and intraluminal catheter surfaces. Previous studies have shown the ability of NO-releasing lock solution present on the inside of the catheter lumen in eradicating biofilm buildup both on the inside and as well as outside of the catheter [60,88]. It is understood that >70% of CRBSIs are extraluminally developed from microorganisms originating on the patient’s own skin flora or the caretaker/healthcare worker’s hands, resulting in a contaminated catheter hub region and infected extraluminal catheter surfaces [89–91]. NO-releasing chemistry and material approaches, such as this DCDI device and catheter-lock solutions, have the advantage due to the diffusion of NO gas through the polymer matrix and are superior to conventional ethanol/antibiotic lock solutions which fail to act on the bacteria present on the extraluminal catheter surfaces. Results obtained in this study revealed a similar antibacterial effect from the NO-releasing DCDI insert where ~97% of viable bacteria were eliminated from the extraluminal catheter surface that was treated with the DCDI as compared to control without any antibacterial insert.

Activation of NO release from SNAP and other photo-responsive NO-releasing composite materials have been previously reported using visible and near-infrared LED lights [92,93,54]. The advantage of RSNO’s sensitivity to light can be exploited to control the amounts of NO released by modifying irradiation time and intensity of light from NO-releasing biocompatible polymer. The influence of light instantly releases NO from the polymer matrix proving the sensitivity of RSNOs to the photolytic feedback. Although, previous studies demonstrated the advantages of having strong control over the NO release levels [94,95], this work of developing the DCDI device validates a practical and functional method to take advantage of the tunable photo-release properties of RSNOs for clinical catheter infection applications. The significant results from both the prevention of bacterial adhesion and in situ disinfection of pre-infected catheter surfaces support the significance of the DCDI device in reducing bacterial contamination on medical devices in clinical settings. The photoinitiated NO release from the DCDI is a potent and locally acting antimicrobial device that is biocompatible, low cost, shelf-stable, and easy to apply. These characteristics make it an ideal choice for integrating with medical devices such as catheters, where the disposable nature of the DCDI device will enable long-term applications of NO to the catheter surfaces both to prevent viable microbial adhesion on catheter surfaces and eradicate biofilms on catheter surfaces to reduce instances of CRBSIs. Therefore, the development of the DCDI device is expected to be a significant step toward preventing microbial contamination in both short- and long-term indwelling catheters that are regularly at a higher risk of contracting an infection (intravenous catheters, urinary catheters, insulin cannulas, peritoneal dialysis catheters, hemodialysis catheter, etc.), resulting in reductions of morbidity, mortality, and associated healthcare costs [96–98].

3.7. Cytocompatibility of DCDI

Over the years, eukaryotic cells have established mechanisms for scavenging the reactive oxygen and nitrogen species mediated by NO which enables them to negate their influence; however, various microorganisms (bacteria and virus) remain vulnerable [99]. The combination of SNAP and light emitted from the DCDI provides broad-spectrum antimicrobial activity as described above. Nonetheless, it was crucial to determine the compatibility of engineered disinfection devices towards mammalian cells for effective in vivo application. To investigate this, the leachates collected over 24 h in DMEM media from the SR, SR-NO, SR-Light, and SR-NO-Light DCDI samples were added and incubated with NIH 3 T3 mouse fibroblast cells for 24 h at 37 °C. The cell cytotoxicity assay was quantified using the CCK-8 cell viability kit. All the samples exhibited >90% viability in the cells over 24 h (Fig. 5F). Experimental designs based on the ISO standards have been previously used to demonstrate the biocompatibility of medical devices. [100] This data is consistent with prior studies that confirmed the biocompatibility of SNAP-based NO-releasing polymer both in vitro and in vivo [84,101]. Together the results from these cytocompatibility studies offer encouraging evidence towards the potential biocompatibility of the light-induced DCDI device.

4. Conclusions

To address the challenges associated with catheter-related bloodstream infections (CRBSIs), a simple, effective, smartphone-compatible, and universal Disposable Catheter Disinfection Insert (DCDI) was fabricated in this study that can both prevent and disinfect indwelling catheters (intravascular, urinary, etc.). The novel DCDI is comprised of a light-sensitive NO donor molecule, SNAP, impregnated in medical-grade silicone rubber tubing that is mounted on a side glow fiber optic. In clinical practice, the DCDI would be attached to the catheter lumen cap and inserted within the lumen of the indwelling catheter. Once the DCDI is inserted, a light source would illuminate the side glow fiber optic by using a simple mobile phone application, providing tunable photo-activated NO release levels in real-time via modulation of light intensity. The modulation of light-activated NO release from the DCDI device can then be used to prevent catheter infections or inserted within an infected catheter to eradicate the colonized bacteria. Among the wavelengths and intensity of light emitted from the side glow fiber optics, the maximum photocatalytic activity of the DCDI releasing physiological NO levels from the SR-NO polymer occurred with the white light (mixture of red (620 nm), green (530 nm), and blue (450 nm) light) at 100% intensity as measured using a chemiluminescence NO analyzer. The antibacterial activity and biocompatibility of the DCDI with real-time control of NO release at physiological levels were also evaluated in vitro. The SR-NO-Light DCDI demonstrated broad-spectrum antibacterial activity eradicating >99% viable S. aureus (Gram-positive) and E. coli (Gram-negative) on the DCDI surface with just 4 h of exposure. Additionally, the insertion of DCDI in an infected model catheter resulted in 97% eradication of S. aureus bacteria in situ exhibiting the potential to eradicate an established catheter infection. The DCDI device presented in this study offers universal compatibility with a wide range of catheters that are currently on the market and used for a range of biomedical applications (e.g., vascular, hemodialysis, urinary catheters). The inexpensive nature of the disposable components of the device, specifically the NO-releasing polymer tubing and the fiber optics (on the order of a few dollars), is expected to be a significant benefit in terms of preventing costly catheter-related infections which are a significant financial burden on the healthcare system. In some cases, the DCDI device technology can also be applied for other applications that use polymer tubings and require a bacteria-free state for efficient function. The ease of synthesis and fabrication, the capacity of the DCDI to be tuned for its NO release levels, its antibacterial efficacy, biocompatibility, smartphone compatibility, ability to withstand conventional hospital sterilization processes, and long-term shelf-life stability at various storage conditions make the DCDI device a promising new solution for the prevention and eradication of catheter-related infections without the need of complicated surgical interventions.