Using the methods previously published by (Liu et al., 2019a), catheters were coated with antimicrobial agents. In brief, mouse and human catheters were coated with auranofin (3 mg/mL and 10 mg/mL), fluconazole (10 mg/mL), and vancomycin (15 mg/mL). Negative control catheters were coated with THF + PU alone, absent any antimicrobial agents. Human (14G x 2”, Exel Safelite Catheters, 26726, Radiopaque teflon material) and mouse (14G x 2”, Surflo I.V. Catheters, SR-OX1451CA, Polyurethane material) catheters were cut into 1 cm segments using a sharp scalpel and submerged in a 20 mL scintillation vial containing 3 mL of THF + PU + auranofin/fluconazole/vancomycin. The setup was placed on a shaker overnight at room temperature. Post incubation, catheters were air-dried in a fume hood overnight and then stored at 4°C until further use.

The fluid flow rate of auranofin-coated and uncoated human and mouse catheters was evaluated using a syringe pump (Harvard Apparatus, Holliston, MA). Assessment entailed providing a constant flow of liquid (phosphate buffered saline (PBS)) and measuring the capture volume.

For both human and mouse catheters, assessment groups included: untreated catheters, THF-coated catheters, PU dissolved in THF catheters, catheters coated with auranofin (3 and 10 mg) suspended in THF + PU coating, vancomycin (15 mg) suspended in THF + PU coating, and fluconazole (10 mg) suspended in THF + PU coating. The flow of PBS through human and mouse catheters was determined by keeping the volume of liquid constant. A 20 mL syringe was filled with 20 mL (the syringe was filled 3 times) of PBS and connected to a catheter adapter that married the catheters to the syringe tip, connecting the devices to the syringe pump ( Supplemental figure 1 ), which was then set to run at a flow rate of 1 mL/min for 10 min at a 50% force level to assess the final volume captured during a fixed period.

By contrast, to quantify the time needed to collect a fixed volume of liquid, a syringe was filled with PBS, locked into the syringe pump, and run for 10 minutes. The flow rate was set at 1 mL/min with 50% force. The volume of PBS collected was measured via an analytical scale and confirmed by measuring volume in a graduated tube.

Female BALB/c mice (n=5/group) were used according to the protocol approved by the Institutional Animal Care and Use Committee. The 1 cm drug-coated catheters were put inside culture tubes containing luminescent S. aureus USA300 (Dai et al., 2013) logarithmic culture and incubated for 2 h with agitation. To implant, the catheters, BALB/c female mice (Charles River, Wilmington, MA.) were anesthetized via intraperitoneal (IP) injection with ketamine (100 mg/kg) and xylazine (10 mg/kg). Mouse flanks were shaved, and the skin was sterilized topically using betadine and isopropyl alcohol. A 5 mm skin incision was made using sterile scissors to create a subcutaneous tunnel for the 1 cm catheter previously seeded with bacteria and the incision site was then closed with surgical staples. Two procedures were performed per animal with placement of two catheters per animal, one on each flank.

Bacterial colonization was analyzed using the live animal imaging system (IVIS Lumina, Series III; Perkin Elmer, Waltham, MA, USA) at various time points following inoculation: 24, 48, and 72 h post insertion. Total photon emission from defined regions of interest within the images of each mouse was quantified using Living Image software (Perkin Elmer, Waltham, MA). After the final imaging time, mice were sacrificed, the catheters were surgically removed, sonicated in 1 mL tryptic soy broth (TSB), and the bacterial load was enumerated after serial dilution and plating on TSB agar plates. Bacteria burden from the experimental catheters was compared using One-way ANOVA using GraphPad Prism Version 6.04 (GraphPad Software, La Jolla, CA).

Both mouse and human catheters were assessed for fungal biofilm formation and accumulation of microbial cells after placing uncoated and coated 1 cm segments in 12-well plates and adding 2 mL of bovine serum (BS) (ThermoFisher Scientific, Waltham, MA) that were then incubated overnight at 37 °C with agitation (150 rpm) in an orbital shaker. Post incubation, the BS was removed through aspiration, and the catheters were washed with 2 mL of PBS. Catheters were then transferred to a fresh plate with 2 mL of spider medium (10 g of Difco nutrient broth; 10 g of Mannitol; 4g of K₂HPO₄; pH to 7.2; autoclave in 1 L of water) and inoculated with 40 µL (OD_{600 =} 0.5) of an overnight culture of constitutively expressed green fluorescent protein (GFP) C. albicans MLR62 (Nobile and Mitchell, 2005). Seeded cultures were incubated for 90 min at 150 rpm on an orbital shaker, allowing fungal cells to adhere to catheter surfaces, then washed with PBS before being transferred to a fresh 12-well plate containing 2 mL of Spider broth. Catheters were allowed to incubate with agitation for 60 hours, allowing the formation of mature biofilms before being gently washed with 2 mL of PBS. While suspended in PBS, the catheters were incubated for 10 min in a sonicating water bath (Model: Branson 2800, SonicsOnline, Richmond, VA, USA) (with an interval of 5 sec after every 30 sec of sonication). A 100 µL aliquot for each sample was transferred to a fresh 96-well plate (Corning, ThermoFisher Scientific, Waltham, MA). The released fungal cells were quantified at 488 nm excitation and 535 nm emission using SpectraMax i3X (Molecular Devices, San Jose, CA). The remaining sonicated slurry was serially diluted in PBS and plated on a YPD agar, then incubated for 18 h before enumerating the colony forming units (CFU).

Both mouse and human catheters were assessed for dual biofilms (S. aureus - C. albicans biofilm) by placing segments in 12-well plates and adding 2 mL of bovine serum (BS) (ThermoFisher Scientific, Waltham, MA). Catheters were incubated overnight at 37°C with agitation (150 rpm) in an orbital shaker. Post incubation, the BS was removed through aspiration, and the catheters were washed with 2 mL of PBS. The catheters were then transferred to a fresh plate 2 mL plate with 40 µL of C. albicans MLR62 (1 × 10⁶ cells/mL) (Nobile and Mitchell, 2005) and S. aureus USA 300 (1 × 10⁷ cells/mL) in 1:1 v/v YPD/BHI (2 mL) media. Seeded cultures were incubated for 90 min at 37 °C at 150 rpm on an orbital shaker, allowing fungal/bacteria cells to adhere to catheter surfaces, then washed with PBS before being transferred to a fresh 12-well plate containing 2 mL of Spider broth. Catheters were allowed to incubate with agitation for 60 hours, allowing the formation of mature biofilms, before being gently washed with 2 mL of PBS. While suspended in 2 mL of PBS, the catheters were sonicated for 10 min (with an interval of 5 sec after every 30 sec of sonication) in a water bath sonicator (Model: Branson 2800, SonicsOnline, Richmond, VA, USA). A 100 µL aliquot for each sample was serially diluted. Fungal cell growth was selected on a YPD plate containing vancomycin (15 mg/mL), and bacterial cells were set on TSB media containing fluconazole (25 mg/mL). The plate was incubated at 37°C for 18 h, and the CFU was enumerated.

Female BALB/c mice (n=5/group) were used according to the protocol approved by the Institutional Animal Care and Use Committee. Mouse catheters (uncoated, THF+PU, auranofin 3 mg + PU 50 mg, auranofin 10 mg + PU 50 mg, and fluconazole 10 mg + PU 50 mg) were treated overnight in 2 mL of FBS and inoculated with 1x10⁴ CFU/mL of C. albicans MLR62 and incubated at 37°C for 90 min (Nobile and Mitchell, 2005). On day 1, the BALB/c female mice (Charles River, Wilmington, MA.) were anesthetized via IP injection with ketamine (100 mg/kg) and xylazine (10 mg/kg). The mouse flanks were shaved, and the skin was sterilized topically using betadine and isopropyl alcohol. A 5 mm skin incision was made using sterile scissors to create subcutaneous tunnels on both animal flanks for 1 cm catheters that were previously seeded with C. albicans MLR62. Upon completion, the incisions were closed with steel wound clips. Control animals were implanted with two drug-free catheters. Post-procedure, the animals were revived with atipamezole (1 mg/10 mg of xylazine used in a 100 PL volume) via IP injection. After catheter implantation and upon animal revival, the mice were given buprenorphine SR at 0.5 to 1 mg/kg via SQ injection.

After the 4-day evaluation period, animals were anesthetized with ketamine-xylazine and euthanized via exsanguination by cardiac puncture to collect blood (~1 mL). Subsequently, the catheters were removed from the euthanized mice and the blood was plated on YPD media to enumerate C. albicans MLR62 CFU. The retrieved catheters were sonicated in 1 mL YPD broth, and the fungal load was enumerated after serial dilution and plating on YPD agar plates ( Supplemental figure 2 ).

PBS was passed through coated catheters and analyzed using various antimicrobial coating agents to determine if the coating process impedes fluid flow. Flow dynamics were assessed for mouse and human catheters coated with auranofin (3 and 10 mg/mL), vancomycin (15 mg/mL), and fluconazole (10 mg/mL). Coated catheters were studied in three aspects, namely: time, volume, and mass.

Fluid flow was assessed by passing PBS fluid through catheters at a constant rate of 1 mL/min using a syringe pump ( Figure 1 ). The total collected volume was measured to see if the THF, PU, or drug coatings impeded fluid. The fluid that passed through the dip-coated catheters (mouse and human) remained consistent compared to the uncoated control catheters, achieving a 60 mL collection volume after 1 h as expected with the 1 mL/min flow rate ( Figures 1A, B ).

Analytical measurement of the total weight of PBS passing through the catheters at a fixed time of 10 min and at a flow rate set at 1 mL/min found that 9.8 and 9.9 g of PBS passed through the mouse and human catheters, respectively ( Figures 1C, D ) within the designated time frame. Again, indicating that the coating process does not impede fluid compared to the non-coated catheters (no statistical difference compared to the control) and that the total volume flowed through the catheters unimpeded. The average blood flow rate under normal conditions can vary for different parts of the body. The arterial flow rate ranges from 3 to 26 mL/min, and the venous flow rate can be 1.2-4.8 mL/min (Klarhöfer et al., 2001). The tested flow rate of 1 mL/min provides proof of concept. We have provided additional evidence that higher flow rates of 5 mL/min and 10 mL/min do not impact fluid flowing through the coated catheters (5 mL/min and 10 mL/min) ( Supplemental Figure 3 ). The catheters were incubated with 3 mL of PEG400 overnight, and the next day they were removed and dried in a sterile chamber for 24 h. Further, the flow rate of 60 mL PBS was analyzed using a Harvard apparatus. The PEG400 coating was also on human and mouse catheters and used as a comparison with PU coated catheters ( Supplemental Figure 4 ). There was no significant change in the flow rate in both catheters.

The time for a fixed volume of PBS to flow through the catheters was tested to determine if the added coating altered the fluid dynamics. A total volume of 10 mL of PBS was passed through the catheters, with a flow rate set at 1 mL/min, and at optimal flow, the total volume would be collected in 10 min. For both the coated and uncoated catheters, the expected 10 mL volume was recovered in the 10 min period ( Figures 1E, F ), indicating that the coating process did not alter the flow rate. Importantly, with the cumulative assessment of volume and time, it appears that neither the carrier PU in THF nor the added antimicrobial drugs altered the flow dynamics compared to uncoated catheters. Although the tested antimicrobial compounds differ in chemistry, size, and hydrophobicity, none adjusted fluid flow based on the tested parameters.

Previous in vitro investigation of the auranofin and vancomycin-coated catheters demonstrated a significant reduction in S. aureus biofilm accumulation on catheters, suggesting this specific antimicrobial drug coating impedes bacterial biofilm formation and accumulation on the devices (Liu et al., 2019a). To advance this early finding, coated catheters were implanted subcutaneously, and the bacterial load was measured at the implant site in live animals, visualized via luminescence imaging. Captured images showed bacterial accumulation when non-coated catheters were implanted ( Figures 2A-E ). Bacteria were also visualized accumulating at the implant site when catheters were coated with PU dissolved in THF. However, bacterial accumulation was reduced when auranofin or vancomycin was included as part of the catheter coatings.

Post implantation, catheters were recovered from the subcutaneous pockets and S. aureus associated with the catheters was enumerated by sonicating the catheter and plating on TSA plates. The uncoated and PU-alone coated catheters accumulated bacteria, reaching 4 x 10⁹ and 1.7 x 10⁹ CFU/mL, respectively. However, a 3 mg/mL auranofin-coated catheter was reduced to 7.4 x 10⁵ CFU/mL, and a 10 mg/mL auranofin-coated catheter reduced bacteria accumulation further to 1.2 x 10⁵ CFU/mL, a 4-log reduction from the controls ( Figure 2F ). The positive control vancomycin demonstrated the most significant reduction of bacteria, with no bacteria recovered from the catheters. While auranofin efficacy was surpassed by vancomycin (a standard-of-care antibiotic) at the tested concentrations, it provides evidence that auranofin can exert potent bacterial inhibition. Although catheters are coated with mg/mL amounts of compounds, previously published data indicate that µg amounts are released. For example, catheters coated with 10 mg/mL auranofin were found to release approximately 37 µg (Liu et al., 2019a).

In prior work, we determined that auranofin coated catheters could reduce the accumulation of S. aureus biofilm in vitro (Liu et al., 2019a). The remarkable nature of auranofin is imparted by further testing the impact on fungal accumulation. Indeed, the antimicrobial nature of auranofin has been shown to inhibit both bacterial and fungal pathogens (Fuchs et al., 2016), making it uniquely suited as a catheter coating that can be exposed to numerous types of microbes. A constitutively expressing GFP C. albicans strain (MLR62) was used to monitor biofilm accumulation on the catheters. The absorbance measurement for C. albicans GFP expression showed a decrease in biofilm formation on auranofin (3 mg and 10 mg) coated catheters compared to the uncoated catheters ( Figures 3A, B ). No GFP was recorded with the 3 mg auranofin-coated mouse catheter, suggesting no detectable accumulation of C. albicans cells.

Enumeration of C. albicans cells on auranofin-coated and uncoated catheters (mouse and human) found a reduction attributed to auranofin coating. When coated with 3 mg and 10 mg auranofin, mouse catheters accumulated 1.6 x 10⁵ and 7.8 x 10⁵ CFU compared to 2.0 x 10⁸ CFU associated with catheters exposed to THF solvent alone ( Figure 3C ). Thus, a 3-log reduction in C. albicans cells was achieved with the auranofin coating with P=0.0229 and P=0.023, respectively. When the dip coating method was applied to human catheters, associated C. albicans cells were also reduced from accumulating on the material. Catheters exposed to THF solvent were found to have 1.6 x 10⁷ CFU compared to 2.5 x 10⁶ (P=0.0052) and 2.8 x 10⁶ (P=0.0049) CFU recovered from catheters coated with 3 and 10 mg/mL auranofin, respectively ( Figure 3D ). Fluconazole-coated catheters (1x10⁴ CFU/mL) also demonstrated a reduction in C. albicans accumulation. Measuring C. albicans biofilm biomass can be challenging because this requires weighing the catheters prior to introducing the C. albicans post-biofilm formation. In this case, there can be variations in the coating volume added to the individual catheters due to slight differences in release volumes during the incubation period that can impact weight. Thus, mass cannot be accurately measured. Therefore, we used C. albicans strain MLR62 which constitutively expresses GFP as a means of measuring live C. albicans cells associated with the catheters. To provide insights into the impact of auranofin on C. albicans biofilm accumulations, C. albicans biofilm biomass changes on silicone pads were demonstrated in the presence of auranofin ( Supplemental Figure 5 ). Treating wells with auranofin demonstrate that C. albicans biofilm accumulation is reduced compared to untreated control wells based on dry weight. Therefore, the result demonstrated that mouse and human catheters coated with auranofin exhibited a significant reduction in C. albicans biofilm formation and accumulation over the 60-hour test period, enough time for mature biofilms to form.

A unique feature of auranofin is its ability to inhibit both bacterial and fungal pathogens (Harbut et al., 2015; Fuchs et al., 2016). To evaluate the ability of auranofin to impact a mixed microbe biofilm, catheters were exposed to a mixed pathogen culture of S. aureus and C. albicans biofilm allowed to mature before being disrupted by sonication and plated on TSA and YPA selection media to enumerate the individual microbes. Applying 10 mg auranofin on human catheters reduced S. aureus accumulation from 5.1 x10⁵ CFU/mL found on non-coated catheters to 6 x10³ CFU/mL but failed to reach significance. However, C. albicans biofilm on the human catheter was reduced from 1.2 x 10⁸ CFU/mL on non-coated catheters to 1.3 x 10⁵ CFU/mL on the 10 mg auranofin coated catheters, an efficient reduction (P=0.0163) as demonstrated with a Student’s t-test ( Figure 4A ). Although both vancomycin and fluconazole impacted microbe accumulation on human catheters, they failed to have cross-species efficacy as seen with high concentrations of auranofin. Fluconazole reduced S. aureus accumulation from 5.1 x10⁵ CFU/mL found on non-coated catheters to 2.5 x10⁴ CFU/mL (P=0.1262) and did not reduce C. albicans accumulation (2.8 x 10⁸ CFU/mL; P=0.4312). Vancomycin coating at 15 mg caused an increase in S. aureus associated with the catheters, reaching 3.4 x 10⁶ CFU/mL. C. albicans accumulation was also not reduced by vancomycin, reaching 2.43 x 10⁷ CFU/mL with the 10 mg coating concentration.

Murine catheters were also evaluated in the same manner. Catheters coated in 10 mg auranofin exhibited S. aureus reduction from 2.3 x 10⁷ CFU/mL to 5 x 10³ CFU/mL (P=0.0005) ( Figure 4B ). C. albicans biofilm was reduced from 2.3 x 10⁷ CFU/mL on non-coated catheters to 6 x 10⁴ CFU/mL when coated with 10 mg of auranofin (P=0.0005) ( Figure 4B ). Thus, both catheters coated with auranofin 10 mg + PU significantly reduced microbe accumulation on the catheters under in vitro conditions that usually promote microbial attachment and mature biofilm formation.

Finding that auranofin-coated catheters demonstrated in vitro reduction of C. albicans biofilm accumulation prompted further evaluation using a subcutaneous murine model. C. albicans cells were allowed to attach to coated catheters before implantation. At 4 days post-implantation, catheters were removed, sonicated to detach C. albicans, and the CFU was enumerated on YPD plates. Both fluconazole (10 mg/mL) and auranofin-coated catheters (10 mg/mL) exhibited a reduction in the C. albicans biofilm, recovering 1.6 x 10⁴ CFU/mL (P=0.0003) and 6.9 x 10⁴ CFU/mL (P=0.0019), respectively, compared to the non-coated catheters (3.9 x10⁵ CFU/mL) (Figure 5). Biofilm reductions depict the efficacy of the coating method and material. The study mainly focuses on testing the efficacy of polyurethane coating with auranofin on catheters. Also, increasing the concentration of auranofin will significantly decrease the biofilm at higher folds in comparison to uncoated catheters. We noted that there was no significant inflammation on the site of catheter insertion in the mouse model. We also collected blood by heart puncture, but no microbe growth was recorded.