Suppression of Biofouling on a Permeable Membrane for Dissolved

Feb 22, 2019 - Matthew Osborne , Aditya Aryasomayajula , Amid Shakeri ... DO sensor designs can employ selectively permeable membranes to isolate DO ...
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Suppression of Biofouling on a Permeable Membrane for Dissolved Oxygen Sensing using a Lubricant-Infused Coating Matthew Osborne, Aditya Aryasomayajula, Amid Shakeri, Ponnambalam Ravi Selvaganapathy, and Tohid F. Didar ACS Sens., Just Accepted Manuscript • DOI: 10.1021/acssensors.8b01541 • Publication Date (Web): 22 Feb 2019 Downloaded from http://pubs.acs.org on February 25, 2019

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Suppression of Biofouling on a Permeable Membrane for Dissolved Oxygen Sensing using a Lubricant-Infused Coating Matthew Osborne a, Aditya Aryasomayajula b, Amid Shakeri b, Ponnambalam Ravi Selvaganapathy a,b, Tohid F. Didar a,b,c* a

School of Biomedical Engineering, McMaster University, 1280 Main Street West, ETB 406, Hamilton, Ontario, Canada, L8S 4K1

b

Department of Mechanical Engineering, McMaster University, 1280 Main Street West, JHE 310, Hamilton, Ontario, Canada, L8S 4L7

c

Institute for Infectious Disease Research, McMaster University, 1280 Main Street West, MDCL 2235, Hamilton, Ontario, Canada, L8S 4K1 *

Corresponding author: send all correspondence to [email protected]

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ABSTRACT Specific ranges of dissolved oxygen (DO) concentrations must be maintained in a waterbody for it to be hospitable for aquatic animals. DO sensor designs can employ selectively permeable membranes to isolate DO from untargeted compounds or organisms in waterbodies. Hence, the DO concentration can be monitored, and the health of the water evaluated over time. However, the presence of bacteria in natural waterbodies can lead to the formation of biofilms that can block pores and prevent analyte from permeating the membrane, resulting in inaccurate readings. In this work, we demonstrate the implementation of a fluorosilane-based omniphobic lubricant-infused (OLI) coating on a selectively permeable membrane and investigate the rate of biofilm formation for a commercially-available DO sensor. Coated and unmodified membranes were incubated in an environment undergoing accelerated bacterial growth, and the change in sensitivity was evaluated after 40, 100, 250, and 500 hours. Our findings show that the OLI membranes attenuate biofouling by 70% and maintain sensitivity after three weeks of incubation, further demonstrating that oxygen transfer through the OLI coating is achievable. Meanwhile, unmodified membranes exhibit significant biofouling that results in a 3.35 higher rate of decay in oxygen measurement sensitivity and an over-70% decrease in static contact angle. These results show that the OLI coating can be applied on commercially available membranes to prevent biofouling. Therefore, OLI coatings are a suitable candidate to suppress biofilm formation in the widespread use of selectively permeable membranes for environmental, medical and fluid separation applications. KEYWORDS: biofouling; bacteria; biofilm; membrane fouling; lubricant-infused coating; dissolved oxygen sensing.

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Natural waterbodies are a critical part of our biosphere and are predicted to be the home to a half-million different eukaryotic species1. These species rely on oxygen intake from dissolved oxygen (DO) gas, and fluctuations in DO concentration levels can be fatal. Diverse fish populations typically require between 7 mg/L and 10 mg/L to thrive. However, excessive pollution can result in hypoxic water conditions with concentrations below 4 mg/L2,3. Handheld DO meters present a reliable and convenient method to monitor the DO concentration in waterbodies and identify whether aquatic species are at risk of hypoxia. Most commercially available DO sensors employ a selectively permeable membrane4. DO is the sole compound capable of diffusing through the membrane, after which it undergoes an electrochemical reduction that leads to a quantifiable signal. By isolating DO from untargeted compounds, the selectively permeable membrane reduces signal noise, leading to higher accuracy and reading stability. In marine environments, planktonic bacteria are capable of detecting and adhering to solid surfaces via electrostatic interactions5, subsequently initiating biofilm formation. Multi-species groups of bacteria physically adhere in clusters on the surface and subsequently produce a viscous extracellular matrix (ECM) containing polysaccharides, proteins, phospholipids, and extracellular DNA (eDNA)6. Reproduction continues as the microcolony matures into a mushroom-shaped biofilm. Bacteria in the bottom layers downregulate their flagella and enter a state of sessile growth7. Finally, newly-produced planktonic bacteria are ejected from the biofilm to form new biofilms elsewhere in the environment. While selectively permeable membranes are an advantageous component in sensor design, they can provide a suitable surface for bacteria adherence. In the case of polluted waters, where DO concentrations are lower than needed for aquatic animals but advantageous for anaerobic bacterial growth, the membranes are at a high risk of undergoing biofouling. When this occurs, a 3 ACS Paragon Plus Environment

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biofilm is capable of blocking pores and preventing the target molecules from diffusing through the membrane, rendering the sensor inaccurate or perhaps inoperable. Furthermore, bacterial ECM are known to consist of acidic compounds that corrode submerged surfaces8. To discourage biofilm formation, surfaces can be coated with blocking agents that inhibit the physical attachment of bacteria. Marine antibiofouling is a longstanding area of interest in scientific and military research due to the corrosive and drag-increasing effects of biofilms on submerged vessels and infrastructure. A number of metallic (e.g. copper, silver) and biocidal nanoparticle coatings have been used9–11, but such cytotoxic chemicals can leach into waterbodies and result in adverse environmental effects12. The food and beverage industry has also employed antibiofouling coatings to limit spoilage and pathogen transfer between items13. These coatings are organic in nature and must be safely digestible in the case of direct contact with food. Similarly, the healthcare and dentistry industries use surface blocking strategies to prevent infections during implantation14,15. Common examples of organic blocking agents for biofouling include inert proteins, organosilanes8, and polyethylene glycol16,17; however, many of these agents have been shown to degrade or desorb over time, meaning that they are unsuitable for long-term use18. Alternatively, there is an emerging class of antibiofouling surface blocking agents called omniphobic lubricant-infused (OLI) coatings. They rely on the capillary forces induced by micro/nano-scale texturing or chemical affinities to lock a thin layer of lubricant over the treated surface19,20. When a foreign liquid is introduced to the surface, the resulting liquid-lubricant interface offers a high degree of repellency and “slipperiness” that prevents adhesion of the liquid as well as any entities suspended in it21–24. Ware et al. has investigated the use of a lubricantinfused nano-wrinkled polymeric surfaces in marine environments and has shown that its antibiofouling properties are effective on nonporous surfaces20. However, nano-wrinkling poses 4 ACS Paragon Plus Environment

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complications in fabrication and membrane permeability that are better addressed by a tethered liquid perfluorocarbon approach first demonstrated by Leslie et al. 25. We have previously used a highly fluorinated lubricant (perfluorodecalin) with a high affinity to a self-assembled monolayer (SAM) of an organosilane comprised of long perfluorocarbon chains, which was achieved using chemical vapor deposition26,27. In this study, we demonstrate the effectiveness of an antiadhesive fluorosilane-based OLI coating in preventing biofouling on the selectively permeable membrane of a commercially available DO sensor (ExStik II DO meter). By accelerating bacterial growth in tap water using yeast extract4, untreated (UT) and OLI membranes were incubated in a highly concentrated bacterial environment for up to three weeks. After set time intervals, biofilm formation was observed and characterized using sensitivity readings, fluorescence microscopy, scanning electron microscopy (SEM), and static contact angle (SCA) measurements. The OLI coating successfully prevented bacterial adhesion more effectively than UT membranes while still allowing DO to diffuse through unhindered. This leads to a more stable sensitivity over time and accurate measurements of DO concentrations. METHODS Materials. The ExStik II DO600 dissolved oxygen meter and DO603 membrane kits were purchased from Extech Instruments (Nashua, USA). Powdered yeast extract was purchased from BioShop (Burlington, Canada). Trichloro (1H,1H,2H,2H-perfluorooctyl) silane (TPFS), 4′,6Diamidino-2-phenylindole dihydrochloride (DAPI), and perfluorodecalin were purchased from Sigma-Aldrich (Oakville, Canada). Phosphate buffered saline (PBS) (0.1 M, pH 7.4) was prepared in-house. Formaldehyde was purchased from Caledon Laboratories (Georgetown, Canada).

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Oxygen plasma activation. Oxygen plasma was used to induce hydroxyl groups on membrane surfaces for covalent attachment of TPFS. Selected membranes were oxidized using medical grade oxygen gas and a Harrick PDC-001 (115V) plasma cleaner for 2 minutes at 450 mTorr. Liquid phase deposition (LPD). Immediately after plasma activation, the membranes were partially submerged in a container containing 5% TPFS in ethanol by volume so that only the exterior surface of the membrane was in contact with the solution. The container was then sealed and the membranes were incubated for 2 hours at room temperature so that the TPFS evenly distributes as a SAM28. Subsequently, they were baked on a hot plate in a fume hood at 60°C for a minimum of 6 hours to promote the formation of covalent bonds and evaporate hydrochloric acid, a by-product of the reaction. Samples were then sonicated in ethanol for 20 minutes to remove any unbound TPFS. Accelerated bacterial growth environment with yeast extract. An environment that stimulates accelerated multi-bacterial growth was prepared in a glass dish using 1 g yeast extract and 100 mL tap water. OLI membranes were first saturated in perfluorodecalin lubricant for 30 seconds, then all membranes were partially submerged in the yeast solution. The container was mostly covered, but air could pass in to avoid mass bacterial death. Membranes (3 OLI and 3 UT) were removed and tested at 40, 100, 250, and 500 hours. Sensitivity readings. The biofilms were fixed with 4% formaldehyde in PBS for 30 minutes. The DO meter equipped with UT or OLI membranes were used to take readings in unstirred tap water at DO concentrations of 4 mg/L, 3 mg/L, 2 mg/L, and 1 mg/L. Tap water was recorded at 4 mg/L, and the subsequent concentrations were achieved by bubbling nitrogen gas in the tap water for 7 seconds, 15 seconds, and 30 seconds respectively. The exact concentrations were verified using a non-biofouled or “clean” UT membrane. Readings were recorded once the sensor exhibited no 6 ACS Paragon Plus Environment

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fluctuations (± 0.01 mg/L) for 60 seconds. The readings were plotted against those given by the clean membrane and fitted with linear trendlines, and the slopes were compared over time. Response times were measured for each membrane at 4 mg/L using a stopwatch. Fluorescence microscopy. Biofouled membranes were stained with 200 µL of 71.5 µM DAPI in PBS for 1 hour and rinsed lightly with distilled water for 5 seconds. Images were taken with a Zeiss AX10 inverted microscope with ApoTome.2 fluorescence functionality. Fluorescence intensity was quantified using ImageJ image processing software. The single highest and lowest values were removed for each condition. Static contact angle measurements. SCA measurements were taken using a Future Digital Scientific Corporation OCA 35 contact angle meter on the air-dried membranes with 1 µL sessile droplets of deionized water. Scanning electron microscopy. Biofouled membranes were cut from their caps and coated in a 36 nm layer of gold. SEM images were obtained using a Tescan VEGA II LSU at 10 kV and 5000x magnification. Bacteria were counted using ImageJ. 16s rRNA sequencing. Samples (1 mL) of biofouling solution after 40 hours of incubation were provided to and analyzed by MOBIX Lab (Sanger Sequencing and Oligo Synthesis Facility) at McMaster University (Hamilton, Canada). Water retention. Dry biofouled membranes were weighed. OLI membranes were soaked in perfluorodecalin for 30 seconds, then all membranes were submerged in fresh tap water for 2 hours. Membranes were removed, tapped vigorously to remove any residual water droplets and weighed. The mass of the thin lubricant layer (14.3 mg) was measured using OLI membranes rinsed to remove excess lubricant and then subtracted from the final masses. 7 ACS Paragon Plus Environment

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Statistical analysis. Data are presented as means ± standard deviation. Statistical significance was assessed using analysis of variance and post-hoc analysis with Tukey’s test, where significant difference was considered as P < 0.05.

Figure 1. Schematic of biofouling on untreated and treated membranes. (a) Progression of biofilm formation, including (i) detection of submerged surface, (ii) bacterial adsorption, (iii) ECM excretion, (iv) mature biofilm, and (v) ejection of planktonic bacteria. (b) In clean water, the selectively permeable membrane allows DO to diffuse through but prevents the diffusion of water and large compounds. (c) Practically, the bacteria living in natural waterbodies adhere to the membrane’s surface over time. They form biofilms that limit particles, including DO, from passing through the membrane. (d) The fluorosilane-based OLI coating prevents bacterial adhesion and

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ensuing biofilm formation but allows dissolved oxygen to diffuse through both the coating and the membrane. RESULTS AND DISCUSSION Figure 1 shows an illustration of biofilm formation on a permeable membrane. A total of 12 membranes were coated using LPD with a 5% TPFS solution as described in the methods. These OLI membranes and 12 UT membranes were used to take calibration readings at DO concentrations of 4 mg/L, 3 mg/L, 2 mg/L and 1mg/L. The readings were plotted in three sets of eight (4 UT and 4 OLI) against a corresponding set of UT readings (i.e. true concentrations), and subsequently fitted with linear trendlines (Figure 2). In the case where UT values were plotted against themselves, the linear trendlines had a unitary slope and overlaid one another. The OLI trendlines, on the other hand, demonstrated a slight average change in slope of 2.3% ± 0.9%. Hence, the OLI coating process has a minimal initial impact on the DO sensitivity.

Figure 2. Calibration of DO sensor with UT and OLI membranes before the biofouling experiment. UT membranes are plotted against themselves, resulting in a single trendline with a 9 ACS Paragon Plus Environment

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unitary slope (green, slope = 1). Each set of OLI membranes (shades of blue, slope = 1.0233 ± 0.0089) show a minor change in slope, confirming that the coating has a minimal effect on the sensor’s sensitivity. Incubation took place in an accelerated bacterial growth broth for varying time periods (40 hours, 100 hours, 250 hours, and 500 hours). A 16s rRNA sequencing analysis of the solution (Table SI 1) revealed that the bacterial composition primarily consists of Bacillus cereus (99.70%). This gram-positive rod-shaped bacteria is widely present in fresh and marine waterbodies, and can thrive in high (aerobic) and low (facultatively anaerobic) oxygen conditions29. The pH of the broth began at 6.52 and was measured at 8.50 after 3 weeks (Figure SI 1). This range is comparable to that of domestic wastewater (pH 6-9)30,31. At each interval, sets of 3 OLI and 3 UT samples were removed and readings were taken as described in the calibration. Figure 3 shows how the slope of each plotted trendline, which reflects the sensitivity, changes with each set of membranes as an absolute percentage. The values obtained using the biofouled membranes were plotted against those taken using an untreated and nonbiofouled membrane. Both UT and OLI curves demonstrate a comparable change in sensitivity between 0 and 100 hours. It is during the first 40 hours that bacterial growth is most active due to the overabundance of yeast extract nutrients in the water. This initial change in sensitivity for OLI membranes may also be attributed to the thin layer of perfluorodecalin lubricant, which has been shown to have very high oxygen solubility32. Afterwards, the UT curve continues to slope upwards at a consistent rate, peaking at a change of 26.0% ± 1.1% at 500 hours. The OLI curve shows no significant difference between 40 hours and 500 hours, generally lying between 4-10%. After 250 hours of biofouling, the DO sensor equipped with OLI membranes produced significantly more accurate readings than the UT membranes. Linear trendlines were added to the UT and OLI sets 10 ACS Paragon Plus Environment

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of data and the ratio of their slopes was calculated to be 3.35. Hence, we conclude that the rate of decay in sensitivity is 3.35 higher for UT membranes than OLI membranes. The response times with increasing biofouling time were recorded for UT and OLI membranes at 4 mg/L (Figure SI 2); however, no significant difference was observed between the coatings at any biofouling time. The average reading took 15 minutes to stabilize (no change ±0.01 mg/L for 1 minute), but variability within each condition was high due to the tendency for the sensor to overshoot and oscillate before stabilizing.

Figure 3. Change in sensitivity of the DO sensor over 500 hours. UT membranes demonstrate a 3.35 times higher rate of decay in sensitivity compared to OLI membranes over a 3-week period. There is no significant difference between UT and OLI membranes up to 100 hours, but UT membranes exhibited a 26.0% change in sensitivity, while OLI membranes were significantly lower (10.3%) by the end of the 21-day experiment. *, **Significant difference in change in sensitivity between UT and OLI readings (P < 0.05).

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All membranes were stained with DAPI, a blue fluorophore that binds to the A-T nucleotides on both the DNA within the fixed bacteria as well as eDNA in the ECM. Fluorescence microscopy was subsequently used to characterize the biofilm distribution (Figure 4a-b). Immature biofilms at the 40-hour mark featured dimmer blue regions of sparser individual bacteria and eDNA, and bright blue clusters of denser bacterial microcolonies. As the density of the biofilms increased with time, the images correspondingly increased in brightness and homogeneity. After 21 days, UT membranes featured significantly denser and uniform biofilms over the entirety of the surface. On the other hand, bacterial adhesion on OLI membranes were significantly dimmer and had numerous areas with little to no growth. Figure 4c-d are SEM images of membranes after 21 days of biofouling at 5000x magnification. The biofilm distribution on the UT membranes is significantly denser and more homogeneous in comparison to the OLI membranes, which agrees with the fluorescence analysis. Accordingly, the OLI membranes exhibit more unhindered surface area for gas permeation. Figure 4e quantifies the surface density of bacteria per square millimeter for UT (6.96  105 cells/mm2) and OLI (2.09 ·105 cells/mm2) membranes as shown in the SEM images. The ratio of these surface densities indicates that uncoated membranes undergo 3.32 times greater biofilm formation than OLI membranes. It can otherwise be stated that the OLI membranes achieved 70% attenuation.

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Figure 4. Characterization of biofilm density and distribution on biofouled membranes. (a, b) Both UT and OLI membranes show an increase in fluorescence intensity with time. The untreated 13 ACS Paragon Plus Environment

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membranes increase in fluorescence intensity at a higher rate and its biofilm distribution is more homogeneous. Scale bar = 200 µm. *, **Significant difference in fluorescent intensity between the UT and OLI samples (P < 0.05). (c) SEM images of UT membranes after 21 days depict a thick and homogeneous biofilm. Scale bar = 10 µm. (d) Meanwhile, isolated bacterial clusters and individual bacteria are sparsely distributed on the OLI membranes after 21 days. (e) Quantification of SEM images. At 500 hours, OLI membranes exhibit 3.32 times less bacteria per square millimeter than UT membranes. *Significant difference in bacterial surface density between UT and OLI membranes (P < 0.001). (f, g) UT membranes, initially hydrophobic (113.9° ± 2.1°), become more hydrophilic as biofouling progresses. After 21 days, biofilm formation leads to very hydrophilic conditions (32.1° ± 11.7°). OLI membranes (initially 119.4° ± 2.9°) maintain hydrophobicity over the 3-week period with only a 21.3° decrease in contact angle. *, **Significant difference in contact angles with biofouling time (P < 0.05). SCA measurements quantify the surface energy and repellency of the membrane surface (Figure 4f-g). Out of the package, the UT membranes are hydrophobic with a SCA of 113.9° ± 2.1°. This is critical to the design of this selectively permeable membranes since it relies on hydrophobicity to enable gas mass transfer while rebuffing water molecules33. The OLI treatment raises the SCA to 119.4° ± 2.9°. However, as they undergo biofouling, the hydrophilicity of the UT membranes drastically increases, resulting in a SCA of 32.1° ± 11.7° after 500 hours. This can be attributed to the hydrophilic nature of the polysaccharides, namely alginate34, that comprise the majority of the ECM composition. The OLI membranes demonstrated markedly less loss in hydrophobicity, finishing with a SCA of 96.3° ± 10.9° after 500 hours. Hence, OLI membranes were able to maintain hydrophobicity due to limited biofilm formation, whereas UT membranes shift from hydrophobic to hydrophilic. In addition to SCA, surface energy was characterized by 14 ACS Paragon Plus Environment

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weighing the water retention due to membrane swelling after 2 hours (Figure SI 3). As the UT membranes undergo more biofouling and increase in hydrophilicity, so does their water retention. On the other hand, the retention of OLI membranes shows no significant change in water retention with increasing biofouling time. As confirmed by fluorescence imaging, SEM and SCA measurements, OLI coated permeable membranes demonstrated a 70% suppression of biofilm formation and effectively maintained repellency. This finding is aligned with results obtained in similar biofouling studies for OLI coatings on nonporous substrates28,35–37. When paired with the DO meter, OLI membranes provided superior long-term reproducibility, demonstrating a peak change in sensitivity of only 10.3% after 500 hours compared to 26% change with commercially available membranes. This was further characterized as a rate of decay in sensitivity that is 3.35 times higher in UT membranes. We also prove that DO can permeate the thin perfluorodecalin layer, and that the fluorosilane does not significantly disrupt the porosity and permeability of the membrane itself. Considering the smaller drop in contact angle on OLI membranes, the fluorosilane SAM was not compromised by significant corrosion or leaching that would have diminished the presence of hydrophobic fluorine on the surface. In previous studies, Epstein et al. demonstrated that an OLI coating was able to prevent over 95% of bacterial attachment for three different strains of bacteria over a 7-day incubation period38. However, in our work, the membranes were not incubated with pre-cultured strains of bacteria, but rather species gained from the ambient air and tap water, which is a closer representation of practical applications. We also demonstrated the OLI coating’s antibiofouling capabilities in a much longer experiment that lasted for three weeks.

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CONCLUSION This work presented the application of OLI coatings on selectively permeable membranes and demonstrates its use in attenuating surface biofouling. In the context of a membrane-based DO sensor, a tethered liquid perfluorocarbon OLI coating successfully suppressed the rate of biofilm formation over a three-week period. This enabled the DO sensor to maintain its sensitivity and provide more accurate readings in comparison to unmodified membranes. These findings present an effective solution that addresses the wide use of selectively permeable membranes in sensing technologies and the need for antibiofouling strategies in numerous areas of research and commercial industries. In addition to water quality monitoring, our developed OLI coating offers strategic surface blocking for DO measurement in healthcare. Hypoxemia, the state of oxygen deficiency in arteries, is a symptom of numerous respiratory disorders and can lead to respiratory failure. Similar to DO in natural waterbodies, dissolved plasma oxygen (roughly 2% of the total oxygen in blood)39 can be monitored in blood by implantable biosensors to evaluate patient health40,41. However, alike to bacteria, blood is capable of interacting with unblocked surfaces. Through the contact coagulation cascade26, clotted blood can cause pore blockages and encapsulation of analyte that would diminish a device’s reliability. In the case of bacterial infection in the bloodstream, which typically occur through the lungs or skin, biofouling may pose additional issues in the device design. OLI coatings are capable of simultaneously repelling bacteria and the plasma proteins responsible for coagulation42. This would enable a sensor to effectively measure DO concentrations in whole blood. A second application within healthcare is in the development of ingestible sensors. These electronic capsules with gas-permeable membranes are ingested by a patient and travel through 16 ACS Paragon Plus Environment

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their gastrointestinal track43,44. When in the gut, the capsules sense oxygen, hydrogen, and carbon dioxide gases to assess the chemical composition of the gut. Considering the high presence of bacteria in the gut, our OLI coating would be a strong candidate to suppress biofilm formation on the gas-permeable membrane and preserve the sensor’s functionality. Furthermore, previous work by Song et al. has demonstrated that OLI coatings are stable for a broad range of pH levels45 and would therefore be suitable for use in the gastrointestinal track. Supporting Information Available The following file is available free of charge. OLI_DO_Sensors_SI.docx. This document includes analysis of the bacterial composition and pH of the biofouling solution, and the response time and water retention exhibited by OLI and untreated membranes. AUTHOR INFORMATION Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

Funding Sources TFD acknowledges support from NSERC Discovery Grant and McMaster start up funds. PRS acknowledges support from the Canada Research Chairs Program and the Discovery Accelerator Supplement from NSERC. Declaration of Interest The authors declare that there are no conflicts of interest. 17 ACS Paragon Plus Environment

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ABBREVIATIONS DO, dissolved oxygen; OLI, omniphobic lubricant-infused; ECM, extracellular matrix; eDNA, extracellular DNA; SAM, self-assembled monolayer; UT, untreated; SEM, scanning electron microscopy; SCA, static contact angle; TPFS, trichloro (1H,1H,2H,2H-perfluorooctyl) silane; DAPI, diamidino-2-phenylindole dihydrochloride; PBS, phosphate buffered saline. REFERENCES (1)

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