Diatom Microbubbler for Active Biofilm Removal in Confined Spaces

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Biological and Medical Applications of Materials and Interfaces

Diatom Microbubbler for Active Biofilm Removal in Confined Spaces Yongbeom Seo, Jiayu Leong, Jun Dong Park, Yu-Tong Hong, Sang-Hyon Chu, Cheol Park, Dong Hyun Kim, Yu-Heng Deng, Vitaliy Dushnov, Joonghui Soh, Simon Rogers, Yi Yan Yang, and Hyunjoon Kong ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b08643 • Publication Date (Web): 14 Aug 2018 Downloaded from http://pubs.acs.org on August 15, 2018

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ACS Applied Materials & Interfaces

Diatom Microbubbler for Active Biofilm Removal in Confined Spaces Yongbeom Seo,1 Jiayu Leong,1,2 Jun Dong Park,1 Yu-Tong Hong,1 Sang-Hyon Chu,3 Cheol Park,4 Dong Hyun Kim,5 Yu-Heng Deng,1 Vitaliy Dushnov,1 Joonghui Soh,1 Simon Rogers,1 Yi Yan Yang,2 & Hyunjoon Kong1,6,* 1

Department of Chemical and Biomolecular Engineering, University of Illinois at Urbana-Champaign,

Urbana, Illinois 61801, United States 2

Institute of Bioengineering and Nanotechnology, 31 Biopolis Way, The Nanos, Singapore 138669,

Singapore 3

National Institute of Aerospace, 100 Exploration Way, Hampton, Virginia 23666, United States

4

Advanced Materials and Processing Branch, NASA Langley Research Center, Hampton, Virginia,

23681, United States 5

Department of Human and Culture Convergence Technology R&BD Group, Korea Institute of

Industrial Technology, Ansan-si, Gyeonggi-do 426-910, South Korea 6

Carle Illinois College of Medicine, Department of Bioengineering, Department of Pathobiology, Carl

R. Woese Institute for Genomic Biology, Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana−Champaign, Urbana, Illinois 61801, United States KEYWORDS: MnO2 Nanosheets, Diatom, Microbubble, Self-locomotion, Biofilm

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ABSTRACT: Bacterial biofilms form on and within many living tissues, medical devices, and engineered materials, threatening human health and sustainability. Removing biofilm remains a grand challenge despite tremendous efforts made so far, particularly when they are formed in confined spaces. One primary cause is the limited transport of anti-bacterial agents into extracellular polymeric substances (EPS) of the biofilm. In this study, we hypothesized that a microparticle engineered to be self-locomotive with microbubbles would clean a structure fouled by biofilm by fracturing the EPS and subsequently improving transports of the antiseptic reagent. We examined this hypothesis by doping a hollow cylinder-shaped diatom biosilica with manganese oxide (MnO2) nanosheets. In an antiseptic H2O2 solution, the diatoms doped by MnO2 nanosheets, denoted as diatom bubbler, discharged oxygen gas bubbles continuously and became self-motile. Subsequently, the diatoms infiltrated the bacterial biofilm formed on either flat or micro-grooved silicon substrates and continued to generate microbubbles. The resulting microbubbles merged and converted surface energy to mechanical energy high enough to fracture the matrix of biofilm. Consequently, H2O2 molecules diffused into the biofilm and killed most bacterial cells. Overall, this study provides a unique and powerful tool that can significantly impact current efforts to clean a wide array of bio-fouled products and devices.

1. INTRODUCTION Biofilms are communities of bacterial cells that are prevalent in many materials such as medical devices, injured/diseased living tissues, household products, and infrastructure.1-11 Multiple microbial cells build the biofilm by accumulating on solid-liquid interfaces and producing a protective matrix of extracellular polymeric substances (EPS), which includes various biomolecules, such as oligopeptides, lipopolysaccharides, proteins, lipids, and DNA.12-16 The resulting biofilms impact public health, product 2 ACS Paragon Plus Environment

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function, and infrastructural sustainability significantly.3-10 About 80% of all medical infections originate from biofilms of pathogens. Therefore, tremendous efforts have been conducted to remove biofilms from the substrates using various antibiotics and disinfectants.17-24 However, bacterial cells residing in biofilms are deemed 100 to 1,000 times more resistant to antibiotics and disinfecting agents than planktonic cells because the EPS matrix limits the transport of the anti-microbial agents and neutralizes them chemically.12-14, 24-25 Furthermore, the biofilm formed in confined spaces are even more challenging to eliminate using conventional methods than that on the open surface because the biofilm matrix becomes even sturdier and denser.26, 27 To this end, we hypothesized that microparticles assembled to generate microbubbles and selfpropel in the anti-bacterial H2O2 solution would act as an active cleaning agent. Due to the small size and active movement, the particles would be able to penetrate the EPS matrix of biofilm and continue to generate microbubbles inside the biofilm. The resulting microbubbles would give rise to a wave of mechanical energy high enough to deform the biofilm. In turn, H2O2 molecules would diffuse into the biofilm matrix and eradicate bacterial cells. We examined this hypothesis by doping manganese oxide (MnO2) nanosheets on the diatom biosilica, which is a skeleton of dead algae. MnO2 nanosheets can generate oxygen (O2) bubbles by decomposing H2O2.28,29 The porous and cylindrical geometry of diatom is advantageous to facilitate the diffusion of H2O2 into MnO2 nanosheets.

2. EXPERIMENTAL SECTION 2.1. Fabrication of the MnO2-diatom bubbler. the MnO2-diatom bubbler was fabricated using diatom particles doped with MnO2 nanosheets. Amine-substituted diatom particles were prepared by the reaction with (3-aminopropyl)triethoxysilane (APTES, Sigma-Aldrich). First, 2 g of diatom particles were added to 60 mL of toluene in a three-necked round-bottomed flask fitted with a thermometer, a 3 ACS Paragon Plus Environment


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reflux condenser, and an N2 gas tube. 0.6 mL of distilled water was added to the mixture and stirred for 2 h at room temperature. Then, 3.4 mL of APTES was added to the mixture, which was taken to reflux for 6 h at 60 oC. Second, the mixture was cooled and washed with toluene, 2-propanol, and distilled water 3 times. The obtained sample was dried in a vacuum desiccator for 2 days. Finally, 0.1 g of amine-substituted diatoms were added to 1 mL of 50 mM potassium permanganate (KMnO4, SigmaAldrich) solution and sonicated for 30 min at room temperature. Then, the samples were filtered and washed with distilled water and ethanol three times. The obtained sample was dried in an oven for 1 day at 60 oC. 2.2. Characterization of the MnO2-diatom bubbler. The optical images and movies of diatom bubblers were obtained with the optical microscope (Leica DMIL). The SEM images and elemental mapping of microparticles were obtained with HITACHI S-4700 microscope operating at 2.0 kV. The TEM images were obtained using JEOL 2100 TEM with an accelerating voltage of 200 kV. The elemental analysis of manganese within the diatom bubbler was conducted by the inductively coupled plasma atomic emission spectroscopy (ICP/AES). The nitrogen adsorption–desorption isotherms were measured at 77 K with the Quantachrome NOVA 2200e surface area and pore size analyzer. The specific surface area was calculated with the Brunauer–Emmett–Teller (BET) equation from the adsorption and desorption curve in the relative pressure (P/P0) range of 0.05-0.30. The pore volume and pore size distribution were calculated from the desorption curve by the Barrett–Joyner–Halenda (BJH) method in the relative pressure P/P0 range of 0.05-0.99. The powder X-ray diffraction data was obtained using the Rigaku Miniflex 600. The size distribution of microparticles was obtained by the image analysis with the ImageJ software. The speed of particles was measured by using the Fiji software. The mass of O2 gas generated from the decomposition of H2O2 was quantified by adding 2-4 mg diatom bubbler to 2 mL of 3 % H2O2 solution in a gas-sampling bottle and measuring the partial pressure of O2 in the gas phase with the sensor (Vernier). 4 ACS Paragon Plus Environment

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2.3. Preparation of microgrooved PDMS substrates. First, the silicon master was fabricated using the photolithography technique. To begin with, the flat silicon wafer was washed and dried. Then, a photoresist (SU-8 2050, MicroChem Corporation) was spin-coated (100 µm thickness) onto the wafer to reach desired thickness. Next, the substrate underwent soft bake, UV exposure, and post-exposure bake. During the UV exposure step, a photo mask with the designed pattern was placed on the wafer to cure only the area of interests. After the curing process, the excess uncured photoresist was washed out. The master was silanized by treatment with a vapor of trichloro (1H,1H,2H,2H-perfluorooctyl)silane, Sigma-Aldrich) to create a hydrophobic surface. Second, PDMS pre-polymer (SYLGARD 184 Silicon Elastomer Kit, Dow Corning) and its curing agent were thoroughly mixed in the 1:10 mass ratio. After degassing, the pre-polymer solution was cast onto the silicon master with the desired feature on it. Then, it was cured at 60°C for 2 hr. The cured PDMS with the desired pattern was gently peeled off from the master. The fabrication of the flat PDMS followed the same procedure. The only difference was that the pre-polymer solution was poured on the flat silicon wafer. 2.4. Biofilm growth on the PDMS substrate. Microbial broths were prepared using LB Broth, Lennox media (Difco™). Escherichia coli (ATCC No. 25922) was grown in the LB Broth at 37 °C under shaking (100 rpm) until they reached the mid-logarithmic growth phase. The concentration of E. coli was adjusted to obtain an optical density of 0.07 at the wavelength of 600 nm on a microplate reader (TECAN, Switzerland). This optical density corresponds to the concentration of McFarland 1 solution (3 × 108 CFU mL−1). E. coli (250 µL, 3 × 108 CFU mL−1). Flat or micro-grooved PDMS substrates were placed on the cell culture media in 24-well plates, while immersing only the groove parts in solution. After 24 h, the PDMS substrates were rinsed once with sterile PBS. Then, 250 µL of 3 × 108 CFU mL−1 fresh E. coli suspension was additionally added to the PDMS substrates. This process was repeated every day for 9 days to allow for the biofilm formation. After 9 days, the PDMS substrates were rinsed with sterile PBS before being used for further analysis. 5 ACS Paragon Plus Environment


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2.5. Biofilm removal from the PDMS substrate. After 9 days culture, biofilms grown on PDMS substrates were treated by replacing the cell culture media with 600 µL of 3% H2O2 solution or the mixture of 3 % H2O2 solution and the MnO2-diatom bubblers. After 30 min, all the solution was removed and gently rinsed once with PBS before the analysis. 2.6. Rheological analysis of the biofilm. Rheological properties of the biofilm samples grown on flat PDMS substrates were measured by strain-controlled oscillatory shear experiments using a rotational rheometer (ARES-G2, TA instrument). Oscillatory strain (amplitude)-sweep tests were performed to determine the storage (Gʹ) and loss modulus (Gʺ) with 25mm diameter parallel plate geometry at a fixed frequency of 1 rad sec-1. The dynamic moduli were measured with the biofilm on the PDMS substrate that is fixed to lower plate of the rheometer. All the analyses were conducted and averaged from the results of five different measurements with different samples. 2.7. Viability assay of the biofilm. The Cell Titer-Blue cell viability assay was performed to quantitatively analyze live E. coli cells attached to the PDMS. E. coli in the LB Broth, Lennox media (Difco™) (100 µL, 3 × 108 CFU mL−1) were seeded onto PDMS substrates cured in 24-well plates and cultured at 37 °C for 9 days. After 9 days, 600 µL of 3% H2O2 solution or the mixture of 3 % H2O2 and the MnO2-diatom bubbler particles was added to the biofilms for 30 min. The surfaces were washed twice with sterile PBS, followed by incubation with 100 µL of PBS and 20 µL of Cell Titer Blue Reagent at 37 °C for 6 h. The fluorescence intensity of resorufin resulting from reduction of resazurin by dehydrogenases of viable cells was determined at the excitation wavelength of 560 nm and the emission wavelength of 590 nm using the microplate reader. The experiment was conducted in replicates of 3. 2.8. Fluorescence imaging of the biofilm. Proteins in the biofilm were stained by incubating them for 1 h at room temperature with 500 µL of 10 mg mL-1 fluorescein isothiocyanate (Sigma6 ACS Paragon Plus Environment

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Aldrich, USA) in 0.1 M NaHCO3 buffer to conjugate the dye onto deprotonated amino groups. Then, the biofilm was washed with PBS before staining polysaccharides in the biofilm. Polysaccharides in the biofilms were stained by incubating the biofilm for 2 h with 500 µL of 250 µg mL-1 fluorescentlylabeled lectin concanavalin A conjugated with tetramethyl rhodamine (Molecular Probes, USA). The chemical binds to α-glucopyranosyl and α-mannopyranosyl sugar residues. Subsequently, the biofilms were washed with PBS before incubating them for 2 h with 500 µL of 300 µg mL-1 fluorescent brightener 28 (MP Biomedicals, LLC, USA), which binds to β-linked polysaccharides. The biofilms were washed with PBS. Then, the samples were mounted on glass slides and observed with a laser scanning confocal microscope (Zeiss LSM 700, Germany, a 200x objective).

3. RESULTS AND DISCUSSION 3.1. Synthesis and characterization of MnO2 nanosheets-doped diatom bubbler. The diatom particles used in this study have the hollow cylinder morphology with 10 µm-diameter and 18 µmlength while the wall exhibits many pores with an average diameter of 500 nm (Figures 1a and S1a). First, the diatom surface was functionalized with (3-aminopropyl) triethoxysilane (APTES) to present amine groups (Figure 1b). Then, MnO2 nanosheets were doped on the amine-functionalized diatom by reducing potassium permanganate (Figures 1b and S1b). The loading amount of MnO2 nanosheets was approximately 2.5 wt%, according to the inductively-coupled plasma-atomic emission spectroscopy analysis. The elemental mapping of Mn in SEM confirms that the MnO2 nanosheets are loaded on the diatom particles (Figure 1c). The high-resolution transmission electron microscopic (TEM) images show that MnO2 nanosheets are located on the porous wall surfaces (Figure 1d). According to the N2 adsorption analysis, the specific surface area and pore volume of MnO2 nanosheets-doped diatom (24 m2 g-1 and 0.057 cm3 g-1) are comparable to those of bare diatom particles (27 m2 g-1 and 0.076 cm3 g-1). This result indicates that most pores of the diatom particle were still open after deposition of MnO2 7 ACS Paragon Plus Environment


nanosheets (Figure 1e). On the other hand, the powder x-ray diffraction pattern of the MnO2 nanosheetsdoped diatom exhibits no difference from bare diatom. This result reveals that the doped MnO2 nanosheets on diatom particles are in an amorphous state (Figure 1f). The MnO2 nanosheet-doped diatom (MnO2-diatom) instantly generated O2 microbubbles in a 3 % H2O2 solution while the diatom without MnO2 nanosheets did not show any bubbles (Figure S2). Interestingly, O2 bubbles are continuously ejected from the hollow hole of the MnO2-diatom as shown in the time-lapse images and movie (Figure 2a and Movie S1). The bubbles showed a uniform diameter of 12 µm which is similar to that of diatom. From this result, we assume that H2O2 molecules in the solution diffuse through nanopores of the diatom wall and quickly decompose to generate O2 gas by reacting with MnO2 nanosheets (Figure 2b). The generated O2 gas bubbles start to nucleate and form microbubbles inside the hollow space of a diatom. As the bubbles build up the pressure, the diatom bubbler continuously jets microbubbles through the hollow channel. The ejection of bubble from the hollow channel allows the continuous diffusion of H2O2 molecules and generation of the microbubbles. Due to the continuous inward diffusion, no microbubbles were formed on the outer surface of MnO2diatoms. This spatial effect on the microbubble formation will be studied systematically in future. Subsequently, the MnO2-diatom moved at a rate of 60 µm s-1 (Figure 2c and Movie S2). The O2 gas generation from the diatom bubbler was also confirmed with the oxygen sensor. The initial reaction rates with different MnO2-diatom concentrations were calculated to determine the order of reaction (Figure 2d and Figure S3). The order of the reaction was approximately 1.2 with respect to the MnO2diatom concentration. 3.2. Active biofilm removal with diatom bubbler. Next, we examined the extent to which the MnO2-diatom bubbler cleans the flat polydimethylsiloxane (PDMS) substrate fouled by the Gramnegative Escherichia coli (E. coli) biofilm. First, the fouled PDMS substrate was exposed to 3% H2O2 solution for 30 minutes. This procedure resulted in macro-sized bubbles with a few hundred 8 ACS Paragon Plus Environment

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micrometers above the biofilm because the catalase in E. coli decomposed H2O2 into O2 gas (Movie S3). Approximately 80% of the biofilm remained on the PDMS substrates when they were examined with microscope images (Figures 3a, 3b, S4, and Movie S3). According to the oscillatory shear test, the storage modulus of the biofilm decreased from 20 to 10 Pa (Figure 3c). However, H2O2-treated biofilm showed a high yield strain, indicating that the structure of biofilm matrix was retained even after H2O2 treatment, suggesting no significant change to the structure. In contrast, the MnO2-diatom bubbler mixed with H2O2 solution generated microbubbles on the fouled PDMS substrate and removed most of the biofilm within 30 minutes (Figures 3a, 3b, S4, and Movie S3). Because there was no remaining biofilm on the substrate, it was not possible to measure the rheological property of the biofilm after the treatment of MnO2-diatom bubbler. We further examined the capability of the MnO2-diatom bubbler to invade and remove the biofilm formed in a confined space. To simulate the biofilm in confined spaces, we used the microgrooved PDMS substrates with controlled widths of 50, 100, 200, and 500 µm and depth of 100 µm. The biofilms were formed exclusively within microgrooves regardless of width. The confinement effect of biofilm formation could be explained by cell-to-cell signaling and interaction among bacterial cell clusters.26,27,30 Most of the biofilms in microgrooves of PDMS substrates were removed 30 minutes after the mixture of 3% H2O2 and the MnO2-diatom was added (Figure 3d Figure S5). In contrast, the biofilms still lingered on the microgrooves after 3% H2O2 solution treatment (Figure 3d and Figure S5). The quantitative analysis of the images revealed that the MnO2-diatom treatment cleaned 8-fold larger surface area than H2O2 solution treatment alone (Figure 3e). The cell viability gradually decreased with the time at the concentration of the MnO2-diatom bubbler being 1.0 mg mL-1 in 3% H2O2 solution (Figure S6a). On the other hand, increasing the concentration of the MnO2-diatom bubblers from 1.0 to 3.0 mg mL-1 decreased the cell viability a lot within 5 min (Figure S6b). As a result, 95 % of E. coli cells in the biofilm lost viability after the MnO2-diatom bubbler treatment, while most of the cells in the 9 ACS Paragon Plus Environment


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biofilm treated only with the H2O2 solution remained viable (Figure 3f). Moreover, the secondary treatment with a fresh mixture of 3% H2O2 and the MnO2-diatom bubblers led 99.9 % of E. coli cells to lose viability. In addition, the MnO2-diatom bubbler was effective to reduce the viability of E. coli in the detached biofilm (Figure S6c). The structure of biofilms in microgrooves before and after the treatment was analyzed with fluorescence images (Figure 4a and S7). The EPS matrix of untreated biofilm consists of β-linked polysaccharides (blue-colored) and α-glucopyranosyl and α-mannopyranosyl sugar residues (redcolored). Extracellular proteins (green-colored) were localized mostly on E. coli cell walls. Even after 3 % H2O2 solution treatment, a large amount of EPS and E. coli remained in the microgrooves, as quantified with the mean fluorescence intensity across the microgrooves. In contrast, the mixture of H2O2 and MnO2-diatom bubblers removed all EPS and E. coli cells, as confirmed with no fluorescence yield from the substrate. This result indicates that MnO2-diatom bubbler treatment could prevent biofilm from recovering because there was no residual amount of EPS and bacterial cells at all after the treatment. 3.3 Mechanism study of active biofilm removal. We monitored the intermediate stage of treatment to address the active cleaning mechanism of the MnO2-diatom bubbler in more detail. The self-locomotive diatom bubblers infiltrated the biofilm and continuously produced microbubbles within the biofilm (Movie S4). These microbubbles merged and burst to deform and fracture the biofilm until they collapsed. As a result, circular- or ellipsoidal-shaped damages could be found in between the biofilm while the diatom particles are observed inside the biofilm matrix (Figure 4b and 4c). In contrast, the biofilm exposed to the H2O2 solution exhibited macro-sized bubbles with a few hundred micrometers stayed outside the substrate due to the diffusion limitation (Figure 4c and Movie S5).


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The energy of microbubbles used to rupture the biofilm was estimated by calculating the change in the surface energy of microbubbles.31 The coalescence of the microbubbles is considered as a process to minimize surface energy.32 During this coalescence, the surface energy is released in the form of mechanical energy. Since initially formed microbubbles have a nearly spherical shape, the mechanical energy is calculated from surface energy difference between initial and final states as follows, ∆ = Σ4π Γ − Σ4π Γ

(1)

where ri, rf, and Γ represent the radius of bubbles at the initial state, the radius of bubbles at the final states, and the surface tension value for the water-air interface, respectively. We calculated the ∆E based on the results observed in optical microscope (Figure 4d and Movie S6). The calculated ∆E was compared to the mechanical work required to deform the biofilm in the observed area. Using the storage modulus (G′) of the untreated biofilm shown in Figure 3c, the required mechanical work per unit volume (Ereq) is given as,

= = ′

(2)

where Υ is the applied strain. According to the calculation (Supporting Information), the mechanical energy attained from the coalescence of microbubbles (3.02 x 10-10 J) far outweighs Ereq (2.03 x 10-12 J). This result indicates that the fusion between microbubbles from the diatom bubbler generates mechanical energy high enough to rupture the biofilm, confirming the experimental observations. We further demonstrated that the MnO2-diatom bubbler could remove the biofilm formed even in complex confined spaces (Figure 4e). The use of 3% H2O2 solution may cause corrosion on the metallic substrates. However, the MnO2-diatom particles decompose H2O2 gradually for the microbubble generation. Consequently, the MnO2-diatom bubbler not only improve anti-bacterial activity of H2O2



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during treatment period but also neutralize H2O2 in the end, which decrease the possibility of substrate corrosion.

4. CONCLUSION In summary, this study addresses that diatoms engineered to generate microbubbles and selfpropel can invade and damage the biofilm (Figure 4f). In the antiseptic H2O2 solution, diatoms doped by MnO2 nanosheets infiltrate the biofilm by microbubble propulsion. The diatom particles invading the biofilm continue to generate microbubbles that merge to deform and ultimately fracture the EPS matrix of biofilm. Then, H2O2 molecules diffuse into the biofilm more actively and induce bacterial cell death more efficiently than those free of the diatom bubbler as observed with the decrease in the cell viability over time. Past studies demonstrated various type of bubble-propelled particles and their applications.3337

However, to the best of our knowledge, this is the first study to assess the potential of active bubbling

particles for cleaning bio-fouled surfaces. In addition, this study utilizes a unique particle system created by hybridizing a H2O2–decomposing MnO2 nanosheets with naturally-derived diatom skeletons. As the diatoms are abundant in nature, the diatoms doped with MnO2 nanosheets can readily be produced on a large scale (Figure S8). Our results offer a promising path toward the effective development of active anti-biofilm system in biomedical and environmental applications.


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FIGURES

Figure 1. Fabrication and characterization of the MnO2 nanosheets-doped diatom. (a) SEM image of diatom. (b) Schematic illustration of fabrication steps for MnO2 nanosheets-doped diatom. (c) Elemental mapping images showing homogenous distribution of MnO2 nanosheets on diatom. (d) High-resolution TEM image of MnO2 nanosheets on diatom. (e, f) Nitrogen adsorption-desorption isotherms (e) and powder X-ray diffraction patterns (f) of diatom and MnO2 nanosheets-doped diatom.

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Figure 2. Self-locomotion of MnO2 nanosheets-doped diatom in H2O2 solution. (a) Time-lapse images of the microbubble generated from the diatom in 3 % H2O2 solution (Scale bar = 10 µm). (b) Schematic illustration of the mechanism by which the MnO2-diatom produces O2 bubbles and self-propels. (c) Tracking of the self-locomotion of MnO2-diatom bubbler added to H2O2 solution (Scale bar = 20 µm). (d) Quantification of O2 gas generated from 3 % H2O2 solution mixed with varying amount of the MnO2-diatom bubbler. Unmodified diatom was included as a control.


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Figure 3. Effects of the MnO2-diatom bubbler on the biofilm formed on flat or microgrooved PDMS substrate. (a, b) Optical (a) and SEM (b) images of E. coli biofilm grown on flat PDMS substrates before and after treatments of 3% H2O2 solution without and with MnO2-diatom. (c) Storage modulus (Gʹ) and loss modulus (Gʹʹ) of the biofilm as a function of the oscillatory strain amplitude. d) Optical image of the biofilm formed on microgrooved PDMS substrates (50 µm width and 100 µm depth) before and after treatments of 3% H2O2 solution without and with MnO2-diatom. e) Quantified biofilm area on microgrooved PDMS substrate. f) The viability of E. coli in the biofilm on the PDMS substrate. The secondary treatment with diatom bubbler led to the death of 99.9 % of cells, which is marked with an asterisk (*).


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Figure 4. Mechanistic study of the active biofilm removal using MnO2-diatom bubbler. (a) Fluorescent images of the β-linked polysaccharides (blue), α-glucopyranoayl/α-mannopyranosyl polysaccharides (red), and extracellular proteins (green) in extracellular polymer substances. (Scale bar = 10 µm). (b) Optical image of the intermediate stage of biofilm removal using MnO2diatom. Red and yellow arrows indicate the cleaned area in microgrooves by microbubbles and the invaded MnO2-diatom particles in biofilm matrix, respectively. (c) Optical images of the biofilm captured after 5 minutes of exposure to 3% H2O2 solution with and without MnO2diatom. (d) Time-lapse image of microbubbles within the grooves. Smaller microbubbles (dot circles) merge into a big bubble (yellow arrows), which collapses eventually. (e) Optical images of (i) the biofilm formed in the PDMS substrate with complex micropattern (i), and the biofilm treated by 3% H2O2 solution without (ii) and with MnO2-diatom (iii). (f) Schematic illustration of the active cleaning mechanism.


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ASSOCIATED CONTENT Supporting Information. Additional details on experimental methods and calculation. Figures showing the morphology and bubbling reaction of MnO2-diatom particles, the treatment of biofilm on flat or microgrooved PDMS substrates by H2O2 solution with and without diatom bubbler, fluorescent microscopic images of biofilm in microgrooves before and after the diatom bubbler treatment, and scale-up synthesis of MnO2-diatom particles. Movies showing bubbling and self-propelling motion of MnO2-diatom particles, biofilm on flat and microgrooved PDMS substrates by H2O2 solution with and without MnO2-diatom bubbler.

AUTHOR INFORMATION Corresponding Author *[email protected] Notes The authors declare no competing financial interest. ACKNOWLEDGMENT We acknowledge the financial support from the National Institutes of Health (Grant 1R01 HL109192 and 1R21 HL131469 to H. K.), National Science Foundation (STC-EBICS Grant CBET-0939511 to H. K.), and Korea Institute of Industrial Technology (JEI40004). J. L. acknowledges the A*STAR Graduate Scholarship (Overseas) from the Agency for Science, Technology, and Research. Y. Y. Y. acknowledges the support from the Institute of Bioengineering and Nanotechnology (A*STAR), Singapore. Electron microscopy was performed


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at the Frederick Seitz Materials Research Laboratory Central Facilities at the University of Illinois. ICP/AES and XRD were conducted at Microanalysis Laboratory (SCS CORES) at the University of Illinois.

REFERENCES (1) Chew, B. H.; Lange, D. Ureteral Stent Symptoms and Associated Infections: a Biomaterials Perspective. Nat. Rev. Urol. 2009, 6, 440. (2) Donelli, G., Biofilm-based healthcare-associated infections. Springer: 2014; Vol. 2. (3) Venkatesan, N.; Shroff, S.; Jayachandran, K.; Doble, M. Polymers as Ureteral Stents. J. Endourol. 2010, 24, 191-198. (4) Nguyen, C. T.; Jung, W.; Kim, J.; Chaney, E. J.; Novak, M.; Stewart, C. N.; Boppart, S. A. Noninvasive in vivo Optical Detection of Biofilm in the Human Middle Ear. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 9529-9534. (5) Donlan, R. M. Biofilm Formation: A Clinically Relevant Microbiological Process. Clin. Infect. Dis. 2001, 33, 1387-1392. (6) Guglielmi, G.; Chiarani, D.; Judd, S.; Andreottola, G. Flux Criticality and Sustainability in a Hollow Fibre Submerged Membrane Bioreactor for Municipal Wastewater Treatment. J. Memb. Sci. 2007, 289, 241-248. (7) Schultz, M.; Bendick, J.; Holm, E.; Hertel, W. Economic Impact of Biofouling on a Naval Surface Ship. Biofouling 2011, 27, 87-98. (8) Wu, H.; Moser, C.; Wang, H.-Z.; Høiby, N.; Song, Z.-J. Strategies for Combating Bacterial Biofilm Infections. Int. J. Dent. Oral Sci. 2015, 7, 1-7. (9) Agarwal, A.; Xu, H.; Ng, W. J.; Liu, Y. Biofilm Detachment by Self-Collapsing Air Microbubbles: a Potential Chemical-Free Cleaning Technology for Membrane Biofouling. J. Mater. Chem. 2012, 22, 2203-2207. (10) Gupta, A.; Das, R.; Tonga, G. Y.; Mizuhara, T.; Rotello, V. M. Charge-Switchable Nanozymes for Bioorthogonal Imaging of Biofilm-Associated Infections. ACS Nano 2018, 12, 89-94.


18

Page 19 of 21 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(11) Tesler, A. B.; Kim, P.; Kolle, S.; Howell, C.; Ahanotu, O.; Aizenberg, J. Extremely Durable Biofouling-Resistant Metallic Surfaces Based on Electrodeposited Nanoporous Tungstite Films on Steel. Nat. Commun. 2015, 6, 8649. (12) Leslie, D. C.; Waterhouse, A.; Berthet, J. B.; Valentin, T. M.; Watters, A. L.; Jain, A.; Kim, P.; Hatton, B. D.; Nedder, A.; Donovan, K. A Bioinspired Omniphobic Surface Coating on Medical Devices Prevents Thrombosis and Biofouling. Nat. Biotechnol. 2014, 32, 1134. (13) Voo, Z. X.; Khan, M.; Xu, Q.; Narayanan, K.; Ng, B. W.; Ahmad, R. B.; Hedrick, J. L.; Yang, Y. Y. Antimicrobial Coatings against Biofilm Formation: the Unexpected Balance between Antifouling and Bactericidal Behavior. Polym. Chem. 2016, 7, 656-668. (14) Flemming, H.-C.; Wingender, J. The Biofilm Matrix. Nat. Rev. Microbiol. 2010, 8, 623. (15) Epstein, A. K.; Pokroy, B.; Seminara, A.; Aizenberg, J. Bacterial Biofilm Shows Persistent Resistance to Liquid Wetting and Gas Penetration. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 9951000. (16) Sahoo, P. K.; Janissen, R.; Monteiro, M. P.; Cavalli, A.; Murillo, D. M.; Merfa, M. V.; Cesar, C. L.; Carvalho, H. F.; Souza, A. A.; Bakkers, E. P. A. M.; Cotta, M. A. Nanowire Arrays as Cell Force Sensors To Investigate Adhesin-Enhanced Holdfast of Single Cell Bacteria and Biofilm Stability. Nano Lett. 2016, 16, 4656-4664. (17) Flemming, H.-C.; Wingender, J.; Szewzyk, U.; Steinberg, P.; Rice, S. A.; Kjelleberg, S. Biofilms: an Emergent Form of Bacterial Life. Nat. Rev. Microbiol. 2016, 14, 563. (18) James, G. A.; Swogger, E.; Wolcott, R.; Secor, P.; Sestrich, J.; Costerton, J. W.; Stewart, P. S. Biofilms in Chronic Wounds. Wound Repair Regen. 2008, 16, 37-44. (19) Bridier, A.; Sanchez-Vizuete, P.; Guilbaud, M.; Piard, J.-C.; Naïtali, M.; Briandet, R. Biofilm-Associated Persistence of Food-Borne Pathogens. Food Microbiology 2015, 45, 167-178. (20) Peterson, B. W.; He, Y.; Ren, Y.; Zerdoum, A.; Libera, M. R.; Sharma, P. K.; Van Winkelhoff, A.-J.; Neut, D.; Stoodley, P.; Van Der Mei, H. C. Viscoelasticity of Biofilms and Their Recalcitrance to Mechanical and Chemical Challenges. FEMS Microbiol. Rev. 2015, 39, 234-245. (21) Pavlovsky, L.; Younger, J. G.; Solomon, M. J. In situ Rheology of Staphylococcus Epidermidis Bacterial Biofilms. Soft Matter 2013, 9, 122-131. (22) Meylan, S.; Andrew, I. W.; Collins, J. J. Targeting Antibiotic Tolerance, Pathogen by Pathogen. Cell 2018, 172, 1228-1238. (23) Stewart, P. S.; Costerton, J. W. Antibiotic Resistance of Bacteria in Biofilms. The Lancet 2001, 358, 135-138.


19


Page 20 of 21

(24) McKay, G.; Nguyen, D. Handbook of Antimicrobial Resistance 2017, 203-229. (25) Feng, G.; Cheng, Y.; Wang, S.-Y.; Borca-Tasciuc, D. A.; Worobo, R. W.; Moraru, C. I. Bacterial Attachment and Biofilm Formation on Surfaces Are Reduced by Small-Diameter Nanoscale Pores: How Small Is Small Enough? NPJ Biofilms Microbiomes 2015, 1, 15022. (26) Gu, H.; Chen, A.; Song, X.; Brasch, M. E.; Henderson, J. H.; Ren, D. How Escherichia Coli Lands and Forms Cell Clusters on a Surface: a New Role of Surface Topography. Sci. Rep. 2016, 29516. (27) Bodelón, G.; Montes-García, López-Puente, V.; Hill, E. H.; Hamon, C.; Sanz-Ortiz, M. N.; Rodal-Cedeira, S.; Costas, C.; Celiksoy, S.; Pérez-Juste, I.; Scarabelli, L.; Porta, A. L.; PérezJuste, J.; Pastoriza-Santos, I.; Liz-Marzán, L. M. Detection and Imaging of Quorum Sensing in Pseudomonas Aeruginosa Biofilm Communities by Surface-Enhanced Resonance Raman Scattering. Nat. Mater. 2016, 15, 1203–1211. (28) Seo, Y.; Leong, J.; Teo, J. Y.; Mitchell, J. W.; Gillette, M. U.; Han, B.; Lee, J.; Kong, H. Active Antioxidizing Particles for On-Demand Pressure-Driven Molecular Release. ACS Appl. Mater. Interfaces 2017, 9, 35642-35650. (29) Yang, G.; Xu, L.; Chao, Y.; Sun, X.; Wu, Y.; Peng, R. Liu, Z. Hollow MnO2 as a TumorMicroenvironment-Responsive Biodegradable Nano-Platform for Combination Therapy Favoring Antitumor Immune Responses. Nat. Commun. 2017, 902. (30) Gu, H.; Hou, S.; Yongyat, C.; Tore, S. D.; Ren, D. Patterned Biofilm Formation Reveals a Mechanism for Structural Heterogeneity in Bacterial Biofilms. Langmuir, 2013, 29, 1114511153. (31) Yasui, K., Bubble Dynamics. In Acoustic Cavitation and Bubble Dynamics, Springer: 2018; pp 37-97. (32) Massey, B. S.; Ward-Smith, J., Mechanics of fluids. Crc Press: 1998; Vol. 1. (33) Li, J.; Rozen, I.; Wang, J. Rocket Science at the Nanoscale. ACS Nano 2016, 10, 5619-5634. (34) Solovev, A. A.; Xi, W.; Gracias, D. H.; Harazim, S. M.; Deneke, C.; Sanchez, S.; Schmidt, O. G. Self-Propelled Nanotools. ACS Nano 2012, 6, 1751-1756. (35) Sánchez, S.; Soler, L.; Katuri, J. Chemically Powered Micro‐and Nanomotors. Angew. Chem. Inter. Ed. 2015, 54, 1414-1444. (36) Li, J.; Shklyaev, O. E.; Li, T.; Liu, W.; Shum, H.; Rozen, I.; Balazs, A. C.; Wang, J. SelfPropelled Nanomotors Autonomously Seek and Repair Cracks. Nano Lett. 2015, 15, 7077-7085.


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(37) Safdar, M.; Khan, S. U.; Janis, J. Progress toward Catalytic Micro- and Nanomotors for Biomedical and Environmental Applications. Adv. Mater. 2018, 1703660.

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Diatom Microbubbler for Active Biofilm Removal in Confined Spaces

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