Cleaning with Bulk Nanobubbles - Langmuir (ACS Publications)

Apr 23, 2016 - Therefore, it is possible that platinum particles are being liberated as nanoparticles and that these nanoparticles may influence our d...
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Cleaning with Bulk Nanobubbles Jie Zhu,† Hongjie An,‡ Muidh Alheshibri,‡ Lvdan Liu,† Paul M. J. Terpstra,§ Guangming Liu,*,† and Vincent S. J. Craig*,‡ †

Department of Chemical Physics, Hefei National Laboratory for Physical Sciences at the Microscale, University of Science and Technology of China, Hefei, PR China 230026 ‡ Department of Applied Mathematics, Research School of Physical Sciences and Engineering, Australian National University, Canberra ACT 2601, Australia § Consumer Technology Research Institute, Wageningen, The Netherlands S Supporting Information *

ABSTRACT: The electrolysis of aqueous solutions produces solutions that are supersaturated in oxygen and hydrogen gas. This results in the formation of gas bubbles, including nanobubbles ∼100 nm in size that are stable for ∼24 h. These aqueous solutions containing bubbles have been evaluated for cleaning efficacy in the removal of model contaminants bovine serum albumin and lysozyme from surfaces and in the prevention of the fouling of surfaces by these same proteins. Hydrophilic and hydrophobic surfaces were investigated. It is shown that nanobubbles can prevent the fouling of surfaces and that they can also clean already fouled surfaces. It is also argued that in practical applications where cleaning is carried out rapidly using a high degree of mechanical agitation the role of cleaning agents is not primarily in assisting the removal of soil but in suspending the soil that is removed by mechanical action and preventing it from redepositing onto surfaces. This may also be the primary mode of action of nanobubbles during cleaning.

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INTRODUCTION Nanobubbles or fine bubbles are small gaseous entities that are found on surfaces and in the bulk when solutions are supersaturated with gas.1,2 They were first proposed to explain unusual long-range attractive forces observed between hydrophobic surfaces3 and later imaged on surfaces using atomic force microscopy.4,5 They are of particular interest as the observed stability of nanobubbles is inconsistent with current theories of bubble dissolution.6,7 Moreover, surface nanobubbles exhibit exceptionally high contact angles that cannot currently be explained.8 Despite the theoretical challenges in understanding nanobubbles, they are already finding applications in a broad range of areas including medicine,9−11 crop production,12 controlling boundary slip,13 and bioremediation.14 The possible advantages of using nanobubbles in a broad range of applications include the ease with which they can be produced, the low cost of materials, and the potential to easily remove them from the system once they have performed their function. Additionally, they may provide solutions to industrial challenges with low environmental impact. It is for this reason that nanobubbles are seen as promising agents for cleaning. Cleaning in a scientific sense is often associated with detergency, but common experience tells us that cleaning does not require the presence of a surfactant or detergent if sufficient mechanical means is employed. Our prior work, as well as the work of others, describes the prevention of both fouling and cleaning using surface nanobubbles.15−17 The role of nanobubbles in preventing fouling is to provide a mechanical barrier to the adsorption of material on the surface. Thus, the regions © 2016 American Chemical Society

of the surface that are decorated with nanobubbles mask the surface from exposure to contaminants. When employed in this manner, the regions of the surface not covered by nanobubbles will still be fouled, so this is not a preferred mode of operation. However, it was shown that the production of nanobubbles on a surface that is already contaminated results in surface cleaning that can result in nearly all contamination being removed.16,17 Nanobubbles in these studies were prepared by electrolyzing water, with the resultant supersaturation of dissolved gas leading to the formation of surface nanobubbles. As the nanobubbles expanded, the advancing three-phase line, which marks the boundary between the nanobubble and the liquid on the surface, removed the contaminant from the surface. It was found that cleaning was more effective with nanobubbles than with sodium dodecylsulfate (SDS) surfactant that is commonly used in detergents and that cleaning was further enhanced when surfactant was also used.16,17 Surfactants clean by reducing the adhesive forces between the contaminant and the surface, whereas surface nanobubbles clean when the advancing three-phase line transfers contaminants from the substrate to the gas/solution interface. The high energy of the interface provides for powerful cleaning action and reflects the comparative strength of capillary forces compared to surface Special Issue: Nanobubbles Received: March 15, 2016 Revised: April 22, 2016 Published: April 23, 2016 11203

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positively buoyant particles of the same size (∼100−250 nm) as particles detected by dynamic light scattering, providing strong evidence for the existence of bulk nanobubbles. When a modulation interference microscope is used, particles can be distinguished on the basis of the refractive index and thereby bubbles (refractive index = 1) can be distinguished from other particles. Bunkin et al. have identified stable nanobubbles in the size range of 250−750 nm using this technique.30 Temperature changes have also been used to produce bulk nanobubbles 290 nm in diameter.31 Increasing the temperature reduces the solubility of dissolved gases, resulting in supersaturation. These experiments were performed with the temperature change being induced in a cell mounted in a light-scattering instrument. The conclusion is that particles produced upon increasing the temperature are nanobubbles as the solubility of most materials other than gas will increase with increasing temperature. Here we test the hypothesis that bulk nanobubbles produced through the electrolysis of aqueous electrolyte solutions enhance the cleaning capacity of the solution. Electrolysis results in the production of hydrogen and oxygen gas as well as a range of other chemical reactions. To control for the effect of chemical changes as opposed to physical changes in the solutions, we compare solutions that have been filtered to remove nanobubbles with those containing nanobubbles in order to determine the effectiveness of the nanobubbles as cleaning agents. Additionally, we use untreated solutions as control samples.

forces. Thus, cleaning with nanobubbles also involves a mechanical component, in addition to surface energetics. A particular advantage of using nanobubbles for cleaning is that there is no need to rinse surfactant residue from the surface after cleaning has been effected. Thus, nanobubbles have been proposed for use in specialist applications such as cleaning semiconductors.18 Furthermore, cleaning with nanobubbles has the potential advantages of reducing the considerable environmental impact of surfactant use, lowering the use of nonrenewable resources, and lessening emissions into the environment. Electrolysis has also been reported to produce nanobubbles in the bulk.19−25 Electrolysis results in the supersaturation of oxygen and hydrogen in the anodic and cathodic streams of the solution, respectively, causing bubbles to be nucleated. Larger bubbles quickly rise to the surface and burst, but particle sizing techniques show persistent nanoparticles of around 100 nm following electrolysis that are not present in control solutions. A number of authors have concluded that the nanoparticles produced by electrolysis are gaseous, as described below. This is consistent with the range of methods used for their production, which all involve supersaturation and the absence of nanoparticles in the control solutions that have been investigated. Kikuchi et al.23 electrolyzed water to generate solutions supersaturated with oxygen. The formation of nanoparticles in these solutions was detected by light scattering, and their concentration was analyzed by measuring the oxygen content. The dissolved oxygen level was analyzed with a dissolved oxygen (DO) meter and the Winkler titration method.26 It was shown that these methods access only the dissolved oxygen, not that present within bubbles, but by the addition of sulfuric acid, the stability of the nanobubbles was disrupted and a subsequent Winkler titration revealed an increase in oxygen concentration. The difference in oxygen concentration was used to determine the concentration of bulk nanobubbles of oxygen produced during electrolysis. Similarly, in previous work Kikuchi et al.19 investigated solutions supersaturated in hydrogen gas. By comparing the results from a dissolved hydrogen meter and chemical analysis, they found that the chemical analysis revealed a higher concentration of hydrogen gas and attributed the difference to the existence of hydrogen nanobubbles. Note that the chemical analysis required acidification. This is thought to remove the charge on the nanobubbles and destabilize them. The production of bulk nanobubbles by mechanical methods of supersaturation has also been reported. Ohgaki et al.27 reported that solutions of bulk nanobubbles consisting of nitrogen, methane, and argon with an average radius of 50 nm were stable for 2 weeks. They were produced by mechanical gas injection, which leads to a supersaturated solution. A scanning electron micrograph of replica films of rapidly frozen and fractured solutions showed a 50-nm-radius cavity. Ushikubo et al.28 reported the production of both oxygen and air-filled nanobubbles by mechanical methods. The size of the nanobubbles was characterized by dynamic light scattering. Additionally, it was claimed that nanobubbles caused an increase in the T1 relaxation time of the solutions as measured by NMR. Kobayashi et al.29 have used an Archimedes microelectromechanical sensor for the detection of nanobubbles. This technique is sensitive to the density of particles; therefore, particles that are positively buoyant are readily distinguished from negatively buoyant particles. Furthermore, the size of the particle is measured independently of the density. Using this technique, nanobubble solutions showed

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MATERIALS AND METHODS

Sodium chloride solutions of concentration 2.1 × 10−3 M were prepared with water that had been purified using either a Milli-Q gradient or an ELGA purelab Chorus 3 system, resulting in a solution with conductivity of ∼250 μs/cm. Nanobubbles were produced by electrolysis using an ec-H2O Nanoclean cell that employs five electrode plates of platinum with a plate spacing of 2.79 mm. The device and control electronics is a proprietry design of Tennant Company. Aqueous solution was treated electrochemically using a cell voltage of ∼24 V at a flow rate of 7.5 mL s−1. This results in the fluid transiting the plates in approximately 13 s. The electrolysis reaction produces oxygen and hydrogen gas in a molar ratio of 1:2 according to the reaction equations shown in Figure 1. Immediately after treatment, numerous visible bubbles are apparent in the solution. Because of buoyancy, the larger bubbles rise to the surface and burst, hence the appearance of the solution changes significantly during the first few minutes after production. Filtering was conducted on selected samples using syringe filters of either a 20 nm (0.02 μm Whatman Anotop 10 inorganic with an Al2O3 membrane) or 450 nm (0.45 μm Millipore Millex-LCR Hydrophilic PTFE membrane) pore size. For cleaning studies, we developed a protocol that could be used to determine if any cleaning observed was due to electrolysis and furthermore could determine the size of the species responsible for such action. This was specifically developed to enable any cleaning due to chemical changes that occur during electrolysis to be separated from cleaning associated with the nanoparticles produced during electrolysis. Note that this is necessary because other reactions will occur in the presence of chloride ions, including the formation of hypochlorous acid. In addition the effluent from the cathode and anode locally will be alkaline and acidic, respectively. It is known that alkalinity in particular can affect cleaning performance. However, the streams are mixed, and the change in pH of the solution upon electrolysis is minimal by the time the solutions are used. Five solutions were studied as depicted in Table 1. A NanoSight (NS300, Nanosight, software Nanosight V3.1) instrument for nanoparticle tracking analysis using a blue laser light source (70 mW, λ = 405 nm) at 25 °C was used to size the 11204

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for cleaning analysis. Silicon wafers were rinsed with water and ethanol and dried with a flow of nitrogen and then cleaned by water plasma treatment at a power of 18 W for 15 min. After these treatments, the silicon wafers were used as hydrophilic surfaces (contact angle ∼0°). Hydrophobic surfaces with a water contact angle of ∼107° were prepared by modifying the silicon wafers with a hydrophobic monolayer. Specifically, after treatment by water plasma, the activated substrates were modified via vapor deposition of dodecyltrimethoxysilane at 60 °C for 6 h and then rinsed with ethanol and water and dried with a flow of nitrogen to form the hydrophobic monolayer.

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RESULTS AND DISCUSSION Characterization of the Electrolyzed Solutions. We have not directly demonstrated that the nanoparticles produced by electrolysis are gaseous, but the balance of the evidence is that they are indeed nanobubbles. Consistent with previous reports of nanoparticle production from solutions supersaturated by electrolysis, we will henceforth refer to them as nanobubbles.19−25 The size and concentration of nanobubbles produced by electrolysis were determined using the Nanosight instrument. Samples of ec-H2O NaCl were filtered through a 450 nm filter before introduction into the NanoSight cell. The NanoSight revealed a population of nanobubbles of concentration (750 ± 50) × 106 particles/mL (Figure 2). The mean

Figure 1. Schematic showing the production of samples for investigation. The electrolysis of 2.1 mM NaCl solutions is used to produce a solution supersaturated with oxygen and hydrogen gases (ec-H2O), which is used as a test solution. Filtration through a 450 nm filter excludes all particles and bubbles larger than 450 nm and produces a second test solution (ec-H2O sub 450 nm). Filtration through a 20 nm filter removes the nanoparticles produced during electrolysis and provides a control solution in which the chemical effects of electrolysis are maintained (ec-H2O sub 20 nm). The untreated electrolyte sample, filtered or unfiltered, was also used as a control solution.

Table 1. Nomenclature for the Solutions Employed to Investigate Cleaning designation

description

NaCl(aq) NaCl(aq) sub 450 nm ec-H2O ec-H2O sub 450 nm ec-H2O sub 20 nm

purified water with 2.1 mM NaCl added purified water with 2.1 mM NaCl added that has been filtered through a 450 nm filter purified water with 2.1 mM NaCl that has been electrolyzed purified water with 2.1 mM NaCl that has been electrolyzed and then filtered through a 450 nm filter purified water with 2.1 mM NaCl that has been electrolyzed and then filtered through a 20 nm filter

nanoparticles and determine the concentration. Each measurement comprised an analysis of five movies, each 60 s long, captured at 25 frames/s. The camera level was set to 14, the threshold to 3, the gain to 366, and the shutter to 31.48 ms, and analysis was conducted using a solution viscosity of 0.888 cP. The Nanosight determines the size of individual particles from their diffusion under Brownian motion using nanoparticle tracking. The concentration is directly determined by counting the number of particles observed in a known volume. The tracking of numerous particles enables the size distribution of particles to be determined. Samples of ec-H2O were filtered through a 450 nm filter before introduction into the NanoSight measurement cell, to ensure that larger bubbles did not interfere with the measurements. A Zetasizer Nano ZS (Malvern) was also used to measure the size of nanoparticles by dynamic light scattering as well as the zeta potential by laser Doppler microelectrophoresis in an ac electric field. Proteins bovine serum albumin (BSA), sourced from Hualvyuan Biotechnology Co., and lysozyme, sourced from Bio Basic Int., were used as model contaminants. The amount of material was measured using spectroscopic ellipsometry (M2000 V, J. A. Woollam, U.S.A.) at two incident angles of 65 and 75° in air and at an incident angle of 75° in aqueous solutions after the adsorption of contaminants. The refractive index of the contaminant layer was estimated to be 1.45 to evaluate the layer thickness. The contact angle measurements were performed using a KSV (Helsinki, Finland) CAM 200 contact angle goniometer. Both hydrophilic and hydrophobic surfaces were prepared

Figure 2. Histogram showing the particle concentration in a bin width of 1 nm obtained using the Nanosight NS300 in ec-H2O sub 450 nm, NaCl(aq) sub 450 nm, H2O sub 450 nm, and ec-H2O sub 20 nm. The total concentration of ec-H2O sub 450 nm was (750 ± 50) × 106 particles/mL.

size of nanobubbles for the data shown was 112 ± 2 nm with a mode of 93 ± 4 nm. The error bars are from repeated measurements on the same sample. A 10−15% degree of variation is typically seen between samples. As a control, NaCl(aq) was filtered with a 450 nm filter before measurement, revealing a concentration of (28 ± 2) × 106 particles/mL. The only difference was that this sample was not passed through the ec-H2O electrolysis cell. Filtering ec-H2O through a 20 nm filter removed the nanobubbles, resulting in a concentration of nanoparticles of (3 ± 1) × 106 particles/mL, which is near the lower limit of the range of the apparatus. Similarly, filtering the laboratory-grade water through a 450 nm filter showed a concentration of nanoparticles of (3 ± 1) × 106 particles/mL. It is also important to ascertain the lifetime or the stability of the nanobubbles that are produced by electrolysis. The stability 11205

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Figure 3. (A) Mean bubble size distribution for ec-H2O sub 450 nm measured using light scattering over 10 h. (B) Histogram showing the particle concentration in a bin width of 1 nm measured using the Nanosight on samples of ec-H2O sub 450 nm over 5 days.

of the solutions was first examined using dynamic light scattering (Zetasizer, Malvern). An ec-H2O sub 450 nm sample was checked at intervals to ascertain the size of the nanobubbles. The cuvette was left in place throughout the measurement. The particle size was found to be stable for 10 h, as shown in Figure 3a. These experiments were hampered by the formation of bubbles on the surface of the cuvette preventing the size of the nanobubbles from being followed for longer periods. To evaluate the stability over a longer period of time, a freshly prepared sample of ec-H2O sub 450 nm was measured at an interval of 1 day using the NanoSight. The syringe was filled with the sample immediately after production and remained undisturbed and sealed for 5 days. Each day, more of the sample was measured. Daily measurements confirmed that the size did not change significantly. Although the mean nanobubble size increased from 96 nm on the first day to 120 nm on the third day, the concentration decreased significantly over this time, as shown in Figure 3b and Table S1. This indicates that the size and concentration of nanobubbles are maintained for approximately 24 h after production. Note that the ec-H2O Nanoclean cell has platinum-coated electrodes, which degrade over time. Therefore, it is possible that platinum particles are being liberated as nanoparticles and that these nanoparticles may influence our data and/or the cleaning processes. A calculation was performed to determine if the concentration of nanoparticles observed could be due to platinum particles based on the known rate at which the electrodes degrade. This calculation revealed that if the nanoparticles produced were solely platinum then all of the platinum in the ec-H2O Nanoclean cell module would be exhausted after less than 30 L had been passed through the unit. This is inconsistent with the lifetime of the unit, which shows that many thousands of liters are produced before the platinum is substantially removed. Furthermore, our experiments conducted over a period of more than a year (and after hundreds of liters have been produced) show a consistent production of nanoparticles in both size and concentration. Additionally, it is observed that in a given sample the concentration of nanoparticles observed decreases steadily over a period of days but the size remains rather consistent. This is not consistent with the presence of platinum particles,

which we would expect to be either stable or grow substantially in size before settling in solution. We can therefore discount the possibility that the electrolysis process is producing a stream of platinum nanoparticles that are interfering in our experiments. At normal pH values, it has long been known that bubbles have a negative surface charge. This is attributed to the preferential adsorption of hydroxide ions.32,33 The zeta potential of ec-H2O sub 450 nm nanobubbles as a function of pH is shown in Figure 4. The isoelectric point (iep) is

Figure 4. Zeta potential of ec-H2O sub 450 nm as a function of pH. The measured zeta potential is consistent with literature reports of the zeta potential made on micrometer-sized bubbles.34

observed at a pH of ∼3.7. This is consistent with the measurement of the zeta potential on larger bubbles, as is the magnitude of the zeta potential in a solution of this concentration of NaCl.34 Cleaning of Contaminated Surfaces with Nanobubbles. The first mode of cleaning that we investigated was whether ec-H2O could remove BSA and lysozyme from a hydrophilic surface. The proteins (BSA and lysozyme) were used as model contaminants because they had been the subject of earlier cleaning experiments using surface nanobubbles.15,17 11206

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Figure 5. (a) Exposure of BSA previously adsorbed on a hydrophilic silicon wafer to both nanobubble and control solutions. (b) Exposure of lysozyme previously adsorbed on a hydrophilic silicon wafer to both nanobubble and control solutions.

Figure 6. Adsorption inhibition study performed using ellipsometry to measure the thickness of adsorbed BSA to a hydrophilic silicon wafer as a function of time. Here, the thickness of adsorbed protein was measured in air. (a) BSA adsorption from NaCl solution (2.1 mM). (b) BSA adsorption from ec-H2O solution. (c) BSA adsorption from ec-H2O sub 20 nm solution. (d) BSA adsorption from ec-H2O sub 450 nm solution.

had no cleaning effect on the lysozyme-coated silicon wafer surface. Preventing Surface Contamination. It is known that effective cleaning often requires mechanical agitation. This agitation will release contaminants from the surface; however, for the cleaning to be effective, it is important that the material is not redeposited. The suspension of soil in the cleaning fluid and the removal of the fluid are important in this respect. In situations where the cleaning process involves short periods of exposure to cleaning solutions of a few seconds, we considered the possibility that the cleaning effect of both detergents and nanobubbles in commercial applications may be achieved by stabilizing contaminants in solution that have been detached by mechanical scrubbing rather than removing contaminants from the surface. Therefore, we decided to evaluate whether ec-H2O can prevent the deposition of material to a surface. To do so, we dissolved different concentrations of BSA in solutions and observed the adsorption of BSA to a surface over time. If the amount of material adsorbing to the surface is reduced, then this would indicate the effective prevention of deposition of the contaminant from solution. Figure 6 shows the adsorption of BSA over a 40 min period from different types of solutions. It is clear that the adsorption from ec-H2O and NaCl is the same at

BSA at a concentration of 1.0 mg/mL dissolved in NaCl solution (2.1 mM) was allowed to adsorb to a clean hydrophilic silicon wafer for 60 min with a wet thickness of ∼2.4 nm in aqueous solution. Then the surface was treated with different types of solutions, and the thickness of the adsorbed BSA layer was measured as a function of time of treatment by ellipsometry in situ in NaCl solution (2.1 mM). To avoid the interference of nanobubbles in the ellipsometry measurements for ec-H2O, ec-H2O sub 450 nm, and ec-H2O sub 20 nm solutions, the relevant solutions were replaced by NaCl solution every 10 min to measure the protein layer thickness. It was found that ec-H2O and ec-H2O sub 450 nm cleaned the surface with similar effectiveness, whereas the control NaCl solution and ec-H2O sub 20 nm solution had no effect. (Figure 5). This fact suggests that the nanobubbles in the size range between 20 and 450 nm contribute to the cleaning effect. This is consistent with the results observed in Figures 2 and 3, where the size of the nanobubbles is distributed around 90 nm. Likewise, lysozyme molecules were adsorbed onto the hydrophilic silicon wafer surface from a solution of 1.0 mg/mL lysozyme for 60 min, producing a wet thickness of ∼1 nm. Lysozyme can also be removed by ec-H2O and ec-H2O sub 450 nm solutions. In contrast, the NaCl solution and ec-H2O sub 20 nm solution 11207

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Figure 7. Adsorption inhibition study performed using ellipsometry to measure the thickness of lysozyme adsorbed to a hydrophilic silicon wafer as a function of time. Here, the thickness of adsorbed protein was measured in air. (a) Lysozyme adsorption from NaCl solution (2.1 mM). (b) Lysozyme adsorption from ec-H2O solution. (c) Lysozyme adsorption from ec-H2O sub 20 nm solution. (d) Lysozyme adsorption from ec-H2O sub 450 nm solution.

H2O sub 450 nm are almost identical and, as described above, reduce the adsorption of BSA. It is therefore apparent that nanobubbles (not larger bubbles or chemical effects) are responsible for the inhibition of BSA adsorption to the surface. Lysozyme is a protein that we have previously found to adsorb strongly to surfaces and is harder to remove from a surface than BSA protein using surface nanobubbles.16 It therefore poses a more significant cleaning challenge. The adsorption of lysozyme to silicon wafers in the presence of nanobubbles was investigated to determine if nanobubbles are also effective at reducing the adsorption of lysozyme. The results found for lysozyme parallel those of BSA and indicate that the nanobubbles in ec-H2O inhibit the adsorption of lysozyme to the surface (Figure 7). The control solutions showed that the chemical changes accompanying electrolysis were not responsible for the inhibition of fouling by lysozyme but rather that nanobubbles smaller than 450 nm were responsible for the inhibition of fouling (Figure S2). We can conclude that nanobubbles contribute to the cleaning process by preventing the redeposition of lysozyme. However, the cleaning process was evident for lysoyme concentrations only up to 0.01 mg/mL, whereas BSA solutions up to a concentration of 0.1 mg/mL showed evidence of adsorption prevention by nanobubbles. Thus, preventing fouling by BSA is more easily achieved than for lysozyme. Hydrophobic Surfaces. The nature of the soiled surface (or substrate), in particular, the wettability, is an important consideration when evaluating the cleaning efficiency. In aqueous solutions, hydrophobic surfaces are generally more difficult to clean than hydrophilic surfaces as water molecules interact strongly with hydrophilic surfaces and therefore will strongly compete with contaminants at the surface. This effect is reduced substantially for hydrophobic surfaces. Therefore, we have also investigated the efficacy of nanobubbles in cleaning hydrophobic surfaces. BSA or lysozyme at a concentration of

the highest concentration of BSA studied (1.0 mg/mL) but at lesser concentrations ec-H2O inhibits the adsorption of BSA (Figure 6a,b). The electrolysis of aqueous NaCl solutions results in the supersaturation of hydrogen and oxygen gas but also drives a number of chemical reactions, including the production of hypochlorous acid from the production of Cl2(g) and its subsequent dissolution. To determine if the cleaning effects of ec-H2O were attributable to chemical reactions as opposed to nanobubbles, we investigated solutions in which the nanobubbles were removed by further filtration. We have established that the nanobubbles are larger than 20 nm and filtering them with a 20 nm filter removes them from solution (Figure 2). This filtration step will not remove the soluble chemical species produced during electrolysis, so solutions of ec-H2O sub 20 nm were investigated as control solutions. In this scenario, if cleaning is effective with ec-H2O and ec-H2O sub 450 nm and not effective in untreated solution or ec-H2O sub 20 nm, then the conclusion would be that the cleaning is due to electrolysis and the objects responsible for the cleaning are larger than 20 nm and smaller than 450 nm in diameter. That is, the cleaning action would be attributable to nanobubbles. The adsorption of BSA from ec-H2O sub 20 nm and ec-H2O sub 450 nm under the same conditions was also measured (Figure 6c,d). In the absence of nanobubbles (ec-H2O sub 20 nm), the adsorption is greater, indicating that the nanobubbles play a direct role in preventing the adsorption of BSA to the surface. The adsorption from NaCl and the adsorption from ecH2O sub 20 nm are compared directly, as is the adsorption from ec-H2O and ec-H2O sub 450 nm (Figure S1). What is found is that the data does not significantly differ between the two control solutions of NaCl and ec-H2O sub 20 nm, indicating that the chemical changes to the solution that take place during electrolysis have a minor effect on the adsorption of BSA. It is also apparent that the results for ec-H2O and ec11208

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Figure 8. (a) Exposure of BSA previously adsorbed on a hydrophobized silicon wafer to both nanobubble and control solutions. (b) Exposure of lysozyme previously adsorbed on a hydrophobized silicon wafer to both nanobubble and control solutions.

Figure 9. Adsorption inhibition study performed using ellipsometry to measure the thickness of adsorbed BSA to a hydrophobized silicon wafer as a function of time. Here, the thickness of adsorbed protein was measured in air. (a) BSA adsorption from NaCl solution (2.1 mM). (b) BSA adsorption from ec-H2O solution. (c) BSA adsorption from ec-H2O sub 20 nm solution. (d) BSA adsorption from ec-H2O sub 450 nm solution.

1.0 mg/mL dissolved in NaCl solution (2.1 mM) was allowed to adsorb to a hydrophobized silicon wafer for 60 min. Then the surface was exposed to a range of solutions (Table 1), and the thickness of the adsorbed protein layer was measured as a function of time by ellipsometry to assess the ability of nanobubbles to remove protein molecules. The results are summarized in Figure 8. It is clear that none of the solutions studied were effective at removing either BSA or lysozyme from a hydrophobic surface. The strong adhesion between the protein molecules and the hydrophobic surface can be attributed to the reconfiguration of protein conformation upon adsorption to maximize the interaction of hydrophobic groups with the surface. These experiments did not involve any mechanical agitation, which is often employed in cleaning processes. Mechanical agitation would be expected to improve the cleaning efficiency. The efficacy of using nanobubbles to prevent the fouling of a hydrophobic surface was also studied as a function of BSA concentration over time to ascertain if ec-H2O can prevent the deposition of material to a hydrophobic surface. To do so, we dissolved different concentrations of BSA in solution and observed the adsorption of BSA to a hydrophobic surface over time. If the amount of material adsorbing to the hydrophobic

surface is reduced in the presence of nanobubbles, then this would indicate the effective stabilization of the contaminant in solution by the nanobubbles. This data is shown in Figure 9. Several observations can be made from this data. At the lowest two concentrations (0.001 and 0.01 mg/mL BSA), ecH2O and ec-H2O sub 450 nm prevent the adsorption of BSA from solution onto the hydrophobic surface. It is apparent that ec-H2O sub 20 nm also prevents the adsorption of BSA on the hydrophobic surface, though this is not as strong as when nanobubbles are present (ec-H2O sub 450 nm). This indicates that the nanobubbles prevent the fouling of the surface and the chemical changes associated with the electrolysis of water may also contribute to the prevention of fouling (Figure S3). At the highest concentration of BSA (1.0 mg/mL), the cleaning influence of the solution is swamped by the high concentration of contaminant in solution and fouling is not reduced. The results of the prevention of lysozyme adsorption on the hydrophobized silicon wafer are similar to those observed for BSA adsorption (Figures S4 and S5). The data above shows that ec-H2O solution has a limited capacity for preventing protein adsorption on hydrophobic and hydrophilic surfaces: at high protein concentration, no inhibition is observed. A possible mechanism by which ec11209

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nanobubbles are gaseous, they will leave no residue. Additionally, the environmental impact of nanobubble production is likely to be significantly less than for the production of surfactant.

H2O could be inhibiting protein adsorption involves the protein adsorbing to the nanobubbles in the bulk. This would decrease the available concentration of protein left in solution for adsorption. In this case, the greater effectiveness in preventing the adsorption of BSA to the substrate would be correlated to a greater affinity of BSA for the nanobubble− solution interface. It is known that a high level of protein coverage on a surface is typically up to 2.0 mg/m2. Given the concentration and size of nanobubbles (Figure 2), we can calculate the total surface area of the nanobubbles per unit volume and therefore the concentration of protein that the nanobubbles can theoretically remove from solution through adsorption (Figure S6). The calculation predicts that nanobubbles should significantly influence the adsorption of protein up to a concentration of ∼1.0 × 10−3 mg/mL. In reality, the influence is observed at protein concentrations 100 times this level. This indicates that this model of protein inhibition is unlikely to be correct unless the actual concentration or surface area of nanobubbles is far greater than our measurements indicate. Indeed, recent work suggests that the concentration of nanoparticles determined by the Nanosight in flow mode is underestimated by up to 50%,35 though this discrepancy is insufficient to explain our results. A direct measure of the surface area of the nanobubbles would be very useful in exploring this discrepancy but is not currently possible. Another possibility is that the configuration of the proteins is altered by their interaction with the nanobubbles such that they are less attracted to the interface. Further research is required to determine the mechanism by which nanobubbles inhibit the adsorption of contaminants. Some industrial cleaning applications are carried out very quickly and utilize significant mechanical energy to remove contaminants, e.g., the scrubbing machines employed to clean hard floors, which require mechanical scrubbing for effective cleaning and involve the cleaning solution being in contact with the floor for only a few seconds. It has been demonstrated in careful studies that the adsorption of surfactant to a surface typically takes more than several seconds.36−38 This indicates that the accepted mechanism of detergency whereby surfactants adsorb at the interface and lower the energy required to remove contaminants cannot be responsible for cleaning in scrubbing machines because on the short time scale that the liquid is in contact with the surface the detergent has insufficient time to adsorb to the surface. We suggest that the accepted model of detergency does not apply to floor scrubbing machines but rather that other mechanisms are at play. In applications where mechanical agitation is important, we propose that the most significant contribution to the removal of a contaminant from a surface is the mechanical energy applied to the surface. The soil removed by mechanical means must be stabilized in solution to prevent redeposition for effective cleaning. Surfactants are effective at stabilizing soils from redeposition, particularly over short time periods as they adsorb to hydrophobic materials to make them hydrophilic. That is, the role of detergents is to prevent the soil from redepositing on the surface. The evidence presented above indicates that nanobubbles may promote cleaning by a similar mechanism. That is, dislodged contaminants may be prevented from redepositing by the action of nanobubbles and are thereby stabilized in solution. The use of nanobubbles in cleaning applications may present some advantages over the use of surfactant solutions. Once cleaning has been carried out, it is necessary to remove surfactant residues from the surfaces being cleaned. Because

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CONCLUSIONS We draw a number of conclusions from this work. The electrolysis of NaCl solutions using the ec-H2O Nanoclean device produces nanoparticles that are stable for approximately 24 h and are most likely to be gaseous. These nanobubbles are ∼90 nm in diameter and are present at a concentration of ∼5 × 108 per mL. The nanobubbles are readily removed using a 20 nm filter. Nanobubbles produced in NaCl solutions have a negative charge at neutral pH. Nanobubbles are able to clean already fouled hydrophilic surfaces, but this was not apparent on hydrophobic surfaces. Nanobubbles are able to prevent the fouling of both hydrophilic and hydrophobic surfaces. This capacity for fouling prevention is overcome as the concentration of contaminant increases and as the affinity of the contaminant for the surface increases. In applications where mechanical agitation is important for rapid cleaning, the dominant role of detergents in enhancing cleaning is in the prevention of the redeposition of soil that has been mechanically removed from the surface. The cleaning action of nanobubbles is likely to be predominantly through the same mechanism.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.6b01004. Size and concentration of nanobubbles in ec-H2O sub 450 nm over 5 days, adsorption inhibition study performed using ellipsometry to measure the thickness of adsorbed BSA and adsorbed lysozyme to both hydrophilic and hydrophobic surfaces as a function of time, and calculated protein adsorption capacity of ecH2O (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (G.M.L.). *E-mail: [email protected] (V.S.J.C.). Notes

The authors declare the following competing financial interest(s): Tennant Company provided financial support and the loan of the ec-H2O Nanoclean electrolysis cells. Tennant Company and their employees were not involved in the data analysis, writing, or preparation of the material presented in this article, nor were they provided with a draft of this article before submission.

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ACKNOWLEDGMENTS V.S.J.C. gratefully acknowledges the funding support of the Australian Research Council through the Discovery program. G.L. gratefully acknowledges the funding support of the National Natural Science Foundation of China (21374110 and 21574121). We thank Åsa Jamting and Victoria Coleman of the Nanometrology group in the Australian National Measurement Institute for access to equipment and assistance during the early stages of this project. We thank Tennant 11210

DOI: 10.1021/acs.langmuir.6b01004 Langmuir 2016, 32, 11203−11211

Article

Langmuir Company for financial support and the loan of the ec-H2O Nanoclean electrolysis cells. Tennant Company and their employees were not involved in the data analysis, writing, or preparation of the material presented in this article, nor were they provided with a draft of this article before submission.

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DOI: 10.1021/acs.langmuir.6b01004 Langmuir 2016, 32, 11203−11211