Colorimetric Detection of Urease-Producing Microbes Using an Ammonia-Responsive Flexible Film Sensor

Urease-producing (ureolytic) microbes have given rise to environmental and public health concerns because they are thought to contribute to emissions of ammonia and to be a virulence factor for infections. Therefore, it is highly important to have the ability to detect such microbes. In this study, a poly(dimethylsiloxane) (PDMS)-based colorimetric film sensor was employed for the detection of urease-producing microbes. The sensor was able to detect the enzyme activity of commercially available urease, as the color and absorbance spectrum of the sensor was observed to change upon being exposed to the reaction catalyzed by urease. The ratio of the absorbance of the sensor at 640 nm to that at 460 nm (A640/A460) was linearly proportional to the amount of urease present. The performance of the sensor was validated by the results of a sensitivity and selectivity analysis towards thirteen different bacterial strains. Based on the development of blue color of the sensor, the tested bacteria were classified as strongly positive, moderately positive, weakly positive, or negative urease producers. The response of the sensor to ureolytic bacteria was verified using the urease inhibitor phenyl phosphorodiamidate (PPDA). Additionally, the sensor achieved the selective detection of ureolytic bacteria even in the presence of non-ureolytic bacteria. In addition, a used sensor could be reverted to its original state by being subjected to simple aeration, and in this way the same sensor could be used at least five times for the detection of bacterial urease activity.


Introduction
Urease (urea amidohydrolase EC 3.5.1.5) is ubiquitously found in plants, fungi, bacteria, algae, and even invertebrates [1,2], where it performs basic biological functions, but it also has widespread implications for the environment, society, and human health due to its catalyzing the consumption of urea and hence causing the production of ammonia. Specifically, this nickel-dependent metalloenzyme catalyzes the hydrolysis of urea (CO(NH 2 ) 2 ) to ammonia (NH 3 ) and carbamic acid (H 2 NCOOH), which itself undergoes a spontaneous hydrolytic decomposition to carbonic acid (H 2 CO 3 ) and another molecule of ammonia ( Figure 1) [2][3][4][5]. As a result of its catalyzing the production of ammonia, urease increases the pH of the surrounding environment.
Urease-producing (ureolytic) microbes are present in various settings. For example, they are found in the soil and water environments. But since urea is a source of organic nitrogen (N) and hence is important for soil fertility (note, for example, that urea-based fertilizers as a nitrogen source accounted for 55% of synthetic fertilizers globally in 2014 [6]), the transformation of urea to ammonia gas due to the actions of ureolytic microbes results in ammonia emissions and in a loss of nitrogen fertilizer for agriculture and has thus created environmental and societal concerns [6]. Perhaps even more importantly, ureolytic microbes are also present in the bodies of humans and other animals, and various pathogenic microbes produce urease in order to utilize urea as a nitrogen source [7]. Urea is the major nitrogenous waste product of most terrestrial animals, including humans; thus, bacteria colonized in human bodies exploit, by hydrolyzing urea, the resulting ammonium product as a nitrogen source [8]. Moreover, urease helps human pathogens, including Helicobacter pylori, Yersinia enterocolitica, and Proteus mirabilis, adapt to acidic environments [7,[9][10][11][12]. The priority pathogen list of the World Health Organization (WHO) includes urease-producing bacteria, and these bacteria utilize urease as a virulence factor to infect and colonize the host [1,2,7,13]. Therefore, urease-producing microbes are of great interest due to the relevance of their enzymatic activity in infections [7,8,14,15].
Urease activity can be measured using commercially available kits, such as the urease activity assay kit produced by Sigma-Aldrich (Saint Louis, MO, USA) and a specifically ammonia assay kit produced by Abcam (Cambridge, UK). These assays, based on the Berthelot method, form a colored product-so a positive response can be detected colorimetrically [16]. The kits are easy to use and provide a sensitive performance for the detection of urease activity but often require sample preparation steps, such as filtration or deproteination prior to testing. In addition, the rapid urease test, also known as the Campylobacter-like organism (CLO) test, is used for providing a diagnosis of Helicobacter pylori infection [17,18]. The test depends on pH, so an increase in the pH in the medium caused by urease from Helicobacter pylori induces a color change of the specimen, specifically from yellow to red. The test allows a simple and rapid diagnosis of Helicobacter pylori [17]. However, the color change is influenced by the pH of the sample environment, so false-negative or false-positive results sometimes occur [19,20].
Urease-producing microbes have also been detected using a pH-independent colorimetric assay [21]; specifically, Santopolo et al. employed gold nanoparticles (AuNPs) for the color signal generation, with AuNP aggregation prevented in the presence of urease and showing a red color but allowed in its absence and showing a blue color, providing a sensitive detection of urease-positive bacteria [21].
In this study, we deployed a three-layered colorimetric film sensor for the detection of ureolytic microbes. The sensor included poly(dimethylsiloxane) (PDMS) and was shown to be responsive to ammonia gas [22], changing its color from yellow to blue due to the reaction catalyzed by urease. The colorimetric signal of the sensor was measured using commercially available urease. The performance of the sensor for the detection of urease activity was validated by carrying out a sensitivity and selectivity analysis against thirteen different microorganisms. Based on their pathogenic potential, thirteen bacteria were selected as model microbes in this study. Urease-producing microbes were identified by the observed changes in the color and absorbance spectrum of the sensor. Finally, the reversibility and reproducibility of the sensor was investigated to determine its amenability to repeated use.

Preparation of a Colorimetric Three-Layered Film Sensor
The colorimetric film sensor consisted of a bromocresol green (BCG)-incorporated PDMS sensing layer, which was sandwiched by the top and bottom protection films of PDMS (Figure 1a), as reported in our previous study [22]. Briefly, PDMS and its curing agent (K1 solution, Gwangmyeong-si, Gyeonggi-do, Korea) (10:1, w/w) were first mixed with a spatula. The pre-mixed PDMS was degassed in a plastic desiccator connected to a vacuum pump (Edwards Ltd., Burgess Hill, UK) to eliminate bubbles. The degassed PDMS of 0.5 g was carefully transferred to a 90-mm-diameter Petri dish plate and spread out on the dish using a spin coater (ACE-200, Dong Ah Trade Co., Seoul, Korea) at 6000 rpm for 15 s. The bottom PDMS layer was fully cured at 70 • C on a hot plate (PC-420D, Corning, New York, NY, USA) for 1 h. The sensing BCG-PDMS layer was prepared by mixing bromocresol green sodium salt (BCG, Sigma-Aldrich) and the pre-mixed PDMS. One hundred microliters (100 µL) of a BCG solution (37.5 mg/mL in deionized (DI) water) was added to 1 g of the pre-mixed PDMS to be a desired BCG concentration of 5 mM. The BCG-PDMS mixture was homogenized by a magnetic stirrer. Bubbles formed during the mixing process were removed using a vacuum pump. The BCG-PDMS mixture of 0.5 g was transferred onto the cured bottom PDMS layer on the Petri dish and spread out at 3000 rpm for 15 s using a spin coater. The same curing process as described above was introduced to form a BCG-PDMS/PDMS layer. Finally, the top layer of PDMS was fabricated on the BCG-PDMS/PDMS layer through the spin-casting of a pre-mixed PDMS of 0.5 g. The spin-casting and curing conditions were identical to the process employed for the fabrication of the bottom PDMS layer. The fully cured PDMS/BCG-PDMS/PDMS film was cut into 1 cm × 1 cm pieces and used for the colorimetric detection of urease activity.

Preparation of Urea Base Agar Media
Urea base (0.5 g peptone (Sigma-Aldrich), 0.5 g dextrose (Sigma-Aldrich), 2.5 g sodium chloride (Samchun Chemicals Co. Ltd., Seoul, Korea), 1 g potassium phosphate monobasic (Sigma-Aldrich), and 10 g urea (Sigma-Aldrich)) was dissolved in 50 mL of DI water, and the resulting mixture was filtered using a syringe filter (0.45 µm, Whatman, Maidstone, UK). Additionally, a mass of 10 g of agar (BD, Franklin Lakes, NJ, USA) was dissolved in 450 mL of DI water, and the resulting solution was autoclaved (MDM-60ST, MDM, Suwon-si, Gyeonggi-do, Korea) at 121 • C for 15 min and then allowed to cool down to 50-55 • C. At this point, the urea base solution was added to the agar solution and this composition was gently mixed. Six milliliters (6 mL) of the resulting liquid mixture were added into respective 20-mL glass vials and, at ambient temperature, allowed to solidify for 40 min. The vials were then closed using screw caps and stored at 4 • C prior to use.

Detection of Urease Enzyme Activity
For each experiment, a piece of the above-described colorimetric film sensor was attached to the inside of the 20-mL glass vial, specifically with the bottom of the piece of the sensor~0.5 cm above the top of the urea base agar medium (Figure 1c). The enzyme urease (Type III, powder, Sigma-Aldrich) was dissolved in DI water, and 10 µL of this solution was loaded onto the surface of a urea medium placed in a vial as described above. The final amounts of added urease were 0.25, 0.28, 0.30, 0.40, 0.50, 0.60, and 0.70 U. A volume of 10 µL of DI water was added into another vial as a negative control (marked as 0.00 U in Figure 2). The vials were tightly capped and incubated (VS-8480MX-04-DT, Visionbionex, Bucheon-si, Gyeonggi-do, Korea) at 30 • C for 4 h. After a given incubation, the sensors were photographed and subsequently detached from the vials and transferred to cuvettes, from which absorbance spectra of the sensors were acquired using a UV spectrophotometer (UV-1800, Shimadzu, Kyoto, Japan) at a wavelength range of 400-800 nm. For each spectrum, the ratio of the absorbance at 640 nm to that at 460 nm (A 640 /A 460 ) was used to determine the amount of urease. The experiments were performed in triplicate. to 50-55 °C. At this point, the urea base solution was added to the agar solution and this composition was gently mixed. Six milliliters (6 mL) of the resulting liquid mixture were added into respective 20-mL glass vials and, at ambient temperature, allowed to solidify for 40 min. The vials were then closed using screw caps and stored at 4 °C prior to use.

Detection of Urease Enzyme Activity
For each experiment, a piece of the above-described colorimetric film sensor was attached to the inside of the 20-mL glass vial, specifically with the bottom of the piece of the sensor ~0.5 cm above the top of the urea base agar medium (Figure 1c). The enzyme urease (Type III, powder, Sigma-Aldrich) was dissolved in DI water, and 10 μL of this solution was loaded onto the surface of a urea medium placed in a vial as described above. The final amounts of added urease were 0.25, 0.28, 0.30, 0.40, 0.50, 0.60, and 0.70 U. A volume of 10 µL of DI water was added into another vial as a negative control (marked as 0.00 U in Figure 2). The vials were tightly capped and incubated (VS-8480MX-04-DT, Visionbionex, Bucheon-si, Gyeonggi-do, Korea) at 30 °C for 4 h. After a given incubation, the sensors were photographed and subsequently detached from the vials and transferred to cuvettes, from which absorbance spectra of the sensors were acquired using a UV spectrophotometer (UV-1800, Shimadzu, Kyoto, Japan) at a wavelength range of 400-800 nm. For each spectrum, the ratio of the absorbance at 640 nm to that at 460 nm (A640/A460) was used to determine the amount of urease. The experiments were performed in triplicate. /A460 values of sensors exposed to various units of urease. The dashed line depicts the A640/A460 value of the sensor not exposed to urease (0.00 U, negative control). Symbols and error bars indicate the mean and standard deviations of biological triplicates. Insets show photographs of the sensors exposed to the various units of urease.

Colorimetric Detection of Ureolytic Bacteria using the Produced Sensor
Escherichia coli (E. coli) was purchased from the Korean Collection for Type Cultures (https://kctc.kribb.re.kr/, (accessed on 7 October 2022), Jeongeup-si, Jeollabuk-do, Korea, Table 1). Twelve different bacterial strains were obtained from Nakdonggang National Institute of Biological Resources (https://fbp.nnibr.re.kr/fbcc/, (accessed on 7 October 2022), Sangju-si, Gyeongsangbuk-do, Korea, Table 1). All of the bacteria were cultured in a nutrient broth (MBcell, KisanBio Co., Seoul, Korea) at 30 °C with shaking at 180 rpm (VS-8480MX-04-DT, Visionbionex) for 24 h. In order to collect cells, the culture broth was centrifuged at 5000 rpm (Avanti j-e, Beckman Coulter, Brea, CA, USA) for 10 min and resuspended with fresh nutrient broth to a cell concentration of 1.5 ± 0.4 × 10 8 − 2.2 ± 0.2 × 10 9 colony-forming units per milliliter (CFU/mL) ( Table 1). It is noting that the pH of the resuspended culture broth of bacteria was in a range of 7.37-7.45. Ten microliters (10 μL) of each culture broth were separately added into the vials containing a sensor and urea agar medium. The vials containing the inoculated media were incubated at 30 °C in static

Colorimetric Detection of Ureolytic Bacteria Using the Produced Sensor
Escherichia coli (E. coli) was purchased from the Korean Collection for Type Cultures (https://kctc.kribb.re.kr/, (accessed on 7 October 2022), Jeongeup-si, Jeollabuk-do, Korea, Table 1). Twelve different bacterial strains were obtained from Nakdonggang National Institute of Biological Resources (https://fbp.nnibr.re.kr/fbcc/, (accessed on 7 October 2022), Sangju-si, Gyeongsangbuk-do, Korea, Table 1). All of the bacteria were cultured in a nutrient broth (MBcell, KisanBio Co., Seoul, Korea) at 30 • C with shaking at 180 rpm (VS-8480MX-04-DT, Visionbionex) for 24 h. In order to collect cells, the culture broth was centrifuged at 5000 rpm (Avanti j-e, Beckman Coulter, Brea, CA, USA) for 10 min and resuspended with fresh nutrient broth to a cell concentration of 1.5 ± 0.4 × 10 8 − 2.2 ± 0.2 × 10 9 colony-forming units per milliliter (CFU/mL) ( Table 1). It is noting that the pH of the resuspended culture broth of bacteria was in a range of 7.37-7.45. Ten microliters (10 µL) of each culture broth were separately added into the vials containing a sensor and urea agar medium. The vials containing the inoculated media were incubated at 30 • C in static conditions. The sensors were periodically sampled during the incubation. The sampled sensors were photographed, and their absorbance spectra were acquired using UV-Vis spectrometry as described above. The time it took for the ammonia-producing reaction to occur was determined based on how long it took for the sensor to turn blue as determined visually. The experiments were carried out in triplicate.

Verification of the Performance of the Sensor
The preculture of Klebsiella pneumoniae (K. pneumoniae) broth was collected by subjecting the broth to centrifugation at 5000 rpm for 10 min and resuspended with fresh nutrient broth to produce a cell concentration of 4.2 ± 2.0 × 10 8 CFU/mL. A volume of ninety microliters (90 µL) of K. pneumoniae culture broth was mixed with DI water to produce a total volume of 100 µL. Ten microliters (10 µL) of the culture solution (referred to as non-treated cells) were added into vials containing film sensors and urea agar media. At the same time, a volume of 90 µL of K. pneumoniae culture was treated with 10 µL of a solution of 0.1 mM phenyl phosphorodiamidate (PPDA, Alfa Aesar, Ward Hill, MA, USA) in DI water. Ten microliters (10 µL) of the PPDA-treated K. pneumoniae cells (referred to as PPDA-treated cells) were separately added into other vials containing sensors and urea agar media. The vials were subjected to static incubation at 30 • C. The sensors were analyzed at incubation durations of 0.0, 2.0, 5.0, 6.0, 7.5, 8.0, 10.0, and 11.0 h; specifically, they were photographed and transferred to cuvettes, from which their absorbance spectra were acquired in the wavelength range of 400-800 nm. All of the experiments were performed in triplicate.

Reusability of the Sensor
For each experiment, 10 µL of Proteus terrae (P. terrae) culture with a cell concentration of 1.4 ± 0.2 × 10 9 CFU/mL were added to a vial containing the colorimetric film sensor and urea agar medium. The vial was then incubated at 30 • C for 4 h, after which the sensor was photographed and its A 640 /A 460 value was measured to determine the activity of urease. After these measurements, the sensor was aerated for 2 d at ambient temperature and its absorbances at 460 and 640 nm were measured again to determine the ability to reuse the sensor. The aerated sensor was reused for the detection of urease activity of P. terrae cells in the same manner as described above. The sensor was used five times. The experiments were performed in triplicate.

Design of the Colorimetric Film Sensor for the Detection of Ureolytic Microbes
A colorimetric film sensor was employed for the detection of ureolytic activity in microbes ( Figure 1). The sensor consisted of three functional layers, all containing flexible PDMS elastomer (Figure 1a). In the middle layer of the sensor, a BCG indicator was incorporated into the PDMS elastomer. Note that BCG is a pH indicator that exhibits a significant colorimetric response, from yellow to blue, according to the pH level: at pH levels between 3 and 4, BCG exhibits a yellow color; as the pH is increased to 5 to 8, the BCG is deprotonated by basic substances, resulting in a change in its structure from a monoanionic form to a dianionic form, and this change in structure leads to the change in color from yellow to blue [22]. This designed sensing layer (i.e., the BCG-PDMS layer), with a thickness of~60 µm, was embedded between the top and bottom of the hydrophobic thin layers of PDMS each with a thickness of~20 µm. Gaseous basic molecules can pass through PDMS [23][24][25]. On the other hand, water cannot penetrate the PDMS of the sensor, and thus false-positive results caused by alkaline solutions are minimized. Our previous study demonstrated that the sensor was able to selectively detect gaseous ammonia over various other basic substances (specifically, NaOH, KOH, and Ba(OH) 2 ) [22].
Urease catalyzes the hydrolysis of urea to ammonia and carbon dioxide (Figure 1b). Therefore, the amount of ammonia increased during the enzymatic hydrolysis of urea by ureolytic (i.e., urease-positive (+)) microbes and, accordingly, the color of the sensor changed from yellow to blue (Figure 1c). On the other hand, the color of the film did not change if the microbes do not have urease activity (i.e., are urease-negative (−)) ( Figure 1c). In this regard, ureolytic microbes were simply detected using our sensor and the colorimetric response was easily identified by the naked eye.
This change in the color of the sensor can be reversed upon its exposure to the fume of acetic acid or aeration [22]. As a result, the original yellow color of the film was regenerated (Figure 1d). The reversibility of the change in color of the film sensor makes it reusable, which reduces detection expenses. In addition, access to skilled technicians and expensive instruments is unnecessary for this detection of ureolytic microbes. The simplicity and reusability of the film sensor makes it affordable.

Validation of the Sensor for the Detection of Urease
The performance of the colorimetric film sensor was validated using a commercially available urease enzyme (Figure 2). The absorbance spectrum of the sensor changed upon the addition of urease (Figure 2a). Specifically, as the amount of urease was increased from 0.25 to 0.70 U, the absorbance at 460 nm (A 460 ) decreased and the absorbance at 640 nm (A 640 ) concomitantly increased. The A 640 /A 460 value was found to be linearly proportional to the amount of urease, with the regression equation y = 1.18 x + 0.26 and r 2 = 0.97 (Figure 2b). The color of the sensor changed from yellow to blue upon it being exposed to the products of the reaction catalyzed by urease and the blue color became distinct in a manner dependent on the amount of urease. The development of blue color by the sensor was clearly identified by the naked eye. The results demonstrated that our pH-sensitive sensor can be used to visually detect urease activity by showing changes in its color (i.e., from yellow to blue) as well as in its absorbance spectrum upon the urease-catalyzed hydrolysis of urea [6].

Colorimetric Detection of Ureolytic Microbes using the Sensor
Urease-producing microbes were detected using the colorimetric film sensor. Of the thirteen tested bacterial strains, nine of them showed urease activity and different reaction times ( Table 1). The sample of the bacterium K. pneumoniae was indicated to be positive for urease, according to changes of the absorbance spectrum of the film sensor as the incubation duration was increased (Figure 3a). Specifically, the sensor exhibited a decrease in absorbance at a wavelength of 460 nm and an increase at 640 nm, which corresponded to the results for the detection of commercially available urease enzymes, as shown in Figure 2a. The color of the sensor began to change from yellow to blue at 6 h of incubation, with the change attributed to the deprotonation of BCG in PDMS, resulting from the hydrolysis of urea by the urease-producing microbes. Accordingly, the A 640 /A 460 value of the sensor also increased with increasing incubation duration for durations of 6 h and greater (Figure 3b). The change to the blue color became clearly apparent to the naked eye after an incubation of 11 h. available urease enzyme (Figure 2). The absorbance spectrum of the sensor changed upon the addition of urease (Figure 2a). Specifically, as the amount of urease was increased from 0.25 to 0.70 U, the absorbance at 460 nm (A460) decreased and the absorbance at 640 nm (A640) concomitantly increased. The A640/A460 value was found to be linearly proportional to the amount of urease, with the regression equation y = 1.18 x + 0.26 and r 2 = 0.97 ( Figure  2b). The color of the sensor changed from yellow to blue upon it being exposed to the products of the reaction catalyzed by urease and the blue color became distinct in a manner dependent on the amount of urease. The development of blue color by the sensor was clearly identified by the naked eye. The results demonstrated that our pH-sensitive sensor can be used to visually detect urease activity by showing changes in its color (i.e., from yellow to blue) as well as in its absorbance spectrum upon the urease-catalyzed hydrolysis of urea [6].

Colorimetric Detection of Ureolytic Microbes using the Sensor
Urease-producing microbes were detected using the colorimetric film sensor. Of the thirteen tested bacterial strains, nine of them showed urease activity and different reaction times ( Table 1). The sample of the bacterium K. pneumoniae was indicated to be positive for urease, according to changes of the absorbance spectrum of the film sensor as the incubation duration was increased (Figure 3a). Specifically, the sensor exhibited a decrease in absorbance at a wavelength of 460 nm and an increase at 640 nm, which corresponded to the results for the detection of commercially available urease enzymes, as shown in Figure 2a. The color of the sensor began to change from yellow to blue at 6 h of incubation, with the change attributed to the deprotonation of BCG in PDMS, resulting from the hydrolysis of urea by the urease-producing microbes. Accordingly, the A640/A460 value of the sensor also increased with increasing incubation duration for durations of 6 h and greater ( Figure 3b). The change to the blue color became clearly apparent to the naked eye after an incubation of 11 h.  (Table 1, Figures  4a-c, S1a,b). In each of these cases, the A640/A460 value of the sensor significantly increased after an incubation duration of ~24 h. The bacterium P. terrae showed strongly positive urease activity, as the color of the sensor exposed to this bacterium rapidly changed to  (Table 1, Figures 4a-c and S1a,b). In each of these cases, the A 640 /A 460 value of the sensor significantly increased after an incubation duration of~24 h. The bacterium P. terrae showed strongly positive urease activity, as the color of the sensor exposed to this bacterium rapidly changed to blue within 3 h, and this change was clearly identified by the naked eye ( Figure 4d). Meanwhile, the strains Enterobacter hormaechei (E. hormaechei) and Enterobacter roggenkampii (E. roggenkampii) showed weakly positive urease activities (Table 1, Figure 4e,f). In each of these cases, noticeable changes in color and A 640 /A 460 of the sensor occurred only after about 3 days of reaction. When each of the bacterial strains C. koseri, E. coli, K. michiganensis, and P. alcalifaciens were tested, the sensor only marginally changed color after an extended reaction duration of 7 d (Table 1 and Figure S1c-f), indicating these four strains to be negative for urease. while, the strains Enterobacter hormaechei (E. hormaechei) and Enterobacter roggenkampii (E. roggenkampii) showed weakly positive urease activities (Table 1, Figure 4e,f). In each of these cases, noticeable changes in color and A640/A460 of the sensor occurred only after about 3 days of reaction. When each of the bacterial strains C. koseri, E. coli, K. michiganensis, and P. alcalifaciens were tested, the sensor only marginally changed color after an extended reaction duration of 7 d (Table 1 and Figure S1c-f), indicating these four strains to be negative for urease. Ureolytic activity has been described as a virulence factor for several bacteria and an emerging pathogenic factor during fungal infection [1,7,8]. The Proteus and Klebsiella species cause bacterial urinary tract infections in association with urease-dependent processes [7]. Urease-positive bacteria can result in the formation of infection stones and gas- Ureolytic activity has been described as a virulence factor for several bacteria and an emerging pathogenic factor during fungal infection [1,7,8]. The Proteus and Klebsiella species cause bacterial urinary tract infections in association with urease-dependent processes [7]. Urease-positive bacteria can result in the formation of infection stones and gastrointestinal colonization [26][27][28][29]. Infection stones surround the pathogens, thus protecting them. Moreover, ureolytic activity in human pathogens is thought to be related to the infectivity or persistence of the microbes [8].

Verification of the Detection of Urease Activity by the Sensor
The colorimetric response of the film sensor to the ureolytic bacteria was verified using the urease inhibitor PPDA ( Figure 5). The cells of the ureolytic bacterium K. pneumoniae not treated with PPDA (referred to non-treated cells) induced a change in the color of the sensor from yellow to blue within 10 h and achieved an increase in the A 640 /A 460 value of the sensor as the reaction duration was increased (black bars of Figure 5). On the other hand, the sensor exposed to PPDA-treated cells (referred to K. pneumoniae + PPDA) showed neither any change in color nor in its A 640 /A 460 value during 12 h of reaction (red bars of Figure 5). This lack of any significant colorimetric change was attributed to PPDA having specifically bound to the active site of urease, thus preventing not only the binding of urea to urease, but also the hydrolysis of urea to ammonia [4,6]. Based on these results, the colorimetric and absorbance spectral changes of the sensor exposed to non-treated cells were attributed to the reaction catalyzed by urease enzymes produced by Klebsiella pneumoniae cells.

Verification of the Detection of Urease Activity by the Sensor
The colorimetric response of the film sensor to the ureolytic bacteria was verified using the urease inhibitor PPDA ( Figure 5). The cells of the ureolytic bacterium K. pneumoniae not treated with PPDA (referred to non-treated cells) induced a change in the color of the sensor from yellow to blue within 10 h and achieved an increase in the A640/A460 value of the sensor as the reaction duration was increased (black bars of Figure 5). On the other hand, the sensor exposed to PPDA-treated cells (referred to K. pneumoniae + PPDA) showed neither any change in color nor in its A640/A460 value during 12 h of reaction (red bars of Figure 5). This lack of any significant colorimetric change was attributed to PPDA having specifically bound to the active site of urease, thus preventing not only the binding of urea to urease, but also the hydrolysis of urea to ammonia [4,6]. Based on these results, the colorimetric and absorbance spectral changes of the sensor exposed to non-treated cells were attributed to the reaction catalyzed by urease enzymes produced by Klebsiella pneumoniae cells.   [40]. Instead of directly measuring bacterial urease activity, those studies used urease as a signal translation agent that catalytically produces ammonium carbonate, which could elevate the pH of the solution. The pH change was monitored by taking voltammetric measurements of the product of the catalyzed reaction (ammonium carbonate) or a pH-responsive chromogenic dye. Werkmeister et al. (2016) reported the use of organic field-effect transistors (OFETs) to sense ammonia for the purpose of detecting the enzymatically catalyzed breakdown of urea [41]. Due to the urease-catalyzed reaction, the OFETs responded to the millimolar levels of urea, but this feature did not extend to the detection of urease-producing microbes. According to our results, the developed colorimetric film sensor showed reliable performance for the detection of urease activity in various bacterial species.

Selective Detection of Urease-Producing Microbes in the Presence of Non-Ureolytic Bacteria
The selectivity of the colorimetric film sensor for the detection of urease-producing microbes was validated using the ureolytic bacterium strain K. pneumoniae. An inoculum of only K. pneumoniae changed both the color and A 640 /A 460 value of the sensor ( Figure 6). That is, the sensor turned blue upon being exposed to an incubation of only K. pneumoniae cells (B in Figure 6) and comparable responses also occurred when K. pneumoniae cells were incubated with non-ureolytic bacteria (C-G in Figure 6). The presence of only non-ureolytic bacteria cells did not induce a change in the color of the sensor (A in Figure 6).
However, it was incapable of achieving a selective detection of pathogenic bacteria [38]. Singh et al. (2019) employed silver-urease interactions for the colorimetric detection of pathogens [39]. Hou et al. (2020) used a 3D magnetic grid and urease catalysis for the rapid detection of Salmonella [40]. Instead of directly measuring bacterial urease activity, those studies used urease as a signal translation agent that catalytically produces ammonium carbonate, which could elevate the pH of the solution. The pH change was monitored by taking voltammetric measurements of the product of the catalyzed reaction (ammonium carbonate) or a pH-responsive chromogenic dye. Werkmeister et al. (2016) reported the use of organic field-effect transistors (OFETs) to sense ammonia for the purpose of detecting the enzymatically catalyzed breakdown of urea [41]. Due to the urease-catalyzed reaction, the OFETs responded to the millimolar levels of urea, but this feature did not extend to the detection of urease-producing microbes. According to our results, the developed colorimetric film sensor showed reliable performance for the detection of urease activity in various bacterial species.

Selective Detection of Urease-Producing Microbes in the Presence of Non-Ureolytic Bacteria
The selectivity of the colorimetric film sensor for the detection of urease-producing microbes was validated using the ureolytic bacterium strain K. pneumoniae. An inoculum of only K. pneumoniae changed both the color and A640/A460 value of the sensor ( Figure 6). That is, the sensor turned blue upon being exposed to an incubation of only K. pneumoniae cells (B in Figure 6) and comparable responses also occurred when K. pneumoniae cells were incubated with non-ureolytic bacteria (C-G in Figure 6). The presence of only nonureolytic bacteria cells did not induce a change in the color of the sensor (A in Figure 6).  The photographic results were consistent with measurements of A 640 /A 460 of the sensor. The A 640 /A 460 value of the sensor increased only in the presence of K. pneumoniae cells (samples B-G in Figure 6) compared to that in the presence of only non-ureolytic bacterial cells (sample A in Figure 6), and these differences were statistically significant (p < 0.05, as determined from an ANOVA analysis). The non-ureolytic bacteria did not affect the A 640 /A 460 value of the film sensor. That is, the A 640 /A 460 values of the sensors exposed to K. pneumoniae cells with non-ureolytic bacteria (bars C-G of Figure 6) were statistically similar (p > 0.05) to that of the sensor exposed to only K. pneumoniae cells (sample B of Figure 6). As expected, the presence of only non-ureolytic bacterial cells induced just a marginal change in the A 640 /A 460 value of the sensor (A in Figure 6).

Reusability of the Sensor
The color of a used sensor reverted from blue to its original yellow when subjected to aeration for 2 d (Figure 7). The sensor was found to be reusable for the detection of urease in P. terrae cells. Photographs and absorbance levels at 460 and 640 nm of the sensors were obtained between the repeated uses. The color of the sensor changed from yellow to blue upon the production of urease by P. terrae cells and the color change responses between the repeated uses were similar. Similarly, the A 640 /A 460 value of the sensor increased after 4 h of incubation to between 1.17 ± 0.05 and 1.35 ± 0.08 between the repeated uses. The used sensor returned to its original state after each of five consecutive deployments.
determined from an ANOVA analysis). The non-ureolytic bacteria did not affect the A640/A460 value of the film sensor. That is, the A640/A460 values of the sensors exposed to K. pneumoniae cells with non-ureolytic bacteria (bars C-G of Figure 6) were statistically similar (p > 0.05) to that of the sensor exposed to only K. pneumoniae cells (sample B of Figure  6). As expected, the presence of only non-ureolytic bacterial cells induced just a marginal change in the A640/A460 value of the sensor (A in Figure 6).

Reusability of the Sensor
The color of a used sensor reverted from blue to its original yellow when subjected to aeration for 2 d (Figure 7). The sensor was found to be reusable for the detection of urease in P. terrae cells. Photographs and absorbance levels at 460 and 640 nm of the sensors were obtained between the repeated uses. The color of the sensor changed from yellow to blue upon the production of urease by P. terrae cells and the color change responses between the repeated uses were similar. Similarly, the A640/A460 value of the sensor increased after 4 h of incubation to between 1.17 ± 0.05 and 1.35 ± 0.08 between the repeated uses. The used sensor returned to its original state after each of five consecutive deployments. The reusability of the sensor was attributable to the aeration having promoted the release of ammonia from the sensor (Figure 1d). The permeability of the PDMS layer allowed for the transport of ammonia gas into and out of the sensor, and thus the sensor was made reusable through simple aeration. Such reusability reduces fabrication and operation expenses, helping to make our sensor affordable.

Conclusions
We designed a colorimetric film sensor that could detect urease activity in microbes. Microbial urease activity was shown to induce a change in the color of the sensor from yellow to blue and this change in color can be simply identified by the naked eye. As a result of this color change property, the sensor can be used as an indicator of the presence of ureolytic microbes in a sample. The blue color development of the sensor allowed us to The reusability of the sensor was attributable to the aeration having promoted the release of ammonia from the sensor (Figure 1d). The permeability of the PDMS layer allowed for the transport of ammonia gas into and out of the sensor, and thus the sensor was made reusable through simple aeration. Such reusability reduces fabrication and operation expenses, helping to make our sensor affordable.

Conclusions
We designed a colorimetric film sensor that could detect urease activity in microbes. Microbial urease activity was shown to induce a change in the color of the sensor from yellow to blue and this change in color can be simply identified by the naked eye. As a result of this color change property, the sensor can be used as an indicator of the presence of ureolytic microbes in a sample. The blue color development of the sensor allowed us to classify ureolytic microbes into strongly, moderately, and weakly positive urease activities by comparing the incubation times of thirteen different microbes. The sensor was found to be easy to use and to be selective and reusable for the detection of ureolytic microbes. We also expect its use to be extended to other applications, such as the monitoring of medical and environmental samples.

Supplementary Materials:
The following supporting information can be downloaded at: https:// www.mdpi.com/article/10.3390/bios12100886/s1, Figure S1: (a-f) Colorimetric detection of ureolytic bacteria using the designed film sensor. A640/A460 values of sensors exposed to indicated bacteria plotted against incubation duration. Insets show photographs of the sensors. Symbols and error bars represent the mean and standard deviations of biological triplicates.