Urease is a metalloenzyme that catalyzes the hydrolysis of urea to yield ammonia and carbon dioxide. It is found in a wide variety of organisms including plants, fungi and bacteria [57]. Stable hollow polyelectrolyte capsules containing urease (Mw 480 kDa) were produced by means of the layer‐by‐layer assembling of PAH and PSS on melamine formaldehyde microcores followed by the core decomposition at low pH [58]. The authors introduced a new method of loading urease enzyme into the particles depending on a finding that capsules were non‐permeable for urease in water and became permeable in a water/ethanol mixture. As shown in Figure 8 (left), confocal microscopy imaging illustrates that FITC‐labelled urease was excluded from the polyelectrolyte shells. The interior of the capsules remained dark and outside the capsule the background containing FITC‐urease was fluorescent. This observation indicated the closed state of the capsules.
Figure 8 (center) shows a confocal fluorescence image of labelled urease with capsules after addition of ethanol (1:1 water/ethanol mixture). In this case, the fluorescence signal localized in the interior of the capsule has the same intensity as the outside background fluorescence. This indicated the penetration of the enzyme into the capsules and requires an open state of the capsule wall. When the capsules were transferred back into water, the polyion shells became closed, and the urease was captured inside, as illustrated in Figure 8 (right). The interior of the capsule is bright and constant over time, and there is no fluorescence signal from the solution.
Thus, urease filled the capsules. The images did not change with time, indicating that urease is
preserved inside the capsules. The authors mentioned that the mechanism of reversible permeability changes in the polyion multilayers is not fully understood. It may be related to the segregation of the polyion network in water/ethanol media. Such segregation might lead to defects in the shell, and pores forming might be large enough for 5‐nm diameter urease globules to penetrate the wall. Returning capsules into pure water causes a relaxation of the polyion walls to a closed structure [58].
Figure 8: Permeation and encapsulation of urease‐FITC into polyion multilayer capsules. Left, in water;
middle, in water/ethanol mixture 1:1; right, the capsule with encapsulated urease again in the water. Top, scheme; bottom, confocal fluorescence images of the capsules [58] “Reprinted with permission from reference 58. Copyright 2001 American Chemical Society.”
A colorimetric assay based on the hydrolysis of urea was used to investigate the activity of free and immobilized urease [59]. The increase in solution pH due to ammonia production during the enzymatic reaction was monitored by the pH sensitive dye, bromcresol purple. The absorbance of this dye at 588 nm increases linearly with pH in the range of 5.8 to 7.5. Figure 9 gives a comparison of the catalytic activity of the urease loaded into the capsules and free urease. Urease encapsulated inside the LbL shell preserved 13% of its activity as compared to free enzyme. This decrease is mainly due to low substrate diffusion into the capsules. The urease activity inside the capsules was also stable as compared to free urease: after 5 days of storage at 7 °C, encapsulated urease completely preserved its activity while free urease kept under the same conditions in aqueous solution lost 45% of its activity. The polyelectrolyte shell protects encapsulated enzymes from proteases and microbes, and provides better stability [58].
Figure 9: Absorbance at 588 nm by 2.9 mL of enzymatic activity assay solution after addition of 0.1 mL of urease‐loaded microcapsule solution and the same test for addition of 0.05 mL of 0.02 mg/mL free urease [58]. “Reprinted with permission from reference 58. Copyright 2001 American Chemical Society."
In a different LbL design [60], urease was loaded on the surface of submicron‐sized polystyrene particles. Urease multilayers were assembled with alternating oppositely charged polyelectrolytes in a predetermined order, utilizing electrostatic interactions for layer growth. Urease, which has a pI at pH 5 and is stable and active between pH 4 and 8.5 [61, 62], was employed in the LbL assembly either as a negative polyelectrolyte at pH 8 deposited in alternation with polycations (PEI or PDDA), or as a positive polyelectrolyte at pH 4.5 and consecutively deposited with the polyanion (PSS). As an added feature, prior to enzyme adsorption, the colloid particles were coated with an additional layer of silica or magnetite nanoparticles in order to enhance their total surface area and promote further enzyme deposition, to give shell architectures of the following sequence:
[PDDA/PSS/PDDA/40‐nm silica/PDDA/(urease/PDDA)1‐4] or [PDDA/PSS/PDDA/12‐nm magnetite/
PDDA/(urease/PDDA)1‐4].
Urease multilayers were first constructed on quartz cell microbalance (QCM) electrodes in order to establish the conditions for suitable multilayer growth. The QCM frequency shift, caused by the deposition of material on the electrode surface, can be related to the adsorbed mass and layer thickness of the material via the Sauerbrey equation [63]. Figure 10 shows the QCM results for the
construction of a PDDA/PSS/PDDA/(urease/PDDA)5 multilayer film on a QCM electrode. The first three polyelectrolyte layers serve as a precursor film to provide a uniform charge and a smooth surface for subsequent urease deposition. A regular stepwise decrease in the QCM frequency was observed (Figure 10) [60].
Figure 10: QCM monitoring (frequency change vs adsorption steps) of urease/PDDA assembly:
[PDDA/PSS/PDDA /(urease/PDDA)5] [60]. “Reprinted with permission from reference 60. Copyright 2001 American Chemical Society.”
The conditions established for the successful assembly of urease multilayers on the planar QCM substrates were subsequently employed to form enzyme multilayer shells on microparticle templates (470 nm PS spheres). The precursor film (PDDA/PSS/PDDA) with an additional outermost silica or magnetite nanoparticle layer provided a better surface for the formation of stable urease multilayer shells. Attempts to deposit urease onto PDDA/PSS/PDDA‐modified PS particles yielded a low enzyme amount in the shells. This finding was attributed to the fact that weakly attached enzyme layers can be removed from the substrate surface by the next incoming polyion via the formation of water‐soluble polyelectrolyte‐enzyme complexes, as was found for histone/DNA interactions [64]. In addition, improved stability with respect to adsorption of glucose oxidase multilayers deposited onto gold nanoparticle layers, compared with glucose oxidase deposited on a less rough substrate, was observed. Hence, a primer nanoparticle layer was deposited on the PS spheres to increase the surface roughness and improve the adsorption stability of urease. The growth of the urease multilayers on the PS particles was followed by microelectrophoresis, where
the zeta potential of the coated latex particles alternates between negative and positive values, corresponding to the sequential adsorption of cationic and anionic species, respectively [60].
The catalytic activity of encapsulated and free urease was measured by monitoring the change in absorbance of the pH‐sensitive dye bromcresol purple, corresponding to the increase in pH due to the hydrolysis of urea giving ammonium hydroxide. The catalytic activity was found to increase with the increase in the number of urease layers deposited on the particles. Both urease/PSS multilayers and urease/PEI multilayers yielded very low catalytic activities (~100 times lower) compared to the urease/PDDA multilayers despite being assembled in a similar fashion (according to the shell architectures).This may be due to the differences in compactness of the multilayers resulting from different polymer conformations on the surface (e.g., PDDA is a linear polycation and PEI is a branched polycation), different substrate diffusion rates and different degrees of blocking active sites of the enzyme. Compared to free urease, investigations revealed that the activity of immobilized urease (in a triple‐layer shell) was 25% of that of free enzyme. This is a reasonable decrease because of substrate diffusion limitations and difficulties in reaching the active centers of immobilized urease. Another characteristic feature of the urease catalytic reaction is a 10‐minutes dead time, during which no product was detected. A similar dead time was observed for low concentrations of free urease and, probably, is connected to accumulation of the reaction product [60]. Similar LbL design and results for urease were also reported by Wang et al. [65].
Moreover, adding a magnetic functionality to the particles was investigated. 12‐nm‐diameter Fe3O4 nanoparticles were deposited in the shell architecture to supply the particles with a magnetic function. The shell sequence was [PDDA/PSS/PDDA/12‐nm Fe3O4/PDDA/(urease/PDDA)1‐4]. The Fe3O4 nanoparticle distribution on the surface of the latex spheres was found to be less uniform than the silica shell and more than monolayer coverage. Nevertheless, the absolute enzymatic activity of the magnetic catalytic particles was similar to that of the corresponding particles with a layer of 40‐nm silica. In addition, on approaching a 0.3 tesla permanent magnet to the tube wall resulted in the collection of all of the modified latex spheres (on a wall region closest to the magnet) in ~30 seconds. Immersing the permanent magnet into the solution containing the magnetic/urease‐coated particles resulted in their collection on the magnet. This added magnetic function is particularly useful in applications where separation and reuse of such particles is required [60].