Enzyme Stability

The oxygen reactivity of PMOs was measured by a time resolved quantification of H₂O₂ formation in 96-well plates (total volume of 200 μL) using a Perkin Elmer EnSpire Multimode plate reader. All reactions were performed in 100 mM sodium phosphate buffer, pH 6.0 at 22°C. Based on preliminary studies ascorbate and CDH were used in concentrations of 30 μM and 0.3 μM (0.025 mg mL^-1), respectively to prevent a limitation in the PMO reduction step. As electron donor for CDH 500 μM cellobiose was used. When ascorbate was used as reductant, it was added to a final concentration of 30 μM and enzyme assays were started by mixing 20 μL of the respective PMO with 180 μL of the ready-made assay solution containing 30 μM ascorbate, 50 μM Amplex Red and 7.14 U mL^-1 peroxidase in 96-well plate wells. In reference experiments without PMO the background signal (H₂O₂ production by CDH) was measured and subtracted from the assays. When CDH was used as reductant, the PMO assays were started by mixing 20 μL of sample solution and 20 μL CDH solution with 160 μL of the reaction mix containing cellobiose, Amplex Red and peroxidase. Initial fluorescence scans of resorufin showed highest signal intensities and lowest interference with matrix compounds when using an excitation wavelength of 569 nm and an emission wavelength of 585 nm for the selected conditions. The stoichiometry of H₂O₂ conversion to resorufin formation is 1:1. By using a high concentration of Amplex Red (50 μM) the linearity of the detection reaction was ensured and the molar ratio of Amplex Red:H₂O₂ was always greater than 50:1
[22 (link)]. The H₂O₂ concentration in the assays was far below 1 μM, which leads to a linear concentration/activity response of horseradish peroxidase, which has a K_M for H₂O₂ of 1.55 μM. The high final activity of horseradish peroxidase (7.14 U mL^-1) assures a fast conversion of the formed H₂O₂ and prevents the final reaction to be rate limiting. Additionally, it prevents the accumulation of H₂O₂, which could have detrimental effects on enzyme stability in the assay. The stability of resorufin fluorescence under these conditions was tested by addition of varying concentrations of hydrogen peroxide (0.1 – 5 μM) to the assay. A stable signal that remained constant throughout the measured period of 45 minutes was observed and maximum signal intensity was reached already during the mixing period before starting the assay. A linear relation between fluorescence and H₂O₂ concentrations in the range of 0.1 – 2 μM H₂O₂ was observed and the slope (28450 counts μmol^-1) was used for the calculation of an enzyme factor to convert the fluorimeters readout (counts min^-1), into enzyme activity. PMO activity was defined as one μmol H₂O₂ generated per minute under the defined assay conditions.

For studying the impact of pH on the proteolytic activity of UcB5, the reacting solutions were fixed to a wide range of pHs by three buffers. Citrate–phosphate buffer to achieve a pH range of 3.0 up to 6.0, Tris–HCl buffer for pHs in the range of 7.0 up to 8.0, and glycine–NaOH buffer to adjust pHs in the range of 9.0 up to 13.0. Azocasein at 1% (w/v) concentration was used and incubation of enzyme–substrate reactions was done at 35 °C.
For studying the pH stability of the pure enzyme, it was incubated for 60 min at 35 °C with various pHs in the range of 5.0 up to 13.0 using the prescribed buffers. The remaining enzymatic activity was assessed at pH 8.0. Furthermore, the isoelectric point of the tested protease was determined by overnight incubation of a concentrated preparation of the pure enzyme at pHs in the range of 3.0 up to 11.0 at 4 °C. Protein precipitation was done at the gravitational force of 10,000×g for 15 min. The protein pellets were quantified according to Bradford^{12 (link)} method. The isoelectric point was expressed as the pH degree where maximal protein precipitation has been done^{13 (link)}.

Small unilamellar vesicles (SUVs) containing DOPC and C8-LacCer were prepared as previously described [19 (link)], with modifications. Briefly, lipids in chloroform were mixed at desired molar ratios (100:0, 95:5 and 80:20) at a total amount of 5 µmol and dried under a stream of nitrogen gas followed by drying in a vacuum desiccator for 2 h. The lipid mixtures were resuspended in a working buffer at 2 mg/mL and homogenized by five cycles of freezing, thawing and vortexing. The lipid suspensions were subjected to tip sonication in pulse mode (1 s on/1 s off) for 15 min with refrigeration to obtain SUVs. The SUV suspension was centrifuged at 12,100× g for 10 min to remove titanium debris (shed from the sonicator tip) and stored at 4 °C under nitrogen gas until use. QCM-D sensors were pre-treated in UV/ozone (30 min), placed in the flow chamber and installed in the flow modules of the Q-Sense E4 apparatus. Following the acquisition of a baseline in HBS buffer (10 mM HEPES, 150 mM NaCl, pH 7.4), SLBs were formed on the sensor surface by the flow of 50 μg/mL liposomes until frequency and dissipation were stable. SLBs were equilibrated first in HBS buffer and then in reaction buffer (RB; 20 mM HEPES, 0.5 mM MnCl₂, 4.4 mM Tris, 1.1 mM TCEP, pH 7.4), which was matched in content with the LgtC storage buffer to avoid solution effects on the QCM-D response upon starting the enzyme reaction. The RB buffer contained MnCl₂, which is required for LgtC activity [30 ], and DTT was added to enhance enzyme stability. Enzyme reactions were initiated in RB containing 250 µM UDP-Gal and desired concentrations of LgtC (1.72 μM or 17.2 μM). SLBs were washed with RB to stop the enzyme reaction and then with HBS for further analysis. A 6.5 µM flow of StxB in HBS was applied, and StxB binding was observed to report for Gb3 in the SLBs.