The unique potential of NMR spectroscopy for monitoring simultaneously the concentrations of several metabolites in complex milieux makes it one of the most powerful techniques available to carry out this type of study.
For NMR measurements, lysates or cell-free extracts were resuspended in 0.15 M NaCl constituted in 5:1 H2O/2H2O buffer mixtures to provide deuterium frequency lock for the spectrometer. Substrate concentrations were 40 mM L-aspartate and 50 mM carbamoyl phosphate, in 0.1 M HEPES buffer (pH 8.0). The reaction was started by adding 100 μl of cell free extract to a total sample volume of 600 μl. To allow for efficient dispensing of the assay mixture into the 5 mm narrow bore NMR tube (Wilmad, Buena, NJ), the lysate or extract suspension and the substrates were mixed in an Eppendorf tube; diluting viscous cell lysates helped to place them into the tube.
Free induction decays were collected using a Bruker AM-500 spectrometer, operating in the Fourier transformation mode. Measurements were carried out at 37°C, and sequential spectra were acquired automatically at 500.11 MHz with presaturation of the water resonance. The instrumental parameters were: spectral width 5347 Hz, memory size 8 K, recycling time 3.5 s, number of transients 144, and pulse angle 50° (8 μs). To improve signal-to-noise, exponential filtering of 1 Hz was applied prior to Fourier transformation. Depending on the viscosity of the sample and the concentrations of the substrates, it is often advisable to employ resolution-enhancing window functions at the expense of losing some signal intensity; for example, a Gaussian window function with typical parameters: -1 Hz line broadening, and 0.19 Gaussian parameter.
The time evolution of the utilisation of substrates and appearance of product was followed by acquiring sequential spectra of the reactions. Progress curves were obtained by measuring the integrals of substrate and product resonances at each point in time. Maximal rates were calculated from good fits (correlation coefficients 3 0.99) of the data to straight lines for 30 min of the reactions.
ACTase activity was also determined using a radioactive assay that measures the incorporation of [14C]carbamoyl phosphate into carbamoyl aspartate (8). Typical enzyme assay conditions were 40 mM L-aspartate, 0.4 mM carbamoyl phosphate containing [14C]carbamoyl phosphate (0.1 μCi μmole -1 ), Tris buffer (0.1 M, pH 8.0), and cell-free extract in a final volume of 200 μl. The reaction was initiated by adding 10 μl of cell-free extract, and incubated for 10 min at 37°C in a water bath. Reactions were terminated by adding 100 μl of 3 M formic acid and heating at 80°C for 6 min, during which time any unreacted [14C]carbamoyl phosphate decomposed into phosphate and carbon dioxide, and the label was evaporated as 14 CO2. Heating for 6 minutes was found to be the optimum; less time was insufficient for complete conversion to 14CO2of the unreacted [14C]carbamoyl phosphate, and longer times were unnecessary. Ten milliliters of scintillation fluid (2,5-diphenyloxazole/toluene; 0.5% w/v) were added, and the radioactive decay measured on a Packard Tricarb scintillation counter (Packard Instrument Co., USA).
A colorimetric measurement of carbamoyl aspartate production using a microtitre protocol was employed for kinetic studies of the enzyme reaction (9), and all measurements were performed in triplicate. The method was based on that of Prescott and Jones (10) which uses a monoxime and antipyrine colour reagent for the detection of CAA. Two parts antipyrine (5 g/l in 50% (v/v) sulfuric acid) were mixed with 1 part monoxime (8 g/l in 5% (v/v) acetic acid). The colour reagent was prepared immediately before use. Owing to the instability of carbamoyl phosphate (11), it is essential that solutions are prepared fresh before each reaction. The reaction mixture contained the same concentrations of substrates and buffer as described above but without the [14C]carbamoyl phosphate, in a final volume of 200 μl. To start the enzyme reaction 50 μl of freshly prepared carbamoyl phosphate was added to the wells, and the plate incubated for 10 min at 37°C. One hundred microlitres of the color reagent was then added to each well to stop the reaction, and the plate incubated in the dark in a water bath at 60°C for 2 h. Else and co-workers (9) recommend covering the plate with adhesive film while incubating; however when this was done water droplets would sometimes form on the underside of the film and mix into the wells. To avoid this problem another microtitre plate was used as a cover. The plate was then allowed to cool to room temperature in the dark for 15 minutes, and the absorbance measured at 450 nm.
This method was adapted in the present study to provide an activity stain for ACTase in native PAGE gels. Native proteins were separated by modifying the procedure of Laemmli (12), where gel electrophoresis was performed under non-reducing and non-denaturing conditions. The gel was then sliced into 1mm sections, and each section placed into a microtitre well containing the substrates of the ACTase reaction. An identical lane of electrophoretically separated proteins were also sliced and then incubated without aspartate, as a negative control. The microtitre plate was then processed as before.
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ACTase activity could be localised to the soluble fraction obtained by freeze-thawing cell suspensions and separating the cell envelope and cytosolic fractions by high-speed centrifugation. The use of freeze-thawing as a technique of cell lysis has several advantages over other methods in the study of enzyme activities. Sonication may not be desirable on account of the heat generated at the tip of the probe, that could result in denaturation of proteins. It has also been shown that sonication gives rise to H. and OH. radicals which can lead to protein fragmentation (13). Similarly, chemical lysis may affect enzyme activity detrimentally by disrupting enzyme complexes, etc. Some techniques, such as sonication and rupturing with glass beads, employ relatively harsh disruption procedures whose action may dislodge membrane-associated proteins, and thus yield positive activity in the soluble fraction. Freeze-thawing is an efficient and relatively 'gentle' process of cell disruption, which appears to open holes in the cell envelope and have minimal effects on its overall integrity, ensuring little shearing of membrane-bound proteins and good recovery of soluble proteins (14, 15). It is important when lysing cells to take into account not only the efficiency of the lysis procedure in breaking open the cells, but also of any effects the method may have on processes one may subsequently wish to study. For these reasons freeze-thawing was adopted as the preferred method for producing cell free extracts for studying ACTase activity in H. pylori.
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