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Continuous enzyme-coupled assay of phosphate- or pyrophosphate-releasing enzymes
 
Antonio S. Guillén Suárez1,3, Alessandra Stefan1,2, Silvia Lemma1, Emanuele Conte1, and Alejandro Hochkoeppler1,2, 3
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Supplementary Material

It is important to note that INT-formazan can be detected at wavelengths compatible with assays dealing with partially purified enzyme preparations. To test this point, we overexpressed E. coli DNA polymerase III catalytic core, containing the α (polymerase), ε (3′-5′ exonuclease), and θ subunits. In particular, soluble proteins were extracted from E. coli TOP10 subjected to overexpression of the core, and the extract accordingly obtained was loaded onto a Q-Sepharose FF column and eluted with a linear NaCl gradient. An aliquot of each collected fraction was then subjected to our coupled assay performed in microplates in the presence of 0.5 mM INT (see Materials and methods). As a reference assay, we determined in parallel the exonuclease activity of ε, under the conditions reported by Hamdan et al. (17). The polymerase and the exonuclease activity peaks were found to overlap satisfactorily (Supplementary Figure S5), indicating that our INT-based microplate assay of polymerase activity is reliable, and can be conveniently used to monitor purification of phosphate- or pyrophosphate-releasing enzymes.

The Malachite green method was also used to test the activity, in the presence of activated calf thymus DNA, of a partially purified preparation of E. coli DNA pol-III (13). After 30 min of incubation at 37°C, we estimated the release of 1 ± 0.15 nmol of pyrophosphate. When the enzyme-coupled assay was performed under the same conditions (in the presence of oxygen as electron acceptor), we observed polymerase activity equal to 0.7 ± 0.04 nmol of pyrophosphate generated in 30 min. Taking into account the comparisons presented here, we propose the PPX coupled assay as a reliable approach to continuously determine the activity of DNA polymerases.

To evaluate the potential use of our enzyme coupled assay over a wide pH interval, we determined the molar extinction coefficient of uric acid at pH 6 and 9. To this aim, we used the previously described Mes-Tris universal buffer (23), which is known to span a wide pH interval at constant ionic strength (23). As Figure 3A-3B show, at both pH values uric acid features maximal absorbance at 290 nm. This observation was confirmed, for both pH values, by inspection of the first-derivative spectra (data not shown). Our estimate of λmax significantly differs from previously reported values, indicating 293 nm as the wavelength of maximal molar absorbance by uric acid (14). However, it should be mentioned that Smith observed a significant shift (from 292 to 295 nm) of λmax, when comparing the spectra of uric acid in phosphate and carbonate buffers (24). Moreover, a concentration dependence shift of the specific molar absorbance of uric acid was reported (25). We therefore propose 290 nm as the λmax for uric acid at pH 6 and 9, in Mes-Tris buffer. In addition, according to the data reported in Figure 3A-3B, we propose 12 × 103 and 12.4 × 103 M−1cm−1 as the ε of uric acid at 290 nm, at pH 6 and 9, respectively.




Figure 3.  Absorption spectra of 25, 50, 75, and 100 μM uric acid at pH 6 (A) and 9 (B). (Click to enlarge)


The activities of Cellulomonas sp. PNPase (18) and bacterial XOD feature pH optimum at 7.5–8.0. On the contrary, the pH optimum of E. coli PPase is equal to 8.5–9.0 (26). Therefore, to further test the pH-dependence of our enzyme-coupled assay, we determined PPase activity as a function of pH, using the Mes-Tris universal buffer. As Figure 4 shows, PPase activity features a maximum at pH 8.5–9.0, in agreement with previous observations reported by Josse (26). In addition, at pH 6 the activity drops to 15% of the maximum (Figure 4). It is interesting to note that Josse, using maleate buffer, reported at pH 6 a relative activity equal to less than 10% of the maximum (26). The difference between these and our observations could be because of our use of a universal buffer, which avoids potentially inhibitory or stimulatory effects exerted by the different buffers used in activity assays performed as a function of pH. Therefore, we propose that our enzyme coupled assay could be used at pH 6–7 upon increasing the concentration of PPase, e.g. to 10–50 mU/mL instead of 2–10 mU/mL, as optimal for assays at pH 7–8.




Figure 4.  Pyrophosphatase (PPase) activity as a function of pH, in the presence of the universal Mes-Tris buffer and 10 mU/mL of PPase. (Click to enlarge)


According to the observations reported here, the PPX enzyme coupled assay was shown to represent a robust and sensitive method to perform activity assays of phosphate- or pyrophosphate-releasing enzymes. Using this coupled assay, enzyme activities can indeed be determined continuously, quantitatively, and over a wide pH range. Moreover, the assay can be performed in microplates to determine the activity of partially purified enzyme preparations.

Competing interests

The authors declare no competing interests.

Correspondence
Address correspondence to Alejandro Hochkoeppler, Department of Industrial Chemistry, University of Bologna, Viale Risorgimento 4, 40136 Bologna, Italy. Email: [email protected]

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