Part:BBa_K5043009
phtAc from M. vanbaalenii Pyr-1
PhtAc codes for phthalate dioxygenase ferredoxin subunit. It is a subunit of ring-hydroxylating oxygenase enzyme complexes which participate in biodegradation of polycyclic aromatic hydrocarbons. [1] This part encodes for the same protein as BBa_J73042. It only differs in synonym codon usage.
Sequence and Features
- 10COMPATIBLE WITH RFC[10]
- 12COMPATIBLE WITH RFC[12]
- 21COMPATIBLE WITH RFC[21]
- 23COMPATIBLE WITH RFC[23]
- 25COMPATIBLE WITH RFC[25]
- 1000COMPATIBLE WITH RFC[1000]
Contents
Introduction
In this project, we aimed to establish a pyrene degradation pathway in Pseudomonas vancouverensis and to integrate it with the organism's native phenanthrene degradation pathway [2]. It is established that PhtAcAd, as electron transfer components, along with a ring-hydroxylating dioxygenase system formed by pdoA2 and pdoB2, aggregate to create a complex exhibiting dioxygenase activity [3, 4, 5]. This complex is capable of converting Phenanthrene-4-carboxylate, an intermediate product of pyrene degradation, into cis-3,4-dihydroxyphenanthrene an intermediate in the native phenanthrene degradation pathway of P. vancouverensis, thereby facilitating the conjunction of the two pathways [4, 5]. Given that this enzyme complex serves as a critical junction within the pathways, we opted to conduct a more detailed characterization. The components of the electron transport chain, PhtAc a ferredoxin reductase and ferredoxin PhtAd, were produced and purified to elucidate their kinetic parameters. Subsequently, we aimed to analyze the complex formation with PdoA2B2 through HPLC analysis.
Enzyme production and purification
PhtAc coding sequence was cloned in pQE bacterial expression plasmid with a cleavable N-terminal, 6x-His tag. It was expressed in E. coli BL21 (DE3). Main cultures were incubated at 37°C until an OD600 of 0.5 was reached. Subsequently, culture was induced with a final concentration of 0.5 mM IPTG and incubated overnight at 30°C. Purification was performed using Immobilized Metal Affinity Chromatography (IMAC). Both production and purification samples were analyzed via SDS-PAGE and Coomassie Staining. Enzyme concentration was determined using the Bradford Assay, PhtAc (7.2 kDA) final yield was 3682.8 µg/ml, with a final concentration of 511.5 µM.
Characterization of ferredoxin reductase PhtAd and ferredoxin PhtAc
The characterization of PhtAd (insert part) activity was performed by evaluating its ability to reduce 2,6-dichlorophenolindophenol (DCPIP), which serves as an electron acceptor [6]. In its oxidized form, DCPIP displays a blue color, which transitions to colorless upon reduction, facilitating the evaluation of PhtAd activity [6]. The decrease in absorbance was monitored at 600 nm [6].
The activity of PhtAc was assessed using a coupled assay. The increase in absorbance resulting from an electron transfer to Cytochrome c signifies an interaction between ferredoxin PhtAc and ferredoxin reductase PhtAd [6]. The increase in absorbance was monitored at 5500 nm [6]. Both reactions needed NADH and FADH as cofactors [6].
Kinetic characterization of PhtAc
Method
Initial measurements were conducted in accordance with the methodology outlined by Wu et al. (2019). However, the assay produced inconclusive results, as no variation in absorption was observed over time. This lack of change was attributed to the high concentration of Cytochrome C (600 µM) utilized in the initial trials. Consequently, we opted to conduct further assays employing varying concentrations of Cytochrome C and NADH to identify the optimal concentrations that would facilitate a measurable increase in absorption. In alignment with the approach taken by Wu et al. (2019), subsequent assays were performed using a PhtAd to PhtAc ratio of 1:3.5. The increase in absorbance was measured over a 30-minute period at 2-minute intervals at 550 nm. After the optimal Cytochrome C concentration was established, further assays were performed varying the NADH concentration to determine the kinetic parameters.
Results
Determination of the optimal substrate concentration
In the initial assay conducted with 250 µM Cytochrome C, no change in absorbance was observed (see Fig. 3A). Considering this outcome, the subsequent assay was performed using 100 µM Cytochrome C in conjunction with a higher concentration of NADH (200 µM). This combination resulted in a measurable increase in absorbance, demonstrating a linear progression over a duration of 26 minutes (see Fig. 3C) indicating the oxidation of Cytochrome C. To validate this outcome, additional assays were conducted utilizing 100 µM Cytochrome C alongside varying concentrations of NADH, both higher and lower. The assay employing 250 µM NADH also demonstrated a linear increase in absorption (see Fig. 3B) in the first 26 minutes. Conversely, the assay with 100 µM NADH (See Fig. 3D) also exhibited a linear increase in absorption, albeit at a reduced rate. Notably, no saturation point was observed at this concentration, in contrast to the higher concentrations, which indicated saturation occurring approximately 30 minutes into the assay.
Fig. 5 Oxidation of Cytochrome c catalyzed by PhtAc, in conjunction with PhtAd, under varying concentrations of Cytochrome c and NADH. The oxidation of Cytochrome C by PhtAc coupled with PhtAd was monitored for 30 min at a constant enzyme concentration and varying amount of substrate. The reaction mixture contained 100 mM Tris/HCl (pH 7.5), 100 – 250 µM NADH, 100 and 250 µM Cyt. C, 0.5 µM FAD, 1 µM PhtAd and 3.5 µM PhtAc. Reaction was performed at room temperature.
Based on the results obtained, we chose a concentration of 100 µM cytochrome C for the subsequent tests, as this concentration showed a measurable increase in absorbance. For the reaction time, we decided to run further reactions for 34 minutes to confirm the saturation point.
Kinetic parameters of PhtAc
For the determination of the kinetic parameters the Assay was runned with different NADH concentrations ranging from 40 µM to 400 µM. Data obtained from these assays were then analyzed using Michaelis-Menten kinetics (See Fig. 6) and a Lineweaver-Burk plot (See Fig. 7) was constructed to provide a linear representation of the enzyme kinetics to determine key parameters such as Vmax and Km.
Fig. 6 Michaelis-Menten Curve for PhtAc. The velocity was calculated utilizing the Lambert-Beer Law with the extinction coefficient of Cytochrome C (ε550 21 mM-1 x cm–1) [6]. A logarithmic trend line was applied as a preliminary approximation.
Fig. 7 Lineweaver-Burk plot. Graphs show the fitted linear function and its correlation coefficient.
The Vmax, Km, kcat, kcat/Km values of the ferredoxin PhtAc were 0.00043 mM/min, 0.31 mM, 0.12 1/min and 3.9 1/mM x min, respectively. Significant deviations from the literature values (Wu et al., 2019) were observed; however, the reaction was conducted with a substantially lower concentration of Cytochrome C and elevated concentrations of enzymes, as the enzymes produced exhibited reduced activity.
Activity assay coupled with HPLC analysis
We successfully characterized and demonstrated the activity of PhtAc and PhtAd both individually and as a complex. This complex functions as an electron carrier for PdoA2B2 [3, 4, 5]. The subsequent step involved conducting an activity assay in conjunction with HPLC analysis with the aim to evaluate the function of PdoA2B2 both independently and in complex with PhtAcAd, forming a tetramer.
Method
Initially, the activity assay was conducted, wherein the enzymes were incubated at 30°C for approximately 15 to 20 minutes, both in the presence and absence of NADH to examine the effect of NADH on enzymatic aggregation. Subsequently, Phenanthrene-4-carboxylate (P4C) was added to initiate the reaction. The reaction mixture was then incubated for 30 minutes at 30°C, after which the reaction was terminated by the addition of 100% methanol. Following this, High-Performance Liquid Chromatography (HPLC) analysis was performed using a Zorbax SB-C18 column, employing an acetonitrile/water gradient at a flow rate of 0.4 ml/min, with the column oven maintained at room temperature. The gradient elution program commenced with an initial mobile phase of 60:40 (v/v) acetonitrile to water, transitioning linearly to 100% acetonitrile over 14 minutes, followed by a return to the initial phase (60:40) after 5 minutes. The total duration for each analysis was 35 minutes. For the negative control, an assay was conducted using denatured enzymes, achieved by heating the enzymes at 95°C for 10 minutes.
Results
Initial activity assays were performed solely with PdoA2 and PdoB2 to form a dimer, thereby validating their enzymatic activity and evaluating their aggregation characteristics. Monomers were incubated together for aggregation in the presence and absence of NADH to determine whether NADH, as a cofactor, influences the aggregation process.
The efficacy of the reaction was assessed by measuring the absorbance of Phenanthrene-4-carboxylate (P4C) and NADH using HPLC. The area under the resulting peaks corresponding to the levels of P4C and NADH was determined, enabling a comparative analysis of the consumption of P4C and NADH against established standards and between the samples. Furthermore, to eliminate potential statistical error in the assessment of our results, we employed triplicate measurements for each sample and calculated their averages. The results are presented in Figure 1. Results indicated that the assays in which NADH was added prior to aggregation exhibited enhanced P4C degradation and, correspondingly, increased NADH consumption compared to the assays where NADH was added after aggregation. However, the observed NADH consumption was relatively low, which may be attributed to the absence of the electron carrier dimer (PhtAcAd) in the sample, thereby limiting NADH consumption.
Fig. 8 Comparative Analysis of Phenanthrene-4-carboxylat (P4C) and NADH Consumption in PdoA2:PdoB2 activity assays. The degradation of P4C by PdoA2:PdoB2 was monitored for 30 min at a constant substrate and enzyme concentration. The reaction mixture contained 100 mM Tris/HCl (pH 7.5), 25 µM P4C, 1 µM PdoA2 and 1 µM PdoB2. The 100 µM NADH were either added after aggregation (a.A) or before aggregation (b.A).
Considering the initial results, the analysis of the tetramer aggregation process and the degradation of Phenanthrene-4-carboxylat was conducted by introducing NADH prior to the aggregation process.
Degradation of Phenanthrene-4-carboxylat by enzyme complex with dioxygenase activity using NADH as a cofactor
The presence of the functional enzyme complex with dioxygenase activity is anticipated to result in a reduction in the quantity of Phenanthrene-4-carboxylate. This reduction can be quantified by determining the decrease in the area of the corresponding peak in comparison to the standard. Given the unknown stoichiometry of this enzyme complex, we conducted assays utilizing two distinct stoichiometric ratios. The first assay employed a 1:1 stoichiometry of all monomers, while the second utilized a ratio of PdoA2:PdoB2:PhtAc3:PhtAd. This latter stoichiometry was selected based on previous kinetic analyses of PhtAc, which were performed with a ratio of three PhtAc molecules to one PhtAd for determining kinetic parameters. This ratio was employed in the present assays to assess its influence on the degradation of Phenanthrene-4-carboxylate and to determine whether the presence of three PhtAc molecules enhanced the efficiency of the electron transfer process, thereby accelerating the breakdown of Phenanthrene-4-carboxylate. However, results showed (see Fig. 2) that the breakdown of Phenanthrene-4-carboxylat was slightly more efficient with the 1:1 ratio of the monomers; this could suggest that a higher ratio of PhtAc does not enhance the electron transfer process. And that the 1:1 ratio may have created a more favorable environment for the interaction of the active site with the substrate.
To confirm the aggregation of the enzyme complex and the successful degradation of P4C by this complex, negative controls were prepared. For this, assays were prepared with denatured enzymes with the stoichiometries described above. Results showed (see Fig. 2) that the peak area of P4C within these assays was higher in comparison with the assays with the active enzyme complex. The comparison of these results indicates that the enzyme complex was successfully formed and demonstrated the capacity to degrade P4C.
Another negative control was conducted by excluding any enzymes from the assay. In comparison to samples containing denatured enzymes, the results showed a reduced peak area in the peak corresponding to Phenanthrene-4-carboxylat, suggesting either a diminished quantity of P4C in the sample or its degradation. However, the possibility of cross-reactivity with other components in the assay can be discounted, as such interactions would also manifest in the other negative controls. This observation indicates the presence of an independent source of error. Potential issues may involve the unintentional introduction of enzymes or inadequate substrate concentration. The presence of enzymes in the assay can be corroborated by taking into consideration the amount of NADH; notably, the peak area of NADH in this sample was diminished suggesting the possibility that a reaction has taken place, and the results are more consistent with those obtained from the assay conducted with a 1:3 enzyme ratio (see Fig. 2). While numerous errors could have arisen, a systematic issue such as cross-reactivity, which would render the enzyme ineffective, can be dismissed by comparing the sample to those with non-functional enzymes. Moreover, as anticipated, the samples containing the active enzyme complex demonstrated a decrease in the peak area associated with NADH relative to the standard (see Fig. 2). This finding corroborates the functionality of the electron carrier dimer, which collaborates with PdoA2B2 in the degradation of P4C. In contrast, the assay involving denatured enzymes at a 1:1 ratio revealed no consumption of NADH, indicating the absence of any reaction.
However, if we observed the results of NADH consumption from the assay involving denatured enzymes, utilizing three molecules of PhtAc in conjunction with a 1:1 ratio of the other enzymes, demonstrate a reduction in NADH consumption, as evidenced by the diminished area of the corresponding peak relative to the standard. This observation indicates the presence of an independent source of error, as the other assay performed with denatured enzymes shows no NADH consumption. Potential issues may include insufficient NADH concentration. If the NADH concentration was low, degradation could have occurred over time, as the activity assays were conducted days prior to the HPLC analysis. The thawing process and exposure of the sample to temperature fluctuations may have contributed to NADH degradation due to its inherent instability. Additionally, the possibility of incomplete denaturation should be considered. If the enzymes were not fully denatured during the assay, residual active enzyme could remain, resulting in NADH consumption. The denaturation conditions may not have been adequate to completely inactivate the enzymes. This is further supported by the observation that P4C consumption was slightly lower in comparison to the sample with denatured enzymes with the 1:1 ratio.
In conclusion, the enzyme complex with dioxygenase activity demonstrated its ability to degrade Phenanthrene-4-carboxylate (P4C), with slightly greater efficiency observed in the assay utilizing a 1:1 ratio of monomers compared to the 1:3 ratio of PhtAc to PdoA2:PdoB2:PhtAd. This suggests that a higher ratio of PhtAc does not enhance electron transfer efficiency. Negative controls, including assays with denatured enzymes and no enzymes, confirmed that the active enzyme complex was responsible for P4C degradation, while the reduction in NADH in some denatured enzyme assays points to possible errors, such as NADH degradation or incomplete enzyme denaturation. Further assays with stricter controls are needed to confirm these findings and minimize potential sources of error.
Fig. 9 Comparative Analysis of Phenanthrene-4-carboxylat (P4C) and NADH Consumption in PdoA2:PdoB2:PhtAc:PhtAd activity assays. The degradation of P4C by the enzym complex was monitored for 30 min at a constant substrate and enzyme concentration. The reaction mixture contained 100 mM Tris/HCl (pH 7.5), 200 µM NADH, 40 µM P4C, 1 µM PdoA2, 1 µM PdoB2, 1 or 3 µM PhtAc and 1 µM PhtAd.
References
1. S.-J. Kim, O. Kweon, R. C. Jones, J. P. Freeman, R. D. Edmondson, and C. E. Cerniglia, "Complete and integrated pyrene degradation pathway in Mycobacterium vanbaalenii PYR-1 based on systems biology," Journal of bacteriology, vol. 189, no. 2, pp. 464–472, 2007, doi: 10.1128/JB.01310-06.
2. Y. Yang, R. F. Chen, and M. P. Shiaris, "Metabolism of naphthalene, fluorene, and phenanthrene: preliminary characterization of a cloned gene cluster from Pseudomonas putida NCIB 9816," Journal of bacteriology, vol. 176, no. 8, pp. 2158–2164, 1994, doi 10.1128/jb.176.8.2158-2164.1994.
3. Pagnout, C., Frache, G., Poupin, P., Maunit, B., Muller, J., & Férard, J. (2007). Isolation and characterization of a gene cluster involved in PAH degradation in Mycobacterium sp. strain SNP11: Expression in Mycobacterium smegmatis mc2155. Research in Microbiology, 158(2), 175–186. https://doi.org/10.1016/j.resmic.2006.11.002
4. Yuan, K., Xie, X., Wang, X., Lin, L., Yang, L., Luan, T., & Chen, B. (2018). Transcriptional response of Mycobacterium sp. strain A1-PYR to multiple polycyclic aromatic hydrocarbon contaminations. Environmental Pollution, 243, 824–832. https://doi.org/10.1016/j.envpol.2018.09.001
5. Krivobok, S., Kuony, S., Meyer, C., Louwagie, M., Willison, J. C., & Jouanneau, Y. (2003). Identification of Pyrene-Induced Proteins in Mycobacterium sp. Strain 6PY1: Evidence for Two Ring-Hydroxylating Dioxygenases. Journal of Bacteriology, 185(13), 3828–3841. https://doi.org/10.1128/jb.185.13.3828-3841.2003
6. Wu, Y., Xu, Y., & Zhou, N. (2019). A newly defined dioxygenase system from Mycobacterium vanbaalenii PYR-1 endowed with an enhanced activity of dihydroxylation of high-molecular-weight polyaromatic hydrocarbons. Frontiers of Environmental Science & Engineering, 14(1). https://doi.org/10.1007/s11783-019-1193-5
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