Research paperCrystal structure analysis of peroxidase from the palm tree Chamaerops excelsa
Graphical abstract
Introduction
Peroxidases (EC 1.11.1.7; donor: hydrogen peroxide oxidoreductase) are heme proteins that catalyze the oxidoreduction of a broad variety of peroxides. Most commonly, peroxidases catalyze the oxidation of organic substrates, while reducing H2O2 to water. This process involves multiple-reactions and a number of intermediate enzyme forms, and is known as the Poulos–Kraut mechanism, which plays a key role in several metabolic responses of all peroxidases [1]. Peroxidases are important for many biological responses and processes, such as defense against pathogenic microorganisms, cell wall formation, and lignification [2].
The peroxidase superfamily includes animal and non-animal peroxidases, and the latter are generally subdivided into three classes, all sharing a similar three-dimensional fold despite their low amino acid sequence identity [3], [4]. Class I includes intracellular enzymes, such as plant ascorbate peroxidases, yeast cytochrome c peroxidases and bacterial catalases. Class II is composed of secreted peroxidases encoded exclusively by fungal organisms, including lignin peroxidase and Mn2+-dependent peroxidases. Finally, Class III consists of secreted plant peroxidases with molecular weights between 28 and 60 kDa [3], [5]. The Class III plant peroxidases perform a number of functions, e.g.: lignin and suberin formation, the cross-linking of cell wall components, and the synthesis of phytoalexins [6].
It is generally accepted that only Class I enzymes are able to form multimers (dimers and tetramers) while the enzymes of other classes are monomeric and glycosylated. At the same time, multimerization is a very common feature of many mammalian peroxidases that exert their functions as dimers [7], [8]. Despite the physiological importance of this phenomenon, little information is available about the multimerization of plant peroxidases. A dimeric structure for recombinant horseradish peroxidase has been reported previously in micellar systems [7] and water solutions [9], but it has not been observed for the native enzyme. Dimerization affects enzyme stability, activity and the immobilization of physical surfaces [9]. The absence of dimeric quaternary structures of native peroxidase was explained in terms of the high degree of its glycosylation. Heavy glycosylation and enzyme dimerization were also proposed to be the molecular events accounting for the greater stability of palm tree peroxidases [10], [11]. Once the peroxidases from the leaves of tropical plants, such as palm trees, were shown to have high stability, it became crucial to study the quaternary structure of these enzymes. The high stability of the palm tree peroxidases makes them ideal for biotechnological applications and of direct interest to the industry. That is why it became crucial to establish their structure as it impacts immobilization. However, the increased stability of proteins is frequently not a consequence of a single mechanism but instead involves a combination of several factors, including disulfide bond formation, multimerization and glycosylation [10], [12].
Structurally, peroxidases are mainly composed of α-helices and can be divided into two domains: one containing the heme group, and the other with a ferriprotoporphyrin prosthetic group, both located inside of a hydrophobic pocket [13].
Class I peroxidases are not glycosylated, whereas the second and third classes are glycosylated peroxidases with 2–8 N-linked glycans known to contribute to the protein stabilization [14]. Similarly, Class I peroxidases do not contain any disulphide bonds or calcium ions. However, all cysteine residues present in Class II and III enzymes form disulfide bonds, conferring those enzymes higher rigidity. Class II peroxidases also show two calcium-binding sites that have important structural and functional roles. Despite the differences between the peroxidase subfamilies, their similar 3D fold is preserved [13].
To date, several peroxidases from tropical palm-trees – Elaies guineensis, Roystonea regia, Trachycarpus fortunei and Chamaerops excelsa (T. fortunei) – have been isolated and characterized [15], [16], [17], [18]. These are structurally and functionally stable enzymes that exhibit higher thermal stability within a broad pH range and in the presence of denaturing agents in comparison to the well-studied horseradish peroxidase (HRP), soybean seed-coat peroxidase (SBP) and anionic peanut (Arachis hypogaea L.) peroxidase. It has been demonstrated that CEP is a highly stable enzyme over a pH-range of 2.5–10.0, with no noteworthy changes in enzymatic activity [19], [20]. Since enzymes that tolerate extreme pH and temperature conditions are important for various biotechnological applications, the commercial use of such peroxidases is of major interest, especially in the biocatalysis industry. Indeed, assays using modified electrodes of adsorbed anionic royal palm tree peroxidase (RPTP), anionic sweet potato peroxidase (SPP) and cationic horseradish peroxidase (HRP-C) as biosensors for the detection of hydrogen peroxide revealed that the RPTP-based electrodes are more sensitive and exhibit a wider linear range and a higher storage stability [21]. Accordingly, the unique catalytic and stability profile of these palm peroxidases is testimony to their potential as biocatalysts and biosensors for other biotechnological applications [22], [23], [24].
Although the three-dimensional structures of a considerable number of peroxidases have been determined, to date only a few of those belong to Class III plant peroxidases: horseradish [25], peanut [26], barley [27], thale cress [28], soybean [29] and royal palm tree [15]. Despite this, the exact structural features responsible for the improved properties of the palm tree peroxidases as compared to other plant peroxidases remain obscure.
In the present work, we solved the three-dimensional X-ray crystallographic structure of a Class III plant peroxidase isolated from the leaves of the palm tree C. excelsa (CEP) and compared it to other available peroxidase structures. Additionally, the new quaternary structure identified for CEP and for royal palm tree peroxidase (RPTP) [15] offers possible explanations for their high thermal stability. Our results provide new insights into the structure–function relationships of plant peroxidases and their quaternary structures.
Section snippets
Enzyme purification
CEP was purified from the palm tree C. excelsa as described previously [30]. Briefly, leaves (1820 g) from a three-year-old palm tree were milled and homogenized in 7.28 l distilled water for 22–24 h at room temperature. Excess material was removed by vacuum filtration and centrifugation (10, 000 g, 277 K for 15 min). Pigments were extracted by phase separation over approximately 20 h at 277 K after the addition to the supernatant of solid PEG at 14% (w/v) and solid ammonium sulfate at 10%
Structure determination
Crystal plates grew after approximately 24 h and reached their maximum size after one week [30]. Despite having a rather thin plate-shaped morphology, these crystals were found to be suitable for X-ray data collection. The crystals of native CEP belonged to the P212121 space group, had a solvent content of 42% and diffracted to 2.6 Å resolution. The final Rfactor and Rfree for the refined structure were 21.8 and 24.6% respectively. The quality of the final model was checked using the MolProbity
Conclusions
Native peroxidase was successfully extracted and purified from the leaves of the palm tree C. excelsa, crystallized, and its structure was solved by protein crystallography. Consistent with other Class III peroxidases, the structure confirmed that CEP is N-glycosylated and revealed that CEP has a typical peroxidase fold.
Detailed structural analysis revealed a possible dimeric assembly, conserved amongst the plant palm peroxidases CEP and PRPT but absent in other known peroxidase structures.
Conflict of interest
The authors declare that they have no competing interests.
Acknowledgments
We are thankful to the Brazilian National Synchrotron Light Source (LNLS) and to the staff members of the MX2 beamline. This work was supported by Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) via research grants 2009/05349-6, 2008/56255-9, 2010/52362-5 and 2012/22802-9 and by Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) via grants # 490022/2009-0 and 373143/2012-5.
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- 1
These authors contributed equally to the work.
- 2
Present address: Instituto de Estudios Biofuncionales, Departamento de Química-Física II, Facultad de Farmacia, Universidad Complutense de Madrid, 28040 Madrid, Spain.