| Citation: |
Changrong Liu, Yao Xiao, Yilin Xiao, Zhiyong Li. 2021: Marine urease with higher thermostability, pH and salinity tolerance from marine sponge-derived Penicillium steckii S4-4. Marine Life Science & Technology, 3(1): 77-84. DOI: 10.1007/s42995-020-00076-6
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Urease (urea amidohydrolase, EC 3.5.1.5) as the first enzyme crystallized from Jack bean and the first enzymatic protein possessing nickel ions in the active site (Dixon et al. 1975), hydrolyzes urea to yield ammonia and carbamate. To date, urease has been found widespread in organisms including archaea (Bhatnagar et al. 1984), cyanobacteria (Carvajal et al. 1982; Collier et al. 1999; Palinska et al. 2000; Rai 1989), bacteria (Benoit and Maier 2011; Cai and Ni 1996; Liu et al. 2017a; Todd and Hausinger 1987; Turbett et al. 1992), fungi (Geweely 2006; Jahns 1995; Lubbers et al. 1996; Mirbod et al. 2002), plants (Das et al. 2002; El-Hefnawy et al. 2014; Hirayama et al. 2000; Menegassi et al. 2008) and invertebrates (Mcdonald et al. 1980).
Urease has been well used in determining the concentration of blood urea in clinical diagnosis (Qin and Cabral 2002), removing the urea lest forming carcinogenic ethyl carbamate in alcoholic beverages (Yang et al. 2015), detecting heavy metal ion pollutants in environmental protection (Preininger 1999; Wittekindt et al. 1996), and eliminating urea in industrial wastewaters (George et al. 1997). The most common sources of urease are from plants, especially the Jack bean (Canavalia ensiformis). But the pH optima of plant ureases are commonly neutral or alkaline and most ureases are not heat-stable, thus, restricting their further developments in some special fields (Das et al. 2002; El-Hefnawy et al. 2014; Hirayama et al. 2000; Kostecka-Madalska and Noculak 1973; Krajewska 2009; Menegassi et al. 2008; Prakash and Bhushan 1997). As such, the exploration of novel ureases with higher thermostability and pH resistance attracts much interest (Qin and Cabral 2002).
Oceans, covering 71% of the surface of the earth, harbor abundant microorganisms including bacteria, fungi, archaea and microalgae that inhabit special marine ecosystems (Feng et al. 2016; Rodriguez-Marconi et al. 2015). Numerous novel natural products and enzymes have been isolated from marine organisms (Simmons et al. 2008). It would be interesting to determine if the marine-derived ureases have different characteristics from those isolated from terrestrial origins; however, studies of marine-derived urease are still very limited (Mobley and Hausinger 1989; Mobley et al. 1995).
Urea is one of the dominant organic nitrogenous compounds in the oligotrophic oceans. Phylogenetically diverse bacterial ureC genes have been detected in sponges by our group (Su et al. 2013), indicating the role of sponge microbial symbionts in the regenerated utilization of urea in the sea waters. Therefore, microbes derived from sponges are important resources for urease. In this study, marine sponge Siphonochalina sp.-derived Penicillium steckii S4-4 with high urease activity was investigated with the aim to produce a urease with novel characteristics.
More than 20 fungi derived from marine sponge Siphonochalina sp. were screened for urease activity. Penicillium steckii S4-4 showed the highest activity and, thus, was selected for further investigation. Both NiSO4·6H2O and urea were added to examining the effect of Ni2+ and urea on the urease production of Penicillium steckii S4-4. As exhibited in Fig. 1, urease activity per milligram mycelium reached a climax when NiSO4·6H2O concentration was above 0.006% and the urease activity was enhanced along with the increased concentration of urea. However, the fungus cultivated in broth with 0.4% urea yielded the maximal biomass and thus showed the highest total urease activity (data not shown). As a result, it was found that 0.006% NiSO4·6H2O and 0.4% urea could increase urease production approximately 2.5 fold.
The protein quantification and urease activity of each purification fraction were assayed and are summarized in Table 1. The final purification recovered 17.54% of the original urease activity and reached a final specific activity of 1542.2 U mg protein-1 which was 61.4-fold of the crude extract activity. The 40-50% saturation fraction of ammonium sulfate precipitation showed the highest specific activity (62.36 U mg-1), followed by ion-exchange column chromatography where two peaks arose and the urease was eluted as a single peak in fraction 25 with 0.4 mol L-1 NaCl (Fig. 2a). During the last step, the elution profile (Fig. 2b) showed four peaks of proteins, and urease activity displayed a maximum at fraction 24 with a single peak.
| Fraction | Specific activity (U mg−1) | Total activity (units) | Total protein (mg) | Purification (fold) | Yield (%) |
| Crude extract | 25.12 | 6596.01 | 262.58 | 1 | 100 |
| Crude enzyme solution | 62.36 | 4946.40 | 79.32 | 2.48 | 75 |
| Hitrap-DEAE fraction | 272.09 | 2345.42 | 8.62 | 10.83 | 35.56 |
| Superdex-200 fraction | 1542.20 | 1156.65 | 0.75 | 61.40 | 17.54 |
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As native-PAGE shows in Fig. 3a, a purple-red colored band homogeneously occurred in each lane. Likewise, the number of protein bands decreased during the processing of separation and only one single protein band with a molecular weight of 47 kDa was observed in the Superdex-200 fraction via SDS-PAGE (Fig. 3b). The relative native molecular mass of P. steckii S4-4 urease was estimated to be approximately 183 kDa by gel filtration (Fig. 4).
The N-terminal amino acid sequence of the purified 47 kDa monomer was (M) GPVLKKTKAAAV. The best alignment for P. steckii S4-4 urease was completed when a two amino acid gap was placed at the third and fourth residue position and a five amino acid gap appeared at the sixth to tenth position (Fig. 5). Overall, ureases from eukaryotic organisms showed a higher affinity to P. steckii S4-4 urease relative to other prokaryotic sources. The urease sequence with the greatest similarity to the P. steckii S4-4 urease was from Ectocarpus siliculosus, a kind of marine alga.
As demonstrated in Fig. 6, the purified urease exhibited an estimated Km and Vmax of 7.3 mmol L-1 and 1.8 mmol urea min-1 mg protein-1, respectively, at 55 ℃, pH 8.5 and salinity 10%. The effects of temperature, pH and salinity on urease activity were evaluated using the Jack bean urease as a control (Fig. 7). It was found that the optimal temperature, pH and salinity for the urease from P. steckii S4-4 were 55 ℃, 8.5 and 10%, respectively, while the Jack bean ureases showed the highest activity at 35 ℃, pH 7.0 and 5% salinity. The Jack bean urease was stable and retained more than 80% of its maximal activity at a temperature lower than 40 ℃ (Fig. 7b), pH 6.0-8.5 (Fig. 7d) or salinity gradient of 0-10% (Fig. 7f). In contrast, the marine urease from P. steckii S4-4 exhibited higher thermostability, broader pH resistance and stronger salinity tolerance. More than 80% of its maximal activity was maintained at temperatures up to 60 ℃, pH 5.5-10.0 or 0-25% salinity (Fig. 7b, d, f).
In this study, the production of the marine urease by sponge Siphonochalina sp.-derived P. steckii S4-4 was enhanced by nickel, similar to previous investigations on bacteria (Liu et al. 2017b) and fungi (Jahns 1995). It is because the maturation of urease is dependent on recruiting nickel ions to the active site which facilitates the enzyme to exhibit higher activity (Benoit and Maier 2011; Rando et al. 1990). However, urease activity could be inhibited at high concentrations of nickel (Mirbod et al. 2002), because the transcription of urease genes is repressed under high nickel levels (Benoit and Maier 2011). The disparity of the additional nickel's effect is due to the difference of the complex growth media where the contents of available nickel and nickel-chelating compounds are not the same (Rando et al. 1990). Urea is an inducer for urease production (Jones and Mobley 1988; Kakimoto et al. 1990; Mobley et al. 1986). The results from this study showed that 0.4% urea addition could improve the urease production about two-fold. Whereas, a higher concentration of urea caused a lower total urease activity; it was speculated that cell growth could be affected due to the changes of nitrogen content in the broth (Yao and Ye 2016).
The chromatographically purified urease showed a specific activity of 1542.2 U mg protein-1, which was higher than all other reported fungal urease (ranging from 62.5 to 1341 U mg protein-1 (Creaser and Porter 1985; Geweely 2006; Jahns 1995; Lubbers et al. 1996; Phillips et al. 1991; Smith et al. 1993) except the urease from Coccidioides immitis (1750 U mg protein-1) (Mirbod et al. 2002). A relatively higher urease-specific activity would benefit the catalytical reaction thus accelerating the response speed of various applications using the enzyme. Furthermore, the urease produced by P. steckii S4-4 showed a high affinity to urea, with a Km of 7.3 mmol L-1 which was higher or close to the Km observed from other fungi, e.g., Aspergillus nidulans (Creaser and Porter 1985), Aspergillus niger (Smith et al. 1993), Coccidioides immitis (Mirbod et al. 2002), Rhodosporidium paludigenum (Phillips et al. 1991), Schizossacharomyces pome (Lubbers et al. 1996), Sporobolomyces roseus (Jahns 1995) and Ustilago violacea (Baird and Garber 1981).
The native molecular mass of urease from P. steckii S4-4 was estimated to be 183 kDa, which is the second smallest size among all the reported purified fungal ureases possessing molecular weights ranging from 172 to 560 kDa (Creaser and Porter 1985; Geweely 2006; Jahns 1995; Lubbers et al. 1996; Mirbod et al. 2002; Phillips et al. 1991; Smith et al. 1993). Nevertheless, the smaller molecular weight may be beneficial to the subsequent X-ray crystallographic analysis or other processes which are related to molecular weights (Mobley and Hausinger 1989). Besides, a relatively small protein would be beneficial to the heterologous expression of the enzyme (Liu et al. 2017b). Only one single band of urease appeared on native-PAGE of each fraction indicating that only a single species of urease existed unless varieties with the same electrophoretic mobility are present simultaneously. Similarly, only one band with a molecular mass of 47 kDa was observed in the final enzyme preparation suggesting that the urease from P. steckii S4-4 only has one subunit. It is widely accepted that the feature of a single type of urease subunit is conservative among all eukaryotes, although the molecular mass of the monomer may differ from one to another, e.g., molecular dimension of 40 kDa for the A. nidulans urease (Creaser and Porter 1985), 72 kDa for the R. paludigenum urease (Phillips et al. 1991), 90 kDa for the Jack bean urease (Balasubramanian and Ponnuraj 2010), and 98.3 kDa for the urease from Cotton (Gossypium hirsutum) Seeds (Menegassi et al. 2008). It is suggested that the P. steckii S4-4 urease is a homogeneous tetramer by combining the results of PAGE with gel filtration. The subunit stoichiometries of other eukaryotic ureases vary from two in S. pome (Lubbers et al. 1996), three in A. niger (Smith et al. 1993), six in A. nidulans (Creaser and Porter 1985), and eight in R. paludigenum (Phillips et al. 1991).
Unlike the terrestrial Jack bean urease, which shows optimal activity at pH 7.0 and is stable at pH 6.0-8.5, marine urease from P. steckii S4-4 is active over a broad pH range of 5.5-10.0 with the maximum activity at pH 8.5, that is higher than most other fungal ureases (Baird and Garber 1981; Creaser and Porter 1985; Geweely 2006; Jahns 1995; Lubbers et al. 1996; Mirbod et al. 2002; Phillips et al. 1991; Smith et al. 1993), implying the P. steckii S4-4 urease has potential applications in alkaline environments. The majority of reported bacterial ureases show maximal activity over the range of 7.0-7.75 (Mobley and Hausinger 1989), and most plant ureases' optimal pH is slightly above neutral (Das et al. 2002; El-Hefnawy et al. 2014; Hirayama et al. 2000; Menegassi et al. 2008; Prakash and Bhushan 1997). Thus, the P. steckii S4-4 urease will be an effective alternative for present neutral or weakly alkaline enzymes, e.g., in the treatment of wastewaters with pH higher than 9.0 (Qin and Cabral 2002).
The urease's optimal salinity and salinity tolerance of urease were first reported in this study. As expected, marine urease from P. steckii S4-4 showed the maximal activity at the salinity of 10% and retained more than 80% of its maximal activity at salinity up to 25%, which was much higher than that for the Jack bean urease. The characteristics are speculated to be related to the acclimation to sea water which has an average salinity of 3.5%, and it may give a hint to the application of marine-derived ureases in high salinity environments, for instance, detecting heavy metal ions and other pollutants in seawaters (Zhylyak et al. 1995).
The marine urease produced by P. steckii S4-4 showed maximal activity at 55 ℃ and was stable up to 60 ℃, exhibiting higher thermostability than the terrestrial urease from Jack bean, which had an optimal temperature at 35 ℃ and was stable below 40 ℃. These features are similar to all other marine origin ureases obtained from R. paludigenum (Phillips et al. 1991), Sporobolomyces roseus (Jahns 1995) and Prochlorococcus marinus (Palinska et al. 2000) that have maximum activity at 50-60 ℃, 65 and 60 ℃, respectively. The marine-derived ureases demonstrate higher thermal stability than the terrestrial origins and thus may be applicable in thermal environments.
In summary, the preliminary differences between the P. steckii S4-4 urease and the terrestrial Jack bean urease referring to thermostability, pH stability and salt tolerance may provide new insight to develop novel ureases from marine resources.
Penicillium steckii S4-4 was isolated from the sponge Siphonochalina sp. in the South China Sea (18.42° N, 109.97° E) and was grown on agar containing 1% peptone, 0.3% beef extract and 0.4% urea prepared in artificial seawater (Kester et al. 1967) at 28 ℃ for five days. The conidia were harvested and inoculated into 100 ml (250-ml shaking flask) liquid medium containing 2% d-glucose, 1% peptone, 0.5% beef extract, 0.5% yeast extract, 0.2% KH2PO4, 0.4% urea and 0.006% NiSO4·6H2O in artificial seawater on a rotary shaker (180 r min-1) at 28 ℃ for four days. A concentration gradient of 0-0.01% NiSO4·6H2O and 0-0.6% urea was added into the broth to evaluate their effects on urease production. The mycelia were harvested from the broth by filtration through Whatman 40 filter paper (GE Healthcare, USA), rinsing thoroughly with the same medium volume Tris-HCl buffer (50 mmol L-1, pH 8.5) three times, then immediately freezing at - 80 ℃ until use.
The frozen mycelium was ground in liquid nitrogen using a mortar and pestle, then the powder was suspended in pre-cold Tris-HCl buffer (50 mmol L-1, pH 8.5) and disrupted with an ultrasonic apparatus (Shanghai Jingxin Experimental Technology, China). Subsequently, the homogenate was centrifuged at 20, 000g for 30 min at 4 ℃, and the clear supernatant was used as the crude extract where ammonium sulfate powder was added slowly with a constant stirring. The precipitated proteins were collected by centrifugation as above, and the part with the highest specific enzyme activity was dissolved in pre-cold Tris-HCl buffer (50 mmol L-1, pH 8.5) which was then dialyzed against the same buffer at 4 ℃ for 24 h. The resulting solution was then centrifuged as stated above, and the clear supernatant was designated as crude enzyme solution followed by loading on a Hitrap DEAE FF column (GE Healthcare, USA), which was performed as the procedure described previously (Mirbod et al. 2002). The eluate fractions with the highest urease-specific activity were pooled and dialyzed against Tris-HCl buffer (50 mmol L-1, pH 8.5). The retentate was then filtered through a 0.45 μm PES membrane (Millipore, USA) to remove particulate materials. The filtrate was referred to as the Hitrap-DEAE fraction which was then concentrated by ultrafiltration through an Amicon Ultra-15 membrane (Millipore, USA) which is permeable to molecules with Mr < 10, 000. The concentrated sample was then filtered using a PES membrane and applied to a Superdex 200 10/300 GL column (GE Healthcare, USA) where 1 ml fractions were collected (Lubbers et al. 1996). The fraction with the highest urease-specific activity was isolated and designated as the Superdex-200 fraction. All procedures of urease isolation and purification were carried out at 4 ℃ unless otherwise stated.
The amount of protein was determined by BCA protein assay kit (Biomiga, USA), with bovine serum albumin as the standard according to the Bradford method (Bradford 1976). Urease activity was measured by monitoring the rate of release of ammonium from urea. The concentration of ammonium in the reaction mixture was determined as indophenol produced by a phenol-hypochlorite reaction at 625 nm (Weatherburn 1967) and the main steps were executed as reported previously (Liu et al. 2017a). Control assays included the detection of ammonium in the reaction mixture at time zero and when using boiled enzyme extract. Urease specific activity in each chromatography fraction was defined as the amount of enzyme per mg of protein in the solution required to hydrolyze 1 μmol urea per min at 55 ℃ and pH 8.5. One unit of enzyme activity was defined as the amount of enzyme required to hydrolyze 1 μmol urea per min at 55 ℃ and pH 8.5 under standard conditions.
The native molecular weight of the purified urease was determined by gel filtration using the Superdex 200 10/300 GL column according to the method reported previously (El-Hefnawy et al. 2014). The column was calibrated using the standard proteins (GE Healthcare, USA) as follows: blue dextran (2000 kDa), thyroglobulin (669 kDa), ferritin (440 kDa), β-amylase (200 kDa), aldolase (158 kDa), bovine serum albumin (66 kDa) and ovalbumin (43 kDa).
Native (non-denaturing) and sodium dodecyl sulfate (denaturing and reducing) PAGE were performed using a modified protocol of Laemmli (1970) on an 8-20% gradient precast protein gel. The urease existence in the native PAGE gel was visualized as described by Sharma et al. (2008), while the proteins were stained with Coomassie brilliant blue G-250 (Blakesley and Boezi 1977). Protein molecule weight marker was purchased from ThermoFisher Scientific.
The purified urease was subjected to SDS-PAGE and transferred by electroblotting to a polyvinylidene difluoride (PVDF) membrane. After being stained with Ponceau S, the protein band was excised and sequenced based on the Edman method using a PPSQ-33A apparatus (SHIMADZU, Japan). Multiple alignments of protein sequences were performed using the CLUSTAL W alignment program (Thompson et al. 1994).
The Km and Vmax for the purified urease were obtained from a Lineweaver-Burk plot using a part of the Superdex 200 fraction (Lubbers et al. 1996). The optima of temperature, pH and salinity for the purified urease and Jack bean urease (Sigma-Aldrich, USA) were obtained by comparing its activity after incubation at different temperature over a range of 25-70 ℃, pH 4.0-10.0 using citric acid-sodium citrate buffer (50 mmol L-1, pH 4-6.5), Tris-HCl buffer (50 mmol L-1, pH 7-9) and sodium carbonate-sodium bicarbonate buffer (50 mmol L-1, pH 9.5-10) or sodium chloride gradient from 0-25%, separately. Analogously, the thermostability, pH stability and the tolerance to salinity of the urease were determined by contrasting the residual activity after pre-incubating the purified enzyme at different temperature for 30 min, pH at 4 ℃ for 6 h, or salinity gradients at 4 ℃ for 6 h, respectively. All tests were conducted in triplicate (Liu et al. 2017a).
This work was supported by the National Key Research and Development Program of China (2018YFC030980504). The authors would like to thank BiotechPack Co. Ltd for the determination of N-terminal amino acid sequence.
CL finished the experiments and wrote the manuscript, YX analyzed the data, YX wrote the manuscript, ZL designed the experiments and revised the manuscript.
The authors declare that they have no conflict of interest.
This article does not contain any studies with human participants or animals performed by any of the authors.
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Figures(7) / Tables(1)
Supported by: Beijing Renhe Information Technology Co.,
Ltd.
support:
info@rhhz.net
| Fraction | Specific activity (U mg−1) | Total activity (units) | Total protein (mg) | Purification (fold) | Yield (%) |
| Crude extract | 25.12 | 6596.01 | 262.58 | 1 | 100 |
| Crude enzyme solution | 62.36 | 4946.40 | 79.32 | 2.48 | 75 |
| Hitrap-DEAE fraction | 272.09 | 2345.42 | 8.62 | 10.83 | 35.56 |
| Superdex-200 fraction | 1542.20 | 1156.65 | 0.75 | 61.40 | 17.54 |
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