Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1
Mingzhu Ding
 -- 2397 2022-08-16 12:48:22 |
2 format correct
Catherine Yang
 Meta information modification 2397 2022-08-17 03:43:15 |

Video Upload Options

Do you have a full video?
Send video materials Upload full video

Confirm

Are you sure to Delete?
Yes No
Cite
If you have any further questions, please contact Encyclopedia Editorial Office.
Zhang, N.;  Ding, M.;  Yuan, Y. Biodegradation of Polyolefins. Encyclopedia. Available online: https://encyclopedia.pub/entry/26204 (accessed on 24 April 2024).
Zhang N,  Ding M,  Yuan Y. Biodegradation of Polyolefins. Encyclopedia. Available at: https://encyclopedia.pub/entry/26204. Accessed April 24, 2024.
Zhang, Ni, Mingzhu Ding, Yingjin Yuan. "Biodegradation of Polyolefins" Encyclopedia, https://encyclopedia.pub/entry/26204 (accessed April 24, 2024).
Zhang, N.,  Ding, M., & Yuan, Y. (2022, August 16). Biodegradation of Polyolefins. In Encyclopedia. https://encyclopedia.pub/entry/26204
Zhang, Ni, et al. "Biodegradation of Polyolefins." Encyclopedia. Web. 16 August, 2022.
Biodegradation of Polyolefins
Edit
This entry is adapted from the peer-reviewed paper 10.3390/microorganisms10081537

Polyolefins, including PE, PP and PS, are composed of C–C and C–H bonds, which are more stable against degradation than ester bonds. During polyolefin biodegradation, C–C and C–H bonds are oxidized. Many types of microorganisms have been isolated from sea water, compost and activated sludge with the capacity for polyolefin biodegradation.With the development of synthetic biology and metabolic engineering strategies, engineered strains could be used to degrade plastics.

polyolefins polyethylene biodegradation

1. Isolated Microorganisms

Numerous of microorganisms with the capacity for polyolefin degradation, including bacteria, fungi and microbial consortia, have been isolated from the environment, such as soil containing plastic waste, the ocean and the guts of plastic-eating worms. These microorganisms are capable of utilizing polyolefins as sole carbon source or can generate depolymerases involved in polyolefin degradation. Therefore, the screening of these microorganisms is vital for the further degradation of polyolefins. Polyolefin-degrading bacteria can be identified by the following procedure. Soil samples collected from an area containing polyolefin waste was mixed with water and shaken [1]. In order to isolate strains from waxworm guts, worm gust need to be isolated and suspended in water [2]. Then, a portion of the suspension was transferred to polyolefin-containing medium. After incubation for several days, the polyolefin-degrading strains would be isolated through gradient dilution the of medium or collecting the strains on the agar media on which PE fragments were spread [2][3][4].

1.1. Single Bacteria

At present, most single strains that are capable of degrading polyolefins are bacteria, which can form biofilms on the surface of polyolefins or destroy the surface of polyolefins. An increased rate of PE degradation was detected after LDPE was pretreated with UV light [5]. Changes in the molecular weight and molecular number of LDPE were also detected after incubation with R. ruber C208 [6]. It was reported that R. ruber C208 is also capable of degrading PS, achieving a weight loss of 0.8% after 8 weeks [7]. Moreover, R. rhodochrous ATCC 29672 exhibited the ability to degrade PP based on the characterization of changes in the metabolic activity of bacteria, such as ATP content, ADP/ATP ratio and cell viability [8].
Biodegradation should not only be determined by weight loss. Chemical properties, as well as changes in Mw (molecular weight) and Mn (molecular number), which can be characterized Fourier transform infrared spectroscopy (FTIR), X-ray photoelectron spectroscopy (XPS), nuclear magnetic resonance (NMR) and gel permeation chromatography (GPC) analyses, are also vital for the measurement of the degree of polyolefin degradation. After the biodeterioration stage, the nature and occurrence of functional groups on the surface of polyethylene substrates are changed, which can be studied by FTIR [9] spectroscopy, XPS [3] and NMR [10]. After the biofragmentation stage, the length of polyolefins is shorted. The result of biofragmentation can be proven by using GPC to measure changes in Mw and Mn [11]. After the biofragmentation stage, degradation products (such alkanes, alkenes, etc.) can be determined by GC-MS (gas chromatography mass spectrometry) [12]. Moreover, the result of bioassimilation can be determined through measurement of dry biomass weight of polyethylene-containing media. Finally, the percent of mineralization can be analyzed by CO2 measurement [13]. Furthermore, scanning electron microscopy (SEM) is another vital method used to characterize the surface features of polyolefins.
As previously reported, some waxworms can chew and eat plastics [3][14][15][16][17]. Enterobacter asburiae YT1 and Bacillus sp. YP1 were isolated from the guts of waxworms with the capability of degrading PE. Over 28-day incubation of the two strains on PE films, the physical properties (tensile strength and surface topography), chemical structure (hydrophobicity and appearance of carbonyl groups), Mw (accompanied by the formation of daughter products) and weight loss were detected [3]. Kyaw et al. [18] incubated Pseudomonas aeruginosa PAO1 with LDPE films. After exposure to P. aeruginosa PAO1, the LDPE sample turned into a mixture of long-chain fatty acids, esters, hydrocarbons, oxygenated chemical compounds predominantly containing aldehydes, ketones, esters and ether groups, unsaturated fatty acids and certain unknown compounds. Given that the structure of polyolefins is similar to that of alkanes, some strains with an ability to degrade alkanes also have an effect on PE degradation. Alcanivorax borkumensis, a bacterial strain isolated from the sea that can utilize alkane as carbon source [19] was found to induce a weight loss of 3.5% in 7 days [20].

1.2. Microbial Consortia

A mixed microbial consortium consisting of two Bacillus sp. and two Paenibacillus sp. was isolated from a landfill site. After incubation with the mixed microbial consortium for 30 days, the weight and the mean diameter of the PE sample were reduced 16.7% and 22.8%, respectively [21]. A microbial consortium consisting of Lysinibacillus xylanilyticus and Aspergillus niger was found to be capable of degrading PE [22]. After UV irradiation, the mineralization percentage increased from 15.8% to 29.5% after incubation for 126 days in soil, and according to FTIR and XRD, the chemical properties also improved. Skariyachan et al. [23] isolated a microbial consortium comprising Brevibacillus sps. and Aneurinibacillus sp. from waste landfills. During a 140-day incubation of the two strains on PE samples, weight loss for LDPE and HDPE strips reached 58.2% and 46.6% respectively, and weight loss for LDPE and HDPE pellets were 45.7% and 37.2%, respectively. As with a single bacterium, UV pretreatment aided in the degradation of polyolefin samples. In addition, the result of GC-MS analysis indicated that PE samples were degraded into cis-2-chlorovinylacetate, tri-decanoic acid and octadecanoic acid. Muenmee et al. [24] pretreated HDPE, LDPE, PP and PS samples with UV for 200 h, then mixed the plastics together. To simulate a landfill environment, the plastics mixture was placed in simulated lysimeters with a synthetic landfill gas (60% CH4:40% CO2) and a microbial consortium composed of Methylocyctis sp., Methylocella sp., Methylobactor sp., Methyloccus capsulatus., Nitrosomonas sp., Nitrosomonas europaea, Nitrobacter winogradskyi, Nitrobacter hamburgensis, Burkholderia sp., Pseudomonas sp. and Xanthobacter sp. After a reaction period of 3 months under semi-aerobic landfill conditions where different aeration rates were supplied, the plastic samples were degraded into hydrocarbon and oxygenated compounds, such as aliphatic alkanes, alkenes, alcohols and esters. The degradation products of different plastic types, such as the degraded products of HDPE, are mainly alkanes (C24H50, C32H66), alkenes (C15H30, C19H38) and some alcohols (C15H32O, C20H42O) were found, while for LDPE, only one alcohol (C11H24O) was found.

2. Engineered Strains

2.1. Hydrolases Capable of Polyolefin Degradation

Most the enzymes capable of degrading PE are oxidoreductases. As shown in Table 1, well-known identified enzymes with the ability to oxidize polyolefins include laccase, manganese peroxidase, alkane hydroxylase and soybean peroxidase. Furthermore, a PS-degrading enzyme called hydroquinone peroxidase was identified from Azotobacter beijerinckii HM121. However, no PP-degrading enzymes have been identified to date. The result of enzymatic degradation of PE is that oxidation groups are introduced into the PE chain, which means polyolefins cannot be oxidized into monomers as the only act of these enzymes. Most PE-degrading enzymes can only perform terminal oxidation (the terminal carbon in polyolefins be oxidized) and subterminal oxidation (the carbon adjacent to the terminal carbon in polyolefins be oxidized) of PE. For example, laccase and manganese peroxidase can perform terminal oxidation, and the AlkB family can degrade n-alkanes, the main component of polyethylene, through either terminal or subterminal hydroxylation reactions [25]. Therefore, an ideal polyolefin-degrading enzyme has high hydroxylation activity against any carbon in the carbon chain so as to achieve the efficient transformation of polyolefin to its oligomer or monomer.
Laccases (EC 1.10.3.2), which belong to the so-called blue-copper family of oxidases, can catalyze the oxidation of a wide range of phenols and arylamines. Laccases, which are glycoproteins, have been reported in higher plants, fungi and bacteria. A laccase was purified from R. ruber C208 with the ability to degrade LDPE. As laccases contain four copper ion bonding sites, copper markedly affects their induction and activity, resulting in PE degradation. mRNA quantification by RT-PCR revealed a 13-fold increase in laccase mRNA levels in copper-treated cultures compared with an untreated control. The addition of copper to C208 cultures containing PE enhanced the biodegradation of PE by 75% [1][26]. A laccase mediator system (LMS) is composed of laccase and some small-molecule compounds that are easily oxidized by laccase, such as HBT, ABTS and DMP. In the process of LMS oxidation, laccase oxidizes the mediator first; then, the oxidized mediator oxidizes the substrate. It was reported that in the presence of a mediator, laccase can oxidize some substrates that it cannot oxidize alone. HBT, which has been used for PE degradation, reacts with non-phenolic models by a radical mechanism involving hydrogen atom abstraction [27]. Some researchers treated polyethylene with LMS using HBT (0.2 mM) as a mediator; after 3 days, the polyethylene membrane exhibited no elongation, and its relative tensile strength decreased by about 60%, which is higher than in the absence of HBT (20%) [28]. Johnnie et al. used laccase from Trichoderma viride fungus and 1-HBT to degrade LDPE. After incubation for 10 days, the weight loss of LDPE came to 2.3% [29].
Cytochrome P450 (CYP, P450), a member of a superfamily of heme–thiolate proteins, is distributed in most living organisms. There are more than 300,000 P450 genes. However, no P450 genes were found in E. coli, which means that E. coil is a good chassis for the heterologous expression of P450 genes. P450 enzymes can identify multiple substrates and catalyze diverse reactions, such as C–H hydroxylation; C=C double-bond epoxidation; heteroatom oxygenation; O-, N- and S-dealkylation; aromatic coupling; and C–C bond cleavage [30]. Because other PE-degrading enzymes can only perform terminal oxidation and subterminal oxidation of PE, the application of an ideal P450 enzyme that can cleave PE into short chains would contribute to the biodegradation of PE.
Alkane, which is composed of C–C bonds and C–H bonds, has a similar structure to that of PE. Therefore, alkane mono-oxygenase enzymes are potential candidates for the degradation of PE. One alkane hydroxylase, namely AlkB, has been reported to degrade PE [30]. AlkB, which was first identified in alkane-consuming Pseudomonas species isolated from oil-contaminated areas, is a membrane-bound, non-heme di-iron monooxygenase [31].
Manganese peroxidase was purified from a lignin-degrading fungus: P. chrysosporium. Manganese peroxidase was first identified as a lignin-degrading enzyme. It was reported that the addition of Mn (II) to nitrogen- or carbon-limited culture medium enhanced PE degradation [32].

2.2. Engineered Chassis for Polyolefin Biodegradation

As shown in Table 1, the microorganisms that are capable of secreting polyolefin biodegrading enzymes are not model organisms, which means that they are difficult to genetically engineer. Two model organisms, E. coli and Y. lipolytica, have been applied to the expression and secretion of polyolefin-biodegrading enzymes to date [33][34][35]. The heterologous expression of polyolefin-degrading enzymes in model organisms can efficiently increase the expression level of polyolefin-degrading enzymes through genetic engineering in model organisms.
E. coli:E. coli is one of the most widely used model microorganisms for production of recombinant proteins. As a model microorganism, E. coli has advantages in many aspects, such as a simple genetic background, ease of genetic modification and simple growth conditions. Engineered E. coli has been used to express alkane hydroxylase to degrade LMWPE. The alkB gene, which was cloned from Pseudomonas sp. E4, was introduced into E. coli BL21. After incubation for 80 days at 37 °C with engineered E. coli, 19.3% of the LMWPE was degraded [13]. The rubredoxin and rubredoxin reductase could help alkane monooxygenase to transfer electron. If the rubredoxin and rubredoxin reductase are co-expressed with alkane monooxygenase, the conversion rate can be increased. Researchers fused and expressed alkB with its coenzyme genes—rubA1, rubA2 and rub—in E. coli BL21. The result indicated that 30.5% of the carbon of LMWPE-1 degraded into CO2 after 78 days [36]. Another study revealed that an alkane-1-monooxygenase (AlkB) in Acinetobacter johnsonii JNU01 degraded PS, and this finding was later confirmed by recombinant alkB in E. coli BL21 [37]. These researchers also expressed alkB2 in E. coli, and the result indicates that alkB2 was more efficient for low-molecular-weight PE biodegradation than alkB1 [33]. A laccase gene isolated from a marine fungus was expressed in E. coli and showed PE degradation ability [38].
Many recent studies have shown that engineered E. coli can be used for laccase heterologous expression and secretion [39][40][41]. Ihssen et al. [34] expressed five novel bacterial laccase-like multicopper oxidases (LMCOs) of diverse origin. However, a potential issue with laccase expression in E. coli is that it is easy for E. coli to form inclusion bodies when expressing extracellular enzymes. Mo et al. [35] expressed three laccases from three different organisms, namely Lac1326 from marine sediment samples, fungal tvel5 laccase from Trametes versicolor and bacterial BPUL laccases from Bacillus pumilus for the purpose of degrading β-estradiol. The result of Western blot analysis indicates that laccase was detected both in vivo and in vitro in E. coli, which means that some laccase stayed in inclusion bodies instead of being secreted in vitro. Given that laccase has been widely used for the biodegradation of PE [10][29][42][43], the heterologous expression of laccase in E. coli is a potential method for the biodegradation of PE.
Yarrowia lipolytica:Y. lipolytica is a Crabtree-negative ascomycete yeast with good protein secretion capacities. Compared to other yeasts, Y. lipolytica lacks α-1,3-mannosyltransferase, a factor that limits the amount of excessive mannosylation of secreted heterologous glycoproteins and constitutes a valuable asset for the production of therapeutic proteins [44]. Y. lipolytica W29 is a wild-type strain with a remarkable characteristically high secretion level of proteins [45]. Y. lipolytica W29 ura302 was obtained through genetic convention of URA3 into ura3-302 in Y. lipolytica W29, and it was able to utilize sucrose and molasses as a carbon source under the control of XPR2 promoter. After the genetic convention of XPR2 into xpr2-322 and AXP1 into axp1-2, which indicates inactivation of alkaline extracellular protease and acid extracellular protease that would degrade foreign extracellular protein, a strain with high heterologous protein production capability called Y. lipolytica Po1f was obtained. A new strain called Y. lipolytica Po1g that carries a pBR322 docking platform was obtained through the integration of PINA300′ plasmid in Y. lipolytica Po1f [46]. Y. lipolytica Po1g was induced with a YLEX kit for expression/secretion of heterologous proteins [47]. A laccase from the white-rot fungus Trametes versicolor was expressed in Y. lipolytica Po1g for the biodegradation of PE. Compared to the yeast secretion signal, the native secretion signal showed higher enzyme activity in the culture medium. The yield of laccase reached 2.5 mg/L (0.23 units/mL) [26]. Laccase has been widely used for the biodegradation of PE, and the heterologous expression of laccase in Y. lipolytica is a promising method of PE degradation.
Table 1. Enzymes capable of polyolefin degradation.
Plastics Enzymes Enzyme Source Pretreatment Experimental Condition Result Reference
PE Laccase R. ruber C208 Unpretreated LDPE film Incubation for 30 days at 30 °C Weight loss: 1.5–2.5%; reduction of 20% in Mw and 15% in Mn [1]
Manganese peroxidase Phanerochaete chrysosporium Unpretreated PE film Incubation for 12 days at 37 °C Mw decreased [32]
Soybean peroxidase Soybean Unpretreated HDPE film Reaction for 2 h at 60 °C Hydrophilicity increased [48]
Alkane hydroxylase Pseudomonas sp. E4 Unpretreated LMWPE sheet Incubation for 80 days at 37 °C Weight loss: 19.3% [13]
Alkane hydroxylase Pseudomonas aeruginosa E7(uniport Q9I0R2) Unpretreated LMWPE film Incubation for 50 days at 37 °C Weight loss: 19.6–30.5% [33]
PS Hydroquinone peroxidase Azotobacter beijerinckii HM121 Unpretreated PS film Incubation for 20 min Mw decreased [49]
Alkane hydroxylase A. johnsonii JNU01 Unpretreated low-molecular-weight PS powder Incubation for 7 days at 28 °C Confirmed by FTIR and SEM [26]

References

  1. Sivan, A.; Szanto, M.; Pavlov, V. Biofilm development of the polyethylene-degrading bacterium Rhodococcus ruber. Appl. Microbiol. Biotechnol. 2006, 72, 346–352.
  2. Divyalakshmi, S.; Athikayala, S. Screening and isolation of polyethylene degrading bacteria from various soil environments. J. Environ. Sci. 2016, 12, 01–07.
  3. Yang, J.; Yang, Y.; Wu, W.-M.; Zhao, J.; Jiang, L. Evidence of Polyethylene Biodegradation by Bacterial Strains from the Guts of Plastic-Eating Waxworms. Environ. Sci. Technol. 2014, 48, 13776–13784.
  4. Koutny, M.; Amato, P.; Muchova, M.; Ruzicka, J.; Delort, A.-M. Soil bacterial strains able to grow on the surface of oxidized polyethylene film containing prooxidant additives. Int. Biodeterior. Biodegrad. 2009, 63, 354–357.
  5. Gilan, I.; Hadar, Y.; Sivan, A. Colonization, biofilm formation and biodegradation of polyethylene by a strain of Rhodococcus ruber. Appl. Microbiol. Biotechnol. 2004, 65, 97–104.
  6. Santo, M.; Weitsman, R.; Sivan, A. The role of the copper-binding enzyme—Laccase—In the biodegradation of polyethylene by the actinomycete Rhodococcus ruber. Int. Biodeterior. Biodegrad. 2013, 84, 204–210.
  7. Ward, P.G.; Goff, M.; Donner, M.; Kaminsky, W.; O’Connor, K.E. A Two Step Chemo-biotechnological Conversion of Polystyrene to a Biodegradable Thermoplastic. Environ. Sci. Technol. 2006, 40, 2433–2437.
  8. Fontanella, S.; Bonhomme, S.; Brusson, J.-M.; Pitteri, S.; Samuel, G.; Pichon, G.; Lacoste, J.; Fromageot, D.; Lemaire, J.; Delort, A.-M. Comparison of biodegradability of various polypropylene films containing pro-oxidant additives based on Mn, Mn/Fe or Co. Polym. Degrad. Stab. 2013, 98, 875–884.
  9. Restrepo-Flórez, J.-M.; Bassi, A.; Thompson, M.R. Microbial degradation and deterioration of polyethylene—A review. Int. Biodeterior. Biodegrad. 2014, 88, 83–90.
  10. Sarmah, P.; Rout, J. Efficient biodegradation of low-density polyethylene by cyanobacteria isolated from submerged polyethylene surface in domestic sewage water. Environ. Sci. Pollut. Res. 2018, 25, 33508–33520.
  11. Montazer, Z.; Najafi, M.B.H.; Levin, D.B. Challenges with Verifying Microbial Degradation of Polyethylene. Polymers 2020, 12, 123.
  12. Yamada-Onodera, K.; Mukumoto, H.; Katsuyaya, Y.; Saiganji, A.; Tani, Y. Degradation of polyethylene by a fungus, Penicillium simplicissimum YK. Polym. Degrad. Stab. 2001, 72, 323–327.
  13. Yoon, M.G.; Jeon, H.J.; Kim, M.N. Biodegradation of Polyethylene by a Soil Bacterium and AlkB Cloned Recombinant Cell. J. Bioremediation Biodegrad. 2012, 3, 145.
  14. Ru, J.K.; Huo, Y.X.; Yang, Y. Microbial Degradation and Valorization of Plastic Wastes. Front. Microbiol. 2020, 11, 442.
  15. Bombelli, P.; Howe, C.J.; Bertocchini, F. Polyethylene bio-degradation by caterpillars of the wax moth Galleria mellonella. Curr. Biol. 2017, 27, R292–R293.
  16. Chalup, A.; Ayup, M.M.; Garzia, A.C.M.; Malizia, A.; Martin, E.; De Cristóbal, R.; Galindo-Cardona, A. First report of the lesser wax moth Achroia grisella F. (Lepidoptera: Pyralidae) consuming polyethylene (silo-bag) in northwestern Argentina. J. Apic. Res. 2018, 57, 569–571.
  17. Kundungal, H.; Gangarapu, M.; Sarangapani, S.; Patchaiyappan, A.; Devipriya, S.P. Efficient biodegradation of polyethylene (HDPE) waste by the plastic-eating lesser waxworm (Achroia grisella). Environ. Sci. Pollut. Res. 2019, 26, 18509–18519.
  18. Kyaw, B.M.; Champakalakshmi, R.; Sakharkar, M.K.; Lim, C.S.; Sakharkar, K.R. Biodegradation of Low Density Polythene (LDPE) by Pseudomonas Species. Indian J. Microbiol. 2012, 52, 411–419.
  19. Sabirova, J.S.; Ferrer, M.; Regenhardt, D.; Timmis, K.N.; Golyshin, P.N. Proteomic Insights into Metabolic Adaptations in Alcanivorax borkumensis Induced by Alkane Utilization. J. Bacteriol. 2006, 188, 3763–3773.
  20. Delacuvellerie, A.; Cyriaque, V.; Gobert, S.; Benali, S.; Wattiez, R. The plastisphere in marine ecosystem hosts potential specific microbial degraders including Alcanivorax borkumensis as a key player for the low-density polyethylene degradation. J. Hazard. Mater. 2019, 380, 120899.
  21. Park, S.Y.; Kim, C.G. Biodegradation of micro-polyethylene particles by bacterial colonization of a mixed microbial consortium isolated from a landfill site. Chemosphere 2019, 222, 527–533.
  22. Esmaeili, A.; Pourbabaee, A.A.; Alikhani, H.A.; Shabani, F.; Esmaeili, E. Biodegradation of Low-Density Polyethylene (LDPE) by Mixed Culture of Lysinibacillus xylanilyticus and Aspergillus niger in Soil. PLoS ONE 2013, 8, e71720.
  23. Skariyachan, S.; Patil, A.A.; Shankar, A.; Manjunath, M.; Bachappanavar, N.; Kiran, S. Enhanced polymer degradation of polyethylene and polypropylene by novel thermophilic consortia of Brevibacillus sps. and Aneurinibacillus sp. screened from waste management landfills and sewage treatment plants. Polym. Degrad. Stab. 2018, 149, 52–68.
  24. Muenmee, S.; Chiemchaisri, W.; Chiemchaisri, C. Enhancement of biodegradation of plastic wastes via methane oxidation in semi-aerobic landfill. Int. Biodeterior. Biodegrad. 2016, 113, 244–255.
  25. Santacruz-Juárez, E.; Buendia-Corona, R.E.; Ramírez, R.E.; Sánchez, C. Fungal enzymes for the degradation of polyethylene: Molecular docking simulation and biodegradation pathway proposal. J. Hazard. Mater. 2021, 411, 125118.
  26. Jolivalt, C.; Madzak, C.; Brault, A.; Caminade, E.; Malosse, C.; Mougin, C. Expression of laccase IIIb from the white-rot fungus Trametes versicolor in the yeast Yarrowia lipolytica for environmental applications. Appl. Microbiol. Biotechnol. 2005, 66, 450–456.
  27. Crestini, C.; Jurasek, L.; Argyropoulos, D.S. On the Mechanism of the Laccase–Mediator System in the Oxidation of Lignin. Chem.—A Eur. J. 2003, 9, 5371–5378.
  28. Fujisawa, M.; Hirai, H.; Nishida, T. Degradation of Polyethylene and Nylon-66 by the Laccase-Mediator System. J. Polym. Environ. 2001, 9, 103–108.
  29. Johnnie, D.A.; Issac, R.; Prabha, M.L. Bio Efficacy Assay of Laccase Isolated and Characterized from Trichoderma viride in Biodegradation of Low Density Polyethylene (LDPE) and Textile Industrial Effluent Dyes. J. Pure Appl. Microbiol. 2021, 15, 410–420.
  30. Yeom, S.-J.; Le, T.-K.; Yun, C.-H. P450-driven plastic-degrading synthetic bacteria. Trends Biotechnol. 2022, 40, 166–179.
  31. Chen, C.-C.; Dai, L.H.; Ma, L.X.; Guo, R.-T. Enzymatic degradation of plant biomass and synthetic polymers. Nat. Rev. Chem. 2020, 4, 114–126.
  32. Iiyoshi, Y.; Tsutsumi, Y.; Nishida, T. Polyethylene degradation by lignin-degrading fungi and manganese peroxidase. J. Wood Sci. 1998, 44, 222–229.
  33. Jeon, H.J.; Kim, M.N. Comparison of the functional characterization between alkane monooxygenases for low-molecular-weight polyethylene biodegradation. Int. Biodeterior. Biodegrad. 2016, 114, 202–208.
  34. Ihssen, J.; Reiss, R.; Luchsinger, R.; Thöny-Meyer, L.; Richter, M. Biochemical properties and yields of diverse bacterial laccase-like multicopper oxidases expressed in Escherichia coli. Sci. Rep. 2015, 5, 10465.
  35. Mo, Y.; Lao, H.I.; Au, S.W.; Li, I.C.; Hu, J.; Yuen, H.M.; Cheong, W.M.; Lo, O.L.I.; Seak, L.C.U. Expression, secretion and functional characterization of three laccases in E. coli. Synth. Syst. Biotechnol. 2022, 7, 474–480.
  36. Jeon, H.J.; Kim, M.N. Functional analysis of alkane hydroxylase system derived from Pseudomonas aeruginosa E7 for low molecular weight polyethylene biodegradation. Int. Biodeterior. Biodegradation 2015, 103, 141–146.
  37. Muhonja, C.N.; Makonde, H.; Magoma, G.; Imbuga, M. Biodegradability of polyethylene by bacteria and fungi from Dandora dumpsite Nairobi-Kenya. PLoS ONE 2018, 13, e0198446.
  38. Gao, R.; Liu, R.; Sun, C. A marine fungus Alternaria alternata FB1 efficiently degrades polyethylene. J. Hazard. Mater. 2022, 431, 128671.
  39. Wang, T.-N.; Zhao, M. A simple strategy for extracellular production of CotA laccase in Escherichia coli and decolorization of simulated textile effluent by recombinant laccase. Appl. Microbiol. Biotechnol. 2016, 101, 685–696.
  40. Grandes-Blanco, A.I.; Tlecuitl-Beristain, S.; Díaz, R.; Sánchez, C.; Téllez-Téllez, M.; Márquez-Domínguez, L.; Santos-López, G.; Díaz-Godínez, G. Heterologous Expression of Laccase (LACP83) of Pleurotus ostreatus. BioResources 2017, 12, 3211–3221.
  41. Guo, H.; Zheng, B.; Jiang, D.; Qin, W.S. Overexpression of a Laccase with Dye Decolorization Activity from Bacillus sp. Induced in Escherichia coli. J. Mol. Microbiol. Biotechnol. 2017, 27, 217–227.
  42. Sowmya, H.V.; Ramalingappa; Krishnappa, M.; Thippeswamy, B. Degradation of polyethylene by Trichoderma harzianum—SEM, FTIR, and NMR analyses. Environ. Monit. Assess. 2014, 186, 6577–6586.
  43. Sowmya, H.V.; Ramalingappa; Krishnappa, M.; Thippeswamy, B. Degradation of polyethylene by Penicillium simplicissimum isolated from local dumpsite of Shivamogga district. Environ. Dev. Sustain. 2015, 17, 731–745.
  44. De Pourcq, K.; Vervecken, W.; Dewerte, I.; Valevska, A.; Van Hecke, A.; Callewaert, N. Engineering the yeast Yarrowia lipolytica for the production of therapeutic proteins homogeneously glycosylated with Man8GlcNAc2 and Man5GlcNAc2. Microb. Cell Factories 2012, 11, 53.
  45. Ogrydziak, D.M.; Nicaud, J.-M. Characterization of Yarrowia lipolyticaXPR2 multi-copy strains over-producing alkaline extracellular protease—A system for rapidly increasing secretory pathway cargo loads. FEMS Yeast Res. 2012, 12, 938–948.
  46. Madzak, C.; Tréton, B.; Blanchin-Roland, S. Strong hybrid promoters and integrative expression/secretion vectors for quasi-constitutive expression of heterologous proteins in the yeast Yarrowia lipolytica. J. Mol. Microbiol. Biotechnol. 2000, 2, 207–216.
  47. Madzak, C.; Beckerich, J.-M. Heterologous Protein Expression and Secretion in Yarrowia lipolytica. In Yarrowia lipolytica: Biotechnological Applications; Springer: Berlin/Heidelberg, Germany, 2013; Volume 25, pp. 1–76.
  48. Zhao, J.C.; Guo, Z.; Ma, X.Y.; Liang, G.Z.; Wang, J.L. Novel surface modification of high-density polyethylene films by using enzymatic catalysis. J. Appl. Polym. Sci. 2004, 91, 3673–3678.
  49. Nakamiya, K.; Ooi, T.; Kinoshita, S. Non-heme hydroquinone peroxidase from Azotobacter beijerinckii HM121. J. Ferment. Bioeng. 1997, 84, 14–21.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
Upload a video for this entry
Information
Subjects: Microbiology
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to https://encyclopedia.pub/register :
Ni Zhang
,
Mingzhu Ding
,
Yingjin Yuan
View Times: 697
Entry Collection: Biopharmaceuticals Technology
Revisions: 2 times (View History)
Update Date: 17 Aug 2022
Table of Contents
    1000/1000
    Hot Most Recent
    Feedback