Your browser does not support the NLM PubReader view.

Go to this page to see a list of supporting browsers.

Effect of microplastics on soil microbial community and microbial degradation of microplastics in soil: A review

Article information

Environmental Engineering Research. 2023;28(6)
Publication date (electronic) : 2023 February 20
doi : https://doi.org/10.4491/eer.2022.716
Kehinde Caleb Omidoyin1, Eun Hea Jho2,orcid_icon
1Department of Agricultural Chemistry, Chonnam National University, 77 Yongbong-ro, Buk-gu, Gwangju 61186, South Korea
2Department of Agricultural and Biological Chemistry, Chonnam National University, 77 Yongbong-ro, Buk-gu, Gwangju 61186, South Korea
†Corresponding author: E-mail: ejho001@jnu.ac.kr, Tel: +82 62 530 2134, Fax: +82 62 530 2139
Received 2022 November 29; Revised 2023 February 2; Accepted 2023 February 19.

Abstract

The mass production, continual usage, and improper disposal of plastic products have resulted in significant environmental pollution. The larger plastic polymers gradually break down into smaller particles called microplastics (<5 mm). Existing studies on the occurrence and ecological impact of microplastics have focused on the aquatic ecosystems, with very little attention given to the soil environment. The soil represents a natural sink for microplastics from sources such as sewage sludge, landfills, plastic mulch from agricultural activities, fertilizers, and municipal wastewater effluent. The current study, therefore, provides an overview of existing knowledge on soil microplastic pollution focusing on the impact of microplastics on soil microbial community and microbial degradation of microplastics in soil to systematically identify knowledge gaps to be filled with further research. Future research challenges to be addressed include detailed monitoring of the sources and distribution of microplastics in soil under different land uses, exploring diverse microorganisms in their natural environments for their microplastic biodegradation potential using cultivation-dependent and independent approaches, understanding the mechanism of ecological impacts of microplastics and contributions of microplastic additives, degradation products, and other adsorbed environmental pollutants on soil microbial community.

Abstract

Graphical Abstract

Keywords: Biodegradation; Microorganism-microplastic interactions; Microplastics; Microplastic additives; Soil microbial community

1. Introduction

Due to their durability, lightweight, and cost-effectiveness, plastics are widely and frequently used synthetic materials [1]. Almost every aspect of human activities has found synthetic plastics useful, and it can be concluded that, in recent years, plastic materials have grown to become indispensable, replacing other materials such as wood, metals, and glass in various applications. Different synthetic plastics have been used in the production of various materials such as polyethylene (PE) for shopping bags, plastic bottles, and toiletry bottles; polystyrene (PS) for food containers, packaging foams, and disposable cups; polyurethane (PU) for sealants, adhesives, and extrusion and injection-molded parts; polyvinyl chloride (PVC) for plumbing pipes and guttering, window frames, and shower curtains; polypropylene (PP) for microwavable containers, drinking straws, and plastic pressure pipe systems; polyethylene terephthalate (PET) for plastic films, engineering components, and carbonated drinks bottles; nylon for gears, bushings, and plastic bearings; polycarbonate for automobile components, plastic lenses, riot shields; and polytetrafluoroethylene (PTFE) for hookup wire, coaxial cables, and gaskets [2].

Since 1950, the production of plastics has rapidly increased on a global scale due to their extensive use. Between 1950 and 2019, the annual global production increased dramatically from 2 million tons to 368 million tons, with about 40% going toward single-use applications [3]. Virgin plastics produced are largely single-use convenience products that are discarded within a short period after use, and they result in a rapid and massive accumulation in the natural environments [4]. It is predicted that, by 2050, up to 26 billion tons of plastic waste will be generated and more than 50% will be discarded into landfills and eventually enter natural environments like oceans, lakes, rivers, cultivated lands, etc., thereby resulting in serious environmental pollution [5]. Undoubtedly, as a result of its wide distribution across the world [6–8], plastic pollution has become a global scale issue, and the fate of plastic waste in the environment is now a subject of increasing concern.

The term “microplastic (MP)” is proposed to be first used by the African scientists named Ryan and Moloney in 1990 in their research article titled “Plastic and other artefacts on South African beaches: temporal trends in abundance and composition” [9]. This term, however, became widely recognized among researchers after a report published by Thompson et al. [10] who examined the abundance of microplastics (MPs) in the sediment of beaches, estuarine, and subtidal around Plymouth in the United Kingdom. Since then, the term “microplastics” has been generally used to describe small particles of plastics [11]. The defining characteristics of MPs are still under debate [12], but it is agreed by most researchers that plastic particles ranging between 100 nm to <5 mm in size are regarded as MPs.

In the past years, most studies on MP pollution have focused on the marine environment, with very little attention drawn to the terrestrial environment, although between 2018 and 2021, more researchers have studied the effect of MPs on the soil ecosystem [13, 14]. Due to the critical role that the soil plays in regulating nutrient cycling, maintaining the biodiversity of organisms, and providing food [15], it is necessary to evaluate the ecological effect of MP pollution in the terrestrial environment, especially the soil [14, 16]. After the findings of Rillig [17] on the detrimental effects of MPs in soil and terrestrial ecosystems, more research interests have been drawn to plastic pollution in soil. Studies have shown that the soil environment receives much more plastic waste than the marine environment [18] and researchers have warned about the ecological effects of plastics and small plastic particles in soil and terrestrial environments [17, 19]. The main effects of MPs in soil are likely to occur at the interface between soil particles and plastic polymers (i.e., plastisphere). Similar to interactions in the plant rhizosphere, the physicochemical properties of MPs could stimulate the diversity and activities of soil microbial communities at the soil-plastic interface. These interactions can favor the proliferation and activities of specific microbial taxa, and lead to the formation of microbial hotspots in the soil [20]. With the increasing rate of MP contamination in most agricultural soils [21], the specific microbial niches at the soil-MP interface (i.e., microplastisphere) are of ecological importance. However, the effect of MPs on soil microbial communities and its corresponding impact on biodegradation in soil remains largely unclear.

Because of their physicochemical properties, which enhance their resistance to degradation, plastics can accumulate in natural ecosystems. High crystallinity, high molecular weight, and absence of functional groups that favor oxidative reaction processes are contributing factors to the non-biodegradability of plastics [22]. Current methods for managing plastic waste including incineration, landfilling, and recycling have associated demerits that could lead to further environmental issues. For instance, more toxic and volatile waste materials such as nitrogen oxides, furans, heavy metals, sulfides, and dioxins, which are considered to have potential carcinogenic effects, are produced when different synthetic plastics are incinerated [23]. Also, cost-ineffectiveness [24] and down-cycling [25] are undesirable consequences associated with recycling synthetic plastics. Landfilling, a widely used method for plastic waste disposal, especially in developing countries, results in a huge accumulation of plastic waste occupying a vast amount of land. Due to these, efforts are recently being made by researchers to explore other environmentally friendly and sustainable approaches to manage plastic waste and decrease environmental pollution caused by plastic waste. Microorganisms have been explored and reported as promising alternative for the degradation MPs. A number of research reports have identified different microbial species capable of degrading MPs. Although most reports available focused mainly on the biodegradation of a single kind of plastic such as PET [26], PE [27], PU [28], PS [29], and PP [30]. More recently, insects in their larva form including waxworm, mealworm, and superworm have also demonstrated the ability to eat, degrade, and mineralize various MPs, though not without the support of the microorganisms residing in their guts [31]. Research ideas focusing on the biodegradation of all main types of MPs are necessary as well as the biological upcycling of plastic waste [32].

Therefore, the aim of this study is to review previous literatures on the impact of MPs on soil microbial communities and biodegradation of MPs in soil. Also, extensively described are the current knowledge and knowledge gaps regarding the interactions between MPs and microbial communities in soil. Additionally, an effort has been made to outline the MP-related factors that affect the structural composition, diversity, and functionality of soil microbial communities. Finally, the challenges that need to be addressed in order to fill the knowledge gaps are itemized, along with future research prospects.

2. Sources of MP Pollution in the Natural Environment

Based on source points and their formation pathway, MPs in the environment can be categorized into two, namely; primary MPs and secondary MPs [8]. Primary MPs are synthesized from manufacturing activities and designed for commercial purposes, such as manufacture of personal care and cosmetic products (e.g., microbeads in hand and facial cleansers, toothpaste, shower gel), appliance manufacturing, industrial abrasives (e.g., air blasting), and textile fibers in clothing (e.g., acrylic fibers) [8, 33]. Primary MPs can be produced from the air-blasting industry as a result of the abrasion of materials during the process of preproduction of resin pellets [34]. Production of secondary MPs, on the other hand, is from physical (e.g., wave strike, abrasion, and water disturbance), chemical (e.g., UV radiation, the freeze-thaw cycle), and biological (e.g., biodegradation) activities, which involve the degradation and fragmentation of larger plastic materials into micro-sized particles [34].

The sources of MPs in the soil include vinyl mulch commonly used in agricultural activities [35], domestic sewage water containing MP beads from biosolids, personal care products, and fibers from clothing materials [36], landfills from industrial and urban centers [37], fertilizers [18], illegal waste dumping, irrigation with wastewater, littering roads and lake water flooding [38], atmospheric MP particles transported over long distances [39], and tire abrasion [40]. These MP particles settle on the soil surface and can penetrate into subsoils via physical and environmental activities. Recently, increasing attention has been drawn to MP pollution in the soil ecosystem from various pathways [41]. Studies have not revealed the transfer or presence of MPs in groundwater; however, there is growing concern over the potential distribution and transportation of MPs into the groundwater system and the hyporheic zone based on previous investigations on the mechanism of transport of MPs [38].

As more attention is drawn to MP pollution in the soil environment, it is necessary to understand the interaction and response of soil microbial communities to MPs in the soil and its effect on MP biodegradation. The process of microbial degradation of MPs in soil is governed by the diverse soil microbial communities that are able to colonize the MP surface, form biofilms, and establish a microenvironment that facilitates MP degradation [42] in the soil environment.

3. Microplastic Pollution and Soil Microbial Community

3.1. Microbial Interactions with MPs in Soil

Microorganisms interact with MPs in different environments, including the soil [43]. As more MPs are introduced in the soil, there is an increasing concern about their ecological effect, especially their effect on soil microbiota. For example, the worldwide use of plastic mulch films in agricultural activities increased from around 4.4 million tons in 2012 to about 7.4 million tons in 2019 [44], and this represents a significant source of MP contamination in the terrestrial environment [45]. Investigations have revealed that several groups of microorganisms including bacteria, fungi, and algae are found attached to MP surfaces [46, 47]. Previous studies have shown that different types of MPs can selectively stimulate the proliferation of specific soil microbial taxa in a microbial community (Table 1). For example, specific soil bacterial and fungal taxa were enriched when exposed to different plastic types (Table 1). It is interesting to note that, in the soil environment, the microbial richness and diversity on MP surfaces are unique and less diverse than those found colonizing natural materials such as wood [48] and the surrounding rhizospheric soil [49]. Due to their uniqueness, the microbial communities on MPs have been referred to as microplastisphere/plastisphere [50].

Response of Soil Microbial Communities to Different Plastics

A few studies have investigated the effect of MPs on the composition and diversity of soil microbial communities; however, their reports remain inconsistent. While some studies have reported that no significant effect was observed on soil microbial community structure [51–53], others reported considerable changes in the abundance and diversity of soil microbial communities exerted by MPs which affect the overall soil function [54]. Yi et al. [55] found that the alpha diversity and soil microbial communities changed significantly with membrane-like PE and fibrous PP, and this resulted in the abundance of Acidobacteria and Bacteroidetes, while Deinococcus-Thermus and Chloroflexi decreased in abundance. In another report, the substrate-induced respiration rate was reduced and significant changes in the root colonization rate of arbuscular mycorrhizal fungi were observed with the addition of MPs, and these suggest that the presence of MPs caused changes in microbial functions [53].

The soil treated with 0.007% low-density polyethylene (LDPE) resulted in three times higher specie turnover rate of bacterial communities than the untreated soil, and the divergence of soil microbial communities increased continually as the LDPE exposure time prolonged [56]. Ng et al. [57] also observed that the addition of LDPE (3% w/w) and PET (0.2% and 0.4% w/w) to the growth medium affected the even distribution and richness of soil microorganisms and soil microbial functions, and 14 unique bacterial genera were predominant and enriched. Liu et al. [58] reported that PP (7% and 28%) had a positive effect on the activities of soil microbiota, while de Souza Machado et al. [59] and Awet et al. [60] observed that polyester (0.05–0.4%), polyacrylic (0.05–0.4%) and PS (1 mg kg−1) showed significant negative effects on soil microbial activities. From these studies, it is quite difficult to draw a general conclusion on the effect of MPs on soil microbial communities as the types, sizes, shapes, and concentrations of MPs, as well as environmental conditions varied in these investigations. The different physical, chemical, and biological properties of the soil types used in these investigations might also have played a role in their findings. However, further research is required to gain deeper insight and validate the effect of co-interaction of soil properties and MP properties on soil microbial communities.

3.2. Factors Influencing the Interaction and Response of Soil Microbial Community to MPs

The structure, diversity, and richness of soil microbial communities can be significantly impacted by MP-related properties. The size of MPs might influence the microbial attachment and colonization as well as the distribution and diversity of soil microbial communities, although existing information on this remains scarce. For example, LDPE powder (150–250 μm) at 2% and 7% (w/w) showed differential tolerance on the bacterial diversity at the genus level [71], but LDPE particles (30 μm) at 0.2% (w/w) had no effect on the microbial community [72]. These findings corroborated the earlier report of Frère et al. [73] who found that MP size, regardless of the MP type, had no significant effect on the composition and diversity of microbial communities attached to them. These, however, contradict the report of Guo et al. [74] who found a significant increase in the richness of soil microbial communities in the presence of PE microfiber of <2 mm long. It was also observed that Actinobacteria was significantly enriched on PE, PS, and PP MPs when compared with their macroplastic forms [75]. These findings indicate that the MP size might play a role in microbial diversity and richness; however, more investigations are needed for a better understanding of the effect of MP size on soil microbial communities.

Apart from the size of MPs, the type of MPs could affect the composition, richness, and diversity of soil microbial communities. The soil samples treated with PE, PS, and PP of the same size (150 μm) and concentration (1% w/w) showed that the microbial communities of the soils treated with PP and PE responded differently from the PS-treated soil, although no major effect was observed in the bacterial diversity among the different MP types [76]. Similarly, the treatment of soil with LDPE particles (150–250 μm) and PS particles (0.33–0.64 μm) did not show significant changes in bacterial diversity [71, 77], whereas the treatment of soil with LDPE particles (678 μm) and PVC particles (15 μm) of the same concentration resulted in a decrease in the Shannon diversity indices of microbial communities [61]. Also, the increases in the relative abundances of Bacteroidetes and Gemmatimonadetes in the PP- and PE-treated soils were greater than that observed in the PS-treated soil [55]. The disparities in these findings can be attributed to the vast and robust microbial diversity in different soils with varying capacities to respond to disturbances [78] and responses of soil microorganisms to single artificial pressure are not always straightforward [79]. The hydrophobicity and roughness of MP surfaces are important parameters that could influence the microbial attachment and colonization of MPs and the resulting microbial communities [80]. Microorganisms colonizing MP surfaces are found to secret several extracellular polymeric substances (EPS), which act as bio-adhesives to enhance their attachment to MP surfaces [80]. Information on the relationship between MP surface hydrophobicity and soil microbial community and their interaction is still limited. However, it was reported that during the early colonization stage, marine bacteria adhere to the hydrophilic carrier interface for biofilm formation [81]. It was observed that hydrophilic groups such as C-O and C=O increased on PE surface in seawater, leading to a significant decrease in the MP hydrophobic properties [82].

As biofilms form and mature on MP surfaces, the bacterial hydrophobicity positively enhances the adhesion of the microbial communities to the MPs [83]. During the MP aging process, MP properties such as surface topography, polarity, surface area, and roughness could change [84]. These physical and structural changes could exert a direct influence on the composition and diversity of the associated microbial communities. For example, Betaproteobacteria was dominant on the smooth PS surface, while Gammaproteobacteria was dominant on the rough PE surface [85]. A rougher MP surface was found to have a positive correlation with the growth rate and density of the adhered microorganisms [83]. During the early biofilm colonization stage, Vibrio crassostreae colonized the smooth PS surface faster (<10 h) than the rough PS surface (6 d); however, rapid decolonization was also observed with the smooth PS surface (i.e., zero after 24 h) [86]. Furthermore, the surface charge on MPs plays a significant role on the abundance and diversity of microbial communities; plastic surface energy of 31–43 mN m−1 was discovered to be ideal for microbial colonization [87].

Plastic additives such as citrate esters, phthalate esters, fatty acid esters, glycerides, and polyhydric alcohols have the potential to leach out during the plastic weathering process [88], and their concentrations might increase in soil over time and cause significant changes to the abundance, diversity, and activities of soil microbial communities. Few studies have investigated the effect of plastic additives on soil microbial community and microcosm activity [89–91]. For instance, soil treated with diethyl phthalate (DEP) and di (2-ethyl hexyl) phthalate (DEHP) at 0.1 mg g−1 had no significant effect on the structural and functional diversity of the soil microbial community [91]. Also, the increasing dibutyl phthalate (DBP) concentration in soil decreased the bacterial diversity and the relative structure, composition, and abundance of soil bacterial communities reflected a successional change as DBP undergoes degradation [90]. Although plastic additives are not often observed to have a profound impact on soil microbiomes at concentrations of environmental relevance [91], more studies on the impact of different categories of plastic additives on soil microbial communities could throw more insight into understanding the microbial colonization pattern of MPs in the natural environment.

Other factors beyond MP-related properties could also play significant roles in the structural composition, diversity, and activities of soil microbial communities. Exposure duration of soil microorganisms to MPs could impact the composition and diversity of the microbial communities associated with MPs [92, 93]. Molecules adsorbed to MPs such as sugars, nucleic acids, proteins, fatty acids, and lipids have also been shown to improve the primary colonization process of microorganisms [94]. Environmental conditions such as nutrient availability (organic and inorganic nutrients), presence/absence of other pollutants, temperature, pH, light intensity, ionic strength, salinity, and biotic factors are critical to the composition, richness, diversity, and activities of microbial communities associated with MPs [48, 95]. Similarly, seasonal variations and geographical locations have been demonstrated to have significant influence on the diversity of microbial communities associated with MPs [95, 96]. So far, the mechanism of colonization of microbial communities on MPs is still largely unknown. When plastic is introduced to the soil, a series of physicochemical changes occur as a result of natural weathering activities [97], which could impact the soil microbial communities. Virgin plastics are normally smooth with a uniform structure and nearly no surface charge [98]. When exposed to sunlight, photooxidation occurs where solar photons with wavelengths within the UV and blue spectrum strike chromophores in the plastic polymer to initiate photooxidative decomposition [97]. The photooxidative weathering process creates radicals that are responsible for chain scissions in the plastic polymer and their reaction with oxygen oxidizes the polymer surface. This results in an increase in hydrophilic groups [30] and carbonyl groups that enhance the primary attachment and colonization of microorganisms on plastic surfaces for microbial community formation and biodegradation [87, 99, 100]. Plastic additives such as plasticizers, UV stabilizers, heat stabilizers, flame retardants, and pigment agents are also partially degraded during photooxidation [97] and might play a critical role in shaping the composition, abundance, and diversity of microbial communities associated with MPs in the soil [52, 99]. Similar to photooxidation, thermal weathering processes also generate radicals through chain scissions at high temperatures, which enhance microbial colonization and biofilm formation on MP surfaces, and biodegradation [101]. Also, MP fragmentation by mechanical stresses in the soil and biogeochemical attacks reduce the hydrophobicity, brittleness, and stiffness of MPs, thereby, enhancing microbial colonization and the formation of plastic-associated microbial communities [27, 97].

4. Microbial Degradation of MPs

Generally, synthetic plastics can resist degradation and persist in the environment for a long period of time. In the natural environment, the degradation of MPs is an integrated process that in corporates physical, chemical, and biological elements [102]. Despite MPs being less susceptible to degradation as compared to other biodegradable materials, a number of microbial strains have been identified with the potential to biodegrade MPs in the natural environment, including soils of plastic-dumping sites, waste of mulch films, marine water, crude oil-contaminated soils, landfills, sewage sludge, and guts of plastic-eating insects [103]. Microorganisms are found everywhere in nature, and they are able to adapt to a variety of environmental conditions, including extreme conditions. Their complex enzymatic system, ability to utilize various organic and inorganic substrates for growth, and ability to adapt to extreme environmental conditions are factors that contribute to their capacity to break down a wide variety of environmental contaminants. Therefore, they represent a sustainable and eco-friendly alternative to the management of plastic pollution in the environment. MPs form a novel ecological niche for microorganisms by providing a support system for colonization and growth, and a source of nutrient for microbial nutritional needs. The attachment of different groups of microorganisms (e.g. bacteria, actinomycetes, fungi, protists, algae, and viruses) to MP surfaces provides them an enabling environment to form biofilms [104]. The result of the microbial enzymatic activities leads to the structural deformation and loss of properties of the MPs [105]. Different categories of microorganisms capable of degrading MPs have been isolated from various environments [106]; however, there is still a knowledge gap about microbial interactions with MPs, which has contributed to the lesser adoption of plastic biodegradation strategies [107]. There are four stages involved in biofilm formation and biofilm-mediated MP degradation [104], and these include 1) microorganism adhesion to MP surface and modification of their surface properties, 2) MP deterioration, in which microbial enzymes speed up degradation and liberate monomers and additives from the MPs, 3) MP fragmentation, where the MPs lose their mechanical stability and become fragile as a result of microbial attack, and 4) assimilation of MPs, which involves the penetration of microbial filaments and water and the subsequent microbial decomposition and use of MPs as a nutrient source.

4.1. Actinomycetes in MPs Biodegradation

Actinomycetes, a group of gram-positive, filamentous bacteria, can be found in various ecological habitats including soil, plant tissues, freshwater, and marine environments [108]. Actinomycetes are recognized for their metabolic versatility and can perform different functions in the environment. They are explored for numerous biotechnological applications such as the production of enzymes, anticancer drugs, antibiotics, and other bioactive metabolites [109]. Some strains of actinomycetes have also been used for the bioremediation of toxic materials, breakdown of resistant carbohydrates (e.g. chitin, cellulose), and recycling of organic carbons [110]. Their ability to produce hydrolytic enzymes and bioactive metabolites enables them to grow on and degrade many complex polymers [111] including MPs [112]. Actinomycetes are known to produce extracellular biopolymers like levan, glycogen, dextran, and N-acetylglucosamine-rich slime polysaccharides, which are assumed to enhance their attachment to MP surfaces for subsequent microbial activities [113]. Actinomycetes such as the genera Streptomyces, Actinomadura, Rhodococcus, and thermophilic species of Thermoactinomycetes have been isolated from different environments with significant plastic degradation potential [112, 114]. Other actinomycetes species that have been reported to be associated with MP degradation are listed in Table 2. From the list, it is evident that limited actinomycetes genera have so far been identified to have MP-degradation potential, predominantly members of Streptomyces, Rhodococcus, and Micrococcus. Reports have also shown that some actinomycetes can form biofilm, similar to other bacterial strains, which is important for their survival and colonization on MPs [115].

Plastic Degradation by Actinomycetes, Biodegradation Assay Conditions, and Applied Detection Methods

4.2. Other Bacteria in MPs Biodegradation

Bacteria are the most abundant group of microbes. They are well recognized for their ubiquity and can be found in various environments such as water, soil, and atmosphere. Numerous bacterial species have been extensively studied for their roles in the biodegradation of complex polymers and bioremediation of environmental pollutants such as metal compounds, crude oil, antibiotics, plastic, and other compounds of environmental concern [136]. With their diversity and metabolic activities, bacterial strains are able to adsorb, desorb, and break down MPs [114, 137]. Different bacterial strains have been reported to utilize MPs as their main carbon source in minimal medium, and with their metabolic activities, they induced significant weight loss and changes to the morphological and chemical structure of MPs (Table 3).

Plastic degradation by bacteria, biodegradation assay conditions, and applied detection methods

Plastic-degrading bacteria have been isolated from different environments such as cold marine environment [138], dumpsites [139], landfills [140], recycling sites [141], and insects’ guts [142]. Most studies on the bacterial degradation of MPs have used pure bacterial cultures isolated from different environments or obtained from culture collections for the degradation of MPs under laboratory conditions. The use of pure bacterial strains can be advantageous when investigating specific metabolic pathways, evaluating different conditions, and/or closely monitoring the process of MP degradation [143]. However, in nature, many bacterial strains act in synergy, forming consortium and constituting a stable microbial community, which ultimately enhances their survival and degradation potential. So far, few studies have focused on MP biodegradation by bacterial consortia [129], it is, therefore, essential that more investigations be conducted to explore the potential of different bacterial consortia in MP biodegradation, as this might lead to greater efficiency due to the metabolic synergism between different bacterial strains.

4.3. Algae in MPs Biodegradation

In the last decades, algae have been extensively studied for their biotechnological applications, especially in the production of biofuels [155, 156]. The ability of different photosynthetic and heterotrophic algae to degrade environmental pollutants, both organic and inorganic, has also been well studied and established [157]. They are able to degrade environmental pollutants by adsorbing, accumulating, or metabolizing them into safer levels [156, 157]. Most studies on the environmental biodegradation of MPs have focused on the potentials of other groups of microorganisms such as bacteria and fungi, and only a few studies have investigated the potential of algae for biodegradation of MPs. Rather, recent studies have focused on the potential of algae in green plastic production [158].

Algal species including Oscillatoria, Spirogyra, Anabaena, Spirulina, and Chlorella have been reported to be found colonizing MP surfaces in terrestrial environments, but it is still inconclusive to confirm that these algal species are able to metabolize the MPs [159, 160]. In a study by Khoironi et al. [161], Spirulina sp. was able to biodegrade PP and PET during 112 d incubation. The authors reported that the tensile strength of PP and PET decreased by 0.1977 MPa d−1 and 0.9939 MPa d−1, respectively. Kumar et al. [162] also reported that Navicula pupula, Scenedesmus dimorphus, and Anabaena spiroides showed degradation potentials on both LDPE and high-density PE (HDPE). From their findings, A. spiroides was able to degrade 8.18% of LDPE after 30 d, representing the most promising algal strain in their study. The reported degradation rate is considerably lower than that of some investigations where bacteria and fungi were used to degrade PE. For instance, Brevibacillus borstelensis, a thermophilic bacteria isolated from soil, was able to degrade 30% of PE films after 30 d incubation [163]. A marine fungus, Zalerion maritumum, was similarly able to degrade 43% of PE pellets after 28 d incubation [164]. Generally, the low MP degradation rate by algae in comparison with bacteria and fungi is quite understandable. Unlike bacteria and fungi, most algae are considered to be photoautotrophic organisms that use atmospheric CO2 as their main source of carbon, and their main energy source is derived from sunlight [165]. Some algae are able to grow under heterotrophic conditions where external carbon and energy sources are utilized under dark conditions [166]. Even though they are able to colonize and assimilate MPs, algae are not metabolically inclined to mineralize MPs [162]; therefore, there is the possibility of plastic accumulation in algae which might be introduced to the food chain [167].

4.4. Fungi in MPs Biodegradation

Fungi are known to possess vast metabolic potentials, including the production of extracellular multienzyme complexes [168], thereby making them microorganisms of interest in MP biodegradation research. Different fungal strains can be found in different natural environments and they play significant roles in maintaining biogeochemical cycles and promoting the transformation of different substances [169]. As highlighted in Table 4, different fungal species and consortia have been reported with MP-degrading potential based on their ability to utilize MPs as their sole carbon or energy source, predominantly members of the genus Aspergillus, Penicillium, Fusarium, Trichoderma, Mucor, and Cladosporium. Some edible fungal species such as Agaricus bisporus, Pleurotus abalones, and Pleurotus ostreatus have also been reported to utilize PE and PS for growth, with changes in laccase activity [170]. Similar to bacteria, fungi are able to adhere to and utilize MPs [171]. Through their metabolic activities, they have the potential to decrease the hydrophobicity of MPs by promoting the formation of chemical bonds like ester, carbonyl, and carboxyl functional groups in MPs [104].

Plastic degradation by fungi, biodegradation assay conditions, and applied detection methods

5. Knowledge Gaps and Future Prospects

The majority of research on MP pollution and its ecological effects has focused on the marine environment and other aquatic ecosystems, and there have been limited investigations on MP pollution in soil thus far. Unlike water, the soil is a unique media with relatively complex and distinctive physical, chemical, and biological characteristics; as a result, it is challenging to investigate the ecological effect of MPs, particularly with regard to soil microorganisms, which are crucial to nutrient cycling and maintenance of soil functions. From this study, it is evident that, although research on MPs in soil is increasing as more attention is drawn to this area, there is still a substantial gap in understanding the interactions of soil microorganisms with MPs as well as the mechanisms by which microorganisms degrade MPs in the soil. The ecological effect of MPs on the structural composition, diversity, and activities of soil microbial communities are far from being fully understood. Therefore, future studies should focus on addressing the following issues. Firstly, it is crucial to monitor the sources and distribution of MPs in the soil. The extent of MP pollution under different land uses and natural environments should be understood. Also, further research is needed to better understand the ecological impact of MPs on soil microbiota and microbial communities. Critical questions to be answered with multiple lines of evidence would include i) what is the mechanism by which MPs affect soil microbial communities?, ii) how do MP properties affect the structural composition, richness, diversity, and activities of soil microbial communities in MP-polluted soils?, iii) would the co-interactions of MP properties and soil properties have a significant impact on soil microbial communities?, iv) do intermediates and products of MP degradation impact the diversity and activities of soil microbial communities?, v) what ecological impact do MP additives and adsorbed environmental pollutants have on the microbial communities in the soil?. Lastly, current literature revealed that information on microbial degradation of MPs has centered on specific groups of microorganisms of limited genera, majorly pure microbial cultures, with a few studies on microbial consortium. This amplifies the necessity to explore the potential of various microbes in their natural environments for MP degradation. As most microorganisms exhibit synergistic interactions in their natural environments, it is suggested that the use of different microbial strains in consortium will result in greater efficiency in MP degradation. Furthermore, the application of omic tools such as metagenomics, genomics, metabolomics, proteomics, and transcriptomics will aid in understanding the biological activities that take place at the genetic and metabolic levels, and the influence of environmental factors during MP biodegradation.

Acknowledgments

This study was funded by the National Research Foundation of Korea (NRF-2021R1A2C4001746).

Notes

Conflict of Interest

The authors declare that they have no conflict of interest.

Author contributions

K.C.O. (PhD student) wrote the manuscript. E.H.J. (Associate professor) wrote and revised the manuscript.

References

1. Chae Y, An YJ. Current research trends on plastic pollution and ecological impacts on the soil ecosystem: A review. Environ. Pollut 2018;240:387–395. https://doi.org/10.1016/j.envpol.2018.05.008 .
2. Amobonye A, Bhagwat P, Singh S, Pillai S. Plastic biodegradation: Frontline microbes and their enzymes. Sci. Total Environ 2021;759:1–16. https://doi.org/10.1016/j.scitotenv.2020.143536 .
3. Plastic Europe. Plastics - the Facts 2020 [Internet] Brussels: Plastics Europe; c2022. [cited 31 October 2022]. Available from: https://plasticseurope.org/knowledge-hub/plastics-thefacts-2020/ .
4. Nielsen T, Hasselbalch J, Holmberg K, Stripple J. Politics and the plastic crisis: A review throughout the plastic life cycle. WIREs Energy Environ 2019;9:1–18. https://doi.org/10.1002/wene.360 .
5. Jambeck JR, Geyer R, Wilcox C, et al. Marine pollution: Plastic waste inputs from land into the ocean. Science 2015;347:768–771. https://doi.org/10.1126/science.1260352 .
6. Van Cauwenberghe L, Devriese L, Galgani F, Robbens J, Janssen CR. Microplastics in sediments: A review of techniques, occurrence and effects. Mar. Environ. Res 2015;111:5–17. https://doi.org/10.1016/j.marenvres.2015.06.007 .
7. E erkes-Medrano D, Thompson RC, Aldridge DC. Microplastics in freshwater systems: a review of the emerging threats, identification of knowledge gaps and prioritisation of research needs. Water Res 2015;75:63–82. https://doi.org/10.1016/j.watres.2015.02.012 .
8. Duis K, Coors A. Microplastics in the aquatic and terrestrial environment: sources (with a specific focus on personal care products), fate and effects. Environ. Sci. Eur 2016;28:1–25. https://doi.org/10.1186/s12302-015-0069-y .
9. Ryan P, Moloney C. Plastic and other artefacts on South African beaches: Temporal trends in abundance and composition. S. Afr. J. Sci 1990;86:450–452.
10. Thompson RC, Olsen Y, Mitchell RP, et al. Lost at sea: where is all the plastic? Science 2004;304(5672):838. https://doi.org/10.1126/science.1094559 .
11. Alimi OS, Fadare OO, Okoffo ED. Microplastics in African ecosystems: Current knowledge, abundance, associated contaminants, techniques, and research needs. Sci. Total Environ 2021;755:1–42. https://doi.org/10.1016/j.scitotenv.2020.142422 .
12. Othman A, Abu Hasan H, Muhamad M, Ismail NI, Abdullah S. Microbial degradation of microplastics by enzymatic processes: A review. Environ. Chem. Lett 2021;19:1–17. https://doi.org/10.1007/s10311-021-01197-9 .
13. Monkul MM, Ozhan HO. Microplastic contamination in soils: A review from geotechnical engineering view. Polymers 2021;13:1–24. https://doi.org/10.3390/polym13234129 .
14. Hur J, Jho EH. Current research trends on the effects of microplastics in soil environment using earthworms: Mini-review. J. Korean Soc. Environ. Eng 2021;43:299–306. https://doi.org/10.4491/KSEE.2021.43.4.299 .
15. Zhu F, Zhu C, Wang C, Gu C. Occurrence and ecological impacts of microplastics in soil systems: A review. Bull. Environ. Contam. Toxicol 2019;102:741–749. https://doi.org/10.1007/s00128-019-02623-z .
16. Hasan MM, Jho EH. Effect of microplastics on the germination and growth of terrestrial plants. J. Korean Soc. Environ. Eng 2022;44:375–382. https://doi.org/10.4491/KSEE.2022.44.10.375 .
17. Rillig MC. Microplastic in terrestrial ecosystems and the soil? Environ. Sci. Technol 2012;46:6453–6454. https://doi.org/10.1021/es302011r .
18. Horton AA, Walton A, Spurgeon DJ, Lahive E, Svendsen C. Microplastics in freshwater and terrestrial environments: Evaluating the current understanding to identify the knowledge gaps and future research priorities. Sci. Total Environ 2017;586:127–141. https://doi.org/10.1016/j.scitotenv.2017.01.190 .
19. Rodriguez-Seijo A, Lourenço J, Rocha-Santos TAP, et al. Histopathological and molecular effects of microplastics in Eisenia andrei Bouché. Environ. Pollut 2017;220:495–503. https://doi.org/10.1016/j.envpol.2016.09.092 .
20. Kuzyakov Y, Blagodatskaya E. Microbial hotspots and hot moments in soil: Concept and review. Soil Biol. Biochem 2015;83:184–199. https://doi.org/10.1016/j.soilbio.2015.01.025 .
21. Steinmetz Z, Wollmann C, Schaefer M, et al. Plastic mulching in agriculture. Trading short-term agronomic benefits for long-term soil degradation? Sci. Total Environ 2016;550:690–705. https://doi.org/10.1016/j.scitotenv.2016.01.153 .
22. Rajendran S, Kannan V, Natarajan K, et al. The role of microbes in plastic degradation. In : Chandra R, ed. Environmental Waste Management 1st edth ed. Florida: CRC Press; 2015. p. 341–370.
23. Verma R, Shankarappa V, Papireddy M, Gowda ANS. Toxic pollutants from plastic waste-A review. Procedia Environ. Sci 2016;35:701–708. https://doi.org/10.1016/j.proenv.2016.07.069 .
24. Gradus R, Nillesen P, Dijkgraaf E, Koppen R. A cost-effectiveness analysis for incineration or recycling of Dutch household plastic waste. Ecol. Econ 2017;135:22–28. https://doi.org/10.1016/j.ecolecon.2016.12.021 .
25. Rahimi A, Garcia J. Chemical recycling of waste plastics for new materials production. Nat. Rev. Chem 2017;1:1–11. https://doi.org/10.1038/s41570-017-0046 .
26. T aniguchi I, Yoshida S, Hiraga K, Miyamoto K, Kimura Y, Oda K. Biodegradation of PET: Current status and application aspects. ACS Catal 2019;9:4089–4105. https://doi.org/10.1021/acscatal.8b05171 .
27. Restrepo-Flórez J-M, Bassi A, Thompson M. Microbial degradation and deterioration of polyethylene – A review. Int. Biodeterior. Biodegradation 2014;88:83–90. https://doi.org/10.1016/j.ibiod.2013.12.014 .
28. Magnin A, Pollet E, Phalip V, Avérous L. Evaluation of biological degradation of polyurethanes. Biotechnol. Adv 2020;39:1–24. https://doi.org/10.1016/j.biotechadv.2019.107457 .
29. Ho BT, Roberts TK, Lucas S. An overview on biodegradation of polystyrene and modified polystyrene: the microbial approach. Crit. Rev. Biotechnol 2018;38:308–320. https://doi.org/10.1080/07388551.2017.1355293 .
30. Arutchelvi J, Muniyasamy S, Arkatkar A, Doble M, Bhaduri S, Uppara P. Biodegradation of polyethylene and polypropylene. Indian J. Biotechnol 2008;7:9–22.
31. Zhang J, Gao D, Li Q, et al. Biodegradation of polyethylene microplastic particles by the fungus Aspergillus flavus from the guts of wax moth Galleria mellonella . Sci. Total Environ 2020;704:1–8. https://doi.org/10.1016/j.scitotenv.2019.135931 .
32. Blank LM, Narancic T, Mampel J, Tiso T, O’Connor K. Biotechnological upcycling of plastic waste and other non-conventional feedstocks in a circular economy. Curr. Opin. Biotechnol 2020;62:212–219. https://doi.org/10.1016/j.copbio.2019.11.011 .
33. Cincinelli A, Martellini T, Guerranti C, Scopetani C, Chelazzi D, Giarrizzo T. A potpourri of microplastics in the sea surface and water column of the Mediterranean Sea. TrAC Trends Anal. Chem 2019;110:321–326. https://doi.org/10.1016/j.trac.2018.10.026 .
34. Khalid N, Aqeel M, Noman A, et al. Linking effects of microplastics to ecological impacts in marine environments. Chemosphere 2021;264:1–13. https://doi.org/10.1016/j.chemosphere.2020.128541 .
35. Farmer J, Zhang B, Jin X, Zhang P, Wang J. Long-term effect of plastic film mulching and fertilization on bacterial communities in a brown soil revealed by high through-put sequencing. Arch. Agron. Soil Sci 2016;63:230–241. https://doi.org/10.1080/03650340.2016.1193667 .
36. Carr SA, Liu J, Tesoro AG. Transport and fate of microplastic particles in wastewater treatment plants. Water Res 2016;91:174–182. https://doi.org/10.1016/j.watres.2016.01.002 .
37. Nizzetto L, Bussi G, Futter MN, Butterfield D, Whitehead PG. A theoretical assessment of microplastic transport in river catchments and their retention by soils and river sediments. Environ. Sci: Processes Impacts 2016;18:1050–1059. https://doi.org/10.1039/c6em00206d .
38. Bläsing M, Amelung W. Plastics in soil: Analytical methods and possible sources. Sci. Total Environ 2018;612:422–435. https://doi.org/10.1016/j.scitotenv.2017.08.086 .
39. Dris R, Gasperi J, Saad M, Mirande C, Tassin B. Synthetic fibers in atmospheric fallout: A source of microplastics in the environment? Mar. Pollut. Bull 2016;104:290–293. https://doi.org/10.1016/j.marpolbul.2016.01.006 .
40. Foitzik M-J, Unrau H-J, Gauterin F, Dörnhöfer J, Koch T. Investigation of ultra fine particulate matter emission of rubber tires. Wear 2018;394–395:87–95. https://doi.org/10.1016/j.wear.2017.09.023 .
41. Ng EL, Huerta Lwanga E, Eldridge SM, et al. An overview of microplastic and nanoplastic pollution in agroecosystems. Sci. Total Environ 2018;627:1377–1388. https://doi.org/10.1016/j.scitotenv.2018.01.341 .
42. Debroy A, George N, Mukherjee G. Role of biofilms in the degradation of microplastics in aquatic environments. J. Chem. Technol. Biotechnol 2021;97:3271–3282. https://doi.org/https://doi.org/10.1002/jctb.6978 .
43. Mammo FK, Amoah ID, Gani KM, et al. Microplastics in the environment: Interactions with microbes and chemical contaminants. Sci. Total Environ 2020;743:1–21. https://doi.org/10.1016/j.scitotenv.2020.140518 .
44. Batista T, Cansado IP, Tita B, et al. Dealing with plastic waste from agriculture activity. Agronomy 2022;12:1–12. https://doi.org/10.3390/agronomy12010134 .
45. Huang Y, Liu Q, Jia W, Yan C, Wang J. Agricultural plastic mulching as a source of microplastics in the terrestrial environment. Environ. Pollut 2020;260:1–6. https://doi.org/10.1016/j.envpol.2020.114096 .
46. Masó M, Garcés E, Pagès F, Camp J. Drifting plastic debris as a potential vector for dispersing Harmful Algal Bloom (HAB) species. Sci. Mar 2003;67:107–111. https://doi.org/10.3989/scimar.2003.67n1107 .
47. De Tender C, Devriese LI, Haegeman A, et al. Temporal dynamics of bacterial and fungal colonization on plastic debris in the North sea. Environ. Sci. Technol 2017;51:7350–7360. https://doi.org/10.1021/acs.est.7b00697 .
48. Oberbeckmann S, Kreikemeyer B, Labrenz M. Environmental factors support the formation of specific bacterial assemblages on microplastics. Front. Microbiol 2017;8:1–12. https://doi.org/10.3389/fmicb.2017.02709 .
49. Yu H, Zhang Y, Tan W. The “neighbor avoidance effect” of microplastics on bacterial and fungal diversity and communities in different soil horizons. Environ. Sci. Ecotechnol 2021;8:1–9. https://doi.org/https://doi.org/10.1016/j.ese.2021.100121 .
50. Zettler ER, Mincer TJ, Amaral-Zettler LA. Life in the “plastisphere”: Microbial communities on plastic marine debris. Environ. Sci. Technol 2013;47:7137–7146. https://doi.org/10.1021/es401288x .
51. Huang Y, Zhao Y, Wang J, Zhang M, Jia W, Qin X. LDPE microplastic films alter microbial community composition and enzymatic activities in soil. Environ. Pollut 2019;254:1–10. https://doi.org/10.1016/j.envpol.2019.112983 .
52. Yan Y, Chen Z, Zhu F, Zhu C, Wang C, Gu C. Effect of polyvinyl chloride microplastics on bacterial community and nutrient status in two agricultural soils. Bull. Environ. Contam. Toxicol 2021;107:602–609. https://doi.org/10.1007/s00128-020-02900-2 .
53. Judy JD, Williams M, Gregg A, et al. Microplastics in municipal mixed-waste organic outputs induce minimal short to long-term toxicity in key terrestrial biota. Environ. Pollut 2019;252:522–531. https://doi.org/10.1016/j.envpol.2019.05.027 .
54. Gao B, Yao H, Li Y, Zhu Y. Microplastic addition alters the microbial community structure and stimulates soil carbon dioxide emissions in vegetable-growing soil. Environ. Toxicol. Chem 2021;40:352–365. https://doi.org/10.1002/etc.4916 .
55. Yi M, Zhou S, Zhang L, Ding S. The effects of three different microplastics on enzyme activities and microbial communities in soil. Water Environ. Res 2021;93:24–32. https://doi.org/10.1002/wer.1327 .
56. Wang J, Huang M, Wang Q, Sun Y, Zhao Y, Huang Y. LDPE microplastics significantly alter the temporal turnover of soil microbial communities. Sci. Total Environ 2020;726:1–8. https://doi.org/10.1016/j.scitotenv.2020.138682 .
57. Ng EL, Lin SY, Dungan AM, et al. Microplastic pollution alters forest soil microbiome. J. Hazard. Mater 2021;409:1–10. https://doi.org/10.1016/j.jhazmat.2020.124606 .
58. Liu H, Yang X, Liu G, et al. Response of soil dissolved organic matter to microplastic addition in Chinese loess soil. Chemosphere 2017;185:907–917. https://doi.org/10.1016/j.chemosphere.2017.07.064 .
59. de Souza Machado AA, Lau CW, Till J, et al. Impacts of microplastics on the soil biophysical environment. Environ. Sci. Technol 2018;52:9656–9665. https://doi.org/10.1021/acs.est.8b02212 .
60. Tsegai A, Kohl Y, Meier F, et al. Effects of polystyrene nanoparticles on the microbiota and functional diversity of enzymes in soil. Environ. Sci. Eur 2018;30:1–10. https://doi.org/10.1186/s12302-018-0140-6 .
61. Fei Y, Huang S, Zhang H, et al. Response of soil enzyme activities and bacterial communities to the accumulation of microplastics in an acid cropped soil. Sci. Total Environ 2020;707:1–9. https://doi.org/10.1016/j.scitotenv.2019.135634 .
62. Zhang X, Li Y, Lei J, et al. Time-dependent effects of microplastics on soil bacteriome. J. Hazard. Mater 2023;447:1–13. https://doi.org/https://doi.org/10.1016/j.jhazmat.2023.130762 .
63. Wang Q, Feng X, Liu Y, et al. Effects of microplastics and carbon nanotubes on soil geochemical properties and bacterial communities. J. Hazard. Mater 2022;433:1–13. https://doi.org/https://doi.org/10.1016/j.jhazmat.2022.128826 .
64. Dong Y, Gao M, Qiu W, Song Z. Effect of microplastics and arsenic on nutrients and microorganisms in rice rhizosphere soil. Ecotoxicol. Environ. Saf 2021;211:1–12. https://doi.org/https://doi.org/10.1016/j.ecoenv.2021.111899 .
65. Fan P, Tan W, Yu H. Effects of different concentrations and types of microplastics on bacteria and fungi in alkaline soil. Ecotoxicol. Environ. Saf 2022;229:1–9. https://doi.org/https://doi.org/10.1016/j.ecoenv.2021.113045 .
66. Li H-Q, Shen Y-J, Wang W-L, Wang H-T, Li H, Su J-Q. Soil pH has a stronger effect than arsenic content on shaping plastisphere bacterial communities in soil. Environ. Pollut 2021;287:1–8. https://doi.org/https://doi.org/10.1016/j.envpol.2021.117339 .
67. Qian H, Zhang M, Liu G, et al. Effects of soil residual plastic film on soil microbial community structure and fertility. Water Air Soil Pollut 2018;229:1–11. https://doi.org/10.1007/s11270-018-3916-9 .
68. Ren X, Tang J, Liu X, Liu Q. Effects of microplastics on greenhouse gas emissions and the microbial community in fertilized soil. Environ. Pollut 2020;256:1–11. https://doi.org/https://doi.org/10.1016/j.envpol.2019.113347 .
69. Temporiti ME, Nicola L, Girometta CE, Roversi A, Daccò C, Tosi S. The analysis of the mycobiota in plastic polluted soil reveals a reduction in metabolic ability. J. Fungi 2022;8:1–20. https://doi.org/10.3390/jof8121247 .
70. Zhou Y, Sun Y, Liu J, Ren X, Zhang Z, Wang Q. Effects of microplastics on humification and fungal community during cow manure composting. Sci. Total Environ 2022;803:1–10. https://doi.org/https://doi.org/10.1016/j.scitotenv.2021.150029 .
71. Rong L, Zhao L, Zhao L, et al. LDPE microplastics affect soil microbial communities and nitrogen cycling. Sci. Total Environ 2021;773:1–11. https://doi.org/10.1016/j.scitotenv.2021.145640 .
72. Li HZ, Zhu D, Lindhardt JH, Lin SM, Ke X, Cui L. Long-term fertilization history alters effects of microplastics on soil properties, microbial communities, and functions in diverse farmland ecosystem. Environ. Sci. Technol 2021;55:4658–4668. https://doi.org/10.1021/acs.est.0c04849 .
73. Frère L, Maignien L, Chalopin M, et al. Microplastic bacterial communities in the Bay of Brest: Influence of polymer type and size. Environ. Pollut 2018;242:614–625. https://doi.org/10.1016/j.envpol.2018.07.023 .
74. Guo QQ, Xiao MR, Ma Y, Niu H, Zhang GS. Polyester microfiber and natural organic matter impact microbial communities, carbon-degraded enzymes, and carbon accumulation in a clayey soil. J. Hazard. Mater 2021;405:1–10. https://doi.org/10.1016/j.jhazmat.2020.124701 .
75. Zhang K, Xiong X, Hu H, et al. Occurrence and characteristics of microplastic pollution in Xiangxi Bay of Three Gorges Reservoir, China. Environ. Sci. Technol 2017;51:3794–3801. https://doi.org/10.1021/acs.est.7b00369 .
76. Sun Y, Duan C, Cao N, et al. Effects of microplastics on soil microbiome: The impacts of polymer type, shape, and concentration. Sci. Total Environ 2022;806:1–11. https://doi.org/https://doi.org/10.1016/j.scitotenv.2021.150516 .
77. Xu M, Du W, Ai F, et al. Polystyrene microplastics alleviate the effects of sulfamethazine on soil microbial communities at different CO2 concentrations. J. Hazard. Mater 2021;413:1–11. https://doi.org/https://doi.org/10.1016/j.jhazmat.2021.125286 .
78. Lee S-H, Sorensen JW, Grady KL, Tobin TC, Shade A. Divergent extremes but convergent recovery of bacterial and archaeal soil communities to an ongoing subterranean coal mine fire. ISME J 2017;11:1447–1459. https://doi.org/10.1038/ismej.2017.1 .
79. Rillig M, Ryo M, Lehmann A, et al. The role of multiple global change factors in driving soil functions and microbial biodiversity. Science 2019;366:886–890. https://doi.org/10.1126/science.aay2832 .
80. Ogonowski M, Motiei A, Ininbergs K, et al. Evidence for selective bacterial community structuring on microplastics. Environ. Microbiol 2018;20(8):2796–2808. https://doi.org/10.1111/1462-2920.14120 .
81. Pinto M, Langer TM, Hüffer T, Hofmann T, Herndl GJ. The composition of bacterial communities associated with plastic biofilms differs between different polymers and stages of biofilm succession. PLoS One 2019;14:1–20. https://doi.org/10.1371/journal.pone.0217165 .
82. Tu C, Chen T, Zhou Q, et al. Biofilm formation and its influences on the properties of microplastics as affected by exposure time and depth in the seawater. Sci. Total Environ 2020;734:1–9. https://doi.org/10.1016/j.scitotenv.2020.139237 .
83. Pompilio A, Piccolomini R, Picciani C, D’Antonio D, Savini V, Di Bonaventura G. Factors associated with adherence to and biofilm formation on polystyrene by Stenotrophomonas maltophilia: the role of cell surface hydrophobicity and motility. FEMS Microbiol. Lett 2008;287:41–47. https://doi.org/10.1111/j.1574-6968.2008.01292.x .
84. Mao R, Lang M, Yu X, Wu R, Yang X, Guo X. Aging mechanism of microplastics with UV irradiation and its effects on the adsorption of heavy metals. J. Hazard. Mater 2020;393:1–9. https://doi.org/https://doi.org/10.1016/j.jhazmat.2020.122515 .
85. Parrish K, Fahrenfeld N. Microplastic biofilm in fresh- and wastewater as a function of microparticle type and size class. Environ. Sci.: Water Res. Technol 2019;5:1–25. https://doi.org/10.1039/C8EW00712H .
86. Foulon V, Le Roux F, Lambert C, Huvet A, Soudant P, Paul-Pont I. Colonization of polystyrene microparticles by Vibrio crassostreae: Light and electron microscopic investigation. Environ. Sci. Technol 2016;50:10988–10996. https://doi.org/10.1021/acs.est.6b02720 .
87. Nauendorf A, Krause S, Bigalke NK, et al. Microbial colonization and degradation of polyethylene and biodegradable plastic bags in temperate fine-grained organic-rich marine sediments. Mar. Pollut. Bull 2016;103:168–178. https://doi.org/10.1016/j.marpolbul.2015.12.024 .
88. Hahladakis JN, Velis CA, Weber R, Iacovidou E, Purnell P. An overview of chemical additives present in plastics: Migration, release, fate and environmental impact during their use, disposal and recycling. J. Hazard. Mater 2018;344:179–199. https://doi.org/10.1016/j.jhazmat.2017.10.014 .
89. Kong S, Ji Y, Liu L, et al. Diversities of phthalate esters in suburban agricultural soils and wasteland soil appeared with urbanization in China. Environ. Pollut 2012;170:161–168. https://doi.org/10.1016/j.envpol.2012.06.017 .
90. Kong X, Jin D, Jin S, et al. Responses of bacterial community to dibutyl phthalate pollution in a soil-vegetable ecosystem. J. Hazard. Mater 2018;353:142–150. https://doi.org/10.1016/j.jhazmat.2018.04.015 .
91. Cartwright C, Thompson I, Burns R. Degradation and impact of phthalate plasticizers on soil microbial communities. Environ. Toxicol. Chem 2000;19:1253–1261. https://doi.org/10.1002/etc.5620190506 .
92. Kirstein IV, Wichels A, Gullans E, Krohne G, Gerdts G. The Plastisphere - Uncovering tightly attached plastic “specific” microorganisms. PLoS One 2019;14:1–17. https://doi.org/10.1371/journal.pone.0215859 .
93. Xu X, Wang S, Gao F, et al. Marine microplastic-associated bacterial community succession in response to geography, exposure time, and plastic type in China’s coastal seawaters. Mar. Pollut. Bull 2019;145:278–286. https://doi.org/10.1016/j.marpolbul.2019.05.036 .
94. Keswani A, Oliver DM, Gutierrez T, Quilliam RS. Microbial hitchhikers on marine plastic debris: Human exposure risks at bathing waters and beach environments. Mar. Environ. Res 2016;118:10–19. https://doi.org/10.1016/j.marenvres.2016.04.006 .
95. Kesy K, Oberbeckmann S, Kreikemeyer B, Labrenz M. Spatial environmental heterogeneity determines young biofilm assemblages on microplastics in Baltic Sea mesocosms. Front. Microbiol 2019;10:1–18. https://doi.org/10.3389/fmicb.2019.01665 .
96. Curren E, Leong SCY. Profiles of bacterial assemblages from microplastics of tropical coastal environments. Sci. Total Environ 2019;655:313–320. https://doi.org/10.1016/j.scitotenv.2018.11.250 .
97. Pickett JE. Weathering of Plastics. In : Kutz M, ed. Handbook of environmental degradation of materials 3rd edth ed. Oxford. William Andrew Publishing: 2018. p. 163–184.
98. Fotopoulou KN, Karapanagioti HK. Surface properties of beached plastics. Environ. Sci. Pollut. Res 2015;22:11022–11032. https://doi.org/10.1007/s11356-015-4332-y .
99. Wei R, Zimmermann W. Microbial enzymes for the recycling of recalcitrant petroleum-based plastics: how far are we? Microb. Biotechnol 2017;10:1308–1322. https://doi.org/10.1111/1751-7915.12710 .
100. Dong M, Zhang Q, Xing X, Chen W, She Z, Luo Z. Raman spectra and surface changes of microplastics weathered under natural environments. Sci. Total Environ 2020;739:1–9. https://doi.org/https://doi.org/10.1016/j.scitotenv.2020.139990 .
101. Volke T, Saucedo-Castañeda G, Gutiérrez-Rojas M, Manzur A, Favela-Torres E. Thermally treated low density polyethylene biodegradation by Penicillium pinophilum and Aspergillus niger . J. Appl. Polym. Sci 2001;83:305–314. https://doi.org/10.1002/app.2245 .
102. Ammalaa A, Batemana S, Deana K, et al. An overview of degradable and biodegradable polyolefins. Prog. Polym. Sci 2010;36:1015–1049. https://doi.org/10.1016/j.progpolymsci.2010.12.002 .
103. Krueger M, Harms H, Schlosser D. Prospects for microbiological solutions to environmental pollution with plastics. Appl. Microbiol. Biotechnol 2015;99:8857–8874. https://doi.org/10.1007/s00253-015-6879-4 .
104. Yuan J, Ma J, Sun Y, Zhou T, Zhao Y, Yu F. Microbial degradation and other environmental aspects of microplastics/plastics. Sci. Total Environ 2020;715:1–9. https://doi.org/10.1016/j.scitotenv.2020.136968 .
105. Miao L, Wang P, Hou J, et al. Distinct community structure and microbial functions of biofilms colonizing microplastics. Sci. Total Environ 2019;650:2395–2402. https://doi.org/10.1016/j.scitotenv.2018.09.378 .
106. Park SY, Kim CG. Biodegradation of micro-polyethylene particles by bacterial colonization of a mixed microbial consortium isolated from a landfill site. Chemosphere 2019;222:527–533. https://doi.org/10.1016/j.chemosphere.2019.01.159 .
107. Alshehrei F. Biodegradation of synthetic and natural plastic by microorganisms. J. Appl. Environ. Microbiol 2017. 58–19. http://pubs.sciepub.com/jaem/5/1/2 .
108. Valan Arasu M, Asha K, Duraipandiyan V, Ignacimuthu S, Agastian P. Characterization and phylogenetic analysis of novel polyene type antimicrobial metabolite producing actinomycetes from marine sediments: Bay of Bengal India. Asian Pac. J. Trop. Biomed 2012;2:803–810. https://doi.org/10.1016/S2221-1691(12)60233-0 .
109. Omidoyin KC, Femi-Ola TO. Isolation and screening of Streptomyces spp from soil samples of Ekiti State University Nigeria for antibacterial activity. Int. J. Res. Innovation Entrepreneurship 2020;1:48–54. https://doi.org/10.15294/ijrie.v1i1 .
110. Gohain A, Manpoong C, Saikia R, De Mandal S. Actinobacteria: diversity and biotechnological applications. In : De Mandal S, Bhatt P, eds. Recent Advancements in Microbial Diversity Amsterdam: Academic Press; 2020. p. 217–231.
111. Sharma M, Dangi P, Choudhary M. Actinomycetes: Source, identification, and their applications. Int. J. Curr. Microbiol. Appl. Sci 2014;3:801–832.
112. Jabloune R, Khalil M, Ben Moussa IE, et al. Enzymatic degradation of p-Nitrophenyl esters, polyethylene terephthalate, cutin, and suberin by Sub1, a suberinase encoded by the plant pathogen Streptomyces scabies . Microbes Environ 2020;35:1–7. https://doi.org/10.1264/jsme2.ME19086 .
113. Petar P, Beaman B, Ravalison M, Boiron P, Rodriguez-Nava V. Nocardia and Actinomyces. In : Yi-Wei T, Andrew S, eds. Molecular Medical Microbiology 2nd ed. Amsterdam: Academic Press; 2015. p. 731–752.
114. Auta HS, Emenike CU, Jayanthi B, Fauziah SH. Growth kinetics and biodeterioration of polypropylene microplastics by Bacillus sp. and Rhodococcus sp. isolated from mangrove sediment. Mar. Pollut. Bull 2018;127:15–21. https://doi.org/10.1016/j.marpolbul.2017.11.036 .
115. Gilan I, Sivan A. Effect of proteases on biofilm formation of the plastic-degrading actinomycete Rhodococcus ruber C208. FEMS Microbiol. Lett 2013;342:18–23. https://doi.org/10.1111/1574-6968.12114 .
116. Sivan A, Szanto M, Pavlov V. Biofilm development of the polyethylene-degrading bacterium Rhodococcus ruber . Appl. Microbiol. Biotechnol 2006;72:346–352. https://doi.org/10.1007/s00253-005-0259-4 .
117. Farzi A, Dehnad A, Fotouhi A. Biodegradation of polyethylene terephthalate waste using Streptomyces species and kinetic modeling of the process. Biocatal. Agric. Biotechnol 2018;17:25–31. https://doi.org/10.1016/j.bcab.2018.11.002 .
118. Bonhomme S, Cuer A, Delort A-M, Lemaire J, Sancelme M, Scott G. Environmental biodegradation of polyethylene. Polym. Degrad. Stab 2003;81:441–452. https://doi.org/10.1016/S0141-3910(03)00129-0 .
119. Kathiresan K. Polythene and plastic-degrading microbes in an Indian mangrove soil. Rev. Biol. Trop 2003;51:629–633.
120. Nowak B, Pająk J, Drozd-Bratkowicz M, Rymarz G. Microorganisms participating in the biodegradation of modified polyethylene films in different soils under laboratory conditions. Int. Biodeterior. Biodegradation 2011;65:757–767. https://doi.org/10.1016/j.ibiod.2011.04.007 .
121. Farzi A, Dehnad A, Shirzad N, Norouzifard F. Biodegradation of high density polyethylene using Streptomyces species. J. Coastal Life Med 2017;5:474–479. https://doi.org/10.12980/jclm.5.2017J7-94 .
122. Rajandas H, Parimannan S, Sathasivam K, Ravichandran M, Lee S. A novel FTIR-ATR spectroscopy based technique for the estimation of low-density polyethylene biodegradation. Polym. Test 2012;31:1094–1099. https://doi.org/10.1016/j.polymertesting.2012.07.015 .
123. Midhun K, Kalyani L, Guntuku G, Dandu S. Biodegradation of low density polyethylene (LDPE) by a new biosurfactant-producing thermophilic Streptomyces coelicoflavus NBRC 15399T. Afr. J. Biotechnol 2015;14:327–340. https://doi.org/10.5897/AJB2014.14224 .
124. Pometto AL, Lee BT, Johnson KE. Production of an extracellular polyethylene-degrading enzyme(s) by Streptomyces species. Appl. Environ. Microbiol 1992;58:731–733. https://doi.org/10.1128/aem.58.2.731-733.1992 .
125. Abraham J, Ghosh E, Mukherjee P, Gajendiran A. Microbial degradation of low density polyethylene. Environ. Prog. Sustain. Energy 2016;36:1–8. https://doi.org/10.1002/ep.12467 .
126. Mor R, Sivan A. Biofilm formation and partial biodegradation of polystyrene by the actinomycete Rhodococcus ruber: biodegradation of polystyrene. Biodegradation 2008;19:851–858. https://doi.org/10.1007/s10532-008-9188-0 .
127. Fontanella S, Bonhomme S, Brusson J-M, et al. Comparison of biodegradability of various polypropylene films containing additives based on Mn, Mn/Fe or Co. Polym. Degrad. Stab 2013;98:875–884. https://doi.org/10.1016/j.polymdegradstab.2013.01.002 .
128. Usha R, Sangeetha T, Muthusamy P. Screening of polyethylene degrading microorganisms from garbage soil. Libyan Agric. Res. Cen. J. Int 2011;2:200–2014.
129. Shah A, Hasan F, Akhter J, Hameed A, Ahmed S. Degradation of polyurethane by novel bacterial consortium isolated from soil. Ann. Microbiol 2008;58:381–386. https://doi.org/10.1007/BF03175532 .
130. El-Shafei H, Alborki AN, Kansoh A, Ali A. Biodegradation pf disposable polyethylene by fungi Streptomyces species. Polym. Degrad. Stab 1998;62:361–365. https://doi.org/10.1016/S0141-3910(98)00019-6 .
131. Jarerat A, Tokiwa Y, Tanaka H. Production of poly(L-lactide)-degrading enzyme by Amycolatopsis orientalis for biological recycling of poly(L-lactide). Appl. Microbiol. Biotechnol 2006;72:726–731. https://doi.org/10.1007/s00253-006-0343-4 .
132. Jarerat A, Tokiwa Y. Poly(L-lactide) degradation by Saccharothrix waywayandensis . Biotechnol. Lett 2003;25:401–404. https://doi.org/10.1023/a:1022450431193 .
133. Jarerat A, Tokiwa Y, Tanaka H. Poly(L-lactide) degradation by Kibdelosporangium aridum . Biotechnol. Lett 2003;25:2035–2038. https://doi.org/10.1023/b:bile.0000004398.38799.29 .
134. Pranamuda H, Tokiwa Y, Tanaka H. Polylactide Degradation by an Amycolatopsis sp. Appl. Environ. Microbiol 1997;63:1637–1640. https://doi.org/10.1128/aem.63.4.1637-1640.1997 .
135. Sukkhum S, Tokuyama S, Kitpreechavanich V. Development of fermentation process for PLA-degrading enzyme production by a new thermophilic Actinomadura sp. T16–1. Biotechnol. Bioprocess Eng 2009;14:302–306. https://doi.org/10.1007/s12257-008-0207-0 .
136. Bakir A, Rowland SJ, Thompson RC. Enhanced desorption of persistent organic pollutants from microplastics under simulated physiological conditions. Environ. Pollut 2014;185:16–23. https://doi.org/10.1016/j.envpol.2013.10.007 .
137. Auta HS, Emenike CU, Fauziah SH. Screening of Bacillus strains isolated from mangrove ecosystems in Peninsular Malaysia for microplastic degradation. Environ. Pollut 2017;231:1552–1559. https://doi.org/10.1016/j.envpol.2017.09.043 .
138. Urbanek AK, Rymowicz W, Mirończuk AM. Degradation of plastics and plastic-degrading bacteria in cold marine habitats. Appl. Microbiol. Biotechnol 2018;102:7669–7678. https://doi.org/10.1007/s00253-018-9195-y .
139. Muhonja CN, Makonde H, Magoma G, Imbuga M. Biodegradability of polyethylene by bacteria and fungi from Dandora dumpsite Nairobi-Kenya. PLoS One 2018;13:1–17. https://doi.org/10.1371/journal.pone.0198446 .
140. Gaytán I, Sánchez-Reyes A, Burelo M, et al. Degradation of recalcitrant polyurethane and xenobiotic additives by a selected landfill microbial community and its biodegradative potential revealed by proximity ligation-based metagenomic analysis. Front. Microbiol 2019;10:1–19. https://doi.org/10.3389/fmicb.2019.02986 .
141. Yoshida S, Hiraga K, Takehana T, et al. A bacterium that degrades and assimilates poly(ethylene terephthalate). Science 2016;351:1196–1199. https://doi.org/10.1126/science.aad6359 .
142. Ren L, Men L, Zhang Z, et al. Biodegradation of polyethylene by Enterobacter sp. D1 from the guts of wax moth Galleria mellonella . Int. J. Environ. Res. Public Health 2019;16:1–11. https://doi.org/10.3390/ijerph16111941 .
143. Janssen PH, Yates PS, Grinton BE, Taylor PM, Sait M. Improved culturability of soil bacteria and isolation in pure culture of novel members of the divisions Acidobacteria, Actinobacteria, Proteobacteria, and Verrucomicrobia. Appl. Environ. Microbiol 2002;68:2391–2396. https://doi.org/10.1128/aem.68.5.2391-2396.2002 .
144. Mahdi M, Ameen R, Ibrahim H. Study on degradation of nylon 6 by thermophilic bacteria Anoxybacillus rupiensis Ir3 (JQ912241). Int. J. Adv. Res. Biol. Sci 2016;3:200–209. http://dx.doi.org/10.22192/ijarbs.2016.03.09.027 .
145. Ambika Devi K, Lakshmi BKM, Hemalatha KPJ. Degradation of low density polythene by Achromobacter denitrificans strain S1, a novel marine isolate. Int. J. Rec. Sci. Res 2015;6:5454–5464.
146. Yang J, Yang Y, Wu WM, 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. https://doi.org/10.1021/es504038a .
147. Pramila R, Ramesh KV. Potential biodegradation of low density polyethylene (LDPE) by Acinetobacter baumannii . Afr. J. Bacteriol. Res 2015;7:24–28. https://doi.org/10.5897/JBR2015.0152 .
148. Syranidou E, Karkanorachaki K, Amorotti F, et al. Development of tailored indigenous marine consortia for the degradation of naturally weathered polyethylene films. PLoS One 2017;12:1–21. https://doi.org/10.1371/journal.pone.0183984 .
149. Shahnawaz M, Sangale MK, Ade AB. Analysis of the plastic degradation products. In : Shahnawaz M, Sangale MK, Ade AB, eds. Bioremediation technology for plastic waste Singerpore: Springer; 2019. p. 93–101.
150. Novotný C, Malachová K, Adamus G, et al. Deterioration of irradiation/high-temperature pretreated, linear low-density polyethylene (LLDPE) by Bacillus amyloliquefaciens . Int. Biodeterior. Biodegradation 2018;132:259–267. https://doi.org/10.1016/j.ibiod.2018.04.014 .
151. Umamaheswari S, Subramani M. GCMS Analysis of Pseudomonas sp., mediated degradation of polystyrene. Ann. Biol. Res 2017;8:8–11.
152. Uscátegui YL, Arévalo FR, Díaz LE, Cobo MI, Valero MF. Microbial degradation, cytotoxicity and antibacterial activity of polyurethanes based on modified castor oil and polycaprolactone. J. Biomater. Sci. Polym. Ed 2016;27:1860–1879. https://doi.org/10.1080/09205063.2016.1239948 .
153. Shimpi N, Borane M, Mishra S, Kadam M. Biodegradation of polystyrene (PS)-poly(lactic acid) (PLA) nanocomposites using Pseudomonas aeruginosa . Macromol. Res 2012;20:181–187. https://doi.org/10.1007/s13233-012-0026-1 .
154. Jeon H, Kim M. Biodegradation of poly(L-lactide) (PLA) exposed to UV irradiation by a mesophilic bacterium. Int. Biodeterior. Biodegradation 2013;85:289–293. https://doi.org/10.1016/j.ibiod.2013.08.013 .
155. Kathirvel B, Mathimani T, Rene E, Shanmugam S, Chi N, Pugazhendhi A. Impact of cultivation conditions on the biomass and lipid in microalgae with an emphasis on biodiesel. Fuel 2020;284:1–8. https://doi.org/10.1016/j.fuel.2020.119058 .
156. Correa D, Beyer H, Possingham H, Fargione J, Hill J, Schenk P. Microalgal biofuel production at national scales: Reducing conflicts with agricultural lands and biodiversity within countries. Energy 2020;215:1–12. https://doi.org/10.1016/j.energy.2020.119033 .
157. Mustafa S, Bhatti HN, Maqbool M, Iqbal M. Microalgae biosorption, bioaccumulation and biodegradation efficiency for the remediation of wastewater and carbon dioxide mitigation: Prospects, challenges and opportunities. J. Water Process Eng 2021;41:1–15. https://doi.org/https://doi.org/10.1016/j.jwpe.2021.102009 .
158. Shanmugam S, Hari A, Kumar D, et al. Recent developments and strategies in genome engineering and integrated fermentation approaches for biobutanol production from microalgae. Fuel 2021;285:1–9. https://doi.org/10.1016/j.fuel.2020.119052 .
159. Sarmah P, Rout J. Algal colonization on polythene carry bags in a domestic solid waste dumping site of Silchar town in Assam. Phykos 2018;48:67–77.
160. Wang F, Zhang X, Zhang S, Zhang S, Sun Y. Interactions of microplastics and cadmium on plant growth and arbuscular mycorrhizal fungal communities in an agricultural soil. Chemosphere 2020;254:1–10. https://doi.org/10.1016/j.chemosphere.2020.126791 .
161. Khoironi A, Anggoro S, Sudarno S. Evaluation of the interaction among microalgae Spirulina sp, plastics polyethylene terephthalate and polypropylene in freshwater environment. J. Ecol. Eng 2019;20:161–173. https://doi.org/10.12911/22998993/108637 .
162. Gopal R. Biodegradation of polyethylene by green photosynthetic microalgae. J. Bioremediat. Biodegradation 2017;8:1–8. https://doi.org/10.4172/2155-6199.1000381 .
163. Hadad D, Geresh S, Sivan A. Biodegradation of polyethylene by the thermophilic bacterium Brevibacillus borstelensis . J. Appl. Microbiol 2005;98:1093–1100. https://doi.org/10.1111/j.1365-2672.2005.02553.x .
164. Paço A, Duarte K, da Costa JP, et al. Biodegradation of polyethylene microplastics by the marine fungus Zalerion maritimum . Sci. Total Environ 2017;586:10–15. https://doi.org/10.1016/j.scitotenv.2017.02.017 .
165. Dineshbabu G, Uma V, Mathimani T, Prabaharan D. Elevated CO2 impact on growth and lipid of marine cyanobacterium Phormidium valderianum BDU 20041–towards microalgal carbon sequestration. Biocatal. Agric. Biotechnol 2020;25:1–7. https://doi.org/10.1016/j.bcab.2020.101606 .
166. Jareonsin S, Pumas C. Advantages of heterotrophic microalgae as a host for phytochemicals production. Front. Bioeng. Biotechnol 2021;9:1–17. https://doi.org/10.3389/fbioe.2021.628597 .
167. Hoffmann L, Eggers SL, Allhusen E, Katlein C, Peeken I. Interactions between the ice algae Fragillariopsis cylindrus and microplastics in sea ice. Environ. Int 2020;139:1–9. https://doi.org/10.1016/j.envint.2020.105697 .
168. Matavuly M, Peter M. Marine fungi: Degraders of poly-3-hydroxyalkanoate based plastic materials. Zb. Matice Srp. Prir. Nauke 2009;116:253–265. https://doi.org/10.2298/ZMSPN0916253M .
169. Chen Y, Stemple B, Kumar M, Wei N. Cell surface display fungal laccase as a renewable biocatalyst for degradation of persistent micropollutants bisphenol A and sulfamethoxazole. Environ. Sci. Technol 2016;50:8799–8808. https://doi.org/10.1021/acs.est.6b01641 .
170. Ong H, Ling Shing W. The growth and laccase activity of edible mushrooms involved in plastics degradation. Curr. Top. Toxicol 2019;15:57–62.
171. Mitik-Dineva N, Wang J, Truong VK, et al. Escherichia coli, Pseudomonas aeruginosa, and Staphylococcus aureus attachment patterns on glass surfaces with nanoscale roughness. Curr. Microbiol 2009;58:268–273. https://doi.org/10.1007/s00284-008-9320-8 .
172. da Luz JMR, da Silva MdCS, dos Santos LF, Kasuya MCM. Plastics polymers degradation by fungi. In : Blumenberg M, Shaaban M, Elgaml A, eds. Microorganisms London: IntechOpen; 2019. p. 261–270.
173. 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. https://doi.org/10.1016/S0141-3910(01)00027-1 .
174. Sowmya HV, Ramalingappa , Krishnappa M, Thippeswamy B. Degradation of polyethylene by Trichoderma harzianum-SEM, FTIR, and NMR analyses. Environ. Monit. Assess 2014;186:6577–6586. https://doi.org/10.1007/s10661-014-3875-6 .
175. Gajendiran A, Krishnamoorthy S, Abraham J. Microbial degradation of low-density polyethylene (LDPE) by Aspergillus clavatus strain JASK1 isolated from landfill soil. 3 Biotech 2016;6:1–6. https://doi.org/10.1007/s13205-016-0394-x .
176. Chaudhary AK, Vijayakumar R. Studies on biological degradation of polystyrene by pure fungal cultures. Environ. Dev. Sustain 2020;22:4495–4508. https://doi.org/10.1007/s10668-019-00394-5 .
177. Yanto D, Krishanti N, Ardiati F, et al. Biodegradation of styrofoam waste by ligninolytic fungi and bacteria. IOP Conf. Ser.: Earth Environ. Sci 2019;308:1–11. https://doi.org/10.1088/1755-1315/308/1/012001 .
178. Brunner I, Fischer M, Rüthi J, Stierli B, Frey B. Ability of fungi isolated from plastic debris floating in the shoreline of a lake to degrade plastics. PLoS One 2018;13:1–14. https://doi.org/10.1371/journal.pone.0202047 .
179. Álvarez-Barragán J, Domínguez-Malfavón L, Vargas-Suárez M, González-Hernández R, Aguilar-Osorio G, Loza-Tavera H. Biodegradative activities of selected environmental fungi on a polyester polyurethane varnish and polyether polyurethane foams. Appl. Environ. Microbiol 2016;82:5225–5235. https://doi.org/10.1128/aem.01344-16 .
180. Cosgrove L, McGeechan PL, Robson GD, Handley PS. Fungal communities associated with degradation of polyester polyurethane in soil. Appl. Environ. Microbiol 2007;73:5817–5824. https://doi.org/10.1128/aem.01083-07 .
181. Khan S, Nadir S, Shah ZU, et al. Biodegradation of polyester polyurethane by Aspergillus tubingensis . Environ. Pollut 2017;225:469–480. https://doi.org/10.1016/j.envpol.2017.03.012 .
182. Ali M, Perveen Q, Ahmad B, et al. Studies on biodegradation of cellulose blended polyvinyl chloride films. Int. J. Agric. Biol 2009;11:577–580.
183. Sumathi T, Viswanath B, Sri Lakshmi A, SaiGopal DV. Production of laccase by Cochliobolus sp. isolated from plastic dumped soils and their ability to degrade low molecular weight PVC. Biochem. Res. Int 2016;2016:1–10. https://doi.org/10.1155/2016/9519527 .
184. Jarerat A, Tokiwa Y. Degradation of Poly(L-lactide) by a fungus. Macromol. Biosci 2001;1:136–140. https://doi.org/10.1002/1616-5195(20010601)1:43.0.CO;2-3 .
185. Karamanlioglu M, Houlden A, Robson GD. Isolation and characterisation of fungal communities associated with degradation and growth on the surface of poly(lactic) acid (PLA) in soil and compost. Int. Biodeterior. Biodegradation 2014;95:301–310. https://doi.org/10.1016/j.ibiod.2014.09.006 .
186. Lipsa R, Tudorachi N, Darie-Nita RN, Oprică L, Vasile C, Chiriac A. Biodegradation of poly(lactic acid) and some of its based systems with Trichoderma viride . Int. J. Biol. Macromol 2016;88:515–526. https://doi.org/10.1016/j.ijbiomac.2016.04.017 .

Article information Continued

©2023 Korean Society of Environmental Engineers

This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/3.0/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

Funded by : National Research Foundation of Korea
Award ID : NRF-2021R1A2C4001746
Funding : This study was funded by the National Research Foundation of Korea (NRF-2021R1A2C4001746).

Table 1

Response of Soil Microbial Communities to Different Plastics

Soil type MP typea MP form and concentrationb Enriched microbial taxa Reference
Bacteria
Field soil Plastic mulching n.a. Cyanobacteria [49]
Cinnamon soil LDPE Fragments, 2000 fragments (kg of soil)−1 Actinobacteria, Bacteroidetes, Gemmatimonadetes, and Proteobacteria [51]
Loamy soil LDPE, PVC Particles, 1% and 5% (w/w) Proteobacteria [61]
River shore soil PE, PP, PA, PS, PET, PVC Particles, 2% (w/w) Actinobacteria [62]
Farmland loamy soil HDPE, PLA Particles, 0.5% and 10% (w/w) Actinobacteria, Proteobacteria, Chloroflexi, Gemmatimonadetes [63]
Rice paddy soil PS, PTFE Particles, 0.25% and 0.5% Chloroflexi and Acidobacteria [64]
Farmland soil PE, PS, PVC Particles, 7% and 14% (w/w) Proteobacteria and Actinobacteria [65]
Tea garden soil PVC Particles, 900 mg (3.5 kg soil)−1 Proteobacteria and Firmicutes [66]
Field soil Plastic film Film, n.a. Proteobacteria, Bacteroidetes, and Cyanobacteria [67]
Field soil PE Film, 5% (w/w) Proteobacteria, Actinobacteria, and Acidobacteria [68]
Fungi
Field soil Plastic mulching n.a. Ascomycota and Basidomycota [49]
Farmland soil PE, PS, PVC Particles, 7% and 14% (w/w) Ascomycota [65]
Field soil PE Film, 5% (w/w) Ascomycota [68]
Grassland soil Plastic fragments Fragments, n.a. Mortierellomycota and Ascomycota [69]
Compost PE, PVC, PHA Particles, 0.5% (w/w) Ascomycota and Basidomycota [70]
a

PE: Polyethylene; PVC: Polyvinyl chloride; PS: Polystyrene; PP: Polypropylene; PET: Polyethylene terephthalate; HDPE: High-density polyethylene; LDPE: Low-density polyethylene; PLA: Polylactic acid; PA: Polyamide; PTFE: Polytetrafluorethylene; PHA: Polyhydroxyalkanoates

b

n.a. information not available

Table 2

Plastic Degradation by Actinomycetes, Biodegradation Assay Conditions, and Applied Detection Methods

Actinomycetes strain Plastic typea Biodegradation assay conditions Biodegradation detection method Polymer reduction (%)b Reference
Streptomyces scabies PET Ten (10) mg of PET (ground granules), 1 mL of Tris-HCl (20 mM, pH 7.5), and 3 μg of the enzyme Sub 1 from S. scabies. Incubation for 15 d at 37°C Measurement of the amount of terephthalic acid (TA) released from PET n.a. [112]
Rhodococcus ruber C208 PE Synthetic media, for 2 months, 150 rpm, at 30°C Weight loss; SEM 7.5 [116]
Streptomyces sp. PET Mineral salt media, for 18 d, 120 rpm at 28°C Weight loss, SEM 68.8 [117]
Rhodococcus rhodochrous ATCC 29672 PE Mineral salt media, for 6 months, at 27°C and 85% humidity FTIR; SEM; GPC n.a. [118]
Micrococcus sp. PE Nutrient broth media, for 1 month Weight loss 6.61 [119]
Bacterial consortia (Arthrobacter viscosus; Micrococcus lylae; M. luteus; Bacillus mycoides; B. cereus; B. pumilus; B. thuringiensis) PE Films buried in soil for 7.5 months, at room temperature Weight loss; FTIR; SEM; elongation at break 17.0 [120]
Streptomyces sp. HDPE Mineral salt media, for 18 d, 120 rpm at 28°C Weight loss; SEM 18.26 [121]
Microbacterium paraoxydans PE (pre-treated with nitric acid) Minimal broth media, for 2 months, 180 rpm, at room temperature Weight loss; FTIR 61.0 [122]
Streptomyces coelicoflavus NBRC 15399T LDPE Mineral salt medium for 4 weeks Weight loss; pH change, trinocular microscope 30 [123]
Streptomyces badius Starch-PE (10 d-heat-treated) 0.6% yeast extract media, for 0.75 months, 125 rpm, at 37°C FTIR; tensile strength at break; GPC n.a. [124]
Streptomyces setonii Starch-PE (10 d-heat-treated) 0.6% yeast extract media, for 0.75 months, 125 rpm, at 37°C FTIR; tensile strength at break; GPC n.a. [124]
Streptomyces viridosporus Starch-PE (10 d-heat-treated) 0.6% yeast extract media, for 0.75 months, 125 rpm, at 37°C FTIR; tensile strength at break; GPC n.a. [124]
Streptomyces sp. LDPE M1 media for 45 d, 120 rpm at 25°C Weight loss; Sturm test, AFM, FTIR 5.2 [125]
Rhodococcus ruber PS Synthetic media, for 2 months, Weight loss; SEM 0.8 [126]
Rhodococcus sp. strain 36 PP Bushnell Haas (BH) media, for 1.25 months, at 29°C Weight loss; FTIR; SEM 6.4 [114]
Rhodococcus rhodochrous ATCC 29672 PP (pre-treated with photo and thermo-oxidation) Mineral media, for 6 months, at 27°C FTIR; 1H NMR; ADP/ATP ratio n.a. [127]
Streptomyces sp. PE Mineral salt medium for 6 months, 150 rpm at 30°C Clear zone test; weight loss. 46.16 [128]
Streptomyces sp. Plastic bag Mineral salt medium for 6 months, 150 rpm at 30°C Clear zone test; weight loss. 35.78 [128]
Micrococcus sp. AF10 PU Films buried in soil, for 6 months, at 30–35°C Clear zone test; SEM; FTIR n.a. [129]
Arthrobacter sp. AF11 PU Films buried in soil, for 6 months, at 30–35°C Clear zone test; SEM; FTIR n.a. [129]
Streptomyces spp. PE (containing 6% starch) Mineral salt medium containing 0.6% yeast extract for 4 weeks, 125 rpm at 30°C Weight loss; tensile strength; percent elongation Inconclusive [130]
Amycolatopsis orientalis (enzyme production) PLA (production of purified enzyme) Incubation for 8h, 140 rpm, at 30°C FTIR, pH variation; enzyme degrading activity n.a. [131]
Saccharothrix waywayandensis PLA Basal media with 0.1% gelatin, for 0.25 months, 180 rpm, at 30°C Weight loss; pH variation 95 [132]
Kibdelosporangium aridum PLA Basal media with 0.1% gelatin, for 0.5 months, 180 rpm, at 30°C Weight loss; pH variation; SEM 97 [133]
Amycolatopsis sp. PLA Basal medium for 14 d, 180 rpm at 30°C Weight loss; plate count; clear zone method 60 [134]
Actinomadura sp. T16-1 (enzyme production) PLA (production of PLA-degrading enzyme) Basal media with 0.2% gelatin, for 96h, 150 rpm, at 50°C Enzyme activity n.a. [135]
a

PE: Polyethylene; PET: Polyethylene terephthalate; HDPE: High-density polyethylene; LDPE: Low-density polyethylene; PS: Polystyrene; PP: Polypropylene; PU: Polyurethane; PLA: Polylactic acid; SEM: Scanning electron microscope; FTIR: Fourier-transform infrared spectroscopy; GPC: Gel permeation chromatography; AFM: Atomic force microscopy; NMR: Nuclear magnetic resonance spectroscopy; ADP: Adenosine diphosphate; ATP: Adenosine triphosphate

b

n.a. information not available.

Table 3

Plastic degradation by bacteria, biodegradation assay conditions, and applied detection methods

Bacterial strain Plastic typea Biodegradation assay conditions Biodegradation detection method Polymer reduction (%)b Reference
Anoxybacillus rupiensis Nylon 6 Chemically defined medium containing 0.5% nylon 6 for 7 d, 180 rpm at 65°C HPLC; FTIR n.a [144]
Bacillus cereus PE; PET; PS; pre-treated with UV Mineral salt media with 0.5 g polymer, for 1.25 months, 150 rpm, at room temperature Weight loss; FTIR; SEM 1.6; 6.6; 7.4, respectively [137]
Bacillus gottheilli PE; PP; PET; PS; pre-treated with UV Mineral salt media with 0.5 g polymer, for 1.25 months, 150 rpm, at room temperature Weight loss; FTIR; SEM 6.2; 3.0; 3.6; 5.8, respectively [137]
Achromobacter denitrificans PE pre-treated with UV and heat (70°C for 3 d) Synthetic nutrient medium with 0.1 g polymer for 2 months, 180 rpm at 30°C Weight loss; tensile strength; NMR; XRD; thermogravimetric analysis; carbon analysis; GCMS 40 [145]
Bacillus sp. YP1 PE Liquid carbon-free media (LCFBM) with 1 g polymer, for 2 months, 120 rpm, at 30°C Weight loss; FTIR; SEM; GPC; tensile strength 10.7 [146]
Enterobacter asburiae YT1 PE LCFBM with 1 g polymer, for 2 months, 120 rpm, at 30°C Weight loss; FTIR; SEM; GPC; tensile strength 6.1 [146]
Acinetobacter baumannii PE Synthetic medium with film for 30 d, 100 rpm at 37°C FTIR; GC-MS; tensile strength n.a [147]
Indigenous Marine Microbial Community bioaugmented with Lysinibacillus sp. and Salinibacterium sp. PE Nutrient broth media enriched with saline water, for 6 months, 120 rpm, at 25°C Weight loss; FTIR; SEM 19 [148]
Bacillus cereus PE Basal medium with polymer strip for 2 months GC-MS n.a [149]
Mesophilic mixed Bacterial Culture (Bacillus sp. and Paenibacillus sp.) PE Basal media with 100 mg of polymer, for 2 months, at 30°C Weight loss; FTIR; SEM 14.7 [106]
Bacillus spp. PE Synthetic media with polymer powder for 16 weeks, 150 rpm at 37°C Weight loss; FTIR; GC-MS 35.72 [139]
Biofilm composed of Pirellulaceae, Phycisphaerales, Cyclobacteriaceae, and Roseococcus PE and PP Dechlorinated tap water, Woods Hole media, for 0.7 months (incubation in tanks in a greenhouse exposed to natural light) DNA extraction, amplification and sequencing (evaluation of the effects of substrate type on microbial communities) n.a [105]
Bacillus amyloliquefaciens PE Bushell Haas media with polymer film for 2 months, 120 rpm at 28°C Weight loss; GPC; FTIR; DSC; TGA; ESI-MS 3.2 [150]
Brevibacillus borstelensis PE Synthetic media with polymer powder for 16 weeks, 150 rpm at 37°C Weight loss; FTIR; GC-MS 20.28 [139]
Bacillus sp. strain 27 PP Bushell Haas media with 0.5 g polymer, for 1.25 months, at room temperature Weight loss; FTIR; SEM; pH variation 4 [114]
Pseudomonas sp. PS Mineral salt media with polymer beads for 1 month, 120 rpm at 37°C GC-MS na [151]
Pseudomonas aeruginosa PU Trypticase soy broth media, for 72 h, at 37°C Weight loss; SEM; tensile strength and elongation at break 2.5 [152]
Escherichia coli PU Trypticase soy broth media, for 72 h, at 37°C Weight loss; SEM; tensile strength and elongation at break 2.4 [152]
Pseudomonas aeruginosa PS-PLA
Nanocomposites		 Minimum salt media, for 0.94 months, at room temperature Weight loss; FTIR; SEM; pH variation 9.9 [153]
Stenotrophomonas maltophilia LB 2–3 PLA Mineral medium, for 1.33 months, at 37°C GPC; FTIR; NMR; tensile strength n.a [154]
a

PE: Polyethylene; PET: Polyethylene terephthalate; PS: Polystyrene; PP: Polypropylene; PU: Polyurethane; PLA: Polylactic acid; HPLC: High-performance liquid chromatography; FTIR: Fourier-transform infrared spectroscopy; SEM: Scanning electron microscope; NMR: Nuclear magnetic resonance spectroscopy; XRD: X-Ray diffraction analysis; GC-MS: Gas chromatography–mass spectrometry; GPC: Gel permeation chromatography; DSC: Differential scanning calorimetry; TGA: Thermogravimetric analysis; ESI-MS: Electrospray ionization mass spectrometry,

b

n.a. information not available

Table 4

Plastic degradation by fungi, biodegradation assay conditions, and applied detection methods

Fungal strain Plastic typea Biodegradation assay conditions Biodegradation detection method Polymer reduction (%)b Reference
Aspergillus flavus PE Sole carbon source medium for 28 d, 150 rpm at 28°C Weight loss; molecular weight shift; surface smoothness change 3.9025 [31]
Aspergillus flavus VRKPT2 PE Synthetic media with mineral oil, for 1 month, at 30°C Weight loss; FTIR; SEM 9.34 [172]
Aspergillus tubingensis VRKPT1 PE Synthetic media with mineral oil, for 1 month, at 30°C Weight loss; FTIR; SEM 6.88 [172]
Aspergillus flavus PE (with 6% starch) Mineral salt medium containing 3% yeast extract for 4 weeks, 125 rpm at 30°C Weight loss; tensile strength; percent elongation 1.2 [130]
Mucor rouxii NRRL 1835 PE (with 6% starch) Mineral salt medium containing 3% yeast extract for 4 weeks, 125 rpm at 30°C Weight loss; tensile strength; percent elongation Inconclusive [130]
Aspergillus oryzae PE Synthetic medium with ampicillin, for 4 months, at 28°C in a shaker incubator Weight loss; FTIR; GC-MS 36.4 [139]
Penicillium simplicissimum YK PE Medium C with 0.5 g polymer, for 3 months, 150 rpm, at 30°C FTIR; GPC n.a. [173]
Zalerion maritimum PE Minimum growth media with 0.130 g of polymer, for 0.94 months, 120 rpm, at 25°C Weight loss; FTIR; NMR 43 [164]
Trichoderma harzianum PE (UV treated) Mineral salt medium for 3 months Weight loss; SEM; FTIR; NMR 40 [174]
Pleurotus ostreatus PE (120 d exposed to sunlight) Mineral medium, for 2 months, at 25°C FTIR; SEM; mechanical properties n.a. [172]
Aspergillus clavatus JASK1 PE M1 media for 90 d, 120 rpm, at 25°C Weight loss; Strum test; FTIR; SEM; AFM analysis. 35 [175]
Cephalosporium sp. (NCIM 1251) PS Mineral salt media, for 2 months, at 120 rpm, at 28°C Weight Loss; FTIR; SEM; pH variation; gel permeation chromatography 2.17 [176]
Mucor sp. (NCIM 881) PS Mineral salt media, for 2 months, at 120 rpm, at 28°C Weight Loss; FTIR; SEM; pH variation; gel permeation chromatography 1.81 [176]
Pestalotiopsis sp. PS (Styrofoam) Malt extract broth, for 1 month, at 25°C Weight loss; FTIR; SEM 74.43 [177]
Ceriporia sp. PS (Styrofoam) Malt extract broth, for 1 month, at 25°C Weight loss; FTIR; SEM 19.44 [177]
Cymatoderma dendriticum PS (Styrofoam) Malt extract broth, for 1 month, at 25°C Weight loss; FTIR; SEM 15.50 [177]
Cladosporiumcladosporioides PU Agar medium Clear zone test; FTIR n.a. [178]
Cladosporium pseudocladosporioides PU Mineral medium for 14 d at 30°C Weight loss; FTIR; GC-MS; SEM 87 [179]
Phoma sp. PU Buried in soil for 5 months Clear zone test; tensile strength n.a. [180]
Aspergillus tubingensis PU Mineral salt medium, for 0.75 months, at 150 rpm, at 37°C FTIR; SEM; tensile strength n.a. [181]
Phanerochaete chyrosporium PVC (blended with cellulose) Soil buried (soil was mixed with municipal sewage sludge), for 6 months Clear zone test; FTIR; SEM n.a. [182]
Cochliobolus sp. PVC Laccase degradation FTIR; SEM; GC-MS n.a. [183]
Tritirachium album PLA Basal media, for 0.5 months, at 180 rpm, at 30°C Weight loss; SEM; pH variation 76 [184]
Thermomyces lanuginosus PLA Wheat grain with mineral salt medium, for 2 months, at 50°C SEM; tensile strength n.a. [185]
Trichoderma viride PLA (plasticized with USE) Liquid Sabouraud medium, for 0.7 months, at 28°C Weight loss; FTIR; gel permeation chromatography; SEM 1.2 [186]
Aspergillus nomius PE M1 media for 45 d, 120 rpm at 25°C Weight loss; Sturm test, AFM, FTIR, GC-MS 4.9 [125]
a

PE: Polyethylene; PS: Polystyrene; PU: Polyurethane; PVC: Polyvinyl chloride; PLA: Polylactic acid; FTIR: Fourier-transform infrared spectroscopy; SEM: Scanning electron microscope; GC-MS: Gas chromatography–mass spectrometry; GPC: Gel permeation chromatography; NMR: Nuclear magnetic resonance spectroscopy; AFM: Atomic force microscopy

b

n.a. information not available