Development of Polyethylene Terephthalate Plastic Eating Pseudomonas Putida A

DEVELOPMENT OF POLYETHYLENE TEREPHTHALATE PLASTIC EATING PSEUDOMONAS PUTIDA

______

A Thesis

presented to

the Faculty of Natural and Applied Sciences

at Notre Dame University-Louaize

______

In Partial Fulfillment

of the Requirements for the Degree

Master of Science

______

RANA MATAR

OCTOBER 2020

ABSTRACT

Pollution has long been a burden to humanity, and among the most prominent types is plastic pollution. Plastics have been an expedient source for human activity and progress, however its large carbon footprint and harmful effects on animals, humans and habitats outweighs its advantages. Despite recycling efforts, plastic’s non bio-degradability continues to harm the environment.

Bioremediation has been the most recent endeavor in managing plastic pollution, where recently discovered microorganisms have shown the capacity to break down different types of plastics. This paper highlights all studies pertaining to the plastic eating bacterium Ideonella sakaiensis with the potential of breaking down

Polyethylene Terephthalate (PET). This degradation is done through the action of its enzyme Polyethylene Terephthalate hydrolase (PETase), producing

Monohydroxyethyl Terephthalate (MHET), which is further broken down into terephthalic acid (TPA) and ethylene glycol (EG) by the action of

Monohydroxyethyl Terephthalate hydrolase (MHETase) enzyme. Among the many types of bacteria capable of breaking down plastics, Pseudomonas Putida has been seen to have an effect on Polyethylene (PE), but no study was conducted to assess its effect on PET plastic. This bacterium is well known for its ability to

ii survive in extreme environments including salt water. This study aims to transform P. putida into a PET degrading bacterium through the use of a constructed pSEVA441-EGFP-PrpsL-PelB-IsPETase-FLAG tag plasmid (also known as pRT2210 plasmid). Transformed P. putida were selected based on their streptomycin (sm) resistance and the formation of a yellowish halo around colonies. The successful production of PETase enzyme could be validated by the western blot analysis, and its effectiveness through the gravimetric weight loss analysis. Anticipated results may indicate that transformed P.putida bacteria are capable of producing PETase enzymes that may be functional against PET plastic in salt water.

Keywords: PET Plastic, PETase, Pseudomonas Putida, bacterial transformation

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ACKNOWLEDGEMENTS

I would like to thank every person who has played a role in helping me accomplish my thesis project.

First, I would like to give a special thank you to my supervisor Dr. Pauline

Aad, if it wasn’t for her, I would have never come up with such a topic. She has been a great mentor and an exceptional guide throughout my thesis work.

In addition, I would like to extend gratitude to Dr. Diala El Khoury who also served as guiding collaborator during my project. She has always shown interest in my work and has always supported and aided me throughout my university courses.

I appreciate the presence of Dr. Diala El Khoury and Dr. Kamil Rahme as committee members.

Second, I would like to thank my best friend and sister Tina Touma. She was present with me from the very first day of university and has always had by back.

She never let me face difficulties alone, and has been and will always be my biggest support system.

Finally, I am beyond grateful for my parents and their never ending love and support. They have always had faith in my abilities and have always pushed me to strive for more.

TABLE OF CONTENTS

ABSTRACT ...... ii

Keywords: PET Plastic, PETase, Pseudomonas Putida, bacterial transformation ...... iii

ACKNOWLEDGEMENTS ...... iv

TABLE OF CONTENTS ...... vi

LIST OF FIGURES ...... viii

LIST OF TABLES ...... ix

LIST OF ABBREVIATIONS ...... x

I. Introduction ...... 1

1. Plastic pollution ...... 1

2. Plastic: a technological miracle that has proved deadly ...... 2

3. Bioremediation of plastic compounds: finding a way out ...... 4

II. Literature Review ...... 7

1. Ideonella Sakaiensis: a newly categorized organism with immense plastic degrading abilities ...... 7

2. PETase analysis and improvement ...... 9

3. Analysis and improvements of the MHETase enzyme ...... 15

4. The biodegradation of Polyethylene plastic by Pseudomonas Putida ...... 17

5. Bacterial Transformation and Types of Vectors...... 19

6. SEVA plasmids (pSEVA) and their Uses ...... 20

III. Materials and Methods ...... 25

1. Materials and Reagents ...... 25

2. Pseudomonas Putida and Cell culture ...... 26

3. Designing the pRT2210 plasmid ...... 27

4. The Transformation of Pseudomonas putida AC 13 by pRT2210 plasmid ...... 28

5. Selecting Transformed Pseudomonas putida Bacteria ...... 30

6. Protein Purification and Western Blotting ...... 30

7. Hydrolysis of PET films by transformed P. putida ...... 32

8. Statistical analysis ...... 33

IV. Results ...... 33

1. Identification of Transformed Pseudomonas putida ...... 34

2. Protein Purification and Western Blot ...... 35

3. Hydrolysis of PET plastic by PETase enzyme from transformed P. putida cells ...... 37

V. Discussion ...... 38

VI. Conclusion ...... 40

VII. References ...... 44

vii

LIST OF FIGURES

Figure 1: PET manufacture...... 4

Figure 2: PET degradation...... 8

Figure 3: Synthesis of indole-based AB monomers (3-6) and polyesters (P3-6) .... 10

Figure 4: A graphical representation of Ideonella sakaiensis, its enzymes and their modification for enhanced activity ...... 17

Figure 5: SEVA structure and nomenclature...... 22

Figure 6: The constructed pRT2210 plasmid that may be used for the transformation of P.putida AC 13...... 34

Figure 7: White and fluorescent colonies...... 35

Figure 8: Anticipated Western Blot Analysis results...... 36

viii

LIST OF TABLES

Table 1: The Seven Types of Plastics and Some of Their Major Characteristics. ... 42

Table 2: Examples of microorganisms (fungi and bacteria) capable of degrading

the seven types of plastic...... 43

LIST OF ABBREVIATIONS

BHETase: bis-(2-hydroxyethyl) EGFP: enhanced green fluorescent protein

terephthalate hydrolase GcIR: glyoxylate carboligase repressor

BHET: bis-(2-hydroxyethyl) terephthalate GRAPE: Greedy Accumulated Strategy for bp: base pair Protein Engineering

BPA: bisphenol A H2O: water

CaCl2: calcium chloride HDPE or PE-HD: high-density polyethylene

Cd: Cadmium Hg: mercury

CD: compact disc His: Histidine

DB: database I.sakaiensis: ideonella sakaiensis

DYKDDDDK: Aspartic acid-Tyrosine- Lysine- IsPETase: Ideonella Sakaiensis polyethylene

Aspartic acid-Aspartic acid-Aspartic acid- terephthalate hydrolase

Aspartic acid-Lysine kDa: Kilodaltons

ECL: enhanced chemiluminescence KH2PO4: potassium dihydrogen phosphate

E.coli: Escherichia coli LB: Luria Bertani

EG: ethylene glycol LCC: leaf-branch compost cutinase

LDPE or PE-LD: low-density polyethylene PETase: polyethylene terephthalate

hydrolase MgSO4: magnesium sulfate

2PET: two ethylene terephthalate MHETase: Monohydroxyethyl

Terephthalate hydrolase PET or PETE: polyethylene terephthalate

MHET: Monohydroxyethyl Terephthalate PETG: polyethylene terephthalate glycol

Na2HPO4: disodium hydrogen phosphate PLA: polylactic acid

NaCl: sodium chloride p-NP: p-nitrophenyl

NdeI: endonuclease isolated from Neisseria PP: polypropylene

denitrificans P. putida: pseudomonas putida

NH4Cl: ammonium chloride p. tricornutum: Phaeodactylum tricornutum

NotI: restriction enzyme isolated from the pSEVA: Standard European Vector bacterium Nocardia otitidis Architecture plasmid

Ori: origin of replication PS: polystyrene

PBS: polybutylene succinate PVC or V: polyvinyl chloride

PE: polyethylene PVDF: Polyvinylidine difluoride

PEF: polyethylene-2,5-furandicarboxylate RIPA: Radio immune-precipitation assay pelB: pectate lyase B S.D: standard deviation

SDS: sodium dodecyl sulfate

SDS-PAGE: Sodium Dodecyl Sulfate–

polyacrelamide gels

SEVA-DB: Standard European Vector

Architecture Database

Sm: streptomycin

SPLamB: maltoporin binding periplasmic

protein

SPMalE: maltose/maltodextrin binding

periplasmic protein

SRP: signal recognition particle

TBST: Tris-Buffered Saline,0.1% Tween

TfCut2: PET degrading cutinase from

Thermobifida fusca

T.fusca: thermobifida fusca

TPA: terephthalate

UV: ultraviolet

xii

I. Introduction

1. Plastic pollution

Plastic pollution is among the most common types of pollution worldwide with a large carbon footprint. The composition and nature of plastic allows it to last for long periods of time (Barnes et al., 2009) and escape complete degradation (Webb et al., 2013), where it only breaks down into smaller pieces of microplastics by UV radiation, photodegradation and oxidation (Biundo et al.,

2018). These microplastics gradually bioaccumulate in different organisms and environments and lead to negative effects on animals, humans, and habitats

(Barnes et al., 2009; Worm et al., 2017) (Biundo et al., 2018). Moreover, levels of plastic pollution in oceans and seas continue to rise due to inappropriate plastic waste management as around 4 to 12 million tons of plastic materials make their way into marine habitats yearly (Jambeck et al., 2015). Plastic’s ability to escape complete degradation exacerbates the situation (Webb et al., 2013) and forces many countries worldwide to implement strategies in order to decrease the manufacturing and use of plastic products.

The versatility of plastic as well as its multipurpose applications and cost- effectiveness (North and Halden, 2013) makes its discontinuous use economically very challenging. Moreover, the lack of appropriate alternatives to plastics

(Kershaw, 2018) has increased the challenge of enforcing such efforts to reduce; reuse options are not always possible since many plastics release toxic elements

(Laskar and Kumar, 2019). As a result, the most effective way to decrease the amount of plastic pollution is to recycle, since a solution to remove plastic effectively and permanently from the environment has not been devised or implemented yet.

Collected plastic for recycling is therefore undergoing a variety of processes to reduce its volume, and strategies are being implemented to decrease the amount of plastics in earth’s environment (Sinha et al., 2010; Rhodes, 2018).

These include incineration, land filling, mechanical recycling, chemical recycling, replacing plastic items with glass, and even bioremediation. These integrative plastic contamination solutions are very challenging, mainly due to the diversity of produced plastic.

2. Plastic: a technological miracle that has proved deadly

Seven different types of plastic are categorized according to their different polymer types. These include polyethylene terephthalate (PET or PETE), high- density polyethylene (HDPE or PE-HD), polyvinyl chloride (PVC or V), low-density polyethylene (LDPE or PE-LD), polypropylene (PP), polystyrene (PS), other resin

2 and undefined multi-materials (So et al., 2016). The properties, characteristics, common application, potential for toxin release and their effects, as well as the recycling possibility of these different types are summarized in Table 1.

Among these seven types of plastics, PET poses a serious problem to the environment, especially due to the fact that it has a general lifetime ranging from

16 to 48 years (Müller et al., 2001). This thermoplastic polyester polymer resin is among the most essential raw materials used in the making of synthetic fibers

(Sinha et al., 2010). PET, of chemical formula (C10H8O4)n, is used to make items such as plastic water bottles, and is manufactured through petroleum terephthalic acid (TPA) and ethylene glycol (EG) monomer polymerization with the use of antimony, titanium- or germanium-based catalysts (Westerhoff et al.,

2008) as seen in Figure 1.

This semi-crystalline resin is naturally colorless and malleable, and its rigidity depends on the manner in which it is processed by, either by ethylene glycol and dimethyl terephthalate transesterification, or by ethylene glycol and terephthalic acid esterification (Begum et al., 2020).

Because of its wide range of use, PET disposal has been a main issue, and bioremediation techniques have been taking center stage in order to find ways to abolish it effectively.

Figure 1: PET manufacture: The process is accomplished by petroleum terephthalic acid (TPA) and ethylene glycol (EG) monomer polymerization (Al-

Sabagh et al., 2016).

3. Bioremediation of plastic compounds: finding a way out

Bioremediation is a process that relies on various agents such as yeast, fungi and bacteria (Bhatnagar and Kumari, 2013) to decompose and eliminate different types of contaminants such as plastic (Gupta and Prakash, 2020). Many such microorganisms have been discovered with the capacity to degrade plastics as briefly summarized in table 2.

One major microorganism is Pseudomonas Putida, a gram-negative bacterium that can live in either soil or water and adapt to harsh environments

(Nikel and de Lorenzo, 2018). It can survive in a pH range between 5.5 and 10

(Reva et al., 2006) with an optimal pH at around 7 (Elsayed and El-Nady, 2013).

These bacteria grow at an optimal temperature of 30 °C, but can survive at temperatures as low as 4°C (Fonseca et al., 2011). P. putida has been found to rarely cause disease in humans (Peter et al., 2017). The major plastic being degraded by P. putida is polyethylene (PE) plastic, as will be described in the next section.

Other plastic degrading organisms include the following: The fungi

Phanerochaete chrysosporium and Engyodontium album (Jeyakumar et al., 2013) and the bacterium Stenotrophomonas panacihumi (Jeon and Kim, 2016) can degrade polypropylene (PP). Moreover, some polyethylene types (LDPE and

HDPE) may be broken down by organisms such as the bacteria Rhodococcus ruber

(Orr et al., 2004) and Brevibacillus borstelensis (Hadad et al., 2005) and the fungus

Penicillium simplicissimum (Sowmya et al., 2015). The fungi Phanerochaete chrysosporium, Lentinus tigrinus, Aspergillus niger and Aspergillus sydowii (Ali et al., 2014) and the bacteria Pseudomonas otitidis, Acanthopleuribacter pedis,

Bacillus cereus (Anwar et al., 2016) have all been found to degrade polyvinyl

5 chloride (PVC). In addition, the bacterium Rhodococcus ruber (Mor and Sivan,

2008) and the fungi Cephalosporium species (Chaudhary and Vijayakumar, 2019) are capable of acting upon polystyrene (PS). Other resins have also been degraded by microorganisms such as Bacillus pallidus (Tomita et al., 2003), Vibrio furnisii (Sudhakar et al., 2007), Cladosporium cladosporioides, Xepiculopsis graminea, and Penicillium griseofulvum (Brunner et al., 2018).

Among the different microbes found to play a role in treating plastic pollution, many bacteria have been discovered with the potential of degrading

PET plastic wastes (Shahnawaz et al., 2019), such as Thermobifida fusca (Müller et al., 2005), Thermobifida halotolerans (Ribitsch et al., 2012), Thermomonospora curvata (Wei et al., 2014), Saccharomonospora viridis (Kawai et al.,

2014),Comamonas testosterone F6 (Gong et al., 2018), Streptomyces species

(Farzi et al., 2019), Nocardia species (Sharon and Sharon, 2012), and Ideonella sakaiensis 201-F6 (Yoshida et al., 2016).

The following section summarizes all studied that were conducted on

Ideonella sakaiensis and Pseudomonas putida; the two key bacteria that will be used in this paper. By the use of genetic engineering methods, PETase enzyme genes will be isolated for I. sakaiensis and will be used in order to transform P. putida into PET plastic eating bacteria in salt water.

II. Literature Review

1. Ideonella Sakaiensis: a newly categorized organism with immense plastic

degrading abilities

In 2016, a group of scientists (Yoshida et al., 2016) discovered a plastic- eating bacterium collected from a microbial consortium in Sakai Japan. Studies conducted on this bacterium proved its relation to the genus Ideonella; however, it was clearly distinguished from other species within the genus and was identified as a novel bacterium termed Ideonella sakaiensis strain 201- F6T.

Ideonella sakaiensis is a gram negative, rod-shaped, non-spore-forming, aerobic bacterium capable of degrading PET. In addition, I. sakaiensis cells are motile and survive in habitats with a pH range from 5.5 to 9.0 (optimal pH between 7 - 7.5), and temperatures ranging between 15 and 42ºC (optimal 30 to 37ºC)

(Tanasupawat et al., 2016; Widyastuti, 2018).

I. sakaiensis is capable of breaking down PET as a carbon source used for its own growth (Widyastuti, 2018). This process is accomplished through the action of two primary hydrolytic α/β-hydrolase fold enzymes known as Polyethylene

Terephthalate hydrolase (PETase) and Monohydroxyethyl Terephthalate hydrolase (MHETase) (Ramirez-Ekner et al., 2017). PETase is a cutinase-like

7 enzyme (Chen et al., 2018) that breaks down PET into mono-(2-hydroxyethyl) terephthalate (MHET), which in turn is broken down into PET educts terephthalate (TPA) and ethylene glycol (EG) by the action of MHETase enzyme

(Palm et al., 2019), as seen in Figure 2.

Figure 2: PET degradation (Bornscheuer, 2016): PET plastic is degraded by

IsPETase enzyme into MHET, which is further broken down into EG and TPA through the action of MHETase enzyme.

2. PETase analysis and improvement

Multiple studies have observed the potential and efficiency of Ideonella

PETase enzyme to degrade plastic, as well as their possible usage for integration in an industrialized remediation program for plastic pollution. These studies are summarized in this section and demonstrate the variable conditions under which such experiments were conducted, the plastic types used as well as the outcome of those experiments.

Two studies were conducted on four indole-based AB-type polyesters: P3,

P4, P5, and P6, synthesized following melt-polycondensation of indole-based hydroxyl-carboxylate (AB-type) monomers with a varied number of flexible methylene units in the backbones (Liu et al., 2018; Sagong et al., 2020; Wang et al., 2020) as seen in Figure 3. Favorable interactions between the polyesters and the enzyme polyethylene terephthalate hydrolase isolated from the bacteria

Ideonella sakaiensis (IsPETase) were reported. Molecular docking simulations showed the matching ligands to a macromolecule (Salmaso and Moro, 2018), and results revealed the successful bio-degrading ability of PETase on the indole- based AB polyesters P3, P4, and P5. However, polymer P6, the most thermally stable and flexible polymer, was not degraded by PETase, mainly due to the

9 increased methylene units present in the polymer backbone and its high degree of crystallinity which impaired enzymatic degradation (Sagong et al., 2020).

Figure 3: Synthesis of indole-based AB monomers (3-6) and polyesters (P3-6)

(Wang et al., 2020)

Further, the I. sakaiensis enzymatic activity of IsPETase was highlighted in various environmental and physiochemical conditions. Salts and glycerol promoted the enzymatic activity of IsPETase; however, organic solvents and detergents inhibited it (Liu et al., 2019). With industrial polyesters such as p- nitrophenyl (p-NP) esters and naphthyl esters on the rise, Kawai et al. (2019) showed that the acyl esters P-nitrophenyl (p-NP) (Kawai et al., 2019) were hydrolyzed by IsPETase while naphthyl esters were hydrolyzed by genetically engineered IsPETase mutants (IsPETaseS93M, IsPETaseW159F, and IsPETaseN241F).

These mutants differed in their hydrophobicity and steric hindrance effects as compared to normal IsPETase (Liu et al., 2019).

Further, research on the PETase structure, crystallization and binding site analysis were conducted in order to improve the PETase/PET binding and degradation. Comparative analysis of the PETase fold structure between different organisms showed that the α/β-hydrolase fold structure of PETase derived from I. sakaiensis showed a more open active site cleft than homologous cutinases such as from Thermobifida fusca (Furukawa et al., 2019) and Fusarium solani (Egmond and de Vlieg, 2000). The study revealed optimized PETase activity through the modification of the binding cleft achieved by the mutation of two-active site residues into conserved amino acids in cutinases. In addition, experimental studies showed that PETase not only degrades PET but may also break down polyethylene-2,5-furandicarboxylate (PEF), a newly synthesized semiaromatic PET substitute (Eerhart et al., 2012) with enhanced barrier characteristics (de Jong et al., 2012). However, PETase was not effective against aliphatic polyesters such as polybutylene succinate (PBS) (Xu and Guo, 2010) and polylactic acid (PLA)

(Drumright et al., 2000), hence proving that this enzyme is an aromatic polyester hydrolase (Austin et al., 2018). Furthermore, the active site of I.

Sakaiensis derived PETase (IsPETase) possesses increased flexibility at room

11 temperature as compared to its thermophilic counterparts as PET-degrading cutinase (TfCut2) from T. fusca (Roth et al., 2014) and a leaf-branch compost cutinase (LCC) derived from metagenomes (Sulaiman et al., 2014). PETase, TfCut2, and LCC are all structurally similar and have a high sequence identity. However,

IsPETase displays near its active site a disulfide bond that controls its flexibility and its removal decreases hydrolase activity and destabilizes the catalytic triad

(Bollinger et al., 2020). Moreover, many residual substitutions of PETase active site enhance the binding of PET and reveal an improved and favorable substrate enzyme interaction. This was revealed through comparative molecular docking of a model substrate made up of two ethylene terephthalate (2PET) repeating units

(Fecker et al., 2018).

The expression of IsPETase was improved by using a secretion system based on the fusion of Sec-dependent and SRP-dependent signal peptides

(maltose/maltodextrin binding periplasmic protein (SPMalE), maltoporin (SPLamB) obtained from E. coli). These peptides are involved in pathways related to translocation across cytoplasmic membranes of bacteria (Dastjerdeh et al., 2019).

This secretion system was successfully inserted into the expression vector PET22b having the IsPETase gene, thus leading to the development of the extracellular enzyme IsPETase derived from E.coli and active against PET films (Seo et al.,

2019). Moreover, enhancement of the enzyme-substrate binding of IsPETase and

PET film was tested via incubating the film with anionic alkyl surfactants such as

- - - + Cn-OSO3 , Cn-SO3 , Cn-COO , Cn-N(CH3)3 before adding the cationic PETase

(Furukawa et al., 2018). These surfactants are organic substances that create negatively charged particles in water (Beringer and Kurz, 2011), and therefore, the positive-negative interaction of PET and PETase leads to the improvement of the action of PET by 120-fold, with the thickness of the film decreasing by 22% after 36 hours at 30 °C (Furukawa et al., 2018).

In addition, similar work revealed that some mutations such as Y58A,

W130A, W130H, and A180I in IsPETase lead to the formation of more space on several binding centers allowing for more efficient substrate binding. Other mutations such as S185H may also improve aromatic properties on the edge of binding pockets, allowing for enhanced enzymatic activity as compared to the wild type enzyme (Liu et al., 2018).

Genetic engineering of a photosynthetic microalga Phaeodactylum triconutum, a marine phytoplankton diatom (Xue et al., 2015), which serves as an expression system for several enzymes (Hempel et al., 2011; D'Adamo et al.,

2019), was used to produce PETase enzyme whose genes were primarily isolated from I. sakaiensis. The aim of this study was to decompose PET by using a

13 microorganism that is capable of surviving in salt water, unlike I. sakaiensis

(Tanasupawat et al., 2016). The genetically modified microalgae P. tricornutum with I. Sakaiensis PETase successfully produced PETase (PETaseR280A-FLAG) and showed catalytic activity against PET and PETG copolymer (Moog et al., 2019).

Furthermore, protein engineering focused on six important residues found around the substrate-binding groove to create new highly efficient PETase mutants expressed in E.coli (Ma et al., 2018). R61A, L88F, and I179F variants were found to have 1.4 fold, 2.1 fold, and 2.5 fold increases, respectively, as compared to wild type PETase. In addition, studies revealed that I179F mutant had the highest PET degradation rate of 22.5 mg per μmol·L−1 PETase per day (Ma et al.,

2018).

Other research focused on the development of a rational protein engineering strategy in order to improve IsPETase’s low thermal stability and PET degrading ability. This strategy relied on highly thermally stable IsPETase variants according to mutation differences, including IsPETaseP181A, IsPETaseS121D/D186H,

IsPETaseS121E/D186H, IsPETaseP181A/S121D/D186H, IsPETaseP181A/S121E/D186H,

IsPETaseS121D/D186H/R280A, and IsPETaseS121E/D186H/R280A. IsPETaseS121E/D186H/R280A variants which possessed a PET degradation activity that was enhanced by 14-fold at 40 °C as compared to wild type IsPETase, and had a melting temperature that was

14 increased by 8.81°C . This variant was manufactured with an extended subsite IIc that provides a hydrophobic and non-protruding cleft (Joo et al., 2018), as well as a stabilized β6-β7 connecting loop that is among the most flexible parts of the enzyme (Son et al., 2019).

In addition, a computational strategy known as greedy accumulated strategy for protein engineering (GRAPE) was employed to optimize cluster mutations in order to develop functional variations and diminishing the risk of joining mutations with opposing outcomes. A new variant (S214H-I168R-W159H-

S188Q-R280A-A180I-G165A-Q119Y-L117F-T140D) called IsPETase-M10 or

DuraPETase possessed a 31°C higher melting point and a 23% enhanced PET degradation capability at moderate temperatures (Cui et al., 2019).

3. Analysis and improvements of the MHETase enzyme

The role of MHETase is to break down MHET into PET educts terephthalate

(TPA) and ethylene glycol. A MHETase variant was successfully generated to also hydrolyze bis-(2-hydroxyethyl) terephthalate (BHET), an intermediate produced from the glycolysis of PET (Yue et al., 2013), into its basic building block TPA.

Creating new MHETase variants allows the breakdown of many partially degradable polymer products such as BHET.

The structure and mode of action of one of I. Sakaiensis’ key enzymes

MHETase consist of an α/β-hydrolase domain with nucleophile His-Acid catalytic triad (Holmquist, 2000) and a lid domain, an additional element involved in structural and functional differences (Mindrebo, Nartey et al. 2016). Structural studies revealed that the lid domain of MHETase allows for MHET hydrolysis; however, MHETase has low activity towards BHET (Yue et al., 2013; Palm et al.,

2019).

Similar investigations relied on the extracellular production of active

MHETase enzyme derived from I. Sakaiensis. Experiments revealed that MHETase not only has a hydrolyzing effect on MHET, but can also play a role similar to that of BHETase and exo-PETase which break down BHET and PET pentamer respectively. Additional research led to the development of a

MHETaseR411K/S416A/F424I variant, which possesses improved PET film degradation activity and an increased BHET activity (Sagong et al., 2020).

Ideonella Sakaiensis’ enzymes may be of high importance within the biodegradation field. The genes of these enzymes or their alteration may be used in several ways in order to progress the elimination of plastic. Recombinant technology may be among the various processes used in order to achieve successful outcomes based on these enzymes. A graphical representation of

Ideonella sakaiensis, its enzymes as well as their modification for enhanced activity is seen in Figure 4.

Figure 4: A graphical representation of Ideonella sakaiensis, its enzymes and their modification for enhanced activity

4. The biodegradation of Polyethylene plastic by Pseudomonas Putida

When it comes to bioremediation, studies have shown that Pseudomonas

Putida has the ability to biodegrade polyethelyne (PE) plastic as the major carbon source. This degradation was accomplished under the following optimum conditions: mineral salt water of pH 6.5, Temperature of 37 °C, and an incubation period of seven days. (Jailawi et al., 2015).

Studies have revealed that, under certain conditions, P. Putida strain KT2440 is capable of metabolizing ethylene glycol. This process is accomplished through the inhibition of the glyoxylate carboligase pathway by the transcriptional regulator

GcIR (glyoxylate carboligase repressor) (Li et al., 2019). Further experiments proved that P. putida has the ability to biodegrade many plastic materials such as: plastic bags, polythene bags, plastic cups, and milk covers, with milk cover being the most degradable substance (Saminathan et al., 2014). Evidence also showed that P. putida cells of strain IRN22 were capable of fragmenting and biodegrading

LDPE (powdered low-density polyethylene plastic) into polyhydroxyalkanoate polymers (Montazer et al., 2019). Another study conducted on P. Putida bacterial cells revealed that certain enzymatic systems that require oxygen have the ability to oxidize PE. This process is accomplished by the help of the monooxygenase hydroxylation system (Wasserbauer et al., 1990). Although many studies have proven the effect of P. Putida in degrading PE plastic, none have shown that it possesses PET degrading characteristics. The use of pseudomonas putida as a vector may be promising in the development of bacteria that break down PET.

5. Bacterial Transformation and Types of Vectors

Bacterial transformation is defined as the genetic alteration of a cell following uptake and integration of exogenous genetic material from its surrounding (Johnston et al., 2014).

It may be a natural phenomenon affected by environmental factors, or it may be induced inside a laboratory (Rahimzadeh et al., 2016). Laboratory-induced cellular transformation plays an important part in recombinant DNA technologies.

Bacterial transformation is achieved by the use of vectors, which are DNA carriers that are able to replicate in a host cell (Staworzynska et al., 2011). Such vectors are characterized by their small molecular size, the presence of an origin of replication termed “Ori”, and marker genes used for the selection of transformed bacteria (Nora et al., 2019).

There are several classes of vectors used such as cosmids, phages and plasmids (Griffiths et al., 1999). The choice of an appropriate cloning vector relies on several factors including insert size, vector size, restriction sites, copy number, cloning efficiency, and the ability to screen for inserts (Bajpai, 2014). Among all types of vectors available, plasmids are the most commonly used in bacterial transformations (Staworzynska et al., 2011).

Plasmids are circular, double-stranded DNA molecules that are normally present in bacteria and in some eukaryotic cells (Shintani et al., 2015) and can replicate independently of the host cell (Wegrzyn and Wegrzyn, 2002). They can range in size from few kb to around 100 kb (Lodish et al., 2000) and can carry up to 15 kb of fragments (Bajpai, 2014). Plasmid vectors must include the following: an origin of replication that is recognized by the cellular replication machinery and also determines the number of copies of a certain plasmid within the cell. In addition, an antibiotic resistance marker gene must be found that allows a selective choice of transformants; and multiple cloning sites that are recognized by different restriction enzymes (Couturier et al., 1988; Nora et al., 2019).

6. SEVA plasmids (pSEVA) and their Uses

Recombinant technological strategies are improving regularly, and plasmids’ use is specifically being refined continuously. Although studies show promising developments, plasmids nomenclature and standardization is not necessarily unified among the community of already existing users and available constructs (Silva-Rocha et al., 2013). A way to overcome such caveats, specific resource to implement a norm for the physical setup of vector plasmids and their nomenclature, the ‘Standard European Vector Architecture’ database (SEVA-DB,

20 http://seva.cnb.csic.es) was developed. This database provides a platform of functional sequences as well as definite available constructs, compatible with the old and novel cloning and DNA assembly techniques.

The many vectors found in the above mentioned database include broad- host-range origins of replication, antibiotic markers, expression systems and reporter genes that end with terminators and specific insertion sites. Under SEVA guidelines, each DNA part of the plasmid vector is edited in a way to reduce flaws in the sequences or function (Silva-Rocha et al., 2013). SEVA plasmids are annotated with a 3 digit number next to “pSEVA”. These numbers represent the selection marker, origin of replication and cargo segment respectively. SEVA plasmids structure and nomenclature are represented in Figure 5:

Figure 5: SEVA structure and nomenclature (Silva-Rocha et al., 2013): (A) SEVA plasmids consist of three elements: cargo (blue), replication origin (green) and an antibiotic marker (magenta). Enzymes that modify functional DNA segments are represented in the same color code. The modules are separated by three permanent parts that are shared between all SEVA vectors: the transcriptional terminators To and T1, and the conjugation origin oriT. (B) SEVA plasmids are named based on a 3-digit number. These numbers represent the selection marker, origin of replication and cargo segment respectively.

One SEVA plasmid capable of replicating in and transforming Pseudomonas bacteria (Pseudomonas putida and Pseudomonas syringae) is pSEVA441, based on previously published studies (Lauritsen et al., 2017; Batianis et al., 2020; Calero et al., 2020) due to its versatility and effectiveness in yield. This plasmid will be used for the experimental transformation of P. putida bacteria within the study.

Plastic is a versatile material whose cost effectiveness has made it very difficult on society to accept and incorporate another bio friendly substance in its place. The accumulation of plastic has led to disastrous amounts of pollution worldwide, and efforts to reduce the amount of such contaminations are being integrated frequently. Recycling is currently the most used form of plastic reprocessing; however, advanced practices need to be found and used. The urge of discovering and incorporating a new form of plastic solution stems from the vast increase of plastic pollution daily.

A promising technique that may prove more efficient than recycling may be the use of organisms in bioremediation techniques. I. sakaiensis is a newly found microorganism whose enzymes have been linked to plastic degradation. This bacterium possesses PETase enzyme which has been found to degrade

PET plastic, one of the major types of plastics responsible for plastic pollution.

Many studies have showed the importance of PETase enzyme and ways to

23 enhance its activity. Thus, I. sakaiensis may be used as a future solution to the world’s most difficult dilemma; however, this bacterium’s inability to survive in oceans and seas may limit its advantages.

Recombinant DNA technology may prove helpful in overcoming such an obstacle. This may be achieved through the use of a second bacterium known as

P. putida. P. putida has been found to survive in salt water and to possess the ability to degrade PE plastic, but no study has shown its effect on PET.

Recombinant DNA technology may be used to transform P. putida into PET degrading bacteria.

Since no work was conducted to show the effect of P. putida on PET plastic, this study was designed to use recombinant DNA technology to investigate the potential of integrating modified PETase genes derived from I. Sakaeinsis into P. putida. The main goal is to transform P. putida into bacteria that can survive in marine environments and also degrade PET. Though these are theoretical approaches displaying in details the techniques and tools required for such an achievement, we also theorized on the potential outcome of such experiments based on similar studies on other microorganisms.

III. Materials and Methods

1. Materials and Reagents

The materials and reagents used for the following experiments along with the suppliers are classified according to the application as following

- For cell culture: Pseudomonas putida AC 13 strain (ATCC 47054), Luria

Bertani (LB) broth medium (containing 10g/L tryptone, 10g/L NaCl, and

5g/L yeast extract), M9 minimal salt medium (containing 22 mM KH2PO4,

48 mM Na2HPO4,·7 H2O, 18.7 mM NH4Cl, 8.6 mM NaCl), Glucose,

MgSO4, thiamine hypochloride, CaCl2, 5-fluorouracil (Sigma-Aldrich

Chemical Co. via Ibra Haddad (Beirut, Lebanon)), Agar plates (TPP

Techno Plastic Products AG via Ibra Haddad (Beirut, Lebanon)).

- For cell transformation: pRT2210 plasmid ordered online from GenSmart

Design, LB broth medium and CaCl2 from Sigma-Aldrich Chemical Co. via

Ibra Haddad (Beirut, Lebanon).

- For selection of transformed bacterial cells: streptomycin (sm) from

Biowest, E.coli BL21, LB agar plates from Sigma-Aldrich Chemical Co.

both via Ibra Haddad (Beirut, Lebanon)

- For protein purification and western blot analysis: Radio immune-

precipitation (RIPA) buffer, Sodium Dodecyl Sulfate–polyacrelamide gels

(SDS-PAGE) buffer, Polyvinylidine difluoride (PVDF) membrane, sponge

and filter papers transfer buffer, Tris-Buffered Saline,0.1% Tween (TBST),

Horseradish peroxidase linked secondary antibodies from Bio-Rad

(Beirut, Lebanon)), anti-FLAG (anti-DYKDDDDK) and anti-

lysozyme, Chemiluminescence (ECL) plus kit from Sigma-Aldrich

Chemical Co. via Ibra Haddad (Beirut, Lebanon)).

- Gravimetric weight loss analysis: PET film from COPAR (Beirut, Lebanon),

SDS and ethanol from Bio-Rad (Beirut, Lebanon).

2. Pseudomonas Putida and Cell culture

The preparation of Pseudomonas Putida cell culture prior to bacterial transformation was performed as previously described by (Graf and

Altenbuchner, 2014). Briefly, a non-pathogenic strain of Pseudomonas Putida AC

13 was obtained, inoculated and maintained in LB broth as a complete growth

26 medium, and stored at 32°C for 12 hours. Streaking was then done on M9 minimal salt supplemented with 0.2 % glucose as a carbon source, 1 mM MgSO4,

−1 6 μM thiamine hydrochloride, 0.1 mM CaCl2, and 20 μg ml 5-fluorouracil). Using a sterile loop, bacterial suspension was taken from LB broth and streaked under sterile conditions on M9 minimal salt medium by the four quadrant technique.

Agar plates were then stored at 30°C for 48 hours.

3. Designing the pRT2210 plasmid

The construction of pSEVA441-egfp-PrpsL-PelB-IsPETase-FLAG-tag plasmid, whose alternative name given was pRT2210 plasmid, was performed by using the

GenSmart Design program (www.genscript.com). Briefly, the backbone DNA sequence of the 3629 bp pSEVA441 cloning vector was inserted first, and then genes were added simultaneously, by inserting their DNA sequence one after another in the forward sequence direction. Genes can be inserted next to each other, taking into consideration the usage of operons by bacteria.

The first restriction site used was NotI (GCGGCCGC) at position 46 bp of the cloning vector. Two nucleotides were added, “G&C”, before the start codon

“ATG”, after which the 1089 bp egfp gene was inserted. The 416 bp PrpsL gene responsible for PrpsL protein (a 30S subunit promoter) was added next, followed

27 by target gene’s IsPETase signal peptidePelB gene of 1037 bp length. The 1083 bp

IsPETase gene was then introduced and finally the 23 bp FLAG-tag sequence. The whole gene sequences were followed by another NotI restriction sites

(GCGGCCGC) at position 3716 bp, and 4 nucleotides were added “GTCG” before the “TGA” stop codon. To note that NdeI restriction site (CATATG) was directlt found before PETase gene.

The whole sequence was checked online by the GenSmart program for any possible errors within the gene. After validating the vector’s integrity, the entire

7225 bp pRT2210 plasmid, consisting of the backbone and gene inserts, was ordered online.

4. The Transformation of Pseudomonas putida AC 13 by pRT2210 plasmid

The already constructed pRT2210 plasmid was used in order to transform cultured Pseudomonas putida AC13 bacteria as described by (Zhang et al., 2016) and (Chakrabarty et al., 1975). Briefly, single colonies of P. putida AC 13 bacteria were isolated from the plated M9 minimal salt agar plate and inoculated into a fresh LB broth medium for 3 hours at 32°C. 1 mL of the bacterial suspension was transferred into a 2 mL sterile micro centrifuge tube by using a sterile pipette. 250

μL of 0.1 M transformation solution CaCl2 was added through a micropipette to the micro centrifuge tube containing the bacterial suspension and mixed by

28 repeatedly pipetting the mixture up and down several times. The cells were chilled on crushed ice for 20 min. The solution was then centrifuged for 5 min at maximum speed, and bacterial pellet was harvested. The pellet was resuspended in another 250 μL of CaCl2, and 0.2 mL of mixture was transferred to a new microtube. 0.1 mL of recombinant pRT2210 fusion solution was added to the microtube and the whole cell-DNA combination was incubated at 0°C for 30 min.

This mixture was then shocked by a heat pulse of 42°C for 2 min, and then transferred to an ice bath for 2 min. 1 mL LB Broth was added to cells and placed on a shaker at 37°C for an hour.

An important note to take into consideration is that the previously described bacterial transformation procedure was primarily performed in E.coli

BL21 strain before transforming P. putida AC-13. This is due to the fact that E.coli is used as a model system in bacterial transformation; these bacteria can rapidly divide every 20 min and have the ability to take up foreign DNA plasmids easily

(Lodish et al., 2000). This experiment was performed first in E.coli according to

(Rahimzadeh et al., 2016) in order to obtain enough quantity before being able to transfect P. putida AC-13 and test the effectiveness of PETase enyme on PET plastic.

5. Selecting Transformed Pseudomonas putida Bacteria

The selection of transformed P. putida bacteria by pRT2210 plasmid was done as described by (Zhang et al., 2016) and (Godeux et al., 2018). Briefly, 150 μL of cellular culture containing streptomycin (sm) resistant cells was pipetted and grown on LB agar plate with 50μg/mL streptomycin antibiotic and incubated for

48 h at 30°C.

Transformed cells were then selected through the observation of a yellow transparent halo produced around the bacteria on the agar plate due to the presence of the egfp gene incorporated within the plasmid and replicated in LB broth before every application. These transformed selected cells were then stored in batches in 70% glycerol at -80C and revived by resuspension in LB broth before each application.

6. Protein Purification and Western Blotting

The western blot analysis was performed in order to validate the presence of protein expression, and assess the presence of functional PETase enzyme produced by transformed P.putida bacterial cells. The process was carried out as described by (Xu et al., 2018) and (Gao et al., 2019). Briefly, 2  105 transformed cells were lysed on ice with a radio immune-precipitation buffer (RIPA), and

30 centrifuged at 13000 g speed at 4°C for 15 min. The supernatant was collected, mixed with sodium dodecyl sulfate–polyacrylamide gels (SDS-PAGE) buffer, and boiled for 5 min. Electrophoresis was performed based on previously published method (Mahmood and Yang, 2012) with a ready-made gel as described by (Lee et al., 2012).

Briefly, the running buffer was poured, and the gel was placed inside the electrophoresis device connected to a power supply. The comb was slowly removed and the marker of 6 μL was loaded carefully into the well, followed by

20 μL of the sample. The gel was run first with a low voltage of 60 V for 5 min (for stacking gel), and then at 140 V for 1 hour (for separating gel).

The gel was transferred onto a polyvinylidine difluoride (PVDF) membrane according to the following steps: Filter sheet was cut to fit the dimensions of the gel along with another PVDF membrane with the same measurements. The sponge and filter papers were soaked in transfer buffer, and the PVDF membrane in methanol. The gel was retrieved and a transfer sandwich was created with the sponge, filter papers, gel PVDF and filter papers in a way that no air bubbles were present. The sandwich is placed in the transfer apparatus along with transfer buffer and put on an ice block. The whole apparatus was transferred in a cold room at a constant current of 10 mA for 90 min.

The membranes were then blocked in 5 % non-fat milk powder for 2 h and probed with primary antibodies (anti-FLAG and anti-lysozyme as a positive control), in Tween 20/TBS (TBST) plus 5% milk overnight at 4°C. Negative control consisted of no proteins and no antibodies. The next day, after washing with TBST three times, PVDF membranes were incubated with Horseradish peroxidase linked secondary antibodies for 1 h at room temperature. Protein bands were then visualized by enhanced chemiluminescence (ECL) plus kit after incubating the membranes with ECL mix for 2 minutes by using a 1000 μL pipette. Results were visualized in a dark room.

7. Hydrolysis of PET films by transformed P. putida

In order to detect transformed P. putida PETase activity on PET, PET films were subjected to laboratory gravimetric weight loss determination as described previously (Ma et al., 2018). Briefly, 1.5 cm × 1.0 cm PET film were obtained, washed with 1% sodium dodecyl sulfate (SDS), 70% ethanol and distilled water, and then incubated in salt water for 10 min. The films were washed, weighed at zero, dried at 50 °C, and then placed again in a salt water bath (pH=8.4) inoculated with transformed P. putida colonies as the treatment group for 48 hours at 25°C. The process was then ended by draining the salt water and

32 exposing the films to 50°C for 10 min. Afterward, the PET films were washed, dried, and subjected to gravimetric weight-loss determination.

Two types of control were used in this experiment. The first negative control group included non-transformed P. putida AC 13 cells isolated from the previously prepared cell culture. These cells were inoculated within the salt water bath following the same experimental conditions as the treatment group. The second positive control consisted of already purified IsPETase enzymes that were added to the salt water bath within the same experimental setup as the other two groups. The PET film’s weights from the three groups were recorded before and after the conduction of the experiment.

8. Statistical analysis

Data were analyzed using PROC GLM in SAS 9.0 and data were reported as mean ± standard error of the mean (SEM). Each experiment was replicated in triplicate samples per run. Student’s t-test was used to determine mean differences and a p value < 0.05 was considered significant.

IV. Results

1. Identification of Transformed Pseudomonas putida

The vector results here as depicted in Figure 6 followed by the white and fluorescent colonies

Figure 6: The constructed pRT2210 plasmid that may be used for the transformation of P.putida AC 13.

After transforming P. putida AC13 cells with pRT2210 plasmid, cells were grown on LB agar plate with streptomycin antibiotic. Vector-containing P. putida cells all exhibit streptomycin resistance and will grow on the agar plate, while the egfp gene containing transformed bacteria may produce a yellow transparent

34 halo around the colony. These colonies will shine with green fluorescence under

UV light, further validating the presence of the egfp gene, adjacent to the

IsPETase, the plastic degrading gene. Expected results may be as seen in figure 7.

Figure 7: White and fluorescent colonies. To the left, vector-containing streptomycin resistant P. putida may grow on agar plate in the presence of streptomycin antibiotics, but do not fluoresce if not transformed. To the right, only successfully transformed bacteria may grow and fluoresce under UV light due to the presence of egfp gene adjacent to PETase gene. (These pictures were obtained in the Microbial Biotechnology course for similar transformation)

2. Protein Purification and Western Blot

In order to validate the production of functional PETase enzyme by P. putida bacteria, cells were lysed and proteins were purified through the process of gel electrophoresis. The detection of the enzymes may be done through

Western Blot analysis. Chemiluminescent bands can be present between 40 and

55 kDa (Moog et al., 2019) indicating FLAG-tagged PETase proteins and thus may prove the formation of these enzymes by the transformed P. putida cells. The anticipated results are depicted in Figure 8.

Figure 8: Anticipated Western Blot Analysis results. Cartoon representation showing from left to right the following: ladder, experimental well with the 40-55 kDa FLAG-tagged PETase protein, positive control consisting of lysozyme detected by anti-lysozyme antibodies, and a negative control with no proteins.

3. Hydrolysis of PET plastic by PETase enzyme from transformed P. putida

cells

The gravimetric weight loss determination was used to reveal functional

Pseudomonas putida PETase activity on PET plastic. After PET films are placed in salt water within the appropriate experimental group, PET film masses may be recorded after 48 hours. The expected results might be as follows regarding the three experimental groups used.

- Group 1: Negative control group that consisted of non-transformed P.

putida AC 13 cells: PET film might show no obvious weight loss, due to the

inability of these non-transformed cells to produce any PETase enzyme.

- Group 2: Positive control group that consisted of already purified PETase

enzymes: PET film might show a significant weight loss due to the

functional PETase enzymatic degrading ability against PET.

- Group 3: Treatment group that consisted of transformed P.putida bacteria:

PET film might as well show a significant weight loss. This could be due to

the ability of transformed P. putida bacteria to produce PETase enzymes

that are functional, in salt water, against PET films.

V. Discussion

The many uses of plastic have been a shield from the pollution that results from the usage of this material. Plastic accumulation in the environment causes serious threat to the well-being of plants, animals, and habitats (Barnes et al.,

2009; Worm et al., 2017; Biundo et al., 2018). For years, the world has been trying to find effective ways to deal with plastic pollutions, but none have proven effective in the long run. PET is among the most common types of plastics, and ways to completely eliminate this material from the environment are still primitive.

A PET plastic eating bacterium, Ideonella Sakaiensis (Yoshida et al., 2016), may prove promising in bioremediation through the action of its enzymes PETase and MHETase (Ramirez-Ekner et al., 2017).

Several studies have been conducted in order to shed light on the importance of such enzymes. Some results revealed the successful bio-degrading ability of PETase on other types of polyesters (Liu et al., 2018; Sagong et al., 2020)

(Eerhart et al., 2012). Moreover, PETase activity was enhanced through the addition of salts and glycerol in moderate amount (Cui et al., 2019), and its degrading ability was promoted through some binding cleft mutations (Bollinger et al., 2020).

Many researches focused on enhancing PETase activity either by using a secretion system based on the fusion of Sec-dependent and SRP-dependent signal peptides (maltose/maltodextrin binding periplasmic protein (SPMalE), maltoporin

(SPLamB) obtained from E. coli), or by relying on certain mutations within

IsPETase gene which lead to the formation of more space on several binding centers (Liu et al., 2018). Moreover, the genetically modified microalgae P. tricornutum with I. Sakaiensis PETase successfully produced PETase

(PETaseR280A-FLAG) that was capable of degrading PET in salt water. (Moog et al., 2019).

Among the many bacteria capable of playing a role in bioremediation,

Pseudomonas Putida has been well known for its ability to survive in extreme environments such as salt water (Nikel and de Lorenzo, 2018). This bacteria has the ability to biodegrade polyethelyne (PE) plastic (Jailawi et al., 2015), and many studies have proved that P.putida may break down items such as plastic bags, polythene bags, plastic cups, and milk covers (Saminathan et al., 2014).

Although research has proved the effect of P. putida in degrading PE plastic, no evidence exists on whether or not it has PET degrading characteristics. One way to eliminate PET from salt water could be by transforming p. putida cells.

The above experiments have been designed in order to prove whether or not P. Putida can be transformed into bacteria capable of breaking down PET in salt water. This may be accomplished by pRT2210 plasmid that could: transform

P. putida cells into bacteria that fluoresce under UV light and grow in the presence of streptomycin, produce efficient 40-55 kD PETase enzymes, and cause a reduction in PET films’ weight in salt water.

VI. Conclusion

The world has long been working towards a solution to eliminate plastics quickly, effectively and safely. Recycling methods have been implemented as such, though none have been fully successful due to the diversity of plastic types, the lack of potential alternatives and the difficulty to remanufacture these plastics into other types. Although PET, the most abundant and widespread used plastic, is commonly recyclable; however its uses pose a serious threat to the environment. The discovery of Ideonella sakaiensis and the potential improvements herein discussed are promising as means to incorporate bioremediation into the problem of plastic pollution. This bacterium has been found to break down PET into TPA and EG through the use of the two enzymes

PETase and MHETase, respectively. These enzymes could be the answer to an age

40 long problem. Bacterial transformation of P. Putida cells by IsPETase gene could lead to the development of bacterial cells capable of surviving in salt water and breaking down PET. The studies that have further investigated applying genetic engineering strategies to these enzymes as listed in this paper show great potential for new engineered organisms or products. These new discoveries may be the gateway to help end the crisis of plastic pollution.

Tables

Table 1: The Seven Types of Plastics and Some of Their Major Characteristics.

Resin IDa / Toxins Potential Toxin Plastic Type Abbreviati Common usage Recyclingb Ref released effect on Beverage bottles, (Keresztes et al., 2009; So Polyethylene PET or Antimony 1 clothing and carpet Cancer +++ et al., 2016; Council, Terephthalate PETE trioxide fiber 2017). HDPE Estrogen- (So et al., 2016; Council, High-density Shopping bags, toys, Hormonal system 2 or PE- mimicking +++ 2017); (Burns, 2008) polyethylene detergent bottles imbalance HD chemicals Credit cards, water BPA, (So et al., 2016; Council, Cancer, allergies, Polyvinyl PVC or pipes , bath curtains, phthalates, 2017) 3 & hormonal ++ chloride V synthetic leather lead, dioxins, (Schuler, 2008; Lithner et imbalances products Hg, & Cd al., 2012; Wu et al., 2019) LDPE Cling film, bread (Maraschin, 2002; So et Low-density Irregular 4 or PE- bags, irrigation pipes, Low toxicity ++ al., 2016; Council, 2017) polyethylene hormone levels LD plastic bags Asthma & (Lithner et al., 2012; So et Straws, plant pots, Polypropylene 5 PP Low toxicity hormonal + al., 2016; Council, 2017) microwave containers imbalance CD cases, coat Nervous system, (Khaksar and Ghazi- Polystyrene 6 PS hangers, disposable Styrene lungs, liver, and +++ Khansari, 2009; So et al., cups and plates immune system 2016; Council, 2017) Reproductive (Schuler, 2008; So et al., system, behavior 2016; Council, 2017) Other /multi- CDs, medical storage 7 - BPA and immune - materials containers, eyeglasses function alteration (a) Resin identification number, (b) Recycling frequency from difficult (-) to occasional (+) or common (+++).

Table 2: Examples of microorganisms (fungi and bacteria) capable of degrading the seven types of plastic.

Resin Fungi capable of degradation Bacteria capable of degradation ID Penicillium funiculosum (Nowak et al., 2011) Thermobifida fusca (Müller et al., 2005) Thermobifida halotolerans (Ribitsch et al., 2012) Thermomonospora curvata (Wei et al., 2014) Saccharomonospora viridis (Kawai et al., 2014) 1 Comamonas testosterone F6 (Gong et al., 2018) Nocardia species (Sharon and Sharon, 2012) Streptomyces species (Farzi et al., 2019) Ideonella sakaiensis 201-F6 (Yoshida et al., 2016) Penicillium simplicissimum (Sowmya et al., 2015) Rhodococcus ruber (Orr et al., 2004) 2 Brevibacillus borstelensis (Hadad et al., 2005) Phanerochaete chrysosporium (Ali et al., 2014) Pseudomonas otitidis (Anwar et al., 2016) Lentinus tigrinus (Ali et al., 2014) Acanthopleuribacter pedis (Anwar et al., 2016) 3 Aspergillus niger (Ali et al., 2014) Bacillus cereus (Anwar et al., 2016) Aspergillus sydowii (Ali et al., 2014) Penicillium simplicissimum (Sowmya et al., 2015) Rhodococcus ruber (Orr et al., 2004) 4 Brevibacillus borstelensis (Hadad et al., 2005) Pseudomonas putida (Montazer et al., 2019) Phanerochaete chrysosporium (Jeyakumar et al., 2013) Stenotrophomonas panacihumi (Jeon and Kim, 2016) 5 Engyodontium album (Jeyakumar et al., 2013) 6 Cephalosporium species (Chaudhary and Vijayakumar, 2019) Rhodococcus ruber (Mor and Sivan, 2008) Cladosporium cladosporioides (Brunner et al., 2018) Bacillus pallidus (Tomita et al., 2003) 7 Xepiculopsis graminea (Brunner et al., 2018) Vibrio furnisii (Sudhakar et al., 2007) Penicillium griseofulvum (Brunner et al., 2018)

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