Identifying and Characterizing Novel Bacilli Capable of Degrading Recalcitrant Polymers

University of Tennessee, Knoxville TRACE: Tennessee Research and Creative Exchange

Doctoral Dissertations Graduate School

12-2019

Breaking the Chain: Identifying and Characterizing Novel Bacilli Capable of Degrading Recalcitrant Polymers

Kyle Bonifer University of Tennessee, [email protected]

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Recommended Citation Bonifer, Kyle, "Breaking the Chain: Identifying and Characterizing Novel Bacilli Capable of Degrading Recalcitrant Polymers. " PhD diss., University of Tennessee, 2019. https://trace.tennessee.edu/utk_graddiss/5738

This Dissertation is brought to you for free and open access by the Graduate School at TRACE: Tennessee Research and Creative Exchange. It has been accepted for inclusion in Doctoral Dissertations by an authorized administrator of TRACE: Tennessee Research and Creative Exchange. For more information, please contact [email protected]. To the Graduate Council:

I am submitting herewith a dissertation written by Kyle Bonifer entitled "Breaking the Chain: Identifying and Characterizing Novel Bacilli Capable of Degrading Recalcitrant Polymers." I have examined the final electronic copy of this dissertation for form and content and recommend that it be accepted in partial fulfillment of the equirr ements for the degree of Doctor of Philosophy, with a major in Microbiology.

Todd Reynolds, Major Professor

We have read this dissertation and recommend its acceptance:

Elizabeth Fozo, Gladys Alexandre, Jennifer Debruyn

Accepted for the Council:

Dixie L. Thompson

Vice Provost and Dean of the Graduate School

(Original signatures are on file with official studentecor r ds.) Breaking the Chain: Identifying and Characterizing Novel Bacilli Capable of Degrading Recalcitrant Polymers

A Dissertation Presented for the Doctor of Philosophy Degree The University of Tennessee, Knoxville

Kyle Sabastian Bonifer

December 2019

ACKNOWLEDGEMENTS

First, I would like to thank my advisor, Dr. Todd Reynolds, for all his help during my six years in graduate school. I have struggled tremendously throughout this process, having had many failed projects and ideas that did not come to successful fruition. Dr. Reynolds has been the most patient and understanding person that I could have hoped for, helping me find my way and allowing me to grow as a scientist. I will always hold him in the highest regard and strive to be as great a scientist and a person as he is.

In graduate school, I have had committee members and collaborators who have always been there to shed new light and provide perspectives on the problems I was trying to solve. My former and current committee members, Drs. Jeffrey Becker and Robert Wheeler, Elizabeth Fozo, Jennifer Debruyn and Gladys Alexandre have always been willing to take my questions and critique my work. I have learned a lot from them and would not have been able to complete this work without their help.

The people who I have worked with in the Reynolds lab have been a driving force for new ideas, collaboration and comedic relief. I would like to thank Bob Tams, Chelsi Cassilly, Tian Chen, Sahar Hasim, Sarah Davis, Andy Wagner, Billy Carver, Elise Phillips and Aeric Zhou for making the thousands of hours we’ve spend together memorable and enjoyable. I would also like to thank all the people in the Microbiology department for contributing to both my scientific and social development. I will miss the camaraderie and council I have had with all my peers, many of whom have always been willing to talk about science and life over beers.

I also want to extend a huge thank you to the love of my life, Tamara Demchenko, who has been with me since the beginning of my journey in graduate school. Whether it be her words of wisdom, love and affection or cleaning up my graphs and posters with her designer magic, she has contributed more to my success than anything else. Without her support and confidence in me, I don’t think I could have finished this process.

Finally, I’d like to send out a thank you to all my friends and family for their support, patience and understanding these past six years. No process like this can be completed by one person and having this group of people in my life has been key to my success.

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ABSTRACT

The development of plastic polymers has revolutionized modern society. These polymers are found ubiquitously in commodities and the production of these plastics is one of the most profitable industries in the world. However, the very characteristics that make plastics so universal and useful is also leading to detrimental ecological impacts. Due to their prevalence, finding novel microorganisms capable of breaking down these plastics is an important task. Another way these problems have are being addressed is by creating plastics that are more amenable to degradation. These plastics contain moieties more easily recognized by enzymes created by microbial life. Due to the increased adoption of these plastics, its important to understand their effect on the biology of microbial life.

In this dissertation we developed a biochemical assay to detect lactic acid that was being released as monomer from the biodegradable polymer polylactic acid due to bacterial degradation. We found a novel isolate of Bacillus pumilus (B12) that is capable of breaking down polylactic acid, with this degradation being able to robustly within 48 hours of incubation. Next we used our assay to observe how changing the nutrient composition allotted to B. pumilus B12 will alter its rate of PLA degradation. We saw increases in degradation when B. pumilus B12 was given maltose and mannitol as a sole carbon source, while observing decreases with adenine, potassium and phosphate supplementations. Next we used proteomic and transcriptomic tools to identify genes differentially expressed in conditions with altered PLA degradation. We found that in conditions with a lowered level of PLA degradation, genes associated with motility, chemotaxis and protein export are all downregulated.

Next, we investigated contamination found in a surfactant rich environment for novel bacteria capable of breaking down polyethylene and other recalcitrant carbon sources. We found six strains of novel Bacilli that were capable of growing on polysorbates, polyethylene glycol and mineral oil-rich in alkenes. Upon incubating one isolate of Bacillus megaterium with polyethylene films, we observed microbial growth, a lowered pH and a loss of tensile strength of the plastic. These two projects contribute new assays, bacterial isolates and insight to the field of polymer degradation.

TABLE OF CONTENTS

Chapter I: Introduction ...... 1

The Costs and Benefits of Man-Made Polymers ...... 2

Microbial-Derived Enzymatic Degradation of Natural Polymers ...... 4

Plastic Polymer Degradation...... 7

Summary ...... 14

References ...... 16

Chapter II: Bacillus pumilus B12 Degrades Polylactic Acid and Degradation is Altered by Changing Nutrient Conditions...... 33

Abstract ...... 35

Introduction ...... 36

Methods...... 37

Results ...... 40

Discussion ...... 71

References ...... 81

Appendix ...... 84

Chapter III: Bacillus pumilus B12 Alters its Gene Expression in Conditions that Reduce PLA Degradation ...... 88

Abstract ...... 90

Introduction ...... 91

Methods...... 92

Results ...... 97

Discussion ...... 115

References ...... 117

Appendix ...... 121

Chapter IV: Novel Bacillus Strains Cultured from Contaminated Polysorbate 40 Show High Tolerance to Various Surfactants with One Utilizing Polyethylene as a Sole Carbon Source ...... 129

Abstract ...... 131

Introduction ...... 132

Methods...... 133

Results ...... 136

Discussion ...... 145

References ...... 154

Appendix ...... 157

Chapter V: Conclusion and Future Directions ...... 165

Conclusion ...... 166

Future Directions ...... 167

References ...... 169

Vita ...... 172

LIST OF TABLES

Table 3. 1. Bacillus pumilus B12 is closely most closely related to other B. pumilus strains based on amino acid similarity...... 98

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LIST OF FIGURES

Figure 1. 1. Examples of the four different classifications of plastics based on biodegradability and production practices...... 9

Figure 1. 2. Examples of biodegradable and non-biodegradable plastic polymer degradation...... 10

Figure 2. 1. High levels of L-lactate release detected from PLA films exposed to Proteinase K...... 42

Figure 2. 2. Altering the temperature does not have any impact on Proteinase K driven L-lactate release from PLA...... 44

Figure 2. 3. Changing the pH of the media does not change detection of L-lactate from PLA films...... 45

Figure 2. 4. Adding glucose has no effect on the efficacy of the L-lactate assay...... 46

Figure 2. 5. Increasing salinity has no effect on L-lactate released from Proteinase K. ... 47

Figure 2. 6. Maximum likelihood phylogenetic tree of 16S rDNA for the environmental isolate B12 compared to other Bacillus spp...... 48

Figure 2. 7. Headspace CO2 concentrations after 16 days of culturing strain B12 in MSM with squares of biodegradable plastic mulch films...... 50

Figure 2. 8. High levels of L-lactate can be detected in 3-week microbial cultures when both glucose and PLA are added...... 52

Figure 2. 9. The addition of glucose significantly increases the release of L-lactate from PLA in microbial cultures...... 53

Figure 2. 10. PLA films show a decrease in elastic strength when incubated with B. pumilus B12...... 54

Figure 2. 11. Scanning electron microscopy shows PLA fracturing around B. pumilus B12 cells...... 55

Figure 2. 12. Bacillus pumilus B12 increases alters L-lactate release from PLA when grown in different carbon sources ...... 59

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Figure 2. 13. Bacillus pumilus B12 48-hour growth curves in different carbon sources.. 60

Figure 2. 14. Release of L-lactate from PLA incubated with B. pumilus B12 is independent of growth on different carbon sources...... 61

Figure 2. 15. Changing the primary nitrogen source to nitrate has no effect on PLA degradation by B. pumilus B12...... 62

Figure 2. 16. Increasing ammonium concentration in the media has no effect on PLA degradation by B. pumilus B12 ...... 63

Figure 2. 17. Increasing CSM concentrations available to B. pumilus B12 leads to a decrease in L- Lactate release from PLA...... 64

Figure 2. 18. Bacillus pumilus B12 has altered growth rates in different concentrations of CSM...... 65

Figure 2. 19. Bacillus pumilus B12 has decreased L-lactate release independent of growth

……...... 66

Figure 2. 20. Adenine is the component of CSM that is driving the decrease L-lactate release from PLA and Bacillus pumilus B12 culture...... 67

Figure 2. 21. B. pumilus B12 decreases L-lactate released from PLA in increased KH2PO4...... 68

Figure 2. 22. Bacillus pumilus B12 increases its growth with increasing concentrations of KH2PO4...... 69

Figure 2. 23. Bacillus pumilus B12 has reduced L-lactate release from PLA films with increasing concentrations of KH2PO4...... 70

Figure 2. 24. Increasing NaCl in the media has no effect on PLA degradation by B. pumilus B12...... 72

Figure 2. 25. B. pumilus B12 changes L-lactate release in K+ when normalized for growth...... 73

Figure 2. 26. Bacillus pumilus B12 growth rate changes with increases concentrations of potassium...... 74

Figure 2. 27. Bacillus pumilus L-lactate release from PLA changes in different concentrations of potassium independent of growth...... 75

Figure 2. 28. B. pumilus B12 increases L-lactate release from PLA films with increasing PO4- when growth is not considered...... 76

Figure 2. 29. Bacillus pumilus B12 changes its growth rate in different concentrations of PO4-...... 77

Figure 2. 30. Bacillus pumilus B12 has reduced L-lactate release from PLA films when there is - increased PO4 ...... 78

Figure S2. 1. Atomic Force Microscopy Data for B. pumilus B12 grown in KH2PO4. .... 85

- Figure S2. 2. Atomic Force Microscopy Data for B. pumilus B12 grown in PO4 ...... 86

Figure S2. 3. Atomic Force Microscopy Data for B. pumilus B12 grown in K+...... 87

Figure 3. 1. Bacillus pumilus B12 is similar to B. pumilus SAFR-032 at both a genomic and amino acid level...... 99

Figure 3. 2. Other strains of Bacillus pumilus have varying levels of PLA degradation as measured by L-lactate release...... 100

Figure 3. 3. Extracellular proteins released from B. pumilus B12 go down when incubated with complete supplement media (CSM) addition to the media...... 101

Figure 3.4. Mass spectrometry results of differentially abundant Bacillus pumilus B12 extracellular proteins...... 103

Figure 3. 5. PCA plot displays the clustering of samples within each condition...... 105

Figure 3. 6. RNA expression heatmap shows high similarity between replicates and divergence between variables...... 107

Figure 3. 7. Bacillus pumilus B12 differentially expresses the majority of its genes in MSM+G+CSM conditions compared to MSM+G conditions...... 108

Figure 3. 8. Number of biosynthesis associated genes differentially expressed sorted by into pathways using KEGG pathway mapper...... 109

Figure 3. 9. Number of metabolism associated. genes differentially expressed sorted by into pathways using KEGG pathway mapper...... 110

Figure 3. 10. Number of degradation associated genes differentially expressed sorted by into pathways using KEGG pathway mapper...... 111

Figure 3. 11. In CSM supplemented conditions, B. pumilus B12 significantly down regulates motility, chemotaxis and protein export pathways...... 113

Figure S3. 1. Extracellular proteins collected from B. pumilus B12 in conditions showing high L- lactate release from PLA films show no distinct band differences from standard MSM+G control...... 122

Figure S3. 2. Amino acid alignment of flagellin proteins identified in B. pumilus B12 show no homology to a previously identified proteolytic domain in C. haemolyticum Flagellin 123

Figure S3. 3. Biotinylation of surface associated proteins of B. pumilus B12 in different growth conditions show no obvious differences...... 125

Figure S3. 4. L-lactate metabolism/transport genes and mass spectrometry “hit” protein genes both show overall decrease in gene expression...... 126

Figure S3. 5. Live dead staining of B. pumilus B12 shows no large increase in dead cells in MSM+G conditions...... 127

Figure S3. 6. Bacillus pumilus B12 is able to take up plasmid DNA and express GFP...128

Figure 4. 1. Contamination of Polysorbate-40 (Tween-40) bottle...... 138

Figure 4. 2. Maximum likelihood phylogenetic tree of 16S rDNA for the environmental NB isolates compared to other Bacillus spp...... 139

Figure 4. 3. Growth analysis reveals a preference for NB strains to grow on Tween-40 over Tween- 80 and Tween-20...... 140

Figure 4. 4. NB isolates are capable of growing on polyethylene glycol in solid and liquid media...... 142

Figure 4. 5. Growth of NB strains in different fatty acids...... 143

Figure 4. 6. A Ratio of colony size to lipase and protease halo formation for NB strains over 7 days...... 146

Figure 4. 7. Live/Dead staining of NB strains grown on PE films show no large decrease in cell viability of the course of 8 weeks...... 147

Figure 4. 8. B. megaterium NB isolates lower the pH of the media when incubated with PE over the course of 8 weeks. Samples were collected in triplicate every 2 weeks for 8 weeks..148

Figure 4. 9. Scanning electron microscopy images of polyethylene inoculated with NB isolates over the course of 8 weeks...... 149

Figure 4. 10. Atomic force microscopy shows a general reduction in tensile strength for polyethylene incubated with NB10 isolate...... 151

Figure S4. 1. Growth on NB strains in 25% Tween-80 over 120 hours...... 158

Figure S4. 2. Example of growth inhibition of NB strains by sodium dodecyl sulfate (SDS)...... 159

Figure S4. 3. Analysis of NB growth on the surfactant Triton-X...... 160

Figure S4. 4. NB isolates can grow on mineral oil as a sole carbon source...... 162

Figure S4. 5. Atomic force microscopy data for NB6, NB12 and NB14 strains grown on polyethylene films for 6 weeks...... 163

Figure S4. 6. Biotinylated extracellular proteins collected from NB9 grown in both MSM+G and Tween...... 164

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CHAPTER 1: Introduction

The Costs and Benefits of Man-made Polymers

The advent of man-made plastic polymers has revolutionized industries ranging from office supplies to aerospace engineering. Plastic’s ability to be molded into objects makes its utility only limited to the imagination of the designers. This fact, coupled with the cheap cost of materials to produce plastics, has created a large market for their creation and implementation. The world’s global plastic industry has steadily grown at an average annual rate of 3% and is currently worth an estimated 1.1 trillion dollars [1]. This has led to an ever-increasing production capacity which has been estimated to be 348 million tons as recently as 2017 [2]. There are two main causes for concern for this massive industry: 1) The core material used for the vast majority of the plastics created is petroleum and 2) the innate resistance to degradation of these polymers leads to an accumulation in the environment. Both drawbacks are having a substantial impact on the health of the planet, effecting terrestrial and aquatic ecosystems through both climate change and waste. With the full ecological impact still unknown, it behooves the industry to begin implementing new technologies to help reduce plastic pollution and to limit the dependency of fossil fuels for production.

Recently, the production of plastics which are either bio-based or biodegradable has been increasing. Bio-based plastics are plastics that are derived from a renewable source such as crops, grasses or microorganisms [3,4,5]. Biodegradable plastics can be broken down rapidly in the environment from a combination of environmental weathering and microbial degradation. The key reason for their ability to be degraded is the recognition of bonds by microbially-based enzymes. This is contrast to non-biodegradable plastics which are high in in carbon-carbon bonds in their monomeric linkages, making them poorly recognized by microbially-synthesized extracellular enzymes [6,7]. The implementation of plastics with both of these characteristics can help alleviate some of the negative consequences of current petroleum-based plastics. Having a renewable source of plastic production will limit dependence on the finite supply of oil, while limiting the negative environmental impact associated with petroleum extraction [8,9,10]. Petroleum extraction alone can lead to the destruction of natural habitats, while the consequences of oil spills has led to some of the most devasting ecological destruction in the last 50 years [11]. The ability for plastics to degrade in the environment will also reduce the rate of plastic accumulation that is currently occurring. It is estimated that 99.5 million tons of plastic accumulate near coastlines each year,

2 with over 8 million tons entering open bodies of water [12,13]. These plastics will typically break down into small to microscopic particles called nurdles which have been discovered everywhere from the from to the ice sheets of Antarctica to the digestive tracts of marine life [14,15,16]. Little research has been done on the impact on terrestrial habitats, however, many of the threats being seen in oceanic environments are thought to exist on dry land as well [17].

While using crops as sources for the polymers may drive up food costs, the carbon footprint of collecting and producing the source materials is far lower than for petroleum-based plastics. For polyhydroxyalkanoates (PHA), the current energy needed for production is estimated to be 50.4 MJ/kg [5]. For Poly-lactic acid (PLA) the energy needed is currently around 55 MJ/kg [3]. However, next generation plastic manufacturing facilities have reported creating a kilogram of PLA for as low as 27.2 MJ, with the hope of reducing that number to 16.6 MJ/kg [3]. This contrasts with the two most common petro-polymers; polyethylene and polypropylene, which have energy costs of 80 and 77 MJ/kg respectively [18]. This reduction in energy production costs, coupled with less detrimental material extraction and product degradation, makes the shift in biodegradable polymers for certain uses attractive. This trend can be seen in the increasing production of the biodegradable polymer’s PLA and PHA from 0.7 million tons in 2014 to over 1.2 million tons by 2019 [2].

Although many of the benefits of polymers may seem attractive, there are several negative environmental effects associated with both the production and degradation of bio-based polymers. The source material of many of the bio-based polymers is from agricultural crops. This could lead to an exacerbation of an already daunting problem of feeding the world’s growing population [19]. Also, the cost of producing bio-based polymers is still higher than its petroleum counterparts, creating an economic barrier to large scale implementation. Additionally, although the reduction in plastics in the environment is positive, the by-products of degradation could still lead to damage to the ecosystems. The carbon that is trapped and resilient to release in inert versions of petro- polymers is broken down and utilized by microbial communities. The degradation of these polymers will result in increased biomass in soil environments [20,21]. This increase will not always have positive impacts, possibly leading to increases in both the acidification and eutrophication [22,23]. Another by-product of increased microbial activity is the production of both carbon-dioxide and methane which are released into the atmosphere. Many of the problems that stem from further adoption of biopolymers are similar to the atmospheric impacts of large- 3 scale agriculture [24]. When contemplating the comprehensive adoption of biopolymers, one must weigh both the positive and negative consequences of the new materials. Regardless of the costs and benefits of both petro-polymers and biopolymers and the microbial interactions with these materials require further investigation. This review will summarize how both naturally occurring and manmade polymers are recognized and degraded by microorganisms, how this degradation is measured and what the challenges are in discovering novel plastic degrading microbes and enzymes.

Microbial-Derived Enzymatic Degradation of Natural Polymers

Biomass Degradation

Biodegradation is the breakdown of large complex molecules to a mixture of smaller metabolites and biomass by bacteria, fungi and other organisms. This process is the driving force behind the recycling of carbon on the planet and serves as the vessel for the remediation of many toxic particles in the environment [25,26]. These processes are vital for sustainable ecosystems for the planet, allowing for the circulation of scarce nutrients and building blocks of life, and allowing recovery from the adverse effects of human industrialization.

Due to large proportions of carbon, nitrogen and other vital elements being trapped in large, complex structures in many terrestrial environments, degradation is necessary for the transport of molecules too large/charged to pass through the cell wall/membrane barrier [27]. This process is aided by abiotic factors such as photolysis, chemical hydrolysis and physical desiccation [28,29]. Next exoenzymes, which are enzymes extracellularly excreted by living organisms, target recognized chemical moieties to further break apart [30,21] . To reduce energetic needs of synthesizing new carbohydrates, lipids and amino acids, most exoenzymatic interactions focus on breaking monomeric molecules off from a larger structure [31]. These enzymes recognize the bonds connecting monomers together, often utilizing H2O to hydrolyze and re-arrange ester, ether, amide, peptide and polysaccharide bonds. Major classes of enzymes included in this process that are relevant to plastic degradation are proteases, lipases and amylases, with individuals in each category varying their level of target promiscuity on proteins, lipids and carbohydrates, respectively [32]. Other classes of enzymes that have been shown to be biologically relevant to

4 plastic degradation specifically include cutinases, tannases and laccases, which all aid in the breakdown of large complex molecular structures for the benefit of cellular life [33,34]

Proteases

Proteases are a major class of proteins that initiate the breakdown of other proteins into their amino acid components. They can be classified in several different ways including where in the amino acid chain they target bonds, their catalytic residues, specificity and proteolytic mechanism. Proteases that target bonds found on the ends of a protein chain are classified as exopeptidases while those that cleave bonds within a peptide chain are endopeptidases. The main difference between these two classes is that exopeptidases will create a monomeric by-product while endopeptidases will result in the creation of monomers and shorter polymer chains. Beyond the location of enzymatic reactivity, the residue that catalyzes the reaction is also important for classification. Most proteases rely on one or more specific amino acid residues in an active site to hydrolyze the peptide bond. These include serine, cysteine (thiol) and threonine proteases [35]. Another class is metalloproteases which uses a metal (usually zinc, manganese or cobalt), to initiate the breakdown of the peptide [36]. These enzyme groups have wide ranges of specificity with some being promiscuous not only against different types of peptides, but other polymeric structures sharing a recognition molecule [37,38]. Two examples of this are Trypsin and Proteinase K, two serine proteases that can degrade various types of proteins. Conversely, proteases can also target a specific amino acid sequences, usually through secondary protein structures reducing the access to the active site [39,40]. Finally, the mechanism behind cleaving the peptide bond varies between protease species. Typically, one of two mechanisms occur, either a one-step reaction or a two-step reaction. In one step reactions, the enzyme activates water through polarization which then attacks the peptide moiety. In two step reactions, the enzyme interacts directly with the substrate to break the bond, with the introduction of water used to free the enzyme from the c- terminal end.

Lipases

Lipids are one of the key building blocks of cellular life, comprising the barriers of membranes and acting as storage molecules for dense amounts of energy [41]. Due to their ubiquity and utility, microbial life has developed an array of enzymes to break down and integrate lipids into the cell. This large group of hydrolytic enzymes are lipases which most commonly target

5 bonds between the polar head and nonpolar tail of lipids. Most commonly the activity is against the ester bond of triacylglycerol (fats) which separates glycerol from the fatty acid chains [42,43]. Being such a large class of esterases means that various enzyme species of lipases exist, with some having broad activity against several different lipid moieties [44] and others being specific to a certain ester position or require a specific type of fatty acid tail for activity [45]. Lipases are molecularly and biochemically diverse molecules with the only known conserved region being a Gly-X-Ser-X-Gly pentapeptide that encapsulates the serine residue that is the active amino acid [46]. With few exceptions, all lipases fall into a range between 19-70 kDa [47,48] and tend to work optimally in an acidic pH range with optimal temperature ranges between 15-70℃ [49].

Amylases

Carbohydrates comprise the cell walls of bacteria (peptidoglycan), fungi (β-glucan and chitin) and plants (chitin) and are also the main carbon input into cell fermentation and respiration. The breakdown of long chains of these molecules is imperative for the survival of all life making the production of amylases ubiquitous. Amylases target the α-1,4,-glyocosidic bonds that hold together sugar molecules in the carbohydrate chains [50]. The end results of the hydrolysis reaction of a carbohydrate such as starch are chains of glucose of differing lengths ranging from maltose and maltotriose to short chain oligosaccharides [51]. Amylases are calcium metalloproteases, requiring a calcium ion to catalyze the reaction [50]. Amylases have varying degrees of promiscuity with α-amylases indiscriminately targeting regions in the polymer chain while β- and γ-amylases target specific locations on the carbohydrate [52].

Other Hydrolases

Like many protease and lipases, other hydrolases target ester bonds that hold molecules together with several of these showing activity against plastic polymers. Two examples of this are cutinases and tannases. Cutinases are enzymes that target the waxy plant material that makes up the cuticle known as cutin [53]. These hydrolases have also been shown to act upon a variety of polymers including triglycerides [53]. This target promiscuity has made them observed candidates for plastic degradation ranging from polyethylene terephthalate (PET) to polybutylene succinate (PBS) [54,55]. Tannases are hydrolytic enzymes that target the ester bonds in digallate to create gallate [56]. Digallates and gallates are common phenolic compounds contained in several plant species including oaks, witch hazel and sumac [57]. Due to the wide range of fungal species that

6 produce tannases, many of these enzymes have been identified with several that can degrade PET [58,59].

One example of an enzyme that does not have hydrolytic activity but has been observed to be involved in plastic degradation are laccases. Laccases are copper containing polyphenol oxidases that have been shown to oxidize both phenolic and non-phenolic lignin and other environmental pollutants [60,61]. Due to the diverse range of complex organic matter laccases are involved in degrading it is no surprise that laccases have been shown to degrade some of the most recalcitrant plastic polymers used in industry including polyethylene (PE) and polyvinylchloride (PVC) [62,63].

Plastic Polymer Degradation

Plastic polymers can be categorized in two major ways: 1) whether they are synthetic or bio-based and 2) whether they are biodegradable or non-biodegradable. Bio-based plastics are those created as natural products by animals, plants, fungi or bacteria [5,21,64]. These molecules are either synthesized as polymers by the organism or released as a monomer which can be processed into polymers through anthropomorphic processing [65]. This contrasts with synthetic polymers’ which are most commonly produced from petroleum precursors [18]. This classification of bio-based or synthetic is separate from the polymer’s susceptibility to degradation. Degradation of a polymer has an amorphous definition, with industries and governments differing on what the standard should be. The European Union classifies a polymer as biodegradable if less than 90% of the original mass of the polymer is left after 6 months [66], while many plastic companies used the ASTM standard of not seeing any visible traces of plastics over the same period as compost of organic materials [67]. Generally, the categorical distinction rests on the timeframe that a polymer will persist in the environment; with short time frames of degradation being considered “biodegradable”.

Of the four combinations of these categories, most plastics produced for commercial use are synthetic and non-biodegradable. This include both high- and low-density polyethylene (36%), polypropylene (21%), and polyvinylchloride (12%) (Figure 1.1) [68]. Altogether, this category makes up approximately 80% of all plastic produced globally [68]. Factoring in both synthetic and

7 bio-based non-biodegradable polymers increases this production number to roughly 92% of global production [68]. Both synthetic and bio-based nonbiodegradables have similar properties that make them more inert and recalcitrant, thus making them poor targets for microbial degradation. Plastics such as polyethylene and polypropylene and polystyrene consist entirely of carbon-carbon and carbon-hydrogen bonds. Lacking elements such as oxygen, nitrogen and sulfur in the molecular pattern diminishes enzymatic recognition [69]. This is the very reason bio-based and synthetic biodegradable polymers are broken down. Ester bonds found in poly-lactic acid (PLA), polyhydroxyalkanoate (PHA) and poly-3-hydroxybutyrate (PHB) allow for dehydration reactions to occur by esterases (Figure 1.1,1.2) [3,4,70]. Starch-based plastics can also be hydrolyzed at their 1,4-glycosidic bonds [71].

Although microbial degradation of “non-biodegradable plastics” is more difficult, it has still been observed and quantified. This process happens through the introduction of functional groups that increase the hydrophilicity of the polymer [72]. One mechanism microorganisms use to alter functional groups for metabolic processing is through oxidation (Figure 1.1). This introduces hydroxyl groups, ketones and aldehydes which are more readily targeted for enzymatic degradation [73,74]. By breaking down the oxidized groups in the chain, the polymer fractures and breaks down, increasing surface area. This surface area then allows for a more rapid degradation. This entire process is aided by abiotic factors including cold and heat stress, freezing and thawing tearing and photodegradation [64,75]. Photodegradation in particular, helps introduce oxygen into the polymer where UV energy creates free radicals [76]. Conversely, plastics treated with antioxidants and certain other plasticizers will reduce degradation by preventing oxidation of the carbon-carbon chain [77,78]. The introduction of these plasticizers along with high molecular weight and low molecular branching are the key reasons for the slow breakdown of commercially produced polyethylene and other nonbiodegradable polymers [72].

Biodegradation of polymers is traditionally measured through a variety of quantitative and qualitative techniques, looking at both changes in the microbial community and the plastic itself. One of the most widely utilized techniques for observing plastic degradation is measuring the dry weight loss of the plastic. This combined with measuring CO2 formation is indicative of plastic films being metabolized by microbial communities: The CO2 is a by-product of respiration and utilizing the long carbon chain as a carbon source [79]. This technique is typically performed

Figure 1. 1. Examples of the four different classifications of plastics based on biodegradability and production practices.

Figure 1. 2. Examples of biodegradable and non-biodegradable plastic polymer degradation. The top pathway illustrates the degradation of polylactic acid (PLA) and the bottom pathway shows the degradation of polyethylene.

10 in tandem with visual evidence that can be collected through scanning electron microscopy (SEM) and fluorescent microscopy. SEM allows for observing fracturing/cracking of the plastic as well as microbial adherence [80,81]. Fluorescent microscopy can be utilized by performing live/dead staining and visualize not only microbial adherence but survival on the plastic surface [82]. To give more quantitative measurements than SEM and fluorescent microscopy, atomic force microscopy (AFM) can be utilized to measure changes in the rigidity and tensile strength of the plastic [81]. The more that the polymer is broken down by abiotic and biotic factors, the more of reduction in rigidity and an increase in elasticity (measured by Young’s modulus) is observed [83]. Other techniques that measure changes in the properties of the plastic include Fourier-transform infrared spectroscopy (FTIR) and nuclear magnetic resonance spectroscopy (NMR) which look at chemical changes occurring to the polymer chain. FTIR and NMR can identify the introduction of different functional groups through new peak shifts from the original sample. Furthermore, change in absorbance intensity in FTIR is a good indicator of the amount of polymer being converted to other chemical moieties [84,85]. Another less common technique used is to radiolabel the carbon that makes up the backbone of the polymer chain and to track the radioactive carbon isotopes [86]. A combination of all these techniques have previously been used to measure the alteration and degradation of plastic polymers by microbial communities. Below are some examples of plastic polymers that have been researched with microbes and enzymes observed to lead to degradation.

Polyethylene (PE)

Polyethylene (PE) is the most common plastic polymer made in the world, accounting for

36% of global production [87]. The structure of these plastics is a long chain of carbons (C2H4)n that either have high or low degrees of side-chain branching. This level of branching has a large effect on the density of plastic and the level of crystallinity that can be attained [88]. As previously stated, PE does not meet the classification of a biodegradable plastic despite the evidence showing both bacterial and fungal species are able to grow on it as a sole carbon source. Multicellular species such as Galleria mellonella and the Indian meal moth larvae have also been shown to degrade PE with the aid of their gut microflora [89,90]. Bacteria observed to break down PE include Pseudomonas, Rhodococcus, Streptomyces, Bacillus, Penicillium and Aspergillus species showing that Gram-negative and Gram-positive organisms as well as fungi are viable options for studying degradation [91,92,93,94,95,96,97,98]. The key issue with studies performed on PE degradation is the lack of biochemical characterization of the process. It has been well supported

11 that abiotic factors will introduce linkage breaks and oxidize the plastic allowing for the attack by hydrolytic enzymes [99]. However, there has been little analysis of a biological mechanism for PE degradation without abiotic oxidation occurring. Furthermore, few studies have been performed on pure PE that has not been pretreated with plasticizers. Many of these plasticizers contain ester bonds, possibly skewing the data and leading to false positives [100,101]. Due to the prevalence of PE pollution in the environment, discovering and characterizing manipulatable biological systems for the degradation of PE is the “white unicorn” of microbial plastic degradation.

Polystyrene (PS)

Polystyrene (PS) is another hydrocarbon-based plastic that has a similar backbone to PE apart from an aromatic ring at one of the two carbon backbone positions. PS is a common plastic found in packaging, especially in its foam form [102]. Although chemically inert, it is readily solubilized by many types of organic solvents [103]. Although no enzyme has been linked to PS degradation, several microbes are able to use the monomer styrene as a sole carbon source, with the metabolic process being well characterized [104]. These bacteria include Pseudomonas, Rhodococcus and Corynebacterium species [105,106,107]. Of the strains known to utilize styrene as a carbon source, Pseudomonas putida and Rhodococcus zopfi have been shown to convert styrene oil into the biodegradable polymer polyhydroxyalkanoate (PHA). The key limiting factor to this process is the need for the PS to be broken down via pyrolysis before degradation or conversion into PHA. This process is energetically costly, so there is a need for discovery of new enzymes capable of breaking down large PS chains [106,108,109].

Polypropylene (PP)

The second most produced plastic in the world is polypropylene (PP), with 56 million tons being produced in 2018 with an estimated 88 million tons being produced by 2026 [110]. PP has a chemical structure like PE, except for a single methyl group branching off every other carbon in the backbone. PP has been shown to break down in a presence of mixed soil communities using weight loss measurements, however no isolated bacteria or enzymes have been identified that can degrade PP [111,112,113]. Something that causes a further reduction in degradation is are treated with phosphites that absorb UV energy and reduce their abiotic degradation [114].

Polyethylene Terephthalate (PET)

Colloquially known as polyester, polyethylene terephthalate (PET) is a common polymer used in many types of fibers used in clothing as well as for packaging similar to PE or PS. The backbone of PET is comprised of terephthalic acid and ethylene glycol that are combined through a process of esterification [115]. Although few, both microbes and enzymes have been identified that can degrade PET. Enzymes from Thermobifida sp. and Thermomonospora sp. produce serine hydrolases that are like many types of esterases [116,117,118].

Polyhydroxyalkanoate (PHA)

Polyhydroxyalkanoates (PHA) is a polyester created internally by microbes in response to non-carbon nutrient depletion as a way of storing essential excess carbon [119]. These polymers are considered elastomers with similar properties to the nonbiodegradable polypropylene. Due to their nature as biosynthetic products within microorganisms, these plastics are readily broken down by said microbes as a source of energy. The distribution of microbes capable of breaking down PHA are wide ranging, including soil, sewage, composting and marine environments [120]. PHA depolymerases has been isolated and characterized from Streptomyces, Pseudomonas, Alcaligenes and Ralstonia isolates among others, with the enzyme typically functioning optimally in a more basic pH [120].

Polybutylene Succinate (PBS)

Polybutylene Succinate (PBS) is an aliphatic polyester that is created from the combination of succinic acid and 1,4, butanediol monomers. PBS is a petroleum-based polymer that has similar mechanical properties to PET but is more amenable to biodegradation. Like many other ester- based polymers, PBS is capable of being acted upon by hydrolytic enzymes. Some examples of microbes observed to break down PBS include a species of the bacteria Amycolatopsis and a fungal isolate of Penicillium. Purified lipases from Candida anarctica and Pseudomonas cepacia were shown to lead to a 40% reduction in plastic mass loss with observable fracturing occurring within 24 days [121].

Polylactic Acid (PLA)

Polylactic acid is the second most common biodegradable polymer found in commercial plastics [122]. PLA is an aliphatic polyester formed from direct condensation of lactic acid

13 monomers or from ring polymerization of lactide [123]. PLA is a rigid thermoplastic which is allows it to be implemented in many scenarios where PS or PET would be used. The problem with this rigidity is the plastic can become brittle without the addition of plasticizers [124,125]. Several species of bacteria and fungi have been observed to degrade PLA in composting conditions incusing Fusarium moniliforme, Penicillium roquefort and Bacillus brevis [126,127]. Purified enzymes such as the serine protease Proteinase K isolated from Tritirachium album and a lipase from Rhizopius delemer have also been shown to break down PLA [128,129]. Although PLA has is classified as biodegradable, this process is slow outside of composting conditions with high temperature ranges [130]. Therefore, finding new enzymes and bacteria capable of degrading PLA in less industrial conditions is important.

Summary

Although microbial plastic degradation is a process that has been studied for decades, there are many questions that have been left unanswered. This is partially due to the recalcitrance and generally poor growth of microbial communities on the most common types of plastic. Also, many of the ways that degradation measurements are taken are slow, inaccurate, qualitative, expensive or low resolution. Other challenges include having plastics that are pure and not contaminated with some type of plasticizer, which can lead false positive identification of plastic degrading microbes. The lack of quick detectable indicators to show degradation also impinge the scientific communities’ ability to study both the biology of the organisms degrading the plastic, as well as the environmental conditions that can alter degradation rates. This makes looking at cellular regulatory mechanisms and their role in the bacteria’s ability to degrade polymers difficult. This also leads to a new question: How many bacteria and fungi can degrade plastics if given the proper conditions?

One of the two main goals of this dissertation was to develop a robust biochemical assay to help detect the degradation of polylactic acid from enzymes and bacterial isolates. After the development of said assay, we employed it to study the biology of an identified plastic degrading strain of Bacillus pumilus (B12) and to determine if nutrient/ environmental conditions influence the rates of degradation. After finding conditions that altered bacterial degradation, we utilized proteomics and transcriptomics to uncover metabolic and physiological differences that correlate

14 with plastic degradation. This work was the first step in understanding the regulatory mechanisms in bacteria that control plastic degradation.

The other goal of this dissertation is to find novel environments that might enrich for microbial communities more adept at degrading recalcitrant polymers. Here we discovered several Bacillus spp. that contaminated a bottle of the non-ionic surfactant, Tween-40. These Bacillus spp. could grow in high concentrations of various surfactants and were able to use several fatty acids, polyethylene glycol and mineral oil as sole carbon sources. The evidence of growth on these recalcitrant carbon sources led us to investigate their growth on high density polyethylene (HDPE). Four of these strains showed growth and metabolic activity on HDPE and the HDPE lost tensile strength over the course of 8 weeks. This work indicates that microbes capable of growing in environments rich in fatty acids and other recalcitrant carbon are good candidates to study as possible plastic degraders.

These projects try to build upon previous research in the field , providing both tools and new microbial isolates to help study the breakdown of plastics. The assay developed can allow for quick and simple identification of both enzymes and microbes that are capable of degrading PLA, a plastic that is becoming more prevalent in industrial applications. Furthermore, we’ve taken the first steps in looking into the regulatory mechanisms that can alter the rate of plastic degradation in Bacillus pumilus. This work is coupled with our investigation of a surfactant environment enriching for microbes that can breakdown HDPE. Studying the isolates from this environment could elucidate novel mechanisms needed to grow in this environment that are beneficial for polymer degradation.

References

1. Company, T. B. R. (2018). Plastics and Polymers Global Market Briefing 2018

2. Bioplastics, E. (2017). 2020 Bioplastics Market Forcast. 10th European Bioplastics Conference in Berlin.

3. Erwin T.H. Vinka, K. R. R. b., David A. Glassnerb, Patrick R. Gruberb. (2003). Applications of life cycle assessment to NatureWorks polylactide (PLA) production. Polymer Degradation and Stability, 80, 403-419.

4. Lu, J., Tappel, R. C., & Nomura, C. T. (2009). Mini-Review: Biosynthesis of

Poly(hydroxyalkanoates). Polymer Reviews, 49(3), 226-248. doi:10.1080/15583720903048243

5. Takeharu Tsugea, Y. K., Yoshiharu Doia. (2004). Microbial synthesis and

Enzymatic degradation of renewable poly[(R)-3-hydroxybutyrate-co-(R)-3 hydroxyhexanoate]. Science and Technology of Advanced Materials, 5, 449-453.

6. Shah, A., Hasan, F., Hameed, A., & Ahmed, S. (2008). Biological Degradation of

Plastics: A Comprehensive Review. Biotechnology advances, 26, 246-265. doi:10.1016/j.biotechadv.2007.12.005

7. Amass, W., Amass, A., & Tighe, B. (1998). A review of biodegradable polymers: uses, current developments in the synthesis and characterization of biodegradable polyesters, blends of biodegradable polymers and recent advances in biodegradation studies. Polymer International, 47(2), 89-144. doi:10.1002/(sici)1097

0126(1998100)47:2

8. Teal, J. M., & Howarth, R. W. (1984). Oil spill studies: A review of ecological effects. Environmental Management, 8(1), 27-43. doi:10.1007/BF01867871

9. Fiori, S. M., & Zalba, S. M. (2003). Potential impacts of petroleum exploration and Exploitation on biodiversity in a Patagonian Nature Reserve, Argentina. Biodiversity

&Conservation, 12(6),1261-1270. doi:10.1023/a:1023091922825

10. Pezeshki, S. R., Hester, M. W., Lin, Q., & Nyman, J. A. (2000). The effects of oil spill and clean-up on dominant US Gulf coast marsh macrophytes: a review.

Environmental Pollution, 108(2), 129-139. doi:https://doi.org/10.1016/S0269

7491(99)00244-4

11. Venturini, N., Muniz, P., Bícego, M. C., Martins, C. C., & Tommasi, L. R.

(2008). Petroleum contamination impact on macrobenthic communities under the influence of an oil refinery:Integrating chemical and biological multivariate data.

Estuarine, Coastal and Shelf Science,78(3), 457-467. doi:https://doi.org/10.1016/j.ecss.2008.01.008

12. Jambeck, J. R., Geyer, R., Wilcox, C., Siegler, T. R., Perryman, M., Andrady, A.,

. . Law,K. L. (2015). Plastic waste inputs from land into the ocean. Science, 347(6223),

768-771.doi:10.1126/science.1260352

13. Eriksen, M., Lebreton, L. C. M., Carson, H. S., Thiel, M., Moore, C. J., Borerro,

J.C., . . .Reisser, J. (2014). Plastic Pollution in the World's Oceans: More than 5 Trillion

Plastic Pieces Weighing over 250,000 Tons Afloat at Sea. PLOS ONE, 9(12), e111913. doi:10.1371/journal.pone.0111913

14. Bergmann, M., Mützel, S., Primpke, S., Tekman, M. B., Trachsel, J., & Gerdts, G. (2019). White and wonderful? Microplastics prevail in snow from the Alps to the Arctic. Science Advances, 5(8), eaax1157. doi:10.1126/sciadv.aax1157

15. Duncan, E. M., Arrowsmith, J. A., Bain, C. E., Bowdery, H., Broderick, A. C.,

Chalmers, T., . . .Godley, B. J. (2019). Diet-related selectivity of macroplastic ingestion in greenturtles (Chelonia mydas) in the eastern Mediterranean. Sci Rep, 9(1), 11581. doi:10.1038/s41598-019 48086-4

16. Scanes, E., Wood, H., & Ross, P. (2019). Microplastics detected in haemolymph

17 of the Sydney rock oyster Saccostrea glomerata. Mar Pollut Bull, 149, 110537. doi:10.1016/j.marpolbul.2019.110537

17. De Souza Machado, A. A., Kloas, W., Zarfl, C., Hempel, S., & Rillig, M. C.

(2018). Microplastics as an emerging threat to terrestrial ecosystems. Global Change

Biology,

24(4), 1405-1416. doi:10.1111/gcb.14020

18. Harding, K., Dennis, J., Von Blottnitz, H., & Harrison, S. (2007). Environmental analysis of plastic production processes: comparing petroleum-based polypropylene and polyethylene with biologically-based poly-β-hydroxybutyric acid using life cycle analysis. Journal of biotechnology, 130(1), 57-66.

19. Meyer, W. B., & Turner, B. L. (1992). Human population growth and global land use/cover change. Annual review of ecology and systematics, 23(1), 39-61.

20. Lin, Y.-B., Xue, Q.-H., & Yan, X. (2008). Effect of mulching mode and wheat root on soil microbial flora. Chin. J. Eco-Agric, 16, 1389-1393.

21. Liu, J., Zhan, A., Chen, H., Luo, S., Bu, L., Chen, X., & Li, S. (2015). Response of nitrogen use efficiency and soil nitrate dynamics to soil mulching in dryland maize

(Zeamays L.) fields. Nutrient Cycling in Agroecosystems, 101.doi:10.1007/s10705-015

96785

22. Chen, D., Lan, Z., Hu, S., & Bai, Y. (2015). Effects of nitrogen enrichment on belowground communities in grassland: Relative role of soil nitrogen availability vs. soil acidification. Soil Biology and Biochemistry, 89, 99-108. doi:https://doi.org/10.1016/j.soilbio.2015.06.028

23. Zhang, Y., Wang, L., Hu, Y., Xi, X., Tang, Y., Chen, J., . . . Sun, Y. (2015).

Water Organic Pollution and Eutrophication Influence Soil Microbial Processes,

Increasing Soil Respiration of Estuarine Wetlands: Site Study in Jiuduansha Wetland.

PLOS ONE, 10(5), e0126951.doi:10.1371/journal.pone.0126951

24. Tilman, D. (1999). Global environmental impacts of agricultural expansion: The need for sustainable and efficient practices. Proceedings of the National Academy of

Sciences, 96(11), 5995. doi:10.1073/pnas.96.11.5995

25. Ritter, W. F., & Scarborough, R. W. (1995). A review of bioremediation of contaminated Soils and groundwater. Journal of Environmental Science and Health .Part

A: Environmental Science and Engineering and Toxicology, 30(2), 333-357. doi:10.1080/10934529509376204

26. Dzionek, A., Wojcieszyńska, D., & Guzik, U. (2016). Natural carriers in bioremediation: A review. Electronic Journal of Biotechnology, 23, 28-36. doi:https://doi.org/10.1016/j.ejbt.2016.07.003

27. Yang, N. J., & Hinner, M. J. (2015). Getting across the cell membrane: an overview for small molecules, peptides, and proteins. Methods in molecular biology

(Clifton, N.J.),1266, 29 53. doi:10.1007/978-1-4939-2272-7_3

28. Mood, S. H., Golfeshan, A. H., Tabatabaei, M., Jouzani, G. S., Najafi, G. H.,

Gholami, M., &Ardjmand, M. (2013). Lignocellulosic biomass to bioethanol, a comprehensive review with a focus on pretreatment. Renewable and sustainable energy reviews, 27, 77-93.

29. Benítez, A., Sánchez, J. J., Arnal, M. L., Müller, A. J., Rodríguez, O., & Morales,

G.(2013). Abiotic degradation of LDPE and LLDPE formulated with a pro-oxidant additive. Polymer Degradation and Stability, 98(2), 490-501. doi:https://doi.org/10.1016/j.polymdegradstab.2012.12.011

30. Arnosti, C. (2010). Microbial Extracellular Enzymes and the Marine Carbon

Cycle. Annual Review of Marine Science, 3(1), 401-425. doi:10.1146/annurev-marine

120709142731

31. Gurung, N., Ray, S., Bose, S., & Rai, V. (2013). A broader view: microbial enzymes and their relevance in industries, medicine, and beyond. Biomed Res Int, 2013,

329121.doi:10.1155/2013/329121

32. Lopez-Iglesias, M., & Gotor-Fernandez, V. (2015). Recent Advances in

Biocatalytic Promiscuity: Hydrolase-Catalyzed Reactions for Nonconventional

Transformations. Chem Rec, 15(4), 743-759. doi:10.1002/tcr.201500008

33. Danso, D., Chow, J., & Streit, W. R. (2019). Plastics: Environmental and

Biotechnological Perspectives on Microbial Degradation. Applied and Environmental

Microbiology, 85(19), e01095-01019. doi:10.1128/AEM.01095-19

34. De las Rivas, B., Rodríguez, H., Anguita, J., & Muñoz, R. (2019). Bacterial tannases: classification and biochemical properties. Applied Microbiology and Biotechnology, 103(2), 603- 623. doi:10.1007/s00253-018-9519-y

35. Oda, K. (2011). New families of carboxyl peptidases: serine-carboxyl peptidases and glutamic peptidases. The Journal of Biochemistry, 151(1), 13-25. doi:10.1093/jb/mvr129

36. Birkedal-Hansen, H., Moore, W. G., Bodden, M. K., Windsor, L. J., Birkedal

Hansen, B.,

37. DeCarlo, A., & Engler, J. A. (1993). Matrix metalloproteinases: a review. Crit

Rev Oral BiolMed, 4(2), 197-250. doi:10.1177/10454411930040020401

38. Hedstrom, L. (2002). Introduction: proteases. In: ACS Publications.

39. Schauperl, M., Fuchs, J. E., Waldner, B. J., Huber, R. G., Kramer, C., & Liedl, K.

R. (2015).Characterizing Protease Specificity: How Many Substrates Do We Need?

PLOS ONE, 10(11),e0142658. doi:10.1371/journal.pone.0142658

40. Renicke, C., Spadaccini, R., & Taxis, C. (2013). A Tobacco Etch Virus Protease with Increased Substrate Tolerance at the P1' position. PLOS ONE, 8(6), e67915. doi:10.1371/journal.pone.0067915

41. Harwood, J. (2012). Lipids in plants and microbes: Springer Science & Business

Media.

42. Laachari, F., El Bergad, F., Sadiki, M., Sayari, A., Bahafid, W., Elabed, S., . . .

Ibnsouda, S. K.(2015). Higher tolerance of a novel lipase from Aspergillus flavus to the presence of free fatty acids at lipid/water interface. African Journal of Biochemistry

Research, 9(1), 9-17.

43. Lee, L. P., Karbul, H. M., Citartan, M., Gopinath, S. C., Lakshmipriya, T., &

Tang, T.-H.(2015). Lipase-secreting Bacillus species in an oil-contaminated habitat: promising strains to alleviate oil pollution. BioMed research international, 2015.

44. Hasan, F., Shah, A. A., & Hameed, A. (2006). Industrial applications of microbial lipases. Enzyme and Microbial technology, 39(2), 235-251.

45. Jensen, R. G., deJong, F. A., & Clark, R. M. (1983). Determination of lipase specificity. Lipids,18(3), 239-252. doi:10.1007/bf02534556

46. Brady, L., Brzozowski, A. M., Derewenda, Z. S., Dodson, E., Dodson, G., Tolley,

S., . . . Menge,U. (1990). A serine protease triad forms the catalytic centre of a triacylglycerol lipase. Nature,343(6260), 767-770. doi:10.1038/343767a0

47. Gricajeva, A., Bendikienė, V., & Kalėdienė, L. (2016). Lipase of Bacillus

Stratosphericus L1:Cloning, expression and characterization. International journal of

Biological macromolecules,92, 96-104.

48. Ramakrishnan, V., Balakrishnan, B., Rai, A. K., Narayan, B., & Halami, P. M.

(2012). Concomitant production of lipase, protease and enterocin by Enterococcus

21 faecium NCIM5363 and Enterococcus durans NCIM5427 isolated from fish processing waste. International Aquatic Research, 4(1), 14.

49. Javed, S., Azeem, F., Hussain, S., Rasul, I., Siddique, M. H., Riaz, M., . . .

Nadeem, H. (2017). Bacterial lipases: A review on purification and characterization.

Progress in Biophysics and Molecular Biology, 132. doi:10.1016/j.pbiomolbio.2017.07.014

50. De Souza, P. M., & de Oliveira Magalhães, P. (2010). Application of microbial α amylase in industry - A review. Brazilian journal of microbiology : [publication of the

Brazilian Society for Microbiology], 41(4), 850-861. doi:10.1590/S1517

83822010000400004

51. Whitcomb, D. C., & Lowe, M. E. (2007). Human pancreatic digestive enzymes.

Dig Dis Sci,52(1), 1-17. doi:10.1007/s10620-006-9589-z

52. Saranraj, P. (2013). Bacterial Amylase: A Review. International Journal of

Pharmaceutical &Biological Archives, 4.

53. Chen, S., Su, L., Chen, J., & Wu, J. (2013). Cutinase: Characteristics, preparation, and application. Biotechnology advances, 31(8), 1754-1767.

54. Carniel, A., Valoni, É., Nicomedes, J., Gomes, A. d. C., & Castro, A. M. d. (2017). Lipase from Candida antarctica (CALB) and cutinase from Humicola insolens act synergistically for PET hydrolysis to terephthalic acid. Process Biochemistry, 59, 84- 90.doi:https://doi.org/10.1016/j.procbio.2016.07.023

55. Maeda, H., Yamagata, Y., Abe, K., Hasegawa, F., Machida, M., Ishioka, R., . . .

Nakajima, T.(2005). Purification and characterization of a biodegradable plastic degrading enzyme from Aspergillus oryzae. Applied Microbiology and Biotechnology,

67(6), 778788.doi:10.1007/s00253-004-1853-6

56. Mehta, M., Muddapur, U., & Priya, V. S. (2013). Fungal production of tannase: a

22 review.Int. J.Sci. Eng. Technol, 2(8), 752-755.

57. Fernandes, F. H., & Salgado, H. R. (2016). Gallic Acid: Review of the Methods of Determination and Quantification. Crit Rev Anal Chem, 46(3), 257-265. doi:10.1080/10408347.2015.1095064

58. Roth, C., Wei, R., Oeser, T., Then, J., Föllner, C., Zimmermann, W., & Sträter, N.

(2014). Structural and functional studies on a thermostable polyethylene terephthalate degrading hydrolase from Thermobifida fusca. Applied Microbiology and Biotechnology,

98(18), 78157823.

59. Aguilar, C. N., Augur, C., Favela-Torres, E., & Viniegra-González, G. (2001).

Production of tannase by Aspergillus niger Aa-20 in submerged and solid-state fermentation: influence of glucose and tannic acid. Journal of Industrial Microbiology and Biotechnology, 26(5), 296-302.doi:10.1038/sj.jim.7000132

60. Hüttermann, A., Majcherczyk, A., Braun-Lüllemann, A., Mai, C., Fastenrath, M.,

Kharazipour,A., . . . Hüttermann, A. (2000). Enzymatic activation of lignin leads to an unexpected copolymerization with carbohydrates. Naturwissenschaften, 87(12), 539-541.

61. Prajapati, H. V., & Minocheherhomji, F. P. (2018). Laccase-A Wonder Molecule:

A Review of its Properties and Applications. Int. J. Pure App. Biosci, 6(1), 766-773.

62. Sumathi, T., Viswanath, B., Sri Lakshmi, A., & SaiGopal, D. (2016). Production of laccase by Cochliobolus sp. isolated from plastic dumped soils and their ability to degrade low molecular weight PVC. Biochemistry research international, 2016.

63. Ojha, N., Pradhan, N., Singh, S., Barla, A., Shrivastava, A., Khatua, P., . . . Bose,

S. (2017). Evaluation of HDPE and LDPE degradation by fungus, implemented by statistical optimization. Scientific Reports, 7, 39515. doi:10.1038/srep39515

64. Swift. (1997). Non-medical biodegradable polymers: environmentally degradable polymers. Handbook of Biodegradable Polymers.Amsterdam:HarwoodAcademic, 473-511.

65. Babu, R. P., O'Connor, K., & Seeram, R. (2013). Current progress on bio-based polymers and their future trends. Progress in biomaterials, 2(1), 8-8. doi:10.1186/2194

0517-2-8

66. Vert, M., Doi, Y., Hellwich, K.-H., Hess, M., Hodge, P., Kubisa, P., . . . Schué, F.

(2012).Terminology for biorelated polymers and applications (IUPAC Recommendations

(2012). Pure and Applied Chemistry, 84(2), 377-410.

67. Alexander, M. (1999). Biodegradation and bioremediation: Gulf Professional

Publishing.

68. Geyer, R., Jambeck, J. R., & Law, K. L. (2017). Production, use, and fate of all plastics ever made. Science Advances, 3(7), e1700782. doi:10.1126/sciadv.1700782

69. Luckachan, G. E., & Pillai, C. K. S. (2011). Biodegradable Polymers- A Review on Recent Trends and Emerging Perspectives. Journal of Polymers and the Environment,

19(3), 637-676. doi:10.1007/s10924-011-0317-1

70. Altaee, N., El-Hiti, G. A., Fahdil, A., Sudesh, K., & Yousif, E. (2016).

Biodegradation of different formulations of polyhydroxybutyrate films in soil.

SpringerPlus, 5(1), 762-762. doi:10.1186/s40064-016-2480-2

71. Gould, J. M., Gordon, S. H., Dexter, L. B., & Swanson, C. L. (1990).

Biodegradation of Starch Containing Plastics. In Agricultural and Synthetic Polymers

(Vol.433, pp. 65-75): American Chemical Society.

72. Mierzwa-Hersztek, M., Gondek, K., & Kopeć, M. (2019). Degradation of

Polyethylene and Biocomponent-Derived Polymer Materials: An Overview. Journal of

Polymers and the Environment, 27(3), 600-611. doi:10.1007/s10924-019-01368-4

73. Iram, S. H., & Cronan, J. E. (2006). The β-Oxidation Systems of Escherichia coli and Salmonella enterica Are Not Functionally Equivalent. J Bacteriol, 188(2), 599-608. doi:10.1128/jb.188.2.599-608.2006

74. Fujita, Y., Matsuoka, H., & Hirooka, K. (2007). Regulation of fatty acid metabolism in bacteria. Mol Microbiol, 66(4), 829-839. doi:10.1111/j.1365

2958.2007.05947.x

75. Hamid, S. H. (2000). Handbook of polymer degradation: CRC Press.

76. Yousif, E. a. H., Raghad. (2013). Photodegradation and photostabilization of polymers, especially polystyrene: review. SpringerPlus, 2, 398-398. doi:10.1186/2193

1801-2-398

77. Sejidov, F. T., Mansoori, Y., & Goodarzi, N. (2005). Esterification reaction using solid heterogeneous acid catalysts under solvent-less condition. Journal of Molecular Catalysis A: Chemical, 240(1-2), 186-190

78. Vieira, M. G. A., da Silva, M. A., dos Santos, L. O., & Beppu, M. M. (2011). Natural- based plasticizers and biopolymer films: A review. European Polymer Journal, 47(3), 254-263. doi:https://doi.org/10.1016/j.eurpolymj.2010.12.011

79. Mohan, K. (2011). Microbial deterioration and degradation of polymeric materials. Journal of Biochemical Technology, 2(4), 210-215.

80. Ikada, E. (1999). Electron Microscope Observation of Biodegradation of Polymers. Journal of environmental polymer degradation, 7(4), 197-201. doi:10.1023/A:1022882732403

81. Gajendiran, A., Krishnamoorthy, S., & Abraham, J. (2016). Microbial degradation of low density polyethylene (LDPE) by Aspergillus clavatus strain JASK1 isolated from landfill soil. 3 Biotech, 6(1), 52-52. doi:10.1007/s13205-016-0394-x

82. Yang, J., Yang, Y., Wu, W.-M., Zhao, J., & Jiang, L. (2014). Evidence of

Polyethylene Biodegradation by Bacterial Strains from the Guts of Plastic-Eating

Waxworms. Environmental science & technology, 48(23), 13776-13784. doi:10.1021/es504038a

83. Roa, J. J., Oncins, G., Diaz, J., Sanz, F., & Segarra, M. (2011). Calculation of

Young's modulus value by means of AFM. Recent Pat Nanotechnol, 5(1), 27-36.

84. Celina, M., Ottesen, D. K., Gillen, K. T., & Clough, R. L. (1997). FTIR emission spectroscopy applied to polymer degradation. Polymer Degradation and Stability, 58(1), 15-31.

85. Sowmya, H. V., Ramalingappa, Krishnappa, M., & Thippeswamy, B. (2014).

Degradation of polyethylene by Trichoderma harzianum--SEM, FTIR, and NMR analyses. Environ Monit Assess, 186(10), 6577-6586. doi:10.1007/s10661-014-3875-6

86. El-Din Sharabi, N., & Bartha, R. (1993). Testing of some assumptions about biodegradability in soil as measured by carbon dioxide evolution. Applied and Environmental Microbiology, 59(4), 1201-1205.

87. Sagel, E. (2012). Polyethylene global overview. Pemex, Mexico.

88. Yan, D., Wang, W. J., & Zhu, S. (1999). Effect of long chain branching on

Rheological properties of metallocene polyethylene. Polymer, 40(7), 1737-1744. doi:https://doi.org/10.1016/S00323861(98)00318-8

89. Yang, J., Yang, Y., Wu, W., Zhao, J., & Jiang, L. (2014). Evidence of

Polyethylene Biodegradation by Bacterial Strains from the Guts of Plastic-Eating

Waxworms. Environmental science & technology, 48. doi:10.1021/es504038a

90. Bombelli, P., Howe, C. J., & Bertocchini, F. (2017). Polyethylene bio-degradation by caterpillars of the wax moth Galleria mellonella. Curr Biol, 27(8), R292-r293. doi:10.1016/j.cub.2017.02.060

91. Tribedi, P., & Sil, A. K. (2013). Low-density polyethylene degradation by

Pseudomonas sp.AKS2 biofilm. Environmental Science and Pollution Research, 20(6),

4146-4153.

92. Orr, I. G., Hadar, Y., & Sivan, A. (2004). Colonization, biofilm formation and biodegradation of polyethylene by a strain of Rhodococcus ruber. Applied Microbiology and Biotechnology, 65(1), 97-104.

93. Lee, B., Pometto, A. L., Fratzke, A., & Bailey, T. B. (1991). Biodegradation of degradable plastic polyethylene by Phanerochaete and Streptomyces species. Appl.

Environ. Microbiol.,57(3), 678-685.

94. Das, M. P., & Kumar, S. (2015). An approach to low-density polyethylene biodegradation by Bacillus amyloliquefaciens. 3 Biotech, 5(1), 81-86.

95. Mukherjee, S., Chowdhuri, U. R., & Kundu, P. P. (2016). Bio-degradation of polyethylene waste by simultaneous use of two bacteria: Bacillus licheniformis for production of bio-surfactant and Lysinibacillus fusiformis for bio-degradation. RSC Advances, 6(4), 2982-2992.

96. Yamada-Onodera, K., Mukumoto, H., Katsuyaya, Y., Saiganji, A., & Tani, Y.

(2001). Degradation of polyethylene by a fungus, Penicillium simplicissimum YK.

Polymer Degradation and Stability, 72(2), 323-327.

97. Volke‐Sepúlveda, T., Saucedo‐Castañeda, G., Gutiérrez‐Rojas, M., Manzur, A., &

Favela

98. Torres, E. (2002). Thermally treated low density polyethylene biodegradation by

Penicillium pinophilum and Aspergillus niger. Journal of Applied Polymer Science,

83(2), 305-314.

99. Arutchelvi, J. M. S., Ambika Arkakar, Mukesh Doble, Sumt Bhaduti, Parasu Veer

Uppara. (2008). Biodegradation of polyethylene and polypropylene. Indian Jounral of

Biotechnology, 7,9-22.

100. Vamsee-Krishna, C., & Phale, P. S. (2008). Bacterial degradation of phthalate

27 isomers and their esters. Indian Journal of Microbiology, 48(1), 19.

101. Kurane, R. (1997). Microbial degradation and treatment of polycyclic aromatic hydrocarbons and plasticizers. Annals of the New York Academy of Sciences, 829, 118

134.

102. Chen, X. Y., Romero, A., Paton-Carrero, A., Lavin-Lopez, M. P., Sanchez-Silva,

L.,Valverde, J.L., . . . Rodrigue, D. (2019). Chapter 7 - Functionalized Graphene

Reinforced Foams Based on Polymer Matrices: Processing and Applications. In M.

Jawaid, R. Bouhfid, & A. e.Kacem Qaiss (Eds.), Functionalized Graphene

Nanocomposites and their Derivatives (pp. 121155): Elsevier.

103. García, M. T., Gracia, I., Duque, G., Lucas, A. d., & Rodríguez, J. F. (2009).

Study of the solubility and stability of polystyrene wastes in a dissolution recycling process. Waste Management, 29(6), 1814-1818. doi:https://doi.org/10.1016/j.wasman.2009.01.001

104. O'Leary, N. D., O'Connor, K. E., & Dobson, A. D. (2002). Biochemistry, genetics and physiology of microbial styrene degradation. FEMS Microbiol Rev, 26(4), 403-417. doi:10.1111/j.1574-6976.2002.tb00622.x

105. Ho, B. T., Roberts, T. K., & Lucas, S. (2018). An overview on biodegradation of polystyrene and modified polystyrene: the microbial approach. Critical Reviews in

Biotechnology, 38(2), 308 320. doi:10.1080/07388551.2017.1355293

106. O'Leary, N. D., O'Connor, K. E., Ward, P., Goff, M., & Dobson, A. D. W. (2005).

Genetic Characterization of Accumulation of Polyhydroxyalkanoate from Styrene in

Pseudomonas putida CA-3. Applied and Environmental Microbiology, 71(8), 4380-4387. doi:10.1128/aem.71.8.4380-4387.2005

107. Tischler, D. (2015). Pathways for the Degradation of Styrene. In D. Tischler

(Ed.), Microbial Styrene Degradation (pp. 7-22). Cham: Springer International

Publishing.

108. Ward, P. G., Goff, M., Donner, M., Kaminsky, W., & O'Connor, K. E. (2006). A

Two Step Chemo-biotechnological Conversion of Polystyrene to a Biodegradable

Thermoplastic.Environmental science & technology, 40(7), 2433-2437. doi:10.1021/es0517668

109. Savoldelli, J., Tomback, D., & Savoldelli, H. (2017). Breaking down polystyrene through the application of a two-step thermal degradation and bacterial method to produce usable byproducts. Waste Management, 60, 123-126. doi:https://doi.org/10.1016/j.wasman.2016.04.017

110. G.B.I. Research (2013). Back to Report Store Home Polypropylene Global

Market to 2020 Developing Regions of Asia-Pacific and Middle East and Africa to Drive

Polypropylene Market Growth.

111. Lakovlev, V. V., Guelcher, S. A., & Bendavid, R. (2017). Degradation of polypropylene in vivo:

112. A microscopic analysis of meshes explanted from patients. Journal of Biomedical

Materials Research Part B: Applied Biomaterials, 105(2), 237-248. doi:10.1002/jbm.b.33502

113. Cacciari, I., Quatrini, P., Zirletta, G., Mincione, E., Vinciguerra, V., Lupattelli, P.,

& Giovannozzi Sermanni, G. (1993). Isotactic polypropylene biodegradation by a microbial community: physicochemical characterization of metabolites produced. Appl

Environ Microbiol, 59(11), 3695-3700.

114. Whiteley, K. S., Heggs, T. G., Koch, H., Mawer, R. L., & Immel, W. (2003).

Ullmann’s Encyclopedia of Industrial Chemistry. John Wiley & Sons, Weinheim, 28, 394

400.

115. Farghal, H. H. (2017). Determination of the antioxidant activity of surface treated

PET films coated with natural extracts.

116. Zimmermann, W., & Billig, S. (2011). Enzymes for the Biofunctionalization of

Poly(Ethylene Terephthalate). In G. S. Nyanhongo, W. Steiner, & G. Gübitz (Eds.),

Biofunctionalization of Polymers and their Applications (pp. 97-120). Berlin, Heidelberg:

Springer Berlin Heidelberg.

117. Chen, S., Tong, X., Woodard, R. W., Du, G., Wu, J., & Chen, J. (2008).

Identification and characterization of bacterial cutinase. J Biol Chem, 283(38), 25854

25862. doi:10.1074/jbc.M800848200

118. Wei, R., Oeser, T., Then, J., Kühn, N., Barth, M., Schmidt, J., & Zimmermann,

W. (2014). Functional characterization and structural modeling of synthetic polyester degrading hydrolases from Thermomonospora curvata. AMB Express, 4(1), 44. doi:10.1186/s13568-014-0044-9

119. Anderson, A. J., & Dawes, E. A. (1990). Occurrence, metabolism, metabolic role, and industrial uses of bacterial polyhydroxyalkanoates. Microbiological reviews, 54(4),

450-472.

120. Jendrossek, D., & Handrick, R. (2002). Microbial degradation of polyhydroxyalkanoates. AnnuRev Microbiol, 56, 403-432. doi:10.1146/annurev.micro.56.012302.160838

121. Wcisłek, A., Sonseca Olalla, A., McClain, A., Piegat, A., Sobolewski, P., Puskas,

J., & El Fray, M. (2018). Enzymatic degradation of poly (butylene succinate) copolyesters synthesized with the use of Candida antarctica lipase b. Polymers, 10(6),

688.

122. PlasticsEurope. (2018). Plastics – The Facts 2018: An analysis of European

Plastics production, demand and waste data.

123. Castro-Aguirre, E., Iniguez-Franco, F., Samsudin, H., Fang, X., & Auras, R.

(2016).Poly(lactic acid)-Mass production, processing, industrial applications, and end of life. Adv Drug Deliv Rev, 107, 333-366. doi:10.1016/j.addr.2016.03.010

124. Södergård, A. M. S. (2010). Industrial Production of High Molecular Weight

Poly(Lactic Acid). Poly(Lactic Acid): Synthesis, Structures, Properties, Processing, and

Applications, 27-41.

125. Murariu, M., Da Silva Ferreira, A., Alexandre, M., & Dubois, P. (2008).

Polylactide (PLA) designed with desired end‐use properties: 1. PLA compositions with low molecular weight ester like plasticizers and related performances. Polymers for

Advanced Technologies, 19(6), 636-646.

126. Torres, A., Li, S., Roussos, S., & Vert, M. (1996). Screening of microorganisms

For biodegradation of poly (lactic-acid) and lactic acid-containing polymers. Appl.

Environ.Microbiol., 62(7), 2393-2397.

127. Tomita, K., Kuroki, Y., & Nagai, K. (1999). Isolation of thermophiles degrading poly(L-lactic acid). J Biosci Bioeng, 87(6), 752-755.

128. Yamashita, K., Kikkawa, Y., Kurokawa, K., & Doi, Y. (2005). Enzymatic

Degradation of Poly(l-lactide) Film by Proteinase K: Quartz Crystal Microbalance and

Atomic Force Microscopy Study. Biomacromolecules, 6(2), 850-857. doi:10.1021/bm049395v

129. Fukuzaki, H., Yoshida, M., Asano, M., Kumakura, M., Mashimo, T., Yuasa, H., .

Yamanaka,H. (1990). In vivo characteristics of low molecular weight copolymers composed of L lactic acid and various dl-hydroxy acids as biodegradable carriers for drug

31 delivery systems. Biomaterials,11(6), 441-446. doi:https://doi.org/10.1016/0142

9612(90)90102-V

130. Gorrasi, G., & Pantani, R. (2013). Effect of PLA grades and morphologies on hydrolytic degradation at composting temperature: assessment of structural modification and kinetic parameters. Polymer Degradation and Stability, 98(5), 1006-1014.

Chapter II: Bacillus pumilus B12 Degrades Polylactic Acid and Degradation is Altered by Changing Nutrient Conditions

Contributions

A version of this Chapter is currently in press at Frontiers in Microbiology. The manuscript is by Kyle S. Bonifer, Xianfeng Wen, Sahar Hasim, Elise K. Phillips, Eric R. Gann, Jennifer M. DeBruyn and Todd B. Reynolds.

This article has additional atomic force microscopy data in different nutrient conditions that were not found in the original publication as well as preliminary data showing the degradation detection of poly-butylene succinate (PBS) films. Contributions of this publication are as follows:

Enrichment and CO2 measurements of B. pumilus B12 were performed by Xianfang Wen and Rachel Dunlap in the DeBruyn Lab. The 16s rDNA extraction was performed by Kyle Bonifer and sequenced by the division of biology sequencing facility with the phylogenetic tree construction on 16s rDNA performed by Eric Gann in the Wilhelm Lab. AFM was performed by Sahar Hasim. Enzyme assay development, growth curves, SEM, and plastic film production for all assays was performed by Kyle Bonifer. Elise Phillips aided in completing some of the enzyme assays and SEM.

Abstract

Poly-lactic acid (PLA) is increasingly used as a biodegradable alternative to traditional petroleum-based plastics. In this study, we identify a novel agricultural soil isolate of Bacillus pumilus (B12) that is capable of degrading high molecular weight PLA films. This degradation can be detected on a short timescale, with significant degradation detected within 48-hours by the release of L-lactate monomers, allowing for a rapid identification ideal for experimental variation. The validity of using L-lactate as a proxy for degradation of PLA films is corroborated by loss of rigidity and appearance of fractures in PLA films, as measured by atomic force microscopy and scanning electron microscopy, respectively. Furthermore, we have observed a dose-dependent decrease in PLA degradation in response to an amino acid/nucleotide supplement mix that is driven mainly by the nucleotide base adenine. In addition, amendments of the media with specific carbon sources increase the rate of PLA degradation, while phosphate and potassium additions decrease the rate of PLA degradation by B. pumilus B12. These results suggest B. pumilus B12 is adapting its enzymatic expression based on environmental conditions and that these conditions can be used to study the regulation of this process. Together, this work lays a foundation for understanding the bacterial degradation of biodegradable plastics.

Introduction

The crisis of plastic pollution in the environment has led to an increased adoption of biodegradable polymers in the commercial plastic industry. Biodegradable polymers are considered a more ecologically friendly alternative to conventional plastic polymers due to a limited accumulation in the environment [1]. Some of these biodegradable polymers, such as poly- hydroxyalanoate (PHA) and polylactic acid (PLA), are derived from renewable biomass [2,3], while other biodegradable polymers are synthesized from fossil fuel feed stocks (e.g. PBAT, PCL). Biodegradable polymers like these are polyesters or co-polyesters, formed with ester linkages that are common to many macromolecules that are both synthesized and degraded by microbial life. This is in contrast with conventional petroleum-based polymers that are rich in carbon-carbon linkages which are resistant to microbial degradation, rendering the polymers more recalcitrant.

PLA is the second most common biodegradable polymer found in commercial plastics [4]. PLA is an aliphatic polyester formed from direct condensation of lactic acid molecules or from ring polymerization of lactide [5]. Several soil bacteria including Bacillus brevis and fungi such as Rhizopus delemer have been identified that are capable of degrading PLA, mainly in composting conditions [6,7]. This degradation is carried out by a variety of proteases and lipases that hydrolyze PLA to form lactic acid [8,9,10]. Most previous research monitoring biodegradation has focused on the production of CO2 as an indicator of cellular respiration and loss of film mass as a gauge for polymer degradation [11,12,13]. This practice tends to be performed over the course of weeks to months and does not lend itself to rapid identification of novel microbial isolates, quick detection of enzymes capable of polylactic acid degradation or speedy categorization of microbial adaptation through changes in gene expression.

Members of the genus Bacillus are spore-forming gram-positive rods which include several species, such as Bacillus subtilis and Bacillus thuringiensis, that are amongst the most widely studied microorganisms in molecular biology. This genus is well known for the large quantities of secreted and surface associated enzymes they can produce. As a result, Bacillus spp. are used heavily in various biotechnological settings [14,15,16]. Bacillus pumilus specifically has been shown to be remarkably resistant to many forms of environmental stresses such as intense ultraviolet light and high concentrations of hydrogen peroxide [17,18]. With the combination of

36 high stress tolerance, genetic tractability and high enzyme expression, B. pumilus is an ideal candidate for further investigation in polymer degradation.

Although several bacterial and fungal strains have been identified that are capable of degrading PLA, very little is understood about the underlying regulatory mechanisms of degradation. Many of these organisms live in dynamic environments where nutrient conditions are subject to temporal change. In order to develop hypotheses concerning genetic regulation, there is a need to develop degradation assays which align closer with the timescale of microbial adaptation. In addition, fundamental experiments identifying the conditions under which degradation occurs are needed in order to build a mechanistic understanding of cellular-level biodegradation. In this study we identify a novel agricultural isolate of Bacillus pumilus (B12) with is capable of degrading high molecular weight PLA films. Our goals were to 1) develop a rapid test for identifying markers of PLA degradation; and 2) determine if changing nutrient supplementation in the media affects PLA degradation rates. Our results suggest B. pumilus B12 is adapting its enzymatic expression based on environmental conditions and that these conditions can be used to study the regulation of this process.

Methods Microbial Enrichment and Growth Conditions Soil samples were collected from agricultural soil (sandy loam; 59.9% sand, 23.5% silt, and 16.6% clay; classified as a fine kaolinitic thermic Typic Paleudults) at the University of Tennessee East Tennessee Research and Education Center in Knoxville, Tennessee, USA. Soils were diluted in sterile water, then inoculated onto petri dishes with squares of biodegradable plastic mulch films containing PLA in minimal salt media (MSM) agar, following the methods described in Bailes et al. (2013) [19]. Minimal salt media (MSM) was composed of 7.5 mM NH4SO4, 11 mM KHPO4, 2.87 mM K2HPO4, 1.7 mM MgSO4 and 17.1 mM NaCl, pH 7.0; 1.5% agar was added to make MSM plates. The biodegradable mulch films included BioAgri® (BioBag Americas, Inc.), comprised of Mater-Bi® and an experimental film comprised of a blend of polylactic acid (PLA) and polyhydroxyalkanotaes (PHA). More details on film composition and properties can be found in Hayes et al.,2017 [20]. Soil slurries were plated and incubated statically at 30°C. Single colonies were selected from plates and re-plated on LB agar until morphologically

37 homogeneous colonies were observed, indicative of a pure culture. To test for degradation abilities, strains were inoculated into 100 mL liquid MSM with UV-sterilized 4 cm x 4 cm squares of BioAgri® or PLA/PHA films. Triplicate cultures were grown in sealed flasks with rubber septa for headspace sampling. Flasks were incubated for 16 days at 30°C in a shaking incubator at 180 rpm. Two controls were included in each experiment: one without microbial innocula (i.e. plastic film only) and one without plastic film (i.e. microbes only). Headspace CO2 concentrations were determined every four days by sampling 0.5 mL headspace gas with a syringe and analyzing on an LI-820 infrared gas analyzer (Licor Inc.) and used as an indicator of plastic film degradation. Plastic pieces were also weighed before and after incubation, and mass loss used as an indicator of degradation. One isolate that exhibited the ability to degrade BioAgri® and PLA/PHA films was identified with 100% gene similarity as Bacillus pumilus via 16S rRNA gene sequencing and chosen for further experimentation. B. pumilus cultures were grown overnight, washed three times 8 with phosphate buffered saline and diluted to an OD600nm of 0.1 (approximately 3.125*10 CFU/mL) for all polylactic acid degradation experiments. Cultures were kept static at 30° C in minimal salt media for 48 hours. When adding complete supplementation media to liquid cultures, MP Biomedicals™ CSM-HIS-LEU-URA Powdered Media (Fisher-MP114531212) was used. CSM media contains various amino acids and adenine. All carbon supplementation experiments used a concentration of 55µM for all sugars added to the minimal salt media. When testing nitrate as the main nitrogen source, NaNH3 was supplemented in place of NH4SO4. For growth curve measurements, OD600 was measured every 30 minutes for 72 hours in a Biotek Cytation5 plate reader. Phylogenetic Analysis of 16S rRNA Gene Sequences Genomic DNA from B. pumilus was isolated through phenol-chloroform extraction. Universal 16S primers 27F and 1522R were used for PCR sequence amplification. Amplicons were Sanger sequenced at the University of Tennessee Genomics Core. Near full length 16S nucleotide sequences were downloaded from NCBI [21]. Multiple sequence alignments were performed in MEGA7 [22] using the MUSCLE algorithm [23]. Maximum likelihood phylogenetic reconstruction was performed in MEGA7 with 500 bootstrap replications to determine confidence values for each node.

PLA Film Creation PLA films were created from polylactic acid beads with an inherent viscosity of 1.8 dl/g and a molecular weight between 80,000-100,000 kDa (Polysciences, Inc.). For L-lactate detection and scanning electron microscopy, 10 mg of PLA were dissolved in 1 mL of chloroform, and the dissolved PLA liquid was deposited into autoclaved flat bottom glass vials that were ~1 cm in diameter. The chloroform was evaporated under nitrogen gas in a fume hood over 1 hour resulting in a film that was cast on the bottom of the vial. Vials were stored at -20˚C with sterile lids to reduce desiccation. L-lactate Detection Assay Detection of L-lactate released into the supernatant was performed using ENZYFLUO L- lactate kit (Bioassay Systems Inc.). Overnight cultures were grown in Lennox Broth (LB) shaking at 225 RPM at 30° C and washed three times with phosphate buffered saline. An optical density of 0.1 was incubated in 1 mL of media on 10 mg/mL PLA films in glass vials for 48 hours. Supernatant was taken off cultures centrifuged at 13,000 RPM for 2 minutes from 10mg/mL PLA films. A 1:1 mixture of supernatant collected from polylactic acid films was added to the L-lactate kit detection solution giving a total volume of 100µL in a 96-well plate. The supernatant/detection solution was incubated in the dark at room temperature for 1 hour and colorimetric changes were measured using a 535/580nm excitation/emission ratio in a Biotek Cytation5 plate reader. Results were normalized to a standard curve generated with L-lactate standards ranging from 0-42µM. All measurements were added to the slope generated by the standard curve, yielding approximate concentrations of L-lactate release. All trials included negative controls containing cultures with no PLA added, as well as PLA films in media only. Measurements were taken with 3 biological replicates each with 3 technical replicates, yielding nine total data points per treatment. Statistical significance was determined either through t-test or with ANOVA based on the number of comparisons make with p-values below .01 considered significant. Scanning Electron Microscopy B. pumilus cultures were incubated at 30℃ for 48 hours with 10mg/mL polylactic acid films in minimal salt media. Films were then washed three times with phosphate buffered saline (PBS) and fixed with 2.5% glutaraldehyde for one hour. Samples were then gradually dehydrated with ethanol anhydrous( 25%, 50% and 100% respectively) and air dried in a fume hood overnight.

Samples were mounted on aluminum stubs, sputter coated with iridium and observed using a 1 keV Zeiss GeminiSEM scanning electron microscope. Atomic Force Microscopy Polylactic acid films 50 mg/mL in thickness were incubated for 48 hours in minimal salt media (MSM). Samples were washed with phosphate buffer saline 5 times to remove excess biomass. Each sample was then imaged using a 5500 PicoPlus atomic force microscopy (AFM) operating with the 1.20.2 operating system (Keysight Technologies Inc. Santa Rosa CA). The instrument was operated in contact mode using MLCT probes with spring constants of 0.01 and 0.03 N/m with a tip radius of 10 nm and Poisson ratio of 0.5. For both imaging and elasticity measurements, the applied force was kept at 3-5 nN. The force distance curves were measured with 8x8 points, with each point being an average of 3 force curves. These measurements were recorded on 2-μm2 areas on the top of plastic for an average of 5 different location. This data was then converted to elasticity measurements using the Keysight software in PicoPlus 5500 AFM [24]. Statistical significance was determined via T-Test with p-values below .01 considered significant.

Results Release of L-lactate is used to Measure Poly-lactic Acid Degradation by Hydrolases To quantify poly-lactic acid (PLA) degradation over relatively short time periods, a commercially available L-lactate detection kit was used. A known PLA degrading enzyme, the serine protease Proteinase K, was chosen as a positive control to test the efficacy of the colorimetric assay [25]. Poly-lactic acid films of 10 mg/mL were created in 96-well plate formats and serially diluted concentrations of Proteinase K were added to each well in minimal salt media (MSM). Following 24 hours of incubation at 30⁰C, up to 100µM concentrations of L-lactate were detected in the supernatant of PLA samples exposed to Proteinase K (Figure 2. 1A.). Proteinase K unit concentrations as low as .05 units/mL could release significant concentrations L-lactate compared to the media control over a 24-hour period. Since Proteinase K is a useful positive control, we wanted to determine its parameters for PLA degradation as detected by this kit. Therefore, a time course of PLA degradation was performed to determine when one unit (releases 1µM of substrate per minute) of Proteinase K saturates the detection limit of the L-lactate detection kit. After 240 minutes of incubation, saturation of detectable L-lactate in the supernatant was

40 observed (Figure 2. 1B.). Thus, incubations for 24 hours were adequate to measure saturated degradation of PLA by this purified hydrolase.

Next, assay conditions were altered to mimic expected variables in media conditions during microbial growth. Altering the temperature between 21°C and 37°C had no effect on the ability of Proteinase K to degrade PLA (Figure 2. 2.). Next, acidification of the media or an addition of carbon was investigated to see if it could alter the sensitivity of our L-lactate detection kit. No significant differences were found between the release of L-lactate in more acidified media, and no differences were seen in lactate levels with the addition of glucose. (Figures 2. 3., 2. 4.). Switching Proteinase K buffer to MSM had no negative effect on enzymatic activity and led to a decrease in background detection of L-lactate. Furthermore, L-lactate release from PLA in the presence of Proteinase K was consistent in biologically relevant levels of salinity (Figure 2. 5.). The results of these initial assays gave us confidence in our positive control, Proteinase K, for measuring PLA degradation. Finally, laboratory conditions of pH, temperature and salinity that microbes would be exposed to did not significantly alter L-lactate release by Proteinase K.

Isolation of a Soil Bacterium, Bacillus pumilus B12, Capable of Degrading Poly-lactic Acid Films

To culture PLA degrading microorganisms, samples were collected from agricultural soils where biodegradable plastic mulch films are used and inoculated into enrichment cultures. Soils were diluted and plated on minimal salt media (MSM) agar containing a square of biodegradable plastic mulch films. Isolate B12 was identified as

Bacillus pumilus based on 16S rDNA gene sequencing (Figure 2. 6.). When incubated in MSM with pieces of BDMs, the strain caused a greater increase in CO2 in the headspace of the cultures compared to uninoculated controls (Figure 2. 7.). Over a 16-day incubation, the mass of plastics was decreased by 9.45% and 2.97% for BioAgri® and PLA/PHA film, respectively. The evolution of CO2 combined with mass loss of plastics indicated that this strain had the ability to biodegrade the plastics.

Next, supernatant samples of microbial cultures grown in the presence of laboratory-generated PLA films were tested for L-lactate release. All cultures were grown under static conditions, limiting the total biomass created. This allowed us to simulate conditions more commonly found in the soil environment. Following a three-week incubation, low levels of L-lactate were detected

Figure 2. 1. High levels of L-lactate release detected from PLA films exposed to Proteinase K. A) 10 mg/mL Poly-lactic Acid (PLA) films were incubated at 30⁰ C for 24 hours with serial dilutions of Proteinase K (n=3). Supernatant samples were collected and added to a colorimetric L-lactate detection kit. MSM+G represents minimal salt media with 55mM glucose supplementation. B) PLA films were incubated with 1.6 total units of Proteinase K with supernatant samples being collected at successive time point.

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Figure 2. 3. Changing the pH of the media does not change detection of L-lactate from PLA films. 10 mg/mL films were incubated for 48 hours in minimal salt media and L-lactate was measured from the supernatant. The pH was adjusted by adding hydrochloric acid to the media prior to incubation. Results are compared to PLA films incubated with our positive control Proteinase K for 24 hours. **** indicates ANOVA p-value ≤.0001 and ns indicates ANOVA p-value ≥ .05.

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Figure 2. 4. Adding glucose has no effect on the efficacy of the L-lactate assay. Black bars represent incubation of minimal salt media (Glu-) and grey bars indicate supplementation with 55 mM glucose (Glu+). Presence or absence of Proteinase K (ProK) and Polylactic acid (PLA) are represented by + and – respectively. ** and **** indicates ANOVA p-value of ≤ .01 and ≤ .0001 respectively.

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0 l l l l l l l l C C C C C C C C a a a a a a a a N N N N N N N N M M M M M M M M m m m m m m m m .1 5 2 9 .1 5 2 9 7 3 5 6 7 3 5 6 1 1

Figure 2. 5. Increasing salinity has no effect on L-lactate released from Proteinase K. 10 mg/mL PLA films were incubated for 24 hours in Minimal salt media with glucose (MSM+G) and L-lactate release was measured from the supernatant. Black bars indicate media alone and grey bars indicate 1 unit of Proteinase K addition. “ns” indicates ANOVA p-value ≥ .05 against MSM+G control (17.1 mM NaCl).

Figure 2. 6. Maximum likelihood phylogenetic tree of 16S rDNA for the environmental isolate B12 compared to other Bacillus spp. This bootstrap consensus tree was constructed with 500 replicates. Bootstrap values were used as confidence values for the nodes. Nucleotide sequences for the other bacteria were retrieved from NCBI. The accession numbers are as follows: Bacillus amyloliquefaciens (HQ874610.1), Bacillus megaterium strain SP4-17 (HQ874615.1), Bacillus licheniformis strain OF3-16 (HQ874614.1), Bacillus amyloliquefaciens strain ATCC 23842 (EU689157.1), Bacillus subtilis strain DSM10 (AJ276351.1), Bacillus licheniformis strain HSP-1 (HQ874617.1), Bacillus subtilis subsp. natto strain NT-2 (HQ874611.1), Bacillus subtilis ATCC 6633 (AB018486.1), Bacillus pumilus strain TLB7-7 (HQ874616.1), Bacillus pumilus strain ATCC 7061 (AY876289.1), Bacillus pumilus strain NBRC 12092 (NR_112637.1), Bacillus pumilus strain CIP 52.67 (NR_115334.1), Bacillus pumilus strain NRRL NRS-272 (NR_116191.1), Bacillus pumilus strain SBMP2 (NR_118381.1), Bacillus cereus strain ATCC11778 (AF290546.1), Bacillus cereus strain ATCC14579 (AF290547.1), Bacillus mycoides strain ATCC 6462 (EF210295.1), Bacillus thuringiensis Bt407 (ACMZ01000011.1), Alicyclobacillus acidocaldarius subsp. rittmannii (AB089859.1)

Figure 2. 6 Continued

35000 Plastic only 30000 Strain only Plastic + strain 25000

2 2 20000

15000 CO ppm

10000

5000

0 BioAgri PLA/PHA

Figure 2. 7. Headspace CO2 concentrations after 16 days of culturing B. pumilus B12 in MSM with squares of biodegradable plastic mulch films. Data are means and standard deviations of triplicate culture flasks.

50 in samples grown with PLA as their sole carbon source. However, when the MSM+PLA cultures were supplemented with glucose, large amounts of L-lactate were detected in the supernatant (Figure 2. 8.). This suggests that this strain can degrade the plastic but generates more esterase activity in the presence of glucose. To confirm that detected L-lactate was not a by-product of glucose fermentation, supernatants from B. pumilus cultures grown in glucose media with and without the addition of a PLA film were tested. After just 48 hours in minimal L- lactate medium, significantly higher concentrations of L-lactate were detected in cultures supplemented with both glucose and PLA (MSM+G+PLA) than cultures supplemented with glucose alone (MSM+G) (Figure 2. 9.). In fact, the levels of L-lactate detected from MSM+G+PLA cultures were comparable to PLA films exposed to our positive control, Proteinase K, over the same time period. Following this result, all standard media had 55µM of glucose or another sugar added to prime PLA degradation. No significant release of L-lactate release was seen for Bacillus subtilis strain 168 (NC_000964.3), suggesting that the active enzymes degrading PLA are either unique to B. pumilus, or not expressed in B. subtilis in the same conditions (Figure 2. 9.). This evidence also supports the conclusion that the high L-lactate levels detected in the supernatant were a product of PLA degradation and not glucose fermentation by B. pumilus B12. This assay is a demonstration that degradation can be detected over a far shorter period (i.e. hours to days) than conventional

CO2 and film mass loss analysis with glucose priming (i.e. days to weeks).

Although we detected L-lactate release from our PLA films, we wanted an independent confirmation that PLA was being degraded. Thus, we utilized atomic force microscopy (AFM) to detect nano-scale changes in the mechanical properties of the poly-lactic acid films. Following a 48-hour incubation, we observed a significant decrease in the Young’s modulus (elasticity) of PLA films when exposed to our microbial isolate, B. pumilus B12, indicating a loss in elastic strength of the polymer (Figure 2. 10.). This loss corresponded to an approximately 3-fold loss in kPA compared to media only control films. To test if the differences seen in PLA elastic strength were caused by activities related to this specific microbe, Bacillus subtilis strain 168 (previously shown to not release L-lactate from PLA films) (Figure 2. 9.) was used as a negative control. No significant differences in the Young’s modulus of the PLA could be detected when incubated with B. subtilis str. 168 in MSM+glucose (Figure 2. 10.). This indicates the loss of tensile elasticity of the plastic film is specifically due to the presence of B. pumilus B12. Next, scanning electron

3 0 0 ****

2 0 0

)



(

- L

1 0 0 ns

0 ) -) ) A ( A L G L ly + P P n ( + + o 2 1 G K ia + e d B ( s s a e 2 lu 1 in m i e B t M m s o u u r S p il P M . m + B u A p L s P lu il c a B

Figure 2. 8. High levels of L-lactate can be detected in 3-week microbial cultures when both glucose and PLA are added. B. pumilus B12 was incubated for 3 weeks in minimal salt medium with PLA films and a primary carbon source of glucose was either absent (-) or present (+). Results were compared to PLA films incubated with Proteinase K for four hours, and PLA incubated in Minimal Salt Media (MSM) as a negative control. **** indicates t-test p-value ≤ .0001 and ns indicates t-test p-value ≥ .05 compared to PLA+MSM media only (-). These assays were performed with three biological and three technical replicates.

1 5 0

****

1 0 0

)

 (

a L

- L 5 0 ns

0 r ) ) G A u G A + L o L P (+ M + -H P S 4 8 + G 6 G M + K 1 + r. (+ 2 M e t 1 S s s 8 a s 6 B M n i 1 i il . + te t tr 2 b s 1 ro u s is B P . l ti B b u s . B

Figure 2. 9. The addition of glucose significantly increases the release of L-lactate from PLA in microbial cultures. All microbial samples were incubated for 48 hours in Minimal Salt Media (MSM) media supplemented with glucose (G). Supernatant samples that led to over saturation of the L-lactate detection assay were diluted 1:10 to ensure more accurate results. **** indicates t- test p-value ≤ .0001 compared to and ns indicates t-test p-value ≥ .05 compared to B. pumilus B. subtilis str. 168 (+) control. These assays were performed with three biological and three technical replicates.

4 0 0 0 0 0

3 0 0 0 0 0 ns

)

(

u d

o 2 0 0 0 0 0

o Y **** 1 0 0 0 0 0

****

) 2 ly 8 L 1 n 6 / 1 s B o . it s tr u G s n il + u is 8 m M l . u S ti ( p b . M u K s e B s . a B in te ro P

Figure 2. 10. PLA films show a decrease in elastic strength when incubated with B. pumilus B12. Shows the average of all cantilever measurements taken over 3 biological replicates. Elastic strength is measured in Young’s modulus Pascals (Pa). All samples were incubated in MSM+ glucose for 48 hours. B. subtilis str.168 (NC_000964.3) was used as a negative control. **** indicates t-test p-value ≤ .0001 and ns indicates ANOVA p-value ≥ .05 indicates against minimal salt media plus glucose (MSM+G) control. These assays were performed with three biological and three technical replicates.

Figure 2. 11. Scanning electron microscopy shows PLA fracturing around B. pumilus B12 cells. A-C) 5,000x magnification of 10mg/mL of PLA film for 48 hours with: media only (A), B. pumilus B12 (B) and Proteinase K (C). D-F) 30,000x magnification of 10 mg/mL PLA films with: media only (D), B. pumilus F26 (E) and Proteinase K (F).

Figure 2. 11. Continued

56 microscopy (SEM) was utilized to visualize B. pumilus B12 adhesion and growth on PLA plastic films. While observing the bacterial growth on PLA, several fractures were seen in cells (Figure 2. 11B. and 2. 11E.). These fractures shared similar features to those observed on PLA with Proteinase K added (Figure 2. 11C. and 2. 11F.). The combination of these fractures and the loss of elastic strength of the plastic film are in concordance with our L-lactate detection assays and provide strong evidence that B. pumilus B12 is degrading the PLA films and degradation is detectable within 48 hours.

Carbon, Organic Nitrogen, Phosphate and Potassium Concentrations in Media Regulate Microbial Degradation of Polylactic Acid

In order to identify environmental conditions that regulate degradation of poly-lactic acid by these microbes, we altered carbon, nitrogen, phosphate and potassium sources, as these are some of the most important nutrients found in the soil environment and are known to regulate microbial growth and enzymatic expression [26]. Since supplementation with glucose increases PLA degradation (Figure 2. 8., 2. 9.), we investigated the effects of altering the primary carbon source on the ability of Bacillus pumilus B12 to degrade PLA. Following a 48-hour treatment with a variety of carbon sources, we saw significant increases in L-lactate release from PLA when changing the primary carbon source from glucose to the disaccharide maltose and the sugar alcohol mannitol (Figure 2. 12.). Reductions in L-lactate release were seen for two non-fermentable carbon sources, glycerol and sorbitol, as well as galactose, however, none of these reductions were significant compared to the same grow conditions lacking PLA (Figure 2. 12.). To determine if these changes were the result of altered growth rates, we determined the amount of L-lactate detection per cell to help visualize the effect of carbon sources on PLA degradation. To accomplish this, we performed 48-hour growth curves in each of the different carbon sources to determine the approximate concentration of cells at the time of L-Lactate measurement (Figure 2. 13.). After taking cell number into account, both maltose and mannitol were shown to have a significant increase in PLA L-lactate released per cell compared to glucose (Figure 2. 14.). Furthermore, sorbitol, glycerol and galactose all showed significant reductions when taking cell number into account (Figure 2. 14.). Since nitrogen is often limiting in soils, and thus, is added to agricultural soils as fertilizer, both nitrogen source and concentration were altered to see the effect on B. pumilus B12 PLA degradation. No notable changes or trends in PLA degradation were observed

57 when increasing the concentration of ammonium sulfate in the media or for sodium nitrate. (Figure 2. 15., 2. 16.).Additionally, no noteworthy changes or trends in L-lactate release were observed when increasing the total nitrogen concentration with nitrate (Figure 2. 15.). Thus, we tested the effects of changing the nitrogen source through the addition of free amino acids. Complete Supplement Media (CSM); which contains most amino acids and adenine, was added to the MSM medium and plastic degradation was quantified through L-lactate release. No significant changes in L-lactate were observed with standard additions of CSM, however as concentrations increased, the amount of L- lactate released by B. pumilus B12+CSM+PLA cultures decreased (Figure 2. 17.). Adding these amino acids slightly increased the overall growth of B. pumilus B12, indicating that the decrease seen in PLA degradation correlates with the microbe’s physiology and not its overall abundance (Figure 2. 18., 2. 19.). To determine if specific amino acids drove this reduction in PLA degradation, individual amino acids or nucleotides bases were added as supplements, and we observed that the reduction in L-lactate release was not being driven by any of the free amino acids in the supplement, but instead the nucleotide base adenine (Figure 2. 20.). These results show that increasing free nitrogen alone does not drive PLA degradation up or down, but the supplementation of adenine specifically reduces degradation.

Phosphate is an essential nutrient, with free phosphate being limited in the soil environment. Fertilizers work to replenish phosphate depleted soils, having a large impact on the phosphate concentrations available for the soil microbiome [27]. Phosphate and potassium concentrations were changed in the MSM media to see if there would be a change in PLA degradation by B. pumilus B12. Potassium is a major cation which has been shown to change kinetics, energy coupling and regulation in microbial life. Both ions have been shown to modulate gene regulation and are possible regulators in the release of enzymes with PLA degradation activity [28,29]. When increasing potassium phosphate monobasic (KH2PO4) concentrations in the media, small but significant decreases were initially seen in L-lactate release (Figure 2. 21.). However, there were stark increases in microbial growth with increased concentrations of potassium phosphate (Figure 2. 22.). After normalizing for increased cell number, we observed a large dose dependent decrease in L-lactate release with increasing potassium phosphate (Figure 2. 23.). Thus, we see effects by both potassium and phosphate. To separate the contributions of each ion, media was supplemented with either potassium chloride (KCl) or sodium phosphate (NaPO4).

Figure 2. 12. Bacillus pumilus B12 increases alters L-lactate release from PLA when grown in different carbon sources. Initial measurements of B. pumilus B12 grown in different primary carbon sources at a concentration of 55µM taken after 48 hours. Red bars indicate the addition of PLA, black bars indicate B12 grown in media only. All **** and ns above red bars indicates ANOVA ****=p-value ≤.0001 ns= not significant against media only control. Above **** indicates p-value ≤.0001 against glucose carbon source. These assays were performed with three biological and three technical replicates.

0 .5 G lu c o s e M an n ito l M a lto se S o rb ito l S tarch 0 .4 S u c ro se

)

m n

0 0 .3

(

c i

t 0 .2 p O

0 .1

0 .0 0 5 0 0 1 0 0 0 1 5 0 0 2 0 0 0 2 5 0 0 3 0 0 0 3 5 0 0 T im e (m in )

Figure 2. 13. Bacillus pumilus B12 48-hour growth curves in different carbon sources. Optical density-based growth curves of B. pumilus B12 in the carbon sources taken over 48 hours. Each growth curve represents twelve biological replicates.

Figure 2. 14. Release of L-lactate from PLA incubated with B. pumilus B12 is independent of growth on different carbon sources. Release of L-lactate from PLA normalized for growth of B. pumilus B12 after 48 hours. ANOVA results ****=p-value ≤ .0001 ns= not significant against standard glucose supplementation.

8 0

6 0

) M

 *

(

e t ns

a ns

t 4 0 ns c

a ns

- L

2 0

****

0 ------ly O 3 O 3 O 3 O 3 O 3 O 3 n - N N N N N N O M M M M M M O 3 m m m m m m N 0 1 2 4 8 0 1 2 4 8 1 M 1 m 0 1 1

- Figure 2. 15. Changing primary nitrogen source to Nitrate (NO3 ) has no effect on PLA degradation by B. pumilus B12. PLA films were incubated for 48 hours with B. pumilus and supernatant L-lactate measurements were taken. **** indicates ANOVA p-value ≤ .0001, * indicates ANOVA p-value ≤ .01 and ns indicates ANOVA p-value ≥ .05 against - MSM+G control. NH3 only represents media without bacterial inoculum.

8 0

6 0

)

M ns

 ns

(

** e

t * a

t 4 0

a L

- L

2 0

****

0 + + + + + 4 4 4 ly 4 4 n H H H H H + O N N N N N M M M M M H 4 m m m m m N .5 0 5 0 5 7 3 4 6 7 M m 5 7

Figure 2. 16. Increasing ammonium concentration in the media has no effect on PLA degradation by B. pumilus B12. PLA films were incubated for 48 hours with B. pumilus and supernatant L-lactate measurements were taken. **** indicates ANOVA p-value ≤ .0001, ** indicates ANOVA p-value ≤ .01 , * indicates ANOVA p-value ≤ .05 and ns indicates ANOVA + p-value ≥ .05 against MSM+G control (7.5 mM NH4 ).

8 0

ns ns ns

6 0

)



(

t a

t 4 0

a L

- **** L **** 2 0 **** ****

M M M M M M M G S S S S S S S + C C C C C C C M x x x x x x x S 1 2 4 8 0 5 0 1 1 2 M

Figure 2. 17. Increasing CSM concentrations available to B. pumilus B12 leads to a decrease in L-lactate release from PLA. L-lactate release after 48 hours from PLA incubated with B. pumilus B12 supplemented with increasing concentrations of complete supplement media (CSM). **** indicates ANOVA p-value ≤.0001 and ns indicates ANOVA p-value ≥.05 indicates against minimal salt media plus glucose (MSM+G) control. These assays were performed with three biological and three technical replicates.

0 .4 M S M + G 2 x C S M 4 x C S M 1 0 C S M

0 .3

)

0 0

6 0 .2

(

D O

0 .1

0 .0 0 1 0 0 0 2 0 0 0 3 0 0 0 T im e (m in )

Figure 2. 18. Bacillus pumilus B12 has altered growth rates in different concentrations of CSM. 48-hour growth curves of B. pumilus B12 with added concentrations of CSM. Each growth curve represents twelve biological replicates.

Figure 2. 19. Bacillus pumilus B12 has decreased L-lactate release independent of growth on CSM. Release of L-lactate from PLA normalized for growth of B. pumilus B12 after 48 hours. **** indicates ANOVA p-value ≤ .0001 against MSM+G.

8 0

6 0

)



(

e t

a 4 0

- L

2 0

****

0 e e e e e e n e e d e e G n n n n a n i n n K + i in i in i i h i in c i i s c n l n s n c n n e o a i y p a A e s M r u io g o l u c o a S y e V r L t le i e d n L th p A t r A i M T e A y l o r h te r y Is a T T n p o M e s r h A P P

Figure 2. 20. Adenine is the component of CSM that is driving the decrease L-lactate release from PLA and Bacillus pumilus B12 cultures. L-lactate release after 48 hours from PLA incubated with B. pumilus B12 with single amino/ nucleic acid supplementations. All supplementations are 10-fold higher than the standard concentration found in complete supplement media. Results have been normalized to account for L-lactate released from nutrient supplementation alone. **** indicates ANOVA p-value ≤ .0001 against MSM+G. These assays were performed with three biological and three technical replicates.

4 0

3 0

)



(

t a

t 2 0

a L

- **** **** L

1 0 ****

4 4 4 4 O O O O P P P P H 2 H 2 H 2 H 2 K K K K M M M M m m m m 2 5 0 1 2 5 1 1 1

Figure 2. 21. B. pumilus B12 decreases L-lactate released from PLA in increased KH2PO4. A) Initial measurements of B. pumilus B12 L-lactate release from PLA taken after 48 hours.

K2HPO4 concentration increases are based off standard minimal salt media with glucose (MSM+G). These assays were performed with three biological and three technical replicates. **** indicates ANOVA p-value ≤ .0001 against MSM+G.

1 .0 1 1 m M K H 2 P O 4 2 2 m M K H 2 P O 4 5 5 m M K H 2 P O 4 1 1 0 m M K H P O ) 2 4 0 .8

n 0

(

t 0 .6

i s

l a

c 0 .4 i

p O

0 .2

0 1 0 0 0 2 0 0 0 3 0 0 0 4 0 0 0

T im e (m in )

Figure 2. 22. Bacillus pumilus increases its growth with increasing concentrations of KH2PO4. Optical density-based growth curves of B. pumilus B12 in the carbon sources taken over 50 hours. Each growth curve represents eight biological replicates.

Figure 2. 23. Bacillus pumilus B12 has reduced L-lactate release from PLA films with increasing concentrations of KH2PO4. Normalized L-lactate release/cell based on data from Figures 2.21. 2.22. . **** indicates ANOVA p-value ≤ .0001 against MSM+G.

Thus, we see effects by both potassium and phosphate. To separate the contributions of each ion, media was supplemented with either potassium chloride (KCl) or sodium phosphate (NaPO4). We further controlled for sodium and chloride ions by supplementing with sodium chloride, which had no effect on L-lactate release by B. pumilus B12 (Figure 2. 24.). Thus, we were able to measure + - the contributions of K and PO4 individually. Increasing the concentration of potassium had no initial effect on L-lactate release by B. pumilus, however as the concentration reached 75 mM, there was a large increase in L- lactate detection (Figure 2. 25.). This increase culminated with a sharp decrease in the growth rate compared to concentrations of 37.5 mM K+ or below (Figure 2. 26.). When normalized for cell number, degradation is reduced at 37.5 mM K+, but increases sharply at 75mM K+ (Figure 2. 27.). The significant increase in L-lactate release from PLA correlates with the lower growth rate suggesting added stress on B. pumilus B12 at high - concentrations of potassium chloride. When altering concentrations of free PO4 (without increasing potassium), we see a concentration dependent increase in the total L-lactate being released from PLA films incubated with B. pumilus B12 (Figure 2. 28). However, B. pumilus B12 - has a higher growth rate in the PO4 supplemented media (Figure 2. 29.) and therefore when the release of L-lactate is normalized for microbial growth, there is a significant decrease in L-lactate release compared to no phosphate controls (Figure 2. 30.). The potassium and phosphate results show a general trend that when B. pumilus B12 is growing more efficiently on specific media, less L-lactate is released from PLA films. Conversely, when microbial growth is stressed by being in minimal media or inhibitory concentrations of K+, L-lactate release increases.

Discussion

The measurement of poly-lactic acid (PLA) degradation has historically relied on techniques that are either time consuming or expensive to perform when running experiments with many variables. This limits the investigatory capabilities into large ranges of changing conditions that PLA is exposed to in different degradation settings. In this study we have shown the ability to measure short term degradation of PLA films from a novel B. pumilus isolate. The assay was stable under biologically relevant parameters, with L-lactate concentrations released from PLA mirroring other quantitative and qualitative measurements of degradation (e.g. CO2 evolution or film mass

1 5 0 ns

) 1 0 0



(

- L 5 0

0 l l l l c C C C a a a a N N N N M M M M m m m m 1 . .2 .5 .1 7 5 5 1 1 3 8 7 1

Figure 2. 24. Increasing NaCl in the media has no effect on PLA degradation by B. pumilus B12. Concentrations were increased from the standard NaCl concentration found in minimal salt media (17.1 mM NaCl). ns indicates ANOVA p-value ≥ .05 against MSM+G control.

1 5 0

**** )

1 0 0



(

L -

L ns 5 0 ns

0 + + + + K K K K M M M M m m m m .5 5 .5 5 7 1 7 7 3

Figure 2. 25. B. pumilus B12 changes L-lactate release in K+ when normalized for growth. Initial measurements of B. pumilus B12 L-lactate release from PLA taken after 48 hours. K+ concentration increases are based off standard minimal salt media with glucose (MSM+G). These assays were performed with three biological and three technical replicates. **** indicates ANOVA p-value ≤ .0001 against MSM+G, ns indicates ANOVA p-value ≥.05 against MSM+G.

Figure 2. 26. Bacillus pumilus B12 growth rate changes with increases concentrations of potassium. Optical density-based growth curves of B. pumilus B12 in different K+ concentrations taken over 48 hours. Each growth curve represents 8 biological replicates.

Figure 2. 27. Bacillus pumilus B12 L-lactate release from PLA changes in different concentrations of potassium independent of growth. Relative release of L-lactate/cell based on growth after 48 hours. **** indicates ANOVA p-value ≤ .0001, *** indicates ANOVA p- value ≤ .001 and ns indicates t-test p-value ≥ .05 against MSM+G control.

2 5 0

2 0 0 **** )

M 1 5 0



(

t ****

t c

a 1 0 0 **

- L

5 0

0 - - - -

O 4 O 4 O 4 O 4 P P P P M M M M m m m m .4 .7 6 9 1 8 4 8 1 2

Figure 2. 28. B. pumilus B12 increases L-lactate release from PLA films with increasing - PO4 when growth is not considered. Initial measurements of B. pumilus B12 L-lactate release - from PLA taken after 48 hours PO4 concentration increases are based off standard minimal salt media with glucose (MSM+G). These assays were performed with three biological and three technical replicates. **** indicates ANOVA p-value ≤ .0001 against MSM+G.

0 .4 - 1 1 .4 m M P O 4 - 2 8 .7 m M P O 4 - 4 6 m M P O 4 - 8 9 m M P O 4

0 .3

)

0 0

6 0 .2

(

D O

0 .1

0 .0 0 1 0 0 0 2 0 0 0 3 0 0 0 T im e (M in )

Figure 2. 29. Bacillus pumilus B12 changes its growth rate in different concentrations of - PO4 . Optical density-based growth curves of B. pumilus B12 taken over 48 hours in different PO - concentration. Each growth curve represents 8 biological replicates. 4

Figure 2. 30. Bacillus pumilus B12 has reduced L-lactate release from PLA films when - there is increased PO4 . Relative release of L-lactate/cell based on growth after 48 hours. **** indicates t-test p-value ≤ .0001 and *** indicates ANOVA p-value ≤ .001 indicates against MSM+G control.

78 loss). The rapid nature of the assay allowed for more robust insight into how changing conditions in microbial growth have large impacts on how B. pumilus B12 degrades PLA.

Bacillus pumilus B12 showed degradation of PLA primarily when there was a supplementation of carbon in the medium, indicating that B. pumilus B12 will degrade PLA even if it is not using it as a sole carbon source. This suggests that hydrolytic activity against PLA is happenstance when scavenging for other nutrient and carbon sources in the environment. This is further supported by Bacillus pumilus B12 not growing on PLA films alone, but only growing on

PLA/PHA and other polymer blends. This data supports the notion that PLA degradation is occurring via a co-metabolism mechanism; with degradation of the polymer being dependent on the presence of a primary carbon source.

Although glucose supplementation allowed for higher amounts of detectable PLA degradation, the general trend in our study suggests that the more amenable the conditions for growth, the less PLA degradation occurs. The two carbon sources which increased L-lactate detection significantly, mannitol and maltose, both lowered the total biomass produced by B. pumilus B12 at the 48-hour timepoint. The same trend can be seen for the increases of lactate + degradation that occurred in response to specific concentrations of KH2PO4, K and PO4- in the media. As growth increased, the relative release in L-lactate released was reduced. Also, as high concentrations of K+ in the media started to impede growth, there was once again an observable increase in detectable L-lactate released from PLA. This data leads to the hypothesis that in more favorable, free nutrient conditions, B. pumilus B12 alters gene expression to reduce extracellular enzymes scavenging for alternative nutrient sources, inhibiting the transcription of hydrolytic enzymes.

When investigating a change in nitrogen sources/concentrations in the media, there was only an impact with adenine supplementation. This combined with the lack of alteration in PLA degradation with increased amino acids, ammonium and nitrate availability suggest that adenine alone is driving a transcriptional change in B. pumilus B12. The specificity to adenine over similar or traditionally favored sources of nitrogen downplays the notion of a nutrient driven response. This suggests adenine is acting as specific ligand for a purine riboswitch, negatively regulating downstream enzymes involved in PLA degradation. Riboswitches have been shown to both repress

79 and increase gene expression in Bacillus subtilis strains, suggesting similar functions in Bacillus pumilus B12 [30,31].

Overall, this study is the first step into exploring polymer degradation by Bacillus pumilus B12 and how degradation changes based on the environment it inhabits. In future work, we will be focusing on both the regulatory mechanisms involved in Bacillus pumilus B12 PLA degradation as well as the enzymes(s) acting upon the polymer. Understanding how the bacterium adapts to changing environments will lead to insight into why degradation rates change in real-world settings. Furthermore, this will garner solutions into synthetically altering the physiological machinery of microbial degradation. Identifying active enzymes(s) against PLA will open biochemical avenues of investigation. Both genetic and chemical modifications to active enzymes combined with controlling regulatory mechanisms can lead to a more effectual microbial polymer degrader.

References

1. Emadian, S. M., et al.. (2017). "Biodegradation of bioplastics in natural environments." 59: 526-536.

2. Singhvi, M. and D. Gokhale (2013). "Biomass to biodegradable polymer (PLA)." RSC Advances 3(33): 13558-13568.

3. Madkour, M. H., et al.. (2013). "PHA Recovery from Biomass." Biomacromolecules 14(9): 2963-2972.

4. Europeon Bioplastics (2017). 2020 Bioplastics Market Forecast. 10th European Bioplastics Conference in Berlin

5. Castro-Aguirre, E., et al.. (2016). "Poly (lactic acid)-Mass production, processing, industrial applications, and end of life." Adv Drug Deliv Rev 107: 333-366.

6. Tomita, K., et al.. (1999). "Isolation of thermophiles degrading poly (L-lactic acid)." J Biosci Bioeng 87(6): 752-755.

7. Fukuzaki, H., et al.. (1991). "In vivo characteristics of high, molecular weight copoly(l- lactide/glycolide) with S-type degradation pattern for application in drug delivery systems." Biomaterials 12(4): 433-437.

8. Oda, Y., et al.. (2000). "Degradation of Polylactide by Commercial Proteases." Journal of Polymers and the Environment 8(1): 29-32.

9. Masaki, K., et al.. (2005). "Cutinase-like enzyme from the yeast Cryptococcus sp. strain S- 2 hydrolyzes polylactic acid and other biodegradable plastics." Appl Environ Microbiol 71(11): 7548-7550.

10. Lim, H. A., et al.. (2005). "Hydrolysis of polyesters by serine proteases." Biotechnol Lett 27(7): 459-464.

11. Hanphakphoom, S., et al.. (2014). "Characterization of poly(L-lactide)-degrading enzyme produced by thermophilic filamentous bacteria Laceyella sacchari LP175." J Gen Appl Microbiol 60(1): 13-22.

12. Ahn, H. K., et al.. (2011). "Biodegradability of injection molded bioplastic pots containing polylactic acid and poultry feather fiber." Bioresource Technology 102(7): 4930-4933.

13. Mihai, M., et al.. (2014). "Formulation-properties versatility of wood fiber biocomposites based on polylactide and polylactide/thermoplastic starch blends." 54(6): 1325-1340.

14. Arrieta, M. P., et al.. (2014). "Disintegrability under composting conditions of plasticized PLA–PHB blends." Polymer Degradation and Stability 108: 307-318.

15. van Dijl, J. and Hecker, M. (2013). "Bacillus subtilis: from soil bacterium to super- secreting cell factory." Microbial Cell Factories 12(1):3

16. Schallmey, M., et al.. (2004). "Developments in the use of Bacillus species for industrial production." Canadian Journal of Microbiology 50(1): 1-17.

17. Link, L., et al.. (2004). "Extreme spore UV resistance of Bacillus pumilus isolates obtained from an ultraclean Spacecraft Assembly Facility." Microb Ecol 47(2): 159-163.

18. Handtke, S., et al.. (2014). "Bacillus pumilus reveals a remarkably high resistance to hydrogen peroxide provoked oxidative stress." PLOS ONE 9(1): e85625.

19. Bailes, G., Lind, M., Ely, A., Powell, M., Moore-Kucera, J., Miles, C., Inglis, D. & Brodhagen, M. Isolation of Native Soil Microorganisms with Potential for Breaking Down Biodegradable Plastic Mulch Films Used in Agriculture. JoVE, e50373, doi:10.3791/50373 (2013).

20. Hayes, D. G., Wadsworth, L. C., Sintim, H. Y., Flury, M., English, M., Schaeffer, S. & Saxton, A. M. Effect of diverse weathering conditions on the physicochemical properties of biodegradable plastic mulches. Polymer Testing 62, 454-467, doi:https://doi.org/10.1016/j.polymertesting.2017.07.027 (2017).

21. Benson DA, Karsch-Mizrachi I, Lipman DJ, Ostell J, Wheeler DL. GenBank. Nucleic Acids Res. 2004;33(Database issue): D34-8.

22. Kumar S., Stecher G., and Tamura K. (2016). MEGA7: Molecular Evolutionary Genetics Analysis version 7.0 for bigger datasets. Molecular Biology and Evolution 33:1870-1874.

23. Edgar, R.C., 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797

24. Hasim, S., Allison, D.P., Retterer, S. T., Hopke, A., Wheeler, R.T., Doktycz, M.J., & Reynolds, T.B. (2017) beta-(1,3)-Glucan Unmasking in Some Candida albicans Mutants Correlates with Increases in Cell Wall Surface Roughness and Decreases in Cell Wall Elasticity. Infect Immum, 85(1)

25 Li, S., & McCarthy, S. (1999). Influence of Crystallinity and Stereochemistry on

The Enzymatic Degradation of Poly(lactide)s. Macromolecules, 32(13), 4454-4456. doi:10.1021/ma990117b

26. Newman, M. M., et al.. (2016). "Changes in rhizosphere bacterial gene expression following glyphosate treatment." Science of The Total Environment 553: 32-41.

27. Bargaz, A., et al.. (2018). "Soil Microbial Resources for Improving Fertilizers Efficiency in an Integrated Plant Nutrient Management System." 9(1606).

28. Schramke, H., et al.. (2017). "Revisiting regulation of potassium homeostasis in Escherichia coli: the connection to phosphate limitation." MicrobiologyOpen 6(3): e00438.

29. Botella, E., et al.. (2011). "Cell envelope gene expression in phosphate-limited Bacillus subtilis cells." 157(9): 2470-2484.

30. Mandal, M. and R. R. Breaker (2004). "Adenine riboswitches and gene activation by disruption of a transcription terminator." Nat Struct Mol Biol 11(1): 29-35.

31. Christiansen, L. C., et al.. (1997). "Xanthine metabolism in Bacillus subtilis: characterization of the xpt-pbuX operon and evidence for purine- and nitrogen-controlled expression of genes involved in xanthine salvage and catabolism." J Bacteriol 179(8): 2540-2550.

Appendix

Figure S2. 1. Atomic force microscopy data for B. pumilus B12 grown in KH2PO4. A) Tensile strength measurements of PLA in different media condition. **** indicates ANOVA p-value ≤ .0001 against MSM+G control. AFM images of PLA films grown with B. pumilus

B12 in 11 mM (B), 55 mM (C) and 110 mM (D) KH2PO4 respectively.

- Figure S2. 2. Atomic force microscopy data for B. pumilus B12 grown in PO4 . A) Tensile strength measurements of PLA in different media condition. **** indicates ANOVA p-value ≤ .0001 against MSM+G control. AFM images of PLA films grown with B. pumilus B12 in 28.7 mM (B), 46 mM (C) and 89 mM (D)PO4- respectively.

Figure S2. 3. Atomic force microscopy data for B. pumilus B12 grown in K+. A) Tensile strength measurements of PLA in different media condition. ****,* and ns indicate ANOVA p-value of ≤ .0001, .01 and ≥.05 against MSM+G control respectively. AFM images of PLA + films grown with B. pumilus B12 in 15 mM (B), 37.5 mM (C) and 75 mM (D) K respectively.

Chapter III: Bacillus pumilus B12 Alters its Gene Expression in Conditions that Reduce PLA Degradation

Contributions

Contributions of this Chapter are as follows: Genomic DNA, RNA and Extracellular and Cellular protein extractions were performed by Kyle Bonifer. DNA sequencing was performed by the University of Indiana Center for Genomics and Bioinformatics with contig construction and gene annotation performed by Kyle Bonifer. Amino Acid comparisons between Bacillus species were performed by Eric Gann and Kyle Bonifer with amino acid alignment performed by Kyle Bonifer. Silver stain gels and biotin Western blots were performed by Kyle Bonifer with mass spectrometry performed by Paul Abraham in the Hettich lab at ORNL. RNA sequencing was performed by HudsonAlpha with data analysis performed by Kyle Bonifer. L-lactate assay was performed by both Kyle Bonifer and Elise Phillips.

Abstract

Poly-lactic acid (PLA) is a biodegradable polymer that is increasingly being used as an alternative to petroleum-based plastics. In this study we build upon previous work in identifying microbes and conditions that degrade PLA to elucidate the active enzymes and regulatory machinery involved in microbial PLA degradation. We performed whole genome sequencing on our previously identified Bacillus pumilus strain, B12, and used proteomic and transcriptomic techniques to detect genes that are associated with PLA degradation. First, we extracted proteins from an organic nitrogen rich condition that had reduced PLA degradation by B. pumilus B12 and observed a reduction in extracellular protein bands. These protein bands disappeared within 48 hours of incubation, aligning with our timepoint for measuring PLA degradation. Our mass spectrometry results yielded a total of 43 protein species with 16 showing differing concentrations, between our two conditions. Amongst these were two ubiquitous proteins with defined proteolytic regions and the flagella associated protein, FlaA. RNA sequencing of the normal and organic nitrogen rich media incubations showed 941 genes downregulated and 1600 genes upregulated in the lower plastic degradation condition. These genes are associated with various cellular metabolic pathways dealing with amino/nucleic/fatty acid biosynthesis and uptake as well as a variety of other physiological pathways. Further expression differences include 34/42 of identified motility genes and 17/22 chemotaxis genes being negatively expressed, indicating that motility plays a role in PLA degradation. Furthermore, two of the major protein export systems, Sec and Tat, are negatively expressed in reduced PLA degradation conditions. These results indicate regulatory changes occurring in Bacillus pumilus B12 that affect the amount of degradation. This research is a steppingstone for identifying physiological features of this Bacillus sp. that are important for plastic degradation.

Introduction

The production of plastic polymers is having a profound impact on the environment. Many of these polymers are derived from petroleum products, making their production as well as longevity problematic [1]. One possible alternative is that these petroleum polymers are bio-based biodegradable plastics that do not require petroleum extraction and are readily degraded in soil environments [2]. This degradation is driven primarily by the production of enzymes from the microbial communities growing at the site [3,4]. The enzymes these microbes produce target the linkages between monomers, allowing for uptake and utilization in the cell [5]. This process can be beneficial to the ecosystem, increasing utilizable carbon for microbes, potentially replenishing eroded soil [5].

PLA is the one of the most common biodegradable polymers utilized in commercial industry settings [6]. PLA is an aliphatic polyester formed from direct condensation of lactic acid molecules or from ring polymerization of lactide [7]. This plastic can be found in many commercial products including eating utensils, 3-D printing, food containers and packaging [8,9,10,11]. Due to the chemical structure of PLA, containing large numbers of ester linkages, microbial derived enzymes have been shown to readily target and hydrolyze the plastic back down into its lactic acid monomers [12,13,14,15,16]. This increases the amount of free L-lactate found in the system, with many species of bacteria utilizing this as a carbon source This process increases the usable carbon in the system, simultaneously reducing the mass associated with the plastic.

A ubiquitous genus found in the soil ecosystem is Bacilli. Members of this gram-positive rod-shaped bacterial genus are renowned for their adaptability to diverse environments and their large assortment of extracellular enzymes [17,18]. This enzymatic production and genetic tractability have made Bacillus subtilis and Bacillus megaterium favored strains in industry as vectors for enzyme production [17,19]. More recently, strains of Bacillus pumilus have been shown to be highly resistant to desiccation, oxidative stress and radiation, and are being investigated for biotechnological applications [20,21,22].

This process of PLA degradation has been observed in many fungal and bacterial species, with degradation of macroscopic plastics occurring readily in composting settings [23,24,25]. However, very little work has been put into understanding how microbes regulate their rate of plastic degradation given altered environmental and growth parameters. Here we continue to

91 investigate a previously identified strain of Bacillus pumilus (B12) that is capable is poly-lactic acid (PLA) degradation. We sought to elucidate regulatory mechanisms and enzymes instrumental in B. pumilus B12 PLA degradation. To accomplish this, we performed whole genome sequencing, protein extraction, RNA sequencing and molecular genetics to help identify genes pivotal in the degradation process. We found significant transcriptomic differences in growth conditions where B. pumilus B12 plastic degradation changes. Namely, differences were observed in expected pathways connected to amino/nucleic acid metabolism and biosynthesis with changes also occurring in motility genes, quorum sensing/chemotaxis and protein export systems. The changes in the expression of genes attributed to cell movement, chemical detection and protein secretion suggest these microbial processes are involved in identifying, colonizing and degrading PLA films.

Methods

Growth Conditions of Bacillus pumilus B12

Bacillus pumilus B12 cultures were grown in static conditions at 30° C in minimal salt media for 48 hours. Minimal salt media (MSM) was composed of 7.5 mM NH4SO4, 11 mM

KHPO4, 2.87 mM K2HPO4, 1.7 mM MgSO4 and 17.1 mM NaCl, pH 7.0. When adding complete supplementation (CSM) media to liquid cultures, MP Biomedicals™ CSM-HIS-LEU-URA Powdered Media (Fisher-MP114531212) was used at a concentration of 6.5 g/L. CSM media contains various amino acids and adenine.

Whole Genome Sequencing and Analysis

Bacillus pumilus B12 genomic DNA was isolated via phenol-chloroform extraction and cleaned using RNase treatment. Genomic samples were prepped with a NEXTflex-Rapid-DNA- Seq kit and multiplex sequencing barcodes were added with the NEXTflex-DNA-Barcode-12 kit. The samples were cleaned with double-sided beadcut and verified with a Qubit2 fluorometer and a 2200 Tapestation bioanalyzer. Samples were pooled and then sequenced using an Illumina NextSeq 500 instrument, resulting in 150x150 paired end reads. These were de-multiplexed using bcl2fastq software before any bioinformatic analysis performed. Raw sequencing reads were trimmed and assembled using the de novo sequencing assembly on CLC Genomics Workbench™ v.12.0 software with a mismatch cost of 2 and a similarity fraction of .95. Contigs resulting from

92 this assembly were separated by size and coverage cutoffs of 500 base pairs and 100x coverage respectively. The 34 resulting contigs were uploaded to the Rapid Annotation using Subsystem Technology (RAST) for annotation and amino acid level comparison to other microbial species using BLASTp. Amino acid sequences generated from RAST were aligned against a known proteolytic domain found in the Flagella of Clostridium haemolyticum [26] using the Clustal Omega Multiple Sequence Alignment tool.

L-lactate Detection Assay.

Detection of L-lactate released into the supernatant was performed using ENZYFLUO L- lactate kit (Bioassay Systems Inc.). Overnight cultures were grown in Lennox Broth (LB) shaking at 225 RPM at 30° C and washed three times with phosphate buffered saline. An optical density of 0.1 was incubated in 1mL of media on 10 mg/mL PLA films in glass vials for 48 hours. Supernatant was taken off cultures centrifuged at 13,000 RPM for 2 minutes from 10 mg/mL PLA films. A 1:1 mixture of supernatant collected from polylactic acid films was added to the L-lactate kit detection solution giving a total volume of 100 µL in a 96-well plate. The supernatant/detection solution was incubated in the dark at room temperature for 1 hour and colorimetric changes were measured using a 535/580 nm excitation/emission ratio in a Biotek Cytation5 plate reader. Results were normalized to a standard curve generated with L-lactate standards ranging from 0-42 µM. All measurements were added to the slope generated by the standard curve, yielding approximate concentrations of L-lactate release. All trials included negative controls containing cultures with no PLA added, as well as PLA films in media only. Measurements were taken with 3 biological replicates each with 3 technical replicates, yielding nine total data points per treatment. Statistical significance was determined via t-test with p-values below .01 considered significant.

Protein Isolation, Siler Staining and Biotinylation

For supernatant samples, a 250 mL culture was centrifuged at 4,000 revolutions per minute (RPM) for 20 minutes and then passed through a 0.22 µm filter in order to remove cells. The supernatant was then acetone precipitated overnight at -20℃ and proteins were collected by spinning the mixture at 10,000 RPM for 1 hour. Proteins concentrations were then measured using a Bradford assay and normalized to 5µg/mL. Proteins were then linearized by boiling for 5 minutes and run on a 10% SDS-page gel and silver stained using a Thermo Scientific™ Pierce™ silver stain kit (Fisher-PI24612).

For biotinylation of cell surface proteins, cell pellets from a centrifuged 250 mL culture were washed 3 times with phosphate buffer saline and biotinylated using EZ-Link™ Sulfo-NHS- LC-Biotinylation Kit). After biotinylation, samples were chemically and physically lysed using cell lysis buffer (1M NaCl, .5% SDS, 50mM Tris 7.4 pH) and 150-212 µm sand grains. Cell/lysis buffer mixtures were shaken using a BioSpec16 mini bead beater three times for 30 seconds. Next, protein concentrations were normalized using Bradford assay, mixed with Laemmli buffer and boiled for 5 minutes. Following protein isolation, normalization and biotinylation, samples were fractionated by a 12% SDS-page gel and transferred to Immobilon®-Fl transfer membranes overnight. Blots were then incubated with a conjugated avidin-GFP probe which detects and tightly binds to the pre-bound biotin, and the blots were washed to remove excess antibody. Blots were then visualized using a LiCor Odyssey CLx in the green fluorescent wavelength spectrum.

Extracellular protein samples for mass spectrometry were isolated from 2 liter cultures centrifuged at 4,000 RPM for 20 minutes and passed through a 0.22 µm filter in order to remove cells. The supernatant was then acetone precipitated overnight at -20℃ and proteins were collected by spinning the mixture at 10,000 RPM for 1 hour. The subsequent 500 mL of precipitated protein was further concentrated using tangential flow filtration (TFF) resulting in approximately 5 mL of supernatant protein at a concentration between 6-7 mg/mL measured by Bradford assay. Each condition was performed in triplicate.

All samples were analyzed on a Q Exactive Plus mass spectrometer (Thermo Fischer Scientific) coupled with a with a Proxeon EASY-nLC 1200 liquid chromatography (LC) pump (Thermo Fisher Scientific). Peptides were separated on a 75 μm inner diameter microcapillary column packed with 40 cm of Kinetex C18 resin (1.7 μm, 100 Å, Phenomenex). For each sample, a 2 μg aliquot was loaded in buffer A (0.1% formic acid, 2% acetonitrile) for 10 min, eluted with a linear 90 min gradient of 2–20% buffer B (0.1% formic acid, 80% acetonitrile), followed by an increase in buffer B to 30% for 10 min, another increase to 50% buffer for 10 min, and concluding with a 10 min wash at 98% buffer A. The flow rate was kept at 200 nL min-1. MS data was acquired with the Thermo Xcalibur software version 4.27.19, a topN method where N could be up to 10. Target values for the full scan MS spectra were 1 x 106 charges in the 300 – 1,500 m/z range with a maximum injection time of 25 ms. Transient times corresponding to a resolution of 70,000 at m/z 200 were chosen. A 1.6 m/z isolation window and fragmentation of precursor ions was performed by higher-energy C-trap dissociation (HCD) with a normalized collision energy of 30 94 eV. MS/MS sans were performed at a resolution of 17,500 at m/z 200 with an ion-target value of 1 x 105 and a maximum injection time of 50 ms. Dynamic exclusion was set to 20 s to avoid repeated sequencing of peptides.

MS raw data files were searched against the Bacillus pumilus B12 proteome generated in RAST annotation. A decoy database, consisting of the reversed sequences of the target database, was appended in order to discern the false-discovery rate (FDR) at the spectral level. For standard database searching, the peptide fragmentation spectra (MS/MS) were analyzed by Proteome Discoverer v2.2 (Thermo Fisher Scientific). The MS/MS were searched using the MS Amanda v2.0 and were configured to derive fully tryptic peptides using settings for high-high MS/MS data: MS1 mass tolerance of 5 ppm and MS2 mass tolerance of 0.02 Da. A static modification on cysteines (iodoacetamide; +57.0214 Da), a dynamic modification on methionine (oxidation; 15.9949) and aspartate and glutamate (methylation; 14.016) were considered. The results were processed by Percolator to estimate q values. Peptide spectrum matches (PSMs) and peptides were considered identified at a q value <0.01.

RNA Isolation and Gene Expression Mapping

Bacillus pumilus B12 RNA was isolated from 500 mL cultures grown in in static conditions at 30℃ for 48 hours. Cultures were centrifuged at 10,000 RPM for 1 minute, the supernatant was removed and 10 mL of Trizol was added to the cell pellet. The Trizol/ Pellet mixture was separated into 200 µL volumes in screw cap tubes where ≤106 µm acid washed beads and 200 µL diethyl pyrocarbonate (DEPC) treated phosphate buffered saline was added. The samples were then shaken using a Biospec bead beater 3 times for 30 seconds in a cold room. Next, samples were spun at 13,000 RPM for 10 minutes, and the supernatant was moved to a new microcentrifuge tube containing 300 µL of ice cold 100% ethanol (EtOH). Ethanol washing of the supernatant fraction was repeated until the upper fraction appeared clear. The supernatant fraction was placed at -80℃ for 20 minutes and spun at 13,000 RPM to pellet the RNA. Pellets were washed with 70% ethanol and allowed to air dry at 37℃ to remove excess ethanol. Samples were then resuspended in RNase free water and combined into a single microcentrifuge tube. RNA samples were then DNase treated using an Invitrogen turbo DNase kit. RNA concentration and integrity were determined using RNA gel and through a 2100 series bioanalyzer (Agilent Technologies, USA).

RNA sequencing was performed by HudsonAlpha™ using an Illumina™ Miseq garnering approximately 38 million reads of 150 bp in length per condition. All bioinformatic analysis was performed in CLC Genomics Workbench™ v.12.0 software. Trimming was performed with a quality score 0.01 and ambiguity set to 2 with trimmed reads below 125 bp were discarded. Reads were mapped to the 43 contigs assembled previously from our Bacillus pumilus B12 strain. Genome annotations were performed using our previously generated annotations list from RAST. To identify differentially expressed genes between our MSM+G and MSM+CSM+G conditions p-values of individual genes were adjusted for false discovery rate and genes with an FDR adjusted p-value= ≤ .05 were deemed significant. Expression values above or below 2 with a significant p- value were considered upregulated and downregulated respectively. For clustering genes into pathways, KEGG IDs from Bacillus pumilus SAFR-032 were matched to amino acid sequences with similarity lower than 1e-10 to B. pumilus B12 and run through the Kyoto Encyclopedia of Genes and Genome (KEGG) to build pathways.

Live/Dead Staining

Cell viability was determined by staining the PE films in a Baclight® live/dead cell viability kit and observed with fluorescent microscopy. Cells were inoculated onto 10 mg/mL PLA films for 48 hours in either MSM+G or MSM+G+CSM conditions. Films were then removed and treated with 3µL of SYTO-9 dye and propidium iodide to determine live and dead cell populations respectively. PLA films were then observed under both red and green emission ranges via fluorescent microscopy.

B. pumilus B12 Cloning

For electroporation, an overnight culture of B. pumilus B12 was reinoculated into 50mL of fresh Lennox Broth (LB) at 0.1 OD and allowed to grow to mid log phase (approximately 1.5

OD). The cells were then washed three times in ddH2O to remove excess salts and resuspended in 500 µL of 30% polyethylene glycol (PEG) 6000. Cells were electroporated with the resistance set to 700 Ω with the field strength set to 12.5 kV/cm with 1µg of DNA added to 100 µl of cells. Cells were then placed in SOC media and allowed to recover for three hours before being placed on LB with 100 µg/mL tetracycline for selection. GFP expression was determined using the green emission range via fluorescent microscopy.

Results

Bacillus pumilus B12 is genetically similar to other fully sequenced B. pumilus strains

In order to properly investigate the proteomic and transcriptional changes between nutrient conditions, we first needed to sequence the whole genome of our novel isolate of Bacillus pumilus. Sequencing results most closely matched a laboratory strain of Bacillus pumilus SAFR-032, with another B. pumilus strain (ATCC 7061) being the next closest match based on BLASTP scores generated from RAST (Table 3. 1). B. pumilus B12 has a genome size of approximately 3.79 Mb with a GC content at 41.3% and 3971 putative coding regions (Figure 3. 1.). These numbers are in the approximate range of several other B. pumilus strains that have been fully sequenced (Table 3. 1.). Furthermore, amino acid alignment of all putative proteins to B. pumilus SAFR-032 , shows most proteins in B. pumilus B12 can A) be found in SAFR-032 and B) have at least an 80% similarity match in amino acid sequence. Although similar to other B. pumilus strains, other Bacillus pumilus strains have wide variance in their ability to degrade PLA films (Figure 3. 2.). Of the 5 other B. pumilus strains tested for PLA degradation only one, 9509.4, showed comparable amounts of PLA degradation to B. pumilus B12 (Figure 3. 2.). These results show that our novel isolate is genetically similar to other B. pumilus strains previously isolated, however degradation of PLA differs. This implies that the mechanisms on PLA degradation may be a change seem at a proteomic or transcriptomic level.

Supernatant Proteins Released by Bacillus pumilus B12 Change in Media Rich with Organic Nitrogen Sources

We next wanted to investigate if there were any differences in the extracellular proteomic profile seen between conditions which previously showed altered levels of PLA degradation (Chapter II). Nutrient conditions rich in amino acids and adenine had previously showed a decrease in PLA degradation while conditions with a Supplemental carbon source of maltose and mannitol showed an increase in degradation (Chapter II) In order to visualize differences in the proteomic profile, we performed silver staining on SDS-page gels in order to detect band differences between conditions. For conditions with an increase in PLA degradation; maltose and mannitol, we saw no visible difference in the protein bands (Supplemental Figure 3. 1.) that were not a result of loading concentrations between lanes. However, there was a pronounced decrease in extracellular protein bands for B. pumilus B12 grown in media supplemented CSM (Figure 3. 3A.). This loss of in

Table 3. 1. Bacillus pumilus B12 is most closely related to other B. pumilus strains based on amino acid similarity. Genome size, GC content and coding regions were collected from the corresponding genomic data on NCBI.

2 A. B.

Figure 3. 1. Bacillus pumilus B12 is similar to B. pumilus SAFR-032 at both a genomic and amino acid level. A) Information gathered from RAST annotation of B12 strain contigs giving an overview of genome size, coding regions and closest phylogenetic match. B) Similarity map of amino acid coding sequences overlaying B. pumilus SAFR-032 to B. pumilus B12. Color corresponds to percent of sequence identity, with white spaces representing coding sequences not found in B. pumilus SAFR-032.

3 0 P L A N o P L A ****

****

2 0

l e

r *

t c

a **

- L 1 0 **

0 B 1 2 M T C C 2 7 A 1 2 5 9 5 0 9 .4 N B 5

Figure 3. 2. Other strains of Bacillus pumilus have varying levels of PLA degradation as measured by L-lactate release. L-lactate released (measured in µM) by four strains of B. pumilus with fully sequenced genomes (MTCC, 27A,

125, 9509.4) and one laboratory strain of Bacillus aerius (NB5). Results show L-

lactate released from 10 mg/mL PLA films incubated with a Bacillus spp. for 48 hours (pink bars) and L-lactate released from Bacillus spp. incubated without PLA

films for 28 hours. ****,**,*,ns indicates t-test p-values ≥.0001, ≥.001, ≥ .01 and ≤.05 respectively.

100

A. B.

Figure 3. 3. Extracellular proteins released from B. pumilus B12 go down when incubated with complete supplement media (CSM) addition to the media. A) Silver stain of extracellular proteins collected from B. pumilus B12 following 48 hours of incubation on 10 mg/mL PLA films. Each lane corresponds to a different concentration of CSM added to the media above 1x which is the standard concentration. B) Silver stain gel of B. pumilus B12 grown in 10x CSM media with 10 mg/mL PLA films taken a different timepoints over the course of a week. For lanes labeled with “supp”, fresh media was supplemented onto the PLA films 24 hours prior to protein collection. No CSM indicates proteins collected from MSM+G media conditions after 48 hours and the “- control” lane is 10x CSM media incubated for 48 hours without any bacterial inoculum.

101 protein bands decrease as the concentration of CSM increases with a noticeable disappearance above and below 55 kDA at 4x standard concentration (MP Biomedicals™). Furthermore, protein band disappearance begins to occur following two days of incubation in the CSM supplemented media (Figure 3. 3B.). At the two-day time point, the most noticeable band loss was noticed between 15-55 kDa, a range in which a large number of hydrolytic esterases including several species of lipases and proteases are typically found [27,28]. Both the concentration and the timeframe of band disappearance matches our previous analysis showing PLA degradation decreasing at these same timepoints and concentrations (Chapter II). This evidence suggests that the CSM media is changing the expression of extracellular proteins which impact B. pumilus B12’s ability to degrade PLA.

Proteomic Sample Analysis Shows a Reduction in Total Extracellular Protein

Next, to see if the extracellular bands changes correspond to changes in enzymes released the could potentially target PLA, we ran mass spectrometry to identify protein species. Proteins were concentrated to 400-fold their native concentration and run in triplicate on a Q Exactive Plus mass spectrometer. We observed a total of 43 protein species in both the MSM+G and MSM+CSM+G conditions based that matched backed to amino acid sequences of our B. pumilus B12 genome. Sixteen of these proteins showed decreases in the CSM supplemented conditions compared to our MSM+G control (Figure 3. 4A., 3. 4B.). Of these 16, several have enzymatic activity, with three possibly containing proteolytic domains capable of recognizing the ester bonds holding together PLA films. Several proteins identified are considered cytoplasmic, with three being involved in glycolysis (enolase, fructose 1,6-biphosphatase and phosphogluconate dehydrogenase). Other putative cytoplasmic proteins that show no hydrolytic activity include nucleotidyltransferase which functions by dephosphorylating nucleotides, adenylosuccinate lyase which breaks a C-N bond in adenylosuccinate to form adenosine monophosphate (AMP) and fumarate, and reductoisomerase which reduces oxygen in ketones and is associated with the isoprenoid biosynthesis in Bacillus spp. [29]. Of the proteins with possible proteolytic domains, two are located within the cytoplasm, with the third found outside of the cell wall. RecA, a major protein involved in DNA maintenance and repair contains a proteolytic region that cleaves the SOS DNA repair repressor LexA, requiring ATP to function [30,31,32]. The other cytoplasmic protease is a Clp-protease which has been shown to be involved in protein maintenance by targeting and degrading misfolded proteins [33]. The final possible location for a proteolytic 102

A. B.

Figure 3. 4. Mass spectrometry results of differentially abundant Bacillus pumilus B12 extracellular proteins. A) Extracellular protein abundances of B. pumilus B12 in MSM+G (Blue) and MSM+G+CSM (RED) conditions for 16 proteins observed to have total abundance differences across three replicates. B) List of corresponding protein identities for protein species detected.

103 domain located outside of the cell could be found the flagella of B pumilus B12. Previous studies have shown that the flagella of dozens of bacteria from an array of phylogenetic lineages contain proteolytic domains integrated between the N and C terminals of certain flagellin [26]. Due to FlaA being identified as a protein with lowered concentrations in our CSM condition, we investigated the B. pumilus B12 FlaA for this proteolytic domain. The FlaA from B. pumilus B12 showed a low alignment score with the conserved proteolytic domain found in Clostridium haemolyticum flagellin (Supplemental Figure 3. 2.). Altogether, our results from the extracellular proteome did not reveal any enzymatic targets that would hydrolyze PLA plastic films outside of the cell leading to degradation, with the two identified proteins having proteolytic activity being found within the cytoplasmic space.

We next turned to protein expression on the surface of the bacteria, hypothesizing that hydrolytic targets were attached to the cell wall rather than excreted into the extracellular space. We biotinylated surface proteins on cells grown in all media conditions where we saw a decrease 4 - + in L-lactate release from PLA films: CSM, KH2PO , PO4 and K , observed them on an SDS-page gel and fluorescently tagged them using an avidin-GFP conjugated monoclonal antibody. Compared to our MSM+G control, we saw no noticeable band changes for any of our conditions (Supplemental Figure 3. 3.). These results coupled with our proteomic data did not yield viable targets of enzymatic activity against the PLA films, emphasizing the need to elucidate difference in genetic expression in our separate PLA degradation conditions.

Gene Expression Differs Dramatically Between Bacillus pumilus B12 Grown in Rich Organic Nitrogen Conditions Compared to Minimal Conditions

Next, to understand how Bacillus pumilus B12 changes its genetic expression in conditions with altering PLA degradation, we performed RNA sequencing and compared them to our MSM+G (PLA degradation) control condition to that supplemented with CSM (reduced degradation). Using Illumina Miseq, we generated 35 million 150bp reads between 4 libraries and mapped them to our previously generated B. pumilus B12 genome. Each library was contained two technical replicates to ensure successful and consistent sequencing. A principal component plot (PCA) was created by using the RNA Analysis Package in CLC Genomic Workbench software to determine the variabilities between our two different growth conditions and our variables. Our

104

Figure 3. 5. PCA plot displays the clustering of samples within each condition. PCA plot was created using CLC Genomics Workbench software (V12.0) to determine the variabilities between replicates and variables.

105

MSM+G and MSM+G+CSM RNA samples formed distinct clusters independent of one another, suggesting that there was little variance between our samples (Figure 3. 5.). To further analyze our sequencing results, clustering heat map was constructed using all gene expression data for all of the 8 samples run. We observed a nearly identical overlap of runs using the same samples and overall close alignment between growth conditions (Figure 3. 6.). Furthermore, the MSM+G and MSM+G+CSM conditions have distinct gene expression profiles from one another and cluster dissimilarly from one another. Also, proteins observed to be differentially abundant in our mass spectrometry results had parallel expression results in our RNA sequencing data (Supplemental 3. 4A.) When observing the expression changes between our two conditions we saw a large shift in the gene expression profile in B. pumilus B12. Compared to MSM+G conditions, B. pumilus B12 upregulates 1681 genes and downregulates 941 genes with 1296 genes expression remaining statistically unchanged (Figure 3. 7A., 3. 7B.). These results show B. pumilus B12 has a vastly different gene expression profile in media supplemented with amino and nucleic acids.

To further understand how B pumilus B12 gene expression affects its cellular physiology and how it may relate to PLA degradation, we grouped genes into pathways using the Kyoto Encyclopedia of Genes and Genome (KEGG) pathway mapper tool. A total of 2076 genes (53.0%) were matched based on predicted amino acid sequences to the KEGG database to be mapped into pathways, giving us a representative look at the changing metabolism and physiology of B. pumilus B12. When looking into biosynthetic pathways in CSM media conditions, we saw large changes in gene expression in expected pathways pertaining to amino acid biosynthesis with far more genes being down regulated in these pathways (Figure 3. 8.). Due to these strains being grown in an amino acid rich environment, a reduction in energetically taxing systems related to making said molecules is expected. B. pumilus B12 has a general upregulation of genes involved in synthesizing fatty acids, peptidoglycan and folate. This could possibly indicate cells dividing more rapidly in the more nutrient rich environment supplemented with CSM as these three pathways are necessary for cell membrane, cell wall and diverse biological functions, respectively. The metabolic pathways show a slight increase in amino acid metabolism with increases also seen in the genes associated with the metabolism of sugars and lipids (Figure 3. 9.). This is in contrast with several other metabolic pathways that decreases the metabolism of nucleic acids, selenocompounds, methane, butanoate, sulfur and porphyrin. L-lactate metabolism and uptake was also generally down regulated, supporting the reduction in L-lactate being released from PLA in

106

Figure 3. 6. RNA expression phylogenetic heatmap shows high similarity between replicates and divergence between variables. A phylogenetic heat map was created using

CLC Genomics Workbench software (V12.0). Each biological replicate was run twice

(designated with either “a” or “b”). Numbers of reads mapped to the B. pumilus B12 genome representing on a sliding colorimetric scale with blue showing lower total mapped reads and red representing high total mapped reads.

107

A. B.

Figure 3. 7. Bacillus pumilus B12 differentially expresses the majority of its genes in MSM+G+CSM conditions compared to MSM+G conditions. A) Pie chart showing number the of genes upregulated, downregulated and unchanged based on a -2 to 2-fold expression change cutoff and an FDR adjusted p-value= ≥.01. B) Volcano plot showing the log10 fold expression change (x-axis) and statistical significance (y-axis) of all genes in MSM+G+CSM conditions compared to MSM+G.

108

D o w n re g u la te d G e n e s

K e to n e s U p re g u la te d G e n e s

C a ro te n o id

U b iq u in o n e

C a rb a p e n e m

M o n b a c tu m y

a P o ly k e tid e Su ga r

t a

P P a n to th e n a te

T e ra p e n o id

F o la te

P e p tid o gly c a n

F a tty A c id

A m in o A c id

0 0

0 5 0 0

0 5

1 1 2 5

1 1 N u m b e r o f G e n e s

Figure 3. 8. Number of biosynthesis associated genes differentially expressed sorted by into pathways using KEGG pathway mapper. X- axis represents the number of significantly differentially expressed genes identified. Orange bars represent genes significantly downregulated and pink bars represents genes significantly upregulated in CSM conditions compared to MSM conditions.

109

D o w n re g u la te d G e n e s

P o rp h y rin U p re g u la te d G e n e s

B u ta n o a te

N ic o tin im id e

Se le n o c o m p o u n d

N u c le ic a c id

M e th a n e y

a Su lfu r

h t

a N itro ge n P

A m in o A c id

P ro p a n o a te

L ip id

G ly o x y la te

P y ru v a te

Su ga r

0 0

0 0 0 0

0 5

1 2 3 5

1 1 N u m b e r o f G e n e s

Figure 3. 9. Number of metabolism associated. genes differentially expressed sorted by into pathways using KEGG pathway mapper. X- axis represents the number of significantly differentially expressed genes identified. Orange bars represent genes significantly downregulated and pink bars represents genes significantly upregulated in CSM conditions compared to MSM conditions.

110

Sty re n e D o w n re g u la te d G e n e s U p re g u la te d G e n e s N a p h th a le n e

K e to n e s

X y le n e

G e ra n io l

a w

h A m in o b e n zo a te s

a P C h lo ro a lk e n e s

B e n zo a te

A ro m a tic s

F a tty A c id s

A m in o A c id s

5 0 0 5 5 0

0 5

1 1 1 2 2 3

N u m b e r o f G e n e s

Figure 3. 10 Number of degradation associated genes differentially expressed sorted by into pathways using KEGG pathway mapper. X- axis represents the number of significantly differentially expressed genes identified. Orange bars represent genes significantly downregulated and pink bars represents genes significantly upregulated in CSM conditions compared to MSM conditions.

111 the MSM+CSM+G conditions (Supplemental Figure 3. 4B.) (Chapter II). When we looked at the changes in biodegradation pathways, most have generally similar numbers of up and downregulated genes, with fatty acid and amino acid degradation pathways both being slightly more downregulated than the MSM+G control condition (Figure 3. 10.). These results show a wide-ranging change across many metabolic and biosynthetic pathways showing the nutrient conditions that are associated with reduced B. pumilus B12 PLA degradation have a distinct metabolic profile from PLA degrading conditions.

We next investigated the expression of pathways associated with both cell motility and protein export. This is due to our mass spectrometry results showing a decrease in both the flagellin protein FlaA as well as a general decrease in extracellular proteins in lower PLA degrading conditions seen in silver stains (Figure 3. 3., 3. 4.). When looking at KEGG pathways designated as cell motility and chemotaxis exhibited large decreases in the gene expression for both of these processes compared to MSM+G (Figure 3. 11A., 3. 11B.). For genes associated with cell motility, we saw 34/42 genes were significantly downregulated with only 4/42 showing a significant increase above PLA degrading conditions. These genes range from flagellar assembly (Fli and Flg family) and motor proteins (MotA and MotB) to chemoreceptors (“Che” gene family) (Figure 3. 10A.). Genes that were clustered into chemotaxis pathways had a large amount of overlap with genes clustered into motility, and the same can be seen in both of the pathways (Figure 3. 10B.). Of the 22 genes, 17 were downregulated, with only 4 being upregulated. Out of the 4 genes shown to be upregulated, 3 are annotated as homologs of the same protein, further narrowing the number of genes associated with chemotaxis that are positively regulated. Finally, examining genes associated with protein export, we once again see a down regulation in conditions with reduced plastic degradation. Specially genes associated with the Sec and Tat protein export systems are both negatively expressed, with 4/5 Sec associated genes and both Tat associated genes significantly reduced compared to MSM+G (Figure 3. 10C.). These results indicate that cell motility and export of proteins via the Sec and Tat secretion systems may be necessary components for PLA degradation in B. pumilus B12.

112

Figure 3. 11. In CSM supplemented conditions, B. pumilus B12 significantly down regulates motility, chemotaxis and protein export pathways. Heat maps showing individual expression values for genes in a given pathway with colorimetric indicator of expression seen on the right of each heatmap. A) Shows motility gene expression, B) shows chemotaxis gene expression and C) shows protein export gene expression.

113

A. M o tility G en e E x p ressio n B. C h em o tax is G en e E x p ressio n

C h e P _ 3 C he R _ 1 C h e R C he B /R fusio n F la A _ 4 C he R _ 2 5 T c y A C he R F lgG F liG F lgF C he B F lM F liC F liM 5 F liG F lin/F liY F liH M o tA F liF M o tB F liA C he A F lgE 0 F liE C he W F liL C he C F lgD C he D C h e W C he Y F la ge lla r_ P ro te in _ 2 C he V F lh A R ib o se T ra nsp o rt S yste m F lgK 0 F liQ C he B C he V M o tA -5 C h e V C he C C h e C C he Y F la ge llin C he D F lgC F lgL P ro tein E x p o rt C h e A F lh B C. F liM F liR S ip W flb D -5 C h e Y se c D F 5 C h e D F liZ le p B F liD F liS lsp A F liN M o tB yid C F liP F liL _ 2 ya jC F la A _ 3 0 ftsY le p B

se c G

se c A

se c Y -5 S R P 5 4

se c E

ta tA Figure 3. 11. Continued ta tC -1 0

114

Discussion

Bacillus pumilus B12 is a bacterial isolate capable of degrading poly-lactic acid (PLA) plastic films. Here we build upon our previous work showing changing nutrient conditions effect its rate of PLA degradation. Our results show that B. pumilus B12 grown on media augmented with complete supplement media (CSM) will lead to a reduction in extracellular proteins released and a change in gene expression. Of the proteins identified to be reduced in our proteomic work, two contain proteolytic domains that have potential promiscuity to break down the ester bonds holding together poly-lactic acid (PLA). Both of these genes are found ubiquitously in microbial life and are found in the cytoplasm suggesting that the for either to be the enzymatic target breaking down PLA, the cell would need to lyse. After performing live/dead staining on B. pumilus B12 growing in different nutrient conditions on PLA, we see no evidence of an increase in dead cells (Supplemental 3. 5.). Therefore, the hydrolytic enzyme acting upon PLA has yet to be elucidated.

When investigating how the cell is changing gene expression in conditions where we observe reduced PLA degradation, we see a wide encompassing transcriptional change, with more than 50% of the genes in the genome having significant differential expression (Figure 3. 5.). When breaking down the results into larger pathways, we see stark changes in the cells expression of motility, chemotaxis and protein export pathways. For motility and chemotaxis, in conditions rich in amino/nucleic acids, the cells may need to swim less in order to gather nutrients needed for replication. With free sources of amino acids as well as glucose, the need to expend energy and resources to scavenge more recalcitrant sources of energy is reduced. Therefore, wasting ATP for flagellar movement would be incongruent with bacterial success. As for protein export, reducing the expression of both the twin arginine translocation (TAT) and the general secretory pathway (SEC), would bring down the number of proteins being released into the environment. Considering, simple sugars and free amino acids are being provided to the bacteria, the reduction on enzyme release to break down large molecular weight sources of these molecules would seem appropriate. Therefore, our working hypothesis is that an increase in free nutrients to the bacteria is decrease in the need for bacterial movement and scavenging in order to survive. This lack of pressure to search for nutrients reduces the release of both enzymes, and the need for cell motility towards a more recalcitrant carbon source. A consequence of this abundant source of nutrient

115 options available, is that B. pumilus B12 reduces the degradation of the high molecular weight polymer, PLA.

Having identified possible mechanisms involved in the regulation of B. pumilus B12 PLA degradation, our next step will be to genetically modify B. pumilus B12 to see if our hypothesis is supported. We have so far been able to introduce genetic information in the form of a plasmid containing GFP into B. pumilus B12 and have plans to modify genetic targets identified in our RNAseq data (Supplemental Figure 3. 6.). We plan to disrupt the function of FlaA and other flagellar genes to see if the function or presence of the main motile element has an effect on B. pumilus PLA degradation. Further, we plan to alter the expression of the TAT protein export systems to see if overexpression rescues PLA degradation in MSM+G+CSM conditions. This work will be the next step in understanding the biological mechanisms of PLA degradation in B. pumilus B12 and help advance both the scientific understand and biotechnological applications of this novel soil isolate.

116

References

1 Fiori, S. M., & Zalba, S. M. (2003). Potential impacts of petroleum exploration and exploitation on biodiversity in a Patagonian Nature Reserve, Argentina. Biodiversity & Conservation, 12(6), 1261-1270. doi:10.1023/a:1023091922825

2 Haider, T. P., Völker, C., Kramm, J., Landfester, K., & Wurm, F. R. (2019). Plastics of the Future? The Impact of Biodegradable Polymers on the Environment and on Society. Angewandte Chemie International Edition, 58(1), 50-62. doi:10.1002/anie.201805766

3 Luckachan, G. E., & Pillai, C. K. S. (2011). Biodegradable Polymers- A Review on Recent Trends and Emerging Perspectives. Journal of Polymers and the Environment, 19(3), 637-676. doi:10.1007/s10924-011-0317-1

4 Amass, W., Amass, A., & Tighe, B. (1998). A review of biodegradable polymers: uses, current developments in the synthesis and characterization of biodegradable polyesters, blends of biodegradable polymers and recent advances in biodegradation studies. Polymer International, 47(2), 89-144. doi:10.1002/(sici)1097-0126(1998100)47:2<89::Aid-pi86>3.0.Co;2-f

5 Shah, A., Hasan, F., Hameed, A., & Ahmed, S. (2008). Biological Degradation of Plastics: A Comprehensive Review. Biotechnology advances, 26, 246-265. doi:10.1016/j.biotechadv.2007.12.005

6 Bandopadhyay, S., Martin-Closas, L., Pelacho, A. M., & DeBruyn, J. M. (2018). Biodegradable Plastic Mulch Films: Impacts on Soil Microbial Communities and Ecosystem Functions. Frontiers in microbiology, 9(819). doi:10.3389/fmicb.2018.00819

7 PlasticsEurope. (2018). Plastics – The Facts 2018: An analysis of European plastics production, demand and waste data.

8 Castro-Aguirre, E., Iniguez-Franco, F., Samsudin, H., Fang, X., & Auras, R.

(2016).Poly(lactic acid)-Mass production, processing, industrial applications, and end of life. Adv Drug Deliv Rev, 107, 333-366. doi:10.1016/j.addr.2016.03.010

117

9 Lunt, J. (1998). Large-scale production, properties and commercial applications of polylactic acid polymers. Polymer Degradation and Stability, 59(1), 145-152. doi:https://doi.org/10.1016/S0141-3910(97)00148-1

10 Gregor, A., Filová, E., Novák, M., Kronek, J., Chlup, H., Buzgo, M., . . . Hošek, J. (2017). Designing of PLA scaffolds for bone tissue replacement fabricated by ordinary commercial 3D printer. Journal of Biological Engineering, 11(1), 31. doi:10.1186/s13036-017-0074-3

11 Auras, R., Singh, S., & Singh, J. (2006). Performance Evaluation of PLA against Existing PET and PS Containers. Journal of Testing and Evaluation, 34(6), 530-536. doi:10.1520/JTE100041

12 Arrieta, M. P., López, J., Ferrándiz, S., & Peltzer, M. A. (2013). Characterization of PLA- limonene blends for food packaging applications. Polymer Testing, 32(4), 760-768. doi:https://doi.org/10.1016/j.polymertesting.2013.03.016

13 Tomita, K., Kuroki, Y., & Nagai, K. (1999). Isolation of thermophiles degrading poly(L- lactic acid). J Biosci Bioeng, 87(6), 752-755

14 Fukuzaki, H., Yoshida, M., Asano, M., Kumakura, M., Mashimo, T., Yuasa, H., . . . Yamanaka, H. (1991). In vivo characteristics of high, molecular weight copoly(l-lactide/glycolide) with S-type degradation pattern for application in drug delivery systems. Biomaterials, 12(4), 433- 437. doi:https://doi.org/10.1016/0142-9612(91)90014-2

15 Oda, Y., Yonetsu, A., Urakami, T., & Tonomura, K. (2000). Degradation of Polylactide by Commercial Proteases. Journal of Polymers and the Environment, 8(1), 29-32. doi:10.1023/a:1010120128048

16 Park, S., & Schumann, W. (2015). Optimization of the secretion pathway for heterologous proteins in Bacillus subtilis. Biotechnology and Bioprocess Engineering, 20(4), 623-633. doi:10.1007/s12257-014-0843-5

17 Masaki, K., Kamini, N. R., Ikeda, H., & Iefuji, H. (2005). Cutinase-like enzyme from the yeast Cryptococcus sp. strain S-2 hydrolyzes polylactic acid and other biodegradable plastics. Appl Environ Microbiol, 71(11), 7548-7550. doi:10.1128/aem.71.11.7548-7550.2005

18 Lim, H. A., Raku, T., & Tokiwa, Y. (2005). Hydrolysis of polyesters by serine proteases. Biotechnol Lett, 27(7), 459-464. doi:10.1007/s10529-005-2217-8 118 van Dijl, J., & Hecker, M. J. M. C. F. (2013). Bacillus subtilis: from soil bacterium to super- secreting cell factory. Microbial Cell Factories, 12(1), 3. doi:10.1186/1475-2859-12-3

Schallmey, M., Singh, A., & Ward, O. P. (2004). Developments in the use of Bacillus species for industrial production. Canadian Journal of Microbiology, 50(1), 1-17. doi:10.1139/w03-076

19 Vary, P. S., Biedendieck, R., Fuerch, T., Meinhardt, F., Rohde, M., Deckwer, W.-D., & Jahn, D. (2007). Bacillus megaterium—from simple soil bacterium to industrial protein production host. Applied Microbiology and Biotechnology, 76(5), 957-967. doi:10.1007/s00253-007-1089-3

20 Handtke, S., et al.. (2014). "Bacillus pumilus reveals a remarkably high resistance to hydrogen peroxide provoked oxidative stress." PLOS ONE 9(1): e85625.

21 Poorna, C. A., & Prema, P. (2006). Production and partial characterization of endoxylanase by Bacillus pumilus using agro industrial residues. Biochemical Engineering Journal, 32(2), 106- 112.

22 Battan, B., Sharma, J., Dhiman, S. S., & Kuhad, R. C. (2007). Enhanced production of cellulase-free thermostable xylanase by Bacillus pumilus ASH and its potential application in paper industry. Enzyme and Microbial technology, 41(6-7), 733-739.

23 Emadian, S. M., Onay, T. T., & Demirel, B. (2017). Biodegradation of bioplastics in natural environments. 59, 526-536. doi:10.1016/j.wasman.2016.10.006

24 Kawai, F. (2010). Polylactic Acid (PLA)-Degrading Microorganisms and PLA Depolymerases. In Green Polymer Chemistry: Biocatalysis and Biomaterials (Vol. 1043, pp. 405- 414): American Chemical Society.

25 Singhvi, M., & Gokhale, D. (2013). Biomass to biodegradable polymer (PLA). RSC Advances, 3(33), 13558-13568. doi:10.1039/C3RA41592A

26 Eckhard, U., Bandukwala, H., Mansfield, M. J., Marino, G., Cheng, J., Wallace, I., . . . Doxey, A. C. (2017). Discovery of a proteolytic flagellin family in diverse bacterial phyla that assembles enzymatically active flagella. Nature Communications, 8(1), 521. doi:10.1038/s41467- 017-00599-0

27 Hasan, F., Shah, A. A., & Hameed, A. (2006). Industrial applications of microbial lipases. Enzyme and Microbial technology, 39(2), 235-251.

119

28 López-Otín, C., & Bond, J. S. (2008). Proteases: multifunctional enzymes in life and disease. The Journal of biological chemistry, 283(45), 30433-30437. doi:10.1074/jbc.R800035200

29 Julsing, M. K., Rijpkema, M., Woerdenbag, H. J., Quax, W. J., & Kayser, O. (2007). Functional analysis of genes involved in the biosynthesis of isoprene in Bacillus subtilis. Applied Microbiology and Biotechnology, 75(6), 1377-1384. doi:10.1007/s00253-007-0953-5

30 Little, J. W. (1991). Mechanism of specific LexA cleavage: autodigestion and the role of RecA coprotease. Biochimie, 73(4), 411-421. doi:10.1016/0300-9084(91)90108-d

31 Little, J. W., Edmiston, S. H., Pacelli, L. Z., & Mount, D. W. (1980). Cleavage of the Escherichia coli lexA protein by the recA protease. Proceedings of the National Academy of Sciences of the United States of America, 77(6), 3225-3229. doi:10.1073/pnas.77.6.3225

32 Tessman, E. S., Tessman, I., Peterson, P. K., & Forestal, J. D. (1986). Roles of RecA protease and recombinase activities of Escherichia coli in spontaneous and UV-induced mutagenesis and in Weigle repair. J Bacteriol, 168(3), 1159-1164. doi:10.1128/jb.168.3.1159- 1164.1986.

33 Baker, T. A., & Sauer, R. T. (2012). ClpXP, an ATP-powered unfolding and protein- degradation machine. Biochim Biophys Acta, 1823(1), 15-28. doi:10.1016/j.bbamcr.2011.06.007

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Appendix

121

Figure S3. 1. Extracellular proteins collected from B. pumilus B12 in conditions showing high L-lactate release from PLA films show no distinct band differences from standard MSM+G control. Protein samples were collected from B. pumilus B12 48-hour cultures incubated either with (+) or without (-) 10 mg/mL PLA films. MSM+Glucose represents standard growth conditions where MSM+Maltose and MSM+Mannitol each represent conditions which increased PLA degradation was observed.

122

Figure S3. 2. Amino acid alignment of Flagellin proteins identified in B. Pumilus B12 show no homology to a previously identified proteolytic domain in C. haemolyticum Flagellin. Samples were aligned using ClustalOmega protein alignment tool with “*” representing a perfect amino acid match,” :” representing a strongly similar amino acid property match and “.” representing a weakly similar amino acid property match.

123

Supplemental Figure 3. 2. Continued 124

Figure S3. 3. Biotinylation of surface associated proteins of B. pumilus B12 in different growth conditions show no obvious differences. Cells were washed and biotinylated prior to protein extraction. Proteins were then blotted and run on an SDS-page gel and transferred to a nitrocellulose membrane. MSM+G is our stand growth media, with all 4 other growth supplementation conditions showing a decrease in PLA degradation in previous studies.

125

A. B.

Figure S3. 4. L-lactate metabolism/transport genes and mass spectrometry “hit” protein genes both show overall decrease in gene expression. A) Shows L-lactate associated gene expression based on colorimetric heat map. B) Shows colorimetric gene expression of proteins previously identified to have lower abundance in mass spectrometry. Values of expression can be seen on the bar right of the map.

126

Figure S3. 5. Live dead staining of B. pumilus B12 shows no large increase in dead cells in MSM+G conditions. A) Represents live cell staining of B. pumilus B12 growing on PLA in MSM+G. B) Represents dead cell staining of B. pumilus B12 growing on PLA in MSM+G. C) Represents live cell staining of B. pumilus B12 growing on PLA in MSM+G+CSM. D) Represents dead cell staining of B. pumilus B12 growing on PLA in MSM+G+CSM.

127

A. B.

Figure S3. 6. Bacillus pumilus B12 is able to take up plasmid DNA and express GFP. A) Fluorescent image of Bacillus pumilus B12 following electroporation transformation of TET-GFP plasmid DNA. B) Plasmid map of TET-GFP Vector.

128

Chapter IV: Novel Bacillus Strains Cultured from Contaminated Polysorbate-40 Show High Tolerance to Various Surfactants and can Utilize Polyethylene as a Sole Carbon Source

129

Contributions

Contributions of this Chapter are as follows: Phylogenetic Tree construction was performed by Eric Gann. Strain isolation, identification, protein isolation, blotting and all growth analysis has been performed by Kyle Bonifer. Analysis of polyethylene degradation by Bacillus strains has been performed by both Kyle Bonifer and Elise Phillips, apart from AFM being performed by Sahar Hasim.

130

Abstract

Surfactants are amphiphilic compounds commonly used in many industrial settings as detergents and to reduce surface tension. These molecules typically contain some type of fatty acid tail and a hydrophilic headgroup. One group of surfactant, polysorbates, contains both a fatty acid tail as well as a polyoxyethylene chain and used as emulsifiers in the pharmaceutical industry. In this study we identify several novel isolates of Bacillus species that were found contaminating a bottle of 100% polysorbate-40 (Tween-40). These strains were separately shown to use polysorbates, fatty acids, PEG and mineral oil (made up of short chain alkenes) as sole carbon sources. Next, because these strains showed the ability to utilize recalcitrant carbon chains as a carbon source, we investigated their ability to degrade polyethylene (PE) films. Over the course of 8 weeks, we observed microbial colonization, increased media acidity and decrease in film elasticity of PE films incubated with one of our Bacillus megaterium isolates, NB-10. This work is the first step in determining if novel Bacilli selected from a surfactant rich environment are viable targets for polyethylene degradation.

131

Introduction

Surfactants are compounds that reduce surface tension between two liquids. The basic structure of a surfactant consists of a charged head group and a hydrophobic tail, mimicking many of the properties of cellular lipids. These molecules will form micelles at a concentration known as the critical micelle concentration (CMC). Due to their amphiphilic nature, surfactants are used as detergents, emulsifiers and dispersants in industrial and laboratory settings [1,2,3,4]. Surfactants can be classified as cationic, anionic or nonionic depending on the charge of their headgroups [5].

One class of surfactants are the polysorbates. These molecules are a combination of a fatty acid tail and an ethoxylated sorbitan, which is a combination of ethylene glycol and sorbitol. This creates a non-ionic surfactant, with the numerical classification based on the number of fatty acid tail groups, with the polyoxyethylene chain being 20 monomers long. Polysorbates are currently being studied for their ability to aid in bioremediation of contaminated soil environments due to their relatively low toxicity, ability to form micelles around pollutants and their observed biodegradation in soil environments. [6,7,8,9,10]. However, surfactants are still capable of destabilizing membranes and linearizing proteins in some bacteria, temporarily inhibiting the growth and biofilm formation in microbial communities [11]. Furthermore, no studies have been performed investigating the effects of high concentrations, above 5% polysorbate [12], on microbial growth in the soil environments they are placed in.

When investigating polysorbate degradation, it has been observed that microbial communities preferentially target the fatty acid tail of the molecule, creating acetate from the long carbon chain [13]. Identifying microbes able to cleave this tail from the larger polysorbate molecule and using it as a sole carbon source could be helpful for studying the degradation of larger polymers. Since fatty acid tails have long hydrophobic chains of carbons, they are similar to polyethylene (PE) and polypropylene (PP) plastic polymers; two of the most widely used plastics in the world. Additionally, polyethylene glycol (PEG), is a long chain polymer that closely resembles an oxidized version of PE [14], containing a long chain of carbon-carbon bonds with interspersed oxygens more easily recognized and targeted by microbial enzymes [15]. Polysorbates containing both of these moieties in a single molecule make them an interesting candidate for enriching for microbial communities capable of degrading plastic polymers.

132

Here we investigate several strains of Bacillus that were observed growing in 100% polysorbate-40 (Tween-40). Subgroups of these Bacilli were shown to be capable of growing on several types of nonionic surfactants including Tween-20, Tween-80 and Triton-X. Furthermore, some of the strains break down polysorbate into its individual components, putatively showing preferential use of polyethylene glycol (PEG) for carbon. Finally, we investigated four isolates classified as Bacillus megaterium for their ability to grow on and utilize polyethylene films (PE) as a sole carbon source. Out of these four strains, one isolate, NB-10, was shown through SEM, AFM, pH measurements and live-dead staining to be growing upon and breaking down the polyethylene film. This work is a great first step for both looking into the community dynamics of a microbial population growing in a surfactant rich environment and for investigating plastic degrading microbial enrichment from these types of environments.

Methods

Initial Isolation, Microscopy and Phylogenetic Analysis

The suspected contaminated bottle of polysorbate-40 was sampled and spread on Lennox broth (LB) agar plates and incubated at 30℃ for 48 hours. Isolated colonies were then transferred to fresh LB agar for further processing. Samples were determined to be axenic through light microscopy and 16s rDNA sequencing. Genomic DNA was isolated through phenol-chloroform extraction. Universal 16S primers 27F and 1522R were used for PCR sequence amplification. Amplicons were Sanger sequenced at the University of Tennessee Genomics Core. Near full length 16S nucleotide sequences were downloaded from NCBI [16]. Multiple sequence alignments were performed in MEGA7 [17] using the MUSCLE algorithm [18]. Maximum likelihood phylogenetic reconstruction was performed in MEGA7 with 500 bootstrap replications to determine confidence values for each node.

Surfactant and Fatty Acid Growth Analysis

Samples that were determined to be monocultures were added to media containing either polysorbate-20, polysorbate-40, polysorbate-80, Triton-X or Sodium dodecyl sulfate (SDS). To check for the ability to grow in the presence of these surfactants, cultures were grown in LB media in increasing concentrations of each surfactant over the course of 48 hours. To determine if the

133 isolates were capable to utilize the surfactants as a sole carbon source, isolates were grown in liquid minimal salt media (MSM) containing 7.5 mM NH4SO4, 11 mM KHPO4, 2.87 mM

KH2PO4, 1.7 mM MgSO4 and 17.1 mM NaCl, pH 7.0 and the surfactants as the sole carbon source at different concentrations and grown for 48-120 hours. Cultures were grown into 200 µL volumes in a 96-well clear plates at approximately 5x106 cells and incubated at 30℃ in non-shaking conditions. Growth measurements were taken via optical density (OD600) every 10 minutes at 30℃ for the duration of the growth analysis in a Biotek Cytation 5 plate reader.

For Triton-X, the surfactant was added to ddH2O and serially diluted from a starting concentration of 20%. These dilutions were then added to 2x standard concentration on Lennox broth (LB) to create consistent nutrient conditions sans the surfactant. Growth curves were performed in 200 µL volumes in a 96-well plate with approximately 5x106 cells per well. Optical density (OD600) was measured every 10 minutes at 30℃ for the duration of the growth analysis in a Biotek Cytation 5 plate reader. For collected sub-populations of bacteria capable of growing in higher concentrations of Triton-X, cells were collected at the end of the 48 our growth analysis, reinoculated in fresh media containing 5% Triton X overnight and spread on LB agar.

For fatty acid growth, 25-100 µM concentrations were put in either liquid MSM or LB media to determine carbon utilization and inhibitory effect respectively. Cell concentrations and measurements are the same as the aforementioned conditions with the exception of oleic acid. For oleic acid, 5x106 cells were grown in 10mL culture flasks shaking at 225 RPM in a 30℃ incubator.

Growth was measured using optical density (OD600) over the course of 120 hours.

Poly-ethylene Glycol and Mineral Oil Growth Analysis

Both the utilization polyethylene glycol (PEG) and mineral oil were determined using agar plate assays with PEG utilization also being performed in liquid media. For solid PEG utilization, minimal salt media (MSM) agar plates were made containing 7.5 mM NH4SO4, 11 mM KHPO4,

2.87 mM K2HPO4, 1.7 mM MgSO4 and 17.1 mM NaCl, 10g of agar, pH 7.0 and 1% of PEG with a molecular weight of 6000. The same nutrient conditions were used for mineral oil, only replacing 1% PEG with 1% mineral oil. Growth was tracked over the course of 7 days at 30℃ for the appearance of colonies. Liquid cultures of PEG contained the same nutrient composition as solid growth assays sans agar. Liquid growth in 1% PEG was grown shaking at 225 RPM at 30℃ and optical density was measured every 24 hours for 96 hours.

134

Lipase/ Protease Activity

Determination of hydrolytic activity against lipid and peptide moieties was performed using tributyrin and casein agar plates, respectively. Casein plates were made using an amalgamation of 5 grams of dry skim milk and 5 grams of pure casein added to 2.5 grams of yeast extract, 1 gram of glucose and 10 grams of agar in a 500mL volume. Methylene blue was mixed into the molten agar at a concentration of 1% prior to pouring plates to increase contrast. Three microliters of 1.0 OD600 of bacterial culture were added to the center of the agar plate. Both colony size and halo formation were measured in triplicate for each bacterial isolate over the course of 7 days. Tributyrin agar plates were made with 2.5 grams of peptone and three grams of yeast extract to grams of agar in 500mL. Five mL of tributyrin and 1% of methylene blue was slowly mixed into the media as the contents were brought to a near boil before being autoclaved. Three microliters of 1.0 OD600 of bacterial culture was added to the center of the agar plate. Both colony size and halo formation were measured using a Cabela’s™ electronic caliper in triplicate for each bacterial isolate over the course of 7 days.

Biotinylation of Surface Proteins

Cell pellets were collected from 48-hour cultures via centrifuging at 10,000 RPM for 30 minutes. Cell were then normalized to a standard concentration of approximately 5*106 cells/mL and biotinylated using the EZ-Link™ Sulfo-NHS-LC-Biotinylation Kit. After biotinylation, samples were chemically and physically lysed using cell lysis buffer (1M NaCl, .5% SDS, 50mM Tris 7.4 pH) and 150-212 µm sand grains. Cell/lysis buffer mixtures were shaken using a Biospec bead beater three times for 30 seconds. Next, protein concentrations were normalized using Bradford assay, mixed with Laemmli buffer and boiled for 5 minutes. Following protein isolation, normalization and biotinylation, samples were run through a 12% SDS-page gel and transferred to nitrocellulose blot paper overnight. Blots were then incubated with a conjugated avidin-GFP which detects and tightly binds to the pre-bound biotin. Blots were visualized using a LiCor Odyssey CLx using the green fluorescent emission channel.

Polyethylene Plastic Film Degradation Analysis

Determination of polyethylene degradation was performed using polyethylene bags cut into 5x5mm squares incubated in 1mL of MSM liquid media. Isolates were inoculated at approximately 5x106 cells/mL and incubated at 30℃ for the duration of the experiment.

135

Timepoints were taken every two weeks over the course of ten weeks where adhesion, viability, pH and plastic integrity measurements were taken. Cell adhesion was measured using scanning electron microscopy (SEM). Films were then washed three times with phosphate buffered saline (PBS) and fixed with 2.5% glutaraldehyde for one hour. Samples were then gradually dehydrated with ethanol anhydrous (25%, 50% and 100% respectively) and air dried in a fume hood overnight. Samples were mounted on aluminum stubs, sputter coated with iridium and observed using a 1 keV Zeiss GeminiSEM scanning electron microscope. Cell viability was determined by staining the PE films in a Baclight® live/dead cell viability kit and observed with fluorescent microscopy. For plastic integrity, PE film elasticity was measured using atomic force microscopy (AFM). Samples were washed with phosphate buffer saline 5 times to remove excess biomass. Each sample was then imaged using a 5500 PicoPlus atomic force microscope (AFM) operating with the 1.20.2 operating system (Keysight Technologies Inc. Santa Rosa CA). The instrument was operated in contact mode using MLCT probes with spring constants of 0.01 and 0.03 N/m and a tip radius of 10 nm and Poisson ratio of 0.5. For both imaging and elasticity measurements, the applied force was kept at 3-5 nN. The force distance curves were measured with 8x8 points, with each point being an average of 3 force curves. These measurements were recorded on 2-μm2 areas on the top of plastic for an average of 5 different locations. This data was then converted to elasticity measurements using the Keysight software in PicoPlus 5500 AFM [19]. Statistical significance was determined via ANOVA with p-values below .01 considered significant. pH was measured using a Denver UltraBasic pH meter and compared to media incubated with PE without bacterial inoculum.

Results

Isolation of Novel Bacillus Isolates

We initially observed an increased opacity forming on the upper fraction of a bottle of unopened Polysorbate-40 (Tween-40) used for proteomic work in our laboratory. Over the course of 6 months, the opacity grew to encompass most of the Tween-40 within the bottle (Figure 4. 1.). Next, we took 1mL of the presumed contaminated Tween and diluted it 1:10 in fresh media and then performed a spread plate on LB agar to produce isolated colonies . Upon isolating the colonies, we performed sequencing and phylogenetic analysis on 16 isolates; deemed “novel

136 bacteria (NB)”, collected from the LB agar plate. Of the 16 colonies, 6 were determined to be single isolates, all contained within the Bacillus genus (Figure 4. 2.). Within the Bacillus genus, 4 were most closely to Bacillus megaterium (NB6, NB10, NB12 and NB14) with one isolate aligning to Bacillus aerius (NB5) and one aligning with Bacillus amyloliquefaciens (NB9) (Figure 4. 2.).

Novel Bacillus Isolates Preferentially Grow on Tween-40 and Tween-80 Over Other Surfactants

Next, we wanted to see if the bacteria now isolated away from their community could grow in high concentrations of polysorbates. Tween is considered a mild detergent with low toxicity towards most cellular life tested. However, the highest concentrations reported in these analysis’ has been 5000 mg/L (5%) [12] Furthermore, these high concentrations were not the sole carbon source allotted to the microbe. With this in mind, we measured our NB strains’ growth over the course of 48 hours in Tween-20, Tween-40 and Tween-80. The polyoxyethylene groups on these three molecules are all consistent with the main difference occurring in the fatty acid tail. For Tween- 20, Tween-40 and Tween-80, the fatty acid tails are lauric acid, palmitic acid and oleic acid, respectively. What we observed over the course of 48 hours was preferential growth on Tween-40 at a concentration of 25% compared to the other two surfactants (Figure 4. 3.). To see marginal levels of growth on Tween-80 for any isolates, measurements had to be extended for 120 hours (Supplemental Figure 4. 1.). These results indicate that our NB strains are capable of growing on surfactants as a sole carbon source, and that they preferentially grow on the same surfactant from which they were isolated.

Novel Bacillus Isolates Grow on Fatty Acids, Polyethylene Glycol, and Mineral Oils as a Sole Carbon Source More Efficiently than Others

Next, to see if the growth on Tween was specific to a subsection of the molecules, we tested the growth of our NB strains on both polyethylene glycol (PEG) and different fatty acids. For PEG, we observed growth for all isolates with the exception on one, NB9, on minimal media plates with PEG-6,000 as a sole carbon source (Figure 4. 4A.). Due to their being lipids in agar plates we also grew our samples in liquid media. The plate data was supported by liquid growth analysis’ over the same time period showing small but increased opacity for all NB strains with the exception of NB9 (Figure 4. 4B.). Next we tested the growth of the NB isolates on oleic acid (OA) and linoleic

137

1 A. B.

Figure 4. 1. Contamination of Polysorbate-40 (Tween-40) bottle. A) Shows the original source of contamination following over one year of growth. B) Shows the contaminated Tween-40 (right) compared to non-contaminated Tween-40 (left).

138

Figure 4. 2. Maximum likelihood phylogenetic tree of 16S rDNA for the environmental NB isolates compared to other Bacillus spp. This bootstrap consensus tree was constructed with 500 replicates. Bootstrap values were used as confidence values for the nodes. Nucleotide sequences for the other bacteria were retrieved from NCBI.

139

A. 25% Tween-40 B. 25% Tween-20

C. 25% Tween-80

Figure 4. 3. Growth analysis reveals a preference of NB strains to grow on Tween-40 over Tween-80 and Tween-20. A) Optical density growth curve for NB strains grown in 25% Tween-40 over the course of 48 hours. B) Optical density growth curve for NB strains grown in 25% Tween-20 over the course of 48 hours. C) Optical density growth curve for NB strains grown in 25% Tween-80 over the course of 48 hours.

140 acid (LA). Due to evidence suggesting several Bacillus spp. are inhibited by linoleic acid [20] we tested both the strain’s ability to grow in the presence of LA and their capability of using it as a sole carbon source. For LA, we observed no inhibition of growth for any of the isolates with the exception of NB9 (Figure 4. 4A.). When 100 µM of LA was the sole carbon source given to the isolates, we only saw a slight increase in growth for NB6 over the course of 24 hours (Figure 4. 5B.). Finally, for OA, we noticed a small but steady growth for all strains tested over the course of 120 hours (Figure 4. 5C.). These results point to the possibility that the NB isolates use fatty acids as a sole carbon source, however further work is required determine concentrations and timescales that are most amenable for growth.

Next, we investigated the Bacillus spp.’ ability to withstand high concentrations of other more toxic surfactants. The two other surfactants tested were Triton-X and sodium dodecyl sulfate (SDS), both of which are considered to have stronger antimicrobial properties [21,22]. For SDS, we saw no growth for any of our isolates at concentrations ranging from .79% to 5% in LB (Supplemental Figure 4. 2.). As for Triton-X, growth was observed in a similar pattern for all isolates (Supplemental Figure 4. 3A.) with the exception of NB9 which showed a significantly longer lag period (Supplemental Figure 4. 3B., 4. 3C.). All strains showed the ability to grow in the presence of Triton-X at concentrations up to 20%, however a significant decrease in growth rate was observed for all isolates (Supplemental Figure 4. 3A, 4. 3B.). To investigate the long lag period associated with NB9, we sub-cultured NB9 grown in 20% Triton-X and reinoculated into fresh LB with 20% Triton-X. Interestingly, we saw a disappearance of the long lag period previously observed, with the growth rate resembling that of the other NB isolates (Supplemental Figure 4. 3D.). This data suggests the existence of subpopulations or suppressors of NB9 cells capable of withstanding high concentrations of Triton-X.

Finally, we investigated the isolates’ abilities to use mineral oil as a sole carbon source. This was performed by using 1% mineral oil which is rich in short chain hydrocarbons and aromatic rings [23]. All of our strains showed colony formation over the course of 7 days on these plates, with more rapid colony formation occurring for two of the B. megaterium isolates (NB 10, 14) (Supplemental Figure 4. 4A., 4. 4B.). These results suggest the Bacillus spp. are capable of utilizing the mineral oil as a sole carbon source.

141

Figure 4. 4. NB isolates are capable of growing on polyethylene glycol in solid and liquid media. A) Tracking the formation of colonies for every 24 hours for 7 days on MSM+1% PEG6000 agar plates. Green indicates visible colony formation while red indicates no visible colony formation. B) Growth in MSM+1% PEG6000 liquid cultures. Optical density was taken every 24 hours for 96 hours in triplicate.

142

A B. .

Figure 4. 5. Growth of NB strains in different fatty acids. A) Optical density measurements of NB strains in LB media with a 100µM supplementation of linoleic acid (LA) measured over 48 hours. B) Optical density measurements of NB strains grown in MSM with a 100µM supplementation of LA measured over 20 hours. C) Optical density measurements of NB strains grown in MSM with 100µM supplementation of oleic acid measured over 120 hours.

143

Novel Bacillus Isolates Have Inverted Lipase and Protease Activity from One Another

To determine if strong hydrolytic activity is correlated with our novel isolates’ ability to grow in surfactants, we measured halo formation on protease and lipase plates. The protease plates contain casein, a large milk protein, whilst the lipase plates are rich in tributyrin, a triglyceride. In order to access these nutrient sources, our strains would need to break down the larger protein and lipid complexes outside of the cell by releasing hydrolyzing enzymes. We observed an inverse relationship between the halo formation and growth on either plate between one strain (NB9) and the other isolates (Figure 4. 6.). On casein rich media, NB9 showed a far greater halo size as a ratio of colony size compared to the other NB strains (Figure 4. 6A.). This observation is the inverse for the tributyrin-rich plates, where NB9 halo formation lagged behind all other NB strains (Figure 4. 6B.).

Bacillus megaterium NB-10 Grows on High Density Polyethylene Films as a Sole Carbon Source

Since our NB isolates are able to grow on recalcitrant carbon sources including short chain carbons as fatty acids, we wanted to determine if they could process and breakdown polyethylene (PE) films. Polyethylene comprises of long, densely packed chains of carbon-carbon bonds that are resistant to both chemical and biological erosion [14]. Our results show the most rapid growth on mineral oil by the two our B. megaterium strains (NB10 and NB14), we decided to analyze NB6, NB10, NB12 and NB14 strains initially for PE adhesion and breakdown. We set up 1mL cultures in minimal salt media (MSM) with PE as the sole carbon source and allowed the cells to grow at 30℃ in static conditions. We then took measurements for each strain every 2 weeks for 8 weeks total in order to see if there were any indicators of microbial growth and film breakdown. To see if cells were continuously viable throughout the experiment, we performed live/dead staining on the PE films at each timepoint. We observed no drastic increase in dead cells over the course of the experiment while seeing a small but steady increase in viable cells over each measurement taken for each of the 4 strains (Figure 4. 7.).

To determine if there is metabolic activity occurring in the media with the release of lactate, acetate or other acid compounds, we took pH measurements for each sample over the course of 8 weeks. For all four stains we observed a decrease between the initial pH reading of roughly 6.6 and the subsequent pH readings (Figure 4. 8.). Within 2 weeks, the pH had dropped below 6.4 and

144 by week four pH readings were between 6.2 and 6.3 (Figure 4. 8.). Readings for the following two weeks stayed consistent with what we observed after four weeks (Figure 4. 8.). These results coupled with our live dead staining indicate that these B. megaterium isolates maintain cell viability and are dropping the pH of the media in the presence of PE as a sole carbon source.

Next we wanted to use the power of scanning electron microscopy (SEM) to better observe colony formation on the plastics over the course of 8 weeks. We were able to observe single cells growing upon the PE film from our earliest time point (2 weeks), out to 8 weeks (Figure 4. 9.). For NB6 and NB10 we saw a qualitative increase in the number of cells adhering to the plastic, while NB12 and NB14 growth stayed more static throughout the time course (Figure 4. 9.). Due to the lack of another carbon source added to the media, the increase in cellular abundance is an indication that the NB strains are using PE as a carbon source for growth.

Finally, to see if the microbial growth we were observing was having an impact on plastic integrity, we performed atomic force microscopy (AFM) on the PE films to measure the change in plastic elasticity over time. The level of plastic elasticity is positively correlated with degradation. The lower the Young’s modulus, the higher the overall level of elasticity (and therefore the lower plastic rigidity) of the plastic film. Out of the 4 strains measured, only one, NB10, showed a significant decrease in plastic rigidity over the course of 6 weeks based on Young’s modulus measurements taken in megapascals (MPa) (Figure 4. 10., Supplemental Figure 4. 5.). This data along with live/dead, pH and SEM analysis lend preliminary evidence that the B. megaterium strain, NB10, is both growing on the PE film, and that growth on the film is reducing the rigidity of the plastic.

Discussion

Surfactants are a recalcitrant carbon sources that are widely used in industrial settings, Moreover, they are being studied for their ability to aid in bioremediation (1,6,7,8,9,10,). Due to the increased attention they are garnering, it is important to identify and characterize microbial life capable of persisting in surfactant rich environments and utilizing the amphiphiles as carbon sources. Furthermore, the hydrocarbon rich moieties located in both the hydrophobic and

145

A. B.

Figure 4. 6. A Ratio of colony size to lipase and protease halo formation for NB strains over 7 days. A) Protease activity measured by halo formation on casein agar plates. Each data point represents three replicates with all measurements taken in millimeters. B) Lipase activity measured by halo formation on tributyrin agar plates. Each data point represents three replicates with all measurements taken in millimeters.

146

Figure 4. 7. Live/Dead staining of NB strains grown on PE films show no large decrease in cell viability of the course of 8 weeks. NB10 strains shown above with the top row consisting of dead cells stained with propidium iodide and with live cells shown below stained with Syto 9 dye.

147

6 .6 N B 6 N B 1 0 N B 1 2 N B 1 4

6 .4

H p

6 .2

6 .0 0 2 4 6 8 1 0 T im e (W e e k s)

Figure 4. 8. B. megaterium NB isolates lower the pH of the media when incubated with PE over the course of 8 weeks. Samples were collected in triplicate every 2 weeks for 8 weeks. The pH of the media prior to inoculation is 6.6.

148

Figure 4. 9. Scanning electron microscopy images of polyethylene inoculated with NB isolates over the course of 8 weeks. Images are were taken every two weeks for all 4 isolates tested. Images are between 2000-5000x magnification. NB10’s 2-week sample was not properly dehydrated and did not yield quality images.

149

Figure 4. 9. Continued

150

1 0

)

P M

( *

u 6

u d

o ***

' 4

o Y 2

0 k k k e e e e e e -w -w -w 2 4 6 0 0 0 1 1 1 B B B N N N

Figure 4. 10. Atomic Force microscopy shows a general reduction in tensile strength for

polyethylene incubated with NB10 isolate. Results shown are the average cantilever measurements of the PE film indicating tensile strength as Young’s modulus in megapascals (MPa). *** and * indicate ANOVA p-value = ≤ .001 and .01 respectively.

Here we show 6 novel isolates of the genus Bacillus that are both capable of growing on the non-ionic surfactant Polysorbate-40 (Tween-40). These organisms preferentially grow on with

151 hydrophilic sections could potentially be used as markers for organisms capable of processing petroleum derived polymers. Tween-40 differ from other Tween molecules, with only changes the type of fatty acid tail is esterified to the polyoxyethylene headgroup. Initial growth analyses on either fatty acids alone or polyethylene glycol (PEG) show most strains can grow on PEG and oleic acid as sole carbon sources. The isolates are also capable of growing on high concentrations of the surfactant Triton-X, while growth in sodium dodecyl sulfate (SDS) is minimal. The main structure in common between Tween and Triton-X, but dissimilar in SDS, is a polyoxyethylene chain. This suggests that our organisms are utilizing this subsection of the surfactant as a carbon source. When investigating the ability to grow on short chain hydrocarbons as a sole carbon source, we saw all of our isolates were able to grow within 5 days on MSM+1% mineral oil plates with some strains able to form colonies within 24 hours. This growth was only able to occur in the presence of oxygen, as isolates grown on mineral oil in an anaerobic chamber showed no growth with 14 days. This data coupled with the isolates ability to grow on PEG suggests it requires the oxidation of the hydrocarbon chain to utilize the carbon. Due to the short chain hydrocarbons and PEG both representing different stages of polyethylene (PE) degradation (chain breakage and oxidation respectively), we cultured our isolates with PE films, seeing growth and markers of film breakdown for one of our isolates, NB10.

The culmination of this data shows the possibility of enriching organisms in polysorbate or PEG alone may aid in finding novel organisms capable of degrading polyethylene and other petro-polymers. More work needs to be done to fully elucidate and characterize the growth dynamics of these microbes. First, a larger panel of fatty acids must be screened for the NB isolates ability to use them as a sole carbon source. Furthermore, different average chain lengths of PEG must be used to see at what point our isolates are no longer able to utilize PEG as a carbon source. PEG becomes more recalcitrant as the polymer chain as it becomes longer [24] and determining if and when there is a threshold can aid in determining targets for PE degradation. For PE degradation analysis, a longer trial testing all 6 strains must be performed to see if the trend observed over the course of 2 months can be extrapolated to an extended period of time. Measurements such as FTIR and CO2 should be utilized to see if there is a change in functional groups during degradation and to measure microbial respiration, respectively.

Due to some of the differences observed in growth on fatty acids and PEG between the different isolates, community analyses should be performed to determine if there are mutualistic 152 or dependent dynamics between the different bacterial strains. Finally, we need to identify the enzymes and pathways at work to break down and utilize the surfactants to better propose and answer biological questions pertaining to these isolates’ physiologies. To accomplish this, we need use the power of -omics as well as develop a genetic system in these strains.

153

References

1. Cheng, M., Zeng, G., Huang, D., Yang, C., Lai, C., Zhang, C., & Liu, Y. (2018). Tween

80 surfactant enhanced bioremediation: toward a solution to the soil contamination by hydrophobic organic compounds. Critical Reviews in Biotechnology, 38(1), 17-30. doi:10.1080/07388551.2017.1311296

2. Farn, R. J. (2008). Chemistry and technology of surfactants: John Wiley & Sons.

3. Kralova, I., & Sjöblom, J. (2009). Surfactants used in food industry: a review. Journal of

Dispersion Science and Technology, 30(9), 1363-1383.

4. Birch, B., & Cockcroft, R. (1981). Analysis of ionic surfactants in the detergent industry using ion-selective electrodes. Ion Sel. Electrode Rev, 4, 1-41.

5. Dave, N., & Joshi, T. (2017). A Concise Review on Surfactants and Its Significance. International Jurnal of Applied Chemistry, 13(3), 663672.

6. Bautista, L. F., Sanz, R., Molina, M. C., González, N., & Sánchez, D. (2009). Effect of different non-ionic surfactants on the biodegradation of PAHs by diverse aerobic bacteria. International Biodeterioration & Biodegradation, 63(7), 913-922.

7. Paria, S. (2008). Surfactant-enhanced remediation of organic contaminated soil and water. Advances in colloid and interface science, 138(1), 24-58.

8. Trellu, C., Mousset, E., Pechaud, Y., Huguenot, D., van Hullebusch, E. D., Esposito, G.,

& Oturan, M. A. (2016). Removal of hydrophobic organic pollutants from soil washing/flushing solutions: A critical review. J Hazard Mater, 306, 149-174. doi:10.1016/j.jhazmat.2015.12.008

9. Khalladi, R., Benhabiles, O., Bentahar, F., & Moulai-Mostefa, N. (2009). Surfactant remediation of diesel fuel polluted soil. Journal of Hazardous Materials, 164(2-3), 1179-1184.

10. Mulligan, C., Yong, R., & Gibbs, B. (2001). Surfactant-enhanced remediation of contaminated soil: a review. Engineering geology, 60(1-4), 371-380.

11. de la Cueva, S. C., Rodríguez, C. H., Cruz, N. O. S., Contreras, J. A. R., & Miranda, J. L.

154

(2016). Changes in bacterial populations during bioremediation of soil contaminated with petroleum hydrocarbons. Water, Air, & Soil Pollution, 227(3), 91.

12. Rodríguez-Escales, P., Borràs, E., Sarra, M., & Folch, A. (2013). Granulometry and surfactants, key factors in desorption and biodegradation (T. Versicolor) of PAHs in soil and groundwater. Water, Air, & Soil Pollution, 224(2), 1422.

13. Yeh, D. H., & Pavlostathis, S. G. (2005). Anaerobic biodegradability of Tween surfactants used as a carbon source for the microbial reductive dechlorination of hexachlorobenzene. Water Science and Technology, 52(1-2), 343-349. doi:10.2166/wst.2005.0537

14. Arutchelvi, J.M. S., Ambika Arkakar, Mukesh Doble, Sumt Bhaduti, Parasu Veer

Uppara. (2008). Biodegradation of polyethylene and polypropylene. Indian Jounral of Biotechnology, 7, 9-22.

15. Shah, A., Hasan, F., Hameed, A., & Ahmed, S. (2008). Biological Degradation of

Plastics: A Comprehensive Review. Biotechnology advances, 26, 246-265. doi:10.1016/j.biotechadv.2007.12.005

16. Benson, D., Karsch-Mizrachi, I., Lipman, D., Ostell, J., & Wheeler, D. (2008). GenBank

Nucleic Acids Res.jan, 1, 33.

17. Kumar, S., Stecher, G., & Tamura, K. (2016). MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Molecular biology and evolution, 33(7), 1870-1874.

18. Edgar, R.C., 2004. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797

19 Hasim, S., Allison, D.P., Retterer, S. T., Hopke, A., Wheeler, R.T., Doktycz, M.J., & Reynolds, T.B. (2017) beta-(1,3)-Glucan Unmasking in Some Candida albicans Mutants Correlates with Increases in Cell Wall Surface Roughness and Decreases in Cell Wall Elasticity. Infect Immum, 85(1)

20. Lee, J.-Y., Kim, Y.-S., & Shin, D.-H. (2002). Antimicrobial Synergistic Effect of

155

Linolenic Acid and Monoglyceride against Bacillus cereus and Staphylococcus aureus. Journal of Agricultural and Food Chemistry, 50(7), 2193-2199. doi:10.1021/jf011175a

21. Diaz De Rienzo, M. A., Stevenson, P., Marchant, R., & Banat, I. M. (2016). Antibacterial properties of biosurfactants against selected Gram-positive and -negative bacteria. FEMS Microbiol Lett, 363(2), fnv224. doi:10.1093/femsle/fnv224

22. Cornett, J. B., & Shockman, G. D. (1978). Cellular lysis of Streptococcus faecalis induced with triton X-100. J Bacteriol, 135(1), 153-160. Retrieved from https://www.ncbi.nlm.nih.gov/pubmed/97265

23. Marinescu, I. D., Rowe, W. B., Dimitrov, B., & Inasaki, I. (2004). 14 - Process Fluids for

Abrasive Machining. In I. D. Marinescu, W. B. Rowe, B. Dimitrov, & I. Inasaki (Eds.), Tribology of Abrasive Machining Processes (pp. 531-585). Norwich, NY: William Andrew Publishing.

24. Kawai, F. (2005). Biodegradation of Polyethers (Polyethylene Glycol, Polypropylene

Glycol, Polytetramethylene glycol, and Others). In Biopolymers Online.

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Appendix

157

1 6 8 N B 5 N B 6 N B 9 N B 1 0 0 .6 N B 1 2 N B 1 4

) B 1 2

(

y t

i 0 .4

p O 0 .2

0 .0 0 2 0 0 0 4 0 0 0 6 0 0 0 8 0 0 0 T im e (m in )

Figure S4. 1. Growth on NB strains in 25% Tween-80 over 120 hours. Optical density was measured every 10 minutes for 120 hours.

158

Figure S4. 2. Example of growth inhibition of NB strains by sodium dodecyl sulfate (SDS). Growth curve shows NB5 grown for 20 hours in different concentrations of SDS. Curves are representative of other strains grown in the same concentrations if SDS.

159

Figure S4. 3. Analysis of NB growth on the surfactant Triton-X. A) Isolate NB5 grown in rich media (LB) with serial dilutions of Triton-X over the course of 24 hours. Growth curves are representative of all other isolates with the exception of NB9. B) NB9 grown in LB with serial dilutions of Triton-X for 24 hours. C) The first 500 minutes of NB9 growth in rich media with serially diluted concentrations of Triton X. D) NB9 and NB9 reinoculated from Triton-X culture in a 24-hour growth comparison in LB with 20% Triton-X.

160

A. B.

C. D.

Figure S4. 3. Continued

161

A. B.

Figure S4. 4. NB isolates can grow on mineral oil as a sole carbon source. A) Growth on MSM+ 1% mineral oil plate after 7 days grown at 30℃. B) Time indicators of colony formation for each isolate taken each day for 7 days. Green represents visible colony formation while red indicates no visible colony formation.

162

1 0

)

(

u 6

s '

g 4

0 k k k k k k k k k e e e e e e e e e e e e e e e e e e -w -w -w -w -w -w -w -w -w 2 4 6 2 4 6 2 4 6 6 6 6 2 2 2 4 4 4 B B B 1 1 1 1 1 1 N N N B B B B B B N N N N N N

Figure S4. 5. Atomic force microscopy data for NB6, NB12 and NB14 strains grown on polyethylene films for 6 weeks. Data shows the average cantilever measurements represented by Young’s modulus in megapascals (MPa). NB6 is represented in Blue, NB12 is represented in orange and NB14 is represented in green.

163

Figure S4. 6. Biotinylated extracellular proteins collected from NB9 grown in both MSM+G and Tween. Proteins were extracted via acetone precipitation after 48 hours of growth in each condition. The two left bands are duplicates of MSM+G growth conditions and the two right bands are duplicates of 25% Tween conditions.

164

Chapter V: Conclusion and Future Directions

165

Conclusion

This dissertation aimed to A) develop new tools that can aid in identifying plastic degrading microorganisms and enzymes B) investigate the physiological mechanisms regulating plastic degradation and C) discover new environments that could enrich for plastic degrading organisms. We have showed here a rapid and accurate assay for the detection of poly-lactic acid breakdown by measuring L-lactate monomers. This assay was capable of detecting degradation in a timescale between minutes and hours compared to other measuring systems for plastic degradation that can take days, weeks or months. This rapid assay allowed us to quickly identify a strain of Bacillus pumilus (B12) that was able to break down PLA films. Furthermore, the efficacy of our assay was verified through other standard measurements and observations including atomic force microscopy (AFM) and scanning electron microscopy (SEM).

The development of our assay allowed us to subject our plastic degrading organism to a battery of environmental conditions to identify settings that would either increase or decrease the level of plastic degradation. We found that in conditions with mannitol and maltose sugar supplementation in place of glucose, we saw an overall increase in PLA degradation. Conversely, conditions rich in adenine, potassium and phosphate reduced B. pumilus B12’s ability to degrade PLA. Having observed these different conditions that could alter plastic degradation allowed us to examine what is changing on a transcriptomic and proteomic level in our organism.

In Chapter III, we investigated what changes were occurring in adenine rich conditions that was changing PLA degradation in B. pumilus B12. We observed a change in the abundance of extracellular proteins using silver staining, with mass spectrometry also showing a general reduction in identified peptides. Although there was no detectable difference in the proteins expressed on the cell surface, RNA sequencing revealed that more than half of the genome was differentially expressed in conditions with adenine supplementation. To help simplify the scope of the data, genes were put into pathways to see if major metabolic systems were altered in reduced PLA degradation conditions. We observed a majority of genes in motility, chemotaxis and protein export systems to be downregulated. Based on this data, we propose a hypothesis that nutrient rich conditions are reducing the need for scavenging and enzyme export.

Finally, in Chapter IV we investigated a contaminated bottle of polysorbate-40 (Tween- 40), to characterize organisms that could survive in a surfactant rich environment. We isolated 6

166 novel Bacillus spp. growing in the amphiphilic environment. Growth analysis showed that the organisms all preferentially grew in Tween-40, with growth also being detected in Tween-80 and to a lesser extent Tween-20. When breaking down the surfactant into its individual parts of polyethylene glycol (PEG) and fatty acids, we observed most species capable of using both as a sole carbon source. Furthermore, all strains were able to use mineral oil, which consists of short chain carbons, as their sole carbon source. Due to PEG, fatty acids and mineral oil mimicking polyethylene (PE) at different stages of degradation, we decided to investigate our Bacillus spp. ability to grow on and break down PE films. Out of the 4 strains we tested, one Bacillus megaterium isolate, NB10, showed the ability to grow, reduce the pH of the medium and reduce the elasticity of the PE film. This work was the first step into investigating the level of degradation capable by these Bacillus spp. and what the underlying mechanisms are that allow for both their survival in surfactant rich environments and polymer degradation.

Future Directions

PLA Project

In order to fully elucidate the genes controlling the breakdown of PLA by Bacillus pumilus B12, we need to be able to genetically modify the organism. So far we have shown that B. pumilus B12 is amenable to the uptake of genetic material in the form of plasmid DNA, however the alteration of chromosomal DNA will be necessary to find both the regulatory and enzymatic targets responsible for PLA degradation. Once the cloning protocol has been established, we can test the putative genes identified through RNA sequencing. A possible alternative would be to develop a mutant screen by creating a transposon mutant library.

The most interesting targets identified through RNA sequencing involve flagellar production/motility, chemotaxis and protein export systems. For the flagella, we need to know if it is the production of the flagella itself, the functionality of the structure or both that is affecting PLA degradation. For functionality, we plan on targeting the MotA gene which is essential for motility in Bacillus [1]. With a gene disruption successfully completed, a reduction in L-lactate release from PLA would be indicative of motor function rather the flagellar presence that is changing PLA degradation. The would show that either a structure on the surface of the flagella

167 needs to be in motion to hydrolyze the plastic or that moving and colonizing are key features in the degradation process. Alternatively, disrupting the production of flagellin (FlaA) or the attachment site by affecting the hook-associated proteins (FlgK, FlgL, FlgE , etc.) would remove the flagella structure altogether. Knocking out any of these components while still having functional MotA, would be evidence that the structure itself is having an effect on the breakdown of PLA.

There were two protein export systems that show a general downregulation in conditions with reduced PLA degradation. These two systems are the general secretion route (Sec) and the twin arginine route (Tat). Both are involved in protein secretion with the Sec releasing a larger number of total proteins in an unfolded confirmation and the Tat system releasing a smaller number of fully folded proteins [2]. Due to the SEC pathway being necessary for growth, disrupting the function of each of the TAT pathway individually could help more accurately pinpoint the hydrolytic enzyme being released. If the TAT pathway shows the desired effect of reduced PLA degradation then the signal sequence located upstream of the N-terminal region of the protein can be used to identify proteins that are secreted by this transmembrane complex [3].

With the amount of co-polymer production and multi origin plastics ending up in the same environment, finding multi-faceted organisms and enzymes capable of degrading several plastics becomes important. With the observation of B. pumilus B12 based PLA degradation now established, we plan on investigating the organism’s ability to degrade other biodegradable plastics, plastic blends and non-biodegradable plastics. Interesting targets to begin with are polybutylene succinate (PBS) and polybutylene adipate terephthalate (PBAT). Both of these plastics are considered biodegradable, with PBS releasing succinate as one monomeric byproduct and PBAT releasing 1,4 butanediol [4,5]. Succinate can be detected via luminescent detection, and 1,4 butanediol has been shown to convert into succinic acid (succinate) through alcohol and aldehyde dehydrogenases in liver metabolism [6,7]. This raises the possibility of detecting degradation byproducts of both types of plastic polymer in a single rapid assay. So far we lack an enzyme to be used as a positive control for assay development.

Finally, with the preliminary evidence of other Bacillus pumilus strains having altered amounts of PLA degradation raises the question: what is different between the Bacillus strains? So far we have tested a handful of other isolates, with plans to test several other Bacillus isolates.

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Once this has been completed, we plan on taking a bioinformatic approach do whole genome comparisons of the different strains. If no distinct differences can be found, then utilizing protocols developed for proteomic and transcriptomic analysis on B. pumilus B12 will be applied. This will give insight on both the genetic differences between strains possibly allowing for quicker identification of hydrolases. Also, it could give a better understanding on the different way strains regulate their hydrolytic enzyme release, further elucidating differences in strain specific gene expression.

Surfactant Degradation Project

For our Bacillus isolates shown to use polysorbate-40 (Tween-40) as a sole carbon source there are multiple experiments that need to be performed to characterize the organisms. First, a larger panel of fatty acids at a wider number of concentrations and for a longer period of time need to be used to see if the NB strains can utilize them as a sole carbon source. Moreover, all isolates need to be investigated for polyethylene degradation with a longer time course set-up to better see temporal changes to the polymer. Also, conditions need to be elucidated that can change the isolates’ abilities to survive in tween to help clarify the mechanisms being used to survive in the surfactant. Beyond these experiments, whole genome sequencing must be performed to better classify and compare these microbes to other Bacillus spp.. Finally, being able to genetically modify these non-model organisms and introduce foreign DNA will help distinguish single isolates from one another. Developing these techniques can allow us to investigate community dynamics between two or more of the NB isolates. We could then reveal any synergistic or dependent interactions occurring by measuring the change in fluorescence in a single emission range (i.e. green, red, yellow). Finally, having a segregating tool like isolate specific fluoresce will allow us to observe microenvironment localization and the changes in single isolate abundances.

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References

1. Kojima, S., & Blair, D. F. (2004). Solubilization and Purification of the MotA/MotB Complex of

Escherichia coli. Biochemistry, 43(1), 26-34. doi:10.1021/bi035405l

2. Natale, P., Brüser, T., & Driessen, A. J. M. (2008). Sec- and Tat-mediated protein secretion across the bacterial cytoplasmic membrane—Distinct translocases and mechanisms. Biochimica et Biophysica Acta (BBA) - Biomembranes, 1778(9), 1735-1756. doi:https://doi.org/10.1016/j.bbamem.2007.07.015

3. Clérico, E. M., Maki, J. L., & Gierasch, L. M. (2008). Use of synthetic signal sequences to explore the protein export machinery. Biopolymers, 90(3), 307-319. doi:10.1002/bip.20856

4. Thirunavukarasu, K., Purushothaman, S., Sridevi, J., Aarthy, M., Gowthaman, M. K., Nakajima-Kambe, T., & Kamini, N. R. (2016). Degradation of poly(butylene succinate) and poly(butylene succinate-co-butylene adipate) by a lipase from yeast Cryptococcus sp. grown on agro-industrial residues. International Biodeterioration & Biodegradation, 110, 99-107. doi:https://doi.org/10.1016/j.ibiod.2016.03.005

5. Shah, A. A., Kato, S., Shintani, N., Kamini, N. R., & Nakajima-Kambe, T. (2014). Microbial degradation of aliphatic and aliphatic-aromatic co-polyesters. Applied Microbiology and Biotechnology, 98(8), 3437 3447. doi:10.1007/s00253-014-5558-1

6. Alves, J., Vidugiris, G., Goueli, S. A., & Zegzouti, H. (2018). Bioluminescent High- Throughput Succinate Detection Method for Monitoring the Activity of JMJC Histone Demethylases and Fe(II)/2-Oxoglutarate-Dependent Dioxygenases. SLAS Discov, 23(3), 242-254. doi:10.1177/2472555217745657

7. Poldrugo, F., & Snead, O. C. (1984). 1,4 Butanediol, γ-hydroxybutyric acid and ethanol:

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Relationships and interactions. Neuropharmacology, 23(1), 109-113. doi:https://doi.org/10.1016/0028-3908(84)90226-0

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VITA

Kyle was born and raised in Nassau County, New York by his parents Veronique and Martin along with his sisters, Ashley and Sabrina. He earned his B.S. in Biology from James Madison University in Virginia where he discovered a passion for conducting experimental research. He decided to attend the University of Tennessee, Knoxville for his Ph.D. where he studied both Candida albicans genetics and Bacillus plastic degradation under Dr. Todd Reynolds until his graduation in 2019. Kyle is currently exploring opportunities to conduct research in industry, government, military and academic labs trying to avoid being the most overeducated unemployed person in America. He is accompanied by his loving wife Tamara and their menagerie of animals in their adventure to the next step in their lives. Kyle hopes to one day become a space smuggler on the fastest hunk of junk in the galaxy accompanied by an 8-foot tall walking carpet.

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