Biodegradation of Polyurethane Under Composting Conditions

BIODEGRADATION OF POLYURETHANE UNDER COMPOSTING CONDITIONS

A thesis submitted to The University of Manchester for the degree of DOCTOR OF PHILOSOPHY in the Faculty of Life Sciences

2013

Urooj Zafar

Table of Content

Table of Content ...... 2 List of Figures ...... 6 Chapter 1 ...... 6 Chapter 2 ...... 6 Chapter 2 Appendix ...... 6 Chapter 3 ...... 7 Chapter 3 Appendix ...... 7 Chapter 4 ...... 7 Chapter 4 Appendix ...... 7 List of Tables...... 8 Chapter 1 ...... 8 Chapter 2 ...... 8 Chapter 2 Appendix ...... 8 Chapter 3 ...... 8 Chapter 3 Appendix ...... 9 Chapter 4 ...... 9 Chapter 4 Appendix ...... 9 Abstract ...... 10 Declaration ...... 11 Copyright statement ...... 11 List of Abbreviation ...... 12 Acknowledgement...... 14 Chapter 1 ...... 16 1. Introduction ...... 16 1.1 Types of plastics ...... 18 1.2 General mechanisms of degradation of plastic polymers ...... 18 1.2. 1 Photo degradation ...... 19 1.2. 2 Thermal degradation...... 19 1.2. 3 Ozone induced degradation ...... 19 1.2.4 Mechano-chemical degradation...... 20 1.2.5 Biodegradation ...... 20 1.3 Biodegradable plastics...... 21 1.4 Polyurethanes ...... 22 1.4.1 Chemical structure ...... 23 1.4.2 Types of polyurethanes ...... 24 1.4.3 Biodegradation of polyurethanes ...... 27 1.5 Composting ...... 34 1.5.1 Bacteria associated with compost ...... 41 1.5.2 Fungi associated with compost ...... 42 1.5.3 Actinomycetes associated with compost ...... 43

1.5.4 Biodegradation under composting conditions ...... 44 1. 6 Techniques used to study microbial ecology ...... 45 1.6.1 Molecular approaches to studying microbial diversity ...... 45 1.6.2 Techniques based on PCR amplification of microbial community DNA/ PCR dependent techniques...... 46 1.6.2. 1 Cloning and characterization ...... 47 1.6.2.4 Terminal Restriction Fragment Length Polymorphism (TRFLP) ...... 53 1.6.2.6 454 pyrosequencing...... 57 1.6.3 Community diversity analysis ...... 60 1. 8 Summary ...... 62 1. 9 Experimental aims ...... 62 1.10 Alternative format ...... 64 1.11 References ...... 66 Chapter 2: Fungal community associated with biodegradation of polyester polyurethane buried under compost at different temperatures ...... 80 2.2 Introduction ...... 83 2.3 Materials and Methods ...... 85 2.3.1 Media composition ...... 85

2.3.1.1 R2A ...... 85 2.3.1.2 Potato dextrose Agar ...... 85 2.3.1.3 Compost Extract ...... 85 2.3.1.4 Polyurethane agar (PUA) ...... 86 2.3.2 Fabrication of polyurethane coupons ...... 86 2.3.3 Burial of polyurethane coupons in compost ...... 87 2.3.4 Recovery of polyurethane samples and enumeration of microbes ...... 88 2.3.5 Microscopic analysis of PU coupons ...... 89 2.3.6 Tensile strength and weight loss determination ...... 89 2.3.7 Extraction, amplification and purification of genomic DNA from the isolated PU degrading fungal colonies ...... 90 2.3.8 ITS rDNA sequencing and identification ...... 91 2.3.9 Extraction and amplification of community genomic DNA ...... 91 2.3.10 Analysis of the fungal community by TRFLP analysis ...... 92 2.3.11 454 Pyrosequencing ...... 93 2.3.12 Bioinformatics and statistical analysis ...... 94 2.3.13 Statistical analysis ...... 95 2.4 Results ...... 98 2.4.1 Compost chemical analysis ...... 98 2.4.2 Visual changes to PU coupons during compost burial at different temperatures ...... 98 2.4.3 Impact of soil and compost burial on the tensile strength of PU ...... 99 2.4.4 Fungal colonisation of PU coupons ...... 103 2.4. 5 Identification of polyester PU degrading fungal isolates ...... 105 2.4.6 Fungal community diversity on the surface of buried polyester PU ...... 110

2.4.7 Biodiversity of fungal communities by pyrosequencing ...... 116 2.5 Discussion ...... 124 2.6 References ...... 131 2.7 Appendix2 ...... 136 Chapter 3: Biodegradation of polyester polyurethane during commercial composting 145 3.1 Abstract ...... 146 3.2 Introduction ...... 148 3.3 Materials and Methods ...... 151 3.3.1 Fabrication of polyurethane coupons ...... 151 3.3.2 Burial of PU coupons in compost ...... 151 3.3.3 Environmental Scanning Electron Microscopy (ESEM) of the surface of PU coupons ...... 152 3.3.4 Tensile strength determination ...... 152 3.3.5 Differential Scanning Calorimetry ...... 152 3.3.6 Viable count determination ...... 153 3.3.7 Extraction, amplification and purification of genomic DNA ...... 153 3.3.9 Extraction and amplification of community genomic DNA ...... 155 3.3.10 Analysis of the fungal community by DGGE ...... 155 3.3.11 Analysis of the fungal community by TRFLP ...... 157 3.3.12 4 Analysis of the fungal community by 454 pyrosequencing ...... 158 3.3.13 Bioinformatics and statistical analysis ...... 159 3.3.14 Statistic analysis ...... 160 3.4 Results ...... 163 3.4.1 Effect of composting on the macroscopic and microscopic features of PU coupons ...... 163 3.4.2 Effect of composting on the percentage elongation and tensile strength of PU ...... 164 3.4.1 Effect of composting on the glass transition and melting temperatures of PU 165 3.4.2 Total viable fungal count and percentage impranil degrading fungi recovered from the surface of PU coupons during composting ...... 169 3.4.3 Total viable and PU degrading fungal colony count from the compost ...... 171 3.4.4 Identification of isolates recovered from compost and surface of PU coupons ...... 171 3.4.5 Comparison of the compost community with the community on the surface of PU ...... 178 3.4.6 454 pyrosequencing ...... 187 3.3 Discussion ...... 193 3.4 References ...... 199 3.5 Appendix A3 ...... 203 Chapter 4: Effect of polyurethane on fungal compost communities...... 207

4.1 Abstract ...... 208 4.2 Introduction ...... 210 4.3 Materials and Methods ...... 212 4.3.1 Compost moisture content ...... 212 4.3.2 Microbial viable count from compost ...... 213 4.3.3 Extraction, amplification and purification of genomic DNA from the isolated PU degrading fungal colonies ...... 213 4.3.4 DNA sequencing and identification of isolated PU degrading fungal colonies 214 4.3.5 Extraction of community DNA from compost ...... 214 4.3.6 Amplification of the community DNA from compost for TRFLP analysis ...... 215 4.3.7 Statistic analysis ...... 216 4.4 Results ...... 217 4.4.1 Fungal colony forming units (CFU) from PU amended compost ...... 217 4.4.2 Identification of dominant fungal isolates enriched in compost amended with Impranil DLN ...... 218 Table 4.1: Isolates recovered from impranil amended compost incubated at 45° and 50°C...... 221 4.4.3 Influence of impranil DLN and impranil DLU on the compost community profile ...... 222 4.4.4 Influence of polyester PU beads on the compost community profile...... 222 4.5 Discussion ...... 229 4.6 References ...... 234 4.7 Appendix A4 ...... 237 Table A4.1: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with Impranil DLU (liquid dispersion of polyether PU)...... 237 Chapter 5: General discussion ...... 243 5.1 General discussion ...... 243 5. 2 References ...... 249

List of Figures

Chapter 1 Figure 1.1: Equation showing formation of PU molecule by condensation of polyol and diisocynate form urethane molecule.)...... 25 Figure 1.2: Equation showing formation of urethane bond...... 26 Figure 1.3: Summary of bacterial species reported as PU degraders and source where known...... 29 Figure 1.4: Summary of fungal species reported as PU degraders and source where known...... 30 Figure 1.5: Summary of the enzymes produced by polyurethane degrading organisms and molecular weights where known...... 33 Figure 1.6: Pictorial presentation of different types of composting systems...... 36 Figure 1.7: Schematic presentation of DGGE principal...... 50 Figure 1.8: Schematic presentation of TRFLP principal...... 55 Figure 1.9: Schematic representation of 454-pyrosequencing principal...... 59 Chapter 2 Figure 2.1: Schematic representation of primers designed for unidirectional reads of amplicons from community genomic DNA...... 96 Figure 2.2: Physical changes to polyester polyurethane coupons after burial at different temperatures...... 100 Figure 2.3: Effect of compost burial on the surface of PU coupons visualised by environmental scanning microscopy...... 101 Figure 2.4: Effect of compost burial on the loss of tensile strength of PU coupons. .... 102 Figure 2.5: Changes in the total fungal viable counts and total fungal PU degrader counts over a 12 weeks incubation period...... 104 Figure 2.6: Relative distribution of the colony morphotypes recovered from the surface of PU coupons...... 107 Figure 2.7: Neighbour joining phylogenetic analysis of putative isolated PU degrading fungi...... 108 Figure 2.8: Principal Component Analysis of microbial community obtained from surface of PU and compost...... 113 Figure 2. 9: TRFLP electropherograms of TRFs from fungal communities from the surface of PU over 12 weeks buried in soil or compost...... 115 Figure 2.10: Calculated rarefaction curves of observed species based on pyrosequencing data...... 118 Figure 2.11: 454 pyrosequencing data obtained for fungal community on week 0 and 12 from compost and surface of PU coupons at 25°, 45 ° and 50°C...... 121 Figure 2.12: Principal component analysis of fungal populations obtained after 454- pyrosequencing...... 123

Chapter 2 Appendix Figure A2.1: Physical changes to polyether polyurethane coupons after burial in compost...... 136

Figure A2.2: ESEM images of polyether polyurethane coupons after burial in compost...... 137 Figure A2.3: Appearance of fungal colonies over 10 days after inoculation...... 139 Figure A2.4: Pictorial presentation of species isolated from culture based technique, on PDA (left) and PUA (right)...... 141 Chapter 3 Figure 3.1: Schematic representation of designed primers for unidirectional reads of amplicon from community genome...... 161 Figure 3.2: The effect of compost burial on the macroscopic features of PU coupons. 166 Figure 3.3: The effect of compost burial on the surface features of PU coupons under ESEM...... 167 Figure 3.5: Phylogenetic analysis of the isolated putative PU degrading fungi recovered from the surface of PU coupons...... 177 Figure 3.6: DGGE profiles from the fungal community in compost and from the PU coupons...... 181 Figure 3.7: Principal component analysis of the fungal community DGGE profiles from compost and from the surface of PU coupons...... 182 Figure 3.8: PCA analysis of the T-RFLP profiles of the fungal community in compost and on the PU surface...... 184 Figure 3.10: Rarefaction curve from 454 pyrosequencing data...... 189 Figure 3.12: Scatter plot of principal component analysis from 454 pyrosequencing of the fungal community from compost and from the surface of PU...... 191 Chapter 3 Appendix Figure A3.1: The 10 m compost pile used for burying PU coupons at the TEG group commercial compost site, Todmorden, UK...... 203 Figure A3.2: The effect of compost burial on the macroscopic features of polyether PU coupons...... 204 Chapter 4 Figure 4.1: Enumeration of fungal viable counts recovered from compost amended with or without PU dispersion (impranil)...... 219 Figure 4.2: Enumeration of fungal viable counts recovered from compost amended with or without PU beads...... 220 Figure 4.3: Principal component analysis of TRFLP profiles from fungal communities in compost amended with impranil DLN or DLU...... 225 Figure 4.4: Principal component analysis of TRFLP profiles from fungal communities in compost amended with polyester PU beads...... 227 Chapter 4 Appendix Figure A4.1: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with Impranil DLU (liquid dispersion of polyether PU)...... 237 Figure A4.2: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with Impranil DLN (liquid dispersion of polyester PU)...... 238 Figure A4.3: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks as control...... 239 Figure A4.4: PCA score for samples amended with impranil DLN...... 242

List of Tables

Chapter 1 Table 1.1: Summary of bacterial species reported as PU degraders and source where known...... 29 Table 1.2: Summary of fungal species reported as PU degraders and source where known...... 30 Table 1.3: Summary of the enzymes produced by polyurethane degrading organisms and molecular weights where known...... 33 Chapter 2 Table 2.1: Sequences of the 10 MIDs used to tag PCR amplicons from each sample for 454 pyrosequencing...... 97 Table 2. 2: Description, identification and frequency of occurrence of fungal morphotypes isolates recovered surface of buried polyester PU...... 109 Table 2.3: Variation in the Shannon-Weaver and Evenness indices derived from TRFLP electropherograms from fungal communities...... 114 Table 2.4: Statistical evaluation of fungal communities from compost and from the surface of PU coupons buried in compost incubated at different temperatures for 12 weeks...... 122 Chapter 2 Appendix TableA2.1: Change in tensile strength of polyether PU dumb-bells after 12 weeks of incubation under soil at 25° and compost at 25°, 45° and 50°C...... 138 TableA2.2: Effect of soil and compost burial on percentage weight loss of PU coupons...... 138 TableA2.3: Fungal colony count obtained from compost and surface of PU coupons after incubation at 25° , 45° and 50°C for 4, 8, and 12 weeks...... 140 TableA2.4: Assigned taxonomy to the OTU clusters of sequences obtained from pyrosequencing data. Total number of sequences obtained from each sample is mention under parenthesis beside the name of the samples...... 142

Chapter 3 Table 3.1: Sequences of the 10 MIDs used to tag PCR amplicons from each sample for 454 pyrosequencing...... 162 Table 3.2: Differential calorimetry data of PU coupons recovered from surface and centre of the compost pile...... 170 Table 3.3: Changes in the total viable fungal CFUs from PU coupons (cfu/cm2) buried at the surface and the centre of a 10 m high compost pile...... 173 Table 3.4: Changes in the total viable fungal CFUs at the surface and centre (cfu/g) of a 10 m high compost pile...... 174 Table 3.5: Frequency of isolates at recovered from polyester PU coupons buried at the surface of a compost heap...... 175 Table 3.6: Percentages homology of putative PU degrading fungal isolates for 28s rDNA region with Blast database...... 176 Table 3.7: Statistical analysis of the community profile analysed by TRFLP...... 183 Table 3.8: comparison of the shared and unique number of TRFs...... 185 Table 3.9: Statistical estimation of pyrosequencing data...... 192

Chapter 3 Appendix Table A3.1: Percentages of OTUs and their taxonomic assignments obtained by 454 pyrosequencing...... 205 Chapter 4 Table 4.1: Isolates recovered from impranil amended compost incubated at 45 and 50 °C...... 221 Table 4.2: Shannon index, Eveness and number of TRFs detected following TRFLP analysis of the fungal community in compost amended with either polyester PU beads, impranil DLN or impranil DLU...... 228 Chapter 4 Appendix Table A4.1: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with 10, 30 and 50 °C% of polyester PU beads...... 240

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Abstract

Institution: The University of Manchester Name: Urooj Zafar Degree Title: PhD Microbiology Thesis Title: Biodegradation of polyurethane under composting conditions Date: 2013

Plastic are a highly durable, lightweight and low cost family of polymeric materials that form an essential and integral component of today’s world. Their continued world-wide large scale manufacture has led them to be a major component of man-made waste. A large proportion of plastic waste is directed to the landfill sites, however their low degradation rates, scarcity of landfill sites and growing water and land pollution problems require alternatives to be developed. Composting is a natural process involving aerobic decomposition of organic wastes by a mixed microbial consortium that involves thermophilic microbes during the process due to the heat generated during decomposition. In this study we investigated the biodegradation of polyurethane under composting conditions. Polyurethanes are heteropolymers with a wide range of applications in the medical, automotive, construction and domestic field and in Europe account for 7% of all plastic manufacture and have been shown to be susceptible to biodegradation, particularly by fungi. In this thesis, it was found that loss in tensile strength of >70% occurs at both mesophilic (25°C) and thermophilic (45° and 50°C) temperatures under laboratory conditions and so is susceptible to degradation at all stages of the composting process. Moreover, polyester PU buried in compost piles at a commercial composting site during the maturation phase of an in silo composting process also underwent substantial degradation. Non-culture based analysis by TRFLP, DGGE and 454 pyrosequencing revealed that the fungal communities colonising the surface of PU was substantially different from the surrounding compost indicating selection of fungi on the PU surface. Pyrosequencing revealed that under laboratory conditions, at 25°C Fusarium solani, and 45°C and 50°C, Candida ethanolica was the dominant organism recovered from the PU surface, whereas at the commercial composting site an unidentified fungal clone and Arthrographis kalrae were the dominant organisms recovered. When the microparticulate polyester PU dispersion impranil was added to compost, a substantial shift in the indigenous fungal population was observed along with an increase in fungal viable numbers, however, addition of larger solid PU had no lasting effect on the surrounding compost community. This study demonstrates that polyester PU is highly susceptible to degradation in during composting and indicates a future potential for directing PU wastes to existing commercial composting processes.

Declaration

No portion of the work referred to in the thesis has been submitted in support of an application for another degree or qualification of this or any other university or other institute of learning

Copyright statement i. The author of this thesis (including any appendices and/or schedules to this thesis) owns certain copyright or related rights in it (the “Copyright”) and he has given The University of Manchester certain rights to use such Copyright, including for administrative purposes. ii. Copies of this thesis, either in full or in extracts and whether in hard or electronic copy, may be made only in accordance with the Copyright, Designs and Patents Act 1988 (as amended) and regulations issued under it or, where appropriate, in accordance with licensing agreements which the University has from time to time. This page must form part of any such copies made. iii. The ownership of certain Copyright, patents, designs, trade marks and other intellectual property (the “Intellectual Property”) and any reproductions of copyright works in the thesis, for example graphs and tables (“Reproductions”), which may be described in this thesis, may not be owned by the author and may be owned by third parties. Such Intellectual Property and Reproductions cannot and must not be made available for use without the prior written permission of the owner(s) of the relevant Intellectual Property and/or Reproductions. iv. Further information on the conditions under which disclosure, publication and commercialisation of this thesis, the Copyright and any Intellectual Property and/or Reproductions described in it may take place is available in the University IP Policy (see http://documents.manchester.ac.uk/DocuInfo.aspx?DocID=487), in any relevant Thesis restriction declarations deposited in the University Library, The University Library’s regulations (see http://www.manchester.ac.uk/library/aboutus/regulations) and in The University’s policy on Presentation of Theses.

List of Abbreviation

BLAST Basic local alignment search tool bp Base pairs Ca. circa cfu Colony forming unit Da dalton DEPC diethylpyrocarbonate DGGE Denaturing gradient gel polymorphism DNA Deoxyribonucleic acid dNTPs Deoxyribo nucleotide triphosphate DSC Differential scanning calorimetry EDTA Ethylenediaminetetraacetate ESEM Environmental scanning electron microscopy FISH Fluorescent insitu hybridisation Impranil DLN Polyester Polyurethane suspension used in Polyurethane Agar Impranil DLU Polyether Polyurethane suspension used in Polyurethane Agar ITS Internal transcribed spacer MC Moisture content Mpa Mega Pascal NCBI National center for biotechnology information No. Number OD Optical density PBS Phosphate buffer saline PCA Principal component analysis PCB Polychlorinated biphenyl PCL Poly (ὲ-caprolactone) PCP Pentachlorophenol PCR Polymerase chain reaction PDA Potato dextrose agar PE polyethylene PET Polyethylene tetraphthalate PHA Polyhydroxy alkanoate PHB Polyhydroxy butyrate PLA Polylactic acid PLFA Phospholipid fatty acid pPVC Plasticized polyvinyl chloride PU Polyurethane PUA Polyurethane agar PVA Polyvinyl alcohol QIIME Quantitative insight into microbial ecology rDNA Ribosomal Deoxyribonucleic acid RNA Ribonucleic acid S.E.M. Standard error mean

SEA Soil extract agar SSCP Single strand conformational polymorphism TAE Tris Acetate EDTA Tc Crystallization temperature Temp. Temperature Tg Glass transition temperature TGGE Temperature gradient gel electrophoresis Tm Melting temperature TRF Terminal restriction fragments TRFLP Terminal restriction fragment length polymorphisms whc Water holding capacity YE Yeast extract

Acknowledgement

Bismillah- i- Rahman- i- Raheem…I would like to present my deepest gratitude to my supervisor Dr. Geoff Robson for all his help and guidance throughout my project. The time and commitment he has contributed to my PhD has made it more productive., especially all his efforts and commitment given to reading my drafts is highly valued. Additionally, I would like to thank Dr Jen Cavet for her help and advice throughout my PhD.

I am deeply thankful to Dr. Ashley Houlden for all his help and answering my question which sometime would not have made much sense. I am also grateful to Higher Education Commission and University of Karachi, Pakistan for financial support.

Thanks to all my past and present members of the Robson group. Special thanks to Mehlika Karamanlioglu whose friendship and advice throughout my PhD has been greatly valued. Additionally, thanks to Adrian Langarica and Naquiddin Zairi for help with methods and letting me borrow their primers and reagents. Also, I would like to mention my colleagues at George Kenyon Hall for making my duties easier.

I would also like to thank friends and family for keeping me just in the right side of crazy. I would like to mention Nasir bhai and Nadeem mamo for pushing me to pursue my PhD and making me able to write this acknowledgement. Also my siblings, Mehwish, Sara, Isra, Maryam, Ali and Umer, and niece Saniha deserve to be mentioned, you guys deserve gifts now, eh! I also would like to mention Vikky for just being there.

Last but not the least I would like to thank Ammi and Abbo for their never ending support, and encouragement. Thanks for keeping me alive for the past twenty something years.

For Ammi and Abbo

Chapter 1

1. Introduction

Since Parkesine was first produced by Alexander Parkes in the 1850s

(Brydson, 1999), a plastic derived from cellulose that could melt upon heating and retain its shape after cooling down, there has been a large range of both natural and synthetic plastics discovered and introduced into our everyday lives. Plastics and plastic components have a number of key physical attributes that make them extremely versatile including flexibility, durability, strength, lightness, corrosion resistance, thermal and electrical insulation, and are highly economical and easy to manufacture (Andrady & Neal, 2009; Rivard et al., 1995). It is estimated that the annual worldwide production of plastic polymers in 2011 was around ca. 235

Mtonne with Europe accounting for 47 Mtonne (Plastics – the Facts 2012).

The large and continual increase in production has led to plastics becoming a major contaminant of terrestrial and aquatic ecosystems and a major environmental pollutant (Mueller 2006). In Europe, 25 Mtonne of plastic waste is generated annually (Plastics – the Facts 2012). Some plastics, such as polyurethanes which are heteropolymers, are susceptible to microbial degradation and mineralization, while many other plastics that contain a carbon-carbon homopolymer backbone such as polyvinylchloride, polystyrenes and polyethylenes are relatively resistant to degradation (Zheng et al., 2005). Plastic waste currently can be recycled, added to landfills, incinerated or biodegraded (Shah et al., 2008b). It is estimated that plastic production has outpaced recycling fivefold and out of 25 Mtonne of plastic waste produced in Europe, only 6.3 Mtonne (~25%) was recycled (Plastics – the Facts

2012). Recycling of plastics is far more difficult than glass, aluminium and paper because there are many chemically different types of plastic polymers that complicates the sorting and reprocessing procedure (Hopewell et al., 2009). There is often no indication of the type of plastic used in the material and even for the same type of plastic, formulations vary widely with different additives being incorporated.

In addition, recycling in many instances means conversion to another product, such as shredding into packaging material rather than closed loop recycling where the material can be used again as a raw material (Hopewell et al., 2009, Jayasekara et al.,

2005). From an economic perspective, recycled plastic has limited uses and market demand is weak (Hopewell et al., 2009).

In Europe in 2011, 41% (10.3 Mtonne) of total plastic waste was directed to landfills (Plastics – the Facts 2012). However, landfill disposal poses a number of problems. Contamination of soil and ground water by organic pollutants produced as breakdown by-products can cause severe environmental problems (Oehlmann et al.,

2009) affecting near-by communities and wild life. Increasingly, finding space for new land fill sites is becoming problematic and existing landfill sites are rapidly becoming full, driving the need to find alternatives for municipal waste management

(Jayasekara et al., 2005). While incineration can be a better option as some hydrocarbon polymers produce a considerable amount of energy when incinerated

(Arvanitoyannis and Bosnea, 2001), some plastics such as PVC can produce hazardous substances including furans and dioxins upon incineration (Jayasekara et al., 2005).

1.1 Types of plastics

Superficially plastics can be classified depending on whether they are derived from natural or synthetic products, on whether the polymers are formed by addition or condensation reactions or on their physical properties which divides them into thermoplastics, elastomers and thermoset plastics (Alauddin et al., 1995).

Thermoplastics are formed by the breaking of double bonds followed by polymerization to form new long chain carbon-carbon polymer backbones (Scott,

1999). Individual chains interact to form threads which upon heating, soften and solidify when they cool down (Zheng et al.,2005). Due to the carbon-carbon homopolymer backbone, they are considered as highly resistant to biodegradation

(Sabev et al., 2006; Zheng et al., 2005; Mueller 2006) although they can be blended with biodegradable components such as starch or polyesters (Amass et al., 1998).

Thermoset plastics are generally formed by condensation between a carboxylic acid and an alcohol or amine to form a long chain polyester or polyamide respectively.

They are heat stable and have a heteropolymer backbone consisting of carbon and other elements such as oxygen or nitrogen (Alauddin et al., 1995; Scott, 1999) and the presence of ester and/or amide bonds leaves these polymers susceptible to biodegradation by microbial enzymatic hydrolysis (Mueller 2006; Allen et al., 1999;

Howard & Hilliard 1999b; Nakajima-Kambe et al., 1995; Vega et al., 1999).

Thermoset plastics cannot be reheated and remoulded.

1.2 General mechanisms of degradation of plastic polymers

Broadly, there are five different mechanisms for the degradation of plastics; photodegradation, thermal degradation, ozone inducible degradation, mechano-

18 chemical degradation and biodegradation (Albertsson and Karlsson, 1988; Grima et al., 2000). These main routes of degradation are summarised below;

1.2. 1 Photo degradation

Plastics are susceptible to photo degradation particularly due to the presence of ultraviolet light (UV) which due to its high level of energy can cause direct cleavage of carbon-carbon bonds in the polymer backbone. Absorption of UV light causes excitation of molecules within the polymer generating singlet molecules which are transformed into peroxy radicals leading to direct cleavage of both saturated and unsaturated carbon-carbon bonds (Singh & Sharma 2008).

1.2. 2 Thermal degradation

Thermal degradation leads to molecular deterioration as a result of changing the physicochemical properties of the plastic and can cause random scission of links either within the polymer backbone resulting in molecular weight reduction or chain end scission of C-C bonds generating volatile end products (Singh & Sharma 2008).

1.2. 3 Ozone induced degradation

The presence of ozone in the air, even in very small concentrations, markedly accelerates the aging of polymeric materials. The interaction between ozone and polymers occurs primarily within the main polymer chain and affects carbon-carbon double bonds, aromatic rings or saturated hydrocarbon links. The reaction proceeds through an oxidation mechanism generating unstable intermediates

19 such as, peroxy radicals which can isomerise with polymer leading to hydrolysis

(Singh & Sharma 2008).

1.2.4 Mechano-chemical degradation

Mechano-chemical degradation can occur under mechanical stress and by strong ultrasonic irradiation. High levels of mechanical force, such as during milling of polymers, can lead directly to breakage within the polymer chain, reducing molecular weight and generating free radicals which can cause further chain scission

(Singh & Sharma 2008).

1.2.5 Biodegradation

The term biodegradation means “fragmentation with loss of mechanical properties or chemical modification through the action of microorganisms”. It ultimately leads to the formation of CO2, H2O and new biomass. Biodegradation can occur both aerobically and anaerobically. Aerobic biodegradation produces CO2 and

H2O while anaerobic conditions results the production of CO2, H2O and CH4 (Grima et al., 2000). Aerobic biodegradation occurs in many terrestrial environments such as soil whereas anaerobic biodegradation occurs in sediments, waterlogged soils and landfills (Shah et al., 2008b). Biodegradation is catalysed by the secretion of microbial extracellular enzymes which depolymerise synthetic polymers leading to the release of monomers and/or oligomers which are utilised by microbial cells for growth (Goldberg, 1995). The important factors in biodegradation are the chemical structure of the plastics, the types of microbes present in the environment and the environmental conditions. Initially, the surface of the polymer is affected by physical

20 and biological factors such as wetting/drying, UV irradiation and fungal penetration

(Griffin, 1980). Release of extracellular enzymes causes progressive polymer hydrolysis and fragmentation into lower molecular weight polymers, oligomers, dimers and monomers. These lower molecular weight breakdown products are then bio assimilated into the cells where they are mineralized and the energy produced is utilized (Grima et al., 2000). The enzymes involved in polymer hydrolysis can be divided into two groups mechanistically, endohydrolases and exohydrolases.

Endohydrolases randomly cleave the internal linkages of the polymer, while, exoenzymes sequentially cleave the terminal monomeric or dimeric units (Wales and

Sagar, 1985). Generally, a combination of both classes leads to complete polymer hydrolysis (Howard 2002).

1.3 Biodegradable plastics

Biodegradable plastics are those that are particularly vulnerable to microbial attack and can be synthesized from synthetic or biological monomers (for example polyvinyl alcohol and polylactic acid) and are used to make many products ranging from eating utensils to product packaging. There is a growing interest in these plastics as they have the potential to be degraded with minimal effect on the environment, however their use is currently limited.

Polyvinyl alcohol is a synthetic plastic whose polymeric backbone is composed of vinyl alcohol. It has several different uses due to its water solubility and thermo plasticity. Its water-soluble film is used for packaging and it is also used as an adhesive and thickener material in shampoos, glues, hair sprays and paper coatings and as a surfactant in eye-drops for contact lenses. It was shown in 1973 that

21 polyvinyl alcohol can be fully degraded and utilized as sole source of carbon by a bacterial strain Pseudomonas O-3 and other bacterial strains have since been identified (Shimao, 2001).

Polylactic acid is a biodegradable thermoplastic with a polymer backbone consisting of lactic acid, which is derived by the fermentation of starch, principally from corn (Singh and Sharma, 2008). It is a natural polymer and extensively used in biomedical applications such as sutures, stents and drug delivery devices but more recently is also used in food packaging materials, compost bags and disposable tableware. It can be degraded by several enzymes such as proteinase K, pronase and bromelian (Shimao, 2001).

Polyester polymers consist of ester functional group and the monomers are attached through ester linkages. Polyesters are of several different kinds that include both naturally occurring chemicals such as polyhydroxyalkonates

(polyhydroxybutyrate) as well as synthetic chemicals such as polycaprolactone.

Polyesters are used in home furnishing such as bed sheets, curtains, in industries for tyre reinforcements, ropes and safety belts. Polyesters are easily hydrolysable by enzymes such as esterases which are ubiquitously found in microorganisms (Shimao,

2001; Singh and Sharma, 2008).

1.4 Polyurethanes

Polyurethanes are synthetic and xenobiotic polymers and have found a wide range of uses in medical, automotive, domestic and industrial fields. Some common products include furnishings, coatings, adhesives, construction materials, fibres,

22 padding, paints, elastomers and synthetic skin. They are durable, resistant to water, oil, solvent, abrasion and electrical current (Sauders and Frisch 1964).

Polyurethanes have gradually replaced earlier polymers in various areas including marine and aircraft coatings and foams in car seats and furnishings

(Howard 2002). In the medical arena, polyurethane is considered as one of the most bio and blood compatible materials known. They have played a major role in the development of many medical devices due to their structural properties, blood and tissue compatibility and resistance to macromolecular oxidation, hydrolysis and calcification (Santerre et al.,2005).

Generally, xenobiotic polymers include plastics that are thought to be less susceptible to degradation because evolution has not had the time to design the enzymes to specifically attack these materials as they have not been in the environment and in contact with microorganisms for long enough (Mueller 2006).

However polyester PU’s are known to be vulnerable to microbial attack (Morton &

Surman 1994) as they contain ester linkages within the backbone of the polymer that are naturally vulnerable to esterases with biodegradation first reported by Darby &

Kaplan (1968).

1.4.1 Chemical structure

Polyurethanes were first synthesized by Otto Bayer in 1937. The main constituents of polyurethanes are isocyanate and polyols. Molecules having two isocyanate groups are called diisocyanate (Dombrow 1965). These diisocyanates are regarded as monomers because they are further polymerized to produce

23 polyisocyanates. They are classified as aromatic and aliphatic diisocyanates.

Diphenylmethane diisocyanate (MDI) and toluene diisocyanates (TDI) are examples of aromatic isocyanates while hexamethylene diisocyanate (HDI) and isophorone diisocyante (IPDI) are the examples of aliphatic PU (Szycher, 1988). Polyols are the second essential component in polyurethane synthesis. Molecules having two hydroxyl groups are called diols, three called triols and so on. Polyurethanes are produced by addition reactions between polyisocyanate (difunctional or higher) and a polyol (Figure 1.1). The overall reaction is a condensation of polyol and diisocyanate, during the reaction between isocyanate and polyol the hydrogen atom of the hydroxyl group is transferred to the nitrogen atom of the isocyanate, resulting in the formation of a urethane bond. The repeat unit is usually synthesized by addition of an alcohol across the carbon-nitrogen double bond of an isocyanate

(Dombrow, 1956). In addition, there are certain chain extenders and cross linkers like glycol moieties that influence the physical properties of the polyurethane polymer.

The formation of the urethane bond is illustrated in Figure 1.2.

1.4.2 Types of polyurethanes

Properties of polyurethanes, such as molecular orientation, crystallinity, cross linking and presence of functional groups vary with different formulations

(Sabev et al., 2006) which ultimately results in differences in the degree of susceptibility to biodegradation (Krasowska et al., 2012). Versatility in polyurethanes depends upon the variations in the R groups and substitution of the amide hydrogen.

Kim and Kim (1998) demonstrated the influence of different types of R group

Figure 1.1: Equation showing formation of PU molecule by condensation of polyol and diisocynate form urethane molecule. R1 is the hydroxyl group containing hydrocarbon chain, R2 is the hydrocarbon chain of polyisocyanate whereas, n represent the number of repetitions (Dombrow 1956; Sauders and Frisch 1964).

Figure 1.2: Equation showing formation of urethane bond. Hydroxyl linkage of the diol compound is broken down. Hydrogen binds with the Nitrogen of diisocyanate, with the breakage of double bonds, thus form the ester linkage of the urethane moiety (Dombrow 1956; Sauders and Frisch 1964).

26 substitution on the degradability of the polymer. They suggested that polyols containing diols are more degradable because they induce less hydrophobicity.

Krasowska et al., (2012) studied the degradation of PU with different chemical compositions under natural weathering (composting) conditions. They suggested that the addition polycaprolactone enhanced the rate of biodegaradation. It has also been reported that PU containing ester linkages are more susceptible than

PU containing ether linkages (Darby & Kaplan 1968; Filip 1979; Barratt et al., 2003,

Krasowska et al., 2012). With regard to the diisocyanate, it has been suggested that aliphatic diisocyanates are more susceptible to biodegradation than those having aromatic groups (Darby & Kaplan, 1968) as aliphatic diisocyanates are more flexible and accessible.

The major advantages in terms of biodegradation with PU is the susceptibility due to the presence of repeating units of urethanes that contains moieties such as urea, esters, ethers and aromatic groups. These moieties are the most reported sites of action of degrading enzymes, like ureases, esterases and proteases

(Filip 1979; Loredo-Treviño et al., 2011).

1.4.3 Biodegradation of polyurethanes

There are a number of reports regarding the biodegradation of PU. Both bacteria and fungi are known to participate in PU biodegradation but most work in this area has focused on bacteria with several strains studied to determine the mechanistic pathway of degradation. However, in the environment, fungi have been reported to be the predominant degraders of PU (Barratt et al., 2003). Table 1.1

27 summarizes the bacterial species and Table 1.2 the fungal species reported as PU degraders.

The number and fungal species reported in the literature as potential PU degraders varies widely reflecting the range of environments where the organisms were isolated from. The majority of fungal species were isolated from soil environments with Aspergillus spp. and Penicillium spp. highly prevalent due to their ubiquitous nature in the environment. Russell et al., (2011) reported on a collection of endophytic fungi that were capable of degrading PU. High degradative activity was reported among several isolates of Pestalotiopsis microspora in both solid and liquid suspension and under both aerobic and anaerobic conditions. The majority of reports in the literature relate to the biodegradation of polyester PU as polyether based PU’s have been shown to be highly recalcitrant (Darby & Kaplan 1968;

Nakajima-Kambe et al., 1999). However, one recent report (Matsumiya et al., 2010) demonstrated that an Alternaria sp.PURDK2, was capable of defacing polyether PU and degradation was linked with secreted urethane-bond degrading enzyme (s).

A number of enzymatic activities have been linked to PU degradation, that differ in their site of action and include esterases, ureases, proteases and polyurethanases (reviewed by Howard 2012; Loredo-Treviño, et al., 2011b) and are summarised in Table 1.3.

Table 1.1: Summary of bacterial species reported as PU degraders and source where known.

Bacterial species Reference Source

Acinetobacter El-Sayed et al., 1996 Oil contaminated soil, USA calcoaceticus Acinetobacter gerneri Howard et al., 2012 Soil, USA

Aeromonas salmonicida Kay et al., 1991 Soil buried PU samples

Alcaligenes denitrificans Kay et al., 1991 Soil buried PU samples

Alicycliphilus spp. Oceguera-Cervantes et al., Decomposed soft foam 2007 Alicycliphilus Oceguera-Cervantes et al., Decomposed soft foam denitrificans 2007 Arthrobacter sp. Shah et al., 2008a Activated sludge, Pakistan

Arthrobacter globiformis El-Sayed et al., 1996 Oil contaminated soil, USA

Bacillus sp. Shah et al., 2008a; Blake et Activated sludge, Pakistan; not al., 1998 specified Bacillus subtilis Rowe & Howard 2002 Mesocosm (not specified)

Bacillus pumilus Nair & Kumar 2007 PU contaminated water from industrial waste sites Comamonas acidovorans Nakajima-Kambe et al., 1995 Soil, Japan

Corynebacterium sp. Kay et al., 1991; Shah et al., Soil buried PU samples Activated 2008a2008 sludge Enterobacter agglomerans Kay et al., 1991 Soil buried PU samples

Methanotrix sp. Varesche et al., 1997 Degraded PU foam

Micrococcus sp. Shah et al., 2008a Activated sludge, Pakistan

Pseudomonas sp. Shah et al., 2008a Activated sludge, Pakistan

Pseudomonas aeruginosa Kay et al., 1991 Soil buried PU samples

Pseudomonas fluorescens Howard & Blake 1998 Soil, USA

Pseudomonas Howard & Hilliard 1999 Soil, USA chlororaphis Pseudomonas maltophilia Kay et al., 1991 Soil buried PU samples

Rhodococcus equi Akutsu-Shigeno et al., 2006 Soil, Japan

Serratia rubidaea Kay et al., 1991 Soil buried PU samples

Staphylococcus Jansen et al., 1991 Not specified epidermidis

Table 1.2: Summary of fungal species reported as PU degraders and source where known.

Fungal species References Source

Acremonium sp. (Stranger-Johannessen, 1985) Degraded PU cable in marine environment

Alternaria dauci (Russell et al., 2011) Ecuadorian rainforest

Alternaria spp. (Stranger-Johannessen, 1985; Degraded PU film in air; not specified; Matsumiya et al., 2010; Russell et Ecuadorian rainforest al., 2011)

Alternaria sp.strain 18-2 (Cosgrove et al., 2007) Garden soil, Manchester, UK

Alternaria alternata (Pommer and Lorenz, 1985) Soil, Germany

Alternaria solani (Ibrahim et al., 2009) Soil, Jordan

Aspergillus sp (Stranger-Johannessen, 1985) Degraded PU cable in marine environment

Aspergillus fischeri (Bentham et al., 1987) Soil John Innes No. 2 compost

Aspergillus flavus (Darby & Kaplan, 1968; Pathirana Not specified;soil, Birmingham, UK; not and Seal, 1984; Pommer and Lorenz, specified; soil from waste disposal site, 1985; Mathur and Prasad, 2012) India

Aspergillus fumigatus (Pathirana and Seal, 1984) Soil, Birmingham, UK

Aspergillus niger (Darby & Kaplan, 1968; Filip, 1979; Not specified Amaral et al., 2012)

Aspergillus terreus (Wales and Sagar, 1985) Not specified

Aspergillus ustus (Pommer and Lorenz, 1985; Not specified;Soil John Innes No. 2 Bentham et al., 1987) compost

Aspergillus versicolor (Darby & Kaplan, 1968; Bentham et Not specified; Soil John Innes No. 2 al., 1987) compost)

Aureobasidium pullulans (Darby & Kaplan, 1968; Crabbe et Not specified; Soil, Washington DC al., 1994)

Bionectria spp. (Russell et al., 2011) Ecuadorian rainforest

Chaetomium globosum (Darby & Kaplan, 1968; Pathirana Not specified; and Seal, 1984)

Cladosporium sp. (Crabbe et al., 1994) Garden soil, Washington DC

Cryptococcus laurentii (Cameron et al., 1987) Not specified

Curvularia senegalensis (Crabbe et al., 1994) Garden soil, Washington DC

Cylindrocladiella parva (Cosgrove et al., 2007) Garden soil, Manchester, UK

Edenia gomezpompae (Russell et al., 2011) Ecuadorian Amazonian rainforest

Exophiala leanselmei (Owen et al., 1995) Soil

Fusarium culmorum (Bentham et al., 1987) Soil John Innes No. 2 compost

Fusarium oxysporum (Pommer and Lorenz, 1985) Soil, Birmingham, UK; not specified

Fusarium solani (Bentham et al., 1987; Crabbe et al., Soil John Innes No. 2 compost; Garden 1994) soil, Washington DC USA

Geomyces pannorum (Barratt et al., 2003; Cosgrove et al., Garden soil, Manchester, UK; Soil John 2007) Innes No. 2 compost

Gliocladium roseum (Pathirana and Seal, 1984) Soil, Birmingham, UK

Lasiodiplodia spp. (Russell et al., 2011) Ecuadorian rainforest

Nectria spp. (Cosgrove et al., 2007; Russell et al., Ecuadorian rainforest; Garden soil, 2011) Manchester, UK;

Nectria gliocladiodes (Barratt et al., 2003) Soil John Innes No. 2 compost

Neonectria ramulariae (Cosgrove et al., 2007) Garden soil, Manchester, UK

Nitrospora spherical (Pathirana and Seal, 1984) Soil, Birmingham, UK

Penicillium spp. (Pathirana and Seal, 1984; Bentham Soil John Innes No. 2 compost et al., 1987)

Penicillium (Pathirana and Seal, 1984; Bentham Soil, Birmingham, UK; Soil John Innes No. chrysogenum et al., 1987) 2 compost

Penicillium citrinum (Pathirana and Seal, 1984) Soil, Birmingham, UK

Penicillium decumbens (Pommer and Lorenz, 1985) Not specified

Penicillium funiculosum (Darby & Kaplan, 1968; Pommer Not specified; and Lorenz, 1985)

Penicillium inflatum (Cosgrove et al., 2007) Garden soil, Manchester, UK

Penicillium notatum (Cosgrove et al., 2007) Soil, Birmingham, UK; Garden soil, Manchester, UK Penicillium (Pommer and Lorenz, 1985; Barratt Soil John Innes No. 2 compost ochrochloron et al., 2003)

Penicillium (Pommer and Lorenz, 1985) Not specified purpurogenum

Penicillium rugulosum (Pommer and Lorenz, 1985) Not specified

Penicillium variabile (Pommer and Lorenz, 1985) Not specified

Penicillium viridicatum (Cosgrove et al., 2007) Garden soil, Manchester, UK

Pestalotiopsis (Russell et al., 2011) Ecuadorian Amazonian rainforest microspora Pestalotiopsis sp. (Russell et al., 2011) Ecuadorian Amazonian rainforest

Phaeosphaeria spp. (Russell et al., 2011) Ecuadorian Amazonian rainforest

Phoma spp. (Stranger-Johannessen, 1985) Degrading PU film in air

Phoma fimenti (Pommer and Lorenz, 1985) Not specified

Plectosphaerella spp. (Russell et al., 2011) Ecuadorian Amazonian rainforest

Pleosporales spp. (Russell et al., 2011) Ecuadorian Amazonian rainforest

Rhizopus stolonifer (Wales and Sagar, 1985) Not specified

Scopulariopsis (Pathirana and Seal, 1984) Soil, Birmingham, UK brevicaulis Scopulariopsis fusca (Pommer and Lorenz, 1985) Not specified

Talaromyces spp. (Pommer and Lorenz, 1985) Not specified

Trichoderma spp. (Darby & Kaplan, 1968; Pathirana Not specified; Soil,Birmingham, UK; and Seal, 1984; Loredo-Treviño, et DIA/UAdeC collection, Spain al., 2011a)

Trichoderma viride (Pathirana and Seal, 1984; Pommer soil, Birmingham, UK; Not specified and Lorenz, 1985)

Table 1.3: Summary of the enzymes produced by polyurethane degrading organisms and molecular weights where known.

Source of Enzyme Mol. Wt Enzyme specificity References

Alicycliphilus spp. Esterases Oceguera-Cervantes et al., 2007

Alternaria solani - Proteases Ibrahim et al., 2009

Aspergillus flavus - Esterases Mathur & Prasad 2012

Bacillus pumilus - Lipases & Esterases Nair & Kumar 2007

Bacillus subtilis 28 Esterases Rowe & Howard 2002

Comamonas acidovorans 42 Esterases/Protease Allen et al., 1999

Comamonas acidovorans 62 Esterases Akutsu et al., 1998

Corynebacterium spp. - Esterases Kay et al., 1993

Curvularia senegalensis 28 Esterases/Proteases Crabbe et al., 1994

Geomyces pannorum - Esterases Cosgrove et al., 2006

Pestalotiopsis microspora - Serine hydrolases Russell et al., 2011

Pseudomonas chlororaphis 63 Esterases/Protease Howard 2002

Pseudomonas chlororaphis 60 Esterases Ruiz et al.,1999

Pseudomonas chlororaphis 65 Esterases/ Protease Ruiz et al., 1999

Pseudomonas chlororaphis 31 Esterases Ruiz et al., 1999

Pseudomonas chlororaphis 27 Esterases Howard & Hilliard 1999b

Pseudomonas fluorescens 29 Proteases Howard & Blake 1998

Pseudomonas fluorescens 48 Esterases Vega, et al., 1999

Rhodococcus equi 55 Amidases/ esterases Akutsu-Shigeno et al., 2006

Staphylococcus epidermidis - Ureases Jansen et al., 1991

Trichoderma spp. - Ureases Loredo-Treviño et al., 2011a

Esterase activity has received considerable attention as esterases are known to hydrolyse the ester linkages of main polymer chain (Akutsu et al., 1998). Both membrane-bound and extracellular esterase activity have been linked to PU hydrolysis (Akutsu et al., 1998; Ruiz et al., 1999; Allen et al., 1999; Vega et al.,

1999) depending on the organism are either inducible or constitutive.

A membrane bound esterase was isolated from Comomonas acidovorans by

Akutsu et al., (1998), that absorbed hydrophobically on to the surface of PU followed by hydrolysis. The enzyme consists of two domains, a surface binding domain (SBD) and a catalytic domain. Both the domains are in close proximity with the hydrophobic SBD enabling binding of the enzyme to PU surface. Rowe & Howard

(2002) suggested that the cell bound nature of the enzyme decreases the competition between PU degraders and non-degraders. An extracellular, soluble esterase of 42 kDa has also been isolated from the same organism (Ruiz et al., 1999; Allen et al.,

1999; Vega et al., 1999) that also causes PU hydrolysis. Howard (2002) and Zheng et al., (2005) suggested that both enzymes play different roles during degradation, the surface bound esterase hydrolyses substrate into soluble metabolites, which are then taken up by the cell for further metabolism while the extracellular enzyme hydrolyses the PU polymer into smaller units making it more accessible to microorganisms.

1.5 Composting

Composting is a managed self-heated, aerobic process that controls biological decomposition and transformation of biodegradable materials into a humus-like substance (compost). It is a natural process that results in the production of CO2 and

H2O with the release of minerals and stabilized organic matter (Arvanitoyannis &

Bosnea 2001; Shah et al., 2008b). The most common composting methods are, windrow composting, passively aerated windrows (PAWS), static piles (with or without forced aeration), enclosed or in-vessel composting and vermicomposting.

Windrow composting is an open air process that involves the formation of long triangular shaped heaps (windrows, 3-6 feet) that are turned periodically to reintroduce oxygen and release heat, water vapour and gases until the composting process has been completed (typically up to 16 weeks, Figure 1.3a). The passively aerated windrow system (PAWS) includes perforated pipes placed at the base of each windrow to promote convective airflow throughout the pile. Static piles are the simplest form and require little management. They are compiled and left undisturbed so aeration and moisture content is dependent on size and proportion of bulking materials. It is simple but relatively slow, typically requiring many months for stabilization. The forced aerated static systems are similar to PAWS piles, but blowers are installed at the ends of perforated pipes. Airflow can be adjusted by changing the frequency and duration of the blower. The aerated piles process achieves substantially faster composting rates (Figure 1.3c). In in-vessel systems, material is contained within closed metal or concrete containers allowing a degree of control over airflow and temperature (Figure 1.3b). Finally, vermicomposting; in which red worms like, Eisenia foetida, actively transform decaying matter into product called worm castings which are rich in organic nutrients for plants (Atlas and

Bartha, 1998; Cooperband, 2002).

a b

The Teg Group composting site, Todmorden UK, photo credit U.Zafar 2012

(Adapted from Cooperband 2002)

Figure 1.3: Different types of composting systems. (a) Steaming compost, windrow (b) in-vessel and (c) static pile with forced aeration. Pictures taken by the author at Teg commercial composting site at Todmorden UK (aandb), picture c is adapted from Cooperband (2002).

The major advantages of composting food and green waste are that it is rapid, cost viable environmentally friendly and diverts waste from landfill sites. Due to the high temperatures that occur during the composting process, as a result of microbial respiration and growth, deterioration of material is accelerated (Kim &

Kim 1998) and simultaneously destroys most of the pathogens present (Shah et al.,

2008b).

There are a number of environmental, physical and chemical factors that affect the process of composting. The main factors are oxygen, moisture content, temperature and the feedstock itself (Grima et al., 2000). Microbial respiration is an indicator of decomposition and stability of the compost product. Aerobic decomposition of organic matter requires O2 and an efficient aeration system is required for effective composting (Greizerstein et al., 1993). Three different methods are commonly used to provide O2 during the composting process, physical turning of the compost pile, connective airflow and mechanical aeration. Compost heaps are designed to have a free air space of around 20-30% of the total volume. Reduction in moisture content increases free air space and leads to improved decomposition

(Epstein, 1997). Ventilation is not only important in supplying O2, but also aids in cooling (Finstein et al., 1983) with sufficient ventilation and temperatures >45°C required for efficient biodegradation (Leonas et al., 1994).

Feedstock composition is an important factor in composting affecting the duration of each phase as well as the microbial species present and microbial diversity (Ryckeboer, et al., 2003b). The C:N ratio is known to affect the process

37 with a ratio of not greater than 40:1 required for sufficient microbial growth (Atlas &

Bartha 1998). Readily decomposable organic matter such as cellulose degrades relatively quickly, for example, cardboard milk cartons were reported to degrade in just 20 days, while 89% degradation was observed for wax coated cardboard 85 days.

On the other hand the recalcitrant decomposable compounds such as lignin have a slow rate of decomposition so the process continues for longer and at a slower rate with the slow release of carbon monomers. Nitrogen content is required for the protein synthesis. On a dry weight basis, bacteria contain 7-11% and fungi 4-6% nitrogen. An increase in the amount of nitrogen in the feedstock results in the release of ammonia, an increase in pH and a reduction in cellular growth (Epstein, 1997).

Moisture content is an important factor in composting with water release occurring from the feedstock during microbial respiration and water loss occurring through evaporation, though the amount of water lost is generally less than that produced. Moisture content and water holding capacity (WHC) of the feedstock are critical factors for microbial development and their metabolic activity. Optimum moisture content is between 50 -60% (Schulze, 1962), below 40% the microbial activity decreases while above 70% anaerobic conditions develop.

Temperature variation plays a critical role in composting. Temperature is not uniform throughout the composting mass, with most heat at the centre of the pile with cooler regions around the surface of the pile. As the composting process occurs, the composition of the microbial community changes from mesophilic to thermophilic organisms as the temperature rises and before mesophiles re-emerge during the maturation (cooling) phase. These phases reflect the activities of

38 successive microbial populations performing the degradation of recalcitrant organic matter. Initially, the temperature increases due to the rapid utilisation of readily degraded carbon within a few days with temperatures rising to between 50° C and

70° C and is maintained for a number of weeks depending on the composting system and feedstock before entering the maturation phase during which the compost cools and more recalcitrant compounds are utilised (Grima et al., 2000). The optimum temperature(s) for the composting process depends on the feedstock and composting system. Most studies suggest temperatures rising to 50-60˚C is often optimal for efficient biodegradation (Epstein 1997; Pagga et al., 1995).

On the basis of temperature variation during the composting process, composting can be divided into four main phases: initial, thermophilic, cooling and maturation. The different temperature phases reflect the relative activities of the microbial population. During the initial phase, substrate is at an ambient temperature

(25-45ºC) and the pH is generally slightly acidic. Mesophilic bacterial and fungal populations are dominant in this phase with only occasional recovery of thermophilic/tolerant fungi and bacteria (Partanen et al., 2010; Hultman et al., 2010).

With the degradation of readily utilised organic compounds, the pH decreases favouring the growth of filamentous fungi and yeasts, however, with high levels of nitrogen ammonification causes an increase in pH favouring bacterial growth. During the early stages of composting, bacteria are thought to be responsible for major decomposition and heat generation. They have high surface area to volume ratio that allows the rapid transfer of soluble substrate into the cells and have a short generation time and contain species that are more thermophilic/thermotolerant than fungi. As the temperature rises above 50°C, the composting process enters the

39 thermophilic stage. It is reported that bacteria and actinomycetes play an important role in degradation during this phase and Goodfellow & Williams, (1983) reported an increase in the degradation of xenobiotic compounds during this phase. Pathogen destruction is also observed in this phase due to the high temperature.

During the thermophilic phase, species diversity decreases with the appearance of thermophilic and/or thermotolerant bacteria, actinomycetes and fungi

(Beffa et al., 1996; Ishii et al., 2000). At temperatures around 50˚C, all three groups are active in the compost, however, as the temperature increases further, the fungal population decreases and actinomycetes and bacteria are predominant. At 70˚C or higher, spore-forming bacteria are dominant. Major degradation of a number of different polymers including cellulose and xylan are reported to occur during the thermophilic phase (Beffa et al., 1996; Ryckeboer et al., 2003b). Ishii et al., (2000) suggested that the increased temperature drastically decreases microbial populations, which affects rate of decomposition; however Beffa et al., (1996) suggest that, higher temperatures (60-70˚C) favour cellulose degradation and cellulolytic organisms appear mainly at the end of the thermophilic phase. As substrate availability becomes limiting, microbial growth decreases, temperature falls and the compost enters the cooling/second mesophilic phase. As the temperature decreases, mesophilic organisms recolonize the compost pile. Microbial diversity increases and more recalcitrant organic compounds are utilised. As the compost cools it enters the maturation phase and the compost becomes stabilised and forms humus (Epstein

1997; Ishii et al., 2000; Ryckeboer, et al., 2003b). Recent studies by Gannes et al.,

(2013) and Xu et al., (2013) using 454 pyrosequencing and qPCR, respectively, have suggested that distinct communities exist at different stages of the composting

40 process including the maturation phase. Previously, Chang and Hudson, (1967) reported that due to the heating phase several fungal species including

Aureobasidium pullulans, Alternaria tinius and Cladosporium herbarum died and did not reappear in the maturation phase.

1.5.1 Bacteria associated with compost

Bacteria have a large surface area to volume ratio which allows rapid transport into cells and some genera, like Bacillus, produce thick walled endospores which are very resistant to heat, radiation and chemical disinfections. These factors contribute towards their dominance at high temperatures. They are present throughout the composting process as active or dormant cells, or as spores although their relative abundance and activity changes. Diversity has been reported to be influenced by the composition of the feedstock and composting conditions although other studies suggest that the feedstock composition has little effect on biodiversity

(Takaku et al., 2006; Vaz-Moreira et al., 2008; Partanen et al., 2010; Gannes et al.,

2013; Xu et al., 2013).

During the early mesophilic phase of composting a rather heterogeneous population of bacteria develops leading to a rapid rise in temperatures. During the mesophilic stage when temperatures are in the range 25-45˚C and the pH is acidic,

Lactobacillus, Leuconostoc and Pseudomonas genera were reported to be dominant with domestic household waste as feedstock (Partanen et al., 2010) while with wood chips and plastic bottle flasks as feedstock, bacteria in the order Bacillales were reported as dominant (Watanabe et al., 2010) including Pseudomonas, Klebsiella and

Bacillus (Strom 1985a; Strom 1985b). Acetobacter are also commonly present

41 because they are known to use substances produced by lactic acid bacteria and yeasts as substrates (Partanen et al., 2010).

With the activity of mesophilic community temperature rises and O2 becomes depleted at certain sites within the compost pile. At this stage, dominance of several Bacillus species was evident (Beffa et al., 1996; Ishii et al., 2000; Dees and

Ghiorse, 2001; Ryckeboer et al., 2003a; Ryckeboer et al., 2003b). In studies by

Strom (1985a; 1985b) 87% of 652 randomly picked colonies were Bacillus, with B. circulans and B. stereothermophillus being the most frequent representative of this genus. During the thermophilic phase, members of the genus Thermus form a dominant group and Beffa et al., (1996) reported that at temperatures in the range 65-

82˚C, populations of the genus Thermus occured at rates 107-1010 cells g-1 dry weight.

Takaku et al., (2006) suggested that during the cooling phase, Protobacterea, principally acinetobacter survive, followed by repopulation by Bacteriodetes

(Chitinophaga and Sporocytophaga). During maturation the community stabilises and other genera including Gram positive bacteria (Cellulomonas, Rhodococcus) and

Gram negative bacteria (Pseudomonas, Havobacterium) reappear (Beffa et al.,

1996).

1.5.2 Fungi associated with compost

Thambirajah & Kuthubutheen (1989) reported the fungal count obtained from Palm press compost contained ca. 106 mesophilic fungal CFU per g in the mesophilic stage and ca. 103-106 thermophilic fungi per g of compost during the thermophilic and maturation phases. A number of genera have been found to be

42 prevalent in compost including Acremonium, Aspergillus, Cladosporium,

Penicillium, Pseudollescheria and Thermomyces species (Anastasi et al., 2005).

Hultman et al., (2010) used molecular based approach and reported that fungal communities develop through different stages of composting, mesophilic fungi are abundant during the mesophilic phase and low pH phase with mucorales as a dominant group but as the temperature rises to around 60°C, diversity declines drastically and Ascomycota yeasts become dominant, including Saccharomyces,

Geotrichum and Candida (Bonito et al., 2010). Thambirajah & Kuthubutheen, (1995) also reported that 60˚C was the upper range limit for the growth of fungi during composting and upon cooling, fungi reappear in the compost, but whether these stem from vegetative cells or spores that survived the composting process or are reintroduced from the environment remains unknown. A wide range of fungal genera has been isolated from composts with thermotolerant and thermophilic genera of particular importance. A number of thermophilic/tolerant fungi have been isolated during different studies. The most commonly cited ones are Aspergillus fumigatus and Humicola lanuginosa from all habitats tested, with Agaricus bisporus,

Chaetomium thermophile, Thermoascus aurantiacus, Mucor pusillus, Talomyces duponti, Stilbella thermophila, Humicola grisea and H. insolense among the most predominant (Fergus, 1964; Straatsma et al., 1994; Ryckeboer et al., 2003b; Hultman et al., 2010).

1.5.3 Actinomycetes associated with compost

The ecology of actinomycetes in relation to compost has been reviewed extensively. Actimomycestes tolerate higher temperatures and pH compared to the

43 fungi and are reported to be found during thermophilic, cooling and maturation phases (Tuomela et al., 2000). Members of the family Streptomycetaceae (Korn-

Wendisch et al., 1995) and some genera containing thermophilic species including

Thermomyces, Thermomonospora and Streptomyces are predominant (Strom 1985a).

Fergus (1964) identified eleven species of thermotolerant/ philic actinomycetes,

Nocardia brasiliensis, Pseudonocardia thermophila, Streptomyces rectus, S. thermpviolaceus, S. thermovulgaris, S. violaceoruber, Thermoactinomyces glaucus,

T. vulgaris, T. fusca, Thermopolyspora polyspora and Thermomonospora curvata.

1.5.4 Biodegradation under composting conditions

With an increased knowledge and understanding of the composting process, there has been a shift away from landfills and incineration towards sophisticated, environmentally friendly composting technology for green and domestic waste in recent years (Williams et al., 1992; Mergaert et al., 1994). Composting has also been considered as a potential route for the degradation of some synthetic polymers. For example, Sasek et al., (2006) described the potential for the degradation of aliphatic and aromatic co-polyesters during composting, while Krasowska et al., (2012) has studied the biodegradation of PU under natural compost piles in a 24 month study and reported significant biodegradation at the end of the study. A number of biodegradable polymers have also been developed. The basic parameters for such polymers are biodegradability, disintegratibility, absence of negative effects on the process of composting and no adverse ecotoxicological effects (Chiellini et al.,

1996). A number of studies have been published dealing with the composting of

44 natural and synthetic polymers (Narayan, 1993; Tuomela et al., 2000; Witt, et al.,

1997).

1. 6 Techniques used to study microbial ecology

Microbial diversity depends upon total number and richness, evenness and distribution of active species. There are two elements that define microbial diversity for any particular environmental niche; first the abundant and metabolically active members and secondly, less abundant, rarer taxa that are either slow growing or metabolically non-active (Trevors, 1998).

1.6.1 Molecular approaches to studying microbial diversity

DNA has been frequently targeted for molecular techniques in order to study microbial diversity in natural communities. DNA is at the core of every form of life and provides accurate information encoded within them. This enables individual species to be discriminated from each other and aid in identification by phylogenetic analysis.

Nucleic acid based techniques require DNA extracted from the entire microbial community of niche in question. However, as DNA cannot be extracted from every cell with the same level of efficiency, for example, due to variation in cell lysis, extracted DNA will not represent exactly the proportion of different species present in the community (Prosser, 2002). A number of studies have sought to improve DNA extraction methods and reduce community bias (Von Wintzingerod et al., 1997; Zhou et al., 1996). For a reliable picture, pure and high amount of DNA is

45 required, covering a large diversity of the community (Stach et al., 2001; Yang et al.,

2006).

Following DNA extraction, PCR is frequently employed to amplify a specific region of the DNA using primers to conserved regions between species although not all DNA based analyses employ PCR (Ranjard et al., 2000).

1.6.2 Techniques based on PCR amplification of microbial community DNA/

PCR dependent techniques

PCR-based genetic fingerprinting techniques aim to provide a global picture of the genetic structure and composition of microbial communities independent of conventional culturing. PCR-based approaches are frequently used in microbial ecology because it requires very small amount of DNA and amplifies into billions of copies in a matter of hours. As a consequence it is highly sensitive and capable of amplifying target DNA even if it is present in very limited numbers (Osborn et al.,

2000). Use of specific primers makes PCR specific to particular sequences in the target DNA, however, while these techniques have many advantages, there are also a number of limitations and caveats associated with PCR (Osborn et al., 2000). DNA extraction is assumed to be equally efficient for all species present in a community and in addition, the presence of inhibitors such as humic acid in soils can also interfere with PCR reactions by masking the detection of DNA and block DNA- enzyme interactions (Steffan et al., 1988) although the use of reagents such as

DMSO and bovine serum albumin can help to negate these inhibitors (Winship 1989;

Von Wintzingerode et al., 1997). Differences in PCR efficiency (bias) of target DNA between different species can also influence apparent diversity due to differing copy

46 numbers of target DNA between species, differences in the G+C ratio of target genes, primer affinity and specificity and formation of chimeras (Von Wintzingerode et al., 1997; Wang and Wang 1996). Following are a few techniques used to study microbial communities:

1.6.2. 1 Cloning and characterization

This approach is used to investigate the diversity of microbial communities by producing a library of clones from target sequences obtained by PCR from extracted DNA. Sequencing of individual isolates then allows identification of uncultured isolates as well as an estimation of their abundance in a community

(Handelsman, 2004).

1.6.2.2 Ribosomal intergenic spacer analysis (RISA)

In this technique, fluorescent primers are used to amplify intergenic spacer regions of community DNA and then run either on polyacrylamide gel (RISA) or in automated sequencer (ARISA) in denatured condition with standard size marker that separates microbial community fingerprints based on the size of PCR amplicons

(Borneman and Triplett, 1997).

1.6.2.3 Denaturing Gradient Gel Electrophoresis (DGGE)

DGGE is widely used to study the genetic diversity of uncharacterized microbial populations. The theoretical principal of DGGE is to separate DNA fragments of similar length but differing in DNA sequence (Muyzer et al., 1993;

Fischer & Lerman, 1983). Separation of PCR products is based on the decreased

47 electrophoretic mobility of a partially melted double-stranded DNA molecule in polyacrylamide gels containing a linear gradient of DNA denaturants (a mixture of urea and formamide). Melting of DNA fragments proceeds in discrete melting domains: the stretches of DNA with identical melting concentration of denaturant.

During migration the helical DNA converted to partially melted molecule, that halts the movement. Since the sequence of a DNA fragment influences the melting behaviour, molecules with different sequences will stop migrating at different positions in the gel. In this manner DNA fragments can be separated according to their sequences. An overview of DGGE is shown in Figure 1.4.

DGGE has been widely applied to investigate community diversity in the field of environmental microbiology and is inferred from the number and position of individual bands. The intensity of the bands directly relates to their quantity in the amplicons, in theory the more intense the bands the more abundant the species within the community (Myers et al., 1985). Identification of individuals within the community can also be performed following band extraction and sequencing or by hybridization with labelled specific oligonucleotide probes (Muyzer and Smalla,

1998).

DGGE has been widely used as a method of microbial community analysis as it has a number of advantages. It is a culture-independent technique enabling the inclusion of species that cannot be readily cultured (Muyzer and Smalla, 1998). It is a highly sensitive technique that can separate single base pair substitutions in a DNA fragment of ~500 base pairs (Fischer & Lerman, 1983). Myers (1985) estimated that theoretically, approximately 95% of all possible single base substitutions should be

48 able to be separated by DGGE. This technique is also capable of detecting relatively minor members of the community present with a resolution of species that make up less than 0.1% of the total population (Van Elsas et al., 2000).

1.6.2.3.1 Application of DGGE in microbial ecology analysis

Due to its broad applicability, DGGE has been extensively used for variable purposes. A few of the main uses are listed below;

1. Studying community complexity

Numerous publications over the past two decades have employed DGGE to study biodiversity in microbial populations. James et al., (2007) studied the community structure of chronic wound specimens that revealed diverse microbial communities and the presence of bacteria, including strictly anaerobic bacteria, not revealed by cultivation methods. The diverse microbial nature of specimens underscores the utility of molecular methods for examining complex microbial communities.

Community DNA from extreme environments has been analysed via DGGE successfully. Muyzer et al., (1995) studied genetic diversity of the microbial community around two hydrothermal vents and managed to identify the dominant members-Thiomicrospira sp. and Desulfovibrio salexigens.

Figure 1.4: Schematic presentation of DGGE principal. Community DNA is first amplified using GC tailed primers then amplified products are separated on the basis of their GC content in a polyacrylamide gel containing increasing amount of denaturant from top to bottom.

2. Studying community variation over time

DGGE is an important tool in studying microbial community succession over time, or the effect of any environmental changes (naturally or artificially induced). More than one sample can be run at a time on gels that enable comparison of the banding pattern of DGGE-generated community profiles of the samples.

Cocolin et al., (2001) used DGGE to monitor the microbial community succession during ripening of natural fermented sausages, while Houlden et al., (2008) investigated the seasonal shifts in rhizosphere microbial populations during plant development. Ishii et al., (2000) used DGGE to study the microbial succession during a laboratory-scale composting process of garbage and reported that discrete bacterial populations could be recovered at different stages of the process. During the mesophilic phase some fermenting bacteria, such as Lactobacillus sp., were present while thermophilic Bacillus sp. appeared during the thermophilic phase with a more complex community developing during the maturation phase.

3. Monitoring the enrichment and isolation of bacteria

Apart from analysing microbial diversity and its succession, DGGE can also be used to study mixed cultures and isolation of desired species. Jin et al., (2012) monitored an enriched microbial community from a polycyclic aromatic hydrocarbon

(PAH) contaminated site. After 16s DGGE profiling and sequencing of individual bands, they managed to identify Alteromonas as the key player in PAH degradation.

Teske et al., (1996) analysed and identified the components of a mixed culture capable of sulphate reduction as Desulphovibrio and Arcobacter. This allowed them

51 to subsequently design specific culture conditions to separate and isolate both strains in pure cultures.

4. Comparison of different microbial communities

DGGE has been widely used to study the variance in microbial populations from different environments/treatments. Gurtner et al., (2000) compared bacterial diversity associated with two different biodeteriorated wall paintings and favoured

DGGE because cultivation strategies require far more sample material than could be obtained from art objects. Cosgrove et al., (2007) compared fungal communities on the surface of biodeteriorated PU with native soil populations when PU coupons were buried under two sandy loamy soils with different pH-acidic (5.5) and basic

(6.7). Microbial community on the surface was less diverse and found to be a subset of the soil community.

5. Monitoring the persistence of specific members of microbial community

DGGE can also be used to study the persistence of particular organisms in the environment, for example when one or more species are added to a sample for bioaugmentation. Cosgrove et al., (2010) used DGGE to monitor the effect of bioaugmentation and biostimulation by fungi on the biodegradation of PU coupons.

Specific PU degrading fungi in augmented soil were found to persist both in the soil and on the surface of PU four weeks after their introduction.

6. Association of organism with a process of interest

DGGE is a cost effective and sensitive technique to imply the association of particular organisms with certain processes. Bonito et al., (2010) used DGGE to study the diversity and identification of fungi associated with organic municipal waste composting. Distinct communities with dominant populations were found at each composting phase. Enwall and Hallin (2009) studied bacterial denitrifier community composition in agricultural soil based on nos Z, encoding the gene for nitrous oxide reductase.

1.6.2.4 Terminal Restriction Fragment Length Polymorphism (TRFLP)

Another culture independent technique commonly used to study microbial diversity is TRFLP. Following DNA extraction from the community of interest, fluorescently labelled primers are used to amplify a region of interest from the community. Following amplification, PCR amplicons are digested with one or more restriction enzymes followed by resolution via capillary electrophoresis. In this way, different community members are discriminated as different TRF sizes in a profile

(Liu et al., 1997; Thies 2007). A schematic representation of TRFLP is shown in

Figure 1.5.

TRFLP is a robust technique, which yields fairly accurate results since an internal sized marker is mixed with the digestion products. This technique has gained popularity since the time it was introduced because of its reproducibility and high fidelity procedures and it yields a greater number of OTUs than many other PCR based fingerprinting methods (Osborn et al., 2000), but it does require careful data interpretation in order to accurately characterize sample diversity (Courtney et al.,

2012).

One approach for TRFLP is to generate profiles from different communities or populations exposed to different environmental conditions or treatments. TRFLP analysis has been used to characterize communities of fungi (Wu et al., 2007; Lord et al., 2002; Dickie & FitzJohn 2007), bacteria (Liu et al., 1997; Kaplan et al., 2001;

Blackwood et al., 2003), archeae (Wu et al., 2006) in numerous environments.

TRFLP has been used to investigate spatial and temporal changes in microbial populations in a range of environments including the human gut (Li et al.,

2007); water (McMahan et al., 2012); forests (Dickie et al., 2002; Burke et al., 2012); agricultural soil (Lukow et al., 2000) and grassland soils (Brodie et al., 2003; Nunan et al., 2005; Robinson et al., 2009); land use gradients (Bissett et al., 2011); different soil types (Singh et al., 2006; Smalla et al., 2007) and compost (Tiquia, 2005).

Intensive surveys of general fungi (Dickie et al., 2002; Brodie et al., 2003) and arbuscular mycorrhizas (Bainard et al., 2011) have also been done. A number of studies have used TRFLP to analyse the effect of contamination on environmental microbial populations including antherecin contamination (Wang et al., 2011), PAH contaminated soil (Muckian et al., 2007), its indirect effect on rhizosphere (Kawasaki et al., 2011).

The surveys mentioned above, have reported the change in microbial community composition across time and space and to develop a better understanding for establishing a link with natural or artificial adaptation in the environment.

Figure 1.5: Schematic presentation of TRFLP principal. Here community DNA is amplified with a fluorescently labeled primer then community amplicons are digested with a restriction enzyme. These digested products are separated in a capillary of DNA sequencer that detects the size of each fragment along with intensity of fluorescence. The fluorescent detector only recognizes the fragments with fluorescent dyes rest of the fragments are ignored. The resultant is an electropherogram, which is an intensity (y-axis) plot of the different sized fragments (x-axis).

Another approach has been to use TRFLP to identify individual species by comparing TRF size against a species database (reviewed by Dickie and FitzJohn,

2007). However, this approach has limitations as electrophoretic mobility of DNA fragments varies when labelled with different fluorophores (Tu et al., 1998) or have a different sequence composition (Kaplan et al., 2001) and only a small percentage of sequences for 16S, 18S or ITS are archived in databases. Misidentification may result from the fact that known sequences in the database have the same sequence polymorphisms as unknown sequences in the sample (Blackwood and Buyer, 2007).

While TRFLP has a number of advantages, as a PCR based technique it has limitations similar to DGGE with regard to DNA extraction, PCR inhibition, PCR efficiency and other as outlined previously (Von Wintzingerode et al., 1997). In addition, erroneous combination of either real peaks or “noise” and loss of rare taxa

(Courtney et al., 2012; Dickie et al., 2002), the potential for more than one species to share the same TRF and generation of multiple TRFs from the same species and inefficient restriction digestion can all lead to errors (Dunbar et al., 2000; Avis et al.,

2006; Dickie and FitzJohn, 2007; Smalla et al., 2007).

1.6.2. 5 Comparison of DGGE and TRFLP

A number of studies have made comparisons between DGGE and TRFLP.

DGGE is more convenient and inexpensive while, TRFLP requires more steps and is more expensive. However, DGGE is less sensitive than TRFLP and uses amplicons that are generally (Horz et al., 2001) <500bp restricting the sequence information obtained which can limit phylogenetic identification (Horz et al., 2001; Muyzer et al.,

1993). Sample number is restricted by gel size, making comparisons between a large

56 number of samples difficult (Nunan et al., 2005; Smalla et al., 2007). The lower sensitivity of DGGE compared to TRFLP means that changes in minor components of the population can go undetected (Tiedje et al., 1999). For example, bacterial diversity associated with the roots of submerged rice plants was far higher using

TRFLP compared to DGGE (Horz et al., 2001). Smalla et al., (2007) studied the population diversity of four agricultural soils and found similar clustering with

TRFLP and DGGE but a higher level of species diversity with TRFLP and a similar conclusion was reached by Moeseneder et al., (1999) in a study of marine bacterial plankton.

1.6.2.6 454 pyrosequencing

454 pyrosequencing is a relatively new technique that enables sequences of all amplified PCR products to be analysed from several samples and has largely replaced clone libraries previously employed for metagenomic analysis. It is a high throughput sequencing technique particularly useful in the detection of rare community members that may represent <1% of the biota (Ronaghi, 2001).

The 454 Pyrosequencing takes advantage of DNA captured beads that contain single stranded template complimentary to the adapter region of amplicon

(ssDNA) which is amplified to millions of copies in an oil emulsion PCR (emPCR).

Beads are distributed on a PicoTiter Plate with 1.6 million wells that each can accommodate a single bead and additional reagents including luciferase, polymerase and ATP sulfurylase. Continuous introduction of each of four dNTPs over the

PicoTiter Plate and incorporation of nucleotide releases pyrophosphate.

Pyrophophate serves as the substrate for luminescence reaction that is recorded with

57 a camera. The record of intensity of incorporation of nucleotide is a flow-gram, like a chromatogram, that reports the order of A, T, G and C residues from DNA sequencing templates. The intensity of light (flow-gram values) corresponds to the homopolymer length for that nucleotide. A schematic representation of pyrosequencing is shown in Figure 1.6.

Buée et al., (2013) employed 454 pyrosequencing to assess the diversity of fungi in forest soils and reported high fungal diversity, especially in OTUs belong to basidiomycetes and ascomycetes. Recently, Gannes et al., (2013) used pyrosequencing to analyse the diversity of bacteria in a composting environment.

They rejected the previous belief that Bacillus was the prominent genus in all stages and reported new species were present in each composting phase.

Figure 1.6: Schematic representation of 454-pyrosequencing principle. Beads with primer to ssDNA are in the 1:1 ratio goes through emPCR that generates beads with millions of copies of ssDNA. The beads, along with Luciferase and PCR reagents are then transferred to Picotitre plates. Free dNTPS are added in cycles and with the addition of nucleotide in to the newly formed DNA strands, ATP is generated, which participate in conversion of Luciferin to Oxyluciferin. With this conversion, emission of light takes place that generates a flowgram which is translated into sequences.

1.6.3 Community diversity analysis

Assemblages of populations within a community can be described by several mathematical indices. These indices, (species diversity indices), describe the species richness and abundance of species within the community. Species diversity indices relate the number of species and the relative importance (proportion) of individual species. Two major parameters for measuring species diversity are species richness and evenness. Species richness measures the number of species within a community but not how many individuals of a species are present, here a particular estimate called Chao I was used. Evenness measures the proportion of individuals among the species present. A widely used measure of diversity is the Shannon-

Weaver index. This index is sensitive to both species richness and relative species abundance (Atlas & Bartha 1998). Rarefaction curve is also an important estimate to assess the species richness for a given number of samples.

When samples have high dimensionality because of the large number of variables per sample, it makes visualization of samples difficult and limits exploration of data. Terminal Restriction Fragment analysis of the microbial communities generates hundreds of TRFs and pyrosequencing generates 1000s of sequences per sample. Such high dimensionality limits visualization, making analysis difficult. Principal component analysis (PCA) is a mathematical algorithm that reduces the dimensionality of the data while keeping most of the variation in the data set. Data are arranged along first principal component, means maximal variation among the variables. Then the second principal component is drawn in the direction uncorrelated to the first component along which the samples show largest variation.

This makes it possible to visualize and assess similarities and differences between samples and determine whether samples can be grouped or not (Ringnér, 2008; Park, et al., 2006).

1. 7 Physico-chemical tests for polymers

There are number of physico-chemical tests available to assess the biodegradation of PU. These tests assess the structural and chemical changes in polymer as result of biodegradation and biodeterioration. The most important tests are tensile strength measurement and differential scanning calorimetry. Tensile strength is the measurement for brittleness; it’s the maximum stress a material can withstand while being stretched before breaking. Differential Scanning Calorimetry

(DSC) is a thermal analysis to study the structural changes in polymer. It is to measure the amount of heat absorbed or evolved from sample under isothermal condition. Both polymer and reference are heated at the same rate. The amount of extra heat absorbed by polymer sample is with reference to the reference material studied. DSC helps to assess the crystallinity, melting and glass transition (Tg) temperature. Tg of a polymer is the “temperature at which molecular segment begin to rotate” (Sauders & Frisch, 1964).

1. 8 Summary

In summary, polyurethanes are xenobiotic plastics that have a broad range of applications with production exceeding 3 Million tonnes per annum. Due to constrictions and environmental concerns surrounding landfill, alternative strategies are actively being pursued for managing waste streams that include recycling, incineration and biodeterioration. Composting is a microbially driven process that undergoes defined temperature transitions with an initial increase in temperature due to rapid metabolism leading to enrichment for thermophiles and in recent years has been widely commercialised for the disposal of domestic green and food waste. PU’s have been reported to be susceptible to microbial attack, especially by fungi, however, the ability and role of fungi in PU degradation in composting environments has not previously been investigated. In this study, the potential of the composting process to biodgrade PU and the role of fungi in this process were investigated using both conventional culturing and molecular non-culture based approaches.

1. 9 Experimental aims

The central theme of this thesis was to examine the biodegradation of PU under composting conditions. This will be addressed in three separate papers each addressing a different aim.

Aims:

 To assess the rate of degradation of PU buried in compost at different temperatures for a period of 12 weeks, to isolate potential fungal degraders and to monitor the temporal changes in the fungal community colonising the surface of PU

 This aim was addressed in chapter two “Biodegradation of PU under composting conditions at different temperatures”. This paper compared the biodegradation of PU buried in compost when incubated at 25°, 45° and 50 C.

Temporal changes in the fungal community on the surface of PU was investigated using both conventional and molecular approaches.

 To evaluate the potential of a widely employed commercial composting system to biodegrade PU and to investigate and identify fungi associated with this process

 This aim was addressed in chapter 3 “Biodegradation of polyurethane during commercial composting”. This paper examined the rate of degradation of PU during maturation of compost in a commercial composting system. Temporal succession of the fungal community on the surface of PU was also monitored using DGGE and

TRFLP and the composition of the community was determined by 454 pyrosequencing.

 To understand the impact of PU contamination on the native fungal population of compost at different temperatures for a period of 12 weeks

 This aim was addressed in chapter 4 “ Effect of polyurethane on fungal compost communities”. This paper assessed the impact of the introduction of PU on the fungal population in compost using both conventional and molecular techniques.

1.10 Alternative format

The thesis is being presented in the alternative format in accordance with the rules of the University of Manchester. The three results chapters presented herein are in manuscript form in the style suitable for a peer-reviewed journal “Bioresource

Technology” to ensure a cohesive body of work. Below are the details of each manuscript, its intended journal and contribution of each author to the work presented.

Chapter 2: Biodegradation of PU under composting conditions at different temperatures.

Authors: Urooj Zafar, Ashley Houlden and Geoff Robson

Intended journal: Applied and Environmental microbiology

Contribution of authors: the work described in the manuscript represents experiments to which I contributed solely. The community pyrosequencing and DNA sequencing of isolates was done by Genomic centre, University of Liverpool and DNA sequencing facility at the University of Manchester, respectively. Dr Ashley Houlden provided help with TRFLP and pyrosequencing analysis. My supervisor Dr Geoff Robson provided advice and guidance on all experimental work. As first author of this paper I was also fully responsible for writing the text of the manuscript. The first draft was produced by myself, which my supervisor then reviewed and commented on. These comments were then synthesised by me into the final version presented here.

Chapter 3: Biodegradation of polyurethane during commercial composting.

Authors: Urooj Zafar, Ashley Houlden, Alan Heyworth, Alberto Saiani and Geoff Robson

Intended journal: Bioresource Technology

Contribution of authors: The work described in the manuscript represents to experiments of which I contributed solely. The community pyrosequencing and DNA sequencing of isolates was done by Genomic centre, University of Liverpool and DNA sequencing facility at the University of Manchester, respectively. Dr Ashley Houlden provided help with TRFLP and pyrosequencing analysis. Dr Alberto Saiani helped in the analysis of DSC results. My supervisor Dr Geoff Robson provided advice and guidance on all experimental work. As first author of this paper I was also fully responsible for writing the text of the manuscript. The first draft was produced by myself, which my co-authors and supervisor then reviewed and provided comments. These comments were then synthesised by me into the final version presented here.

Chapter 4: Effect of Polyurethane on fungal communities.

Authors: Urooj Zafar and Geoff Robson

Intended journal: Environmental microbiology

Contribution of authors: The work described in the manuscript represents experiments to which I contributed solely. Sequencing facility at the University of Manchester did DNA sequencing of isolates. My supervisor Dr Geoff Robson provided advice and guidance on all experimental work. As first author of this paper I was also fully responsible for writing the text of the manuscript. The first draft was produced by myself, which my supervisor then reviewed and provided comments. These comments were then synthesised by me into the final version presented here.

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Chapter 2: Fungal community associated with biodegradation of polyester polyurethane buried under compost at different temperatures

Urooj Zafar, Ashley Houlden and Geoff D. Robson

Faculty of Life Sciences, Michael Smith Building, University of Manchester, Manchester M13 9PT, UK

2.1 Abstract Plastics form an essential role in the modern world due to their low cost, insulating properties, light weight and durability. However, accumulation of plastic waste in the environment is responsible for unique and long lasting effects and managing waste plastics is increasingly expensive. Polyurethanes (PU’s) are a high volume plastic that make up ca. 7% of the total plastic production in Europe with demand increasing every year. PU’s are heteroplymers and polyester PU’s in particular have been extensively reported as susceptible to biodegradation in the environment, particularly by the fungi. In this study, we investigated the impact of composting on PU’s as composting is a microbially rich process that is increasingly being used for the processing of green and food waste as an economically viable alternative to landfills.

PU coupons were incubated for 12 weeks in fresh compost from the maturation stage at 25°, 45° and 50°C to emulate the mesophilic, maturation and thermophilic stages of the composting process. Incubation at all temperatures caused significant physical deterioration of the polyester PU coupons at all temperatures and was associated with extensive fungal colonisation and loss in tensile strength. By contrast, no loss in tensile strength was detected in polyether PU coupons over the 12 week incubation period. A number of fungal species were isolated from the surface of polyester PU coupons at 25°C while only Aspergillus fumigatus and Thermomyces lanuginosus was recovered at 45° and 50°C respectively. TRFLP and pyrosequencing of the fungal communities on the PU surface and in the surrounding compost revealed that the population on the surface of PU was different from the community observed in compost suggesting enrichment and selection on the surface of PU coupons. The most dominant fungi identified from surface of PU coupons by pyrosequencing were

Fusarium solani and Bionectria ochroleuca at 25°C, while at both 45° and 50°C

Candida ethanolica and an unidentified fungal clone were dominant. This preliminary study suggests that the composting process has the potential to biodegrade PU waste if optimised further in the future.

2.2 Introduction

Polyurethanes are synthetic plastics with a wide range of applications in the medical, automotive and industrial fields (Kaplan et al., 1968; Howard, 2012).

Polyurethanes are formed by the condensation of polyisocyanate and polyols and are heteropolymers (Dombrow, 1956; Krasowska et al., 2012). 7% of the total plastics used in Europe (47 Million tonnes) in 2011 were polyurethanes (Plastics – the Facts

2012). A large proportion of plastic waste is directed to landfill sites; however its low degradation rates, scarcity of landfill sites and growing water and land pollution problems require alternative waste management strategies to be developed

(Jayasekara et al., 2005; Oehlmann et al., 2009). However, polyester PUs are known to be vulnerable to microbial attack (Morton & Surman 1994) as they contain ester linkages within the backbone of the polymer that are naturally vulnerable to esterases as first reported by Darby & Kaplan (1968). In contrast, polyether PU’s, which contain ether, within the polymer backbone are reported to be far more recalcitrant

(Darby & Kaplan, 1968; Pommer and Lorenz, 1985)

The development of large-scale commercial composting facilities for the treatment of green and food waste has been helpful in reducing landfill and meeting recycling goals (Epstein, 1997). Composting is a managed self-heated, aerobic process that controls the biological decomposition and transformation of biodegradable materials into a humus-like substance (compost). It is a natural process that results in the production of CO2, H2O, minerals, and stabilized organic matter

(Arvanitoyannis & Bosnea, 2001; Shah, et al., 2008). The major advantages of composting are that it is rapid, relatively inexpensive and environmentally friendly. It is a natural degradation process similar to degradation in the soil but during

83 composting a considerable amount of heat is produced as a result of microbial respiration that accelerates the rate of deterioration (Kim & Kim 1998; Shah et al.,

2008). Thus, temperature is the single most important factor affecting the microbial population, growth and activity (Epstein, 1997). Microbiological development progresses in defined temperature phases. The initial rise in temperature from ambient to 45°C is called the mesophilic phase. The thermophilic phase is dominated by thermophilic microbes and temperature can reach 70°C or higher. Once readily metabolised substrates have been utilized, microbial activity and temperature decreases and eventually approach ambient temperature (Ishii et al., 2000). Extreme changes in temperature makes composting an ecologically complex system and distinct diverse microbial populations can be recovered at different phases.

Previously, fungi have been shown to be the dominant microorganisms involved in the biodegradation of polyester PU when buried in soil (Barratt et al.,

2003; Cosgrove et al., 2007; 2010) and a number of fungal species have been isolated and identified that are capable of degrading PU (Mathur & Prasad 2012; Russell et al., 2011; Cosgrove et al., 2007; Barratt et al., 2003; Pathirana & Seal 1984; Darby &

Kaplan 1968). In this chapter we investigated the potential of the composting process to deteriorate PU by comparing the rate of biodegradation when buried in compost at different temperatures representing the mesophilic and thermophilic stages. We report that polyester PU undergoes significant degradation under composting conditions while polyether PU appeared largely unaffected. The results indicate that composting may therefore represent an alternative waste management route for polyester PU.

2.3 Materials and Methods

2.3.1 Media composition

2.3.1.1 R2A

R2A (pH 7.2, Difco USA) was used for the recovery and enumeration of bacteria from compost and surface of degrading polyurethane. It was prepared according to manufacturer’s instructions.

2.3.1.2 Potato dextrose Agar

PDA (pH 5.6, Oxoid UK) was used for the recovery and enumeration of fungi from compost and surface of degrading polyurethane. This medium was also used for sub- culturing to isolate pure fungal strains and for routine fungal culture maintenance. It was prepared according to manufacturer’s instructions.

2.3.1.3 Compost Extract

Compost extract was used for testing the effect of immersion and the presence of soluble compost components on the tensile strength of PU and in compost extract agar for enumeration of fungal colony forming units (CFU). Compost extract was prepared according to Alef & Nanniperi (1995). Dried compost (500 g, The Compost

Shop, Orrel Hill Lane, UK ) was mixed with 1 L of distilled water by shaking at 150 rpm for 30 min, autoclaved at 121°C for 30 min, filtered through two layers of J- cloth. After adjusting the volume of the filtrate to 1 L with sterile distilled water, filtrate was sterilised at 121°C for 20 min. Compost extract agar was prepared by adding glucose (1 g), peptone (0.2 g), yeast extract (0.1 g), K2HPO4 (0.4 g),

MgSO4.7H2O (0.05 g), sodium deoxycholate (0.5 g/l), compost extract (100 ml) and agar (15 g) to 900 ml of distilled water and autoclaved at 121°C for 15 min.

2.3.1.4 Polyurethane agar (PUA)

Polyurethane agar (PUA, pH 6.7) was used to recover polyurethane degrading microorganisms from compost and from the surface of polyurethane coupons. PUA

(Crabbe et al., 1994) was prepared by using impranil as the sole carbon source.

Impranil is an anionic aliphatic polyurethane dispersion free from organic solvents and emulsifiers. Impranil DLN is a dispersion of polyester PU and Impranil DLU is a dispersion of polyether PU. PUA was prepared by adding 20 ml of 50X phosphate

-1 -1 buffer (50 gl K2HPO4 and 25 gl KH2PO4) to 950 ml distilled water containing 1 g

(NH4)2SO4 0.02 g yeast extract, 7.5 ml impranil DLN, 0.5 g sodium deoxycholate (to induce compact colony growth) and 15 g agarose and sterilised at 121°C for 20 min.

-1 After cooling to 50°-55°C, 10 ml of filter sterilised (0.22 µm) MgSO4.7H20 (50 gl )

-1 and 1 ml of filter sterilised (0.22 µm) trace elements solution (MnCl2.4H2O 2 gl ,

-1 -1 -1 CuCl2.2H2O 28 mgl , ZnCl2 22.0 mgl , CaCl2.2H2O, 27 mgl , Na2Mo4.2H2O 26

-1 -1 mgl and FeCl2.6H2O; 150 mgl ).

2.3.2 Fabrication of polyurethane coupons

Polyester and polyether polyurethane (PU) coupons of 16 x 16 x 1 cm (total surface area 518.4 cm2) were fabricated from PU beads (Ellastollan® 685 A10 and

Ellastollan ® 1180 A10 respectively) by melting at 180°C at 8x105 Pascal in a compression moulding machine (Moore, Birmingham, UK). Dumb-bells (total length

5 cm, width at the end 1.9 cm with 19 cm gauge length) were cut from the coupons

86 using a molder cutter (Wallace Test equipment, UK). Rectangular coupons 4 x 4 cm were cut with scalpel blades.

2.3.3 Burial of polyurethane coupons in compost

Fresh mature compost (temperature ca. 65°C, The Compost Shop, Orrel Hill Lane,

UK) and soil (Online turf, Lancashire, UK) were sieved through a 4 mm mesh prior to use. Percentage water holding capacity (WHC) of the compost and soil was measured following the modified protocol of Alef and Nannipieri (1995). Five replicates (10 g) were dried at 55°C to constant weight. Percentage WHC was calculated using the following equation:

( ) ( )

( )

Moisture content of the compost was determined according to Barratt et al., (2003).

Five replicates (10 g) were weighed (fresh weight) and then dried at 55°C to constant weight (dried weight). Percentage moisture content was calculated using the following equation:

( ) ( )

( )

Percentage moisture content of compost and soil was adjusted to 40% and 20% respectively, by the addition of sterile distilled water.

Plastic containers (25x20x10cm) were cleaned using 70% (v/v) ethanol before filling

2/3 of height with compost. Dumb-bells and rectangular coupons were weighed and

87 buried vertically at 1cm apart such that the tops of the coupons were approximately 3 cm below the surface. Samples were recovered after 1, 4, 8 and 12 weeks. Moisture content of the compost and soil was monitored periodically by weighing the containers every 2 days and maintained by the addition of sterile water using a plant spray.

2.3.4 Recovery of polyurethane samples and enumeration of microbes

Polyurethane samples were recovered from the compost using forceps and microbial biomass recovered from the rectangular coupons according to Cosgrove et al.,

(2007). Briefly, rectangular coupons were first agitated in sterile PBS (Phosphate buffer saline) pH 7.4 for 3 min manually, placed into a Petri dish (90 mm) with 20 ml

PBS and biomass scraped from both sides of the coupons using a sterile blade and the suspension agitated briskly for 5 min. An aliquot of 1 ml was used to enumerate the total microbial viable count by plating onto compost extract agar [containing 50

µg/ml chloramphenicol (to suppress bacterial growth and enable fungal enumeration) and R2A (Oxoid, UK, section 2.3.1.1) and 250 µg/ml nystatin (to suppress fungal growth and enable bacterial enumeration)] following serial dilution in PBS. PU degrading microbes were enumerated by plating onto polyurethane agar (PUA, section 2.3.1.5, Crabbe et al., 1994) supplemented with chloramphenicol (50 µg/ml) or nystatin (250 µg/ml) and containing 0.75% (v/v) impranil (a polyurethane dispersion, Bayer, Newbury, UK) as a sole carbon source. Plates were incubated for up to 1 week at the same temperature the PU samples had been incubated. As fungal colonies on PUA were difficult to differentiate morphologically, individual colonies were transferred onto potato dextrose agar (PDA), incubated for up to 1 week at the

88 same temperature the PU samples had been incubated and grouped into different morphotypes.

2.3.5 Microscopic analysis of PU coupons

Coupons recovered after 12 weeks were washed with sterile distilled water followed by 70% (v/v) ethanol, air dried and observed under environmental scanning microscopy (ESEM, FEI Quanta 2000 Netherlands) to visualize the surface of the coupons. ESEM examination of coupons was done at low pressure (Torr) and room temperature with distance covered from 1000 to 5 µm scale range. Coupons were directly put on the sample stage in ESEM without processing or dehydrating.

Microscopy was done with the assistance of Dr. Patrick Hill, School of Chemical

Engineering and Analytical Science, University of Manchester.

2.3.6 Tensile strength and weight loss determination

The tensile strength of PU dumb-bells (5 replicates) was determined using a Tinius

Ohlsen H5KT-0586 (UK) with cross head speed of 1.5 cm/min. Tensile strength was measured as the maximum load (N) required to break PU dumb-bells. In addition, the weight of PU dumb-bells was determined weekly after first washing thoroughly in distilled water.

In addition to unburied controls, PU coupons were also autoclaved at 121°C for 5 min and completely immersed in filter sterilized (0.22 µm) compost or soil extract in sterile 7 ml tubes and incubated at 25°, 45° and 55°C for 12 weeks.

2.3.7 Extraction, amplification and purification of genomic DNA from the isolated PU degrading fungal colonies

Genomic DNA was extracted from the mycelium of fungal colonies according to

Feng, et al., (2010). Mycelium/ spores (ca. 20 mg) were collected from the surface of confluent PDA cultures with a sterile toothpick and placed into 1.5 ml centrifuge tubes containing 0.5 g of 0.5 mm diameter glass beads. After addition of 0.65 ml lysis buffer (100 mM Tris-HCl, pH 8.0; 50 mM EDTA, pH 8.0; 1% (w/v) SDS; 10

μg ml−1 RNase A), tubes were homogenized twice for 30 sec and centrifuged for 2 min at 13000 rpm. After centrifugation, 500 μl of supernatant was transferred into a new tube containing 100 μl of potassium acetate buffer (3.0 M, pH 5.5). The tube was inverted several times and centrifuged for 2 min at 13000 rpm, 500 μl of supernatant was transferred into a new tube containing 500 μl of isopropanol, inverted several times and centrifuged for 2 min 13000 rpm. The supernatant was removed and the DNA pellet washed with 750 μl of 70% (v/v) ethanol. After centrifugation for 30 s, ethanol was removed and the DNA pellet air-dried for 5–10 min. The DNA was dissolved in 50 μl sterile distilled water and stored at -20°C until required.

Isolates were identified according to Webb et al., (2000) using the Internally

Transcribed Region (ITS) of rDNA amplified using the fungal universal primers

ITS1 (5’-TCCGTAGGTGAACCTGCGG-3’) and ITS4 (5’-

TCCTCCGCTTATTGATATGC-3’) (White et al., 1990). The reaction mixture contained genomic DNA (20-100 ng), primers (0.2 µM), MgCl2 1.5 mM, 1x NH4 reaction buffer, 200 µM of each of dNTPs and 1U Taq Polymerase (Bioline UK).

The PCR consisted of initial denaturation 94°C for 3 min, 35 cycles, denaturation at

94°C for 1 min, annealing at 56°C for 1 min and extension at 72°C for 1 min. Final extension was 72°C for 3 min. PCR amplicons varied between ca. 500-575 bp depending on the species. Amplified PCR products were visualized by electrophoresis through a 1.2% (w/v) agarose gel in the presence of TAE buffer (40 mM, pH 8.6) with Hyperladder I (Bioline UK) at 100V for 1 h and PCR products purified using the QIAquick ® PCR purification Kit (Qiagen UK) according to the manufactures instructions.

2.3.8 ITS rDNA sequencing and identification

Purified samples were sequenced by the in house sequencing facility (Faculty of Life

Sciences, University of Manchester, UK) using an ABI Prism® 3100 Genetic analyser (Applied Biosystems USA). Sequencing results were viewed using FinchTV v1.4.0 software (Geospiza Inc.). Nucleotide sequences were interrogated using the

BLAST (Basic Local Alignment search Tool) algorithm at the National Center for

Biotechnology Information website (www.NCBI.nlm.nih.gov) and phylogenetic trees were compiled by the Mega 5 alignment tool and CLUSTALW programme with 500 bootstrapping value.

2.3.9 Extraction and amplification of community genomic DNA

Genomic DNA was extracted from biomass obtained from the surface of PU coupons and from compost using the Powersoil DNA isolation kit (MO BIO Laboratories,

USA) according to the manufacturer’s instructions except that during the lysis step tubes were homogenized twice for 30 s at 5000 rpm. The concentration of eluted

DNA was measured using a NanoDropTM 1000 and samples stored at -20°C until required.

2.3.10 Analysis of the fungal community by TRFLP analysis

For fungal community analysis by TRFLP, the fungal ITS1-5.8s-ITS2 rDNA region was amplified using the fluorescent labelled primers, ITS5-FAM (fam-

GGAAGTAAAAGTCGTAACAAGG) and ITS4- HEX (hex-

TCCTCCGCTTATTGATATGC). A 50 µl PCR mix was made using 1xNH4 buffer,

1.5 mM MgCl2, 200 µM of each dNTPs, 0.5U Taq polymerase (Bioline UK), 0.2 µM primers and BSA 100 µg/ml (New England Biolabs UK). DNA template was Ca. 50-

100 ng per PCR reaction. The PCR regime consisted of: initial denaturation at 94°C for 10 min, then 35 cycles of 94°C for 1 min, 54°C for 1 min and 72°C for 1 min, with a final extension 72°C for 10 min. Amplified DNAs were verified by electrophoresis through a 1.5% (w/v) agarose gel with a Hyperladder I (Bioline UK) at 100V for 1 h. Three PCR amplicon replicate samples were pooled and purified by ethanol precipitation. Ethanol (100%) was added to the pooled DNA samples in 2:1 ratio and incubated overnight at -20 °C for better yield. Tubes were centrifuged at

13000 rpm for 30 minutes at 0 °C and the supernatant discarded. Ice cold 70% (v/v) ethanol was added to the pellet and gently mixed by inverting the tubes several times and then centrifuged at 13000 rpm at 0°C for 10 min. The supernatant was discarded and the pellet left to air dry overnight at room temperature. The pellet was dissolved in 10 µl of sterile DPEC water and DNA concentration measured using a

NanoDropTM 1000 (Thermofisher Scientific Inc., USA). To produce a mixture of variable length end-labelled ITS rDNA fragments, PCR products (1.5 µg) were digested with 0.5 U Hhal (Fermentas, UK) at 37°C overnight in Tango buffer (10 µl).

Digested products (0.5 µl) were mixed with 9.25 µl Hi-Di formamide (ABI, UK) and

0.25 µl GS500LIZ (ABI, UK) in a 96 well PCR plate and products separated and

92 analysed in-house on an ABI Prism® 3100 Genetic analyser (Applied Biosystems

USA) (Kawasaki et al., 2011).

The size of the fragments was determined using Peak Scanner™ Software Version

1.0 (Applied Biosystems), using peak height detection of 50 fluorescent units The output was further analyzed with online T-Align programme

(http://inismor.ucd.ie/~talign/) to generate a consensus profile of TRFs sizes between the technical duplicates and to compare the profiles between the samples (Smith et al., 2005). Shannon index (Ĥ) and evenness (e) were measured for each T-RFLP according to Tiquia (2005). Principal Component Analysis was employed to cluster the samples based on Boolean characters (0, 1), which corresponds to the presence or absence of TRFs from each TRFLP pattern. Principal Component Analysis was performed using MVSP version 3.13g (copyright © 1985-2003 Kovach computing services).

2.3.11 454 Pyrosequencing

Fusion primers (Table 2.1) were designed with an adapter (blue) and key “TCAG”

(red) sequence with ITS5 and ITS4 primer sequences (Figure 2.1). Forward primers also had one of 10 bp unique Roche multiplex identifiers (MID, Table 2.1), which were used to tag PCR amplicons from each sample. Sequence of forward primer

(Primer A): 5’-CCATCTCATCCCTGCGTGTCTCCGACTCAG-{MID}-

{GGAAGTAAAAGTCGTAACAAGG}-3’.Reverse primer (Primer B): 5’-

CCTATCCCCTGTGTGCCTTGGCAGTCTCAG-{

TCCTCCGCTTATTGATATGC }-3’ (Rosche, 2011a).

PCR was conducted using the High Fidelity PCR system (Roche, USA). 50 µl reaction mixture contained 1xNH4 buffer, 1.5 mM MgCl2, 200 µM of each dNTPs, 1

U polymerase , 0.2 µM primers (HPLC purified) BSA 100 µg/ml (New England

Biolabs), 2 µl DMSO an ca. 50-100 ng of DNA template per PCR reaction. The PCR regime used was similar to TRFLP, except 30 cycles were used to reduce chimera formation. PCR products were verified by running on a 1% (w/v) agarose gel (100 V for 45 min) containing 0.005% ethidium bromide, visualised under a UV transilluminator, and products excised from the gel (ca. 575-700 bp expected size range) using a sterile rectangular blade, DNA was extracted using Gel extraction kit

(Qiagen, UK) according to the manufacturer’s instructions. Products were further purified through column purification (Qiagen, UK) and amplicons from three triplicate samples pooled in equal concentration to give a final concentration of

10ng/µl. Pooled samples were sent for 454 Titanium platform pyrosequencing to the

Centre for Genomic Research, University of Liverpool, UK.

2.3.12 Bioinformatics and statistical analysis

Sequence data processing was performed with MacQIIME version 1.6.0 following the procedure similar to that of Caporaso et al., (2010). After splitting the libraries and denoising using default settings (Reeder and Knight, 2010), sequences were grouped into OTUs at a similarity level of 97% using uclust (default) against

UNITE/QIIME 12_11 ITS reference database, any sequence which did not hit the reference database were subsequently clustered denovo. A representative set of

OTUs was generated and the taxonomy of each set was assignment of OTUs was performed using same database (UNITE+INSD). OTUs defined as 97% sequence

94 similarity threshold were used for rarefaction curve, PCA and beta diversity analysis

(chao1 and Shannon- weaver index) using QIIME.

2.3.13 Statistical analysis

To determine the statistical significance, data were subjected to Analysis of Variance with the significance threshold set at a P value of 0.05 (JMP basic version 9.0.2

Figure 2.1: Schematic representation of primers designed for unidirectional reads of amplicons from community genomic DNA. There was an adapter (blue) and key “TCAG” (red) sequence, forward primers also had a unique Roche multiplex identifiers, MID (yellow). (Adapted from Roche pyrosequencing guidelines, accessed June’2013).

Table 2.1: Sequences of the 10 MIDs used to tag PCR amplicons from each sample for 454 pyrosequencing.

ID MID Sequence ID MID Sequence

MID1 ACGAGTGCGT MID6 ATATCGCGAG

MID2 ACGCTCGACA MID7 CGTGTCTCTA

MID3 AGACGCACTC MID8 CTCGCGTGTC

MID4 AGCACTGTAG MID10 TCTCTATGCG

MID5 ATCAGACACG MID11 TGATACGTCT

(Adapted from Roche pyrosequencing guidelines, accessed June 2013).

2.4 Results

2.4.1 Compost chemical analysis

Moisture content of soils has previously been found to have a marked influence on fungal growth and degradation of PU (Barratt et al., 2003). Water holding capacity was determined to be 71% and 40% and the moisture content was ca. 30% and 15% for compost and soil respectively. Sterile distilled water was added to increase moisture content to 40% and 20% in compost and soil respectively and maintained during the 12 weeks incubation period by periodically adding sterile distilled water to replace that lost through evaporation.

2.4.2 Visual changes to PU coupons during compost burial at different temperatures

To determine physical deterioration of PU coupons when buried in compost,

PU coupons were buried and incubated at 25°, 45° and 50°C. Coupons were recovered after 4, 8 and 12 weeks (Figure 2.1). For comparative purpose, coupons were also buried in soil at 25°C. Initially PU coupons were transparent, flexible and had a smooth surface. At 25°C after incubation in soil or compost, the surface of PU coupons appeared rough after 4 weeks, visible cracks appeared after 8 weeks that increased in number and deepened after 12 weeks. White patches of mycelial growth were also clearly visible on the PU surface after 8 weeks and became extensive after

12 weeks (Figure 2.1a & b). At 45° and 50°C, PU coupons became opaque after 4 weeks and fungal mycelium covered ca. 50-90% of the surface. Coupons were completely opaque after 8 weeks and by the end of 12 weeks, complete discolouration and surface cracking were observed (Figure 2.1c & d).

To further examine the effects of burial on the surface of PU coupons,

ESEM was conducted on PU coupons after 12 weeks of incubation (Figure 2.3).

Cracking of the PU surface was observed at all temperatures but was more extensive at 45° and 50°C (Figure 2.3). By contrast, no physical changes were observed in polyether PU coupons after 12 weeks burial at different temperatures (Appendix 2.7-

Figures A2.1 and A2.2).

2.4.3 Impact of soil and compost burial on the tensile strength of PU

In order to quantify the extent of biodegradation during burial, tensile strength of buried PU dumb-bells were determined periodically following incubation at 25°, 45° and 50°C over 12 weeks. In order to distinguish between microbiological effects and potential effects of chemical components in soil and compost, PU dumb- bells were also immersed in sterile soil and compost extracts (Figure 2.4). Tensile strength of PU dumb-bells buried in compost or soil at all temperatures after 12 weeks showed major loss (p< 0.05) in tensile strength (>75%) compared to dumb- bells immersed in sterile soil or compost extract. There was slight variation (p>0.05) in the tensile strength loss at different temperatures, however, the variation in loss of tensile strength amongst replicate coupons increased with increase in temperature

(Figure 2.4 e). No considerable physical degradation was seen in polyether PU dumb-bells after 12 weeks under any condition (Appendix 2.7-Table A2.1).

To further assess biodegradation, dry weight loss was determined after 12 weeks of burial (Appendix 2.7-Table A2.2). Despite the major reduction in tensile strength and physical disruption of the surface of PU, weight loss in all cases was

<1% and was not significant (p>0.05).

Incubation time (Weeks)

0 4 8 12

Figure 2.2: Physical changes to polyester polyurethane coupons after burial at different temperatures. Physical deterioration of polyester coupons buried in soil at (a) 25°C and compost at (b) 25°C (c) 45° and (d) 50°C for 0, 4, 8 and 12 weeks. The white and cream discolouration visible on the surface is fungal mycelium. The scale bar represents 4 cm.

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Figure 2.3: Effect of compost burial on the surface of PU coupons visualised by environmental scanning microscopy. PU coupons were recovered after 12 weeks of burial in compost at unburied (a), 25°C (b), 45°C (c), 50°C (d) and changes in the surface properties visualised by environmental electron microscopy. Prominent cracks can be seen on the surface of the PU. Scale bar represents 20 µm.

101

a b

80 80 70 70 60 60 50 50 40 40 30 30 20 20

10 10 Tensile Tensile strength (MPa) 0 0 0 1 4 8 12 0 1 4 8 12

c 80 d 80

70 70 60 60 50 50 40 40 30 30 20 20

10 10 Tensile Tensile strength (MPa) 0 0 0 1 4 8 12 0 1 4 8 12 Incubation time (Weeks) e

Figure 2.4: Effect of compost burial on the loss of tensile strength of PU coupons. PU coupons were buried in soil or compost and loss of tensile strength measured over a 12 week period. Control coupons were immersed in sterile soil or compost extract () and buried () in soil at 25°C (a), compost at 25°C (b), compost at 45°C (c), compost at 50°C (d). The results are the means of 5 replicates ±SEM. The variation in replicate PU coupons was higher at 45°C and 50°C compared to 25°C (e).

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2.4.4 Fungal colonisation of PU coupons

To investigate fungal colonisation of PU coupons following burial, total fungal viable counts and putative fungal PU degrader counts recovered from the surface of coupons were enumerated on CEA and PUA plates respectively (Figure

2.5A). Total fungal viable and putative fungal degrader counts in the compost and soil environments in which the PU coupons were buried were also enumerated

(Figure 2.5B). In order to determine the minimum incubation period for plates required before no further fungal colonies emerged, CFUs were determined daily for up to 10 days (Section 2.7-Figure A2.3). At 25°C, no further colonies emerged after

6 days incubation and at 45° and 50°C, no further colonies were visible after 4 days.

Total viable counts in soil and in compost in which the coupons were buried remained similar for any temperature over 12 weeks. At 25°C, total viable counts in soil and compost were similar, however counts in compost were significantly

(P<0.05) lower at 45°C and 50°C, temperatures at which only thermotolerant and thermophilic fungi would grow (Dix and Webster, 1995). When total viable counts and putative PU degrader counts were compared, PU degraders composed ca 40-70% of all colonies recovered (Figure 2.5B) indicating that a large proportion of the viable fungal community was putative degraders. Total fungal viable counts from the surface of PU coupons buried in soil and compost at 25°C increased up to week 8 and then remained similar. Total viable counts from the PU surface also increased up to week 8 when incubated 45°C and 50°C, but decreased at week 12. A comparison of total viable counts and putative degrader counts demonstrated that ca. >70% of fungi colonising the PU surface were putative degraders (Figure 2.5A).

103

1E+07

1E+06

1E+05

1E+04

1E+03

1E+02

1E+01 Viablefungal count (CFU/cm2, log10axis) 1E+00 1 4 8 12 Incubation period (weeks) B

1E+07

1E+06

1E+05

1E+04

1E+03

1E+02

1E+01 Viablefungal count (CFU/g,axis) log10

1E+00 0 1 4 8 12 Incubation period (weeks)

Figure 2.5: Changes in the total fungal viable counts and total fungal PU degrader counts over a 12 weeks incubation period. (A) Total viable and putative PU degrading fungal counts (cfu/cm2) recovered from the surface of polyester PU coupons. (B) Total viable and putative PU degrading fungal counts in compost/soil (cfu/g compost/soil). PU coupons were buried in soil at 25°C (■ and ■ total and PU degraders), compost at 25°C (■ and ■ total and PU degraders), 45°C (■ and ■ total and PU degraders) and 50°C (■ and ■ total and PU degraders). Data represent the mean of three replicates ±SEM.

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2.4. 5 Identification of polyester PU degrading fungal isolates

Fungal colonies growing on PUA plates were difficult to distinguish morphologically. Therefore, colonies were transferred onto PDA and incubated up to

5 days. Differences in development of pigments, sporulation and colonial morphology on PDA enabled colonies to be grouped into different morphotypes and quantified (Figure 2.6). 4-5 isolates were randomly chosen, total genomic DNA was extracted, the ITS1-5.8s-ITS2 region of rDNA amplified by PCR, sequenced and sequences used to interrogate the NCBI database (Table 2.2). All the isolates were then aligned with the published sequences and a phylogenetic tree was constructed

(Figure 2.7). Although the two environments were very different (soil and compost), a number of common species were isolated at 25°C from the surface of PU from both soil and compost with only an Alternaria sp (morphotype 1) and Volutella ciliata

(mophotype 5) uniquely recovered from the surface of soil buried PU (Figure 2.6).

Fusarium, Penicillium, Trichosporon, Geomyces and Bionecteria (morphotypes, 6, 7,

4, 3 and 2 respectively) were consistently recovered from the surface of the PU coupons from both soil and compost (Figure 2.6 and Table 2.2). However, the relative proportions of the species recovered from the surface of PU differed between soil and compost and changed over the twelve weeks incubation period. From the surface of soil buried PU at 25°C, the proportion of the different species recovered was relatively constant except that Volutella ciliata could not be recovered after week 4, Geomyces pannorum could not be recovered after week 8 and Bionectria ochroleuca was the dominant species by week 12. The proportion of species recovered from the surface of PU buried in compost was also variable over the twelve week incubation period. Penicillium spp. that was dominant at only week 1

105 and from week 4-12 Fusarium solani/oxysporum was the dominant phenotype. At

45° and 50°C, Aspergillus fumigatus and Thermomyces lanuginosus respectively were the only species recovered at any time point. For morphotypes 6 and 7, subsequent sequencing of random isolates revealed that they were composed of more than one single species (a Penicillium madriti, P. chrysogenum and P. roseopurpureum for morphotype 6 and Fusarium solani and F. oxysporum for morphotype 7) (Table 2.2).

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100%

80%

60%

40%

20%

0% percentage eachof mophotype 1 4 8 12 1 4 8 12 Soil compost Incubation period of PU coupons (weeks)

Figure 2.6: Relative distribution of the colony morphotypes recovered from the surface of PU coupons buried over 12 weeks at 25°C. Polyester PU coupons were buried in soil at 25°C and in compost at 25°, 45° and 50°C for 12 weeks and viable fungal colonies recovered periodically from the surface of the coupons on PUA. Following transfer onto PDA, colonies were classified into different morphotypes and the percentage of fungal colonies recovered for each fungal morphotype calculated. A total of seven major morphotypes were recovered. Morphotype 1 (■, Alternaria sp), 2 (■, Bionectria ochroleuca), 3 (■, Geomyces pannorum), 4 (■,Trichosporon moniliformis), 5 (■,Volutella ciliata), 6 (■, Penicillium sp.), 7 (■, Fusarium sp.). At 45°C and 50 °C only Aspergillus fumigatus and Thermomyces lanuginosus respectively were recovered.

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Figure 2.7: Neighbour joining phylogenetic analysis of putative isolated PU degrading fungi. Fungi isolated and identified from the surface of PU coupons (Table 2.2) (● etic analysis with closely related species using Neighbour Joining Phylogenetic Tree (bootstrap corrected with 500 samples).

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Table 2. 2: Description, identification and frequency of occurrence of fungal morphotypes isolates recovered surface of buried polyester PU.

NCBI Morhotype Closest match in NCBI database Frequency on Temp. (˚C) Homology % accession (morphotype number) PUA plates* Environment# number

KF314689 Brown coloured, septate (5) Volutella ciliata ++ 25° s 100 JX996129 Yellowish cottony mycelium (7) Fusarium solani +++ 25° c+s 100 JX996130 Olive green, filamentous Aspergillus fumigatus +++ 45°+50° c 100

JX996131 Yellowish cottony mycelium (7) Fusarium oxysporum +++ 25° c+s 100 JX996132 Yeast like (4) Trichosporon moniliforme +++ 25° c+s 99 JX996133 Dark green colour, granular (6) Penicillium madriti ++ 25° c 99

JX996134 Dark green colour, granular (6) Penicillium chrysogenum ++ 25° c+s 99 JX996135 White cottony (2) Bionectria ochroleuca ++ 25° c+s 99 JX996136 Dark green colour, granular (6) Penicillium roseopurpureum + 25° c 99

JX996137 Grey (1) Alternaria sp. ++ 25° s 98 JX996139 Brown, floccose, radial edges (3) Geomyces pannorum ++ 25° c+s 100 JX996140 Peach, felt like Thermomyces lanuginosus +++ 50° c 100

All the isolates recovered had 99-100% similarity with the published isolates in the NCBI database. (* +++ = dominant, ++ frequent, +

occasional). #(c =compost, s=soil).

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2.4.6 Fungal community diversity on the surface of buried polyester PU

TRFLP was used to study the temporal change in the fungal communities on the surface of polyester PU during burial in soil at 25°C and in compost at 25°C,

45°C and 50°C. TRF electropherograms were compared using Principal Component

Analysis (PCA) and displayed as a scatter graph for each separate temperature for both the communities on the PU surface and in the surrounding compost/soil (Figure

2.8). At all temperatures, the fungal communities in soil and in compost separated into two distinct groups (soil and compost) indicating two distinctive communities

(Figure 2.8a). Within soil/ compost, at all temperatures, the community showed change after incubation (week0-4) then it was relatively stable over 12 weeks.

However, the communities colonising the PU surface were separate from the compost community but did not cluster together, indicating that the community on the PU surface was different from the surrounding soil or compost but was not stable and slightly changed over the 12 week period (Figures 2.8 a, b and c).

However, at 50°C the clustering was much weaker than at 45°C on the PU surface suggesting greater community changes over time at the higher temperature (Figure

2.8c).

TRFLP profiles generated from compost, soil and PU coupons were subjected to statistical analysis and the total number of TRFs and their relative abundance was used to calculate Shannon-Weaver index and evenness using MVSP as indicators of diversity and equitability respectively (Table 2.3). When soil or compost was incubated at 25°C, Shannon index increased from week 0 to week 4 and then remained approximately constant indicating an initial increase in species

110 diversity. The Shannon index from the surface of PU remained approximately constant over 12 weeks but was significantly lower (P<0.05) indicating the community on the surface was far less diverse than the surrounding compost or soil.

At 45° and 50°C, the Shannon Weaver index decreased at week 4 and then remained approximately constant indicating selection for thermophilic and thermotolerant species and a consequent reduction in species diversity. Again, the Shannon weaver index from the PU surface was significantly (P<0.05) lower than the surrounding compost (Table 2.3). Evenness remained approximately constant in soil and compost and on the surface of PU at all temperatures indicating no emergence of dominant species in the community over time, however, the evenness value from the PU surface was lower than the surrounding soil or compost indicating a less even distribution in the surface community.

The distribution and relative intensity of TRFs recovered from the surface of

PU coupons demonstrated that while culture-based analysis suggested a relatively low species diversity, particularly at 45° and 50°C, where only Aspergillus fumigatus and Thermomyces lanuginosus were recovered respectively, more than 30 TRFs were detected with no particular dominant species present (Figure 2.9).

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A TRFLP PCA score for fungal succession at 25°C

1.09

0.87

0.65

0.44 Group I 0.22

0.00 Group II Axis 2 (24%) 2 Axis -0.22 Group III -0.44

-0.65

-0.87

-1.09 -1.09 -0.87 -0.65 -0.44 -0.22 0.00 0.22 0.44 0.65 0.87 1.09

Axis 1 (46%)

PCA case scores B 8.0 6.4

4.8

3.2

1.6 Group I

0.0 Axis 2 (27%) 2 Axis -1.6

-3.2 Group II -4.8

-6.4

-8.0 -8.0 -6.4 -4.8 -3.2 -1.6 0.0 1.6 3.2 4.8 6.4 8.0

Axis 1 (40%)

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C TRFLP PCA score for fungal succession at 50°C 8.0

6.4

4.8

3.2

1.6

0.0

Axis 2 (23%) 2 Axis Group I -1.6

-3.2 Group II

-4.8

-6.4

-8.0 -8.0 -6.4 -4.8 -3.2 -1.6 0.0 1.6 3.2 4.8 6.4 8.0

Axis 1 (37%)

Figure 2.8: Principal Component Analysis of microbial community obtained from surface of PU and compost. Scatter plot for Principal Component Analysis of TRFLP pattern obtained from the surface of the buried polyester PU coupon and compost/soil at (A) 25°C and (B) 45 and (C) 50°C after week 4, 8 and 12. Fungal community in compost/soil and on the surface of PU showed obvious demarcation. PU surface under compost; week 4 (●), week 8 (●) and week 12(●), compost; week 0 (▲) week 4 (▲), week 8 (▲) and week 12(▲), PU surface under soil; week 4 (♦), week 8 (♦) and week 12(♦) and soil; week 0 (■) week 4 (■), week 8 (■) and week 12(■).

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Table 2.3: Variation in the Shannon-Weaver and Evenness indices derived from TRFLP electropherograms from fungal communities.

Shannon-Weaver Evenness Index Samples (temperature and Compost PU Compost PU weeks) or soil surface or soil surface 25°C soil 0 2.9 n/a# 0.7 n/a#

25°C soil 4 3.4 2.5 0.7 0.6

25°C soil 8 3.7 2.7 0.8 0.6

25°C soil 12 3.2 2.4 0.8 0.7

# # 25°C compost 0 2.8 n/a 0.8 n/a 25°C compost 4 3.5 2.6 0.8 0.7 25°C compost 8 3.3 2.5 0.8 0.7

25°C compost 12 3.4 2.5 0.8 0.8

45°C compost 0 2.8 n/a# 0.7 n/a#

45°C compost 4 2.2 1.4 0.8 0.6

45°C compost 8 1.9 1.1 0.7 0.7

45°C compost 12 2.1 1.5 0.6 0.6

50°C compost 0 2.9 n/a# 0.7 n/a#

50°C compost 4 2.0 1.1 0.6 0.6

50°C compost 8 2.0 1.8 0.7 0.6 50°C compost 12 2.1 1.7 0.7 0.7

Shannon’s index and Evenness were calculated for fungal communities from soil, compost and from the surface of buried PU coupons. # No fungi were detected from the PU surface prior to burial.

114

79.5

Relativentensity i (%)

51.55

54.25

65.38

71.26

94.79

128.8

106.18

111.88

134.82

155.01

334.4

165.24

176.19

248.93

391.6

268.82

315.69

321.23

326.38

345.83

369.27 438.92

TRF size (bp)

Figure 2. 9: TRFLP electropherograms of TRFs from fungal communities from the surface of PU over 12 weeks buried in soil or compost. Polyester PU coupons were buried in soil or compost and incubated at 25°C, or in compost and incubated at 45°C and 50°C over 12 weeks. Genomic DNA extracted from the fungal community from the PU surface was subjected to TRFLP analysis to generate TRF electropherograms. PU buried in soil at 25°C for 4 ■, 8 ■ and 12 ■ weeks; PU buried in compost at 25°C for 4 ■, 8 ■ and 12 ■weeks; PU buried in compost at 45°C for 4 ■, 8 ■ and 12 ■ weeks; PU buried in compost at 50°C for 4 ■, 8 ■ and 12 ■ weeks.

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2.4.7 Biodiversity of fungal communities by pyrosequencing

A total of 94,502 sequences were obtained after discarding low quality and short reads. OTU defined at 97% sequence similarity threshold were used to generate rarefaction curve. The curve leveled off in almost all samples indicated that the libraries provided an adequate sampling of fungal diversity. The number of observed species ranged between 14 and 51, with the highest number detected at 25°C (Figure

8a). Chao1 index, an estimator of OTU richness, was higher at 25°C in the compost and lowest in 50°C compost followed by population of the surface of PU coupons incubated at 45° and 50°C (Table 3). In compost at 25°C, Geomyces pannorum was the dominant species prior to incubation and after 12 weeks (49.6% and 42.2% respectively) while other species declined or were not detected (Thielavia sp.,

Arthrographis kalrae, Pseudallescheria boydii, Arthrobotrys flagran and

Doratomyces nanus). After 12 weeks, other species appeared at levels >1% that were not detected or present at less than 1% prior to incubation (Scytalidium thermophilum, Fusarium solani, Mortierella sp., Pseudallescheria fimeti and

Thermomyces lanuginosus). Incubation of compost at 45°C and 50°C for 12 weeks led to the development of a very different community compared to compost prior to incubation due to the selection of thermophilic and thermotolerant species. Three species, Emericella rugulosa, an unidentified fungal clone A and Scytalidium thermophilum were present at both 45°C and 50°C and accounted for 81% and

88.9% of all sequences respectively although Scytalidium thermophilum was predominant at 45°C and Emericella rugulosa at 50°C (Figure 9).

Pyrosequencing revealed clear differences in the fungal communities in the compost compared to the surface of the PU coupons at each temperature (Figure 2.11). At

1 16

25°C, the population of Geomyces pannorum in compost was 42.4% but on the surface was present at < 1% with Fusarium solani dominating at 53%. At 45°C and

50°C, the fungal community on the surface of PU were similar to each other and dominated by Candida ethanolica (60.3% and 57.6% respectively) an unidentified fungus clone A (32.3% and 29.0% respectively) and Penicillium paneum (2.4% and

2.5% respectively). Candida rugosa (4.9%) was the only species present at >1% that was only found only at 50°C. The compost community at 45°C was dominated by

Scytalidium thermophilum (68.1%) wheras at 50°C Emericella rugulosa was dominant (60.8%). PCA analysis of the OTU’s also demonstrated that the compost community at 45°C and 50°C clustered together and a second distinct cluster composed of the communities on the surface of PU at 45° and 50°C (Figure 2.12).

PCA analysis also showed that incubation of compost at 45° and 50°C had a greater effect on the fungal community than incubating at 25°C, but nonetheless, the community on the surface of PU at 25°C was different from the surrounding compost.

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Figure 2.10: Calculated rarefaction curves of observed species based on pyrosequencing data. Compost week 0 (−−−), compost incubated for 12 weeks at; 25˚C (−−−), 45˚C (−−−) and 50˚C (−−−), surface of PU coupons after 12 weeks incubated at 25˚C (−−−), 45˚C (−−−) and 50˚C (−−−).

118

Day 0 Geomyces pannorum ■ (49.6%)

Pseudallescheria boydii ■ (22.3%)

Arthrographis kalrae ■ (20.8%)

Arthrobotrys flagran ■ (2.5%)

Thielevia sp.■ (1.7%)

Doratomyces nanus ■ (1.0%)

Other ■ (2.2%, 14 OTU’s)

25°C compost Geomyces pannorum ■ (42.4%)

Thielevia sp.■ (14.7%)

Scytalidium thermophilum ■ (13.0%)

Fusarium solani ■ (9.1%)

Mortierella sp.■ (6.5%)

Pseudallescheria fimeti ■ (3.5%)

Arthrographis kalrae ■ (2.3%)

Thermomyces lanuginosus ■ (1.2%)

Others ■ (7.4%, 36 OTU's)

25°C PU Fusarium solani ■ (53.0%) Bionectria ochroleuca ■ (17.4%)

Alternaria sp.■ (16.6%)

Pleosporales sp.■ (2.5%)

Fusarium redolens ■ (2.3%)

Volutella ciliata ■ (2.1%)

Arthrographis kalrae ■ (1.7%)

Unidentified soil fungus clone A ■ (1.6%)

Others ■ (1.8%, 11 OTU's)

119

45°C compost Scytalidium thermophilum ■ (68.1%)

Sordariomycete sp ■ (11.0)

Unidentified fungus (c) ■ (8.4%)

Emericella rugulosa ■ (4.5%)

Aspergillus fumigatus ■ (2.1%)

Scopulariopsis brevicaulis ■ (2.1%)

Others ■ (3.9%, 15 OTU's)

45°C PU

Candida ethanolica ■ (60.3%)

Unidentified fungus clone A ■ (32.3%)

Penicillium paneum ■ (2.4%)

Unidentified fungus clone B ■ (1.6%)

Others ■ (3.3%, 14 OTU's)

120

50°C compost Emericella rugulosa ■ (60.8%)

Unidentified fungus clone A ■ (20.8%)

Scytalidium thermophilum ■ (7.3%)

Thermomyces lanuginosus ■ (5.0%)

Unidentified Ascomycota ■ (1.8%)

Arthrographis kalrae ■ (1.5%)

Candida ethanolica ■ (1.3%)

Others ■ (1.8%, 4 OTU's)

50°C PU

Candida ethanolica ■ (57.6%)

Unidentified fungus clone A ■ (29%)

Candida rugosa ■ (4.9%)

Penicillium paneum ■ (2.5%)

Unidentified fungus clone B ■ (1.3%)

Others ■ (4.6%, 15 OTU's)

Figure 2.11: 454 pyrosequencing data obtained for fungal community on week 0 and 12 from compost and surface of PU coupons at 25°, 45 ° and 50°C. Community on the surface of PU coupons is different from the compost and change was observed in compost from week 0 to 12. At 25°C Fusarium solani and 45 and 50°C Candida ethanolica was the dominant taxa. There were different cluster of sequences assigned as unidentified fungal clone which are differentiated as “A” and “B”.

121

Table 2.4: Statistical evaluation of fungal communities from compost and from the surface of PU coupons buried in compost incubated at different temperatures for 12 weeks.

Samples after Shannon index Chao1 index 12 weeks

Compost initial 2.3 30.9 (time 0)

Compost 25°C 3.0 63.2

PU surface 25°C 2.7 29.4

Compost 45°C 1.8 32.1

PU surface 45°C 1.8 27.1

Compost 50°C 1.0 15.5

PU surface 50°C 1.8 29.7

ITS amplicons from the initial compost (compost 0) and from compost and PU surface (PU) after 12 weeks of incubation at 25°, 45° and 50˚C were subjected to 454 pyrosequencing and the relative distribution amongst the detected OTU’s used to calculate the Shannon and Chao1 indices.

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Figure 2.12: Principal component analysis of fungal populations obtained after 454- pyrosequencing. ITS amplicons from the initial compost and from the PU surface buried in compost after 12 weeks of incubation at 25°, 45° and 50°C were subjected to 454 pyrosequencing and principal component analysis. Compost at week 0 (▼), compost and PU coupons after week 12 at 25°C (▲,), compost and PU coupons after week 12 at 45°C (,●), compost and PU coupons after week 12 at 50°C (▲,▲).

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2.5 Discussion

Polyurethanes are a diverse group of plastics with a wide range of applications ranging from packaging materials, shoe soles coatings and paints and are therefore common environmental pollutants (Howard, 2002). Polyester PU’s are known to be susceptible to microbial enzymatic degradation through hydrolysis of the polymer (Pathirana and Seal, 1984; Morton and Surman, 1994; Howard, 2002) and it has been shown previously, that fungi are the dominant microbes responsible for colonisation and biodegradation of PU’s in soil environments, particularly when the moisture content falls between 20 and 70 % (Barratt, et al., 2003;Cosgrove et al.,

2007, 2010)

Composting is a natural process involving the aerobic decomposition of organic wastes by a mixed microbial consortium that also involves thermophilic microbes appearing due to the increase in heat that occurs during microbial respiration and in recent years has been increasingly used commercially as a waste management treatment for green and food wastes (Ryckeboer, Mergaert, Vaes, et al.,

2003). In this study, we investigated whether microbes, and in particular fungi present in compost, had the capacity to degrade PU and whether selection of particular communities on the surface of PU were associated with this process. For this purpose, we incubated PU coupons for up to 12 weeks in mature compost at

25˚C (to emulate the maturation phase) and at 45˚C and 50˚C (to emulate the thermophilic stage). Coupons were also buried in soil at 25°C to enable the rate of degradation in soil to be compared to compost. Significant colonisation by fungal mycelia was clearly observed in all cases along with significant macroscopic changes to the PU (Figure 2.2) and severe cracking of the PU surface and fungal hyphae were

124 visible under environmental scanning electron microscopy (Figure 2.3). Barratt et al.,

(2003) also observed a high coverage of polyurethane film by fungal hyphae and spores and extensive surface damage in soil buried polyester PU. Despite these major changes to the integrity of the coupons, little loss in dry weight was observed.

Weight loss over 12 weeks was not therefore a reliable indicator for polyester PU biodegradation. Barratt et al., (2003) and Pathirana & Seal (1984) also reported no change in weight loss.

Significant changes were observed in the tensile strength of buried polyester

PU. Tensile strength is defined as the load at break divided by the original cross- sectional area. A number of studies have analysed the rate of degradation of PU by analysing loss in tensile strength as it is a sensitive measure of polymer integrity

(Barratt et al., 2003; Bentham, et al., 1987; Cosgrove et al., 2007, 2010; Dale &

Squirrell 1990; Ibrahim et al., 2009; Kay et al., 1991; Mathur & Prasad 2012). Umare

& Chandure (2008) studied the degradation of polyester PU under three different treatments; alkaline hydrolysis, enzymatic hydrolysis and soil burial and reported that soil burial gave the greatest level of degradation. Previously Barratt et al., (2003) buried polyurethane in soil microcosms and reported a reduction in tensile strength of 60% after 44 days. In another study, Cosgrove et al., (2007) investigated the rate of degradation of polyester PU in situ in the environment and found a ca. 95% reduction in tensile strength after 5 months. In our study, percentage loss in tensile strength of buried polyester PU coupons at 25°, 45° and 50°C in compost was 90, 80 and 70%, respectively suggesting that in compost, PU degradation occurs to a similar extent at all phases of the composting process, albeit by different fungal species and that the extent of degradation in compost was similar to that observed in soil. Loss in

125 tensile strength has been shown to be due to the secretion of extracellular enzymes that degrade the ester and urethane linkages of polyurethane causing polymer chain scission (Pathirana and Seal, 1984; Dale and Squirrell, 1990; Akutsu et al., 1998;

Nakajima-Kambe et al., 1999). A fall in tensile strength was also observed when polyester PU coupons were incubated for the same length of time in sterile compost extract, although to a much lower extent. Aquino et al., (2010) studied hydrolysis of polyester PU over a wide temperature range (10°-70°C) and found that PU was stable at 50°C but significant polymer hydrolysis occurred at 70°C after 177 days of incubation. In another study, Zuidema et al., (2009) also reported significant hydrolysis of PU at two different temperatures (37°C and 60°C) after 3 years. Thus, while polyester PU is vulnerable to hydrolysis it is a slow process and does not account for the large drop in tensile strength observed when buried in soil or compost.

By contrast, no change in tensile strength was observed after 3 months burial under any condition for polyether PU. Polyether PU is known to be far more recalcitrant to degradation compared to polyester PU (Rutkowska et al., 2002).

Polyether PU are moderately to highly resistant to fungal attack than polyester PU because of the presence of high molecular weight branched polyol chains (Darby &

Kaplan 1968; Filip, 1985). Jansen et al., (1991) reported that polyether PU degradation occurred very slowly when Staphylococcus epidermis was used as a test organism and polyether based PU also found to be highly resistant to anaerobic bacterial degradation (Urgun-Demirtas et al., 2007). Krasowska et al., (2012) reported that polyether based PU only turned yellow brownish when incubated in a compost pile. However, Matsumiya et al., (2010) reported that an Alternaria sp.

126 could degrade polyether polyurethane physically and enzymatically and released metabolites were detected in the media indicating physical degradation. This appeared to be due to the hydrolysis of urea and urethane bonds releasing polyols and polyisocyanates. The difference in degradation could be due to the difference in composition of polyether PU types. In our study, polyether PU was incubated for 12 weeks and may require a much longer incubation period before a significant change in tensile strength is observed.

The fungal species isolated and cultured from the surface of polyester PU at

25˚C in both soil and compost after 12 weeks were the same (Bionectria ochroleuca, a Penicillium sp, Fusarium oxysporum/solani and Trichosporon moniliformis) although recovered to different extents. The only exception was Geomyces pannorum which was recovered from the PU surface in compost but not in soil after week 12 although it was recovered at weeks 1, 4 and 8. Many of the fungi recovered have been reported to be associated with PU degradation previously (Pathirana and Seal,

1984; Pommer and Lorenz, 1985; Bentham et al., 1987; Crabbe et al., 1994; Barratt et al., 2003). Thus, culturing suggested limited species diversity on the surface of PU when buried in both soil and compost, but cultivation is known to be unreliable and to under report the true diversity of microbial communities as only a highly limited proportion of species (estimated to be <1-5%) can grow on selected synthetic media

(Magnuson & Lasure, 2000; Salar & Aneja, 2006).

TRFLP has previously been successfully used to analyse species diversity in fungal communities from a wide variety of environments (Dickie & FitzJohn,

2007; Lord et al., 2002; Wu et al., 2007). In this study, TRFLP analysis indicated far

127 greater species diversity on the surface of PU than that reported by culturing alone

(Figure 2.9) particularly at 45° and 50°C, where only one species in each case was recovered by cultivation. PCA analysis of TRFLP electropherograms also demonstrated clear differences in the communities colonising the surface of PU compared to the surrounding environment indicating selection on the PU surface

(Figure 2.8). Previously, Cosgrove et al., (2007) also reported the polyester PU buried in two soil types using DGGE showed different microbial communities.

To examine the diversity of the fungal communities on the surface of PU buried in compost in more detail, pyrosequencing analyses were performed with total genomic DNA qextracted from mycelium colonising the surface of the PU coupons after 12 weeks burial at 25°, 45° and 50°C and compared to the surrounding compost fungal community (Figure 2.11). PCA analysis of the pyrosequencing data were broadly congruent with the PCA analysis of the TRFLP electropherograms in demonstrating that the fungal community on the surface of PU was different from the surrounding compost community at each temperature, and that the PU surface community at 45° and 50°C were broadly similar (Figure 2.12).

454 pyrosequencing is a high throughput sequencing technique particularly useful in the detection of rare community members that may represent <1% of the microbiota (Ronaghi, 2001). Recently 454 pyrosequencing demonstrated that fungal diversity is rich in soil with detection of rare OTUs that were previously under estimated (Buée et al., 2013; Danielsen et al., 2012). At 25°C, Fusarium solani,

Bionectria ochroleuca and Alternaria sp. accounted for 53%, 17.4% and 16.6% of the sequences recovered. After 12 weeks, both F. solani and B. ochroleuca were also

128 the dominant organisms recovered by cultivation (Figure 2.6). F. solani has previously been associated with the degradation of polyester PU (Bentham et al.,

1987, Crabbe et al., 1994). Similarly, B. ochroleuca, was also identified as a dominant organism from the surface of polyester PU buried in soil after pre- treatment with impranil DLN (a polyester PU dispersion) following cloning and sequencing of a major band in DGGE (Cosgrove et al., 2010). Interestingly, it was also isolated from soil as a major degrader of poly (butylene succinate) films, a biodegradable polyester polymer suggesting it posseses the ability to efficiently degrade esters within long chain polymers (Mei et al., 2012). Alternaria has also been ubiquitously found in soil and compost and several species have been extensively reported as polyurethane degraders (Pommer and Lorenz, 1985;

Stranger-Johannessen, 1985; Cosgrove et al., 2007; Ibrahim et al., 2009; Matsumiya et al., 2010; Russell et al., 2011)

While the most dominant species cultivated from the surface of PU at 25°C were amongst the most dominant identified by pyrosequencing, at 45° and 50°C there was little correlation. At 45° and 50°C, A. fumigatus and T. lanuginosus were the only fungi isolated via the culture-based technique. However, pyrosequencing revealed that in both cases, Candida ethanolica was the dominant organism on the surface of PU. C. ethanolica was first isolated as an ethanol tolerant yeast with a limited ability to ferment sugars and is closely related to the Pichia group (Rybarova et al., 1980). It has, however, been reported to be amongst a group of yeasts identified from a clone library which dominated the early stages of composting in a commercial compost facility (Hultman et al., 2010). Also dominant was an unidentified fungal clone A also present at both 45°C and 50°C that has been found

129 in soils but has yet to be isolated or taxonomically assigned. The third most dominant taxon was Candida rugosa comprising 4.9% of the community on the surface of PU coupons at 50°C. Gautam et al., (2007) have demonstrated that a lipase enzyme from

Candida rugosa was able to cause significant PU degradation. Penicillium paneum was also detected on the surface of PU coupons and is closely related to Penicillium roquiforti and has been reported to grow under highly acidic environments (Chitarra et al., 2004). Together, C. ethanolica and unidentified fungus clone A accounted for

>85% of the sequences recovered with the remaining sequences belonging to 14 to

16 other species (Section 2.7- Table A2.4). While A. fumigatus and T. lanuginosus were detected by pyrosequencing on the surface of PU, they accounted for <0.35% of the sequences, despite appearing to be the only organisms on the surface through conventional cultivation.

In summary, this study has demonstrated that polyester PU is susceptible to fungal biodegradation in compost under both thermophilic (thermophilic stage) and mesophilic (maturation phase) conditions and that positive selection for rare taxa from the existing compost community on the PU surface occurs. Thus, there may be the potential for directing polyester PU waste through existing commercial composting processes, thereby reducing the burden on landfill sites.

130

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2.7 Appendix2

Figure A2.1: Physical changes to polyether polyurethane coupons after burial in compost. Polyether PU coupons unburied (a) buried under compost at 25° (b), 45° (c) and 50° C (d) for 12 weeks. No change was observed on the surface of the coupons.

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Figure A2.2: ESEM images of polyether polyurethane coupons after burial in compost. Polyether PU coupons unburied (a) buried under compost at 25° (b), 45° (c) and 50° C (d) for 12 weeks. No change was observed on the surface of the coupons.

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TableA2.1: Change in tensile strength of polyether PU dumb-bells after 12 weeks of incubation under soil at 25° and compost at 25°, 45° and 50°C.

Tensile strength Treatment (MPa)

control 51.93±2.3

25°C soil 55.12±6.3

25°C compost 55.33±4.3

45°C compost 55.33±10.3

55°C compost 54.64±5.1

TableA2.2: Effect of soil and compost burial on percentage weight loss of PU coupons.

Treatment % Weight loss

25°C soil 0.57±0.40

25°C compost 0.12±0.19

45°C compost 0.45±0.19

50°C compost 0.99±0.23

Percentage weight loss buried polyurethane dumb-bells after twelve weeks of burial under compost & soil at 25, 45 and 50°C.

138

8.0

7.0

6.0

5.0

4.0

3.0

2.0 log log cfu/gm of compost fungi from 1.0

0.0 1 2 3 4 5 6 7 8 9 10 DAYS

Figure A2.3: Appearance of fungal colonies over 10 days after inoculation. Fungal biomass recovered from PU coupons was plated onto media and colonies appearing counted over 10 consecutive days. Viable count from 25°C at SEA, Impranil clearing count on PUA at 25°C, Viable count from 45°C at SEA,  Impranil clearing count on PUA at 45°C, Viable count from 50°C at CEA and  Impranil clearing count on PUA at 50°C).

139

TableA2.3: Fungal colony count obtained from compost and surface of PU coupons after incubation at 25° , 45° and 50°C for 4, 8, and 12 weeks.

Weeks Media 25°C soil 25 °C compost 45°C compost 50°C compost 6 5 3 3 0 SEA 3 ± 0.2 X10 A 5 ± 0.1 X10 A 4 ± 0.8 X10 A 8 ±1.7 X10 A 6 5 3 3 PUA 1 ±0.1 X10 A 2 ±0.5 X10 C 3 ±0.3 X10 A 6 ±0 X10 B % 41.8 38.2 89.4 70.4 5 5 4 4 1 SEA 7 ±0.02 X10 B 3 ±0.8 X10 B 1±0.1 X10 B 1 ±0.07 X10 A, B 5 5 3 2 PUA 3 ±0.2 X10 B 2±0.2 X10 C 2 ±0.5 X10 A 3 ±0 X10 B % 48.3 61.2 17.0 2.4 5 5 4 3 , 4 SEA 4 ±0.3 X10 B 1±0.01 X10 B, C 2 ±0.1 X10 B 9 ±0.1 X10 A B PUA 3 ±0.1 X105 1 ±0.07 X105 2 ±0.06 X104 4 ±0.8 X102 B C B B % 76.3 95.4 64.7 4.6 5 5 4 3A, 8 SEA 5 ±0.1 X10 B 2 ±0.4 X10 B, C 2 ±0.09 X10 B 9 ±0.8 X10 B 5 5 4 2 PUA 3 ±0.3 X10 B 2 ±0.4 X10 C 1 ±0.2 X10 B 4 ±0.8 X10 B % 69.8 76.7 66.4 4.6 5 5 4 3 12 SEA 5 ±0.2 X10 B 6 ±0.2 X10 B, C 3 ±0.08 X10 B 7 ±1.7 X10 A, B 5 5 4 2 PUA 4 ±0.02 X10 B 5 ±0 X10 C 2 ±0.3 X10 B 4 ±0 X10 B % 76.2 83.0 61.5 5.1 Weeks Media 25°C soil 25 °C compost 45°C compost 50°C compost

3 2 1 SEA 2 ±0.2 X10 A 2±0.1 X10 A 6 ±0 A 3 ±0 A

3 2 PUA 1±0.3 X10 A 1±0.2 X10 A 1±0.1 A 0±0 A % 77.9 74.4 12.8 16.7

3 3 4 SEA 3±0.4 X10 A 2±0.1 X10 B 87±11 B 26±2 A, B

3 3 PUA 3±0.3 X10 A 2±0.2 X10 A 88±4.2 B 20±0.3 A, B % 96.3 93.2 100.0 77.2

4 3 8 SEA 4±0.2 X10 B 2±0.7 X10 B 33±2 C 35±2 B

4 3 PUA 3±0.4 X10 B 2±0.04 X10 B 20±9.2 A, C 32±1.1 B % 96.3 93.1 59.7 92.9

4 3 12 SEA 2±0.1 X10 B 3±0.5 X10 B 5±1 A 8±1 A

4 3 PUA 2±0.5 X10 B 3±0.2 X10 B 5±0.4 A 8±1.2 A % 100.0 98.9 100.0 92.0

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Figure A2.4: Pictorial presentation of species isolated from culture based technique, on PDA (left) and PUA (right).

141

TableA2.4: Assigned taxonomy to the OTU clusters of sequences obtained from pyrosequencing data. Total number of sequences obtained from each sample is mention under parenthesis beside the name of the samples.

Taxon DAY 0 25°C 25°C 45°C 45°C 50°C 50°CPU PU PU

Unidentified soil fungi (a) 0.87 0.05 32.3 20.7 29

Unidentified soil fungi (b) 1.6 1.33

Unidentified soil fungi (c) 8.39

Unidentified ascomycota 0.20 0.11 0.05 1.75 0.35

k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Capnodiales;f__Mycosphaerellaceae;Other 0.23 0.04

k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Incertae_sedis;f__Eremomycetaceae;g__Arthrographis;s_kalra 20.76 2.27 1.73 1.50 e

k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;g_Pseudallescheria;s_fimeti 2.54

k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__Pleosporaceae;g__Alternaria 0.02 16.61

k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Pleosporales;f__unidentified;g__unidentified 0.85

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Incertae_sedis;g__Thermomyces;s 0.04 1.16 0.30 0.15 5.00 0.07

_lanuginosus

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Aspergillus;s_fumigatus 0.01 0.29 2.09 0.35 0.75 0.35

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Emericella;s_rugulosus 0.35 4.47 60.75 0.77

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Eurotium 0.05

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Paecilomyces 0.02 0.03

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Penicillium;s_paneum 0.81 0.05 0.19 2.41 2.46

142 142

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Sagenomella 0.05

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Talaromyces 0.25 0.28

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Onygenales;f__Incertae_sedis;g__Myceliophthora 0.05 0.07 0.05 0.21

k__Fungi;p__Ascomycota;c__Incertae_sedis;o__Incertae_sedis;f__Incertae_sedis;g__Ilyonectria 0.04

k__Fungi;p__Ascomycota;c__Lecanoromycetes;o__unidentified;f__unidentified;g__unidentified 0.08

k__Fungi;p__Ascomycota;c__Leotiomycetes;o__Incertae_sedis;f__Incertae_sedis;g__Geomyces; s_pannorum 49.62 42.44 1.10 0.41 0.25

k__Fungi;p__Ascomycota;c__Orbiliomycetes;o__Orbiliales;f__Orbiliaceae;g__Arthrobotrys;s_flagrans 2.46 0.24 0.32 0.04 0.30 0.28

k__Fungi;p__Ascomycota;c__Pezizomycetes;o__Pezizales;f__Incertae_sedis;g__Cephaliophora 0.60 0.84

k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;f__Dipodascaceae;g__Galactomyces 0.01 0.12 0.60 0.28

k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;fIncertae_sedis;g__Candida;s_ethanolica 0.17 0.08 0.78 60.3 1.25 62.53

k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;f__Incertae_sedis;g__Yarrowia 0.40 0.42

k__Fungi;p__Ascomycota 0.01 0.02 11.04 0.25

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Bionectriaceae;g__Bionectria;s_ochroleucia 0.35 17.36

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Clavicipitaceae;Other 0.39

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Cordycipitaceae;g__Lecanicillium 0.12 0.08

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Cordycipitaceae;g__Simplicillium 0.02 0.01

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Hypocreaceae;g__Acrostalagmus 0.03

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g_fusarium;s_solani 0.04 9.14 52.96 0.26 0.07

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g__Fusarium;s_sp. 0.14 2.25

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g__Neonectria 0.15

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g__Volutella;s_cilliata 0.05 2.06

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;Other 22.31 0.14 0.15 0.05 0.14

143 143

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Cephalotrichum 0.97 0.32

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Doratomyces 0.16 0.11 0.63

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Microascus 2.09

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Petriella 0.02

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Pseudallescheria 0.08 3.51 0.03

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Scedosporium 0.66 0.34 0.20 0.28

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;Other;Other

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__Chaetomiaceae;g__Corynascus 0.01 0.68 0.04 0.07

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__Chaetomiaceae;g__Thielevia 1.67 14.70 0.24

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__Sordariaceae;Other 0.01 0.29

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__unidentified;g__unidentified 0.06 0.37 0.50 0.07

k__Fungi;p__Ascomycota;c__Sordariomycetes;o__unidentified;f__unidentified;g__unidentified 0.05

k__Fungi;p__Ascomycota;c__Leotiomycetes;o__unidentified;f__unidentified;g__Scytalidium;s_thermophilum 0.42 12.96 0.04 68.08 0.20 7.25 0.07

k__Fungi;p__Basidiomycota;Other;Other;Other;Other 0.34 0.84

k__Fungi;p__Basidiomycota;c__Agaricomycetes;o__Agaricales;f__Agaricaceae;g__unidentified 0.05 0.11

k__Fungi;p__Basidiomycota;c__Agaricomycetes;o__Agaricales;f__Bolbitiaceae;g__Conocybe 0.03

k__Fungi;p__Basidiomycota;c__Tremellomycetes;o__Filobasidiales;f__Filobasidiaceae;g__Cryptococcus 0.03

k__Fungi;p__Basidiomycota;c__Tremellomycetes;o__Tremellales;f__Trichosporonaceae;g__Trichosporon 0.05

k__Fungi;p__Zygomycota;c__Incertae_sedis;o__Mortierellales;f__Mortierellaceae;g__Mortierella;s_sp. 6.45 0.03

k__Fungi;p__Zygomycota;c__Incertae_sedis;o__Mucorales;f__Lichtheimiaceae;g__Lichtheimia

k__Fungi;p__Zygomycota;c__Incertae_sedis;o__Mucorales;f__Mucoraceae;g__Rhizopus 0.03

144

Chapter 3: Biodegradation of polyester polyurethane during commercial composting

Urooj Zafar1, Alan Heyworth2, Alberto Saiani3, Ashley Houlden1 and Geoff D.

Robson1

1Faculty of Life Sciences, Michael Smith Building, University of Manchester,

Manchester M13 9PT, UK

2The TEG group plc, Chorley, Lancs, PR7 7NA, UK

3School of Materials, Faculty of Engineering and Physical Sciences, University of

Manchester, Manchester M13 9PL, UK

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3.1 Abstract

Plastics are an integral part of today’s society and have a large and diverse range of applications due to their durability, low cost, strength and thermal and electrical insulating properties. In Europe alone, 25 million tonnes of plastic waste was generated in 2011 with 41% directed to landfill sites. However, their low degradation rates coupled with an increasing scarcity of landfill sites and growing water and land pollution problems require alternatives to be developed. Composting is a natural process involving the aerobic decomposition of organic wastes by a mixed microbial consortium that involves thermophilic microbes during the process due to the heat generated during decomposition. In this study we have selected polyurethane (PU) as a model plastic to investigate the biodegradation during the maturation stage of a commercial composting process. PU has a wide range of applications in the medical, automotive and industrial fields and constitutes ca. 7% of all plastics produced in Europe. PU coupons were buried for up to 28 days in the centre and at the surface of a 10 m high compost pile formed from the output of an in silo composting process for green and food waste. Fungal communities colonising

PU coupons were compared with the native compost communities using culture based and molecular techniques. Putative PU degrading fungi were ubiquitous in compost and rapidly colonized the surface of PU coupons. The most dominant isolates recovered from PU were Thermomyces lanuginosus, Candida rugosa and

Aspergillus fumigatus, also Malbranchea cinnamomea and Lichtheimia sp.were also frequently recovered. Denaturing Gradient Gel Electrophoresis (DGGE), Terminal

Restriction Fragment Length Polymorphism (TRFLP) and 454 pyrosequencing analysis revealed that with time as the temperature decreased from 70°C to 42°C at

146 the surface of the compost pile, fungal diversity in the compost and on the surface of the PU coupons increased. The fungal community on the PU coupons was different from the surrounding compost with an unidentified fungal clone A and

Arthrographis kalrae as the dominant species on the PU surface. The community was associated with physical deterioration of the PU coupons leading to deep pits and cracking of the PU coupons accompanied by a loss in both tensile strength

(>70%) and % elongation at break (>50%).

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3.2 Introduction

The volume of petro-chemical based plastics used in commercial applications has grown enormously world-wide since their first wide scale introduction over 50 years ago. In Europe alone in 2010, the total plastic production was 57 million tons, which in the same year generated 24.7 million tonnes of plastic waste (Plastics – the Facts 2012). Such large volumes of waste require a number of different waste management strategies with recycling, incineration and disposal to landfill sites amongst the most widely implemented (Shah et al., 2008; Thompson et al., 2009). In 2010, 6.0 million tonnes were recycled and 10.4 million tonnes were directed to landfills (Plastics-the facts 2012). However, these current strategies have a number of limitations. Not all plastics can be recycled and those that are often have limited uses with weak market demand, are associated with high cost and are difficult to separate into different plastic chemistries (Hopewell et al., 2009).

Incineration of some plastics is associated with the production of hazardous substances (Jayasekara et al., 2005) while directing waste to landfill sites leads to the contamination of soil and ground water with plasticizers and other additives and is increasingly expensive due to reduced availability of landfill sites (Oehlmann et al.,

2009). Thus, there is a need to develop alternative strategies for municipal plastic waste management.

In 2011/12 in the UK, 6.7 Million tonnes of waste was collected by local authorities and 43% of household waste was recycled (DEFRA, 2013) including waste directed to commercial composting facilities. Composting is a natural self- heating process with an increase in temperature occurring due to microbial growth

148 and respiration as organic substances utilised (Finstein and Morris, 1975).

Composting therefore goes through a range of temperatures starting from a mesophilic phase with increasing temperature leading to a thermophilic phase before cooling and entering a maturation phase. Polyurethanes (PU) are a group of heteropolymer plastics formed by the condensation of polyisocyanate and polyols and have a wide range of applications in the medical, automotive, building and industrial sectors (Krasowska et al., 2012). PUs are known to be vulnerable to microbial attack (Morton and Surman 1994) as they contain ester linkages within the backbone of the polymer that are naturally vulnerable to esterases as first reported by

Darby & Kaplan (1968). Previously, it has been reported that fungi are the predominant microbes responsible for the biodegradation of PU in situ (Barratt et al.,

2003) and fungal PU biodegraders have been isolated from soil (Cosgrove et al.,

2007) and at least some, have been shown to have the potential to accelerate PU biodegradation (Cosgrove et al., 2007). Chapter 2 of this thesis reported that a number of fungal isolates are able to degrade impranil (liquid dispersion of PU) including mesophilic, thermotolerant and thermophilic fungi. Pyrosequencing and

TRFLP data suggested that the community colonising the surface of PU coupons was different from the community in the bulk compost suggesting selection for PU degraders.

The present study investigated the potential of a commercial composting process to degrade PU. PU coupons were introduced into compost heaps formed from the output of a commercial in silo green waste composting facility (TEG group,

UK). We report the physical changes in PU during this process and monitored the diversity and development of the fungal community over the surface of the buried

149 polyester PU coupons using both culture-based and molecular analysis (DGGE,

TRFLP and 454 pyrosequencing). The results reported here indicate that considerable degradation of polyester PU occurs during the maturation phase of the composting process due to deterioration by fungi. Enrichment of the fungal community occurred on the surface of PU coupons with the appearance of rare taxa.

This study demonstrates that commercial composting has the potential to be an alternative process for PU waste disposal.

150

3.3 Materials and Methods

3.3.1 Fabrication of polyurethane coupons

Polyester and polyether polyurethane (PU) coupons of 16 x 16 x 0.1 cm

(total surface area 518.4 cm2) were fabricated from PU beads (Ellastollan® 685 A10 and Ellastollan ® 1180 A10) by melting at 180°C at 8 Barr pressure in a compression moulding machine (Moore, Birmingham, UK). Dumb-bells (total length 5 cm, width at the end 1.9 cm with 19 cm gauge length) were cut from the coupons using a moulder cutter (Wallace Test equipment, UK). Rectangular coupons (33.6 cm2) were cut with scalpel blades.

3.3.2 Burial of PU coupons in compost

In situ biodegradation of PU coupons was carried out at a commercial composting site (The TEG group, Todmorden, UK). Input material consisted largely of domestic green and food waste with material added daily to the top of silo cages and mechanically removed from the base of the silos periodically. Material added to the top of the silos takes ca. 14 days to reach the bottom and the temperature within the silos ranged from ca. 60° to 75°C. Material removed from the silos is used to form large (ca 8-10 m tall) compost piles and allowed to mature further without turning for ca. 28 days during which the temperature gradually declines (Appendix-figure 3.).

Polyester and polyether PU coupons were buried at approx. 0.3 meters (surface samples) and 4 meters (centre samples) from the top of a 10 m tall compost pile, as it was formed. After 2, 14 and 28 days, temperature at the centre and surface of the compost pile was determined and three replicate coupons were recovered. Coupons were cut into two halves; one half was used to collect the biomass from the surface,

151 dumb-bells and small pieces (20 mg) were cut from the other half for tensile strength and Differential Scanning Calorimetry (DSC) analysis respectively. Control coupons were placed in sealed sterile Petri plates and incubated at 45°C and 55°C for up to 6 months.

3.3.3 Environmental Scanning Electron Microscopy (ESEM) of the surface of PU coupons

Coupons recovered from the pile were washed with sterile distilled water followed by 70% (v/v) ethanol, air dried and observed under environmental scanning microscopy (ESEM, FEI Quanta 2000 Netherlands) to visualize the surface of the coupons at low pressure and room temperature.

3.3.4 Tensile strength determination

Physical deterioration of polyurethane coupons was measured by the loss in tensile strength and percentage elongation at break. Dumb-bells (total length 5 cm, width at the end 1.9cm with 19 cm gauge length) were cut from the coupons using a moulder cutter (Wallace Test equipment, UK) and tensile strength and percentage elongation at break determined with a crosshead speed of 1.5 cm/min (Tinius Olsen H5KT-

0586, UK).

3.3.5 Differential Scanning Calorimetry

Differential Scanning Calorimetry (DSC) was used to monitor changes in the physical structure of buried PU coupons by determining the glass transition temperature. Compost particles were removed from the surface of the coupons with a soft brush, briefly rinsed with water and wiped with 70% (v/v) ethanol. Samples (10

152 mg) were excised, hermetically sealed and analysed over a temperature range of -90 to 220°C at a rate of 20°C/min (DSC Q100, TA instruments, USA).

3.3.6 Viable count determination

For cultivation and molecular analysis, biomass from the surface of PU coupons was recovered according to Cosgrove et al., (2007) 2, 14 and 28 days after burial.

Coupons were first agitated in 20 ml sterile phosphate buffered saline (PBS, section

2.3.1.6) for 3 min to remove loosely attached particles, placed into a Petri dish and both sides scraped using a sterile razor in the presence of 10 ml of PBS. Triplicate samples were pooled, agitated for 5 min and 1 ml of the biomass suspension used for microbial viable count determination. The remaining suspension was centrifuged at

3000 X g for 30 min at 4°C, the supernatant discarded and the pellet used for DNA extraction.

The biomass suspension was serially diluted in PBS and plated out onto Potato

Dextrose Agar (PDA Formedium, England and section 2.3.1.2) and R2A (Oxoid

England, section 2.3.1.1) agar plates to determine the total viable count of fungi and bacteria respectively. Polyurethane agar (PUA, Crabbe et al., 1994 and section

2.3.1.5) was used for the enumeration of polyurethane degrading microorganisms with Impranil (a polyurethane dispersion, Bayer, Newbury, UK), as a sole carbon source. Filter sterilized (0.22 µm) chloramphenicol (50 µg/ml) and nystatin (250

µg/ml) was used to inhibit bacterial and fungal growth respectively.

3.3.7 Extraction, amplification and purification of genomic DNA

Genomic DNA was extracted from the mycelium of fungal colonies according to Feng et al., (2010). Mycelium/ spores (ca. 20 mg) were collected from

153 the surface of confluent PDA cultures with a sterile toothpick and placed into 1.5 ml centrifuge tubes containing 0.5 g of 0.5 mm diameter glass beads. After addition of

0.65 ml lysis buffer (100 mM Tris-HCl, pH 8.0; 50 mM EDTA, pH 8.0; 1% (w/v)

SDS; 10 μg ml−1 RNase A), tubes were homogenized twice for 30 s and centrifuged for 2 min at 13000 rpm. After centrifugation, 500 μl of supernatant was transferred into a new tube containing 100 μl of potassium acetate buffer (3.0 M, pH 5.5). The tube was inverted several times and centrifuged for 2 min at 13000 rpm and 500 μl of supernatant was transferred into a new tube containing 500 μl of isopropanol, inverted several times and centrifuged for 2 min 13000 rpm. The supernatant was removed and the DNA pellet washed with 750 μl of 70% (v/v) ethanol. After centrifugation for 30 s, ethanol was removed and the DNA pellet air-dried for 5–10 min. The DNA was dissolved in 50 μl sterile distilled water and stored at -20°C until required.

Isolates were identified according to Webb et al., (2000) using the

Internally Transcribed Region (ITS) of rDNA amplified using the fungal universal primers ITS1 (5’-TCCGTAGGTGAACCTGCGG-3’) and ITS4 (5’-

The PCR consisted of initial denaturation 94°C for 3 min, 35 cycles, denaturation at

94°C for 1 min, annealing at 56°C for 1 min and extension at 72°C for 1 min. Final extension 72°C for 3 min. PCR amplicons varied between ca. 500-575 bp depending on the species. Amplified PCR products were visualized by electrophoresis through a

1.2% (w/v) agarose gel in the presence of TAE buffer (pH 8.6, section 2.3.1.7) with a

154

Hyperladder I (Bioline UK) at 100V for 1 h and PCR products purified using the

QIAquick ® PCR purification Kit (Qiagen UK) according to the manufactures instructions.

3.3.8 ITS rDNA sequencing and identification

Purified samples were sequenced by the in house sequencing facility

(Faculty of Life Sciences, University of Manchester) using an ABI Prism® 3100

Genetic analyser (Applied Biosystems USA). Sequencing results were viewed using

FinchTV v1.4.0 software (Geospiza Inc.). Nucleotide sequences were interrogated using the BLAST (Basic Local Alignment search Tool) algorithm at the National

Center for Biotechnology Information website (www.NCBI.nlm.nih.gov) and phylogenetic trees were compiled by the Mega 5 alignment tool and CLUSTALW programme with 500 bootstrapping value.

3.3.9 Extraction and amplification of community genomic DNA

Genomic DNA was extracted from biomass obtained from the surface of PU coupons and from compost using Powersoil DNA isolation kit (MO BIO

Laboratories, USA) according to the manufacturer’s instructions except that during the lysis step tubes were homogenized twice for 30 s at 5000 rpm. The concentration of eluted DNA was measured using a NanoDropTM 1000 and samples stored at -

20°C until required.

3.3.10 Analysis of the fungal community by DGGE

For community analysis by DGGE, genomic DNA was amplified using primers GM2 (5′-CTGCGTTCTTCATCGAT-3′) and JB206

(5′CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGGAAGTAAA

155

AGTCGTAACAAGG-3′) (modified from White et al., 1990) which amplify the ITS

1 region of fungal rDNA as previously described (Cosgrove et al., 2007). A 50 µl

PCR mix was made using 1xNH4 buffer, 1.5 mM MgCl2, 200 µM of each dNTPs,

0.5U Taq polymerase (Bioline UK), 0.2 µM primers and BSA 100 µg/ml (New

England Biolabs UK). DNA template was Ca. 50-100 ng per PCR reaction. To minimize PCR bias three replicates for each sample were run. The PCR conditions employed consisted of initial denaturation at 94°C for 5 min; 20 cycles at 94°C for

30 s, annealing for 30 s at 59° to 49°C with the annealing temperature reduced by

0.5°C at each cycle and extension at 72°C for 45 s with a final extension at 72°C for

5 min (Cosgrove et al., 2007). PCR products were verified by electrophoresis through a 1.5% (w/v) agarose gel with a Hyperladder I (Bioline UK) at 100V for 1 h.

Amplicons from three replicates were pooled and purified using a Qiagen purification kit according to the manufacturer’s instructions and DNA concentration measured using a NanoDropTM 1000 machine (Thermofisher Scientific Inc., USA).

DGGE was performed according to Cosgrove et al., (2007) except that gels were run for 16 h. DGGE amplicons were separated using the D-Code universal mutation detection system (Bio-Rad, UK). Gels measured 16 X 16 cm and contained 10%

(v/v) bisacrylamide in 1X TAE buffer with a denaturing gradient of 25% to 55%. Ca.

500 µg of DGGE PCR products per lane were run in 1x TAE buffer at 60°C for 16 h at 42 V. Gels were stained with SyberGold (Molecular Probes, Netherlands) according to the manufacturer’s instructions and visualised under UV light. The position and intensity of the bands was quantified using Phoretix ID Advanced V5.00 software (Nonlinear dynamics, UK). Principal Component Analysis was performed using Multi-variate Statistical Package (MVSP) version 3.13g (copyright © 1985-

156

2003 Kovach computing services). In order to normalise and allow comparisons to be made between different gels, pooled PCR products amplified from the DNA of

Malbranchea cinnamomea, Lichtheimia sp., Aspergillus fumigatus, Acremonium flavum and Thermomyces lanuginosus were included as a species ladder on every gel.

3.3.11 Analysis of the fungal community by TRFLP

For fungal community analysis by TRFLP analysis, fungal ITS1-5.8s-ITS2 rDNA region was amplified using the fluorescent labelled primers, ITS5-FAM (fam-

GGAAGTAAAAGTCGTAACAAGG) and ITS4- HEX (hex-

TCCTCCGCTTATTGATATGC). PCR reaction conditions were the same as DGGE

PCR mixture, with the PCR regime of: initial denaturation at 94°C for 10 min, then

35 cycles of 94°C for 1 min, 54°C for 1 min and 72°C for 1 min, with a final extension 72°C for 10 min. Amplified DNAs were verified by electrophoresis through a 1.5% (w/v) agarose gel with a Hyperladder I (Bioline UK) at 100V for 1 h.

Three PCR amplicon replicate samples were pooled and purified by ethanol precipitation. Ethanol (100%) was added to the pooled DNA samples in 2:1 ratio and incubated overnight at -20 °C for better yield. Tubes were centrifuged at 13000rpm for 30 minutes at 0 °C and the supernatant discarded. Ice cold 70% (v/v) ethanol was added to the pellet and gently mixed by inverting the tubes several times, centrifuged at 13000 rpm at 0°C for 10 min, the supernatant discarded and the DNA pellet air died overnight at room temperature. The DNA pellet was dissolved in 10 µl of sterile

DPEC water and DNA concentration measured using a NanoDropTM 1000

(Thermofisher Scientific Inc., USA). To produce a mixture of variable length end- labelled ITS rDNA fragments, PCR products (1.5 µg) were digested with 0.5 U Hhal

15 7

(Fermentas, UK) at 37°C overnight in Tango buffer (10 µl). Digested products (0.5

µl) were mixed with 9.25 µl Hi-Di formamide (ABI, UK) and 0.25 µl GS500LIZ

(ABI, UK) in a 96 well PCR plate and products separated and analysed in-house on an ABI Prism® 3100 Genetic analyser (Applied Biosystems USA) (Kawasaki et al.,

2011). The size of the fragments was determined using Peak Scanner™ Software

Version 1.0 (Applied Biosystems), using peak height detection of 50 fluorescent units The output was further analyzed with online T-Align programme

(http://inismor.ucd.ie/~talign/) to generate a consensus profile of TRFs sizes between the technical duplicates and to compare the profiles between the samples (Smith et al., 2005). Shannon index (Ĥ) and evenness (e) were measured for each T-RFLP according to Tiquia (2005). Principal Component Analysis was employed to cluster the samples based on the presence, absence and relative intensities of TRFs from each TRFLP pattern using MVSP version 3.13g (copyright © 1985-2003 Kovach computing services).

3.3.12 4 Analysis of the fungal community by 454 pyrosequencing

Extracted DNA samples that were used for TRFLP and DGGE were subjected to 454 pyrosequencing. Fusion primers (Figure 3.1) were designed with an adapter sequences (blue) and key “TCAG” (red) with ITS1-5.8S-ITS2 sequences. Forward primers also had one of the 10 bp length unique Roche multiplex identifiers (MID-,

Table 3.1) to tag PCR amplicons from each sample. Sequence of forward primer

(Primer A): 5’-CCATCTCATCCCTGCGTGTCTCCGACTCAG-{MID}-

{GGAAGTAAAAGTCGTAACAAGG}-3’.Reverse primer (Primer B): 5’-

CCTATCCCCTGTGTGCCTTGGCAGTCTCAG-{

158

TCCTCCGCTTATTGATATGC }-3’. Unidirectional longer reads were used for sequencing because it is more informative than obtaining a mixture of forward and reverse reads (Anonymous, 2011a).

PCR reagents were used from PCR high fidelity PCR system (Roche, USA). A 50 µl reaction mixture contained 1xNH4 buffer, 1.5 mM MgCl2, 200 µM of each dNTPs, 1

U polymerase, 0.2 µM primers (HPLC purified), DMSO 2ul and BSA 100 µg/ml

(New England Biolabs) with approx. 50-100 ng of DNA template per PCR mixture.

The PCR regime used was the same as for TRFLP, except that 30 cycles were used to avoid chimera formation. PCR products were separated on a 1% (w/v) agarose gel containing 0.005% ethidium bromide at 100 V for 45 min. Gel bands (575-700 bp) were excised using a sterile rectangular blade and DNA was extracted using Gel extraction kit (Qiagen, UK) according to the manufacturer’s instructions and purified by column purification (PCR purification kit, Qiagen UK). Quantification was performed in-house using a fluorometer-based Qubit system. After individual quantification, amplicons were pooled in equal concentration with a final concentration of 10 ng/µl in 10 µl mixture. Pooled samples were subjected to 454

Titanium platform pyrosequencing by the Centre of Genomic Research, University of Liverpool, UK.

3.3.13 Bioinformatics and statistical analysis

159

UNITE/QIIME 12_11 ITS reference database, any sequence which did not hit the reference database were subsequently clustered denovo. A representative set of

OTUs was generated and the taxonomy of each set was assignment of OTUs was performed using same database (UNITE+INSD). OTUs defined as 97% sequence similarity threshold were used for rarefaction curve, PCA and beta diversity analysis

(chao1 and Shannon- weaver index) using QIIME.

3.3.14 Statistic analysis

To determine the statistical significance, data were subjected to Analysis of

Variance with the significance threshold set at a P value of 0.05 (JMP basic version

160

Figure 3.1: Schematic representation of designed primers for unidirectional reads of amplicon from community genome. There was an adapter (blue) and key “TCAG” (red) sequence, forward primers also had a unique Roche multiplex identifiers, MID (yellow). (Adapted from CGR Guideline 454 amplicon pyrosequencing, centre of Genomic research pyrosequencing, university of Liverpool, accessed May’2013).

161

Table 3.1: Sequences of the 10 MIDs used to tag PCR amplicons from each sample for 454 pyrosequencing.

ID MID Sequence ID MID Sequence

MID1 ACGAGTGCGT MID6 ATATCGCGAG

MID2 ACGCTCGACA MID7 CGTGTCTCTA

MID3 AGACGCACTC MID8 CTCGCGTGTC

MID4 AGCACTGTAG MID10 TCTCTATGCG

MID5 ATCAGACACG MID11 TGATACGTCT

(Adapted from Roche pyrosequencing guidelines, accessed June 2013).

162

3.4 Results

3.4.1 Effect of composting on the macroscopic and microscopic features of PU coupons

Polyester and polyether PU coupons were buried in the centre and 0.4 m from the surface of a 10 m tall compost heap during its formation at a commercial composting plant (TEG group, UK). The compost pile was formed from output material from an in silo composting vessel composed of domestic green and food waste and had an initial temperature of 68°C. Coupons were recovered from the surface and centre of the compost pile, analysed macroscopically and under ESEM and compared to unburied controls. Unburied polyester PU coupons had a smooth surface and were transparent (Figure 3.2A). Polyester PU coupons recovered from the centre of the compost pile had become brown in colour but the surface remained smooth (Figures 3.2B,D and F). However, polyester PU coupons recovered from the surface of the compost pile had become opaque, with deep cracks visible toward the centre of the coupons. Discolouration and cracks were evident after 14 days and became deeper after 28 days (Figure 3.2 C,E and G respectively). Surface examination of polyether PU coupons showed no visible surface changes or discolouration (Section 3.7 Figure A3.2).

Environmental scanning electron microscopy of the surface of unburied PU coupons and of coupons recovered from the centre and surface of the compost pile are shown in Figures 3.3 A, B and C respectively. Coupons recovered from the centre of the pile showed slight wearing and the presence of a bacterial population on

163 the surface but no fungal hyphae or cracks were observed (Figure 3.3 B). By contrast, coupons recovered from the surface of the compost pile had deep cracks and pits on the surface and was associated with the presence of bacteria, yeast and filamentous fungi (Figure 3.3 C).

3.4.2 Effect of composting on the percentage elongation and tensile strength of PU

Temperature variation at the surface and centre of the compost pile was measured at 0, 2, 14 and 28 days. After 28 days, temperature at the centre remained similar (67°C) to that measured on the first day (68°C) but at the surface it had decreased to 42°C (Figure 3.4A).

Physical degradation of PU coupons was analysed by measuring percentage elongation at break (Figure 3.4B) and the tensile strength (force to break) (Figure

3.4C). Polyester PU recovered from the surface of the compost pile after 14 days, showed a significant reduction (p<0.05) in both percentage elongation (ca. 50%) and tensile strength (>70%) but no further significant reduction was observed after 28 days (p>0.05). PU coupons recovered from the centre of the compost pile showed a similar reduction in tensile strength but no significant change (p>0.05) was found in percentage elongation (Figure. 3.4 B and C).

Polyether PU dumb-bells after 28 days from surface and centre of compost pile showed no significant change (P>0.05) in either tensile strength or percentage elongation (50.1 ±4 to 43.3 ±3 MPa and 1277±65 to 1192±41 respectively)(results not shown).

164

The rate of degradation of polyester PU was enhanced when buried under compost. Polyester PU buried at the surface of the compost pile showed biodegradation with >70% loss in tensile strength and 50% loss in percentage elongation but the samples buried at centre of the compost showed partial degradation because only tensile strength was affected. Polyether PU showed degradation but at a very slower rate.

3.4.1 Effect of composting on the glass transition and melting temperatures of PU

Differential Scanning Calorimetry (DSC) was performed to determine if compost burial influenced the glass transition temperature (Tg) or melting temperature (Tm) of polyester PU. Tg is related to the amorphous region of the semi- crystalline structure of PU and is the temperature at which the semi-crystalline regions change from a hard crystalline to a rubber-like semi-crystalline state

(Sandler, et al., 1998). There was a significant (P<0.05) decrease seen in the Tg value of PU coupons recovered from centre of the compost pile after 14 days from -

22.49°C to -35.16°C but did not fall further after 28 days, however, Tm was not significantly affected (Table 3.2). By contrast, coupons from the surface of the compost pile showed only a small decrease in Tg after 14 and 28 days (-24.46°C and

-24.39°C respectively) but a large and significant (P<0.05) decrease in Tm from

151.4° to 113.7° and 118.7°C after 14 and 28 days respectively.

165

A B A

B C

D E

F G

Figure 3.2: The effect of compost burial on the macroscopic features of PU coupons. PU coupons were buried 0.4 m from the surface and at the centre of a 10 m tall compost pile and recovered periodically over 28 days. PU coupons were recovered after 2 days from the centre (B) and surface (C), 14 days from the centre (D) and surface (E) and after 28 days from the centre (F) and surface (G) of the compost pile. The unburied control coupon prior to burial is shown in (A) for comparison. PU coupons from the surface of the compost pile showed deep cracks and discolouration but PU coupons from centre of the compost pile showed discolouration only.

166

A a b

B C

Figure 3.3: The effect of compost burial on the surface features of PU coupons under ESEM. PU coupons were buried 0.4 m from the surface and at the centre of a 10 m tall compost pile and recovered after 28 days from the centre (B) and the surface (C) of the compost pile. An unburied control coupon is shown for comparison (A). Coupons were buried in the compost pile during its formation at a commercial composting plant (TEG group, UK). The compost pile was formed from output material from an in silo composting vessel composed of domestic green and food waste and had a temperature of 68°C. Coupons from the centre of the pile had bacteria and the surface showed weathering but PU coupons from the surface of the compost pile displayed surface cracking, fungal hyphae and bacterial cells.

167

80 H A

C) ° 40

Temperature Temperature ( 0

1400 1200

B 1000

800 600

400

elongation elongation Percentage Percentage 200 0 80 70 60 50 C 40 30 20 10 0

Tensile strength Tensile strength (Mpa) 0 2 14 28 Incubation period (Days) Figure 3.4: Changes in the temperature of the compost pile and in the tensile strength of buried PU coupons over a 28 day period. Coupons were buried at 0.4 m depth from the surface and at the centre of a compost pile at a commercial composting site. (A) Change in temperature 0.4 m from the surface (■) and at the centre (■) of the compost pile. (B) Percentage elongation of PU coupons buried 0.4 m from the surface (■) and at the centre (■) of the compost pile. (C) Tensile strength of PU coupons buried 0.4 m from the surface (■) and at the centre (■) of the compost pile. Unburied control coupons were incubated at 55°C (■). Temperature decreased at the surface of compost pile and PU coupons buried at the surface showed significant reduction in tensile strength and % elongation. PU buried at the centre of the compost pile showed reduction in tensile strength only. Results are the means of 9 replicates ± SEM.

168

3.4.2 Total viable fungal count and percentage impranil degrading fungi recovered from the surface of PU coupons during composting

In order to enumerate the total viable fungi and the percentage of PU degraders on the surface of PU coupons following compost burial, biomass from the surface of PU coupons was serially diluted and plated onto potato dextrose agar

(PDA, for total fungal counts) and impranil agar (PUA, to observe halo formation) and incubated at 37°, 45°, 50° and 55°C (Table 3.3). Total CFUs recovered from the surface of PU buried at the surface of the compost pile increased when plates were incubated at 37° or 45°C. However, when plates were incubated at 50 or 55°C, CFUs initially increased but then fell after 28 days. This could be due to either the decrease in the temperature of the compost pile or increase in the number of uncultivable fungal isolates. CFUs recovered from the PU surface buried at the centre of the compost pile increased over 28 days at all incubation temperatures, however the number of CFUs recovered was much lower compared to those recovered from PU buried at the surface of the compost pile. When CFUs recovered from the surface of

PU coupons were enumerated on PUA, all colonies produced a zone of clearing suggesting that all were capable of degrading impranil. When compared to CFUs enumerated on PDA, the number of CFUs recovered on impranil medium was generally lower although this was not always the case (Table 3.3).

Enumeration of fungi from polyester PU buried at the surface of the compost pile suggests that the population size increased over the 28 day period of burial and as the temperature of the surface of the compost pile decreased. PU coupons from the centre of the compost pile did not show any significant variation in population size throughout the period of incubation.

169

Table 3.2: Differential calorimetry data of PU coupons recovered from surface and centre of the compost pile.

PU coupons Tg (°C) Tm (°C) recovered from Surface Centre Surface Centre compostUnburied pile PU -22.5±0.5 151.4±5

Day 14 -24.5±0.9 -36.1±2 113.7±7 150.9±3.5

Day 28 -24.4±1.5 -36.0±3 118.7±4 150.9±8

Glass transition temperature (Tg) and melting temperatures (Tm) of PU coupons recovered after 14 and 28 days from the surface and centre of the compost piles. A significant (P<0.05) decrease was seen in the Tg of PU buried in the centre of the compost pile for 14 and 28 days and a significant decrease (P<0.05) was observed in Tm of PU buried at the surface of the compost pile for 14 and 28 days. Results are the means of 3 replicates ± SEM

170

3.4.3 Total viable and PU degrading fungal colony count from the compost

In addition to determining the CFUs on the surface of PU coupons, the fungal CFUs were also determined in the compost surrounding the coupons (Table

3.4). With the exception of plates incubated at 37°C few fungi were detected before

14 days, fungi were recovered on both PDA and impranil medium at similar numbers indicating a prevalence of thermophilic and thermotolerant fungi at both the surface and centre of the compost pile.

3.4.4 Identification of isolates recovered from compost and surface of PU coupons

Fungal isolates growing on PUA were difficult to distinguish morphologically due to lack of pigmentation and sporulation on this medium.

Therefore, isolates were subcultured onto PDA and grown at the same temperature as the original PUA plate for up to 5 days. Subsequently, colonies were grouped into a number of different colony phenotypes and randomly picked isolates from each group subjected to 28s rDNA sequencing and sequences used to interrogate the

NCBI database. The identification of the isolates was further confirmed with ITS1-

5.8s-ITS2 rDNA region (Table 3.6).

Diversity in the fungal community recovered from PU coupons buried at the surface of compost pile was dependent on the incubation temperature. At 37°C,

Acremonium flavum and Candida rugosa, were consistant mesophilic species with dominant Arthrographis kalrae on day 28. At 45°C on day 2, the biomass obtained from the surface of buried PU coupons was dominated by Aspergillus spp. and on day 28, a mixed community of Leichthemia sp., Aspergillus fumigatus with occasional isolates of Malbranchea cinnamomea and Emericella nidulans were

171 recovered. Malbranchae cinnamomea and Aspergillus fumigatus were also recovered from 50°C but the major population at 50° and 55°C was Thermomyces lanuginosus at all time points (Table 3.5).

PU coupons recovered from the centre of the compost pile were mainly dominated by Acremonium flavum and Candida rugosa when plates were incubated at 37°C, Aspergillus fumigatus at 45°C and Thermomyces lanuginosus at 50°and

55°C on day 28 (Table 3.5).

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Table 3.3: Changes in the total viable fungal CFUs from PU coupons (cfu/cm2) buried at the surface and the centre of a 10 m high compost pile.

Total viable and Impranil clearing fungi recovered from the surface of PU coupons

Surface of the compost pile Centre of the compost pile

Burial Viable Impranil Viable Impranil Temp time count clearing count clearing (°C) (days) (CFU/cm2) (CFU/cm2) (CFU/cm2) (CFU/cm2)

37 2 <1 <1 <1 <1

14 165±12 178±27 15 ±<1 8 ±<1

28 1131±58 263±93 60±49 6±3

45 2 7±2 7±<1 <1 <1

14 5183±54 125±20 17±8 15±11

28 2007±230 386±72 49±56 4±5

50 2 5±1 <1 <1 <1

14 3335±1482 93±12 19±<1 <1

28 1944±232 211±49 93±8 19±4

55 2 2±<1 <1 <1 <1

14 2306±943 132±1 22 ±2 <1

28 415±97 137±<1 31±5 13±5

Results are the means of 3 replicates ± SEM. All colonies recovered on impranil agar (PUA) produced halos indicating degradation.

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Table 3.4: Changes in the total viable fungal CFUs at the surface and centre (cfu/g) of a 10 m high compost pile.

Total viable and Impranil clearing fungi in compost

Surface of the compost pile Centre of the compost pile

Total viable Impranil Total viable Impranil Temp Time count clearing count clearing (°C) (day) (CFU/g) (CFU/g) (CFU/g) (CFU/g)

37 0 >1 >1 >1 >1

2 >1 >1 >1 >1

14 6.0±0.5X104 >1 1.4±0.3X104 1.3±0.3X104

28 10±8X104 25±3X104 28±1X104 5.5±0.9X104

45 0 10±2X104 3.2±0.7X104 1.7±0.3X104 3.7±0.7X104

2 10±4X104 19±1X104 17.5±0.4X104 4.7±0.4X104

14 200±30X104 2.5±0.1X104 6±2X104 2.7±0.08X104

28 80±10X104 13±2X104 14±4X104 8.6±0.2X104

50 0 10±2X104 2.6±0.8X104 11±0.22X104 2.7±0.8X104

2 12±4X104 2.7±0.3X104 65±0.77X104 65±1X104

14 90±10X104 28±4X104 6±2X104 45±2X104

28 84±9X104 6.2±0.55X104 140±20X104 4.0±0.2X104

55 0 2.2±0.3X104 2.9±0.8X104 1.5±0.3X104 2.7±0.8X104

2 7±3X104 17±0.3X104 5±1X104 2.3±0.2X104

14 4±2X104 4.6±0.7X104 1.0±0.7X104 1.2±0.1X104

28 28±2X104 13±1X104 4.5±0.1X104 2.7±0.4X104

Results are the means of 3 replicates ± SEM. All colonies recovered on impranil agar (PUA) produced halos indicating degradation.

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Table 3.5: Frequency of isolates at recovered from polyester PU coupons buried at the surface of a compost heap.

Day (temp. °C) Frequency Closest match 2 (37) +++ Candida rugosa 2 (45) +++ Aspergillus fumigatus 2 (50) +++ Thermomyces lanuginosus 2 (55) +++ Thermomyces lanuginosus 14 (37) +++ Arthrographis kalrae 14 (37) + Acremonium flavum 14 (37) + Candida rugosa 14 (45) +++ Aspergillus fumigatus 14 (45) +++ Lichtheimia sp. 14 (45) + Malbranchea cinnamomea 14 (50) +++ Thermomyces lanuginosus 14 (50) + Aspergillus fumigatus 14 (50) + Malbranchea cinnamomea 14 (55) +++ Thermomyces lanuginosus 28 (37) +++ Arthrographis kalrae 28 (37) + Acremonium flavum 28 (37) + Candida rugosa 28 (45) +++ Aspergillus fumigatus 28 (45) +++ Lichtheimia sp. 28 (45) + Emericella nidulans 28 (45) + Malbranchea cinnamomea 28 (50) +++ Thermomyces lanuginosus 28 (50) + Aspergillus fumigatus 28 (50) + Malbranchea cinnamomea 28 (55) +++ Thermomyces lanuginosus

Coupons were buried at 0.4 m depth from the surface of a 10 m tall compost pile during its formation at a commercial composting plant (TEG group, UK). Isolates were recovered from the surface of the PU coupons 2, 14 and 28 days of burial by plating onto PUA agar and incubating at 37°, 45°, 50° and 55°C. Representatives of different morphotypes were identified by 28s rDNA sequencing. Relative abundance of each species is shown; + occasional, ++ frequent and +++ dominant.

175

Table 3.6: Percentages homology of putative PU degrading fungal isolates for

28s rDNA region with Blast database.

Isolates Closest match in NCBI % Accession ID database number homology

Teg 1 Emericella nidulans 100 JQ966573

Teg 2 Lichtheimia sp. 99 JQ966578

Teg 11 Aspergillus fumigatus 100 JQ966572

Teg 5 Candida rugosa 100 JQ966580

Teg 8 Thermomyces lanuginosus 100 JQ966575

Teg 16 Acremonium flavum 99 JQ966574

Teg 14 Malbranchea cinnamomea 99 JQ966576

Teg 17 Arthrographis kalrae 99 JQ966577

Fungal isolates showed 99-100% homology with the published sequences (accession number shown) from the NCBI database. With the exception of Acremonium flavum and Arthrographis kalrae, identities of the isolates were further confirmed by ITS1- 5.8s-ITS2 rDNA region sequencing.

176

Figure 3.5: Phylogenetic analysis of the isolated putative PU degrading fungi recovered from the surface of PU coupons buried in compost. V3 (28s rDNA) sequences from the isolates were aligned to closely related species from the NCBI database and used to form a neighbour joining phylogenetic tree (Bootstrap with 1000 samples). Isolates from this study are marked (■).

177

3.4.5 Comparison of the compost community with the community on the surface of PU

DGGE and TRFLP were used to study temporal changes in the fungal community in the compost pile and on the surface of buried polyester PU coupons.

DGGE revealed that in the compost, the fungal population diversity was very low initially when the pile was first formed from the output material of the in silo vessel

(day 0) when temperature was above 65°C and only three distinct bands were seen.

The number of bands recovered from the centre of the compost pile remained similar over 28 days and the temperature remained above 65°C. However, as the temperature declined at the surface of the pile, the number of visible DGGE bands from day 14 to

28 increased (Figure 3.6).

Comparing the DGGE bands obtained from the compost samples and from the PU coupons, the diversity (band number) was lower on the PU surface. While some bands in the compost and from PU coupons co-migrated (bands 4, 5 and 6), some of the bands from PU coupons were not detected in the surrounding compost.

When comparing the compost with PU coupon surface, more bands were observed in the former sample. There were a few bands that were not seen in the compost (bands

7, 8, 9 and 10). Band 1 and 3 appeared to be the most common among the compost samples (Figure 3.6).

Boolean data (presence/absence of bands on DGGE gel, 0 or 1) of the community fingerprints obtained after DGGE were subjected to Principal

Component Analysis (Figure 3.7). Samples clustered into three 3 groups. Group I contained community profiles from the compost on day 0, and from compost and PU coupons at the centre of the pile at 14 and 28 days, group II was from compost

178 samples from the surface of the compost pile (14 and 28 days) and group III contained samples obtained from the PU coupons recovered from the surface of the compost pile (14 and 28 days).

TRFLP profiles of the fungal communities were aligned to identify shared and unique components of the fungal community using the T-align programme.

Fungal diversity within the fungal community was determined using the Shannon index and Evenness statistical indices based on the number of T-RFs and their relative abundance (proportional peak area) in each sample (Table 3.7). The number of TRFs recovered from compost and from the surface of PU coupons taken from the centre of the compost pile remained relatively constant over 28 days. In contrast, the number of TRFs detected from the compost and PU coupons buried at surface of the compost pile increased over 28 days. Interestingly, after day 28 the number of TRFs detected from the PU coupons buried at the surface of the compost pile was two-fold higher than that detected in the surrounding compost (Table 3.7).

A significant increase in the Shannon index was observed in the compost samples from the surface of the pile from day 2 to day 14 corresponding to an increase in the diversity of the population and correlated with a decrease in the temperature of the compost. Shannon index from compost on day 0 and from the surface of PU coupons taken from the centre of the compost pile was ≤ 2.5 indicating a stable fungal population over the course of study. The assemblage of population

(evenness) and diversity on PU coupon remained fairly stable. An increase in the

Evenness value (E) was observed from day 2 to day 28 in surface compost samples indicating that different members of the community are stabilising and getting

179 proportional over time. Shannon index from the PU coupons decreased slightly from day 14 to day 28 while evenness increased (T able 3.7).

Comparison of the unique TRFs in TRFLP profiles of compost and PU coupons from the surface of compost pile, showed 69.4 and 73% of the TRFs were only found on the surface of PU coupons and were not shared with compost at that particular time. A total of 61 unique TRFs were detected only on PU (Table 3.8).

TRFLP profiles were subjected to Principal Component Analysis based on the presence, absence and peak heights of TRFs (Figure 3.8). TRFLP pattern generated using HhaI for microbial community diversity, accounted for 55% of total variance. While the fungal community profiles of compost clearly differed from the

PU samples buried at the surface of the compost pile after 14 and 28 days, there was less separation amongst the other samples.

180

Figure 3.6: DGGE profiles from the fungal community in compost and from the PU coupons. DGGE profile of fungal communities from the compost and the surface of buried PU coupons. Distinct community profiles were evident in compost and surface of PU coupons. (0,14 and 28 indicates the day of incubation, “c” represents centre of the compost pile, “s” surface of the compost pile, “cPU” and “sPU” indicates community from PU coupons recovered from centre and surface of the compost pile, respectively, while M represents reference marker. M comprised of DNA obtained from Malbranchea cinnamomea, Acremonium flavum, Lichtheimia sp., Aspergillus fumigatus and Thermomyces lanuginosus. Bands represented with numeric are common between samples.

181

DGGE PCA for fungal community

1.0

0.8 Group II 0.6

0.4 Group I

0.2 Axis 2 (25%)

0.0

-0.2 Group III

-0.4

-0.6 -0.6 -0.4 -0.2 0.0 0.2 0.4 0.6 0.8 1.0

Axis 1 (34%)

Figure 3.7: Principal component analysis of the fungal community DGGE profiles from compost and from the surface of PU coupons. Coupons were buried in the centre and at 0.4 m depth from the surface of a 10 m tall compost pile during its formation at a commercial composting plant (TEG group, UK). Total genomic DNA was extracted periodically and subjected to DGGE and analysed by PCA. Fungal community profiles obtained from compost during the formation of the pile Day 0 (●), from the surface of compost pile day 14 () and day 28 ( centre of the compost pile day 14 ( ), from the PU coupons buried at the surface of the compost pile (day 14 (▲) and day 28 (▲ coupons buried at the centre of the compost pile (day 14 (▼) and day 28 (▼ Transition in community was observed with the maturation of compost at the surface of compost pile.

182

Table 3.7: Statistical analysis of the community profile analysed by TRFLP.

Day TRFs Shannon (std.dev) Evenness (std.dev)

Surface Centre Surface Centre Surface Centre

Compost

0 26 2.16±0.56 0.58±0.05

2 8 8 1.8 ±0.76 1.70 ±0.37 0.65 ±0.09 0.73 ±0.05

14 42 8 2.58 ±0.35 2.12 ±0.47 0.65 ±0.04 0.87 ±0.02

28 32 12 2.78 ±0.35 2.18 ±0.26 0.75 ±0.03 0.83 ±0.04

Surface of PU coupons

14 85 10 2.81 ±0.52 1.99 ±0.42 0.59 ±0.05 0.79 ±0.08

28 63 13 2.71±0.41 1.50 ±0.46 0.61 ±0.03 0.72 ±0.05

Estimation of number of TRFs, Shannon index and Evenness of the fungal community profile from compost and surface of PU coupons buried at the surface and centre of compost pile. The number of TRFs and Shannon index increased with the period of incubation in compost and PU coupons recovered from the surface while no significant increase was seen in the ones recovered from centre of compost pile.

183

TRFLP PCA score for fungal community

3.00

2.40

1.80

1.20

0.60

Axis 2(18%) Axis Axis 2(18%) 0.00

-0.60

-1.20

-1.80 -1.80 -1.20 -0.60 0.00 0.60 1.20 1.80 2.40 3.00

AxisAxis 1(37%) 1(37%)

Figure 3.8: PCA analysis of the T-RFLP profiles of the fungal community in compost and on the PU surface. PU coupons were buried in the centre and at 0.4 m depth from the surface of a 10 m tall compost pile during its formation at a commercial composting plant (TEG group, UK). Total genomic DNA was extracted periodically and subjected to TRFLP and analysed by PCA. Fungal community profiles obtained from compost during the formation of the pile (● Day 0); compost from surface of compost pile after 2 (), 14 (), and 28 () days; compost from centre of compost pile after 2 (), 14 (  surface of the compost pile after 14 (▲) and 28 (▲ co ▼).

184

Table 3.8: comparison of the shared and unique number of TRFs.

Time in No. of total TRFs No. of unique TRFs % of unique TRFs No. of dominant TRFs Ratio of dominant

days compost PU compost PU compost PU compost>1% peak areaPU (% compostTRFs PU

14 (S) 42 85 16 59 38.1 69.4 13peak (73.6) area covered)12 (66.9) 0.3 0.1

28 (S) 32 63 15 46 46.9 73.0 12(52.6) 8 (74.1) 0.4 0.1

14 (C) 8 10 4 6 50 60 6(94) 9 (95.1) 0.9 1

28 (C) 12 13 7 8 58.3 61.5 12(94) 11 (95.2) 0.9 0.9

TRFs generated from Hhal digested compost and biomass PU coupons from the surface of the compost pile. The number of total TRFs obtained from PU coupons was two-fold than from compost. Ratio of number of TRFs with % intensity >5 to total number was calculated and their combine peak area is presented, that contributes > 50% of the total peak intensity.

1 85 185

40 day 0 compost

35 day 14 compost

30 day 14 PU coupons

25 day 28 compost

20 day 28 PU coupons

Fluorescence Fluorescence intensity 10

35.3

79.6

39.43

47.08

53.01

57.78

63.09

70.76

75.05

88.28

95.17

104.22

114.06

120.88

124.85

133.98

140.74

149.86

159.12

169.31

259.7

184.11

192.95

199.67

203.99

208.75

245.24

254.01

409.4

264.26

273.26

279.76

295.15 350.25 TRFs 379.29

Figure 3.9: Electropherogram of TRFs detected compost and from the surface of PU coupons. Arrows shows unique TRFs only obtained on the surface of PU coupons with >1% intensity. Communities from compost; day 0 (), day 14 () and day 28 () and from PU coupons; day 14 () and day 28 () buried at the surface of the compost pile.

1 86 186

3.4.6 454 pyrosequencing

454 pyrosequencing was used to analyse in detail the diversity and identity of the fungi present at the surface of the compost pile and on the surface of PU coupons after 28 days. ITS1-5.8s-ITS2 region of the community was sequenced and sequences obtained were denoised and classified into unique OTUs. Total 55,430 sequences were obtained after denoising the samples, which were divided into 49 different OTUs that were shared between compost on day 0 and 28 and on the surface of PU. Rarefaction curves levelled off and showed saturation of the sequences (Figure 3.10). The Shannon index after 28 days on the surface of PU surface was less with more Chao1 then compost (Table 3.9). The representative species in each sample were clustered into 8, 15 and 22 species (Section 3.7 Table

A3.1).

Sequences were associated with three phyla; Ascomycota, Basidiomycota and

Zygomycota with the Ascomycota the most dominant phylum (Figure 3.11) On day 0,

Candida ethanolica (91.9%) was the most dominant fungal species present.

However, compost after day 28 days the dominant OTU detected was an unidentified fungal clone A (56.2%), which was also the dominant species detected on the PU surface (80.4%). The compost community contained 8 OTU’s with at >1% with 7

OTU’s at less than 1%, while the community on the surface of PU only contained 4

OTU’s at >1% but has 20 other OTU’s present at <1% (Figure 3.11). Thermomyces lanuginosus, Aspergillus fumigatus, Arthrographis kalrae and Emericella nidulans that were cultured from the surface of PU were also detected by pyrosequencing but at a low abundance.

187

The scatter plot for principal component analysis of the 454 sequencing data samples were divided into 2 groups, group 1 as compost day 0 and group 2 contained day 28 compost and the surface of PU community (Figure 3.12).

188

Figure 3.10: Rarefaction curve from 454 pyrosequencing data. Number for observed specie in compost at day 0 (−−−), day 28 (−−−) and surface of PU on day 28 (−−−) levelled off suggesting no of sequences library.

189

Teg Day 0 Teg day 28 compost A B C Teg day 28 PU

Candida ethanolica  (91.9%) Unidentified fungal clone A  (56.2%) Unidentified fungal clone A  (80.4%) Candida rugosa  (5.5%) Sordariales sp. (15.1%) Arthrographis kalrae  (9.7%) Sordaria fimicola  (1.9%) Scedosporium proloficans  (10.6%) Unidentified fungal clone B  (4.4%) Others  (0.9%, 5 OTU’s) Arthrobotrys vermicola  (6.3%) Thermomyces lanuginosus  (1.0%) Pseudallescheria boydii  (4.9%) Others  (4.5%, 20 OTU’s) ) Unidentified fungal clone B  (2.9%) Talaromyces thermophilus  (1.8%) Thermomyces lanuginosus  (1.8%) Others  (1.0%, 7 OTU’s)

Figure 3.11: Fungal diversity and relative abundance from compost and from the surface of PU by 454 pyrosequencing. PU coupons were buried at 0.4 m depth from the surface of a 10 m tall compost pile during its formation at a commercial composting plant (TEG group, UK). Genomic DNA was extracted from samples and ITS1-5.8S-ITS2 PCR amplicons subjected to 454 pyrosequencing. Data represent the % of the total returned sequence for each identified OTU. A) Fungal population from compost during formation of the compost pile (Day 0), B) fungal population from compost from the surface of the compost pile after 28 days C) fungal population from the PU coupons buried at the surface of compost pile after 28 days. OTU’s present at <1% of the returned sequence were grouped as “others” (■) with the number of OTU’s in parenthesis.

1 190

Figure 3.12: Scatter plot of principal component analysis from 454 pyrosequencing of the fungal community from compost and from the surface of PU. Fungal community profile from compost on day 0 (●), compost on day 28 (●) and from the surface of PU coupons () on day 28.

191

Table 3.9: Statistical estimation of pyrosequencing data.

Samples Shannon-index Observed species Chao1

Compost Day 0 0.6 8.2 9.8

Compost Day 28 2.1 15.6 17.2

PU surface Day 28 1.2 22.0 28.7

Shannon index, number of observed species and Chao 1 estimation of fungal community from 454 pyrosequencing obtained on Day 0 and day 28 compost and on day 28 from surface of PU coupons.

192

3.3 Discussion

Commercial composting has become an effective strategy for diverting green and food waste from landfill sites as it is cost effective, benign and has little negative impact on the environment (Chiellini et al., 1996; Shah, Hasan, Hameed, et al., 2008). In the previous chapter (chapter 2), we found that polyester polyurethanes, which have previously been shown to be susceptible to microbial degradation in soils, particularly by fungi, (Barratt et al., 2003; Cosgrove et al., 2007) are susceptible to degradation by the mesophilic and thermophilic/tolerant fungal population in compost. Moreover, despite the reduction in biodiversity at 45° and

50°C due to the selection of only thermophilic and thermotolerant fungi, degradation still occurred at similar rates to mesophilic (25°C) temperatures. However, only one fungal species was predominantly isolated from the PU surface at 45° and 50°C

(chapter 2). To further investigate the potential for polyurethanes to be degraded in composting waste treatment facilities, PU coupons were buried 0.4 m from the surface (surface samples) and at the centre (centre samples) of a 10 m tall compost pile. It was formed from green and food waste that had passed through an in silo- composting vessel and recovered after 2, 14 and 28 days.

Previous studies have reported a loss in tensile strength of polyester PU of ca. 60% after 1.5 months burial in lab based soil microcosms (Barratt et al., 2003) while a 95% loss in tensile strength was observed after 5 months burial in neutral and acidic soil in the environment (Cosgrove et al., 2007). In this current study, we demonstrated that the tensile strength of polyester PU decreased by ca. 75% when buried for 4 weeks in a commercial composting pile and was associated with deep

193 cracks and pits in the PU surfaces (Figure 3.3). Chiellini et al., (1996) studied the biodegradation of blends of PCL and PET under different environmental conditions including full scale composting, soil burial, bench scale aerobic degradation and exposure to axenic cultures and estrolytic enzymes. Their data suggested that among the environments compared, full-scale composting was the most efficient method for degradation. Composting offers advantages over soil for biodegradation including an enhanced opportunity for co-metabolism (Williams et al., 1992) and a rich organic matrix (Kästner and Mahro, 1996). Tuomela et al., (2000) reviewed biodegradation of lignin in a compost environment and concluded that elevated temperatures during the thermophilic phase of composting are essential for rapid degradation of lignocellulosic complexes and are mainly degraded by thermophilic microfungi and actinomycetes at the optimum temperature of 40-50°C.

The samples from the middle and the surface of the compost pile both showed discolouration and a decrease in the tensile strength though more severe deterioration and cracking was evident from the surface samples. While the decrease in surface samples was associated with a decrease in percentage elongation at break

(a measure of softness of the material), there was no change in percentage elongation at break for samples buried in the middle of the pile. DSC also revealed differences in the PU coupons from the two burial points (Table 3.2). The glass transition temperature (Tg) was significantly lower in samples buried at the centre of the compost pile compared to the surface buried samples and unburied controls (ca. -

36°C compared to –ca -24.5°C and -22.5°C respectively). Tg is the temperature at which the polymer changes from a rigid, hard crystalline state to a soft semi- crystalline state containing unordered amorphous regions and is affected by a number

194 of factors including the presence of side groups and interactions between polymer chains (Sandler et al, 1998). Therefore, samples at the centre of the pile underwent different molecular changes to those at the surface where Tg decreased only slightly.

This was also apparent in the differences measured in the melting temperature (Tm) of the polymer. While surface buried PU showed a reduction in Tm compared to unburied controls, samples buried at the centre were unaffected. Tm represents the temperature at which the semi-crystalline state becomes wholly amorphous and suggests that the crystalline regions of the polymer of samples buried at the surface were affected more than samples buried at the centre of the compost pile. These differences between the samples at the centre and surface of the compost pile probably reflect that due to the temperature difference between the centre and the surface of the compost pile, bacteria and actinomycetes will be actively growing at the centre while at the surface where temperatures decline, fungi will be more active.

Huang and Roby (1986) reported that PU degradation proceeded in a selective manner, with the amorphous regions being degraded prior to crystalline regions

(reviewed by Howard 2012). The amorphous regions in the structure of PU contribute to the tensile strength, stiffness and Young’s modulus, while the crystalline regions contribute to elongation, elasticity, softness and degradability

(Krasowska et al., 2012).

We selected windrow phase of the composting where the compost begins the maturation stage after the in silo phase as the samples would be undisturbed and provide an opportunity to compare the variability in temperature with time and also the clear demarcation in the active and dormant state of fungi at the surface and centre of the pile respectively. At the centre of the compost pile, the temperature

195 remained >60°C, a temperature at which fungi are unable to grow, whereas at the surface of the compost pile, temperatures fell to 42°C after 28 days and abundant fungal growth was apparent (Finstein and Morris, 1975; Beffa et al., 1996;

Ryckeboer, Mergaert, Coosemans, et al., 2003). When the temperature cools down, the fungal population starts recolonizing the compost (Ryckeboer et al., 2003 a & b).

Hassen et al., (2001) studied the relationship of physicochemical conditions and microbial population present in compost of municipal solid waste. They reported that at ca.60°C, the count of yeasts and filamentous fungi decreased from 106 to

103cells/g, thus while unable to grow and causing loss in viability, significant numbers survive and are able to recolonise the compost once the temperature begins to cool.

On the PU coupons from the surface of the compost pile, the fungal viable count increasing significantly with time and we were able to identify 7 different fungal isolates. The ITS region of rDNA was amplified for the identification of isolates (Peay et al., 2008) but only a few isolates were identified on the basis of the sequence of the 28s rDNA region because of the failure of ITS primers. The most dominant isolate obtained from plates incubated at 50-55°C was Thermomyces lanuginosus (Kane and Mullins, 1973) (synonym Humicola lanuginosus) and is a dominant thermophile from mushroom and wheat straw compost (Chang and Hudson

1967; Anastasi et al., 2005). T. lanuginosus has been well characterized and known to produce a variety of thermostable proteoleases, amylases, xylanases, ureases and lipases (Allen et al., 1999; Howard and Hilliard, 1999b; Ruiz et al., 1999;

Maheshwari et al., 2000; Matsumiya et al., 2010) some of which have been reported to participate in PU degradation. Aspergilllus fumigatus (Kane and Mullins 1973,

196

Anastasi et al., 2005) and Emericella have previously been isolated as potential PU degraders (Barratt, et al., 2003). 454 pyrosequencing data suggested that

Thermomyces, Emericella, Lichthemia and Aspergillus are present but as a minor proportion of the total population, while the dominant fungus was identified as

Arthrographis kalrae (Figure 3.11). Arthrographis kalrae is a common pathogen ubiquitously found in soil and compost and has reported as having urease activity

(Liu, 2011).

The diversity of fungal populations were analysed by DGGE, TRFLP and

454 pyrosequencing, suggested that distinct communities are present on PU coupons which are not found in compost at the same time point suggesting selection for PU degraders. Previously, Cosgrove et al., (1997) compared the community of PU coupons with soil and reported that PU community is the subset of the soil population because of the enrichment of the specie that colonizes and /or degrades

PU.

On a comparison of two techniques, DGGE and TRFLP, the latter was found to be more sensitive as has been previously reported (Moeseneder et al., 1999; Nunan et al., 2005; Smalla et al., 2007). PCA scatter plots from the two techniques (Figure

3.7 and 3.8) however gave similar results, suggesting that with prolonged incubation and decrease in temperature, diversity increased due to the proliferation of thermophilic and thermotolerant fungi.

This is the first study to compare the dynamics of population on the basis of culture dependant and independent techniques in compost and on the surface of polyester PU coupons. This study suggests that the rate of degradation is enhanced

197 under thermophilic and early maturation stage of commercial composting and that thermophilic and thermotolerant fungi have the capacity to cause significant PU degradation. Thus, existing commercial composting systems designed to treat green and food waste have a potential to be modified to allow the input of PU’s as a possible alternative waste disposal strategy.

198

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Hassen, a, Belguith, K., Jedidi, N., Cherif, A., Cherif, M., Boudabous, A., 2001. Microbial characterization during composting of municipal solid waste. Bioresource Technology 80, 217–225. Hopewell, J., Dvorak, R., Kosior, E., 2009. Plastics recycling: challenges and opportunities. Philosophical Transactions of the Royal Society of London. Series B, Biological sciences 364, 2115–2126. Howard, G.T., 2012. Polyurethane biodegradation, in: Singh, S.N. (Ed.), Microbial Degradation of Xenobiotics. Springer Berlin Heidelberg, Berlin, Heidelberg, pp. 371–394. Howard, G.T., Hilliard, N.P., 1999. Growth of Pseudomonas chlororaphis on a polyester polyurethane and the purification and characterization of a polyurethanase- esterase enzyme. International Biodeterioration and Biodegradation 43, 7–12. Huang, S.J., Roby, M.S., 1986. Biodegradable polymers poly(amide-urethanes) [1]. Journal of Bioactive and Compatible Polymers 1, 61–71. Jayasekara, R., Harding, I., Bowater, I., Lonergan, G., 2005. Biodegradability of a selected range of polymers and polymer blends and standard methods for assessment of biodegradation. Journal of Polymers and the Environment 13, 231–251. Kane, A.B.E., Mullins, J.T., 1973. Thermophilic fungi in a municipal waste compost system. Mycologia 65, 1087–1100. Kästner, M., Mahro, B., 1996. Microbial degradation of polycyclic aromatic hydrocarbons in soils affected by the organic matrix of compost. Applied Microbiology and Biotechnology 44, 668–675. Kawasaki, A., Watson, E.R., Kertesz, M. a., 2011. Indirect effects of polycyclic aromatic hydrocarbon contamination on microbial communities in legume and grass rhizospheres. Plant and Soil 358, 169–182. Krasowska, K., Janik, H., Gradys, A., Rutkowska, M., 2012. Degradation of polyurethanes in compost under natural conditions. Journal of Applied Microbiology 125, 4252–4260. Liu, D., 2011. Arthrographis, in: Molecular Detection of Human Fungal Pathogens. CRC press. USA, p. 167. Maheshwari, R., Bharadwaj, G., Bhat, M.K., 2000. Thermophilic fungi: their physiology and enzymes. Microbiology and Molecular Biology Reviews 64, 461– 488. Matsumiya, Y., Murata, N., Tanabe, E., Kubota, K., Kubo, M., 2010. Isolation and characterization of an ether-type polyurethane-degrading micro-organism and analysis of degradation mechanism by Alternaria sp.Journal of Applied Microbiology 108, 1946–1953.

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Moeseneder, M.M., Arrieta, J.M., Muyzer, G., Winter, C., Herndl, G.J., 1999. Optimization of terminal-restriction fragment length polymorphism analysis for complex marine bacterioplankton communities and comparison with denaturing gradient gel electrophoresis. Applied and Environmental Microbiology 65, 3518– 3525. Morton, L.H.G., Surman, S.B., 1994. Biofilms in Biodeterioration-a review. International Biodeterioration and Biodegradation 203–221. Nunan, N., Daniell, T.J., Singh, B.K., Papert, A., Mcnicol, J.W., Prosser, J.I., 2005. Links between plant and rhizoplane bacterial communities in grassland soils, characterized using molecular techniques. Applied and Environmental Microbiology 71, 6784–6792. Oehlmann, J., Schulte-Oehlmann, U., Kloas, W., Jagnytsch, O., Lutz, I., Kusk, K.O., Wollenberger, L., Santos, E.M., Paull, G.C., Van Look, K.J.W., Tyler, C.R., 2009. A critical analysis of the biological impacts of plasticizers on wildlife. Philosophical Transactions of the Royal Society of London. Series B, Biological sciences 364, 2047–2062. Peay, K.G., Kennedy, P.G., Bruns, T.D., 2008. Fungal Community Ecology : Bioscience 58, 799–810. Plastics – the Facts 2012. An analysis of European plastics production , demand and waste data for 2011, 2012. . Reeder, J., Knight, R., 2010. Rapidly denoising pyrosequencing amplicon reads by exploiting rank-abundance distributions. Nature Methods 7, 668–669. Ruiz, C., Hilliard, N.P., Howard, T., 1999. Purification and characterization of two polyurethanase enzymes from Pseudomonas chlororaphis. International Biodeterioration and Biodegradation 43, 43–47. Ryckeboer, J., Mergaert, J., Coosemans, J., Deprins, K., Swings, J., 2003. Microbiological aspects of biowaste during composting in a monitored compost bin. Journal of Applied Microbiology 94, 127–137. Ryckeboer, J., Mergaert, J., Vaes, K., Klammer, S., 2003. A survey of bacteria and fungi occurring during composting and self-heating processes. Annals of Microbiology 53, 349–410. Sandler, S.R., Karo, W., Bonesteel, J., Pearce, E.M., 1998. Polymer synthesis and characterisation: A laboratory manual. Academic Press, Cambridge UK. Shah, A.A., Hasan, F., Hameed, A., Ahmed, S., 2008. Biological degradation of plastics: a comprehensive review. Biotechnology Advances 26, 246–265. Smalla, K., Oros-Sichler, M., Milling, A., Heuer, H., Baumgarte, S., Becker, R., Neuber, G., Kropf, S., Ulrich, A., Tebbe, C.C., 2007. Bacterial diversity of soils assessed by DGGE, T-RFLP and SSCP fingerprints of PCR-amplified 16S rRNA

201 gene fragments: do the different methods provide similar results? Journal of Microbiological Methods 69, 470–479. Smith, C.J., Danilowicz, B.S., Clear, A.K., Costello, F.J., Wilson, B., Meijer, W.G., 2005. T-Align, a web-based tool for comparison of multiple terminal restriction fragment length polymorphism profiles. FEMS Microbiology Ecology 54, 375–380. Tiquia, S.M., 2005. Microbial community dynamics in manure composts based on 16S and 18S rDNA T-RFLP profiles. Environmental Technology 26, 1101–1113. Tuomela, M., Vikman, M., Hatakka, A., It, M., 2000. Biodegradation of lignin in a compost environment : a review. Bioresource Technology 72, 169–183. Webb, J.S., Nixon, M., Eastwood, I.M., Greenhalgh, M., Robson, G.D., Handley, P.S., 2000. Fungal colonization and biodeterioration of plasticized polyvinyl chloride. Applied and Environmental Microbiology 66, 3194–3200. White, T.J., Bruns, T.D., Lee, S., Taylor, J., 1990. Analysis of phylogenetic relationshipd by amplificaion and direct sequencing of ribosomal RNA genes, in: Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocol: a Guide to Methods and Applications. Academic Press, New York, US, pp. 315–322. Williams, R.T., Ziegenfuss, S., Wayne, E., Way, W., Chester, W., 1992. Composting of explosives and propellant contaminated soils under thermophilic and mesophilic conditions. Journal of Industrial Microbiology 9, 137–144.

202

3.5 Appendix A3

Figure A3.1: The 10 m compost pile used for burying PU coupons at the TEG group commercial compost site, Todmorden, UK. Initial temperature of the pile was 68°C.

203

Figure A3.2: The effect of compost burial on the macroscopic features of polyether PU coupons. Polyether PU coupons were buried at a depth of 0.4 m (surface) and 7 m (centre) from the surface of a 10 m tall compost pile and recovered after two weeks. A) Unburied control. B) Surface coupon C) Centre coupon.

204

Table A3.1: Percentages of OTUs and their taxonomic assignments obtained by 454 pyrosequencing. Day 0 (2983) % k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;f__Incertae_sedis;g__Candida;s_ethanolica 91.89 k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;f__Incertae_sedis;g__Candida;s_rugosa 5.50 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__Sordariaceae;g_soradria;s_fimicola 1.68 Unidentified fungi (a) 0.77 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Penicillium;s_paneum 0.07 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;Other 0.03 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Scedosporium 0.03 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__unidentified;g__unidentified 0.03 Day 28 (17770) Unidentified fungi (a) 56.17 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__unidentified;g__unidentified 15.03 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Scedosporium;s_proloficans 10.57 k__Fungi;p__Ascomycota;c__Orbiliomycetes;o__Orbiliales;f__Orbiliaceae;g__Arthrobotrys;s_vermicola 6.34 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g_Pseudallescheria;s_boydii 4.89 unidentified fungal clone (b) 2.92 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Talaromyces;s_thermophillum 1.81 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Incertae_sedis;g__Thermomyces;s_lanuginosus 1.24 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;Other;Other 0.32 k__Fungi;p__Ascomycota;c__Leotiomycetes;o__unidentified;f__unidentified;g__Scytalidium;s_thermophillum 0.32 k__Fungi;p__Basidiomycota;c__Agaricomycetes;o__Agaricales;f__Agaricaceae;g__unidentified 0.19 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Petriella 0.14 k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Incertae_sedis;f__Eremomycetaceae;g__Arthrographis;s_kalrae 0.03 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g_fusarium;s_solani 0.03 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Aspergillus;s_fumigatus 0.01 Day 28 PU (34660) Unidentified fungal clone A 80.35 k__Fungi;p__Ascomycota;c__Dothideomycetes;o__Incertae_sedis;f__Eremomycetaceae;g__Arthrographis;s_kalrae 9.71 Unidentified fungal clone B 4.39

05 205

k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Incertae_sedis;g__Thermomyces;s_lanuginosus 1.04 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Emericella;s_rugulosa 0.86 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Talaromyces;s_thermophillum 0.78 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Aspergillus;s_fumigatus 0.72 k__Fungi;p__Basidiomycota;c__Agaricomycetes;o__Agaricales;f__Agaricaceae;g__unidentified 0.69 k__Fungi;p__Ascomycota;c__Leotiomycetes;o__unidentified;f__unidentified;g__Scytalidium;s_thermophillum 0.37 k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;f__Dipodascaceae;g__Galactomyces 0.30 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__unidentified;g__unidentified 0.17 k__Fungi;p__Basidiomycota;c__Tremellomycetes;o__Tremellales;f__Trichosporonaceae;g__Trichosporon 0.16 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;Other 0.14 k__Fungi;p__unidentified;c__unidentified;o__unidentified;f__unidentified;g__unidentified 0.09 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Scedosporium 0.08 k__Fungi;p__Zygomycota;c__Incertae_sedis;o__Mucorales;f__Lichtheimiaceae;g__Lichtheimia 0.04 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Microascales;f__Microascaceae;g__Microascus 0.03 k__Fungi;p__Ascomycota;c__Saccharomycetes;o__Saccharomycetales;f__Incertae_sedis;g__Candida 0.03 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Eurotium;s_sp. 0.02 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Hypocreales;f__Nectriaceae;g_fusarium;s_solani 0.01 k__Fungi;p__Ascomycota;c__Pezizomycetes;o__Pezizales;f__Incertae_sedis;g__Cephaliophora 0.01 k__Fungi;p__Ascomycota;Other;Other;Other;Other 0.00 k__Fungi;p__Ascomycota;c__Eurotiomycetes;o__Eurotiales;f__Trichocomaceae;g__Penicillium;s_paneum 0.00 k__Fungi;p__Ascomycota;c__Leotiomycetes;o__Incertae_sedis;f__Incertae_sedis;g__Geomyces;s_pannorum 0.00 k__Fungi;p__Ascomycota;c__Sordariomycetes;o__Sordariales;f__Chaetomiaceae;g__Corynascus 0.00

2 06

206

Chapter 4: Effect of polyurethane on fungal compost communities

Urooj Zafar and Geoff D. Robson

Faculty of Life Sciences, Michael Smith Building, University of Manchester, Manchester M13 9PT, UK

207 4.1 Abstract

Polyurethanes are a highly versatile group of plastics that have a broad range of applications and in Europe alone annual production exceeds 3 Million tonnes. Polyester PU’s are particularly susceptible to microbial degradation where fungi are the major organisms involved. Previous studies have investigated the degradation of PU’s in composting systems and shows promise as an alternative route for waste disposal. This study investigated the influence that diverting PU waste might have an effect on the indigenous fungal compost community. Compost was amended with a microparticulate PU dispersion and solid PU and the temporal change in total viable counts and in community diversity using TRFLP assessed at

25°, 45° and 50°C. Both polyester and polyether PU dispersions increased total viable counts at all temperatures but was much higher with polyester PU whereas solid polyester PU had no impact. In the presence of microparticulate dispersion of polyester PU, in addition to Aspergillus fumigatus and Themomyces lanuginosus, which were also present in unamended compost, Humicola grisea and Malbranchea cinnamomea were also identified as putative PU degraders. PCA analysis of

TRFLP’s revealed that the fungal community in compost amended with either polyester or polyether microparticulate dispersions formed a discrete group separate from unamended compost at each temperature. However, addition of solid polyether

PU caused only a temporary shift in the community which returned to resemble the unamended community after 8 weeks suggesting the temporary perturbation may have been caused by the physical disruption of the compost by the solid PU rather than the presence of PU itself. The data suggest that while microparticulate PU’s can shift the community diversity and enrich for particular species due to the high surface

208 area to volume ratio and homogeneous distribution, solid PU waste is unlikely to have a major impact on the surrounding fungal compost community.

209 4.2 Introduction

Contamination of the environment by plastics and their disposal has become increasingly problematic worldwide. In Europe in 2011, 41% (10.3 Mtonne) of total plastic waste was directed to landfills (Plastic Europe 2011). However, shortage of landfill capacity and long-term risks of contamination of soils and groundwater by additives and breakdown products (Hopewell et al., 2009; Oehlmann et al., 2009) have led to alternative waste management strategies to be developed including recycling and incineration although these alternatives also pose significant challenges. For example, many plastics are not amenable for recycling (for example thermoset plastics) or contain a range of polymers and additives which increase recycling costs considerably (Hopewell et al., 2009). In recent years, green and domestic waste has increasingly been diverted to commercial composting sites as it is relatively cost efficient, rapid, environmentally benign and reduces the burden on landfill sites (Areikin et al., 2012). Composting has also been considered as a potential route for the degradation of some synthetic polymers. For example, Sasek et al., (2006) described the potential for the degradation of aliphatic and aromatic co- polyesters during composting, while Krasowska et al., (2012) has studied the biodegradation of PU under natural compost piles in a 24 month study and reported significant biodegradation. While fungi have previously been shown to be the principal microbial group involved in PU degradation in soil (Barratt et al., 2003;

Cosgrove et al., 2007) and many studies have previously isolated fungi associated with PU degradation (Pathirana and Seal, 1984; Bentham et al., 1987; Kay et al.,

1991; Crabbe et al., 1994; Barratt et al., 2003; Cosgrove et al., 2007, 2010), few studies have investigated the potential for composting to biodegrade PU’s.

210 Previously, we investigated the impact of compost burial on PU degradation under controlled laboratory conditions (Chapter 2) and investigated the extent of PU degradation in a commercial composting process (Chapter 3). In order to investigate the impact of introducing PU into composts on the fungal communities, in this study we introduced both solid and a liquid dispersion of PU into compost and study its impact on fungal communities at 25°, 45° and 50°C using both culture based and non-culture-based techniques (TRFLP). We report that the addition of both solid PU induced a temporary shift while PU dispersion caused both an increase in the fungal population and its diversity.

211 4.3 Materials and Methods

4.3.1 Compost moisture content

Fresh mature compost (temperature ca. 65°C) was obtained from The

Compost Shop, (Orrel Hill Lane, UK) and sieved through a 4 mm mesh prior to use.

Percentage water holding capacity (WHC) of the compost was measured following the modified protocol of Alef and Nanniperi, (1995). Five replicates (10 g) were saturated with water, weighed and then dried at 55°C to constant weight to determine the percentage WHC, calculated using the following equation:

( ) ( )

( )

Moisture content of the compost was determined according to Barratt et al.,

(2003). Five replicates (10 g) were weighed (fresh weight) and then dried at 55°C to constant weight (dried weight). Percentage moisture content was calculated using the following equation:

( ) ( )

( )

Percentage moisture content of compost was adjusted to 40%, by the addition of sterile distilled water with a plant spray. For solid polyester PU amendment, PU beads (Ellastollan® 685 A10, 1.21 g/cm3) were mixed with compost to give a final percentage (w/w) of 0%, 10%, 30% and 50% (w/w). To amend the compost with a

PU dispersion, 10 g of Impranil DLN and Impranil DLU (liquid dispersion of polyester and polyether polyurethane respectively, Bayer, UK) was mixed

212 thoroughly with 90 g of compost. Unamended and amended composts were incubated in plastic containers (16 x 12 x 5 cm) at 25°, 45° and 50°C for 12 weeks and the moisture content maintained between 37% and 40% by weighing the containers every two days and adding sterile distilled water with a plant spray to replace water lost through evaporation.

4.3.2 Microbial viable count from compost

Amended and unamended compost samples (1g) were serially diluted in phosphate buffer saline (PBS) and plated out onto compost extract agar (CEA)

(Barratt et al., 2003, section 2.3.1.4) and PUA (Crabbe et al., 1994, section 2.3.1.5) for the enumeration of total fungal viable count and percentage polyurethane fungal degraders. Filter sterilised chloramphenicol was added to the agar to a final concentration of 50 µg/ml to inhibit bacterial growth.

4.3.3 Extraction, amplification and purification of genomic DNA from the isolated PU degrading fungal colonies

Colonies that produced halos on PUA were sub cultured onto PDA. Total genomic DNA was extracted from 5-7 days old fungal colonies, according to Feng

(2010) except that 0.5 g of glass beads (0.5 mm diameter) were used instead of a single bead and mycelium was homogenized twice for 30s at 5000 rpm. Isolates were identified according to Webb et al., (2000) using the rDNA Internally Transcribed

Spacer Region (ITS) with universal primers ITS1 (5’-

TCCGTAGGTGAACCTGCGG-3’) and ITS4 (5’-TCCTCCGCTTATTGATATGC-

3’) (White et al., 1990). The reaction mixture contained the DNA sample (20-100 ng), primers (0.2 µM), MgCl2 1.5 mM, 1x NH4 reaction buffer, 200 µM of each of dNTPs and 1U Taq Polymerase (Bioline UK). The PCR consisted of initial

213 denaturation 94°C for 3 min, then 35 cycles, denaturation at 94°C for 1 min, annealing at 56°C for 1 min and extension at 72°C for 1 min. Final extension 72°C for 3 min. PCR products of ITS1, 5.8S and ITS2 rDNA sized ca. 496-576 bp.

Amplified DNAs were verified by electrophoresis through a 1.2% (w/v) agarose gel with a Hyperladder I (Bioline UK) at 100V for an hour. PCR product was purified using the QIAquick ® PCR purification Kit (Qiagen UK) following the manufacturer’s instructions.

4.3.4 DNA sequencing and identification of isolated PU degrading fungal colonies

Purified samples were sequenced by the in house sequencing facility using an

ABI Prism® 3100 Genetic analyser (Applied Biosystems USA). Sequencing results were viewed using FinchTV v1.4.0 software (Geospiza Inc.). The nucleotide sequence was interrogated using the BLAST (Basic Local Alignment search Tool) algorithm at the National Center for Biotechnology Information website

(www.NCBI.nlm.nih.gov). The phylogenetic trees were compiled by the Mega 5 alignment tool and CLUSTALW programme with 500 bootstrapping value.

4.3.5 Extraction of community DNA from compost

Genomic DNA was extracted from triplicate compost samples using a

Powersoil DNA isolation Kit (MOBio Labs, Inc.) according to the manufacturer’s instructions except that during the lysis step tubes were homogenized twice for 30 s at 5000 rpm. The concentration of eluted DNA was measured using a NanoDropTM

1000 and samples stored at -20°C until required.

214 4.3.6 Amplification of the community DNA from compost for TRFLP analysis

Fungal ITS1-5.8s-ITS2 rDNA genes were amplified from compost community DNA using the fluorescent labelled primers, ITS5-FAM (fam-

GGAAGTAAAAGTCGTAACAAGG) and ITS4- HEX (hex-

TCCTCCGCTTATTGATATGC). A 50 µl PCR mix was made using 1xNH4 buffer,

1.5 mM MgCl2, 200 µM of each dNTPs, 0.5U Taq polymerase (Bioline UK), 0.2 µM primers and BSA 100 µg/ml (New England Biolabs). DNA template was approx. 50-

100 ng per PCR mixture. To minimize PCR bias three replicates for each sample were run. PCR regime was: initial denaturation at 94°C for 10 min, then 35 cycles of

94°C for 1 min, 54°C for 1 min and 72°C for 1 min, with a final extension 72°C for

10 min. Amplified DNAs were verified by electrophoresis through a 1.5% (w/v) agarose gel with a Hyperladder IV (Bioline UK) at 85V for an hour. The three replicates were pooled and purified by ethanol precipitation. Final DNA pellet was dissolved in 10 µl of sterile DPEC water and DNA concentration measured using a

NanoDropTM 1000 (Thermofisher Scientific Inc., USA).

Amplified DNAs were verified by electrophoresis through a 1.5% (w/v) agarose gel with a Hyperladder IV (Bioline UK) at 85V for 1 h. Amplicons from three replicates were pooled and purified using Qiagen purification kit. The final concentration of 30 µl eluted DNA was measured using a NanoDropTM 1000 machine and the ND-10003.1.0 software (Thermofisher Scientific Inc., USA).

To produce a mixture of variable length end-labelled ITS rDNA fragment, digestion mixture (10 µl) was prepared with labelled PCR products (1.5 µg), restriction endonucleases Hhal 0.5 U (Fermantas, UK), Tango buffer in MilliQ water

215 and incubated at incubated at 37°C overnight (as per manufacturer’s instruction). The digested products (0.5 µl) were mixed with 9.45 µl Hi-Di formamide (ABI) and 0.05

µl GS500LIZ (ABI) in a 96 well PCR plate, analysed in-house on an ABI Prism®

3100 Genetic analyser (Applied Biosystems USA) (Kawasaki et al., 2011).

The size of the fragments was determined using Peak Scanner™ Software

Version 1.0 (Applied Biosystems), using peak height detection of 30 fluorescent units The output was further analyzed with online T-Align programme http://inismor.ucd.ie/~talign/ to generate a consensus profile of TRFs sizes between the technical duplicates and to compare the profiles between the samples (Smith et al., 2005). Shannon index (Ĥ) and evenness (e) were measured for each T-RFLP

(Tiquia, 2005). Principal Component Analysis was employed to cluster the samples based on the presence, absence and relative intensity of TRFs from each TRFLP pattern using MVSP version 3.13g (copyright© 1985-2003 Kovach computing services).

4.3.7 Statistic analysis

To determine the statistical significance, data were subjected to Analysis of

Variance with the significance threshold set at a P value of <0.05 (JMP basic version

216 4.4 Results

4.4.1 Fungal colony forming units (CFU) from PU amended compost

Total fungal counts (CEA) and total PU degrading fungal counts (PUA) were determined periodically over 12 weeks in compost, unamended and amended, with PU incubated at 25°, 45° and 50°C. In the absence of impranil, the total fungal count on CEA remained similar at 25°C but at 45° and 50°C 10 fold rise was seen, from week 0 to week 1 then remain stable till week 12 (Figure 4.1A). The fungal counts on impranil DLN agar (polyester PU dispersion) also showed the same trend over 12 weeks for each temperature except counts were ca. ten fold lower at 50°C.

The fungal counts on CEA and PUA indicate that the majority of fungi present were impranil DLN degraders.

When compost was amended with impranil DLN (liquid dispersion of polyester PU), total fungal counts recovered on CEA and on DLN PUA media were significantly (P<0.05) higher (ca. 100-1000 fold) compared to unamended compost for each temperature indicating the presence of impranil DLN increased fungal growth (Figure 4.1B). Again, counts recovered on DLN PUA compared to CEA indicated the majority of fungi recovered were PU degraders. Interestingly, while the total fungal counts on CEA at 25°C remained similar over time, the counts on DLN

PUA increased after 1 and 4 weeks.

For compost amended with impranil DLU (liquid dispersion of polyether

PU), no fungal counts were recovered on DLU PUA agar. However, total viable counts on CEA agar were higher then unamended compost at each temperature

(Figure 4.1 C). Thus, both impranil DLN and DLU appeared to be utilized for growth

217 increasing the total viable fungal population at each temperature although it was not possible to isolate any fungi on DLU agar. Colony count suggested impranil DLU does not have negative effect on fungal population.

Unlike the PU dispersion, the fungal count obtained from compost amended with PU beads did not show any significant difference (P>0.05) between unamended and PU bead amended compost at any temperature (Figure 4.2).

4.4.2 Identification of dominant fungal isolates enriched in compost amended with Impranil DLN

Addition of Impranil DLN increased the colony count significantly at 45° and 50 °C. At 45°C, two dominant isolates were recovered and identified as

Rhizomucor meihei and Aspergillus fumigatus. However, at 50°C, Themomyces lanuginosus was the dominant isolate recovered with Aspergillus fumigatus,

Malbranchea cinnamomea and Humicola grisea present in much lower numbers.

Corynascus vermicosus and Scytalidium thermphilum were occasionaly found but only on CEA media and were not recovered from PUA (Table 4.1). On unamended compost, only Aspergillus fumigatus at 45°C and Thermomyces lanuginosus at 50°C were recovered.

218 1.E+09 A 1.E+08 1.E+07 1.E+06 1.E+05 1.E+04 1.E+03 1.E+02

1.E+01 1.E+00 B 1.E+09 1.E+08 1.E+07 1.E+06 1.E+05 1.E+04 1.E+03 1.E+02 1.E+01

1.E+00 Fungalviable count(cfu/gm, log scale) C 1.E+09 1.E+08 1.E+07 1.E+06 1.E+05 1.E+04 1.E+03 1.E+02 1.E+01 1.E+00 0 1 4 8 12 Incubation period (weeks)

Figure 4.1: Enumeration of fungal viable counts recovered from compost amended with or without PU dispersion (impranil). Compost was incubated up to12 weeks at 25°C, 45°C or 50°C without (A) or after amendment with impranil DLN (liquid dispersion of polyester PU) (B) or impranil DLU (liquid dispersion of polyether PU) (C). Total fungal counts were enumerated on CEA and impranil degrading counts were enumerated on PUA following incubation at the same temperature as the compost. (CEA 25°C , 45°C  and 50°C , PUA 25°C , 45°C  and 50°C  . In (C), no colonies were recovered on impranil DLU agar plates.

219 1.E+09 1.E+08 1.E+07 1.E+06 1.E+05

1.E+04 scale) 1.E+03 1.E+02 1.E+01

viablefungal (cfu/gm,log count 1.E+00 0 4 8 12 0 4 8 12 0 4 8 12 25 45 50 Temperature (°C)

Figure 4.2: Enumeration of fungal viable counts recovered from compost amended with or without PU beads. Compost was incubated for up to12 weeks at 25°C, 45°C or 50°C without and after amendment 10%, 30% and 50% (w/v) polyester PU beads. Total fungal counts were enumerated on CEA and impranil degrading counts were enumerated on PUA following incubation at the same temperature as the compost. Unamended compost (figure 4.1A) suggested no significant difference with PU beads uamended compost at all temperatures. (10% PU CEA , 10% PU PUA , 30% PU CEA , 30% PU PUA , 50% PU CEA  and 50% PU PUA ).

220 Table 4.1: Isolates recovered from impranil amended compost incubated at 45° and 50°C.

Isolates Incubation Clearance on PUA

temperature (°C)

Aspergillus fumigatus 45 +

Rhizomucor meihei 45 -

Themomyces lanuginosus 50 +

Aspergillus fumigatus 50 +

Malbranchea cinnamomea 50 +

Humicola grisea 50 + Corynascus vermicosus 50 -

Scytalidium thermphilum 50 -

+ represents clearance and - represents undetectable clearance of impranil on PUA. The unamended compost only had Aspergillus fumigatus and Thermomyces lanuginosus at 45° and 50 °C.

221

4.4.3 Influence of impranil DLN and impranil DLU on the compost community profile

In order to investigate whether the addition of microparticulate polyester and polyether PU (impranil DLN and impranil DLU respectively) caused a change in the compost fungal community, the ITS1-5.8s-ITS2 region of rDNA was used for

TRFLP analysis and the position and height of TRFs generated compared in different samples by PCA (Figure 4.3). In unamended compost, the fungal population at 25°C grouped closely together indicating little change in the community profile, whereas a marked shift was seen when compost was incubated at 45° and 50°C after 1 week and thereafter little further change was seen. Addition of either polyester PU

(impranil DLN) or polyether PU (impranil DLU) caused a shift in the fungal community profiles in at all temperatures but clustered together at all time points indicating a rapid shift in the profiles in response to impranil but then remained constant over the twelve weeks. Addition of impranil DLN increased the Shannon index and the number of detected TRFs at 45° and 50°C but had little impact at 25°C whereas impranil DLU had no effect at any temperature (Table 4.2).

4.4.4 Influence of polyester PU beads on the compost community profile

The effect of polyester PU beads on the fungal community in compost was assessed with TRFLP technique after 4, 8 and 12 weeks of incubation at 25°, 45° and

50°C (Figure 4.3A, B and C respectively). At 25°C, fungal communities broadly separated into two groups. Group I contained communities from week 1 and week 4 and group II contained communities from week 8, week 12 and control compost samples (unamended) with the community on day 0 (Figure 4.3A). At 45°C and

222 50°C, the community of day 0 was distinct from the incubated compost, and again the amended compost at week 1 and 4 clustered separately from weeks 8, 12 and unamended compost although the difference between them was not as great as that seen at 25°C (Figure 4.3B and C respectively). Addition of PU beads had little impact on either the Shannon index or on the total number of TRFs detected compared to the unamended compost (Table 4.2).

223 TRFLP PCA score for compost amended with impranil at 25C A 7.0

5.6 01

04 4.2 Group II 08 2.8 Group I 012 1.4 00

u4 0.0 Axis 2 (4.4%) 2 Axis n4 -1.4 n8

n12 -2.8 u1 -4.2 n1

-5.6 u8

u12 -7.0 -7.0 -5.6 -4.2 -2.8 -1.4 0.0 1.4 2.8 4.2 5.6 7.0

Axis 1 (89.9%) TRFLP PCA score for compost amended with impranil at 45 C B 7.0

5.6 01

04 4.2 Group II 012 2.8 Group I 08

1.4 00

n1 0.0

Axis 2 (25.5%) 2 Axis u 12

-1.4 u 8

n 8 -2.8 u1 -4.2 u 4

-5.6 n 12

n 4 -7.0 -7.0 -5.6 -4.2 -2.8 -1.4 0.0 1.4 2.8 4.2 5.6 7.0

Axis 1 (52.3%)

224

C TRFLP PCA score for compost amended with impranil at 50C

7.0

5.6 01

012 4.2

04 2.8 Group I 08

1.4 00

0.0 Group II n4 Axis 2 (30.5%) 2 Axis u4 -1.4 n12 -2.8 u1

-4.2 n8

-5.6 u8

u12 -7.0 -7.0 -5.6 -4.2 -2.8 -1.4 0.0 1.4 2.8 4.2 5.6 7.0

Axis 1 (53.5%)

Figure 4.3: Principal component analysis of TRFLP profiles from fungal communities in compost amended with impranil DLN or DLU. Untreated compost (control) and compost amended with impranil DLN (polyester PU) or impranil DLN (polyether PU) were incubated for 12 weeks at 25°C (A), 45°C (B) or 50°C (C). Total genomic DNA was extracted periodically and subjected to TRFLP and the TRF position and peak height analysed by PCA. Unamended compost prior to incubation (day 0,●) and after 1 (♦), 4(♦), 8(♦) and 12(♦) weeks. Compost amended with DLN after 1 (▼), 4(▼), 8(▼) and 12(▼) weeks. Compost amended with impranil DLU after 1 (), 4(), 8() and 12() weeks. For each time point, three replicate samples were used and the TRFLP PCR amplicons pooled prior to analysis.

225 A 9.0

7.2

5.4

3.6

1.8 Group II Group I

0.0 Axis 2 (14.3%) 2 Axis

-1.8

-3.6

-5.4

-7.2

-9.0 -9.0 -7.2 -5.4 -3.6 -1.8 0.0 1.8 3.6 5.4 7.2 9.0

Axis 1 (64.9%)

B 9.0

7.2

5.4

3.6 Group II 1.8

0.0 Group I Axis 2 (18%) 2 Axis

-1.8

-3.6

-5.4

-7.2

-9.0 -9.0 -7.2 -5.4 -3.6 -1.8 0.0 1.8 3.6 5.4 7.2 9.0

Axis 1 (68%)

226

C 9.0

7.2

5.4

3.6 Group I 1.8 Group II

0.0 Axis 2 (31.6%) 2 Axis

-1.8

-3.6

-5.4

-7.2

-9.0 -9.0 -7.2 -5.4 -3.6 -1.8 0.0 1.8 3.6 5.4 7.2 9.0

Axis 1(37.8%)

Figure 4.4: Principal component analysis of TRFLP profiles from fungal communities in compost amended with polyester PU beads. Untreated compost (control) and compost amended with polyester PU beads were incubated for 12 weeks at 25°C (A), 45°C (B) or 50°C (C). Total genomic DNA was extracted periodically from the surrounding compost and subjected to TRFLP and the TRF position and peak height analysed by PCA. Unamended compost prior to incubation (day 0, ●) and after 1(), 4(), 8() and 12 () weeks. Compost amended with 10% (w/w) PU beads after 1(▼), (▼), (▼) and 12 (▼) weeks. Compost amended with 30% (w/w) PU beads after 1(▲), 4(▲), 8(▲) and 12(▲) weeks. Compost amended with 50% (w/w) beads after 1(), 4(), 8 () and 12 () weeks. For each time point three replicate samples were used and the TRFLP PCR amplicons pooled prior to analysis.

227

Table 4.2: Shannon index, Evenness and number of TRFs detected following TRFLP analysis of the fungal community in compost amended with either polyester PU beads, impranil DLN or impranil DLU.

25 °C 45 °C 50 °C Number Sample S I E TRF S I E TRF S I E TRF 1 0%0 2.8 0.8 43 2.8 0.8 43 2.8 0.8 43 2 0% w1 2.9 0.7 55.0 2.2 0.5 31.0 3.1 0.8 37.0 3 0% w4 3.2 0.8 60.0 2.2 0.5 43.0 3.2 0.8 46.0 4 0% w8 2.8 0.7 53.0 3.0 0.7 38.0 2.8 0.8 31.0 5 0% w12 3.0 0.7 50.0 2.0 0.5 39.0 2.4 0.7 28.0 6 10% w1 3.5 0.8 70.0 2.8 0.8 34.0 3.6 0.9 31.0 7 10% w4 3.8 0.8 67.0 2.8 0.7 47.0 3.1 0.8 49.0 8 10% w8 3.1 0.8 53.0 1.6 0.4 47.0 2.8 0.8 36.0 9 10% w12 3.1 0.7 74.0 2.2 0.5 40.0 2.7 0.7 39.0 10 30% w1 2.9 0.8 39.0 2.8 0.7 51.0 3.5 0.8 37.0 11 30% w4 3.2 0.7 65.0 2.7 0.7 41.0 2.9 0.7 37.0 12 30% w8 3.0 0.7 73.0 1.8 0.5 53.0 3.2 0.8 38.0 13 30% w12 3.4 0.7 77.0 2.7 0.6 52.0 3.0 0.7 33.0 14 50% w1 3.1 0.7 67.0 2.6 0.7 52.0 3.6 0.9 34.0 15 50% w4 3.2 0.7 77.0 2.7 0.7 40.0 2.4 0.7 37.0 16 50% w8 2.6 0.6 64.0 2.0 0.5 47.0 3.0 0.7 30.0 17 50% w12 3.1 0.7 61.0 2.0 0.5 49.0 3.1 0.8 37.0 18 DLNw1 3.3 0.8 67.0 3.5 0.8 70.0 ND ND ND 19 DLN w4 3.3 0.9 63.0 3.7 0.9 65.0 3.5 0.9 56.0 20 DLN w8 3.1 0.8 59.0 3.9 0.9 63.0 4.0 0.9 74.0 21 DLN w12 2.9 0.8 63.0 3.9 0.9 73.0 3.6 0.9 54.0 2w12w12 22 DLU w1 2.5 0.7 56.0 2.8 0.8 30.0 3.3 0.9 30.0 23 DLU w4 2.5 0.9 59.0 2.5 0.8 21.0 3.0 0.8 37.0 24 DLU w8 2.9 0.9 51.0 2.0 0.6 28.0 1.9 0.7 32.0 25 DLU w12 2.6 0.8 53.0 2.7 0.8 33.0 3.2 0.9 35.0

Untreated compost (control) and compost amended with 10%, 30% or 50% (w/w) polyester PU beads, impranil DLN or impranil DLU were incubated for 12 weeks at 25°, 45° or 50°C. Total genomic DNA was extracted periodically from the surrounding compost and subjected to TRFLP and the number of detected TRFs, Shannon index and Eveness determined. For each time point three replicate samples were used and the TRFLP PCR amplicons pooled prior to analysis. ND=not determined, SI=Shannon index, E=Evenness, TRF=number of detected Terminal restriction fragments.

228 4.5 Discussion

Previous chapters (chapters 2 and 3) have suggested that the native fungal community in compost has the capacity to degrade polyester PU under thermophilic and mesophilic conditions and that polyester PU also undergoes degradation under commercial composting conditions. Therefore, composting may offer an alternative waste treatment strategy for disposal of waste PU. While it has been shown in the previous chapters (chapter 2 and 3) that a diverse community different from the surrounding compost colonizes and degrades PU, if PU is to be diverted into commercial composting streams, it is important to understand what impact this will have on the indigenous fungal compost population. In this study, the effect of the addition of a microparticulate dispersion of polyester PU (impranil DLN), polyether

PU (impranil DLU) and polyester PU beads on the indigenous fungal compost community under thermophilic and mesophilic temperatures was investigated by studying changes in the total viable fungal population and in the diversity of the fungal community using TRFLP analysis.

Addition of impranil DLN and DLU increased the total viable fungal count in the compost at all temperatures by ca. 10 to 100 fold indicating utilization of the microparticulate polyester polyether PU as a carbon source (Figure 4.1B), whereas the total fungal viable count in unamended compost remained stable (Figure 4.1A).

Moreover, the increase in the fungal viable population was rapid and occurred within

1 week and for impranil DLN amended compost, began to decline after 12 weeks, suggesting a significant proportion of impranil DLN had been utilized. Fungal viable counts on impranil DLN PU agar were only slightly lower than total fungal viable

229 counts suggesting the majority of the fungi present have the capacity to degrade polyester PU even in the native unamended compost. Previous studies have also demonstrated that the majority of fungi in soils are also capable of degrading impranil DLN (Barratt et al., 2003; Cosgrove et al., 2007). However, Cosgrove et al.,

(2007) reported that the addition of impranil DLN alone to soil did not cause a significant increase in the total fungal viable count but did lead to a >10 fold increase if added together with yeast extract, while yeast extract alone had little effect. The dependency of presence of yeast extract for impranil DLN to affect growth may be due to a requirement for additional nutrients that were low in soil whereas compost is rich in organic residues and may be sufficient to allow impranil DLN utilization. The addition of DLN impranil to compost also changes the species isolated at 45°C and

50°C. In unamended compost, only A. fumigatus and Thermomyces lanuginosus were isolated at 45° and 50°C respectively, whereas when amended with impranil

DLN, other fungal species were also isolated suggesting enrichment. While impranil

DLU also caused an increase in the fungal viable count of 10 fold, no fungi were recovered on impranil DLU agar. However, impranil agars (either DLN or DLU) are mineral salts media with no additional complex nutrients, which may be required for impranil DLU utilization. Polyether PU has been reported to be far more recalcitrant to microbial degradation due to the ether linkages being more resistant to enzymatic hydrolysis (Darby & Kaplan, 1968; Filip, 1985; Rutkowska et al., 2002; Urgun-

Demirtas et al., 2007; Krasowska et al., 2012). Growth on impranil DLU agar plates may require far longer incubation periods to be detected.

In order to investigate whether the addition of microparticulate PU had an impact on the fungal community diversity, TRFLP was used to study any temporal

230 changes in the indigenous fungal population (Figure 4.3) following addition of impranil DLN and impranil DLU. In unamended compost, the fungal population at

25°C grouped closely together with day 0 indicating little change in the community profile, whereas a marked shift was seen when compost was incubated at 45° and

50°C due to the selection of thermtolerant and thermophilic fungi. This shift in the population occurred after 1 week and the population remained similar up to 12 weeks. Addition of polyester PU (impranil DLN) caused a marked and rapid shift in the fungal community profile in compost at all temperatures and clustered together, indicating that the community changed rapidly following impranil DLN addition and then remained stable and correlates with the rapid increase seen in the total viable counts. Thus impranil DLN causes both a rapid increase in the total viable fungal population and a shift in the community profile. Interestingly the community at week 12 in the presence of impranil DLN was positioned slightly away from the profiles at weeks 1, 4 and 8 and correlates with the slight decline seen in the total fungal viable counts and may indicate that at this stage, a significant proportion of the PU had been utilized and the community profile was changing in response

(Appendix 4.7 Figure A4.4). Surprisingly, impranil DLU (polyether PU) had a highly similar impact on the fungal compost community and clustered tightly with the impranil DLN communities despite having no significant impact on the total viable fungal count. While many studies report that polyether PU is relatively recalcitrant

(Darby & Kaplan, 1968; Filip, 1985; Rutkowska et al., 2002; Urgun-Demirtas et al.,

2007; Krasowska et al., 2012), and we reported previously that there was no loss in tensile strength in polyether PU coupons buried in compost for 12 weeks at 25°, 45°

231 or 50°C (Chapter 2), some studies have reported degradation (Matsumiya et al.,

2010).

When the impact of the addition of polyester PU beads on the fungal compost community was examined by TRFLP, at each temperature there was a shift in the community profile after 1 and 4 weeks, however at 8 and 12 weeks the community was similar to compost without beads (Figure 4.4). After 12 weeks of incubation, the PU beads were still intact and therefore the shift in the community away and then back to the unamended profile was not due to the complete degradation of the beads. Unlike the addition of microparticulate impranil DLN, addition of the PU beads had no significant impact on the total fungal viable count

(Figure 4.2.). The shift in the population due to the addition of beads at 1 and 4 weeks only may reflect the initial perturbation and disruption of the compost by the physical presence of the beads rather than due to any substrate impact and that by week 8, the community had recovered. A wide range of minor perturbations are known to rapidly but transiently effect natural soil bacterial communities although this has not been investigated previously for fungi (Bressan et al., 2008; Berga et al.,

2012). PU buried in soil and in compost rapidly becomes colonized by fungi which degrade the surface (Barratt et al, 2003; Cosgrove et al., 2007; 2010, Chapter 2 and

3), the fact that no permanent change occurred in the surrounding community despite the continued presence of the beads after 12 weeks, suggests that any released breakdown products are utilized by the fungal community attached to the PU surface and do not diffuse into the wider environment.

232 Colony count of PU beads amended compost did not show any significant difference at particular temperature among different percentages of PU, also diversity via culture based method was similar with the unamended compost at 45° and 50°C.

TRFLP does not reveal any change in compost amended with beads because being insoluble. The population on beads during the incubation were grouped together with population of compost with unamended compost on PCA. Since in previous chapters we have found out that the population on the surface of PU is different from native and organisms are growing on the surface only with no effect on native compost population. The short term shift in the community structure was most pronounced at

25°C but far less pronounced at 45° and 50°C which may reflect the fact that the community composition is far more diverse at lower temperatures where mesophiles predominate and far more restricted at the elevated temperatures where only thermotolerant and thermophilic species can grow. The addition of macroscopic PU waste to composts may therefore have little impact on the overall structure of the fungal compost community in the long term though may cause a temporary shift in structure.

In summary this study demonstrates that the addition of macrospcopic polyester PU to composts appears to have no long lasting effect on the fungal community profile but that extended periods will be required for total degradation while the addition of microparticulate PU, due to its greater surface to area ratio and homogenous distribution in the compost, is degraded more rapidly leading to an increase in the fungal viable count and a shift in the community profile. However it is not yet known if the community profile would return to that of unamended compost after all the PU has been utilised for growth.

233 4.6 References

Alef, K., Nanniperi, P., 1995. Methods in applied soil microbiology and chemistry. Academic Press, London,U.K. Anonymous, 2011. Plastics - the Facts 2011 An analysis of European plastics production , demand and recovery for 2010. Areikin, E., Horne, J., Scholes, P., Mines, U., Briggs, L., Brown, B., Dyson, B., Resource, W., 2012. A survey of the UK organics recycling industry in 2010. Barratt, S.R., Ennos, A.R., Greenhalgh, M., Robson, G.D., Handley, P.S., 2003. Fungi are the predominant micro-organisms responsible for degradation of soil- buried polyester polyurethane over a range of soil water holding capacities. Journal of Applied Microbiology 95, 78–85. Bentham, R.H., Morton, L.H.G., Allen, N.G., 1987. Rapid assessment of the microbial deterioration of polyurethanes. International Biodeterioration 23, 377–386. Berga, M., Székely, A.J., Langenheder, S., 2012. Effects of disturbance intensity and frequency on bacterial community composition and function. PloS one 7, 1–11. Bressan, M., Mougel, C., Dequiedt, S., Maron, P.-A., Lemanceau, P., Ranjard, L., 2008. Response of soil bacterial community structure to successive perturbations of different types and intensities. Environmental microbiology 10, 2184–2187. Cosgrove, L., McGeechan, P.L., Handley, P.S., Robson, G.D., 2010. Effect of biostimulation and bioaugmentation on degradation of polyurethane buried in soil. Applied and Environmental Microbiology 76, 810–819. Cosgrove, L., McGeechan, P.L., Robson, G.D., Handley, P.S., 2007. Fungal communities associated with degradation of polyester polyurethane in soil. Applied and Environmental Microbiology 73, 5817–5824. Crabbe, J.R., Campbell, J.R., Thompson, L., Walz, S.L., Schultz, W.W., 1994. Biodegradation of a colloidal ester-based polyurethane by soil fungi. International Biodeterioration and Biodegradation 33, 103–113. Darby, R.T., Kaplan, A.M., 1968. Fungal susceptibility of polyurethanes. Applied Microbiology 16, 900–905. Feng, J., Hwang, R., Chang, K.F., Hwang, S.F., Strelkov, S.E., Gossen, B.D., Zhou, Q.A., 2010. An inexpensive method for extraction of genomic DNA from fungal mycelia. Canadian Journal of Plant Pathology 32, 396–401. Filip, Z., 1985. Microbial degradation of polyurethanes, in: Seal, K.J. (Ed.), Biodeterioration and Biodegradation of Polymers. Biodeterioration Society, New York, US, pp. 51–55.

234 Hopewell, J., Dvorak, R., Kosior, E., 2009. Plastics recycling: challenges and opportunities. Philosophical Transactions of the Royal Society of London. Series B, Biological sciences 364, 2115–2126. Kawasaki, A., Watson, E.R., Kertesz, M. a., 2011. Indirect effects of polycyclic aromatic hydrocarbon contamination on microbial communities in legume and grass rhizospheres. Plant and Soil 358, 169–182. Kay, M.J., Morton, L.H.G., Prince, E.L., 1991. Bacterial degradation of polyester polyurethane. International Biodeterioration and Biodegradation 27, 205–222. Krasowska, K., Janik, H., Gradys, A., Rutkowska, M., 2012. Degradation of polyurethanes in compost under natural conditions. Journal of Applied Microbiology 125, 4252–4260. Matsumiya, Y., Murata, N., Tanabe, E., Kubota, K., Kubo, M., 2010. Isolation and characterization of an ether-type polyurethane-degrading micro-organism and analysis of degradation mechanism by Alternaria sp.Journal of Applied Microbiology 108, 1946–1953. Oehlmann, J., Schulte-Oehlmann, U., Kloas, W., Jagnytsch, O., Lutz, I., Kusk, K.O., Wollenberger, L., Santos, E.M., Paull, G.C., Van Look, K.J.W., Tyler, C.R., 2009. A critical analysis of the biological impacts of plasticizers on wildlife. Philosophical Transactions of the Royal Society of London. Series B, Biological sciences 364, 2047–2062. Pathirana, R.A., Seal, K.J., 1984. Studies on polyurethane deteriorating fungi. Part 1. Isolation and characterization of the test fungi employed. International Biodeterioration 20, 163–168. Rutkowska, M., Krasowska, K., Heimowska, A., Steinka, I., Janik, H., 2002. Degradation of polyurethanes in sea water. Polymer Degradation and Stability 76, 233–239. Sasek, V., Vitásek, J., Chromcová, D., Prokopová, I., Brozek, J., Náhlík, J., 2006. Biodegradation of synthetic polymers by composting and fungal treatment. Folia Microbiologica 51, 425–430. Smith, C.J., Danilowicz, B.S., Clear, A.K., Costello, F.J., Wilson, B., Meijer, W.G., 2005. T-Align, a web-based tool for comparison of multiple terminal restriction fragment length polymorphism profiles. FEMS Microbiology Ecology 54, 375–380. Tiquia, S.M., 2005. Microbial community dynamics in manure composts based on 16S and 18S rDNA T-RFLP profiles. Environmental Technology 26, 1101–1113. Urgun-Demirtas, M., Singh, D., Pagilla, K., 2007. Laboratory investigation of biodegradability of a polyurethane foam under anaerobic conditions. Polymer Degradation and Stability 92, 1599–1610.

235 Webb, J.S., Nixon, M., Eastwood, I.M., Greenhalgh, M., Robson, G.D., Handley, P.S., 2000. Fungal colonization and biodeterioration of plasticized polyvinyl chloride. Applied and Environmental Microbiology 66, 3194–3200. White, T.J., Bruns, T.D., Lee, S., Taylor, J., 1990. Analysis of phylogenetic relationshipd by amplificaion and direct sequencing of ribosomal RNA genes, in: Innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocol: a Guide to Methods and Applications. Academic Press, New York, US, pp. 315–322.

236 4.7 Appendix A4

Table A4.1: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with Impranil DLU (liquid dispersion of polyether PU).

DLU Week 0 Week 1 Week 4 Week 8 Week 12

5 5 5 5 4 25 4.3 X10 a 2.3X10 b 2.5 X10 b 2.4X10 b 2.4 X10 b

4 3 4 3 2 45 2.3X10 c 2.5X10 c 1.8 X10 c 6.1X10 c 8.2X10 c

4 2 3 4 3 50 °C 4.2 X10 c 8.8X10 c 1 X10 c 1.7X10 c 1.2 X10 c

Total fungal count was enumerated to observe the effect of Impranil DLU addition on microbial count and ecology. Trend observed was seen similar as of control (exposure to temperatures without Impranil amendment- Table 1).

237 Table A4.2: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with Impranil DLN (liquid dispersion of polyester PU).

DLN 25 °C 45 °C 50 °C

Week CEA PUA % CEA PUA % CEA PUA %

5 5 4 4 4 4 0 4.3 X10 a 1.6 X10 a 37.2 3.8 X10 a 3.4 X10 a 89.5 6.9 X10 a 5.2 X10 a 75.4

6 4 6 5 6 5 1 1.5 X10 b 4.4 X10 a 29.3 1.2 X10 c 3.1X10 d 25.8 1.2 X10 c 3.9 X10 d 32.5

5 5 6 5 6 5 4 9.7X10 c 2.8 X10 d 28.9 1.4 X10 c 2.6 X10 d 18.6 1.2 X10 c 3.1 X10 d 25.8

5 5 6 5 6 5 8 9.5 X10 c 2.6 X10 a,d 48.0 1.2 X10 c 2.1 X10 d 17.5 1.1 X10 c 4.6 X10 d 41.8

4 4 4 3 3 3 12 3.2 X10 a 1.7 X10 a 53.1 1.2 X10 a 2.2 X10 a 18.3 2.4 X10 a 1.1X10 a 47.1

Total and Impranil degrading count was enumerated to observe the effect of Impranil DLN addition on microbial count and ecology. Significant rise in cfu/gm was seen at 45 and 50 °C after Impranil DLN addition. The total count and Impranil degrading count at higher temperature showed similar count as at 25 °C.

238

Table A4.3: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks as control.

Unamended compost

Week 25 °C 45 °C 50 °C

CEA PUA % CEA PUA % CEA PUA %

5 5 3 3 3 3 0 3.9 X10 a 1.6 X10 c 41 3.3 X10 d 1.5 X10 45.5 5 X10 d 2.0 X10 d 40

5 5 4 3 3 2 1 2.6 X10 b 1.6 X10 c 61.5 1.3 X10 e 2.2X10 d 16.9 1.2 X10 d,e 2.5 X10 d 20.8

5 5 4 4 3 2 4 2.1X10 b,c 1.3 X10 c 61.9 2.4 X10 e 1.5 X10 e 62.5 9.0 X10 d,e 4.2 X10 d 4.7

5 5 4 4 3 2 8 2.5 X10 b,c 1.8 X10 c 72 1.8 X10 e 1.2 X10 e 66.7 9.1 X10 d,e 4.2 X10 d 4.6

5 5 4 4 3 2 12 2.1X10 b,c 1.7 X10 c 70.8 2.2 X10 e 2.0 X10 e 90.9 5.0X10 d,e 4.5X10 d 9

Total and Impranil degrading count was enumerated to observe the effect of temperature on microbial count and ecology. Effect was observed just after exposure (week 1), which remain unchanged till week 12.

239 Table A4.4: Enumeration of cfu/gm of compost incubated at 25, 45 and 50 °C for 12 weeks after amendment with 10, 30 and 50 °C% of polyester PU beads.

10% 30% 50% Temp. weeks CEA PUA CEA PUA CEA PUA

6 6 6 6 6 6 25 4 2 (0.3)X10 a 1 (0.2)X10 a 2 (0.1)X10 a 1 (0.4)X10 a 2 (0.1)X10 a 1 (0.2)X10 a

6 5 5 5 5 5 8 1 (0)X10 a 5(0)X10 a 2 (0.2)X10 a 3 (0.4)X10 a 6 (0.4)X10 a 1 (0)X10 a

6 5 6 6 6 6 12 2 (0.2)X10 a 5 (0.6)X10 a 5 (0.6)X10 a 2 (0.2)X10 a 3 (0.3)X10 a 2 (0.2)X10 a

3 3 4 3 4 4 45 4 5 (1)X10 b 9 (5)X10 b 1 (0.2)X10 b 3 (0.2)X10 b 9 (2)X10 b 7 (2)X10 b

3 3 4 4 4 4 8 7 (0.7)X10 b 3 (0.9)X10 b 3 (0.70X10 b 2 (0.5)X10 b 2 (0.2)X10 b 2 (4)X10 b

4 4 4 4 3 3 12 1 (0.1)X10 b 1 (0.2)X10 b 8 (0.9)X10 b 5 (0.6)X10 b 3 (0.4)X10 b 1 (0.2)X10 b

3 3 3 3 3 3 50 °C 4 6 (0.7)X10 b 2 (0.7)X10 b 4 (2)X10 b 5 (0.2)X10 b 7 (0.4)X10 b 1 (0.4)X10 b

4 3 3 3 3 3 8 1 (0.1)X10 b 8 (0)X10 b 7 (0.4)X10 b 2 (0.2)X10 b 6 (0.2)X10 b 3 (0.2)X10 b

3 3 3 3 3 3 12 6 (0.6)X10 b 4 (0.5)X10 b 6 (0.7)X10 b 2 (0.2)X10 b 3 (0.3)X10 b 1 (0.1)X10 b

Total and Impranil degrading count was enumerated to observe the effect of addition of polester PU on microbial count and ecology. No major effect was seen on total and Impranil degrading fungal count after amendment. Count observed from 45 and 50 °C were significantly different from count at 25 °C.

240 45

4.06

3.24 2.43 25 1.62

0.81 Axis 2 (28%) 2 Axis

0.00

-0.81

-1.62

-2.43 -2.43 -1.62 -0.81 0.00 0.81 1.62 2.43 3.24 4.06 Axis 1 (62%) A 5.15

4.12

3.09

2.06

1.03 Axis 2 (16%) 2 Axis

0.00

-1.03

-2.06

-3.09 -3.09 -2.06 -1.03 0.00 1.03 2.06 3.09 4.12 5.15

Axis 1(70%)

B 45

4.06

3.24

2.43

1.62

0.81 Axis 2 (28%) 2 Axis

0.00

-0.81

-1.62

-2.43 -2.43 -1.62 -0.81 0.00 0.81 1.62 2.43 3.24 4.06

Axis 1 (62%)

241

50 C 4.0

3.2

2.4

1.6

0.8

0.0 Axis 2 (19%) 2 Axis

-0.8

-1.6

-2.4

-3.2 -3.2 -2.4 -1.6 -0.8 0.0 0.8 1.6 2.4 3.2 4.0 Axis 1 (52%)

Figure A4.1: PCA score for samples amended with impranil DLN ( for week 1 (), week 4 (), week 8 () and week 12 () and DLU (▼) for week 1 (▼), week 4 (▼), week 8 (▼) and week 12 (▼). With out control samples were different in between DLN and DLU with major transition observed in impranil DLN amended compost at all temperatures.

242 Chapter 5: General discussion

5.1 General discussion Polyurethanes are synthetic high volume plastics that have a very broad range of applications due to their physicochemical characteristics ranging from footwear, medical devices, flooring, furniture, adhesives, sealants and automotive interiors (Howard, 2002). In Europe alone, production amounted to 3.29 Million tonnes in 2012 and accounted for 7% of all plastics manufactured (Plastics – the

Facts 2012). As a result, a large proportion of PU’s enter the waste streams every year with a large proportion directed to landfill sites, which are becoming an increasingly expensive and limited resource. Polyester PUs have been shown to be highly susceptible to microbial degradation due to the presence of ester and urethane linkages in the polymer which are vulnerable to enzymatic hydrolysis (Darby &

Kaplan 1968; Morton & Surman 1994;Krasowska et al., 2012) and more recently it has been reported that fungi are the principal organism responsible for PU biodeterioration (Barratt et al., 2003; Cosgrove et al., 2007). The aims of this thesis were to investigate if PU was susceptible to biodegradation in compost environments; to determine the rate and extent of PU degradation; to characterise the fungi colonising and deteriorating PU under laboratory conditions and in situ at a commercial composting facility and to evaluate the potential for the composting process to manage PU wastes.

Chapter two investigated the succession of microbial colonization and degradation of PU following burial in compost under laboratory conditions. Polyester

PU was found to be highly degradable under compost at 25°, 45° and 50°C with a

243 large decrease in tensile strength that occurred at a similar rate as that observed in soil at 25°C. Thus polyester PU is susceptible to fungal colonisation and degradation at both mesophilic and thermophilic stages of the composting process. Polyether PU however did not show any reduction in tensile strength over the 12 weeks incubation period. While known to be more recalcitrant to biodegradation due to the presence of ether rather than ester linkages (Darby & Kaplan, 1968), nonetheless there are some reports of significant fungal deterioration of polyether PU and longer incubation periods may be necessary to determine whether the polyether PU is truly recalcitrant

(Matsumiya et al., 2010). In addition, while the majority of fungi present in compost and soil were shown to be capable of polyester PU degradation, this may be far more restricted for polyether PU. In this study, no fungi were recovered from compost on polyether PU plates. As the diversity of fungi is known to vary widely in composts, a much broader range of composting materials should be used to assess polyether PU degradation (Ryckeboer, Mergaert, Vaes, et al., 2003).

454 pyrosequencing using the ITS region of rDNA was also employed to study the community structure of fungi on the surface of PU during deterioration in compost. The community structure on the surface of PU coupons was very different from the community in the surrounding compost. Dominant isolates recovered from the surface of PU at 25°C included Fusarium solani, Bionectria ochroleuca and an

Alternaria sp. whereas at 45° and 50 °C Candida ethanolica and an unidentified fungal clone were dominant. Among these, Fusarium solani and Alternaria species have been reported previously as PU degraders (Pommer and Lorenz, 1985;

Stranger-Johannessen, 1985; Bentham et al., 1987; Crabbe et al., 1994; Cosgrove et al., 2007; Ibrahim et al., 2009; Matsumiya et al., 2010; Russell et al., 2011). A

244 commercial lipase from Candida rugosa, which was also detected on the PU surface at 50°C has been reported to cause PU degradation (Gautam, Bassi, Yanful, et al.,

2007). While many of the organisms detected on the surface of PU at 25°C were also isolated by cultivation, at 45° and 50°C this was not the case as only Aspergillus fumigatus and Thermomyces lanuginosus were isolated. 454 pyrosequencing is a very powerful tool for investigating the structure and diversity of fungal communities and although widely employed in the study of bacterial systems has to date only been applied to fungal communities in a limited number of studies (Danielsen et al., 2012;

Buée et al., 2013). Targeted attempts to isolate Candida ethanolica and Penicillium paneum that were dominant at thermophilic temperatures would enable future studies to investigate the enzymes involved in PU degradation and infrared spectroscopy and

HPLC of degrading materials in the presence of these isolates would enable the pathways of PU degradation to be elucidated.

The results from chapter 2 suggested that even with low diversity at 45° and

50° C, the rate of degradation of PU was still substantial. However, these studies were conducted in microcosms under laboratory conditions and it has been reported that the results from the small scale studies cannot be directly applied to full-scale composting processes (Hultman et al., 2010). Therefore, in chapter 3, the degradation of PU was investigated in a commercial composting facility (TEG group, UK). The commercial process in use was in silo composting followed by maturation in large

(10 m tall) compost heaps for 4 weeks. PU coupons were buried at the centre and 0.4 m from the surface of the compost pile and after 28 days showed > 70% decrease in the tensile strength. A number of thermophilic/tolerant isolates were recovered from the surface of buried PU buried at 0.4 m from the surface of the compost pile where

245 the temperature had declined to ca. 45°C over the 28 day period whereas no fungi were recovered from the centre of the pile where the temperature remained >65°C preventing active fungal growth. Both TRFLP and 454 pyrosequencing again indicated that community colonising the PU surface was different from the surrounding compost. The two most dominant fungi detected from the surface of PU coupons was an unidentified fungal clone and Arthrographis kalrae comprising ca.

90% of the population.

While significant loss in tensile strength was observed in PU buried at the centre and surface of the compost pile, significant differences were seen on the effects on Tg and Tm. This may reflect bacterial activity in the centre of the pile where temperatures prevent fungal growth and differing modes of enzymatic action on PU between bacteria and fungi (Wales and Sagar 1985). Further investigation into the structural changes in PU under these two conditions employing infrared spectroscopy, molecular weight determination and detection of released breakdown products by HPLC would enable the degradation pathways and enzymes sites of action to be better characterised.

As only the post silo windrows phase of the process was investigated, it would be necessary to also determine the effect of the in silo composting process on the structure and integrity of PU and how this subsequently also impacts on degradation in the windrows stage. Moreover, longer windrows incubation periods and the effect of regularly physically turning the compost piles should also be investigated to determine the time required for complete degradation.

If PU’s were to be diverted into composting systems, it is important to

246 understand what influence this may have on the surrounding community structure as this may have an impact on compost degradation and quality or lead to the prevalence of the opportunistic pathogen A. fumigatus which may have health implications by increasing spore concentration around composting sites. In Chapter

4, the influence of adding microparticulate PU and solid PU on the compost fungal community was investigated at 25°, 45° and 50°C for a period of 12 weeks.

The microparticulate dispersion impranil had a profound effect on total viable counts and fungal diversity. In the presence of impranil, additional fungal species were isolated including Humicola grisea and Malbranchea cinnamomea, that were also identified in Chapter 3 as a potential PU degrader. All of these isolates have been extensively isolated from compost previously (Finstein and Morris, 1975;

Ryckeboer, Mergaert, Vaes, et al., 2003; Hultman et al., 2010) but have not been reported as PU degrader except Aspergillus fumigatus (Pathirana and Seal, 1984).

Interestingly, while polyether coupons showed little degradation in chapter 2, microparticulate impranil DLU (polyether PLU) also increased the fungal viable count suggesting that perhaps degradation is much lower and only affected the numbers of viable fungi due to the particle’s small size. Far longer incubation periods and studies on the physical structure of polyether PU will need to be undertaken to fully characterise how recalcitrant the material is.

While microparticulate polyester PU increased both population size and diversity, solid PU only caused a slight and temporal change in the surrrounding community, possible due to intial perturbation of the system caused by the physical inclusion of macroscopic PU. Thus, prelimanary studies suggest that solid PU has

247 little effect on the surrounding compost but further work will need to be conducted to verify this.

Overall, this study provides prelimary evidence that composts have the potential to incorporate PU’s into the process and away from other more damaging waste disposal systems. However, experiments looking at comparing different composting systems and on changing parameters within existing processes will need to be undertaken to fully evaluate if PU’s can be efficiently and economically treated through composting.

248 5. 2 References

Barratt, S.R., Ennos, A.R., Greenhalgh, M., Robson, G.D., Handley, P.S., 2003. Fungi are the predominant micro-organisms responsible for degradation of soil- buried polyester polyurethane over a range of soil water holding capacities. Journal of Applied Microbiology 95, 78–85. Bentham, R.H., Morton, L.H.G., Allen, N.G., 1987. Rapid assessment of the microbial deterioration of polyurethanes. International Biodeterioration 23, 377–386. Buée, A.M., Reich, M., Murat, C., Morin, E., Nilsson, R.H., Uroz, S., Martin, F., Mur, C., 2013. 454 Pyrosequencing analyses of forest soils reveal an unexpectedly fungal diversity. New Phytologist 184, 449–456. Cosgrove, L., McGeechan, P.L., Robson, G.D., Handley, P.S., 2007. Fungal communities associated with degradation of polyester polyurethane in soil. Applied and Environmental Microbiology 73, 5817–5824. Crabbe, J.R., Campbell, J.R., Thompson, L., Walz, S.L., Schultz, W.W., 1994. Biodegradation of a colloidal ester-based polyurethane by soil fungi. International Biodeterioration and Biodegradation 33, 103–113. Danielsen, L., Thürmer, A., Meinicke, P., Buée, M., Morin, E., Martin, F., Pilate, G., Daniel, R., Polle, A., Reich, M., 2012. Fungal soil communities in a young transgenic poplar plantation form a rich reservoir for fungal root communities. Ecology and Evolution 2, 1935–1948. Darby, R.T., Kaplan, A.M., 1968. Fungal susceptibility of polyurethanes. Applied Microbiology 16, 900–905. Finstein, M.S., Morris, M.L., 1975. Microbiology of solid waste composting. Advance and Applied Microbiology 19, 113–149. Gautam, R., Bassi, a. S., Yanful, E.K., Cullen, E., 2007. Biodegradation of automotive waste polyester polyurethane foam using Pseudomonas chlororaphis ATCC55729. International Biodeterioration and Biodegradation 60, 245–249. Howard, G.T., 2002. Biodegradation of polyurethane: a review. International Biodeterioration and Biodegradation 49, 245–252. Hultman, J., Vasara, T., Partanen, P., Kurola, J., Kontro, M.H., Paulin, L., Auvinen, P., Romantschuk, M., 2010. Determination of fungal succession during municipal solid waste composting using a cloning-based analysis. Journal of Applied Microbiology 108, 472–487. Ibrahim, I.N., Maraqa, A., Hameed, K.M., Saadoun, I.M., Maswadeh, H.M., Nakajima-kambe, T., 2009. Polyester-polyurethane biodegradation by Alternaria solani , isolated from northern Jordan. Advances in Environmental Biology 3, 162– 170.

249 Krasowska, K., Janik, H., Gradys, A., Rutkowska, M., 2012. Degradation of polyurethanes in compost under natural conditions. Journal of Applied Microbiology 125, 4252–4260. Matsumiya, Y., Murata, N., Tanabe, E., Kubota, K., Kubo, M., 2010. Isolation and characterization of an ether-type polyurethane-degrading micro-organism and analysis of degradation mechanism by Alternaria sp.Journal of Applied Microbiology 108, 1946–1953. Morton, L.H.G., Surman, S.B., 1994. Biofilms in Biodeterioration-a review. International Biodeterioration and Biodegradation 203–221. Pathirana, R.A., Seal, K.J., 1984. Studies on polyurethane deteriorating fungi. Part 1. Isolation and characterization of the test fungi employed. International Biodeterioration 20, 163–168. Plastics – the Facts 2012. An analysis of European plastics production , demand and waste data for 2011, 2012. . Pommer, E.H., Lorenz, G., 1985. The behaviour of polyester and polyether polyurethanes towards microorganisms, in: Seal, K.J. (Ed.), Biodeterioration and Biodegradation of Polymers. biodeterioration society, New york, pp. 77–86. Russell, J.R., Huang, J., Anand, P., Kucera, K., Sandoval, A.G., Dantzler, K.W., Hickman, D., Jee, J., Kimovec, F.M., Koppstein, D., Marks, D.H., Mittermiller, P. a, Núñez, S.J., Santiago, M., Townes, M. a, Vishnevetsky, M., Williams, N.E., Vargas, M.P.N., Boulanger, L.-A., Bascom-Slack, C., Strobel, S. a, 2011. Biodegradation of polyester polyurethane by endophytic fungi. Applied and Environmental Microbiology 77, 6076–6084. Ryckeboer, J., Mergaert, J., Vaes, K., Klammer, S., 2003. A survey of bacteria and fungi occurring during composting and self-heating processes. Annals of Microbiology 53, 349–410. Stranger-Johannessen, M., 1985. Microbial degradation of polyurethane products in service, in: Seal, K.J. (Ed.), Biodeterioration and Biodegradation of Polymers. Biodeterioration Society, New york, pp. 264–267. Wales, D.S., Sagar, B.F., 1985. The mechanism of polyurethane biodeterioration, in: Seal, K.J. (Ed.), Biodeterioration and Biodegradation of Polymers. New york, pp. 56–69.

250