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Biodegradation of low-density polyethylene by fungi
Submitted in total fulfilment of the requirements for the degree of
Doctor of Philosophy
by
Sasi Kiran Kumar Kanchi
Faculty of Science, Engineering and Technology
Swinburne University of Technology
October 2015
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Abstract
The accumulation of recalcitrant plastics in the environment, particularly
polyethylene, is a major threat to the ecosystem. Among the various types of
polyethylene, low-density polyethylene (LDPE) is the most commonly used.
Unfortunately, the rate of production and consumption of polyethylene is outweighs the
rate of its disposal. Although several approaches such as incineration, landfill treatment
and recycling have been proposed, these approaches have been either too costly or only
partially effective. An alternative solution, involving the biodegradation of LPDE, has
been proposed.
Current understanding of the biodegradation of LDPE suggests that it requires a
considerable amount of time for the process to be completed. Previous studies on LDPE
describe the importance of oxidising the LDPE to make it suitable for microbes to
degrade. In this study, an attempt was made to understand the biodegradation process
after oxidation, in order to develop strategies for improving the efficiency and reducing
the time required for this process to be completed.
LDPE samples were treated with fungal isolates that were collected from river
and landfill sources. Their intrinsic properties were assessed before and after fungal
treatment, and their surface characteristics were determined by atomic force microscopy
(AFM) and scanning electron microscopy (SEM). Modifications of functional groups
were evidenced by Fourier transform infrared spectroscopy (FT-IR) and Raman
spectroscopy. Changes in crystallinity were monitored by X-ray diffraction (XRD). A
relatively inexpensive staining technique was proposed to quantify the differences
between untreated LDPE and fungal-treated LDPE.
Fungal-treated LDPE showed characteristic features that differed significantly
from untreated LDPE. Changes in the crystallinity, buoyancy and colour of LDPE were
observed following fungal treatment. The fungal strain isolated was identified as
Fusarium oxysporum by18S rRNA gene sequencing. The hydrophobicity of this isolate
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and other fungal varieties was measured to estimate their ability to attach to LDPE.
These fungi were classified according to their microscopic and macroscopic features.
Various factors affecting biodegradation were monitored. Salts, alcohols and
sugars were screened for their effects on the biodegradation process. The effects of pH,
temperature, oxidation, co-metabolites and biofilm formation were also determined, as
well as the weight losses of LDPE samples during the course of biodegradation.
This work reports for the first time the ability of Fusarium oxysporum to degrade
LDPE. Alcohols and sugars were shown to accelerate the biodegradation process, along
with salts, such as MnCl2. The ambiguity regarding the necessity for biofilm formation
during biodegradation was clarified. It was demonstrated that formation of biofilm was
not necessary for biodegradation to occur. In fact, biofilm formation was shown to slow
this process. The effects of co-metabolites, such as monosachcharides, disachccharides
and polysachcharides, were described in detail for the first time. Further, the reasons for
the enhanced biodegradation of LDPE in the presence of co-metabolites were
elaborated.
Biodegradation of LDPE with fungal extracts was also performed in order to
determine the possible factors affecting the activity of proteins that may participate in
this mechanism. The proteins present in fungal extracts were identified by Sodium
dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE). In this study,
laccase from Fusarium oxysporum was identified of playing an important role in LDPE
biodegradation. In addition, attempts to biodegrade LDPE with enzymes in vitro were
described, and the effects of ethanol and sucrose on the oxidation capacity of laccase
were also determined.
In the final section of this thesis, factors governing the biodegradation rate are
discussed. Physical factors, such as surface roughness and crystallinity, are elaborated.
Structural factors, such as tertiary carbon atoms and polymer chain length, are also
discussed in detail. Various suggestions are made to increase the biodegradability of
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LDPE in its native and oxidised states to encourage biodegradation in natural
environments, landfills and laboratory fermenters.
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Acknowledgements
My first thanks goes to Professor Enzo Palombo, my primary supervisor. He is a
kind and highly professional supervisor. He is the inspiration and motivation behind this
project. He always had time to discuss about the project.
I would also like to thank Dr François Malherbe, my co-supervisor. His vast
experience and knowledge about polymer science certainly helped in improvising my
thesis. His support and encouragement were vital to develop this thesis.
I would like to thank all laboratory technicians namely Chris, Soula, Savi and
Ngan. I wish to thank Dr. Peter Mahon for his help regarding Raman spectroscopic
analysis. I also wish to convey my thanks to Dr. Hayden Webb for his help regarding
atomic force microscopy.
My thanks also go to all the colleagues, friends of Swinburne University of
technology. I would like to thank my family members for their support and
encouragement.
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Declaration
I, Sasi Kiran Kumar Kanchi, declare that this thesis is my original work. It does not
contain any material that was previously published, except where due reference is made.
I also declare that this work was edited by professional editors for its grammatical
mistakes.
Signature
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Contents
Abstract ............................................................................................................................ ii
Acknowledgements .......................................................................................................... v
Declaration ...................................................................................................................... vi
Contents ......................................................................................................................... vii
List of Tables ................................................................................................................. xv
List of Figures ............................................................................................................... xvi
List of Abbreviations..................................................................................................... xx
List of Symbols and Units ............................................................................................ xxi
Chapter 1 Introduction .................................................................................................. 1
1.1. Introduction ............................................................................................................ 2
1.2. Aim of this study .................................................................................................... 3
Chapter 2 Literature review .......................................................................................... 6
2.1. Chapter overview ................................................................................................... 7
2.2. Introduction ............................................................................................................ 7
2.2.1. PE .................................................................................................................... 7
2.2.2. HDPE .............................................................................................................. 7
2.2.3. LDPE ............................................................................................................... 8
2.2.4. LLDPE ............................................................................................................ 9
2.3. Production of LDPE ............................................................................................. 12
2.4. Oxidation of LDPE .............................................................................................. 13
2.4.1. Photo-oxidation ............................................................................................. 13
2.5. Traditional methods to dispose of LDPE ............................................................. 15
2.5.1. Landfill .......................................................................................................... 15
2.5.2. Incineration ................................................................................................... 15
2.5.3. Recycling ...................................................................................................... 15
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2.6. Attempts to change the nature of LDPE .............................................................. 16
2.6.1. Bioplastics ..................................................................................................... 17
2.6.2. Starch-based LDPE ....................................................................................... 17
2.6.3. Cellulose-based LDPE .................................................................................. 17
2.6.4. Lactic acid-based LDPE ................................................................................ 18
2.6.5. Lignin-based LDPE ....................................................................................... 18
2.7. Oxo-PE ................................................................................................................. 19
2.8. Degradable PE ...................................................................................................... 20
2.9. Various standards of biodegradation .................................................................... 21
2.10. Terminology of biodegradation .......................................................................... 22
2.11. Mode of biodegradation ..................................................................................... 23
2.12. Types of LDPE biodegradation .......................................................................... 25
2.13. Previous attempts to degrade LDPE .................................................................. 25
2.13.1. The high molecular weight of LDPE .......................................................... 25
2.13.2. The hydrophobic nature of LDPE ............................................................... 26
2.13.3. The 3-D structure of LDPE ......................................................................... 26
2.14. Surface characterisation techniques ................................................................... 27
2.14.1. FT-IR ........................................................................................................... 27
2.14.2. Raman spectroscopy ................................................................................... 29
2.15. Surface visualisation .......................................................................................... 32
2.15.1. AFM ............................................................................................................ 32
2.15.2. SEM ............................................................................................................ 33
2.16. Crystallinity of LDPE ........................................................................................ 34
2.16.1. XRD ............................................................................................................ 34
2.17. Classification of fungi ........................................................................................ 35
2.17.1. Macroscopic classification .......................................................................... 36
2.17.2. Microscopic classification ........................................................................... 36
2.17.3. DNA sequencing-based classification ........................................................ 36
2.18. Microbial adhesion to hydrocarbons test ........................................................... 37
2.19. Dissolved carbon dioxide content ...................................................................... 38
ix
2.20. Methylene blue test ............................................................................................ 38
2.21. Fusarium oxysporum .......................................................................................... 39
2.22. Antioxidants ....................................................................................................... 41
2.23. Biofilm ............................................................................................................... 41
2.24. Co-metabolism ................................................................................................... 43
2.24.1. Pre-exposure to an analogue compound ..................................................... 43
2.24.2. Enzyme induction by structurally unrelated compounds ............................ 44
2.24.3. The role of readily degradable compounds ................................................. 44
2.25. Laccase ............................................................................................................... 45
Chapter 3 Materials and methods ............................................................................... 47
3.1. Overview .............................................................................................................. 48
3.2. Introduction .......................................................................................................... 48
3.3. Chemicals and reagents ........................................................................................ 49
3.3.1. LDPE pellets ................................................................................................. 49
3.3.2. HDPE ............................................................................................................ 50
3.4. Preparation of LDPE for biodegradation ............................................................. 50
3.4.1. Photo-oxidation ............................................................................................. 50
3.4.2. Chemical oxidation ....................................................................................... 50
3.4.3. Abiotic oxidation ........................................................................................... 50
3.5. Disinfection of LDPE ........................................................................................... 50
3.6. Isolation of fungi .................................................................................................. 50
3.6.1. Sterilisation ................................................................................................... 51
3.6.2. pH determination and adjustment ................................................................. 51
3.6.3. Measuring weight loss .................................................................................. 51
3.6.4. Isolation of fungal samples from landfill ...................................................... 51
3.6.5. Isolation of fungal samples from river water ................................................ 51
3.6.6. Isolation of fungal samples from leachate .................................................... 52
3.7. Storage of fungi .................................................................................................... 53
3.7.1. Storage on PDA slants .................................................................................. 53
3.7.2. Storage under mineral oil .............................................................................. 53
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3.7.3. Storage under sterilised water ....................................................................... 53
3.7.4. Selection of fungi .......................................................................................... 53
3.7.5. MATH test .................................................................................................... 55
3.7.6. Estimation of attached protein ...................................................................... 56
3.8. Fungal culturing techniques ................................................................................. 57
3.8.1. Shaker flask cultures ..................................................................................... 57
3.8.2. PDA culture................................................................................................... 57
3.8.3. PDA broth ..................................................................................................... 57
3.8.4. Subculturing .................................................................................................. 57
3.8.5. Isolation of Fusarium strains ......................................................................... 58
3.9. Fungus classification ............................................................................................ 58
3.9.1. Slide cultures ................................................................................................. 58
3.9.2. Extraction of fungal DNA ............................................................................. 59
3.9.3. PCR ............................................................................................................... 59
3.10. Assessing fungal effects on LDPE ..................................................................... 61
3.10.1. FT-IR ........................................................................................................... 61
3.10.2. Raman spectroscopy ................................................................................... 61
3.10.3. SEM ............................................................................................................ 61
3.10.4. AFM ............................................................................................................ 62
3.10.5. XRD ............................................................................................................ 62
3.10.6. Methylene blue test ..................................................................................... 63
3.11. Spore counting ................................................................................................... 64
3.12. Factors affecting biodegradation ........................................................................ 64
3.12.1. Effect of fungal micronutrients ................................................................... 65
3.12.2. Effect of co-metabolites .............................................................................. 65
3.12.3. Effect of oxidation on the biodegradation of LDPE ................................... 65
3.12.4. Rate and extent of biodegradation .............................................................. 65
3.13. Biodegradation with cell-free extracts ............................................................... 66
3.13.1. Estimation of the protein quantity in fungal extracts .................................. 66
3.13.2. Estimation of the carbohydrate quantity in fungal extracts ........................ 66
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3.14. Assessment of oxidation of LDPE by FT-IR ..................................................... 67
3.15. Comparative biodegradation of HDPE and LDPE ............................................ 67
3.16. Biodegradation of LDPE with additives ............................................................ 67
3.17. Transmembrane inserts ...................................................................................... 68
3.17.1. Transmembrane experiments ...................................................................... 68
3.17.2. Gel filtration chromatography ..................................................................... 69
3.17.3. SDS-PAGE.................................................................................................. 70
3.18. Effect of metal salts on LDPE biodegradation by cell-free extracts .................. 72
3.19. Detection of extracellular enzymes .................................................................... 72
3.19.1. Enzymatic screening using the plate assay technique ................................. 72
3.20. Biodegradation of LDPE with laccases from Fusarium oxysporum .................. 74
3.20.1. Gel filtration separation of laccase .............................................................. 74
3.20.2. Spectrophotometric assay of laccase activity.............................................. 74
3.20.3. ABTS standard solution .............................................................................. 74
3.20.4. Biodegradation with laccase ....................................................................... 75
3.20.5. Effect of laccase on LDPE oxidation .......................................................... 75
3.21. Effect of co-metabolite additives on laccase oxidation capability ..................... 75
3.21.1. Effect of sucrose .......................................................................................... 75
3.21.2. Effect of ethanol .......................................................................................... 75
3.21.3. Effect of manganese, copper, ferrous and zinc chloride on laccase
oxidation ........................................................................................................ 76
Chapter 4 Examining fungal effect on LDPE ............................................................. 77
4.1. Overview .............................................................................................................. 78
4.2. Selection of fungi ................................................................................................. 78
4.2.1. Dissolved carbon dioxide content ................................................................. 79
4.2.2. Growth rate of fungi ...................................................................................... 80
4.2.3. Microbial adhesion test results ...................................................................... 80
4.2.4. Protein concentration of biofilm ................................................................... 82
4.2.5. Weight loss of LDPE .................................................................................... 83
4.3. Classification of fungi .......................................................................................... 83
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4.3.1. Results of slide culture .................................................................................. 84
4.3.2. DNA sequencing ........................................................................................... 84
4.4. Characterisation of fungal-treated LDPE ............................................................. 85
4.4.1. FT-IR analysis ............................................................................................... 87
4.4.2. Raman spectroscopy ..................................................................................... 89
4.5. Surface visualisation analysis .............................................................................. 91
4.5.1. Scanning electron microscopy ...................................................................... 91
4.5.2. AFM .............................................................................................................. 93
4.6. Crystallinity measurements .................................................................................. 96
4.6.1. X- ray diffraction .......................................................................................... 96
4.6.2. Methylene blue test ....................................................................................... 98
4.7. The visual properties of the LDPE samples ....................................................... 100
4.8. Biodegradation of LDPE with Irganox® ........................................................... 100
4.8.1. Weight loss .................................................................................................. 100
4.8.2. Amount of attached protein ......................................................................... 100
4.8.3. SEM results ................................................................................................. 101
4.8.4. Discussion and conclusion for biodegradation of LDPE with Irganox® ... 103
4.9. Conclusion .......................................................................................................... 103
4.9.1. Chemical aspects of degradation ................................................................. 103
4.9.2. Physical aspects of biodegradation ............................................................. 104
Chapter 5 Factors affecting biodegradation ............................................................. 107
5.1. Overview ............................................................................................................ 108
5.2. Introduction ........................................................................................................ 108
5.3. Optimisation of biodegradation .......................................................................... 109
5.3.1. Effect of micro nutrients (manganese, copper, iron and zinc ions) ............ 109
5.3.2. Effect of temperature .................................................................................. 112
5.3.3. Effect of pH ................................................................................................. 113
5.3.4. Effect of nitrates and phosphates ................................................................ 114
5.3.5. Oxidised LDPE treatment with other Fusarium isolates ............................. 115
5.4. Rate of weight loss ............................................................................................. 116
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5.5. Biodegradation of HDPE and LDPE .................................................................. 118
5.6. Effect of oxidation .............................................................................................. 120
5.7. Comparative degradation of LDPE by different oxidation methods ................. 120
5.8. Importance of biofilm formation in the biodegradation of oxidised LDPE ....... 122
5.8.1. Transmembrane experiments ...................................................................... 122
5.8.2. Difference in weight loss of LDPE samples ............................................... 123
5.8.3. Growth of fungi in transmembrane wells ................................................... 124
5.8.4. FT-IR analysis of LDPE pellets .................................................................. 124
5.9. Conclusion .......................................................................................................... 125
Chapter 6 Laccase and co-metabolism ...................................................................... 129
6.1. Chapter overview ............................................................................................... 130
6.2. Introduction ........................................................................................................ 130
6.3. Biochemical analysis of fungal extract .............................................................. 130
6.4. Gel filtration chromatography of Fusarium oxysporum extracts ....................... 130
6.5. SDS-PAGE analysis of Fusarium oxysporum extracts ...................................... 131
6.6. Identification of fungal enzymes using plate assays .......................................... 132
6.7. Biodegradation with laccase .............................................................................. 134
6.8. Effect of co-metabolites ..................................................................................... 134
6.8.1. Effect of monosaccharides .......................................................................... 134
6.8.2. Effect of disaccharides ................................................................................ 136
6.8.3. Effect of polysaccharides ............................................................................ 138
6.8.4. Effect of methanol, ethanol and propanol ................................................... 140
6.8.5. Summary of effect of co-metabolites .......................................................... 141
6.9. Effect of laccase ................................................................................................. 142
6.9.1. Effect of manganese, copper, iron (II) and zinc chloride on laccase
oxidation ...................................................................................................... 143
6.9.2. Effect of co-metabolism and laccase .......................................................... 145
6.9.3. Effect of sucrose and ethanol ...................................................................... 145
6.9.4. Summary ..................................................................................................... 146
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Chapter 7 Conclusion and future studies ................................................................. 148
7.1. Conclusion .......................................................................................................... 149
7.2. Suggestions for future study............................................................................... 151
References .................................................................................................................... 153
xv
List of Tables
Table 2-1: Physical and chemical properties of HDPE ..................................................... 8
Table 2-2: Physical and chemical properties of LDPE ..................................................... 8
Table 2-3: Physical and chemical properties of LLDPE ................................................... 9
Table 2-4: Assignment of IR absorption peaks for LDPE (Gulmine et al. 2002) ........... 28
Table 2-5: Wave numbers and their corresponding portions .......................................... 32
Table 2-6: Classification table of Fusarium oxysporum ................................................. 40
Table 3-1: Types of LDPE pellets and sheets used in this study .................................... 48
Table 3-2: PCR mix ........................................................................................................ 59
Table 4-1: Radial growth rates of fungi in mm ............................................................... 80
Table 4-2: Weight losses (in mg) of oxidised LDPE measured after biodegradation .... 83
Table 4-3: Intensities of emission by LDPE at 891 cm-1 ................................................ 91
Table 4-4: The αa portion of LDPE types, depending on relative intensity of
emission ......................................................................................................... 91
Table 4-5: Weight loss of LDPE pellets (mg) with Irganox® ...................................... 100
Table 5-1: Radial diameter of fungi before and after incubation in mm (± 2 mm) ...... 124
Table 7-1: Various microbes isolated from marine environment ................................. 151
xvi
List of Figures
Figure 1-1: Thesis structure ............................................................................................. 5
Figure 2-1: Schematic representation of the three main varieties of PE: A) LDPE, B)
HDPE, C) LLDPE .......................................................................................... 10
Figure 2-2: PE formation ................................................................................................ 12
Figure 2-3: Schematic representation of the auto-oxidation process (Peacock 2000) .... 14
Figure 2-4: Biodegradation by bulk degradation (A) and by surface erosion (B) .......... 24
Figure 2-5: Schematic diagram of an FT-IR spectrometer ............................................. 29
Figure 2-6: Schematic representation of Raman spectrometer (Begum et al. 2010) ...... 31
Figure 2-7: Schematic representation of the location of the ITS1 and ITS4 primer
sites (Kües 2007) ............................................................................................ 37
Figure 3-1: Schematic representation of methylene blue test ......................................... 63
Figure 3-2: Transmembrane set-up. A) Basic set-up showing the placement of the
semi- permeable membrane inside the transmembrane well; B) Complete
set-up showing the placement of the medium, LDPE pellets and fungal
suspension ...................................................................................................... 69
Figure 4-1: Weight loss of oxidised LDPE ..................................................................... 78
Figure 4-2: Concentration of dissolved carbon dioxide in g/L ....................................... 79
Figure 4-3: Fungal hydrophobicity ................................................................................. 81
Figure 4-4: Concentration of protein in biofilm .............................................................. 82
Figure 4-5: Micrographs of fungi isolated from landfill: (a) mycelia, (b) groups on
conidia, (c) and (d) micro-conidia (40 x magnifications). ............................. 84
Figure 4-6: Agarose gel electrophoresis of amplified DNA fragment. Lane 4 shows
a pale band of fungal DNA. Lane 1 shows the DNA ladder. ......................... 85
Figure 4-7: FT-IR spectrum of LDPE varieties. Untreated LDPE; Oxidised LDPE;
Oxidised control LDPE (control) and fungal-treated LDPE .......................... 86
Figure 4-8: Differences between functional groups in LDPE subjected to various
treatments ....................................................................................................... 88
Figure 4-9: Crystallinity of LDPE samples ..................................................................... 89
xvii
Figure 4-10: Raman spectra of LDPE ............................................................................. 90
Figure 4-11: Scanning electron micrographs. (a) untreated LDPE, (b) oxidised
LDPE, (c) oxidised control LDPE and (d) fungal-treated LDPE (at 1 µm
resolution). ..................................................................................................... 92
Figure 4-12: Scanning electron micrograph showing Fusarium mycelia and conidia .... 93
Figure 4-13: Average surface roughness of various LDPE samples............................... 94
Figure 4-14: AFM image of four types of LDPE: untreated LDPE (a), oxidised
LDPE (b), oxidised LDPE (control) (c) and Fungal-treated LDPE (d). ........ 95
Figure 4-15: XRD of untreated LDPE ............................................................................ 96
Figure 4-16: XRD of oxidised LDPE.............................................................................. 97
Figure 4-17: XRD of fungal-treated LDPE ..................................................................... 98
Figure 4-18: Optical density at 662 nm of various resultant solutions ........................... 99
Figure 4-19: Scanning electron micrographs. (a) Untreated LDPE, (b) thermally
oxidised LDPE, (c) Oxidised LDPE (control) and (d) fungal-treated
LDPE (at 1µm resolution) ............................................................................ 102
Figure 4-20: Effect of oxidation on LDPE .................................................................... 104
Figure 4-21: A schematic representation of changes in crystallinity of LDPE during
biodegradation process ................................................................................. 106
Figure 5-1: Effect of micro nutrients on LDPE biodegradation with mycelia (A) and
cell-free extract (B) of Fusarium oxysporum; 1) MnCl2, 2) CuCl2, 3)
FeCl2, 4) ZnCl2. ............................................................................................ 110
Figure 5-2: Thermo-oxidation of LDPE by metal ions (Wright 2001) ......................... 112
Figure 5-3: Effect of temperature on weight loss .......................................................... 112
Figure 5-4: Effect of pH on biodegradation Fusarium oxysporum and its cell-free
extract ........................................................................................................... 114
Figure 5-5: Effect of 1) KNO3, 2) NaNO3, 3) KH2PO4 and 4) NaH2PO4 ................. 115
Figure 5-6: Comparative biodegradation by Fusarium oxysporum (1) and other
Fusarium strains (2, 3, and 4) ....................................................................... 116
Figure 5-7: Weight loss of LDPE by Fusarium oxysporum and its cell-free extract
over seven weeks .......................................................................................... 117
xviii
Figure 5-8: Average weight loss (%) following oxidation and fungal treatment for
LDPE and HDPE.......................................................................................... 119
Figure 5-9: Effect of oxidation period on biodegradation in terms of weight loss
(%). ............................................................................................................... 120
Figure 5-10: Effect of oxidation method on biodegradation of LDPE. ........................ 121
Figure 5-11: Transmembrane LDPE biodegradation .................................................... 123
Figure 5-12: FT-IR spectrum of LDPE sample from transmembrane well and
oxidised LDPE (control) .............................................................................. 125
Figure 5-13: Imaginary LDPE biodegradation set up by fermentation ......................... 126
Figure 5-14: Schematic representation of biodegradation by surface erosion of
LDPE ............................................................................................................ 128
Figure 6-1: GFC of Fusarium oxysporum extracts ....................................................... 131
Figure 6-2: SDS-PAGE of the concentrated Fusarium oxysporum extract (Lane 1).
M=molecular weight marker ........................................................................ 132
Figure 6-3: Identification of laccase secretion by Fusarium oxysporum ...................... 133
Figure 6-4: Effect of monosaccharides on LDPE biodegradation with mycelia (A)
and with cell-free extract (B) of Fusarium oxysporum. ............................... 135
Figure 6-5: Effect of disaccharides on LDPE biodegradation with mycelia (A) and
with cell-free extract (B) of Fusarium oxysporum. ...................................... 137
Figure 6-6: Effect of polysaccharides on LDPE biodegradation with mycelia (A)
and with cell-free extract (B) of Fusarium oxysporum. ............................... 139
Figure 6-7: Effect of alcohols on LDPE biodegradation with mycelia (A) and with
cell- free extract (B) of Fusarium oxysporum. ............................................. 141
Figure 6-8: The effect of period of incubation with laccase on oxidation, as reflected
by the carbonyl index ................................................................................... 143
Figure 6-9: Effect of manganese, copper, ferrous and zinc chloride on oxidation of
LDPE (measured by the carbonyl index) ..................................................... 144
Figure 6-10: Effect of concentration of sucrose on LDPE oxidation (measured by
the carbonyl index) ....................................................................................... 145
xix
Figure 6-11: Cumulative effect of co-metabolism on laccase-induced biodegradation
of LDPE ....................................................................................................... 147
xx
List of Abbreviations
ABTS 2,2'-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid)
AFM Atomic force microscopy
AGRF Australian Genome Research Facility
APS Ammonium persulphate
ASTM American Society for Testing and Materials
BSA Bovine Serum Albumin
EPS Extracellular polymeric substance
GFC Gel filtration chromatography
HDPE High-density Polyethylene
LDPE Low-density Polyethylene
LLDPE Linear Low-density Polyethylene
MATH Microbial Adhesion Test for Hydrocarbons
PDA Potato dextrose agar
PDB Potato dextrose broth
PE Polyethylene
PET Polyethylene tetrapthalate
PS Polystyrene
PU Polyurethane
SDS Sodium dodecyl sulphate
SEM Scanning electron microscopy
UV Ultra-violet
XRD X- ray diffraction
xxi
List of Symbols and Units
Angstrom A
Centigrade C
Centimetre cm
Millimetre mm
Micrometre m
Da Dalton
Hertz H
Kilohertz kH
Hour h
Minute min
Second sec
Kilojoule KJ
Megajoule MJ
Litre L
Millilitre mL
Microlitre L
Kilogram kg
Gram g
Milligram mg
icrogram g
Kilopascal kPa
Megapascal mPa
Mole M
Millimole mmol
Micromole mol
S Svedberg
N Newton
CHAPTER 1
INTRODUCTION
Introduction
2
1.1. Introduction
Polyethylene (PE) is a miracle material of the plastics industry with the largest
tonnage worldwide (Brydson 1999). PE usage, particularly of low-density polyethylene
(LDPE) is growing daily. LDPE surpasses other PE types in many properties, making it
extremely popular for a wide range of applications. LDPE film is strong, durable,
thermally stable, odour free, heat sealable and resists chemical and biological attack
(Fellows 2000). In addition, the production cost of LDPE is low compared with other
types of PE. These factors have encouraged manufacturers to produce LDPE in vast
quantities to make a great variety of goods such as plastic bags, detergent bottles,
containers, wrapping films, soil mulch films and pipes. It was estimated that in 2011,
global LDPE production was 19.1 million tonnes, and this is expected to grow to 22
million tonnes in 2015 (Merchant Research and Consulting Ltd 2014).
Unfortunately, the increase in LDPE production and consumption has not been
compensated with proper disposal methods, resulting in its accumulation worldwide. Its
accumulation is negatively affecting terrestrial, freshwater and marine ecosystems, as
well as the organisms that live in these habitats. Marine animals mistakenly consume
plastic bags made from LDPE as a food product and die due to their inability to digest
it. Consumed LDPE enters the food chain when these animals are eaten by humans or
other animals (Knight 2013). LDPE-made plastic bags are also often mistakenly
ingested by grazing animals, which clogs their intestines and results in death by
starvation (Hosetti 2006).
Different types of LDPE disposal methods for have been developed with a
limited degree of success. For example, burying plastic in landfills, using incineration
and recycling consumed plastics. The landfill-disposal method requires empty land and
results in the emission of large volumes of toxic gases (Ebnesajjad 2012). These
emissions make the environment surrounding the landfill uninhabitable. Further, buried
plastics require a great deal of time to biodegrade completely (Stevens 2002). Post-
consumer recycling of plastics is not financially profitable and does not address the
problem of the increasing volume of plastics (La Mantia 2002). The incineration of
commercially available LDPE (along with its additives) is a controversial procedure
Introduction
3
because it results in the emission of polyaromatic hydrocarbons, carbon dioxide,
carbon monoxide, hydrogen chloride, hydrogen cyanide and phosgene into the
environment (Peacock 2000).
The degradation of LDPE by microbes is strongly recommended for their proper
disposal. This involves the conversion of the carbon atoms in recalcitrant plastics into
biological components that are harmless to the environment. Macrobiological and
microbiological degradation methods have been considered for the disposal of plastics
(Anderson et al. 1995). Macrobiodegradation involves the degradation of plastics by
insects, birds and other animals. Microbiological degradation involves LDPE
biodegradation by microbes, including fungi and bacteria (Hasan et al. 2007; Pramila et
al. 2012). The by-products of biodegradation (e.g., humus, carbon dioxide and
water) are generally eco-friendly and are able to be readily assimilated by living
organisms (Premraj & Doble 2005). Thus, biodegradation provides a viable alternative
for LDPE disposal.
Literature is available on the bacterial and fungal biodegradation of LDPE.
However, few efforts have been made to identify and isolate the types of microbes that
are suitable for the biodegradation of LDPE. As the degradation process depends on the
microbial strain employed, isolating a ‘perfect’ microbe is essential. The factors that
affect microbial biodegradation must be understood in detail to accelerate the process.
There is limited literature available on the enzymes that are involved in LDPE
biodegradation. The biodegradation mechanism of LDPE and its rate kinetics are not
known. In addition, co-metabolism of LDPE is not completely understood, although it
has been proven a major biodegradation accelerating factor (Volke-Seplveda et al.
2002).
1.2. Aim of this study
The aim of this study is to address the problem of LDPE disposal and how best
to accelerate its biodegradation. Biodegradation of LDPE, performed by either bacteria
or fungi, is a safer disposal method than incineration or landfill burying. However,
LDPE biodegradation by bacteria has been studied more extensively than fungal
biodegradation (Hadad et al. 2005; Chatterjee et al. 2010; Pramila et al. 2012).
Introduction
4
The biodegradation of LDPE requires an ideal microbe that is compatible with
the semi-anaerobic conditions that mimic the landfill environment. It must be
genetically stable and should possess tolerance for carbon dioxide, methane and other
biodegradation products. Most importantly, it should cause the highest possible weight
loss of LDPE in the lowest time possible. In this study, fungi possessing the
aforementioned characteristics will be isolated. In addition, the results of fungal
biodegradation of LDPE will be presented. While various strains of fungi were shown to
be capable of LDPE degradation, only Fusarium oxysporum will be studied in detail
due to its efficacy in this process.
The primary objective of this study is to identify methods to accelerate the
biodegradation of LDPE in natural or landfill environments. The rate of biodegradation
will be studied to identify any possible accelerating factors. To measure the extent of
biodegradation and the oxidation of polymers, a simple and inexpensive staining
technique is developed. Additionally, an attempt will be made to identify, isolate and
study enzymes that participate in biodegradation by Fusarium oxysporum (see Figure 1-
1).
The secondary objective is to check the effect of the co-metabolism of LDPE by
fungi in the presence of sugars and alcohols. Along with this, the reasons behind the
increase in LDPE biodegradation by co-metabolism will be investigated. Additionally,
the role of laccases in LDPE biodegradation will further be studied. The requirement for
biofilm formation during the course of biodegradation will be explored, as will the
mechanism of biodegradation and its rate kinetics.
In the following chapters, the present understanding of LDPE biodegradation
and the prerequisites for this method are presented. An overview of the methods and
materials that will be used in this study is provided, and the experimental results
describing the methods by which the selected fungi degrade LDPE are detailed. Finally,
the factors affecting LDPE biodegradation by these fungi are presented. After the
experimental results are described and discussed, methods that can be used to enhance
LDPE biodegradation are outlined. This work improves the current understanding of
LDPE biodegradation processes, and provides an important contribution towards the
development of new types of LDPE materials and better disposal processes.
Introduction
5
Figure 1-1: Thesis structure
Literature review Why LDPE is not biodegradable? What are the factors influence rates of biodegradation? What is the mode of biodegradation? Is bio-film formation
necessary? Why co-metabolism encourages LDPE biodegradation?
Factors affecting biodegradation Micro nutrients
that encourage LDPE biodegradation
Effect of oxidation, presence of nitrates and phosphates
Degradation reaction kinetics
Conclusion and future studies Conclusion of factors that influence biodegradation and suggestions to
increase rate of biodegradation Conclusion of necessity of bio-film formation, effect of micro nutrients,
oxidation, reaction mode and its kinetics Laccase and its influence on oxidation and co-metabolism of LDPE Modifications needed to manufacture LDPE to encourage its
biodegradation
Examining fungal effect on LDPE LDPE surface
degradation by fungi
Functional groups degradation by fungi
Physical and chemical aspects of biodegradation
Material and methods Methods to activate LDPE for biodegradation and to isolate
fungi that can do biodegradation Surface characterisation and functional group analysis of
LDPE after fungal treatment Methods to check enzymes of biodegradation Methods to check effect of co-metabolites, laccase and salts
on LDPE
Laccase and co-
metabolism Identification
and separation of laccase
Laccase mediated biodegradation
Laccase influence on LDPE oxidation and co-metabolism
CHAPTER 2
LITERATURE REVIEW
Literature review
7
2.1. Chapter overview
In this chapter, a detailed overview of the literature and prior research
underpinning this study will be presented. A detailed explanation of LDPE as a material
and its potential threat to the global environment is elaborated. The necessity for
improving the biodegradation of LDPE is stressed, using statistical evidence. Various
standards of biodegradation are elaborated.
In the second part of the literature review, previous research is drawn upon to
analyse the reasons for the bioresistance of LDPE. The mechanism of PE oxidation is
detailed to explain the molecular interactions that might participate in its
biodegradation. The importance of biofilm during the course of biodegradation is
described. Along with this, the role of the laccases and their importance in
biodegradation is also detailed. A detailed account of co-metabolism is provided, and
the possible mechanisms that occur during biodegradation are explained.
2.2. Introduction
2.2.1. PE
PE [IUPAC name polyethene or poly(methylene)] is a long-chain synthetic
resin obtained through the polymerisation of ethylene (C2H4) monomers. In its simplest
form a PE molecule consists of chains of covalently linked carbon atoms with a pair of
hydrogen atoms attached to each carbon atom (–CH2–). These chain ends are terminated
by methyl groups (–CH3) (Peacock 2000). PE plastic is characterised by toughness, low
moisture absorption, good chemical resistance, good electrical resistance, a low
coefficient of friction and ease of processing (Rosato 2004). It is the most widely used
plastic, with an annual production of approximately 80 million tonnes (Piringer &
Baner 2008). Depending on their density, these compounds are classified as high-
density polyethylene (HDPE), LDPE and linear low-density polyethylene (LLDPE).
2.2.2. HDPE
This is a high-density version of PE (0.941–0.965 g/cc), with a molecular
weight ranging from 5,000 to 250,000 Da (Aaron et al. 2010). It has a limited number
Literature review
8
of branches in its structure, allowing the polymer chains to pack closely together,
resulting in a dense, highly crystalline material (Carraher 2003). As HDPE exhibits low
swelling characteristics it is commonly used to pack juices, soft drinks and other food
materials. HDPE is comparatively easier to recycle than LDPE (La Mantia 2002). Like
the other PEs, HDPE is resistant to biodegradation.
Table 2-1: Physical and chemical properties of HDPE
Physical properties Chemical properties Rigid, hard, impact-resistant, more crystalline,
more resistant to shrinkage, water and stress
cracking than LDPE
Resistant to chemical corrosion
and hydrophobic in nature
2.2.3. LDPE
This is the low-density version of PE (0.919–0.955 g/cc) (Hilado 1998). Though
its chemical structure is similar to that of HDPE, unlike it, LDPE possesses high
frequency of branching with more tertiary carbon atoms in its structure. This branching
prevents the close approach of polymer molecules and results in decreased
crystallinity (Peacock 2000). This material is relatively soft, flexible and yet tough.
The most popular application of LDPE is foil, from which carrier bags, packaging
material and agricultural plastic are made. It is estimated that 500 billion tons of LDPE
are produced in the form of plastic bags annually (Knight 2013). Another important use
of LDPE is in soil mulching, where it is used as a covering material to prevent the
evaporation of water from the soil and maintain the moisture level during cultivation.
Table 2-2: Physical and chemical properties of LDPE
Physical properties Chemical properties Semi-rigid, translucent, low water absorption
rate, corrosion-resistant, soft surface and low
tensile strength
Generally chemically inert but
combustible at high temperature
Literature review
9
2.2.4. LLDPE
LLDPE is a linear polymer, with significant numbers of short branches (Scheirs
2009). LLDPE possesses higher tensile strength and higher impact and puncture
resistance than LDPE. It is very flexible and elongates under stress (Robertson 2012).
It has good resistance to chemicals and to ultraviolet (UV) radiation. LLDPE possesses
a narrow heat sealing range, making its processing difficult. LLDPE is very cheap
compared to other types of plastic such as nylon, poly(ethylene terephthalate) (PET)
and polystyrene (PS). It is used in manufacturing plastic wrap, stretch wrap, pouches,
Table 2-3: Physical and chemical properties of LLDPE
Physical properties Chemical properties Extremely flexible with tear, dart and impact
resistance
Melt index lower than LDPE (115–
130oC)
Literature review
10
Figure 2-1: Schematic representation of the three main varieties of PE: A) LDPE, B)
HDPE, C) LLDPE
PE can be further sub-classified depending on the frequency and types of
branching. These sub-classifications include ultra-high molecular weight PE, medium-
density PE and ultra-low molecular weight PE (PE WAX). Ultra-high molecular weight
PE is generally used to manufacture high stress resistant components like bearings,
gears and artificial joints. PE WAX is used to manufacture emulsions and polishes.
Among the PE varieties mentioned above, LDPE is the most useful and widely
used variety, in the form of plastic bags. It has been estimated that somewhere between
500 billion to a trillion plastic shopping bags are used every year (Islam 2008). Along
with this, Global demand for LDPE is expected to grow at around 2.6 % (Sagel &
HDPE
LDPE
LLDPE
Literature review
11
Pemex 2012). Adding to this, developing countries such as India and China are
expected to consume more LDPE in the future (Platts McGraw Hill Financial 2014).
Australia produces around 13×105 tonnes of plastic per annum, mostly in the
form of plastic bags, while it consumes 7 billion plastic bags annually (Brown
2003). The United States of America (USA) uses approximately one billion plastic
bags annually, resulting in 300,000 tons of landfill waste (Environmental Protection
Agency 2012).
The next largest use of LDPE is in agriculture for soil mulching. Mulching films
are used to suppress weeds, reduce the loss of moisture from soil, decrease the use of
chemicals in weed control, reduce water consumption and to speed up crop development
(Schettini et al. 2012). It has been estimated that the global consumption of LDPE
mulching films in horticulture is around 700,000 tons per year (Espi et al. 2006).
After consumption, plastic bags are generally discarded, creating an ecological
menace. Once discarded, they either enter landfills or marine ecosystems. Lightweight
plastic grocery bags are more harmful due to their propensity to be carried away by
wind and cause aesthetic damage to their surroundings. Moreover, removing these bags
from the streets is expensive and time-consuming. Discarded plastic bags are often eaten
by birds and cattle, resulting in their death (Norton 2005). Plastic bags also clog
stormwater drains and cause floods (Wehr 2011).
Discarded plastic bags also end up in the oceans and cause severe damage to
marine ecosystems. The obvious adverse effect associated with plastic bag debris in the
oceans is aesthetic. As LDPE has a lower density than water, it floats on the ocean
surface, creating a visual menace (Majumdar 2007). Marine wildlife often consumes
plastic bags, either inadvertently in the process of feeding, or deliberately because they
mistake the plastic bags for food. For example, whales and sea turtles often mistake
plastic bags for squid or jellyfish and ingest them (Schuyler et al. 2012). This ingested
plastic may lead to starvation or malnutrition as marine debris collects in the animal’s
stomach. Marine life can become entangled in plastic debris, and thus become ensnared.
This entanglement can lead to suffocation, starvation and drowning, as well as increased
vulnerability to predators or other injury. Plastic bags are generally made with a variety
Literature review
12
of additives such as plasticisers, fillers and antioxidant pigments, some of which prove
toxic (Jana & Banerjee 1999). As plastics break down, the microscopic fragments
(microplastics) generated can be consumed by fish and thus enter the food chain
(Meenakshi 2012). In addition, marine debris can harm important components of the
economy, including marine tourism, fishing and navigation.
Moreover, the LDPE used for mulching contributes to the pollution problem.
This is in general known as ‘white pollution’ (Hardwick & Gullino 2010). Most of the
mulching film degrades within one year after usage, but the rest of it accumulates in
arable land and pollutes both the ecological environment and the landscape (Stevens
2002). In 2004, it was estimated that 143,000 tons of plastic mulch were disposed of in
the US, either in landfill or by being burned on site, releasing carcinogens in to the air
(Shogren & Hochmuth 2004). In addition, LDPE can be found in other agricultural
operations as silo bunker covers, silage bags, haylage covers, greenhouse covers, bale
wrap and row covers which all contributing to the soil pollution problem. Furthermore,
in recent years, LDPE use has extended to many industries, ranging from the
manufacturing of common household goods to medical devices (Lambert et al. 2001).
2.3. Production of LDPE
LDPE is produced by high pressure polymerisation of ethylene gas. In an
autoclave reactor, ethylene is pressurised to more than 138 MPa and heated to more
than 150 °C. To these monomers units, a small amount of initiator (oxygen or peroxide)
is added to activate the ethylene monomers. This addition initiates the polymerisation
process of ethylene, yielding PE (Yam 2010).
(CH2=CH2) Catalyst and high temperature (-CH2-CH2-)n
Figure 2-2: PE formation
LDPE possesses the strongest carbon-carbon bond (C–C) in its backbone.
Disruption of this bond requires higher energy. The dipole moment between these
carbon atoms is negligible, making the polymer inert towards chemical and biological
substances.
Literature review
13
2.4. Oxidation of LDPE
LDPE biodegradation is normally achieved by the oxidative formation of
functional groups in its polymer matrix. This is primarily achieved by photo-oxidation
with UV radiation. LDPE can be deteriorated by photo-oxidation in natural weathering
conditions (Andrady 2003).
2.4.1. Photo-oxidation
Photo-oxidation of LDPE involves the excitation of the polymer by UV
radiation (λ=100–400 nm). At first, the photon energy of UV radiation is absorbed by
impurities or chemical constituents of the polymer (chromophores). Then these
chromophores participate in oxidative reactions to form free radicals. These reactions
are collectively called Norrish-type reactions. The photo-oxidation of LDPE comprises
four stages: initiation, propagation, branching and termination.
Initiation: involves the absorption of photon energy by chromophores in the
LDPE, which leads to the formation of free radicals in the LDPE matrix.
Chain propagation: involves the reaction with oxygen to produce peroxy and
alkyl radicals. These compounds interact with hydrogen cations to form polymer
hydroperoxide.
Chain branching: involves the formation of oxy radicals and hydroxyl radicals
of the polymer.
Chain termination: involves the reaction of the free radicals generated as
described above with each other to produce non-radical compounds.
Oxidation leads to the formation of reactive groups such as ketones, alcohols,
carboxylic acids and dicarboxylic acids. It results in a decrease in the average molecular
weight of LDPE (Cheremisinoff 1989) and also leads to the formation of alkyl radicals
such as ~CH2 –CH2* and ~CH2-CH2*-CH2~ (Ravve 2012). Due to the presence of free
radicals in the polymer matrix, oxidation of LDPE also occurs during the post-
irradiation period. The rate of photo-oxidation depends upon the availability of oxygen
in the LDPE sample. It can be described by Figure 2-3.
Literature review
14
PH
+ h
H* + P*
O2
POO*
+ PH
POOH + P*
P* + H2O + PO* PO* + OH*
+ PH + PH
POH + P* + PH
H2O + P*
Initiation
Propagation 1
Branching
Propagation 2
Termination: POO* + POO* POOP+O2
P* + P* P-P
PO*+H* POH
P* +H* PH
Figure 2-3: Schematic representation of the auto-oxidation process (Peacock 2000)
Literature review
15
2.5. Traditional methods to dispose of LDPE
Currently, there are three methods of reducing the quantity and environmental
impact of LDPE waste are in practice. They are burying LDPE in land fill, incineration and
recycling. These methods are detailed below.
2.5.1. Landfill
Landfills are the physical facilities used for the disposal, compression and
embankment fill of LDPE in the surface soils of the earth (Ebnesajjad 2012). Burying
LDPE in the land fill is a traditional disposal method. At first landfill was traditionally
selected to dispose LDPE, because of its low cost. Later it became highly expensive
because the cost of available landfill sites rose. LDPE made plastic bags will take many
years to completely degrade that were disposed in to landfills (Stevens 2002). During
their degradation they produce harmful by products in to the surrounding. Along with
this, the leaching of additives of LDPE additionally damages the environment.
2.5.2. Incineration
Incineration is the combustion of waste LDPE material, converting it into carbon
dioxide and water. This was the most popular disposal method in the 1970s
(Goodship 2007). In this process, thermal energy from LDPE is recovered and can be
used to generate electricity or fuel pellets (Vlachopoulos 2009). It has been calculated
that combustion of 1 kg of LDPE yields around 43.3 MJ, which corresponds to 0.25
kg of pit coal (Arvanitoyannis 2010).
Unfortunately, incineration of commercially available LDPE (which contains
additives) produces toxic products, including phosgene, hydrogen chloride, hydrogen
cyanide and the greenhouse gases carbon dioxide and carbon monoxide (Peacock
2000). These compounds can not only cause respiratory difficulties in humans but also
damage the ecosystem. Incineration of LDPE and many other plastics is still in
practice, contributing to air pollution and presenting a severe threat to the environment.
2.5.3. Recycling
Recycling is a reprocessing method for turning waste plastic into useful
products. Recycling of LDPE, especially in the form of plastic bags, is in practice.
Literature review
16
Recycling can be classified as either pre-consumer recycling (primary recycling) and
post-consumer recycling (secondary recycling) (La Mantia 2002). Primary recycling
involves the recycling of scrapped and damaged LDPE during manufacturing. It is
generally practiced on the premises of manufacturing sites. For secondary recycling,
discarded plastic bags or mulch films are taken to another site for processing. LDPE is
generally recycled along with LLDPE, as they share similar properties and uses.
Though recycling is widely practiced it has several disadvantages. Recycling
involves heating polymers, which may cause oxidation of the polymer chains and alter
the properties of its constituent molecules (Moeller 2008). The quality of the LDPE
generated in this way decreases with the number of cycles performed. Recycled LDPE
films exhibit decreased tensile strength, decreased impact resistance and glow, which
diminish the quality and commercial importance of the product. In addition, the LDPE
recycling process is expensive, complicated and labour-intensive (Farag 2013). LDPE
bags usually contains commercial additives that make the recycling process less
profitable, while removing these additives compromises the quality of the recycled
product (La Mantia 2002). In practice, used LDPE mulch films have organic matter
attached to them, which interfere with recycling. Further, the presence of minute
quantities of polyurethanes (PU) and pesticide residues hampers the chemical processes
involved in recycling. Recycled products are seldom approved for applications
involving contact with food. These problems limit opportunities for the recycling of
LDPE. Sorting plastic types is another practical problem associated with recycling (La
Mantia 2002). On top of these problems, recycling does not address the core issue of the
ever-increasing volumes of LDPE.
2.6. Attempts to change the nature of LDPE
As the recalcitrance of LDPE to biodegradation stems from its chemical
composition, several attempts have been made to change this, which have led to the
invention of modified versions of LDPE. These can be classified as bioplastics, oxo-
PEs and biodegradable PEs. Among these, bioplastics were the first to be manufactured
and are now in considerable demand.
Literature review
17
2.6.1. Bioplastics
Bioplastics are carbon-based polymers with biodegradable components in their
structure that initiate oxidation and further degradation. The biological components of
bioplastics are generally derived from carbohydrates or vegetable oils (e.g., starch,
cellulose and pectin) (Stevens 2002). Soil disposal of these polymers favours biological
activity on their biological component, which leads to their biodegradation. The rate and
extent of biodegradation of bioplastics is dependent on factors such as the amount of the
biological component, the type of component and its distribution throughout the
polymer.
2.6.2. Starch-based LDPE
Starch is a plant-produced hygroscopic, translucent natural carbohydrate.
Bioplastics made of starch are semi-crystalline materials, composed of destructured
starch and plasticisers (Domb et al. 1997). The most widely used bioplastics are made
from starch as they are relatively inexpensive compared to other bioplastics (Timings
2004). Starch can easily be mixed with LDPE to produce bioplastics (Fellows 2000).
The composition and nature of starch-based LDPE can be modified by changing the
concentration of starch or by esterification, etherification and grafting (Kalia et al.
2011). The main idea behind these starch-based biopolymers is that upon
disposal in bioactive soil, the starch degrades rapidly, leaving behind the porous
polymer matrix (Kalia et al. 2011).
The main problem with starch bioplastics arises from the hygroscopic nature of
starch and its poor mechanical stability (Tucker et al. 2004). As the esters of starch are
more resistant to water, esterification of starch was introduced as a solution to this
problem. This resulted in manufacturing bioplastics with starch esters, or of blends of
starch with LDPE (Arvanitoyannis et al. 1998). Thus, manufactured starch-LDPE
composites are generally more expensive than traditional polymers.
2.6.3. Cellulose-based LDPE
Another class of LDPE bioplastics are made from cellulose. These composites
are stronger than their starch-based analogues. The hydroxyl groups of cellulose exhibit
Literature review
18
differences in regiochemistry and polarity, making it highly reactive to form LDPE-
cellulose composites (Kabasci & Stevens 2013). Thus, formation of esters and ethers of
cellulose is relatively easier than in the case of starch-based LDPE. Cellulose esters and
ethers that are linked with LDPE have been recognised to have better mechanical
quality. They also show better dyeing capacity and improved physical properties
(Medina-Gonzalez et al. 2012). Cellulose–LDPE composites are mostly used for
packaging. They are good for heat sealing and saving moisture, but are not tear and
moisture resistant. This limits their usage and demand in the market.
2.6.4. Lactic acid-based LDPE
Polylactic acid (PLA) is one of the preferred materials for bioplastics. It is
formed in a polycondensation reaction, in which the hydroxylic and carboxylic groups
of lactic acid (monomer) reacts with each other to form PLA (polymer) (Auras et al.
2011). PLA is not mechanically resilient, but its mechanical properties can be improved
by blending it with LDPE (Auras et al. 2011). As the microbial production of lactic acid
is cost-effective this is preferred over its chemical synthesis from polymerisation of
lactic acid. PLA forms composites with starch, caprolactone and chitosan, resulting in a
range of physical properties. These composites are non-volatile and odourless, and
hence are preferred over other bioplastics.
In terms of appearance, PLA-based LDPE composites are attractive for
packaging. Their hydrolytic properties make this composite useful in making adhesives
and binders. Along with this, it can be used in heat-resistant applications such as
manufacturing microwavable containers and electronic equipment. Compared to
standard LDPE, this bioplastic is more flexible but also more expensive.
2.6.5. Lignin-based LDPE
Lignin is an amorphous, aromatic plant-derived polymer that is relatively
inexpensive for manufacturing bioplastics. Lignin possesses low mechanical strength in
its native form, while blends of it with LDPE show high mechanical strength (Brenes
2006). Due to its phenolic chemical nature it can be easily modified and blended with
LDPE powder by a melt-blending process. Moreover, lignin acts as an absorber of UV
Literature review
19
radiation and stabilises LDPE polymer (Brenes 2006). Like many other bioplastics,
lignin-based LDPE is also an expensive composite.
Despite their environmental appeal, LDPE bioplastics also have several serious
shortfalls. They are generally more expensive than traditional LDPE (Tolinski 2011).
Also, they have inferior mechanical and physical qualities, making them less preferred
by the manufacturing industry.
2.7. Oxo-PE
Oxo-PE is typically PE to which a prodegradant (metal salts) has been added to
accelerate the reaction of LDPE with atmospheric oxygen (Robertson 2012). Upon
disposal in bioactive soil, the constituent photo-reactive metal ions react to UV radiation
to induce chain scission of the polymers. This disintegrates the high molecular weight
fractions of the polymer backbone into low molecular weight carbon chains. These
chains are then small enough to be metabolised by the soil microbiota. The most
commonly used metals in these polymers are the trace metal ions zinc, manganese and
iron (Koutny et al. 2006); these are known as prodegradants.
Oxo-PEs are designed to maintain the useful properties of plastic throughout
their lifetime and be competitive with traditional materials. Various types of oxo-
PEs are available with a range of prodegradants, including photo-degradable,
biodegradable and photo-biodegradable agents (Scott 2003). Most of these comprise
transition metal ions, which tend to form much lower molecular mass scission chains.
Following activation by metal ions, bacterial species such as Nocardia asteroides and
Rhodococcus rhodochrous degrade the low molecular weight fractions of the carbon
chains (Scott 2003). Oxo-PEs can also be recycled, during which process the metal ions
are neutralised.
These polymers have certain disadvantages. Accumulation of prodegradants in
the environment changes its natural composition and is sometimes fatal for
microorganisms (Thomas et al. 2010). Moreover, these plastics are relatively expensive
and as they take longer to biodegrade are not suitable for landfills (Thomas et al.
2010). Indeed, the extent of biodegradation of oxo-plastics is unclear. According to the
Literature review
20
American Society for Testing and Materials (ASTM) D6400 standard, this kind of
plastic is classified as non-biodegradable (Koyikkal 2013).
Oxo-PEs contain metals that compromise the recycling process when mixed
with normal plastics. Moreover, partially degraded oxo-plastics and their products can
attract and store pre-existing toxic products in the environment. If these polymers enter
the food chain, they may adversely affect human health. As a further harmful effect on
the environment, LDPE with prodegradants has been shown to produce 90 % carbon
dioxide upon disposal in landfills (Thomas et al. 2010).
2.8. Degradable PE
These PEs contain reactive groups or unsaturated carbon bonds in their polymer
chains that increases their susceptibility to biological attack. These reactive groups
(carbonyl, ester, or aldehyde) are introduced into the polymer matrix during
polymerisation. For example, carbonyl groups are introduced into the polymer chain by
co-polymerisation. In this method, ethylene monomers are allowed to react with carbon
monoxide to form carbonyl groups in the polymer backbone (Bremer 1982).
Alternatively, carbonyl groups can be introduced into the side chains by the co-
polymerisation of ethylene with vinyl ketone monomers (Sitek et al. 1976). In addition,
introduction of diene content into LDPE increases its biological susceptibility. The rate
of degradation is dependent on the diene content of the polymer (Scott 2007). Further,
introduction of unsaturation into LDPE can be achieved by new generation Ziegler–
Natta catalysts (Lemaire et al. 1991).
Another version of degradable LDPE is hydrolysable LDPE. This can be
prepared by the introduction of ester groups into the chain by co-polymerising ethylene
monomers with 2-methylene-1,3-dioxepane in the presence of toluene (Bailey et al.
1990). Alternatively, ester groups can be introduced into the main chains of the polymer
by reacting ethylene monomers with 2-methylene-1,3-dioxepane and carbon
monoxide in the presence of a free radical initiator (Austin 1994).
Literature review
21
Degradable LDPE is successful in satisfying the reduced plastic visibility
criterion. However, upon degradation it releases polymer fragments and additives into
the soil, which raises important ecological concerns (Roy et al. 2011).
2.9. Various standards of biodegradation
The ASTM is recognised worldwide for setting the standards for testing
materials. The D6400 standard describes the degradation of plastics and plastic products
under aerobic composting conditions (Niaounakis 2013). In this standard the rate and
products of biodegradation are also mentioned and described. The intention is to set the
standard for the labelling of biodegradable plastics. According to these standards, all
biodegradable products that are composed of a single polymeric material must be 50 %
mineralised after 6 months, while those composed of various materials must be
mineralised to 90 % (Rudnik 2008) in 6 months. Along with this, the by-products of
biodegradation must be non-toxic and should not discourage plant growth in the
compost.
The Commonwealth of Australia follows similar standards for labelling
biodegradable plastics (Standards Australia 2006). These are similar to the European
standards (EN13432), except that they include an animal eco-toxicology test. In these
standards, the materials must pass characterisation, biodegradability (aerobic),
disintegration and eco-toxicology tests and a recognisability test. The characterisation
tests examine the material for the possible release of harmful chemicals into the
environment, while the biodegradability tests involve measurement of the material’s
emission of carbon dioxide, methane and other chemicals upon degradation. These tests
also examine the alterations caused by biological action on the test material. The
disintegration test checks the propensity of the tested material to fall apart upon
biodegradation. The tested material must disintegrate into fragments smaller than 2 mm
in 12-week pilot-scale composting bins under aerobic conditions. The eco-toxicity tests
measures the toxicity exhibited by soil containing the biodegraded products (that is, the
compost). This is done by testing the toxicity to earthworms and plants.
In the case of animal eco-toxicity, the morbidity rates and weights of surviving
earthworms are measured, whereas in the plant eco-toxicity test, biomass and
Literature review
22
germination rates are used as indicators. The recognisability test addresses the proper
labelling of the product. According to these standards, a minimum of 90 %
biodegradation of plastic materials must be achieved within 180 days of composting,
and the original plastic must contain more than 50 % organic matter. Oxo-PE plastics
require more time to biodegrade and thus they are not considered biodegradable.
The biodegradation of polymers is represented by these equations (Chiellini et
al. 2001):
Aerobic biodegradation
C polymer + O2 CO2 + H2O + C residue + salts
Anaerobic biodegradation
C polymer CO2 + H2O + C residue + salts + CH4
The rate of biodegradation must be consistent within a prescribed pathway. The
Deborah number (D) was proposed to distinguish between biodegradable and non-
biodegradable polymers.
D = time of degradation/human lifetime
Non-biodegradable polymers have higher D values than biodegradable polymers
(Domb et al. 1997).
2.10. Terminology of biodegradation
Biodegradation is a broad term without a precise definition. In simple terms,
biodegradation can be defined as a natural process by which organic chemicals are
converted to simpler compounds and mineralised (Ren 2011). In 1992, an international
workshop on biodegradability was held in Annapolis, Maryland, US, to define the terms
of biodegradation (Adsantorian 1992). The participants agreed to these important
points:
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23
Materials that are manufactured as biodegradable must mention the
specific disposal method (e.g., sewage treatment, denitrification or
anaerobic sludge treatment) to be used.
The rate of degradation of a biodegradable material should be
consistent with the disposal method and other components of the
biodegradation pathway to control its accumulation.
The biodegradation pathway should lead to the formation of
intermediate products like biomass and humic acid, while the final
products should be carbon dioxide and water.
The biodegradation pathway should not be negatively influenced by
products formed due to the degradation.
According to ISO 472:1988, biodegradable plastics are designed to undergo a
change in chemical structure under specific environmental conditions, facilitated by
naturally occurring microorganisms. This results in a loss of some properties as
measured by standard tests and methods appropriate to plastics. The Japan BioPlastics
Association proposed that polymers can be classified as biodegradable plastics if they
can be changed into lower molecular weight compounds in a process that involves
metabolism by naturally occurring organisms in at least one step.
The latest definition was provided by ASTM in standard D-5488-94d.
According to this document, biodegradation is defined as a process in which the
decomposition of a material occurs predominantly by the enzymatic action of
microorganisms that convert these materials into carbon dioxide, methane, water,
inorganic compounds and biomass (National Institute of Industrial Research 2006). The
biodegradation process can be measured by standard tests over a specified time,
reflecting the available disposal conditions.
2.11. Mode of biodegradation
Biodegradation of solid polymers like LDPE occurs generally by two methods
i.e. surface erosion method or bulk degradation method (Ratner et al. 2012). In surface
erosion method, the polymer starts to degrade from the exterior portion to interior
leading to the thinning of it with time. In this type of biodegradation, molecular weight
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of polymer remains constant. Rate of surface biodegradation is generally follows zero
order kinetics and depends on the available surface area of polymer (Kumbar et al.
2014).
In bulk erosion, polymer biodegradation occurs throughout the polymer matrix
at the same time. Molecular weight decreases with the increasing time and the matrix
dimensions remains constant till the total mechanical failure. In this type of
biodegradation polymer allows penetration of water into the bulk of material (Ratner et
al. 2012). Usually bulk erosion follows first order kinetics (Bader & Putnam 2014).
Figure 2-4 schematically indicates surface and bulk biodegradation of LDPE.
Figure 2-4: Biodegradation by bulk degradation (A) and by surface erosion (B)
The determination of whether a material undergoes surface erosion or bulk
erosion depends on various factors. They are linkage between monomers, method of
chain scission of polymer back bone, mechanism of hydrolysis of monomer units, glass
transition temperature, surface-to-volume ratio and porosity of polymer (Domb &
Kumar 2011). Primarily it depends on hydrophobic nature of polymer matrix. Following
A. Bulk erosion
B. Surface erosion
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25
this polymer architecture and its thickness also plays an important role in determining
its biodegradation mechanism (Ratner et al. 2012).
2.12. Types of LDPE biodegradation
Biodegradation of LDPE can be classified as macrobiological and
microbiological. Some reports suggest that certain insects can secrete fluids that degrade
PE films (Anderson et al. 1995). This is an example of macro (passive) biodegradation
that leads to the deterioration of PE particle size. However, it is a rare process and its
impact is negligible. Active biodegradation by microbes is comparatively faster, with
bacteria and fungi commonly used. Bacteria prefer simple carbon sources (i.e., glucose)
for metabolic purposes. In a selective environment in which carbon sources are
restricted, bacteria and fungi can consume polymers. This process is made more
efficient by the isolation, enrichment and study of microbes that degrade polyolefins
such as LDPE.
2.13. Previous attempts to degrade LDPE
Attempts to biodegrade LDPE in its native form have been made with little or no
success. LDPE has been found to emit only 0.2 % carbon dioxide when incubated in
bioactive soil for more than 10 years (Albertsson 1977; Albertsson & Banhidi 1978). In
other words, the biodegradation of LDPE is almost negligible over 10 years. In order to
explain this, several closely related reasons have been proposed. In this section of the
literature review, these reasons are explained and analysed.
2.13.1. The high molecular weight of LDPE
LDPE’s resistance towards biodegradation stems from its high molecular
weight. It has been shown that low molecular weight paraffins (e.g., PE) are readily
degraded by microbes (Albertsson & Karlsson 1993), while their biodegradability
decreases with increasing molecular weight (Potts et al. 1973). It has been demonstrated
that PE chains with weights of more than 450 Da (C32) are not biodegradable and cannot
be used as a carbon source by microbes (Peacock 2000). In the case of hydrocarbons,
n- alkanes and branched alkanes with C10–C20 are easily biodegradable, while longer
chains of more than 20 carbon atoms are difficult to biodegrade (Speight &
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26
Arjoon 2012). This indicates that as the chain length increases, resistance towards
biodegradation also increases.
2.13.2. The hydrophobic nature of LDPE
The hydrophobicity of LDPE is another reason for its resistance to
biodegradation. In general, microbes are attracted to hydrophilic surfaces or substrates
due to the charge on their cell membranes. This results in the formation of a biofilm,
which is believed to be the initial phase of biodegradation. Hydrophilic polymers adsorb
water into their structure, encouraging the formation of biofilm by microbes and
initiating biodegradation. Thus, like proteins or carbohydrates, hydrophilic polymers are
easily biodegradable. In contrast, as LDPE is hydrophobic, it resists biodegradation
(Hatada et al. 1997).
2.13.3. The 3-D structure of LDPE
The cross-linked chains of these polymers are also responsible for their
resistance to biodegradation. This type of structure prohibits biological entities from
entering the bulk of the polymer, suppressing biological activity, and hence
biodegradation. Branched hydrocarbon chains are more resistant to biodegradation than
their linear chain isomers (Vandecasteele 2008). Likewise, non-crystalline synthetic
polymers are generally more biodegradable than crystalline ones (Ahmed et al. 2012).
Other factors, such as the surface properties of polymers and their
stereochemistry, also influence biodegradation. Polyhydroxybutyrate with atactic side
chains is less biodegradable than its isotactic analogue (Kemnitzer et al. 1992). It
has also been observed that rough surfaces are more biodegradable than smooth ones
(Moore & Saunders 1998). Biodegradation depends on the utilisation of the polymer by
various microbial species. It has been demonstrated that bacterial consortia were more
efficient than a single Aspergillus strain (Mark 2007). Observation of LDPE buried in
soil indicated an increase in the average molecular weight. This indicates that the smaller
polymer chains were biodegraded by microbial consortia, leaving high molecular
weight polymeric chains intact.
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2.14. Surface characterisation techniques
In order to trace LDPE biodegradation, it is essential to characterise the surface
of LDPE by surface characterisation techniques. These techniques will provide the most
useful data regarding the mechanisms of biodegradation. Techniques for the physical
and chemical characterisation of LDPE surfaces are discussed below.
2.14.1. FT-IR
FT-IR is a non-destructive, micro-analytical technique that is used to check
chemical composition and bonding arrangements of LDPE. It is accepted by the ASTM
to analyse polymers like LDPE (Lampman 2003). FT-IR uses infrared energy to
produce vibrations within the molecular bonds of the polymer. During this excitement,
transition from one vibration state to another occurs at characteristic frequencies,
revealing the structure of the sample (Bower & Maddams 1992). Molecules exhibit
different vibration states depending on their chemical bonding. These vibrations can be
classified as either stretching or bending vibrations.
Stretching vibrations: In this mode, the distance between the two atoms on the
bond remains the same while they vibrate with the atoms of the bond remaining in the
same bond axis. Stretching vibrations are sub-classified as symmetrical and non-
symmetrical stretching. In the case of symmetrical stretching, both atoms move in or out
simultaneously, while in asymmetrical stretching one atom moves in, while the other
moves out. In the case of the LDPE spectrum, symmetrical and asymmetrical bond
stretching can be observed with –CH2 approximately at 2,855 cm-1 and 2,920 cm-1
respectively (Gulmine et al. 2002).
Bending vibrations: In this vibration type, the atom position changes relative
to the original bond axis. These vibrations occur in four types:
Scissoring: In this mode of vibration, the movement of atoms is in the opposite
direction, with a change in both bond angles and axes relative to the central atom.
Rocking: In this type of vibration, movement of atoms takes place in the same
direction, with a change in their bond axes with respect to the central atom.
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Wagging: In this movement, both the atoms move simultaneously above and
below with respect to the common atom.
Twisting: In this movement, both atoms move above and below at different
times with respect to the common atom (Mohan 2004).
LDPE polymers show all of these vibrational states, most of which can be
identified by FT-IR. Table 2.4 shows the vibration modes of atoms of LDPE and their
signal intensities from its FT-IR spectrum.
Table 2-4: Assignment of IR absorption peaks for LDPE (Gulmine et al. 2002)
Band (cm-1) Assignment Intensity of signal
2,919 CH2 asymmetric stretching Strong
2,851 CH2 symmetric stretching Strong
1,473 and 1,463 Bending deformation Strong
1,377 CH3 symmetric deformation Weak
1,366 and 1,351 Wagging deformation Medium
1,306 Twisting deformation Weak
1,176 Wagging deformation Very weak
731–720 Rocking deformation Medium
FT-IR may operate in either the transmission or the reflection mode. The
apparatus consists of an IR source, an interferometer, a sample holder and an IR
detector (see Figure 2-5). The IR source produces infrared radiation that exits through
an aperture and impinges on an interferometer, which then produces an interferogram
signal that has all the IR frequencies encoded in it. The resultant beam from the
interferometer enters the sample compartment, reflecting off or being transmitted
through the sample. Finally, the beam passes to a detector which is designed to measure
the interferogram signal generated from the sample (Smith 2011).
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29
Figure 2-5: Schematic diagram of an FT-IR spectrometer
Though FT-IR is a qualitative technique it can be used to derive quantitative
data from the spectrum. The crystallinity of LDPE and the absorption intensities of its
functional groups can be calculated using FT-IR. Changes in these functional group
intensities may reveal the chemical reactions that occur during biodegradation. These
functional group intensities can be calculated using the following formulae:
Keto carbonyl bond (C=O) index = I1715/I1465
Ester carbonyl bond index = I1740/I1465
Vinyl bond index = I1640/I1465
The percentage crystallinity can be calculated from the FT-IR spectrum using
the following formula (Zerbi et al. 1989):
Where Ia and Ib are the absorbance values determined from the bands at 1,474
and 1,464 cm-1 or 730 and 720 cm-1. According to Kaci et al. (1999), using 730 cm-1 and
720 cm-1 is preferable to using 1,474 cm-1 and 1,464 cm-1.
2.14.2. Raman spectroscopy
Raman spectroscopy is used to identify weaker signals that are not observed in
the FT-IR spectrum of LDPE. This technique offers a completely non-destructive option
Interferometer Interferogram Sample
Detector Computer, FFT Spectrum
Source
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for surface analysis (Koenig 1999). Beyond general characterisation of LDPE, Raman
spectroscopy also probes crystallinity and indicates its molecular weight.
Raman spectroscopy relies on the Raman effect. When monochromatic light is
focused on a molecule, it can be either scattered or absorbed. Most of the scattered light
will have the same frequency as the incident light (Rayleigh scattering). However, a
small fraction of the monochromatic light (~1 in 107 photons) will be scattered
inelastically at frequencies that are different from the incident photons (Atkins & de
Paula 2013). The energy difference between the incident and scattered photons is
proportional to the vibrational energy of the molecule. This process of energy exchange
between scattering photons and the incident photons is called the Raman effect. Raman
spectroscopy selectively collects these inelastically scattered photons.
Raman instrumentation consists of a microscope, an excitation source (laser),
filters, slits, a diffraction grating, the necessary optics, a detector and analytical software
(see Figure 2-6). A laser light (green or red) impinges on neutral density filters that
decrease the intensity of the laser beam that passes through them. The laser beam then
passes through a spatial filter, which focuses the beam on the sample. This light is then
directed to the microscope attached to the sample. Once the laser beam impinges on the
sample, both elastically and inelastically scattered light emerges from it. A holographic
filter removes elastically scattered light on the return path. Inelastically scattered light is
then separated by the diffraction grating into discrete wavelengths, and each frequency
is measured simultaneously. The final Raman signal is directed to the detector,
producing the resultant spectrum of the sample (Begum et al. 2010).
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Figure 2-6: Schematic representation of Raman spectrometer (Begum et al. 2010)
Raman spectra can also be used to obtain quantitative data on LDPE. It has been
observed that the intensity of emission at 890 cm-1 inversely indicates the approximate
molecular weight of LDPE (Lobo & Bonilla 2003) (i.e., if the emission intensity at 890
cm-1 is high, the approximate molecular weight of LDPE is low and vice versa). Also,
the amorphous proportion quantity of LDPE can be calculated by measuring relative
intensities at 1,080 nm using Strobl et al.’s formula (Strobl & Hagedorn 1978):
αa = I1,080/0.79
Raman spectroscopy can be used as a complementary technique to FT-IR.
Vibrational modes of atoms, such as scissor vibrations and rocking vibrations, are
barely visible in FT-IR spectra while they are very clear in Raman spectra. Likewise,
asymmetric stretch is barely visible in Raman spectra, while it is clearly visible in FT-
IR spectra (Koenig 1999). Table 2.5 summarises the assignments of Raman bands of
LDPE (Sato et al. 2002).
Microsco
pe Holograp-
hic filters Slit
Diffraction Grating
Dove
CCD
Camera
Laser
Mirror
Rejection
Filter
Spatial
filter
Sample
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Table 2-5: Wave numbers and their corresponding portions
Wave number in cm-1 Nature of peak Features
1,437 Amorphous CH2 bending vibration
1,292 Crystalline CH2 twisting
1,126 Crystalline Symmetric C–C stretching
1,062 Crystalline Asymmetric C–C stretching
897 Amorphous C–C stretching (short branches)
2.15. Surface visualisation
Fungal treatment may result in permanent changes on the LDPE surface.
Visualising these changes in detail could increase our understanding of the
biodegradation process. The atomic force microscopy (AFM) and scanning electron
microscopy (SEM) techniques that were employed to characterise LDPE surface are
discussed in the sections below.
2.15.1. AFM
AFM is a surface scanning technique that allows mapping and measuring of the
topography of materials such as LDPE (Dorobantu & Gray 2010). The basic principle
of AFM is that a probe is maintained in close contact with the sample surface with the
help of a physical mechanism. As the probe scans the surface of the sample, the change
in sample surface and probe distance is monitored and recorded (Haugstad 2012). The
most commonly used type probes are micro-fabricated silicon (Si) or silicon nitride
(Si3N4) cantilevers with integrated tips. AFM can be performed in two types of scanning
modes: the contact mode (static mode) and the semi-contact mode. In the contact mode,
the tip or probe is in constant contact with the sample surface. The probe maintains a
constant force between the tip and the surface of the sample. In the semi-contact mode,
or tapping mode, the tip of the probe ‘taps’ the surface of sample. Here, the probe is
allowed to oscillate at a value that is close to its resonant frequency. When the probe
oscillates near the sample surface, the amplitude of this oscillation decreases. Thus, as
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the surface is scanned by the probe, the oscillatory amplitude decreases or increases,
depending on the topography of sample (Bowen & Hilal 2009).
AFM instrumentation consists of a sample mount, a laser source, a
discriminator, a detector and the output (McMahon 2008). The laser is focused on the
tip that is attached to the cantilever. When the tip moves over the surface, the cantilever
also moves along, and with it the laser as well. This deflection is recorded by a
photodiode detector and is converted into the topographical information of the sample.
AFM produces topographical information that is displayed by using colour maps
for height. It can also be used to measure the sample elasticity by pressing the tip of the
cantilever into the sample and measuring its deflection. AFM is used to measure the
stiffness and crystallinity of samples.
2.15.2. SEM
SEM is a powerful surface characterisation technique that utilizes focused beams
of electrons to obtain information. When primary electrons from the electron source
bombard the sample, it emits x-rays, secondary electrons (SE) and backscattered
electrons (BS). Among each of these electron types, secondary electrons are emitted
from the topmost part of the sample surface (Reed 2005). Thus generated secondary
electrons are used to produce an image of the sample surface. In order to produce these
secondary electrons, samples must have electrical good conductance. However, since
LDPE has low electrical conductance, they were sputter-coated with gold, which has
high electrical conductance.
SEM instrumentation consists of an electron gun, or emitter, condenser lenses,
deflection coils, a final lens, an electron detector, an amplifier and a screen. The
electron gun emits primary electrons that are condensed by the condenser lenses. These
are then deflected in deflector coils and enter the final lens. The deflected electrons are
rastered around the sample. Secondary electrons generated from the sample surface are
then detected by the electron detector. The secondary electron signals are amplified and
projected on the screen.
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SEM is used to obtain qualitative data from samples. It is used to check the
surface feature of LDPE (topography), shape and size of particles that make up the
LDPE surface (morphology) and the elements and compounds of the constituent
molecules on the surface of LDPE of samples (composition).
2.16. Crystallinity of LDPE
Crystallinity is one of the key factors that influence biodegradation of polymers.
LDPE consists of both amorphous and crystalline portions, making it semi-crystalline in
nature (Vasile & Pascu 2005). In crystalline LDPE, the matrix molecules are arranged in
a regular and organised manner. Adjoining these portions, the molecules are arranged at
random and in a disorganised state, referred as the amorphous portions (Gowariker et al.
1986). The crystallinity of a polymer is a relative expression of its crystalline content
with respect to its amorphous content. The percentage crystallinity influences the
physical properties of polymers, such as their hardness, modulus, tensile strength,
stiffness, crease and melting point (Salamone 1996). Crystallinity of LDPE primarily
depends on the extent of its chain branching. As more branches are introduced,
disruption between the chains becomes greater, resulting in a decrease in its
crystallinity. The crystallinity of LDPE is generally calculated using the X-Ray
diffraction (XRD) technique and.
2.16.1. XRD
The XRD technique is a non-destructive method that employs x-rays to scan the
polymer matrix. X-rays are electromagnetic radiation with wavelengths in the range of
0.5 to 2.5 A (Guinier 1994). Due to their smaller wave lengths, x-rays can penetrate
deep into materials, such as LDPE, and provide information about the bulk structure.
Like many other rays, x-rays primarily interact with electrons in atoms. When
the photons in x-rays collide with electrons, some photons from the incident beam will
be deflected away from their incidental angle. If the wavelength of these scattered x-
rays does not change, the process is called elastic scattering (Thompson scattering)
(Moore 2008). These elastically scattered photons are used in the XRD method. During
their scattering, these photons interfere with each other, cancelling their paths. This
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pattern of scattering and the interference of the photons depend on the crystal structure
of the sample.
XRD data can be interpreted by using Bragg’s law (2dsinθ=nλ). In this formula
θ is the scattering angle of the x-rays; λ is the wavelength while n is an integer
representing the order of the diffraction peak (Suryanarayana & Norton 1998). This law
can be applied for any materials with periodic distribution of electron density, as they
scatter incidental x-rays. In other words, Bragg’s law applies to molecules or collections
of molecules, colloids, proteins, virus particles and polymers (LDPE).
XRD instrumentation consists of an x-ray generator (a cathode ray tube), a
sample holder and a detector. Electrons are generated by heating a filament, and then
they are accelerated towards a target and bombard it. This bombardment generates x-
rays from the target material. These materials are generally made up of Cu, Fe, Mo and
Cr (Suryanarayana & Norton 1998).
XRD can be used to check crystallinity, the crystalline microstructure of LDPE
and its orientation. The percentage of the crystalline portion of LDPE can be measured
by calculating the total area under the crystalline peaks and dividing this by the total
area of all peaks. The crystalline peaks can be identified by comparing XRD of LDPE
with that of archived graphs. The formula used to check the percentage crystallinity is
given below.
Percentage crystallinity = (total area of crystalline peaks)/ (total area of all peaks) or
Percentage crystallinity = IC × 100 (Jaffe et al. 2012).
IC+IA
2.17. Classification of fungi
The biodegradation process can be best understood by classifying the microbes
employed in it. There are a number of approaches that have been utilised for
taxonomical characterisation of fungi. These are mainly macroscopic (colonial),
microscopic and DNA sequencing-based methods (Campbell et al. 2013).
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2.17.1. Macroscopic classification
At first, fungal colonies on agar plates are characterised by observing their
macroscopic features. These include the following: the colour and tint of the colony on
both sides of the culture plate; the odour of the culture; the surface structure of the
colony (cottony, crustaceous, embedded, velvety); the pattern of the colony (arachnoid,
flowery, radiate, zonate); the margins of the colony (irregular or smooth); the growth of
the colony (restricted or spreading); the colour and nature of any pigment exuded
(colour or watery) (Watanabe 2010). By observing these patterns basic information
about fungi can be obtained.
2.17.2. Microscopic classification
Next classification can be done by observing the microscopic features of fungi.
The visual characteristics of the mycelia are one of the key features of all fungi. The
presence or absence of conidia, their shape, size and location on the mycelium helps in
identifying fungi. For example, Fusarium produces characteristic sickle-shaped conidia
that allows its identification (Engelkirk & Duben-Engelkirk 2008).
2.17.3. DNA sequencing-based classification
Both macroscopic and microscopic features are not enough to classify fungi in
detail. This can be achieved by sequencing conserved DNA regions of the fungal
genome. The nuclear-encoded ribosomal RNA genes (rDNA) have been extensively
used for this purpose (Berbee & Taylor 1992; Swann & Taylor 1993). These rDNA
genes are arranged in tandemly repeated units within the fungal genome.
Each repeated unit contains genes that encode the small subunit (18 S), the 5.8 S
and the large subunit (25–28 S) of the ribosome. Each repeated unit is separated by one
or more intergenic spacer (IGS) regions. Within the repeated units, the 5.8 S RNA
coding portions are flanked by conserved sequences known as the internal transcribed
sequences (ITS).
The internal transcribed sequence regions of rDNA have been shown to be
particularly useful for the molecular identification of fungi due to their consistent
evolutionary rate. The ITS1 and ITS4 regions are highly conserved in fungi, making
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37
them universal primers to identify fungal strains (White et al. 1990). The sequences of
these ITS, and the flanking SSU and LSU rRNA genes (see Figure 2-7) make them ideal
targets for DNA primer sites to amplify the total ITS region.
Figure 2-7: Schematic representation of the location of the ITS1 and ITS4 primer sites
(Kües 2007)
2.18. Microbial adhesion to hydrocarbons test
This test was used to examine the cell surface hydrophobicity of the microbial
(fungal) species that were participating in biodegradation. It is based on partitioning
microbial cells at an aqueous-hydrocarbon interface (Saini 2010). In this technique and
aqueous suspension of microorganisms is mixed with hydrocarbons (hexadecane or
toulene), and the hydrophobic cells tend to bind more to the hydrocarbons. The original
microbial adhesion to hydrocarbons (MATH) test was developed by Rosenberg
(Rosenberg 1984) for bacteria. Later it was adopted to measure the hydrophobicity of
fungi (Smith et al. 1998; Holder et al. 2007). In this method differences in absorbance
between the aqueous and hydrocarbon phases are measured at 470 nm. The
hydrophobicity of fungal cells was calculated using the formula below:
Hydrophobicity = (A470 of blank) - (A470 of hexadecane treated sample)
A470 of control
However, the hydrophobicity of microbes can be changed depending on the
nutritional conditions they are grown under. It has been observed that carbon deficiency
in the nutritional medium results in changes in the hydrophobic nature of bacteria
(Sanin et al. 2003). Similarly, fungal species demonstrate changes in their surface
properties depending on carbon availability in the media (Kulakovskaya &
Kulakovskaya 2013).
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2.19. Dissolved carbon dioxide content
Aerobic biodegradation of polymers releases water, carbon residues, biomass
(humus) and carbon dioxide as by-products (Mohan & Srivastava 2011). Therefore, the
degree of aerobic biodegradation can be measured by calculating the total amount of
carbon dioxide gas released from the polymer. There are two types of carbon dioxide
measuring methods in practice.
One set of carbon dioxide evolution methods depends on the respiration of
microbes that was employed for biodegradation. During the incubation period, microbes
produce carbon dioxide as a result of their respiration. Thus produced carbon dioxide is
measured and taken as a confirmation of biodegradation. The Sturm test (Sturm 1973) is
one of the widely used respirometric method to quantitate the carbon dioxide. However,
this test method has two major limitations. First, it can be conducted only for 1 month,
whereas most of the biodegradation experiments must be conducted for longer time
periods. Second, the conditions of the Sturm test are not conductive to growth of
filamentous fungi such as Fusarium (Palmisano & Barlaz 1996). These problems limit
the usage of Sturm test for evaluating the biodegradation of LDPE by Fusarium.
Other types of CO2 evolution measurement techniques rely on the solubility of
carbon dioxide in water. Upon its release from polymer, carbon dioxide readily
dissolves in water, resulting in formation of carbonic acid (H2CO3). This acid can be
measured and used as an indication of the evolved carbon dioxide content. However,
this method cannot be used as a direct indication of biodegradation as the salts in the
microbial media change the solubility of the carbon dioxide produced.
2.20. Methylene blue test
Changes on the surface of LDPE caused by biodegradation can be observed by
FT-IR, SEM, AFM and Raman spectroscopy. However, all these techniques are either
expensive or cumbersome to perform. In order to overcome this problem, a simple
methylene blue staining technique is proposed. Though using staining techniques to
locate oxidised sites on polyolefins is not new (Da Costa et al. 1990), they have never
been used to quantify biodegradation.
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This technique exploits differences in the affinities between methylene blue and
LDPE pellet types and measures these differences spectrophotometrically. According to
Beer-Lambert’s law, when monochromatic light travels through a solution, its
absorbance (A) depends on the solute concentration (c) and the distance (d) that the
light travels thorough the solution (d) (A = ε c d) (Kotz et al. 2009). When LDPE pellets
are incubated with methylene blue solution, the dye attaches to the oxidised portions of
LDPE. This attachment decreases the concentration of the methylene blue solution and
results in a decrease in the absorbance of the solution. When these absorbances are
measured at appropriate wavelengths the differences indicate the extent of
biodegradation. In general, biodegradation results in a decrease in oxidised sites on
LDPE and hence the absorbance of the resultant solution would be less than that of a
solution tested with intact LDPE pellets.
2.21. Fusarium oxysporum
Fusarium oxysporum is an anamorphic fungal species that includes both
pathogenic and non-pathogenic strains (Gordon & Martyn 1997). It is primarily a plant
pathogen, with many forms that cause wilt disease, resulting in extensive crop damage
every year (Narayanasamy 2013). Based on their host range, these plant pathogenic
forms are grouped into formae speciales and are named after them. For example
Fusarium oxysporum that attacks tomatoes is called F. oxysporum f.sp. lycopersici and
that which attacks ginger is known as F. oxysporum f.sp. zingiberi.
This fungus has distinctive macroscopic features that allow its preliminary
identification. On agar plates Fusarium oxysporum grows at a rapid rate. Its colonies
have a cottony surface and possess purple aerial mycelia. The reverse side of the agar
plate is generally colourless or dark purple. Under the microscope it exhibits
characteristic sickle-shaped macroconidia along with foot-shaped basal cells (Reiss et
al. 2011).
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Table 2-6: Classification table of Fusarium oxysporum
Kingdom Fungi
Phylum Ascomycota
Order Hypocreales
Class Sordariomycetes
Family Nectriaceae
Genus Fusarium
Species Oxysporum
Fusarium oxysporum can cause root infection in plants optimally at 30 °C, while
growing optimally at 28 °C. It is a soil-borne pathogen and can survive for extended
periods of time in the soil during unfavourable conditions. When favourable conditions
arise the fungus can germinate and penetrate the host roots. At first, it enters the plant’s
vascular system and uses the xylem vessels to rapidly colonise the host. This physical
occupancy of Fusarium oxysporum provokes the characteristic wilt symptoms
(Fusarium wilt) in its host (Beckman 1987).
Fusarium oxysporum secrets a number of hydrolytic enzymes that can degrade
host cell walls during the penetration process (Gupta 2012). These include chitin
synthase, protein kinase, pectate lyase and endo-polygalacturonase, among others.
These are collectively known as cell wall degrading enzymes (CWDE). They play an
important role in at least two phases of Fusarium wilt; first, at the time of penetration
through the root cortex of the host, and second, during colonisation by spreading
upward through xylem vessels (Beckman 1987). It has been shown that CWDE
secretion by Fusarium oxysporum can be induced by the presence of host cell walls
(Jones et al. 1972). Further, CWDE secretion is repressed by catabolites generated
during the degradation of the host cell walls (Cooper & Wood 1973). As pectin is the
most important component of plant cell walls, the pectate lyase of the CWDE plays an
important role in its pathogenesis (Beckman 1987). Pectate lyase activity can be
detected inside tomato root and stem tissues infected by Fusarium oxysporum
f.sp.lycopersici (Roncero 2003). Along with plants, certain strains of Fusarium
oxysporum can attack humans and cause severe health issues, such as keratitis and
Literature review
41
mycosis (Reiss et al. 2011). Fusarium oxysporum has a unique capability to produce
long chain hydrocarbons depending on its growth condition (Ladygina et al. 2006).
It is important to note that most of the LDPE degrading fungi so far isolated
possess cell wall degrading mechanisms. For example, Aspergillus niger and
Phanerochaete chrysosporium possess mechanisms that can attack plant cell walls and
LDPE, as does Fusarium oxysporum (Arutchelvi et al. 2008; De Vries & Visser 2001;
Aro et al. 2005).
2.22. Antioxidants
Antioxidants are added to LDPE during manufacturing. Their function is to
prevent oxidation of the polymer during manufacturing, usage and at any other stage of
the polymer’s life. These compounds are classified as primary and secondary
antioxidants. Primary antioxidants are used to protect the finished polymer (aromatic
nitro compounds, Irganox®). Secondary antioxidants are used during processing of
polymers (organic phosphites, thioethers) (Vasile & Pascu 2005).
Irganox® is a sterically hindered phenolic antioxidant that is added during the
polymerisation process. Irganox® has the ability to react with free radicals and
render them inert. It also prevents the production of free radicals in the LDPE matrix.
The compound is denoted as AH in the scheme below. The hydrogen atom becomes
attached to phenolic compounds, with little dissociation energy. The free radical A*
formed in this process acts as a scavenger of other free radicals.
ROO* + AH ROOH + A*
ROO* + A* ROOA
2.23. Biofilm
Biofilms are matrix-enclosed microbial accretions that adhere to biological or
non-biological surfaces (Madsen 2011). The formation of biofilm is thought to be the
initial stage of biodegradation or biodeterioration (Mohan & Srivastava 2011).
Formation of biofilm can achieved by the action of either exo- or endo-depolymerase
enzymes (Jayasekara et al. 2005). Fusarium has been shown to form biofilm on a wide
Literature review
42
range of substrates, including soft contact lenses (Chang et al. 2006) and also on
plumbing systems (Short et al. 2011).
Defining ‘biofilm’ is not an easy task, as it refers to a wide range of structures.
One definition by Costerton et al. (1995) is that ‘biofilms are complex communities of
microorganisms attached to the surface or interface enclosed in an exopolysaccharide
matrix of microbial and host origin to produce spatially organised 3-D structures’(Jass
et al. 2003). The formation of biofilms by microbes can be observed in porous
materials, reservoirs, plumbing systems, pipelines and on separation membranes.
Generally, these films adhere to solid substrates and exist at water–solid, water–air or
solid–air interfaces. In the case of solid–water biofilms, high amounts of humic
substances have been observed. It was observed that plastics like polyamides encourage
the formation of biofilms on them in river waters (Flemming 1997).
The composition of biofilm varies with the type of microorganisms and
polymers involved in its formation. Biofilms are made of accumulated microorganisms,
extracellular polymeric substances (EPS), cations, colloidal and dissolved components
(Wingender et al. 1999). They may contain proteins, nucleic acids, lipids (Nielsen et al.
1997) and phospholipids (Takeda et al. 1998). Along with these compounds, biofilms
may also consist of organic components like acetyl, succinyl or pyruvyl groups and
inorganic components like sulphates (Wingender et al. 1999). Along with these, DNA
has even been identified in some bacterial biofilms (Sutherland 2001).
Biofilms are secreted by various mechanisms. These include active secretion of
EPS by microbes, shedding of cell surface material by microbes, cell lysis of attached
microbes and absorption from the environment (Wuertz et al. 2003). In gram negative
bacteria (e.g., Neisseria gonorrhoeae) the spontaneous release of integral cellular
components has been noted (blebbing) (Jordan et al. 2004).
Once microbes adhere to the surface of the polymer via a biofilm, they start
degrading it by various mechanisms (Flemming 1998). In the case of hydrocarbon
biodegradation, it has been observed that microbes produce superficial structures that
help them to attach to the substrate (Volke-Seplveda et al. 2002). It has previously been
observed that bioerosion of photo-degradable PE resulted in the loss of molecular
Literature review
43
weight only at the point of interaction between the polymer and the biofilm. In this case,
the decrease in molecular weight was confined to the surface of the polymer
(Bonhomme et al. 2003). In contrast, biodegradation has been observed in shaking
cultures where biofilm formation is not possible or is minimal. In this study, an attempt
has been made to investigate the importance of biofilm formation by the isolated
Fusarium oxysporum strain in the biodegradation of LDPE. A method involving
separating Fusarium oxysporum from LDPE pellets was devised to examine the
requirement for biofilm formation, and is detailed in Section 3.13.
2.24. Co-metabolism
The term co-metabolism refers to the simultaneous degradation of two
compounds, where the degradation of the secondary compound depends on the presence
of the primary compound (Neilson & Allard 2008). More precisely, this is ‘the
transformation of non-growth substrate in the obligate presence of a growth substrate or
another transferable compound’ (Dalton et al. 1982).
Co-metabolism can be observed in natural ecosystem as it accommodates the
simultaneous degradation of various recalcitrant compounds. Co-metabolism has been
demonstrated in pure and mixed cultures for a wide range of compounds, including
insecticides, fungicides and surfactants (Bitton 2011). Co-metabolism generally
encourages the biodegradation of recalcitrant organic compound like LDPE
(McCutcheon & Schnoor 2004). Though the actual reasons are not clearly understood
for this, several possible explanations are outlined below.
2.24.1. Pre-exposure to an analogue compound
Co-metabolism of recalcitrant compounds by microbes can occur because of
similarities in the molecular structures between primary and secondary metabolites. For
example, bacteria grown on phenol or naphthalene are able to oxidise 4-chlorophenol
(Loh & Wang 1997). Unlike 4-chlorophenol, naphthalene and phenol do not possess
any chlorine atoms in their atomic structure. In this case, bacteria do not use the energy
derived from this reaction for their growth or metabolism. Rather, the similarity in the
structures of these compounds induces the production of bacterial enzymes that can
Literature review
44
oxidise 4-chlorophenol (Spokes & Walker 1974). Likewise, the mineralisation of
chrysene and benzopyrene can be achieved by bacteria grown on phenanathrene (Aitken
et al. 1998).
2.24.2. Enzyme induction by structurally unrelated compounds
In some cases, structurally unrelated compounds may cause the production of
enzymes that lead to biodegradation. For example, Pseudomaonas butanovora generally
degrades butane. It can also partially degrade chloroform because of the production of
monooxygenase from it (Hamamura et al. 1997). Further, Burkholderia cepacia
produces toluene-2-monooxygenase, which can attack toluene and has also been shown
to degrade a number of ethers (Hur et al. 1997). In this case, biodegradation results from
the induction of enzymes that are not specific for their substrates.
2.24.3. The role of readily degradable compounds
Simple compounds, like glucose (a monosaccharide) and arginine (an amino
acid), encourage the degradation of complex structures. This phenomenon is as yet
unexplained. The biodegradation of fluorobenzoates by mixed bacterial cultures has
been found to be enhanced by the addition of glucose (Horvath & Flathman 1976). This
might be due to the increased cell density of the bacteria in the presence of glucose.
Here, glucose causes passive degradation of fluorobenzoates by acting as a growth
factor for bacterial cells. Moreover, the biodegradation of pentachlorophenol was
increased by adding readily degraded substrates like glucose, succinate and glutamate to
the media for the growth of Flavobacterium species (Topp et al. 1988).
In contrast, the biodegradation of recalcitrant compounds was decreased by the
addition of glucose. For example, adding glucose to the growth medium repressed the
biodegradation of 2,4,6-trichlorophenol by Pseudomonas pickettii (Kiyohara et al.
1992). Likewise, the biodegradation of phenol in lake water was repressed after the
addition of glucose (Rubin & Alexander 1983). The reasons for this suppression have
not been detailed. This study attempts to understand the effect of co-metabolism on
LDPE biodegradation by Fusarium oxysporum. Monosaccharides, disaccharides and
polysaccharides were screened for their impact on biodegradation, and the effect of
alcohols was also examined.
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45
2.25. Laccase
In this study, laccase secreted by Fusarium oxysporum was shown to play an
important role in LDPE biodegradation. Laccase encourages LDPE oxidation, and thus
results in increased biodegradation by fungi. Here, the laccases from various sources
and their impact on biodegradation are outlined.
The laccases are copper-containing enzymes capable of oxidising a wide range
of substrates, including phenolic compounds, non-phenolic compounds, lignin and
environmental pollutants. These oxido-reductases can also oxidise molecular oxygen to
water (Lee et al. 2002) by an electron transfer mechanism (Sakurai 1992). The
molecular mass of laccase ranges between 50 and 130 kDa (Morozova et al. 2007).
More than 100 forms of laccase have been purified and several have been
characterised (Kunamneni et al. 2008). In general, laccase holoenzymes are dimers or
tetramers, and are covalently linked with carbohydrate moieties. They contain four
copper ions in three different ionic states, with these ions playing an important role in
the oxidation of substrates.
Laccases, particularly those from Basidiomycetes, were identified as having
depolymerising capacity for lignin. Lignin contains phenyl propanoid units linked by
C–C and C–O bonds. Laccases catalyse electron transfer between these phenolic
propanoid groups and molecular oxygen. These enzymes can also degrade plastic
wastes with olefin units (Xu 2000). In conjunction with mediators of electron transfer,
laccase can oxidise biphenol and alkyl phenol derivatives. They can also degrade
organic pollutants (Dehghanifard et al. 2013) and recalcitrant pollutants (Shraddha et al.
2011). Laccases have been reported to oxidise alkenes (Niku-Paavola & Viikari 2000),
carbazole and flourene (Bressler et al. 2000).
The production of laccase is influenced by various factors. For example, the
addition of copper in minute quantities increases its production by 50 % compared to a
control (Li et al. 2011). Other potential inducers of laccase production from fungi
include the addition of ethanol (Lee et al. 1999) or xenobiotic compounds (Vahala &
Lantto 2001) to the cultivation media. Along with these, the addition of aromatic
compounds, such as lignin, veratryl alcohol and xylidine (Bollag & Leonowicz 1984),
Literature review
46
as well as the ratio of carbon to nitrogen in the growth medium, also induces laccase
production (Vahala & Lantto 2001).
These properties of laccase suggest that it may be able to oxidise LDPE. Thus, it
was of interest to examine the effect of laccase from Fusarium oxysporum on LDPE
oxidation, and as well as the factors influencing this oxidation. For example, alcohol
and sucrose were found to increase LDPE biodegradation. Therefore, the influence of
these factors on the oxidation of LDPE by laccase from Fusarium oxysporum was
examined in this work.
.
CHAPTER 3
MATERIALS AND METHODS
Materials and methods
48
3.1. Overview
This chapter provides an overview of the methodology that was used to assess
the impact of fungi on LDPE. Experiments were designed to characterize fungi and
LDPE, as well as the interactions that occurred between them.
3.2. Introduction
LDPE was incubated with fungi, and changes in its physical and chemical
properties were observed by these methods. FT-IR and Raman spectroscopy were used
to follow changes in the superficial functional groups participating in biodegradation.
XRD was used to determine the crystallinity of the LDPE before and after
biodegradation. AFM and SEM were used to determine the effects of biodegradation on
the surface structure and morphology of LDPE.
Weight loss was a preliminary test for biodegradation. Dissolved carbon dioxide
content was also considered. Most experiments were conducted in triplicate to obtain
reliable data. LDPE was characterised in four different forms: untreated LDPE, oxidised
LDPE, control LDPE and test LDPE. The untreated form was neither oxidised nor
treated using any chemical method. Oxidised LDPE was subjected to both chemical and
photo-oxidations. Control LDPE (or oxidised control LDPE) was used to measure the
effects of the cultivating media. It was processed in the same ways as the test LDPE,
such as through disinfection, weight loss and air-drying. The test LDPE was subjected
to fungal treatment.
Table 3-1: Types of LDPE pellets and sheets used in this study
Sample Features
Untreated LDPE (u LDPE) Used as it was received
Oxidised LDPE (o LDPE) Oxidised by both UV and chemical treatment
Control LDPE or oxidised control
LDPE (c LDPE)
Used to test the impact of Czapek’s-Dox
medium on oxidised LDPE
Test LDPE or fungal-treated LDPE
(t LDPE)
Oxidised LDPE treated with Fusarium
oxysporum or with Mucor #1
Materials and methods
49
The fungus Fusarium oxysporum, classified based on microscopic and
macroscopic methods, was used in the testing and observation of LDPE biodegradation.
Untreated LDPE was tested as disks, powders or pellets. Fungal DNA was extracted and
the classification confirmed by sequencing its 18S rDNA fragment. This strain was used
in almost all experiments, with the exception of testing the sheets of LDPE with
Irganox®. For these tests, Mucor # 1 fungus was used. In this case the effects of
biodegradation were determined using scanning electron microscopy and weight loss of
the LDPE. Mucor # 1 was isolated from land fill. Mucor # 2 was isolated form leachate
and Mucor # 3 was isolated from river sources (see Section 3.6.4 to 3.6.6).
3.3. Chemicals and reagents
All chemicals used were of analytical grade and were purchased from Sigma-
Aldrich, unless otherwise specified. Fungal growth media components were purchased
from Oxoid and Difco. Sephadex G-25 with bead sizes in the range 50 to 150 µm and
Sephadex G-75 with bead sizes of 20–50 µm, used for gel filtration, were also
purchased from Sigma-Aldrich.
3.3.1. LDPE pellets
Spherical LDPE pellets of approximately 0.3 g each (ERMEC 590, BCR
certified material) were purchased from Sigma-Aldrich. No colouring agents, stabilisers,
plasticisers or any other additives were added to the pellets.
3.3.1.1. LDPE in disc form
The LDPE pellets were compressed into wafers using a FT-IR disc-making
apparatus.
3.3.1.2. LDPE in powdered form
Small portions of the LDPE pellets were dissolved in xylene at 80 °C.
The powder obtained was then washed with ethanol and allowed to dry.
Materials and methods
50
3.3.1.3. LDPE with additives
Another set of LDPE test materials with the antioxidant Irganox® added was
obtained from common packing materials. They were cut off from this packaging
material using hand scissors.
3.3.2. HDPE
The semi-spherical and translucent form of HDPE was purchased from Sigma-
Aldrich. The average weight of these pellets was 0.3 g. No colouring agents, stabilisers,
plasticisers or other additives were added to these pellets.
3.4. Preparation of LDPE for biodegradation
3.4.1. Photo-oxidation
Photo-oxidation of PE samples was performed by incubation in a UV chamber
for 250 h at a wavelength of 250 nm or 478.51 kJ/mol.
3.4.2. Chemical oxidation
PE samples were washed and then chemically oxidised by incubating them in
HNO3 (99 %) and heating for 3 d at 80 °C.
3.4.3. Abiotic oxidation
LDPE samples, with heat stabilisers, were subjected to abiotic oxidation by
incubating them at 100 °C in a hot air oven for more than 60 days.
3.5. Disinfection of LDPE
After oxidation, all samples were disinfected by immersion in ethanol (70 %)
with constant shaking for 15 min. They were then dried in a laminar airflow hood and
stored in sterile bottles.
3.6. Isolation of fungi
Six species of fungi were isolated from soil collected at a municipal landfill site
(Brooklyn, Victoria), the Yarra River (Kew, Victoria) and landfill leachate (Brooklyn,
Materials and methods
51
Victoria). The fungal isolation procedures are described below (see Sections
3.6.4–3.6.6). The isolated fungi were classified depending on their macroscopic and
microscopic features. Two of the isolated fungal strains were used in the biodegradation
studies with LDPE (without additives) and (LDPE with additives). These were
Fusarium oxysporum and Mucor, respectively.
3.6.1. Sterilisation
All equipment and media were sterilised by autoclaving at 121 °C (101 kPa)
for 15 minutes. Fungal sample inoculations into conical flasks were performed under
sterile conditions in a laminar airflow hood.
3.6.2. pH determination and adjustment
Czapek’s-Dox medium was adjusted to pH 7 using sterile HCl (1 M) or sterile
NaOH (1 M).
3.6.3. Measuring weight loss
LDPE pellets were weighed before and after fungal treatment using a Sartorius
balance. The accuracy of these measurements was ±0.02.
3.6.4. Isolation of fungal samples from landfill
Fungi were isolated from landfill soil using the direct pour plate technique
(Gupta et al. 2012). First, soil samples were collected from landfill at a depth of
1 m. Debris was removed from the samples, and they were then weighed.
Approximately 0.1 g of soil was mixed with Potato dextrose agar (PDA) and this was
then poured into Petri plates. The sample was treated with streptomycin (0.5 g/L) to
prevent bacterial growth. After the plates had solidified, they were incubated at 25 °C
for 5 days. The fungal colonies were then observed and subcultured onto PDA plates.
3.6.5. Isolation of fungal samples from river water
Fungal samples were isolated by the serial dilution method (Kango 2010).
Water samples were collected from the Yarra River in sterile universal bottles (50 mL).
The samples were diluted 1:10 by adding 1 mL of each water sample to 9 mL of
distilled water and shaking this in a rotary shaker for 30 minutes. These samples
Materials and methods
52
were then further diluted to 1:100 by adding 5 mL of the 1:10 sample to 95 mL of
distilled water.
From this sample, 100 µL was transferred onto PDA plates and spread evenly
using an L-shaped spreader. Plates were then incubated for 5 days at 25 °C.
3.6.6. Isolation of fungal samples from leachate
Leachate samples were collected from landfill (Brooklyn, Victoria) and stored at
4 °C. Before processing the leachate sample, solids were decanted. The clarified
leachate (10 mL) was transferred to sterilised conical flasks and processed as for the
river water samples.
In total, 60 different cultures (mycelia) were obtained on PDA. From these
cultures, mycelia mats were carefully removed and inoculated into modified Czapek’s-
Dox liquid media (NaNO3, 3.0 g/L; MgSO4.7 H2O, 0.5 g/L; FeSO4.7 H2O, 0.01 g/L;
K2HPO4, 1.0 g/L) and onto modified Czapek’s-Dox agar (the above medium with 20
g/L agar added) (Pitt & Hocking 2009).
3.6.7. Isolation of LDPE degrading fungi
Fungi that degrade LDPE were obtained by using LDPE as the only source of
carbon in the growth medium. The fungi that were obtained from the river water, soil
and leachate samples were incubated with oxidised LDPE pellets in modified Czapek’s-
Dox media. The pellets were first disinfected and weighed as described above (±0.02)
(see Section 3.5.3). Then they were added to culture flasks along with Tween-80 (1
%) (a surfactant) and streptomycin (0.5 g/L) (an antibiotic). Glycerol (0.01 L/L) was
also added to the culture flasks in order to establish the initial fungal culture. These
culture flasks were incubated at 30 °C with rotation at 160 rpm for an initial 6
days. The resultant mycelia were removed by filtration through Whatman’s No. 1 filter
paper and added to another batch of the medium without glycerol. This time, incubation
was continued for 45 days with medium replacement at regular intervals.
Six different conical flasks showed growth of mycelia. These fungi were
separated and enriched using PDA. The cultures were cultivated in large quantities and
preserved for further usage.
Materials and methods
53
3.7. Storage of fungi
The isolated fungi were stored on PDA slants under mineral oil and under water.
The procedure for these storage methods is described in the following sections.
3.7.1. Storage on PDA slants
PDA slants in McCartney bottles were inoculated with the fungal samples.
These were incubated at 30 °C for 5 days in the dark. After incubation, the bottles were
tightly wrapped in aluminium foil and stored at 4 °C.
3.7.2. Storage under mineral oil
PDA slants were prepared and inoculated with fungal strains at 30 °C for 5 days.
After incubation, sterilised paraffin oil was poured over the top of each slant, at a depth
of not more than 1 cm. The samples were stored at 4 °C.
3.7.3. Storage under sterilised water
Fungal samples were collected and stored under sterilised water (4 mL). They
were incubated at room temperature for more than 5 days before they were used.
3.7.4. Selection of fungi
Only six flasks showed mycelial growth on modified Czapek’s-Dox media.
These were then subcultured. For this, 10 mL of the fungal suspension was inoculated
into modified Czapek’s-Dox medium. After incubation for 45 days, the LDPE pellets
were removed and measured to determine any weight loss.
3.7.4.1. Determination of weight loss of oxidised LDPE
After 45 days of incubation, oxidised LDPE pellets were collected and washed
with sodium dodecyl sulphate (SDS) 2% v/v for 4 h. Then SDS was removed by
washing the pellets with double distilled water. The adsorbed moisture on the pellets
was removed by drying them in an oven at 60 °C overnight. The resultant oxidised
LDPE was weighed using a Sartorius balance with ± 0.02 mg accuracy.
Materials and methods
54
3.7.4.2. Growth rate of fungi
Mycelial growth rate was determined on modified Czapek’s-Dox agar
containing oxidised LDPE in powdered form. The data were recorded as an average
value of 4 days’ growth using the following formula (Roberts 2004):
{[G (d6) – G (d5)] + [G (d5) – G (d4)] + [G (d4) – G (d3)] + [G (d3) – G (d2)]} ÷ 4 (1)
Where G (dx) = Average growth rate of the mycelia in a day, in mm
For the calculation, if the mycelia overgrew and reached the edges of the Petri
plate before Day 6, then the Day 5 data was used.
3.7.4.3. Analysis of dissolved carbon dioxide in LDPE culture flasks
The dissolved carbon dioxide in the culture flasks was determined by titrating
carbonic acid against sodium hydroxide (Gopalan & Sugumar 2008). The reagents used
and the procedure followed are listed below.
3.7.4.4. Reagents
Standard sodium hydroxide solution (0.02 N)
This was prepared by dissolving sodium hydroxide (NaOH) in 0.8 g/L of
distilled water.
Standard potassium hydrogen phthalate (0.02 M)
This solution was prepared by dissolving potassium hydrogen phthalate
(C8H5KO4) in 4.085 g/L of distilled water.
Sodium thiosulphate solution
This solution was prepared by dissolving of sodium thiosulphate (Na2S2O3)
2.5 g in 100 mL of distilled water.
Indicators
Methyl orange and phenolphthalein indicators were used.
Materials and methods
55
3.7.4.5. Procedure
First, fungi were filtered from the culture media using Whatman’s No.1 filter
paper. The filtrate (100 mL) was then added to fresh beakers. To these, one drop of
sodium thiosulphate solution and two drops of methyl orange indicators were added.
This solution was titrated against the NaOH solution until a yellow colour was observed.
After this, two drops of phenolphthalein indicator was added to the filtrate, and the
titration was continued until a light pink colour was observed. The average volume of
NaOH consumed was noted. All six fungal cultures were tested in triplicate.
The formula for calculation of dissolved carbon dioxide is as follows:
(2)
WhereV1 = volume of water sample in mL, V2 = volume of NaOH in mL and N =
normality of the NaOH solution.
3.7.5. MATH test
Fungal cell surface hydrophobicity was measured essentially as described by
Smith et al. (Smith et al. 1998). In this procedure, hexadecane and PUM buffer
(potassium, urea and magnesium sulphate) were used as hydrophobic and hydrophilic
solvents respectively. PUM buffer was prepared by adding K2HPO4 (22.2 g), KH2PO4
(7.26 g), urea (1.8 g), and MgSO4.7H2O (0.2 g) to 1 L of distilled water and adjusting
the pH to 7.1. The LDPE pellets were incubated with isolated fungal strains in shaking
flasks of nutrient broth for 3 days (Holder et al. 2007). PUM buffer was then used to
wash the fungi that had attached to LDPE pellets. The suspension obtained (3 mL) was
taken and its optical density was adjusted to 0.4 at 470 nm. This suspension was
dispensed into acid-washed glass tubes, to which 300 µL of hexadecane was added. The
mixture was then vortexed for 30 seconds and allowed to stand at room temperature for
15 minutes. From this mixture, the upper hexadecane phase was carefully removed
without disturbing the aqueous solution below. For removing the remaining hexadecane
supernatant, the tubes were incubated at 5 °C for a few minutes and then kept at room
temperature for 15 minutes. The absorbance was calculated using a UV-Vis
spectrophotometer at 470 nm with cell-free buffer as a blank.
Materials and methods
56
3.7.6. Estimation of attached protein
The protein concentration in the biofilm that formed on the LDPE pellets was
measured using the Bradford method. LDPE pellets were collected after 45 days of
fungal incubation as described below (see Section 3.7.1). Biofilm proteins that were
attached to LDPE were denatured by boiling in 5 mL of NaOH (0.5 mol/L) at 100 °C
for 30 minutes. The denatured proteins were collected by centrifugation at 2,348 g for
10 minutes. The supernatant was removed and the protein pellet was subjected to the
same procedure again (Hadad et al. 2005). Both the supernatants were combined and
the protein concentration was estimated using the Bradford reagent method (Walker
2002).
3.7.6.1. Bradford reagent method
The Bradford reagent method was followed for protein estimation. The reagents
used and the procedure followed are listed below.
3.7.6.2. Reagents
Coomassie Brilliant Blue G-250, ethanol (95 % w/v), perchloric acid (3.5 %
w/w), and Bovine Serum Albumin (BSA) were used.
3.7.6.3. Preparation of dye
Coomassie Brilliant Blue G-250 (100 mg) was dissolved in 50 mL of ethanol.
To this, 100 mL of perchloric acid was added and the solution was diluted with 1 L of
distilled water. This reagent was filtered using Whatman’s No.1filter paper.
3.7.6.4. Preparation of standard protein BSA
The standard protein BSA was prepared by adding 0.01 g of BSA to100 mL of
distilled water.
3.7.6.5. Procedure
BSA standards were prepared in decreasing amounts (100 µg, 80 µg, 60 µg, 40
µg and 20 µg). Along with these standards, one blank test tube was prepared using only
100 µL of double distilled water. Protein obtained from the isolation process was added
Materials and methods
57
to a separate test tube. Coomassie Brilliant Blue dye (5 mL) was added to all test tubes,
which were then vortexed to mix the solutions. After a few minutes of incubation at
room temperature, the absorbance of each sample was measured at 595 nm. The
concentration of protein was determined by comparison to the standard curve.
3.8. Fungal culturing techniques
The techniques described below were used to cultivate and identify the fungi
used in the biodegradation studies.
3.8.1. Shaker flask cultures
This was the most frequently used fungal-LDPE culturing technique that was
used in this study. Conical flasks with 250 ml of modified Czapek’s-Dox liquid media
were incubated at 30 °C with fungi and shaken at 160 rpm for 45 days. The medium
was replenished at regular intervals by filtering out the mycelia using Whatman’s
No. 1 filter paper. After 45 days, the consumed media solution was filtered and
discarded. The LDPE along with the filtered mycelia was added to the new batch of
sterilised media.
3.8.2. PDA culture
In this technique, molten PDA was poured into sterile Petri plates and allowed to
solidify in a laminar airflow hood. Fungi were plated directly as mycelia or as agar
blocks and incubated in a humidified incubator at 30 °C.
3.8.3. PDA broth
PDA broth was used to cultivate fungi in large quantities for storage purpose.
This was prepared according to the manufacturer’s protocol (Zimbro 2009).
3.8.4. Subculturing
Subculturing was performed to enrich the isolated fungal cultures. For this
technique, small pieces of agar blocks from a PDA slant (1 cm×1 cm) were excised and
transferred to PDA plates. These plates were incubated at 30 °C and used as working
cultures.
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Subculturing was also performed in shaking flasks to yield large quantities of
fungi for storage. For this, 1 L of liquid medium was prepared in 2 L conical flasks
plugged with non-absorbent cotton wool. These were shaken slowly (at 50 rpm) at 30
°C for 10 days.
3.8.5. Isolation of Fusarium strains
Fusarium strains were isolated using malachite green as a selective agent. Soil
samples were processed as described in Section 3.5.4. These fungal isolates were then
further incubated in Czapek’s-Dox liquid medium supplemented with 2.5 mg/L of
malachite green. The contents of this medium are described in Section 3.5.6. In this
instance, sucrose (30 g/L) was added as a carbon source.
3.9. Fungus classification
Fungi isolated by the methods described above (see Section 3.6) were screened
for their efficiency. All the fungi were classified to the genus level using the slide
culture technique. From this, Fusarium oxysporum was identified as being capable of
degrading LDPE efficiently in terms of plastic weight loss. This fungus was further
classified to the strain level by sequencing its 18 S rDNA gene.
3.9.1. Slide cultures
This technique was used to observe isolated fungal strains microscopically.
First, the bottom of a Petri dish was covered with filter paper and wetted with distilled
water. Then a microscope slide was mounted on the paper with the help of a V-shaped
glass rod. Next, a 5 mm square slice of the PDA block was placed on the glass slide.
Using a sterilised loop, an inoculum of the isolated fungus was inserted into this block,
which was then covered with a glass cover slip. After that the Petri plate was covered
and incubated at 30 °C for 5 days.
After incubation, the set-up was taken out of the incubation chamber and
the agar block was discarded. The cover slip and glass slides were rinsed with 95
% ethanol. After the ethanol had evaporated, lactophenol blue was added. A new
glass slide was added to the cover slip, and the glass slide was covered with a new cover
slip. These preparations were then observed under a microscope (Patil & Muskan 2009).
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3.9.2. Extraction of fungal DNA
Fungal DNA was extracted using the DNeasy kit (Qiagen 2010) according to the
manufacturer’s protocol and amplified by the polymerase chain reaction (PCR) method
as follows.
3.9.3. PCR
Amplification of isolated DNA was performed using the PCR technique
(Sambrook & Russell 2001). For this, commercially available Mango mix (Bioline) was
used. This mixture contains Mango Taq DNA polymerase, ultra-pure dNTPs, red and
orange reference dyes and MgCl2 (see Table 3-2). The total 50 µL PCR mix was
prepared along with the primers.
Table 3-2: PCR mix
Component Quantity (µL)
5X Mango Taq reaction buffer 10
50 mM MgCl2 4
100 mM dNTP mix 1
DNA template ng 1
ITS1 and ITS4 of 0.1 µM 0.5
PCR grade water 33
3.9.3.1. Primers for PCR
The primers ITS1 (5’-TCCGTAGGTGAACCTGCGG-3’) and ITS4 (5’-
TCCTCCGCTTATTGATATGC-3’) were used to amplify the ITS region (see Figure 2-
7).
3.9.3.2. PCR cycle
The initial denaturation step was performed at 94 °C for 5 minutes, followed
by 30 amplification cycles each at 94 °C for 1 minute, 54 °C for 1 minute and 72 °C
for 1 minute. A final amplification was done at 72 °C for 10 minutes (Liu 2010).
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3.9.3.3. Electrophoresis of DNA
The amplified DNA was separated using agarose gel electrophoresis
(Sambrook & Russell 2001). Running buffer (TAE buffer) (5X concentration) was
prepared by dissolving 24.2 g of Tris (Tris hydroxymethyl aminomethane), 5.71 mL of
glacial acetic acid and 10 mL of 0.5 M EDTA in 800 mL of distilled water. The pH
was adjusted to 8.0 and the volume made up to 1 L with distilled water.
3.9.3.4. Preparation of TAE buffer
TAE buffer was diluted to working concentration (1X) and to this was added 0.5
g/mL ethidium bromide.
3.9.3.5. Preparation of loading solution
The loading solution was prepared by adding glycerol (10 %) and bromophenol
blue (0.025 %) to distilled water.
3.9.3.6. Agarose gel electrophoresis procedure
Agarose (2 %) was dissolved in TAE buffer and the mixture was heated in a
microwave until the agarose dissolved. Ethidium bromide was added to the molten
agarose solution to a final concentration of 0.5 g/mL. This was then poured into an
electrophoretic tray with an appropriate comb. The gel was allowed to cool until
solidified, at which point the comb was removed without disturbing the wells.
The gel was placed into a horizontal electrophoresis apparatus and covered with
TAE buffer. The amplified DNA, along with its loading solution (in a 1:5 ratio) was
then loaded into the wells. A 100 bp DNA ladder was used as a standard. DNA samples
were electrophoresed at 100 V for approximately 1 hour. The amplified DNA was
subsequently observed using a UV transilluminator.
3.9.3.7. Purification of DNA
In order to obtain clear classification of the isolated fungi, DNA was purified
from each strain. The DNA band that was obtained by electrophoresis was excised from
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an agarose gel using a sterile scalpel. Then gel slice was then placed into a 1.5 mL
microtube. The appropriate sized bands were excised and purified using the QIAquick
Gel Extraction Kit (QIAGEN). The purified DNA was then submitted to the
Australian Genome Research Facility (AGRF) for dideoxy nucleotide sequencing. The
results were analysed using the Bio-Edit program and the data were submitted to the
NCBI database. The identified strain was assigned the accession number JN711444.
3.10. Assessing fungal effects on LDPE
The effect of fungi on LDPE was assessed using the FT-IR, XRD, AFM and
SEM techniques. Along with these techniques, the methylene blue staining technique
was also employed to examine fungal effects on LDPE samples.
3.10.1. FT-IR
All LDPE samples were analysed by FT-IR to identify the functional groups that
participated in biodegradation. For this first, LDPE samples were frozen with liquid
nitrogen and pulverised using a mortar and pestle. Next, they were weighed and mixed
with potassium bromide (1 % w/w) and compressed to form thin wafers. Untreated
LDPE was compressed (without mixing with potassium bromide) to form a film. These
samples were analysed in the range of 4,000 cm-1 to 450 cm-1 using a Perkin-Elmer FT
2000 instrument. The results were normalised and analysed using spectrum for windows
software.
3.10.2. Raman spectroscopy
This technique was performed to identify functional groups that cannot be
clearly identified by FT-IR analysis. For this study, a Horiba Jobin Yvons Lab Ram
800 HR instrument was used with an excitation wavelength of 532 nm. This
spectrometer has a grating with 1,800 lines/mm and has a 200 μm confocal hole with a
50x lens. The spectral acquisition time was set for 2 seconds.
3.10.3. SEM
This technique was used to examine the effect of fungi on LDPE biodegradation
and also to assess the biofilm formed on the LDPE surface during the incubation with
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fungal samples. First, LDPE pellets were incubated with isolated fungi for 5 days on
PDA plates. Then the pellets were washed twice with distilled water to remove
any unattached fungi. Next, they were incubated overnight at 30 °C to remove
moisture. Then they were submitted to SEM to determine if a biofilm had been
established.
The other set of LDPE samples were analysed to determine the surface
modifications caused by fungal treatment. In this instance LDPE samples were
incubated with fungi as described above to facilitate biofilm formation. After the
incubation, the established biofilm was washed off with SDS. Then, LDPE samples
were sputter-coated with gold dust and then submitted to SEM. For this, a Zeiss Supra
40 VP field emission scanning electron microscope was used.
3.10.4. AFM
AFM scans were performed at room temperature using an Innova scanning
probe microscope (Veeco, Bruker, US). All LDPE samples were analysed using
pPhosphorus-doped silicon cantilevers (MPP-31120-10, Veeco, Bruker) in tapping
mode. The spring constant was set at 0.9 N/m, and a resonance frequency at ~20 kHz.
LDPE pellets were mounted on the stage with the help of removable gum. AFM scans
were performed perpendicular to the cantilever axis at the speed of 1 Hz. The resulting
topographical data were processed with first-order horizontal and vertical levelling
before performing a roughness analysis. In order to minimise the effect of noise on peak
counts, surfaces were smoothed using a convolution algorithm, and the minimum
threshold for definition of peaks was set at 10 % of the average roughness above the
mean height. Topographical analysis was performed using the software Nanoscope
(v1.10, Veeco). The roughness parameters included the average roughness (Sa) and root
mean square (rms).
3.10.5. XRD
LDPE samples were examined by XRD using a Bruker D8 instrument. All
samples were positioned onto the sample loading plate located in the XRD chamber.
Samples were arranged on the sample holder and subjected to x-rays and analysed at an
angle of 2θ while the value of this ranged from 5 ° to 85 °.
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3.10.6. Methylene blue test
This test was performed to quantify biodegradation by means of staining of
LDPE samples. The LDPE samples were added to methylene blue solution (1 % w/v)
and then they were boiled for 5 minutes in a water bath. Next, they were allowed to cool
for 5 minutes at room temperature. The optical density of the resultant solutions was
measured at 662 nm using a spectrometer.
Oxidised LDPE Biodegraded LDPE
Methylene blue solution
Resultant methylene blue
Solution
Figure 3-1: Schematic representation of methylene blue test
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3.11. Spore counting
Spore counting was performed to determine the amount of Fusarium oxysporum
used in some experiments. For this, Fusarium oxysporum was cultivated on acidified
PDA to induce spore formation. This medium was prepared by adding 3.9 g of acidified
potato dextrose broth (PDB) to 100 mL of distilled water along with 2 g of agar.
Fusarium oxysporum (Mycelia) were also added to the medium, along with 0.1 mL of
lactic acid (25 %) with constant stirring. Then it was incubated for one week. After this,
the PDA plates were flushed with 5 mL of distilled water and scraped from the plates.
This resulted in a mixed solution of spores and mycelia, which was subsequently
filtered through four layers of cheesecloth to separate out the mycelia. The resulting
filtrate was centrifuged at 3,000 rpm for 5 minutes, and then supernatant was discarded
(Bisen 2014). The pellet was then resuspended in distilled water (5 mL) and the number
of spores was determined using a haemocytometer.
The spore suspension (0.01 mL) was loaded into a haemocytometer and was
allowed to stand for 2 minutes. The number of spores on the four corners of the
haemocytometer counting area was then counted, ignoring those spores touching the top
and left borders (Bisen 2014). The formula used to determine the spore count was:
Spore count = average spore count per square of the four corner squares counted ×104
spores/mL
Using this formula, it was found that 1 mL of spore solution contains 106
spores. Spore suspension solution (4 mL) was cultivated in Czapek’s-dox broth (1 L)
at 30 °C for 7days. Then it was stored in an aluminium foil wrapped flasks for further
use.
3.12. Factors affecting biodegradation
Various properties of biodegradation with the isolated fungal strain, Fusarium
oxysporum, were investigated. Specifically, the effect of micronutrients, co-metabolites,
and oxidation and the rate of biodegradations were investigated. This was done by
inoculating 20 mL of Fusarium oxysporum mycelia that was obtained from spore
solution (see Section 3.11). In general, weight loss of LDPE was considered as a
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biodegradation parameter. The dissolved carbon dioxide content was used to
substantiate the results.
3.12.1. Effect of fungal micronutrients
The effects of Fusarium oxysporum micronutrients were screened for their
impact on biodegradation. These micronutrients were added in the form of chloride
salts; namely, copper chloride (CuCl2), manganese chloride (MnCl2), ferrous chloride
(FeCl2) and zinc chloride (ZnCl2). These salts were added to the Czapek’s-Dox medium
in increasing concentrations (0.25 mmol/L, 0.50 mmol/L and 0.75 mmol/L) and then
incubated with Fusarium oxysporum for 45 days at 30 °C. These experiments were
conducted in triplicate along with appropriate controls. After the first 7 days of
incubation, the dissolved carbon dioxide was measured (see Section 3.7.4.3). Following
this, the weight loss of the LDPE pellets was measured after 45 days of incubation.
3.12.2. Effect of co-metabolites
The biodegradation of oxidised LDPE was tested, alone and with other carbon
sources (co-metabolites) including monosaccharides, disaccharides and polysaccharides
at 0.25 %, 0.50 %, 0.75 % and 1 % (w/v). In addition, the effects of sugars and alcohols
(methanol, ethanol and propanol) were also tested at concentrations of 0.25 %, 0.50 %,
0.75 % and 1 % v/v. Each of these components was tested in triplicate with appropriate
controls.
3.12.3. Effect of oxidation on the biodegradation of LDPE
This experiment was conducted to determine the effect of extent of oxidation on
LDPE biodegradation. Photo-oxidation of LDPE pellets was performed in a UV
chamber at 260 nm for 24, 48 and 72 h. After each of these periods of incubation, the
LDPE pellets were removed from the chamber and submitted to 45 days of incubation
with Fusarium oxysporum as mentioned above (see Section 3.8.1). Their weight loss
was then measured to indicate the extent of biodegradation.
3.12.4. Rate and extent of biodegradation
This experiment was conducted to determine the rate of biodegradation of LDPE
samples. Oxidised LDPE pellets were weighed and subjected to biodegradation as
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described in the previous sections (see Section 3.8.1). In this process, oxidised LDPE
pellets were removed every week from the conical flasks with fungi. The microbial
biofilm was washed off with SDS and the sample was weighed and noted. Incubation
was continued for more than 90 days, and the weight loss over time was plotted to
calculate the rate and extent of biodegradation.
3.13. Biodegradation with cell-free extracts
Fungal cell-free extracts were prepared from mycelia of Fusarium oxysporum
(20 mL) that was obtained from spore solution (see Section 3.11). The mycelia were
removed by filtration through Whatman’s No.1 filter paper followed by 1.2 µm and
0.45 µm Millipore filters. The filtrate was then freeze-dried and stored at 4 °C. For
use in biodegradation experiments, the filtrate was dissolved in an antibiotic solution
[streptomycin (0.5 g/L)], with the pH adjusted to 7. Deionised water was added to make
up the final volume of solution to 100 mL. One of the filtrate was denatured by
autoclaving at 121 °C and served as a control, while the remainders were used in the
biodegradation experiments.
The weight of the pre-treated LDPE was measured as described in the previous
section and noted (see Section 3.6.3). The disinfected LDPE pellets were placed in 20
mL of the cell-free extract and agitated for 45 days at 30 °C.
3.13.1. Estimation of the protein quantity in fungal extracts
This was done using the Bradford method described earlier.
3.13.2. Estimation of the carbohydrate quantity in fungal extracts
The carbohydrate concentration in the fungal extracts was measured using the
Dubois method (Dubois et al. 1956). One mL of the fungal extract solution was mixed
with 1 mL of 5 % phenol and 5 mL of concentrated sulphuric acid. A solution of
glucose (0.1 mg/mL) along with phenol (1 mL) and sulphuric acid (5 mL) was used as a
control. These solutions were allowed to stand for 20 minutes and then optical density
measurements were taken at 490 nm.
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3.14. Assessment of oxidation of LDPE by FT-IR
This experiment was conducted to assess the effect of oxidation and whether it is
confined to the surface or if it penetrated the LDPE matrix. The LDPE pellets were cut
into two, and two small pieces were extracted, one from the centre and another from the
surface of the pellet. These were ground into a powder with KBr (1 % w/w) and were
compressed to make a thin wafer, which was then analysed by FT-IR. The carbonyl
index was also measured and noted using this formula:
COi = Optical density of absorption band at 1,640–1,840 cm -1: Optical density of
absorption band at 1,463 cm-1.
3.15. Comparative biodegradation of HDPE and LDPE
This experiment was conducted to determine the impact of density on LDPE
biodegradation. HDPE and LDPE pellets were photo-oxidised for 10 days, after which
the samples were weighed and disinfected. Both were incubated with Fusarium
oxysporum at 30 °C for 45 days. Biodegradation took place for 45 days, with regular
replenishment of the culture broth. After 45 days, the extent of biodegradation was
assessed by measuring the weight loss of the pellets.
3.16. Biodegradation of LDPE with additives
Commercial non-coloured LDPE sheets were purchased from a local
supermarket. These sheets contained the antioxidant Irganox®. The sheets were
oxidised by heat treatment in a hot air oven for 30 days, disinfected and added to
modified Czapek’s-Dox broth and agar. The samples were then inoculated with various
fungi isolated from river, landfill and leachate sources using the processes outlined in
Section 3.6. Biodegradation was observed by measuring weight loss of the sheets as
described in above section. The biofilms were washed off from these sheets using SDS
(See Section 3.7.4.1). The sheets were then observed under a scanning electron
microscope.
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3.17. Transmembrane inserts
These experiments were conducted to establish the necessity of biofilm
formation by Fusarium oxysporum during biodegradation. In order to physically
separate fungi from oxidised LDPE pellets, polyester transmembrane inserts (Corning)
with a pore size of 0.4 µm were used.
3.17.1. Transmembrane experiments
Transmembrane permeable supports were used to cultivate Fusarium
oxysporum. These permeable supports can be inserted into transmembrane plates. Each
plate contained six wells.
Each well of a transmembrane plate was filled with 1.5 mL of Czapek’s-Dox
medium, and a pellet of oxidised LDPE was added as a carbon source. Next, the
transmembrane was inserted into the well along with another oxidised LDPE pellet, and
coated with 10 µL of fungal suspension. The plate was held closed with cellophane tape
and incubated at 30 °C for 45 days. The medium was replenished regularly, and
distilled water was kept in the incubator to minimise evaporation. After 45 days, the
weight loss of the LDPE and the growth rate of the fungi were observed and noted.
Each of the isolated fungi was tested individually, along with the appropriate controls.
The experimental set-up is illustrated in Figure 3-2.
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Figure 3-2: Transmembrane set-up. A) Basic set-up showing the placement of the semi-
permeable membrane inside the transmembrane well; B) Complete set-up showing the
placement of the medium, LDPE pellets and fungal suspension
3.17.2. Gel filtration chromatography
Gel filtration chromatography was performed to fractionate fungal metabolites
in the transmembrane experiments. Sephadex G-75 was used as a stationary phase for
the separation.
3.17.2.1. Preparation of filtration buffer (Tris-HCl buffer)
Tris-HCl buffer was used as a filtration buffer. This was prepared by
dissolving 121.1 g of Tris in 800 mL of distilled water. The pH was slowly
adjusted to 7.5 by adding 40 mL HCl to the Tris solution. To this, blue dextran was
added as an indicator.
3.17.2.2. Procedure
In a fresh beaker, 400 mL of Tris-HCl buffer was added to 300 ml of the
Sephadex G-75 powder. This mixture was allowed to swell for 3 h. Any fine
particles that did not mix with the gel bed were decanted. Next, a column was clamped
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vertically and cotton wool was placed at the bottom. An initial 5 cm of buffer was
poured into the column, followed by the gel solution. This was allowed to settle for 1
hour. The buffer was then poured into the column and washed twice. Extra buffer on top
of the column bed was carefully removed, leaving only 0.5 cm of buffer above the gel in
the column. As soon as the blue dextran entered the last portion of the column, the tap
was opened and samples were collected at 3 mL/minute. The optical density of the
samples was measured at 254 nm using a UV spectrophotometer and the absorbance
was plotted.
3.17.3. SDS-PAGE
The fractions from gel filtration were centrifuged at 10,000 rpm or 9,391 g for
10 minutes and the supernatant was discarded. The pellet was mixed with sample buffer
(see below) and subjected to SDS-PAGE according to the Mini-PROTEAN® Tetra Cell
Instruction Manual (Bio-Rad 2013).
3.17.3.1. Preparation of sample buffer
Sample buffer (10 mL) at a 3X concentration was prepared by adding 2.4 mL of
Tris-HCl (1 M) at pH 6.8, 3 mL of 20 % SDS (prepared by dissolving 20 g of SDS in
100 mL of distilled water), 3 mL glycerol, 1.6 mL of β-mercaptoethanol and 0.006 g of
bromophenol blue.
3.17.3.2. Preparation of running buffer
A 10X running buffer was prepared by mixing Tris base (30.3 g) and glycine
(144 g) in distilled water at pH 8.3. The buffer was made up to 1 litre and to this was
added 10 g of SDS. The buffer was diluted to working concentration (1X) prior to use.
3.17.3.3. Gel fixing solution
A gel fixing solution was prepared by mixing methanol, acetic acid and distilled
water in a ratio of 50:10:40.
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3.17.3.4. Destaining solution
A destaining solution was prepared by mixing methanol, acetic acid and distilled
water in a ratio of 45:10:45.
3.17.3.5. Coomassie Brilliant Blue stain
The stock solution was prepared by mixing 12 g of Coomassie Brilliant Blue in
300 mL of methanol. The solution was acidified by adding 60 mL of acetic acid. The
working solution was prepared by adding 30 mL of concentrated stock to 500 mL
methanol and 400 mL of distilled water. This was again acidified by adding 100 mL of
acetic acid.
3.17.3.6. Preparation of polyacrylamide gels
Acrylamide-bisacrylamide solution was prepared by dissolving 30 %
acrylamide and 0.8 % bisacrylamide in distilled water and storing in the dark at 4 °C.
An ammonium persulphate (APS) solution (25 mg/100 mL) was made up in distilled
water.
The lower resolving gel (10 %) was prepared by adding 2 mL of acrylamide-
bisacrylamide, 3 mL of Tris-Cl (pH 8.8) and 38 µL of 20 % SDS to 2.43 mL of
distilled water. To initiate polymerisation, 36 µL of 10 % APS and 5 µL of TEMED
(Tetra Methyl Ethylene Diamine) were added to the solution. The gel was cast between
glass plates and then carefully washed with ethanol.
The upper stacking gel (4 %) was made by adding 660 µL of acrylamide-
bisacrylamide, 630 µL of Tris-Cl (pH 6.8) and 25 µL of 20 % SDS to 3.6 µL of distilled
water. Prior to casting, 25 µL of 10 % APS and 5 µL of TEMED were added and an
appropriate comb was inserted. After the stacking gel had polymerised, the comb was
carefully removed. The 3X sample buffer was diluted by mixing 25 µL of buffer with
75 µL of distilled water. Broad-range non-stained proteins (Bio-Rad) were used as
markers.
Electrophoresis was conducted for approximately 1 hour at 200 V. The gel was
removed from the plates and incubated in the gel fixing solution for about 30 minutes.
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Following this, the gel was added to the staining solution and incubated for a further 30
minutes. Destaining was performed by incubating the gel in the destaining solution for
3 h with shaking. Finally, the gel was carefully removed from the solution and
photographed.
3.18. Effect of metal salts on LDPE biodegradation by cell-free extracts
Fungal micronutrients (CuCl2, FeCl2, MnCl2 and ZnCl2) were assessed for their
effects on the biodegradation of oxidised LDPE. All salts were tested at 0.5 mmol/L,
with the pH of solutions adjusted to 7. These experiments were also conducted in
triplicate.
3.19. Detection of extracellular enzymes
The extracellular enzymes produced by the Fusarium oxysporum isolate were
identified using the plate assay technique and a titration method. The NCBI database
suggests that Fusarium oxysporum secretes a number of extracellular enzymes,
including amylase, cellulase, chitinase, and laccase. Screening was conducted for each
of these.
3.19.1. Enzymatic screening using the plate assay technique
These experiments were conducted in order to identify enzymes secreted by
Fusarium oxysporum in the presence of LDPE. First, mycelia of Fusarium oxysporum
was collected from stored PDA slants and inoculated into Czapek’s-Dox liquid medium
using a sterilised loop. To this, pulverised oxidised LDPE powder (0.1 mg/100 mL) was
added. This mix was incubated for 5 days at 30 °C (Kahangi et al. 2012). After
incubation, the resultant mycelium was filtered out with Whatman’s No. 1 filter paper.
It was then point inoculated into the media described below and the resulting
cultures were incubated overnight at 30 °C.
3.19.1.1. Screening for amylase
This was performed by incubating Fusarium oxysporum with starch amended
medium. This medium was prepared by adding 1 g glucose, 0.1 g yeast extract, 0.5 g
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peptone, 20 g starch and 16 g agar to 1 litre of distilled water. The pH was adjusted to 6,
and the mixture poured into Petri dishes. When the plates had solidified, they were point
inoculated with Fusarium oxysporum and incubated for 24 h. After incubation, the
plates were flooded with a solution of 1 % I2 and 2 % KI. A clear zone surrounding the
colonies after incubation was considered as a positive reaction.
3.19.1.2. Screening for cellulase
This was done by incubating Fusarium oxysporum with sodium
carboxymethylcellulose. The medium was prepared by dissolving 0.1 g of yeast extract,
0.5 g of peptone and 16 g of agar in 1 litre of distilled water containing Na-
carboxymethylcellulose (0.5 %). After incubation with Fusarium oxysporum, the plates
were flooded with Congo red. They were then destained by adding 1M of NaCl (20 mL)
for 15 minutes. A clear zone surrounding the colonies was considered a positive
reaction.
3.19.1.3. Screening for chitinase
This was done by incubating Fusarium oxysporum in chitin-amended medium.
The medium contained 4 g of chitin, 0.7 g of K2HPO4, 0.3 g of KH2PO4, 0.5 g of
MgSO4.5H20, 0.01 g of FeSO4.7H20 and 20 g of agar in 1 litre of distilled water. After
inoculation and subsequent incubation with Fusarium oxysporum the plates were
examined. A clear zone surrounding the colonies was considered as a positive reaction
(Maria et al. 2005).
3.19.1.4. Screening for laccase
This was done by incubating Fusarium oxysporum with α-naphthol amended
media. Glucose yeast peptone media was supplemented with 0.005 % α-naphthol
and the pH was adjusted to 6. These plates were then point inoculated with
Fusarium oxysporum and incubated for 24 h. The appearance of a blue colour
around the colonies after 24 h of incubation was considered a positive reaction
(Maria et al. 2005).
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3.20. Biodegradation of LDPE with laccases from Fusarium oxysporum
Fusarium oxysporum was grown on glucose peptone media for more than 5
days. Mycelia were removed by vacuum filtration through Whatman’s No. 1 filter paper
and the resultant filtrate was concentrated by freeze drying. The proteins in the solution
were precipitated by adding solid ammonium sulphate to obtain 80 % saturation. The
solution was stirred gently for 1 hour at 4 °C and left overnight. The precipitate was
collected by centrifugation at 8,000 g for 1 hour at 4 °C (Patrick et al. 2009).
3.20.1. Gel filtration separation of laccase
For gel filtration, Sephadex G-25 was used as a stationary phase and 100 mmol
of sodium chloride was used as a mobile phase. Thirty fractions were collected at a
flow rate of 1 mL/minute and the absorbance of each was measured at 280 nm. High
absorption fractions were separated and concentrated using Amicon Ultra-2 mL
centrifugal filters according to the manufacturer’s protocol. The separated fractions
were stored in 1 mL of 5 mmol bis-Tris-HCl buffer at pH 6.5. Next, 0.1 mL aliquots
were taken, mixed with 1 mL of 0.1 M α-naphthol (dissolved in 96 % ethanol) and
incubated at 30 °C. The development of a blue colour was considered as a positive
reaction for the presence of the enzyme.
3.20.2. Spectrophotometric assay of laccase activity
The absorption of the reaction mixture was measured at 420 nm. One unit of
enzyme activity is defined as the amount of enzyme that can oxidise one µM of 2, 2'-
azino-bis (3-ethylbenzthiazoline-6-sulfonic) acid (ABTS) in 1 minute.
3.20.3. ABTS standard solution
The ABTS standard solution was prepared by dissolving 1 mmol of ABTS
(ε=3.6104 cm-1 mL) in Na2HPO4/citric acid buffer (0.1 M) at pH 3.0 and 30 °C.
Concentrated protein fraction (laccase) from the gel filtration was used to
determination the specific activity of the enzyme (Cavallazzi et al. 2004). The gel
filtration fractions (100 µL) were mixed with 900 µL of ABTS standard solution and
allowed to stand for 5 minutes. The concentration of the laccase was calculated as 84
U/mL.
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3.20.4. Biodegradation with laccase
Laccase was purified from fungal extracts by gel filtration. To the extract, 100
mmol phosphate buffer was added to make up the total volume to 10 mL (Patrick et
al.2009). LDPE pellets were immersed in this buffer at a pH of 7 at a temperature of 30
°C. LDPE pellets were treated with SDS 2 % (v/v) overnight to remove attached
protein. Pellets were then washed with distilled water in order to remove SDS. The
washed, treated pellets were then weighed. Along with weight loss, dissolved carbon
dioxide concentrations were also calculated.
3.20.5. Effect of laccase on LDPE oxidation
LDPE pellets were photo-oxidised by incubating under UV light for 24 h. The
photo-oxidised pellets were then incubated with laccase in phosphate buffer (at pH 7
and at 30 °C). This incubation was continued for 2, 4, 8 and 12 h. The incubated
samples were then subjected to FT-IR analysis. The effect of oxidation was measured in
terms of the carbonyl index A1,712/A1,465.
3.21. Effect of co-metabolite additives on laccase oxidation capability
3.21.1. Effect of sucrose
Photo-oxidised LDPE pellets were incubated with laccase in phosphate buffer
(pH 7 at 30 °C) and incubated for 24 h. Increasing concentrations of sucrose at 0.25 %,
0.50 %, 0.75 % and 1 % (w/v) were added to these samples prior to incubation. After
incubation the pellets were analysed by FT-IR.
3.21.2. Effect of ethanol
Photo-oxidised LDPE pellets were incubated with laccase in phosphate buffer at
pH 7 and at 30 °C. Increasing concentrations of 0.25 %, 0.50 %, 0.75 % and 1 % v/v
ethanol was added to the samples and the reaction was continued for 24 h. The resultant
pellets were analysed by FT-IR.
Materials and methods
76
3.21.3. Effect of manganese, copper, ferrous and zinc chloride on laccase
oxidation
Photo-oxidised LDPE pellets were incubated with laccase in phosphate buffer at
pH 7 at 30 °C. The chloride salts MnCl2, CuCl2, FeCl2 and ZnCl2 were added to this
buffer at concentrations of 0.25 mmol/L, 0.50 mmol/L and 0.75 mmol/L, and the
samples were incubated for 24 h. After incubation, the samples were treated with SDS
as described above and subjected to FT-IR analysis.
CHAPTER 4
EXAMINING FUNGAL EFFECT ON LDPE
Examining fungal effect on LDPE
78
4.1. Overview
This chapter analyses the physical and chemical changes of LDPE surface
caused by fungi. A description of fungi employed for this process is provided. An
attempt to elucidate possible degradation mechanisms by fungi is also described.
4.2. Selection of fungi
Six different fungi were able to survive on oxidised LDPE as the only carbon
source. Based on microscope observations, they were classified as belonging to four
genera: Mucor, Fusarium, Aspergillus and Penicillium. Three of the observed fungi
belonged to the genus Mucor. Among these, fast-growing fungi were selected according
to the parameters of weight loss of LDPE and growth rate of the fungi. Weight losses
were measured by removing attached biofilms using SDS (see Section 3.7.4.1; see
Figure 4-1).
Figure 4-1: Weight loss of oxidised LDPE
0
2
4
6
8
10
12
14
16
Mucor # 1 Mucor # 2 Mucor # 3 Fusarium Aspergillus Penicillium
Ave
rage
wei
ght l
oss (
%)
Examining fungal effect on LDPE
79
Fusarium has shown to be ‘good performing’ fungus in the terms of weight loss
of LDPE. It was also observed growing at faster rates on modified Czapek’s-Dox plates
and in liquid media. Fusarium was shown to degrade not only LDPE, but also other
plastics (Umamaheswari & Murali 2013). Thus it was selected as a preferred fungus
among others for biodegradation of LDPE.
4.2.1. Dissolved carbon dioxide content
The fungi growing on LDPE samples were checked for dissolved carbon dioxide
content, out of 100 mL of culture medium incubated with fungi for one week, using the
procedure outlined in Section 3.7.4.3. Figure 4-2 shows that Fusarium incubated conical
flasks exhibited the highest carbon dioxide content, followed by Mucor #1. The other
strains, Mucor #2, Mucor #3, Aspergillus and Penicillium did not differ markedly from
the control.
-0.5
0
0.5
1
1.5
2
2.5
Control Mucor # 1 Mucor # 2 Mucor # 3 Fusarium Aspergillus Penicillium
Aver
age
conc
entr
atio
n of
CO
2 in
g/L
Figure 4-2: Concentration of dissolved carbon dioxide in g/L
It was observed that biodegradation produces carbon dioxide in to the medium
(Mohan 2011), resulting in a decrease in the dissolved oxygen concentration. Fusarium
Examining fungal effect on LDPE
80
oxysporum has demonstrated the capability to grow under low O2 concentrations (Pitt &
Hocking 2009), making it an appropriate fungus to study LDPE biodegradation.
4.2.2. Growth rate of fungi
Fungal growth rates were observed on plates containing Czapek’s-Dox medium.
The detailed procedure was provided in Section 3.7.4.2. The observed growth rates are
demonstrated in Table 4-1.
Table 4-1: Radial growth rates of fungi in mm
Growth rate of fungi
[mm/day]
Average
growth
rate
Mucor #1 3.2 ±1.2
Mucor #2 2.8 ±1.2
Mucor #3 2.5 ±0.3
Fusarium 6.5±0.3
Aspergillus 2.5 ±0.3
Penicillium 2.7±0.4
Fungal growth was measured at 30 °C and pH 7. Though all fungal strains were
able to grow, Fusarium showed the fastest growth among the tested species. An
increase in the optical density by mycelia was also noted in the Czapek’s-Dox broth,
indicating its efficiency in LDPE biodegradation.
4.2.3. Microbial adhesion test results
Hydrophobicity values describe the efficiency of fungi in attaching to
hydrophobic surfaces like LDPE. The tests were performed on sheets of oxidised LDPE.
The results were calculated after one week of incubation, and are shown in Figure 4-3.
The hydrophobic index was calculated using Equation (4.1):
Examining fungal effect on LDPE
81
(4.1)
0
0.2
0.4
0.6
0.8
1
1.2
Mucor # 1 Mucor # 2 Mucor # 3 Aspergillus Fusarium Penicillium
Fung
al h
ydro
phob
ocity
in a
rbitr
ary
units
Figure 4-3: Fungal hydrophobicity
It is apparent in Figure 4-3 that Fusarium was the most efficient at attaching to
LDPE of the strains examined here, followed by Mucor #1. Fusarium exhibited 150 and
178 units’ higher hydrophobicity than Mucor #1 and #2, respectively. This indicates
that Fusarium can more strongly adhere to the LDPE pellets than the above mentioned
strains.
Most fungi generally exude small amphiphillic proteins called hydrophobins.
These proteins can render hydrophilic nature to hydrophobic structures like LDPE and
vice versa (Ritz 2011). Depending on its nutrient availability, Fusarium is capable of
producing these hydrophobins (Fuchs et al. 2004). During the above experiment, as the
LDPE was used as a sole carbon source, Fusarium might have secreted hydrophobins,
increasing their surface hydrophobicity.
Examining fungal effect on LDPE
82
4.2.4. Protein concentration of biofilm
Figure 4-4 shows the protein concentration of biofilm. The quantity of protein in
Fusarium biofilm was slightly higher than in other types of fungi. Fusarium biofilm had
a 1 µg/L and 3 µg/L higher protein content than Mucor #1 and #2 biofilm protein
contents, respectively.
0
5
10
15
20
25
30
Mucor # 1 Mucor # 2 Mucor # 3 Aspergillus Fusarium Penicillium
Con
cent
ratio
n of
pro
tien
(µg/
L)
Figure 4-4: Concentration of protein in biofilm
Fungal biofilm content depends on various environmental factors, including a
limited availability of nitrogen, high levels of oxygen, low temperature and low pH,
nutrient deprivation and desiccation (Ahimou et al. 2007). As in the above experiments,
carbon source for fungi is limited so fungi have to produce a high quantity of biofilms
to degrade LDPE as an environmental stress response. High protein content of
Fusarium oxysporum in its biofilms indicates that it is actively secreting LDPE
degrading enzymes or hydrophobins, which are proteins in nature.
Examining fungal effect on LDPE
83
4.2.5. Weight loss of LDPE
The Fusarium strain was isolated and tested to verify its ability to cause weight
loss of oxidised LDPE (see Section 3.7.4.1). The results are presented in Table 4-2. The
weight loss (%) was not consistent in the flasks, but was observed to average 17 % (± 3)
after 45 days’ incubation.
Table 4-2: Weight losses (in mg) of oxidised LDPE measured after biodegradation
Sample Weight loss [%]
Untreated 0
1 17.09
2 17.51
3 20.17
4 14.09
4.3. Classification of fungi
Microscopic observations of fungi were undertaken using the slide culture
technique outlined in Section 3.9.1. In micrographs of (a) and (b), mycelia of Fusarium
can be observed. A basic structural examination revealed the presence of sickle shaped
micro-conidia (see Figure 4-5 [c] and [d]), which is a characteristic of Fusarium
(Hospenthal & Rinaldi 2007). These fungal types were isolated from soil on oxidised
LDPE pellets.
Examining fungal effect on LDPE
84
4.3.1. Results of slide culture
a
b
c
d
Figure 4-5: Micrographs of fungi isolated from landfill: (a) mycelia, (b) groups on
conidia, (c) and (d) micro-conidia (40 x magnifications).
4.3.2. DNA sequencing
Fungal DNA was extracted and amplified, as described in Section 3.9.3. The
amplification process showed a 480-base pair DNA fragment (see Figure 4-6).
Examining fungal effect on LDPE
85
Figure 4-6: Agarose gel electrophoresis of amplified DNA fragment. Lane 4 shows a
pale band of fungal DNA. Lane 1 shows the DNA ladder.
The amplified DNA was sequenced by the Australian Genome Research Facility
(AGRF). The sequence was analysed using Bio-Edit® software. The sequence was
submitted to NCBI, and the accession number JN711444 obtained.
4.4. Characterisation of fungal-treated LDPE
To identify the impact of fungal activity on LDPE, several techniques were used,
including FT-IR spectroscopy, Raman spectroscopy, methylene blue test and XRD. It
was observed that oxidised LDPE and oxidised LDPE (control) exhibited similar
properties of crystallinity according to Raman spectroscopy. They also exhibited similar
FT-IR spectra (see Figure 4-7).
Examining fungal effect on LDPE
86
10
00
20
00
30
00
40
00
Ox
idis
ed
LD
PE
10
00
20
00
30
00
40
00
0.0
0.5
1.0
1.5
2.0
Ox
idis
ed
LD
PE
(c
on
tro
l)
10
00
20
00
30
00
40
00
Fu
ng
i tr
ea
ted
LD
PE
10
00
20
00
30
00
40
00
0.0
0.5
1.0
1.5
2.0
Un
tre
ate
d L
DP
EAbsorbance in arbitrary units
Wa
ve
nu
mb
er
(cm
-1)
Figu
re 4
-7: F
T-IR
spec
trum
of L
DPE
var
ietie
s. U
ntre
ated
LD
PE; O
xidi
sed
LDPE
; Oxi
dise
d co
ntro
l
LDPE
(con
trol
) and
fung
al-tr
eate
d LD
PE
Examining fungal effect on LDPE
87
4.4.1. FT-IR analysis
Untreated LDPE, oxidised LDPE, oxidised LDPE (control) and fungal-treated
LDPE were analysed by FT-IR, as described in Section 3.10.1. This technique was used
to identify the functional groups that participated during fungal degradation. Changes in
the crystallinity of the LDPE samples after these four treatments were observed,
according to the formula in Section 2.14.1.
It can be observed from the above spectra that oxidised LDPE contained various
functional groups not observed in untreated LPDE spectra, indicating the formation of
new functional groups. Most new functional groups were also seen in fungal-treated
LDPE, but with different peak intensities. This suggests that Fusarium oxysporum
caused degradation of these functional groups, leading to the biodegradation of LDPE.
It can also be observed that most peaks were present in both oxidised LDPE and
oxidised LDPE (control), but with slight differences in their respective absorption
intensities. Untreated LDPE exhibits broader peaks than those of other types of LDPE.
This may be due to differences in its mode of preparation. Untreated LDPE was
prepared by compressing it to a wafer by a disk-making mechanism, resulting in a
comparatively thicker sample giving broader peaks. Additionally, KBr was not added to
this sample.
Untreated LDPE did not exhibit any bands in the region of 1705–1740 cm-1,
while oxidised LDPE showed a signal at 1715 cm-1, indicating the presence of saturated
aliphatic ketone functional group (C=O) in the polymer chains (Rabek 1995). It is
indicating that oxidation of LDPE resulted in replacement of two hydrogen atoms with
oxygen attached to carbon molecules. The same functional group was observed in the
oxidised LDPE (control). Fungal-treated LDPE showed decreased absorbance for this
signal. This indicated that C=O was subjected to enzymatic activity by the isolated
Fusarium oxysporum. Along with these, the keto carbonyl, ester carbonyl, vinyl bond
and internal double bond indices were observed to have decreased intensities in fungal-
treated LDPE. Their functional group intensities were calculated using the formulae
cited earlier (Section 2.14.1). The results are presented below (see Figure 4-8).
Examining fungal effect on LDPE
88
-0.5
0
0.5
1
1.5
2
Untreated LDPE Oxidised LDPE Oxidised Control LDPE
Fungal-treated LDPE
Keto carbonyl bond index Ester carbonyl bond index
Vinyl bond index Internal double bond index
Figure 4-8: Differences between functional groups in LDPE subjected to various
treatments
Oxidation resulted in the appearance of various functional groups in the LDPE
structure. The functional groups formed can be represented in decreasing order of their
relative intensities, as follows:
RCOR1 > RCOOR1 > CH2=CH2
It was observed that the ester group is the easiest to degrade functional group by
Fusarium oxysporum, followed by the keto and vinyl groups. The biodegradation
pattern can be expressed as:
RCOOR1 > RCOR1 > CH2=CH2
The percentage of crystallinity was calculated from the FT-IR measurements
using Zerbi et al.'s (1989) formula as described in Section 2.14.1. Figure 4-9 shows the
values obtained for the four LDPE samples.
Examining fungal effect on LDPE
89
60
62
64
66
68
70
72
74
Untreated LDPE Oxidised LDPE Oxidised control LDPE
Fungal-treated LDPE
Perc
enta
ge o
f cry
stal
linity
Figure 4-9: Crystallinity of LDPE samples
It can be seen from these results that fungal-treated LDPE contained a greater
crystalline fraction than other LDPE types. It can be explained by the oxidation and
subsequent fungal treatment of LDPE samples. Oxidation results in chain termination of
LDPE, thereby increasing the amorphous fraction of the sample (Scheirs 2000). As
amorphous portions are more susceptible to biodegradation than crystalline portions
(Kroschwitz 1989), it can be expected that amorphous fraction was first consumed by
Fusarium oxysporum, resulting in the crystalline fraction as a residue.
4.4.2. Raman spectroscopy
Raman spectroscopy was performed to monitor changes in the functional groups
not observed by FT-IR. All three types of LDPE pellet were screened, with the results
showing clear differences between them (see Table 4-3 and Figure 4-10).
The untreated LDPE sample exhibited a peak at 1439 cm-1, indicating a semi-
crystalline nature of tested polymer (Lobo & Bonilla 2003). A band at 1296 cm-1 was
also observed in the untreated LDPE spectrum, indicating the presence of a twisting
vibration of –CH2. Further peaks at 1126 cm-1 and 1062 cm-1 indicated the presence of a
stretching vibration of –CH2. At these wave numbers, the untreated LDPE showed a
Examining fungal effect on LDPE
90
higher intensity. Additionally, a twisting vibration was observed at 1295 cm-1. Oxidised
and fungal-treated LDPE exhibited a decrease in bands’ sharpness, corresponding to a
decrease in the densities of their –CH2 groups. The crystalline peaks at 1463 cm-1, 1441
cm-1 and 1418 cm-1 changed to a broad peak at 1440 cm-1 in the oxidised LDPE.
1439 cm-1
1296 cm -1
1062 cm-1
0
50
100
150
200
250
100 350 600 850 1100 1350 1600 1850
Inte
nsity
of e
mis
sion
in a
rbitr
ary
units
Wave number in /cm
Untreated LDPE
Oxidised LDPE
Fungal treated LDPE
Figure 4-10: Raman spectra of LDPE
As shown in Figure 4-10, and following the method given in Section 2.14.2, the
intensity of emission of the fungal-treated LDPE spectra was shown to decrease from
811 to 895 cm-1. In contrast, the intensity of emission of oxidised LDPE was observed
to be relatively higher than those of the untreated and fungi-treated LDPE. This
indicated that short chains were created by oxidation, which were then degraded by the
fungi.
The intensity of emission at 890 cm-1 inversely indicates the approximate
molecular weight of LDPE (Lobo & Bonilla 2003) (see Table 4-3). Thus, depending on
the emission intensity, the oxidation process may decrease the molecular weight of
LDPE. Fungal-treated oxidised LDPE shows an increase in molecular weight.
Examining fungal effect on LDPE
91
Table 4-3: Intensities of emission by LDPE at 891 cm-1
Untreated
LDPE
Oxidised LDPE Fungal-treated LDPE
61.3 94.1 81.3
The amorphous proportion of the different LDPE types was calculated using
strobes formula, as described in Section 2.14.2 and shown below, in Table 4-4.
Table 4-4: The αa portion of LDPE types, depending on relative intensity of emission
Untreated
LDPE
Oxidised LDPE Fungal-treated
LDPE
74.9 118.6 79.8
These results suggest that the oxidation process increases the amorphous content
of the LDPE pellets. The amorphous portions can be attacked by fungi, causing a
decrease in the amorphous nature of LDPE and a corresponding increase in the
crystalline fraction of the pellets. These results were in agreement with those from the
FT-IR spectroscopic analyses, presented above.
4.5. Surface visualisation analysis
AFM and SEM were used to visualise the fungal-treated LDPE after fungal
treatment. The processes are detailed in Section 2.15.
4.5.1. Scanning electron microscopy
The result is presented in Figure 4-11 (d). The fungal-treated samples, in
particular, lost their surface roughness. In contrast, the oxidised LDPE (see Figure 4-11
[b]) and oxidised control LDPE (see Figure 4-11 [c]) surfaces were rougher than the
control (see Figure 4-11 [a]).
Examining fungal effect on LDPE
92
a
b
c
d
Figure 4-11: Scanning electron micrographs. (a) untreated LDPE, (b) oxidised LDPE,
(c) oxidised control LDPE and (d) fungal-treated LDPE (at 1 µm resolution).
Figure 4-12, shows the biofilm formed by Fusarium oxysporum. Along with this
mycelia and conidia of Fusarium oxysporum can be observed on the LDPE surface. It
was noted that biofilm formation depends on factors like the surface spatial structure of
LDPE and its roughness (Sitarska 2009). As the surface of LDPE becomes rough
because of oxidation, it allowed the attachment of isolated fungal strain Fusarium
oxysporum. This adhesion is stabilised by the hydrophobic nature of its cell wall
surface, which allowed its growth on the LDPE surface. Thus, isolated fungi caused the
biodegradation of LDPE.
Examining fungal effect on LDPE
93
Figure 4-12: Scanning electron micrograph showing Fusarium mycelia and conidia
4.5.2. AFM
AFM scans of an LDPE surface degraded over 45 days were conducted under
the same conditions described in Section 3.10.4. All four types of LDPE (untreated
LDPE, oxidised LDPE, oxidised LDPE [control] and fungal-treated LDPE) were
analysed for their topography. The resultant roughness values were presented as average
route mean square (RMS) values in Figure 4-13. These values were obtained from 3
different regions on the same pellet and by calculating the average value of it.
Examining fungal effect on LDPE
94
0
20
40
60
80
100
120
140
160
180
200
Untreated LDPE Oxidised LDPE Oxidised LDPE (control)
Fungal-treated LDPE
RM
S ro
ughn
ess
(nm
) (av
erag
e va
lue)
Figure 4-13: Average surface roughness of various LDPE samples.
It is evident from the graph above that fungal-treated LDPE exhibited low
surface roughness compared to the other LDPE types. Oxidised LDPE (control) showed
the highest surface roughness. The surface roughness of oxidised LDPE increased after
45 days of incubation in the fungal cultivation medium, suggesting that Fusarium
oxysporum attacks the rough surfaces of LDPE and degrades it.
4.5.2.1. AFM images of LDPE
The surface roughness of all four LDPE sample types were analysed by AFM.
Samples were prepared and analysed, as described in Section 3.10.4.
4.5.2.2. Comparison of AFM results by LDPE type
As shown in Figure 4-14, the surface of untreated LDPE (a) was semi-rough,
while oxidation increased the surface roughness of oxidised LDPE (b). The addition of
Czapek’s-Dox medium and incubation for 45 days at 30 °C increased the surface
roughness of oxidised LDPE (control) (c). Copper of Czapek’s-Dox has been found to
form complexes with oxidised LDPE (Sack et al. 1985), enhancing its susceptibility to
biodegradation. Thus functional groups were created by oxidation and increased surface
Examining fungal effect on LDPE
95
roughness by incubation with media salts, which caused biodegradation by fungi (d). It
was already known that biodegradation is best obtained by increasing the available
surface area for microbial activity (Moore & Saunders 1998). This finding is in line
with the observations of Sudhakar et al. (2008) in seawater-incubated LDPE samples.
a
b
c
d
Figure 4-14: AFM image of four types of LDPE: untreated LDPE (a), oxidised LDPE
(b), oxidised LDPE (control) (c) and Fungal-treated LDPE (d).
From the above visual analysis, it can also be noted that biodegradation causes a
smoothing of LDPE surface. From the SEM and AFM results it was observed that
fungus was causing the erosion of peaks or protrusions, created by oxidation of LDPE.
This might be because the rougher surfaces of LDPE were hydrophilic in nature. These
hydrophilic rougher portions facilitate the attachment of fungus (or its extracellular
products) onto the LDPE. This results in the degradation of rougher surfaces first,
resulting in smoother surfaces.
Examining fungal effect on LDPE
96
4.6. Crystallinity measurements
Untreated, oxidised and fungal-treated LDPE samples were submitted to XRD
studies to estimate the changes in crystallinity occurring during the biodegradation
process.
4.6.1. X- ray diffraction
All three samples (untreated LDPE, oxidised LDPE and fungal-treated LDPE)
were scanned by X-ray diffraction (see Figures 4-15–4-17). Deconvolution of the
graphs, curve fitting and area under peaks were calculated using Origin software. The
percentage of crystallinity was measured by formula, provided in Section 2.16.1.
0
20
40
60
80
100
5 15 25 35 45 55 65 75
Inte
nsity
in a
rbitr
ary
units
2 theta
Untreated LDPE
Figure 4-15: XRD of untreated LDPE
Untreated LDPE showed characteristic peaks at 21.6 and 23.18. These peaks
derived from the crystalline portion of the LDPE sample. A further peak can be
observed at 7.8, indicating that this sample was semi-crystalline.
Examining fungal effect on LDPE
97
0
20
40
60
80
100
5 15 25 35 45 55 65 75
Inte
nsity
in a
rbitr
ary
units
2 theta
Oxidised LDPE
Figure 4-16: XRD of oxidised LDPE
Oxidised LDPE showed characteristic peaks at both 21.6 and 23.18. The
intensity of the 21.6 peak is the same as for pure LDPE. The peak at 23.18 was lower
than for untreated LDPE, indicating a reduction in crystallinity.
Examining fungal effect on LDPE
98
0
20
40
60
80
100
5 15 25 35 45 55 65 75
Inte
nsity
in a
rbitr
ary
units
2 theta
Fungal-treated LDPE
Figure 4-17: XRD of fungal-treated LDPE
Fungal-treated or biodegraded LDPE showed different peaks from oxidised
LDPE. The intensity of the peak at 23.18 was reduced to seven units, although the
amorphous halo was not significantly different. The decrease in the amorphous portion
was relatively higher than the decrease in the crystalline portion. In fact, the total
percentage of crystallinity was found to decrease from 27 % for untreated LDPE to 25%
for oxidised LDPE, yet this increased to 28 % for the fungal-treated LDPE. This
indicates that the amorphous portions had been created by oxidation and then
subsequently degraded by fungi. These results are in agreement with those observed by
FT-IR, Raman spectroscopy (see Section 4.5.1 and Section 4.5.2). As XRD deals with
the bulk of material (LDPE pellet), the changes in crystallinity were minimal.
4.6.2. Methylene blue test
The biodegradation of the LDPE pellets was measured by staining using
methylene blue. The optical density of the resultant solution was measured at 662 nm
(see Figure 4-21). The procedure was detailed in Section 3.10.7.
Examining fungal effect on LDPE
99
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
Untreated LDPE Oxidised LDPE Fungal-treated LDPE
Aver
age
OD
at 6
62 n
m
Figure 4-18: Optical density at 662 nm of various resultant solutions
Untreated LDPE does not have any charged groups on its surface, and thus does
not react with methylene blue. Thus, as expected, the absorption by the resultant
solution was higher than the resulting solutions obtained with oxidised and fungi-treated
samples. Oxidised LDPE pellets were shown to possess functional groups on their
surface (keto, ester and vinyl groups), which reacted with the methylene group of dye to
form complexes. This reaction led to a decrease in the absorption by methylene blue in
the resultant solution. Although fungal-treated LDPE possessed all of the above-
mentioned functional groups, they were present in lower concentrations, indicating that
the biodegradation reactions appear to target charged functional groups on the oxidised
polymer. Also, biodegradation process reduced the surface area and the quantity of
polar groups present on oxidised LDPE. Thus, the attachment of methylene blue to
fungal-treated LDPE was comparatively less than to the oxidised LDPE pellets. This
resulted in the high absorbance of the resultant solution. This technique is easy, and can
be performed in minimal time.
Examining fungal effect on LDPE
100
4.7. The visual properties of the LDPE samples
During this research, differences in the properties of untreated LDPE, oxidised
LDPE and fungal-treated LDPE were observed. Visually, oxidised LDPE was pale
yellow, while untreated LDPE was white. The colour of oxidised LDPE was uniform,
while biodegraded (fungal-treated) LDPE had white patches on its surface. Fungal-
treated LDPE appeared opaque when compared with non-degraded LDPE types.
4.8. Biodegradation of LDPE with Irganox®
4.8.1. Weight loss
The weight loss of LDPE sheets after incubation with fungi was observed and
tabulated (see Section 3.7.4.1). The weight loss (%) was not consistent as the available
surface area of LDPE sheets (see Section 3.3.1.3) was different. However, an overall
weight loss of 17 % (±3) was observed after 45 days of incubation at 30 °C (see Table
4-5). This weight loss was not continuous, and it appeared to be reaching its maximum.
Further biodegradation was not observed despite the media being replenished every ten
days.
Table 4-5: Weight loss of LDPE pellets (mg) with Irganox®
Before incubation After incubation Weight loss (%)
25.4 (control) 25.4 0
24.1 20 17.01
26.3 22.5 14.45
25.5 21.6 15.29
26.5 22.9 13.58
4.8.2. Amount of attached protein
LDPE with Irganox® was incubated with Mucor #1 for 45 days, to calculate
weight loss. After incubation, the attached protein concentration was determined as 12
µg/L by the Bradford method, described in Section 3.7.6.
Examining fungal effect on LDPE
101
4.8.3. SEM results
Degradation was visualised by SEM of the surface of LDPE samples. Untreated
LDPE was clear and showed no surface modifications (see Figure 4-22 [a]). Conversely,
thermally oxidised LDPE showed worm-like patterns across its surface (see Figure 4-22
[b]). The same patterns were observed on the surface of the oxidised LDPE (control)
(see Figure 4-22 [c]). The fungal-treated LDPE showed clear patterns of degradation,
where the worm-like patterns had been cleared from some parts of the surface (see
Figure 4-22 [d]). This observation shows that the rough surface was created by
oxidation, subsequently degraded by Mucor #1. This observation is similar to that of
LDPE that had not been treated with Irganox®.
Examining fungal effect on LDPE
102
a
b
c
d
Figure 4-19: Scanning electron micrographs. (a) Untreated LDPE, (b) thermally
oxidised LDPE, (c) Oxidised LDPE (control) and (d) fungal-treated LDPE (at 1µm
resolution)
Examining fungal effect on LDPE
103
4.8.4. Discussion and conclusion for biodegradation of LDPE with Irganox®
The overall weight loss observed was no greater than 17 %. Biodegradation of
LDPE samples was evidenced by the presence of biofilm formed by the fungi. LDPE
with Irganox® can be degraded with Mucor #1 after a long oxidation period by thermal
oxidation. Anti-oxidant effect can be pacified by prolonged periods of thermal oxidation
and subsequent autoclaving of LDPE. Other groups have successfully isolated bacteria
from autoclaved LDPE that were with Irganox® (Chatterjee et al. 2010).
Scanning electron micrographs displayed characteristic worm-like
biodegradation patterns. These structures were observed on the top of thermally treated
LDPE sheets. Similar patterns were observed by Otake et al. (2003) on the surface of
LDPE samples that had been buried in soil for 32 years. The authors described them as
the result of thermal oxidation and possible biodegradation by the bioactive soil. On the
other hand, this pattern appears to be crystals revealed by the thermal oxidation of
LDPE surface (Olley & Bassett 1982). It can be concluded that biodegradation after
oxidation of LDPE with Irganox® is possible.
4.9. Conclusion
Isolated fungal strain Fusarium oxysporum did appear capable of degrading
LDPE and utilising it as a carbon source. Degradation appeared to occur preferentially
at the higher, more exposed or rougher points on the LDPE surface. This mode of
rougher peak degradation resulted in measurably smoother surfaces. This degradation
resulted in both chemical and physical changes in LDPE.
4.9.1. Chemical aspects of degradation
Biodegradation of untreated LDPE was not observed, as it does not have any
chemically reactive groups on its surface. Its oxidation created functional groups (such
as COOH and -CH=CH) that react with fungal products to allow degradation and cause
weight loss of LDPE. The formation of reactive groups on oxidised LDPE also results
in rougher surfaces (peaks), which also encourage the biodegradation process.
Oxidation of LDPE created short chain fragments in its matrix. Oxidation
resulted in the formation of carboxylic acids, aldehydes and alcohols on LDPE matrix.
Examining fungal effect on LDPE
104
Thus, formed carboxylic acids can react with acetyl Co-A synthetase to form
complexes, and can enter the β-oxidation cycle directly (Albertsson et al. 1987).
Aldehydes and alcohols formed by oxidation enter terminal oxidation mechanism,
causing its biodegradation.
Figure 4-20 below explains the formation and subsequent degradation of
functional groups on LDPE during the biodegradation process, described above.
H20 and O2
-CH2-CH2-COOH- + CH2- C-CH2
o
-CH2-CH2-CH2-CH2-CH2-CH2-CH2-
Enters into the -oxidation cycle
Norrish type 1 and type 2 degradation mechanism
Figure 4-20: Effect of oxidation on LDPE
4.9.2. Physical aspects of biodegradation
Initially, LDPE was in a semi-crystalline mode, containing both amorphous and
crystalline phases. When oxidised it resulted in the formation of small molecular
fragments due to chain termination. Oxidation also generated free radicals, further
leading to auto-oxidation. This phenomenon caused a decrease in the crystalline fraction
of LDPE. However, this change occurs primarily on the surface of LDPE pellet.
Findings from previous studies regarding the percentage of crystallinity of
LDPE after biodegradation are inconsistent. For example, Sudhakar et al. (Sudhakar et
Examining fungal effect on LDPE
105
al. 2008) observed a decrease in the crystallinity of seawater-treated LDPE, while
Weiland et al. (1995) observed an increase in crystallinity (compared with the untreated
control) after biodegradation. This was attributed to the degradation of the amorphous
content, leaving the crystalline portion, and resulting in an LDPE sample possessing a
higher percentage of the crystalline fraction. In the present study, it was observed that
changes in the percentage of crystallinity depend primarily on the time of incubation
with the respective microbe.
Figure 4-24 depicts changes in the LDPE crystallinity caused by oxidation and
its subsequent biodegradation. Initially, untreated LDPE owing to the nature of semi
crystalline nature exhibit both amorphous and crystalline regions in its polymer matrix.
When it is undergoing the oxidation process, few crystalline regions were converted
into amorphous regions. As these portions were susceptible to fungi, they were
preferentially degraded by fungi resulting in LDPE weight loss. Thus, the resultant
LDPE has more crystalline portions its matrix, increasing its percentage of crystallinity.
Examining fungal effect on LDPE
106
Untreated LDPE
Oxidised LDPE
Fungal-treated LDPE
Figure 4-21: A schematic representation of changes in crystallinity of LDPE during
biodegradation process
CHAPTER 5
FACTORS AFFECTING BIODEGRADATION
Factors affecting biodegradation
108
5.1. Overview
In this chapter effect of physical and chemical factors in a view to accelerate
LDPE biodegradation is presented. Biodegradation with Fusarium oxysporum
mycelium and with its cell-free extracts are detailed in order to better understanding of
the process. Following this, essentiality of bio-film formation is also scrutinised.
5.2. Introduction
In Chapter 4, it has been shown that Fusarium oxysporum possesses the ability
to biodegrade LDPE. Although the fungus exhibited high growth rate and a
hydrophobic cell wall, the rate of biodegradation was shown to be slow. In order to
increase this rate (optimisation), it is essential that the factors controlling the reaction be
understood. Researchers have undertaken vigorous study into the biodegradation of oil
spills in the environment in order to optimise the process. Many factors have been
proven to influence microbial degradation of oil, including temperature, micronutrients,
oxygen and microbial community dynamics (Das & Chandran 2011). Temperature
plays an important role in the biodegradation of hydrocarbons, as it influences both the
chemistry of oil and physiology of microbial flora (Okoh 2006). Further, it has been
proven that microorganisms require micronutrients such as copper and ferrous
(Newman et al. 2006) and macronutrients such as nitrogen and phosphorus (Xu &
Obbard 2004; Tyagi et al. 2011) for the optimisation of biodegradation. Along with this,
studies have shown that hydrocarbon contamination alters the overall microbial
population and its dynamics at the contaminated site (Liang et al. 2011). In this chapter,
all of the above factors were scrutinised with the view of optimising the biodegradation
of LDPE.
In this section most of the experiments were conducted in two stages. In first
stage, LDPE was attempted to biodegraded with mycelia of Fusarium oxysporum. In the
next stage, LDPE was degraded with cell-free extracts of Fusarium oxysporum.
This was done in a view to separate factors that influence biodegradation by
encouraging Fusarium oxysporum growth (passive manner) with that of the factors
that encourage biodegradation mechanism of Fusarium oxysporum (active manner).
Factors affecting biodegradation
109
Preparation of cell- free extracts was detailed in Section 3.13. In most experiments
biodegradation was assessed in terms of weight loss and dissolved carbon dioxide
concentration.
5.3. Optimisation of biodegradation
LDPE biodegradation was performed with the use of micronutrients (copper,
ferrous, manganese and zinc) in their chloride form. Further, macronutrients
(phosphates and nitrates) were also used in sodium and potassium salt forms.
Biodegradation of LDPE was also performed at varying temperatures and pH in order
that the optimum conditions for LDPE biodegradation be ascertained.
5.3.1. Effect of micro nutrients (manganese, copper, iron and zinc ions)
Micronutrients are elements essential for fungal growth which are needed in
only micro quantities (Griffin 1996). Depending on the forma specialis, it was noticed
that Cu+2, Fe+2, Mn+2 and Zn+2 have been found to encourage or discourage Fusarium
oxysporum growth (Ahmed 2011, Sanjeev and Eswaran 2008). This experiment was
conducted to check the effect of these micro nutrients of Fusarium oxysporum on LDPE
biodegradation in two stages. At first biodegradation (weight loss) was observed with
Fusarium oxysporum mycelia followed by its cell-free extract. All micronutrients were
added in their chloride form. It was observed that, the ion chloride (Cl2-) can inhibit
Fusarium oxysporum (Lichtfouse & Navarrete 2009) at higher concentrations.
However, in the current work, as the low amounts of Cl2- were used their inhibitory
effect on Fusarium oxysporum is negligible.
Manganese, copper, ferrous and zinc chlorides were added to Czapek’s-Dox
media at increasing concentrations of 0.25 mmol/L, 0.50 mmol/L and 0.75 mmol/L.
The weight loss of LDPE samples kept in these media was measured after 45 days
(see Section 3.8.1). All experiments were performed in triplicate. A control was set up
without any nutrients. The resultant average weight loss of the control was observed as
14%. This experiment was conducted with mycelia and with cell-free extract of
Fusarium oxysporum.
Factors affecting biodegradation
110
0
2
4
6
8
10
12
14
16
18
Control 1 2 3 4
Aver
age
wei
ght
loss
(%)
0.25 mmol/L
0.50 mmol/L
0.75 mmol/L
A
0
2
4
6
8
10
12
14
16
18
Control 1 2 3 4
Aver
age
wei
ght l
oss
(%)
0.25 mmol/L0.50 mmol/L0.75 mmol/L
B
Figure 5-1: Effect of micro nutrients on LDPE biodegradation with mycelia (A) and
cell-free extract (B) of Fusarium oxysporum; 1) MnCl2, 2) CuCl2, 3) FeCl2, 4) ZnCl2.
Factors affecting biodegradation
111
Manganese in the form of Mn2+ had a significant influence on biodegradation in
both experiments i.e with mycelia and with cell-free extracts. In general Fusarium
oxysporum is inhibited by increasing concentrations of manganese chloride (Ahmed
2011). However, in this case weight loss with manganese chloride is relatively high,
especially when Fusarium oxysporum used in the mycelial form. This indicates that
Mn2+ activates the enzymes that participate in biodegradation mechanism by Fusarium
oxysporum (active manner).
Cupric chloride was also observed to have a positive effect on the
biodegradation of LDPE especially in the case of cell-free extracts. This implies that
enzymes that participate in biodegradation were activated by Cu2+. Along with this
copper possesses the capability of stabilising the oxidation of polyethylene by
preventing the formation of longer chains in the oxidised LDPE matrix (Vasile & Pascu
2005). This stabilization increases the biodegradation by Fusarium oxysporum.
During this research, it was observed that a slight increase in ferrous (Fe2+)
concentration increased the growth rate of Fusarium oxysporum. Indeed, Fe2+ can
severely impact the growth patterns of certain forma specialis of Fusarium oxysporum
(Woltz & Jones 1971). This implies that an increase in weight loss in the presence of
Fe2+ can be explained by an increase in Fusarium biomass. In the case of cell-free
biodegradation with Fe2+ as the weight loss is not high (compared with other nutrients).
So it can be concluded that Fe2+ encourages biodegradation only by encouraging growth
of Fusarium oxysporum (passive manner).
In contrast to the above ions Zn2+ was found not to encourage biodegradation of
LDPE either with mycelia or with cell-free extracts. This observation indicates that Zn2+
is not participating in biodegradation of LDPE. It was also found that most forma
specialis were inhibited by increasing concentrations of Zn2+ ions (Sanjeev & Eswaran
2008).
In conclusion it can be noticed that Mn2+ has a high impact on LDPE
biodegradation compared with Cu2+ and Fe2+. This indicates most of the ions with +2
valencies can encourage biodegradation by acting as co-enzymes. Co-enzymes are the
Factors affecting biodegradation
112
components of enzymes that stabilises the 3-D structure of enzymes (Bettelheim et al.
2009) and facilitates the substrate (LDPE)-enzyme attachment. On the other hand, the
above used metal ions with a +1 or +2 valency state can react with polymers and
promote thermo- oxidative degradation. This increases availability of susceptible
substrate for biodegradation by Fusarium oxysporum. The reaction can be described as:
ROOH + M+ M + + + RO* + OH-
OR
ROOH + M ++ M+ + ROO* + H+
Figure 5-2: Thermo-oxidation of LDPE by metal ions (Wright 2001)
5.3.2. Effect of temperature
These experiments were performed to figure out the optimum temperature for
biodegradation by Fusarium oxysporum. LDPE samples were incubated with Fusarium
oxysporum was observed at 20 °C, 25 °C, 30 °C and 35 °C as described above. The
experiment was conducted in triplicate and the average weight loss (%) was
determined (see Figure 5-3). This experiment was also done in two stages.
0
2
4
6
8
10
12
14
16
18
20
20 25 30 35
Aver
age
wei
ght
loss
(%)
Temperature ( C)
Weight loss with myceliumWeight loss with cell-free extract
Figure 5-3: Effect of temperature on weight loss
Factors affecting biodegradation
113
Fungal biodegradation increases with increasing temperature. Increased
temperatures increase the extent of oxidation on the LDPE surface, which results in the
generation of groups that are more susceptible to biodegradation by fungal enzymes. As
the reaction temperature increases enzymes coagulate and lost their specific 3-d
confirmation. This results in enzyme inactivation, leading to reduced weight loss. An
increased temperature also causes polymerisation of smaller chains and formation of
larger chains (Sudhakar et al. 2008). This leads to an increase in average molecular
weight. As the high molecular weight fragments are less susceptible for biodegradation
they result in the less biodegradation.
5.3.3. Effect of pH
These experiments were performed in order to figure out the optimum pH for
fungal degradation Fungi were incubated in Czapek’s-Dox media at pH values ranging
from 6 to 9. The experiment was conducted in triplicate and the average weight loss (%)
was determined. This experiment was also conducted in both stages.
As shown in Figure 5-4, alkaline conditions (pH 8) are most favourable for the
biodegradation of LDPE. This could be due to the release or titration of formed carbonic
acid in the alkaline conditions, the removal of which pushes the reaction towards the
right hand side of equilibrium. However, beyond a pH of 9 Fusarium was observed not
to grow, with weight loss being less than at pH 7 or 8. Further, this indicates that the
enzymes responsible for biodegradation act optimally at pH 8. They denatures at lesser
pH of 6 and higher pH of 8.
Factors affecting biodegradation
114
0
2
4
6
8
10
12
14
16
6 7 8 9
Aver
age
wei
ght
loss
(%)
pH
Weight loss with mycelium
Weight loss with cell-free extract
Figure 5-4: Effect of pH on biodegradation Fusarium oxysporum and its cell-free
extract
5.3.4. Effect of nitrates and phosphates
During the research, it was observed that increasing or decreasing the nitrate and
phosphate concentrations of Czapek’s-Dox media affected weight loss of LDPE. These
experiments were conducted to find the exact salt/ion of the nitrate and phosphate salts
that present in media. All salts (KNO3, NaNO3, KH2PO4 and NaH2PO4) were added at
1M concentration to potato dextrose broth (100 mL). This experiment was also
conducted with Fusarium oxysporum mycelium and its cell-free extracts.
Factors affecting biodegradation
115
0
2
4
6
8
10
12
14
16
18
20
1 2 3 4
Aver
age
wei
ght
loss
(%)
Weight loss with mycelia
Weight loss with cell-free extract
Figure 5-5: Effect of 1) KNO3, 2) NaNO3, 3) KH2PO4 and 4) NaH2PO4
The results shown in Figure 5-5 confirmed that nitrates were more effective than
phosphates in promoting the biodegradation of LDPE. Nitrates and phosphates
generally discourage growth of Fusarium oxysporum (Löffler et al. 1986; Woltz &
Engelhard 1973; Der 2013) while in contrast they encouraged biodegradation of LDPE.
So it can be concluded that nitrates and phosphates facilitate enzymatic attack on LDPE
biodegradation in active manner.
5.3.5. Oxidised LDPE treatment with other Fusarium isolates
These experiments were conducted to know whether or not other species of
Fusarium possess LDPE degradation capability. Fusarium species are known for their
ability to degrade cellulose and pectin. Several other groups have suggested that specific
strains of Fusarium can degrade oxidised LDPE (Zahra et al. 2010; Hasan et al. 2007).
It is thus essential to determine whether all Fusarium strains can degrade
oxidised LDPE.
Soil samples were collected and processed as described in Section 3.6.4. The
isolated fungi were incubated in Czapek’s-Dox liquid media supplemented with
Factors affecting biodegradation
116
malachite green (2.5 mg/L), as described in Section 3.8.5. These newly isolated fungi
were then added to oxidised LDPE. This test was performed in triplicate, using three
newly isolated Fusarium strains as well as the previously isolated Fusarium oxysporum
strain.
As shown in Figure 5-6, not all Fusarium isolates are effective for
biodegradation. Fusarium possess high number of species and strains that are both
pathogenic and non pathogenic in nature. This vast variation produces changes in their
reactions towards LDPE.
0
2
4
6
8
10
12
14
16
1 2 3 4
Aver
age
wei
ght
loss
(%)
Figure 5-6: Comparative biodegradation by Fusarium oxysporum (1) and other
Fusarium strains (2, 3, and 4)
5.4. Rate of weight loss
This experiment was conducted to check the biodegradation rate by Fusarium
oxysporum. It was done in two stages i.e with mycelia and with cell-free extracts of
Fusarium oxysporum. Biodegraded LDPE pellets were recovered after every 168 h (or
7 days) and treated with sodium dodecyl sulphate (SDS). The detail procedure was
given in Section 3.7.4.1. A steady rate of weight loss was observed until five weeks of
Factors affecting biodegradation
117
incubation, after which no further measurable change in weight was observed (see
Figure 5-7).
25
26
27
28
29
30
1 2 3 4 5 6 7
Wei
ght o
f oxi
dise
d L
DPE
in m
g
Period of incubation in weeks
Weight loss with mycelium
Weight loss with cell-free extract
Figure 5-7: Weight loss of LDPE by Fusarium oxysporum and its cell-free extract over
seven weeks
In both experiments a gradual and steady decrease in the weight of the LDPE
pellets was observed. It implies that biodegradation happens by surface erosion of pellet
(Alexander 1999). Rate of LDPE biodegradation depends on factors like its
composition or structure of back bone, degradation media, crystallinity, surface
morphology of LDPE and molecular weight of it (Jenkins & Stamboulis 2012). Bond
scission within the polymer chain depends on the total chain length as well as on the
position of the bond within the chain (Ziff & McGrady 1986). As the LDPE consists of
–CH2 in its polymer back bone, gradual removal of it leading to the uniformal weight
loss in a defined time frame. This indicating that weight loss of LDPE is majorly
because of removal of –CH2 by fungi.
It was also observed that continued incubation (more than 45 days) did not result
in a further decrease in the weight of the pellet, indicating that total weight loss reaches
Factors affecting biodegradation
118
a plateau. This may be because, after the removal of functional groups by the fungi, the
remaining components cannot interact well with the fungi or fungal products. It is
suggesting that oxidation is confined to the surface of LDPE pellets. This phenomenon
indicates that fungal treatment results in a decrease in the total surface energy of the
sample.
5.5. Biodegradation of HDPE and LDPE
This experiment was conducted to understand the effect of branching (tertiary
carbon atoms) of LDPE on its biodegradation. HDPE features a less branched structure
compared with LDPE. Therefore, comparative biodegradation of HDPE and LDPE
might provide an understanding of the impact of branching on LDPE biodegradation.
This experiment is also conducted in two stages.
HDPE and LDPE were incubated in a UV chamber and photo-oxidised for one
week before being subjected to fungal treatment for 45 days. The experiment was
conducted in triplicate and the average weight loss (%) was determined and both with
mycelium and with cell-free extracts.
Factors affecting biodegradation
119
0
2
4
6
8
10
12
14
16
18
Weight loss with mycelium Weight loss with cell-free extract
Aver
age
wei
ght
loss
(%)
LDPEHDPE
Figure 5-8: Average weight loss (%) following oxidation and fungal treatment for
LDPE and HDPE
Comparative biodegradation of LDPE and HDPE indicated that HDPE was more
prone to biodegradation. LDPE’s resistance to biodegradation may be explained by
its having more branches compared to HDPE, with the number of tertiary carbon atoms
in a hydrocarbon reducing its susceptibility to biodegradation. It was already proven
that branched hydrocarbons are more recalcitrant than their non-branched homologues.
HDPE has higher density and crystalline portions than LDPE (see Section 2.2.1). The
above both experiments were proven that crystallinity and densities are not rate
determining factors. It is also indicating that tertiary carbon atoms that are linked with
other carbon chains cannot participate in biodegradation. Enzymes that participate in
LDPE biodegradation cannot attach to these structures and hence cannot degrade LDPE.
It is interesting to note that both LDPE and HDPE can be degraded by the same fungal
strain, indicating that polymer branching has a greater effect on resistance than
crystallinity.
Factors affecting biodegradation
120
5.6. Effect of oxidation
In order to find out the effect of oxidation on biodegradation, samples of LDPE
were incubated in a UV light chamber for increasing amounts of time (7–10 days),
followed by fungal treatment for 45 days. The average weight loss for each UV
exposure time was determined. This experiment was also done in two stages.
The increase in weight loss over time shown in Figure 5-9 indicates that LDPE
biodegradation can be encouraged by high UV dosage. UV light absorption by oxidised
LDPE resulted in generation of more free radicals in the LDPE matrix. This caused
auto-oxidation of LDPE, resulting more oxidised units to react with fungal enzymes.
0
2
4
6
8
10
12
7 8 9 10
Aver
age
wei
ght
loss
(%)
Period of photo-oxidation indays
Weight loss with mycelium
Weight loss with cell-free extract
Figure 5-9: Effect of oxidation period on biodegradation in terms of weight loss (%).
5.7. Comparative degradation of LDPE by different oxidation methods
This experiment was performed to figure out what type of oxidation method
is apt for increased biodegradation of LDPE. Batches of LDPE pellets were incubated
in a UV light chamber for seven days, in a hot air oven at 100 °C for seven days
and in HNO3 and H2SO4 solutions and heated as described in Section 3.4.2. These
Factors affecting biodegradation
121
batches of LDPE pellets were then subjected to fungal treatment for 45 days after
which the average weight loss was determined. This experiment was also done in two
stages.
0
2
4
6
8
10
12
Thermal treatment Photo treatment Chemical treatment
Aver
age
wei
ght
loss
(%)
Weight loss with mycelium
Weight loss with cell-free extract
Figure 5-10: Effect of oxidation method on biodegradation of LDPE.
Among the three oxidation methods, the chemical oxidation method had the
greatest impact, attributable to the fact that the acids used (i.e., nitric and sulphuric acid)
could penetrate into the crystalline portions of the LDPE. Especially, nitric acid can
attack polymer molecules in the stressed crystalline portions (Peggs 1990). Thus,
chemical oxidation may shorten the polymer chains in the crystalline portions as well as
the amorphous portions. These fragments are then further degraded by fungi.
In the case of photo-oxidation, the generated free radicals might be confined to
the amorphous portions, which form a continuum around the crystalline portions and
can be easily oxidised. By contrast, the crystalline portions of polyethylene are resistant
to oxygen diffusion (Grassie & Scott 1988) and hence might not been oxidised
by photo-oxidation. Heat treatment might increase the crystallinity of LDPE by
increasing the attachment of short fragments of LDPE. It was observed that heat
Factors affecting biodegradation
122
treatment or thermal oxidation increases the crystallinity of LDPE (Sudhakar et al.
2008). Therefore, this might not be an effective oxidation method for biodegradation.
5.8. Importance of biofilm formation in the biodegradation of oxidised LDPE
5.8.1. Transmembrane experiments
This experiment was conducted to know the importance of biofilm formation
during biodegradation by Fusarium oxysporum. The detailed methodology was
described in Section 3.17.1.
LDPE weight loss was measured in a transmembrane plate containing fungi. The
weight loss of LDPE kept in Czapek’s-Dox media was higher (4.19% on average) than
the LDPE on the membrane (see Figure 5-11). This difference is explained by the
availability of oxygen to the polymer. The LDPE inside the transmembrane wells did
not accommodate biofilm and the dissolved oxygen in the media allowed for
biochemical reaction (biodegradation). In contrast, a biofilm was formed by fungi on
the LDPE on top of the transmembrane, preventing the diffusion of oxygen and
resulting in less biodegradation (as measured by weight loss). This implies that oxygen
is vital for biodegradation.
Factors affecting biodegradation
123
5.8.2. Difference in weight loss of LDPE samples
0
2
4
6
8
10
12
14
1 2 3 4 5
Aver
age
wei
ght
loss
(%)
Sample number
LDPE on semi-permeable membrane
LDPE in the transmembrane well
Figure 5-11: Transmembrane LDPE biodegradation
From the above experiments it can be understood that biofilm formation might
be one of the reasons for lower biodegradation rates of LDPE in nature. Similar results
were observed when starch was mixed with polyethylene (Pometto et al. 1993). In the
case of compost-based biodegradation, it was observed that LDPE that was exposed to
the air degraded faster than LDPE that was imbedded in the soil (Johnson et al. 1993).
This suggests that biodegradation is best achieved in conditions of high aeration,
and recommends against the use of soil-based biodegradation for biodegradation at an
industrial scale. In addition, it also suggests creating types of LDPE that inhibit the
formation of biofilms and thus potentially increase the rate of biodegradation. This can
be achievable by modifying additives like antioxidants and antistatic agents to inhibit
carbohydrate attachment to the polyethylene.
Factors affecting biodegradation
124
5.8.3. Growth of fungi in transmembrane wells
Fungi were observed growing in the inoculated wells after 45 days of incubation
and the results are presented in Table 5-1. Attack of LDPE by fungi on the
transmembrane produced metabolites that can be further degraded or assimilated,
contributing to fungal growth. The below results demonstrates that growth of fungi on
the semi-permeable membranes.
Table 5-1: Radial diameter of fungi before and after incubation in mm (± 2 mm)
Before incubation After incubation
3 8
3.5 7
3 6
3.2 5
3 5
5.8.4. FT-IR analysis of LDPE pellets
FT-IR was performed in order to check the biodegradation of LDPE samples
from the transmembrane wells. LDPE from transmembrane well was used as a test
sample where oxidised LDPE (that was not incubated with Fusarium oxysporum)
served as a control.
A decrease in signal corresponding to the carbonyl moiety (at 1715 cm-1)
and many other functional groups can be observed in the FT-IR spectrum shown in
Figure 5-12, suggesting that enzymes of Fusarium oxysporum can attack functional
groups like C=O and cause biodegradation. A decrease in the carbonyl index (0.58) was
observed from the FT-IR spectrum, indicating that biodegradation by Fusarium
oxysporum was by extracellular enzymes that can degrade the carbonyl groups of
polymeric chains. Thus, biodegradation can be achieved in-vitro.
Factors affecting biodegradation
125
Figure 5-12: FT-IR spectrum of LDPE sample from transmembrane well and oxidised
LDPE (control)
5.9. Conclusion
Biodegradation of LDPE with cell-free extracts was possible with various fungi,
with Fusarium oxysporum being particularly effective on untreated LDPE in the
absence of antioxidant. The ability of cell-free extracts to biodegrade LDPE indicates
that microbial enzymes can attack and consume oxidised LDPE. The biodegradation
pattern in cell-free extracts is similar to that of biodegradation by fungal cells.
Biodegradation can be accelerated by adding Mn2+, Cu2+, Fe2+ ion that acts as
co-enzymes. Biodegradation happens by surface erosion. Biodegradation discouraged
by presence of tertiary carbon atoms in the polymer matrix. Chemical oxidation method
is highly effective for biodegradation and it can convert crystalline regions into
amorphous regions and cause higher biodegradation. Process of biodegradation can be
accelerated by providing optimum temperature and pH depending on the microbe
employed. Addition of nitrates and phosphates increases the rate of biodegradation. All
Factors affecting biodegradation
126
these factors can be used to increase biodegradation of LDPE. Figure 5-13 indicates
a LDPE biodegradation set up that can accelerate the process.
Figure 5-13: Imaginary LDPE biodegradation set up by fermentation
The cumulative effect of these factors on biodegradation can be used to
accelerate the biodegradation process. Chemically oxidised LDPE can be added to a
land fill or fermentor to provide the optimal conditions for biodegradation. Fungi that
cause biodegradation of LDPE can also be added to the fermentor, along with low
concentrations of nitrates and Cu2+ (growth factor). Temperature and pH can be adjusted
depending on the fungal consortia used. Thus, it is possible to increase the rate of
LDPE biodegradation.
Based on the results presented above, and results from Chapter 4 it can be
concluded that the biodegradation of LDPE can be positively influenced by the
following factors: surface roughness, number of branches, type of oxidation products
formed and percentage of crystallinity.
LDPE
Processing to increase surface area
and
Chemical oxidation
Fermentor
Co2 and other
gaseous
products
Fusarium
oxysporum
at pH 8 and 35°C
Factors affecting biodegradation
127
Biodegradation can be achieved by fungal strain Fusarium oxysporum and is
not dependent on a cell-wall-based degradation mechanism. Biofilm formation can be
used as a criterion for the isolation of microbes that can degrade oxidised LDPE, but the
formation of biofilms is not essential for degradation to occur. Indeed, the inhibition of
biofilm formation might increase the rate of biodegradation. The availability of oxygen
to the polymer surface is a crucial rate deciding factor of biodegradation. These
observations including that were observed in chapter 4 indicates that certain additives
can increase biodegradation rate of LDPE if they were added during the processing of
polymer. Inclusion of carbohydrate resisting compounds can stop the formation of
biofilm on the surface and increases its biodegradation rate.
The cumulative results of chapter 4 and 5 provide information about the
mechanism of biodegradation. According to the results of AFM and SEM (see Section
4.5.1 and 4.5.2) and rate of biodegradation (see Section 5.4) it can be observed that
biodegradation of the tested LDPE occurs by surface erosion method. Biodegradation
results in the smoothening of LDPE. As the biodegradation resulting in the smoothening
of LPDE, it can be concluded that resistance for continuation of biodegradation arises
from both because of lacking of reactive groups and also because of non permeation of
water into matrix of LDPE. Figure 5-14 depicts the surface erosion of LDPE by
Fusarium oxysporum.
Factors affecting biodegradation
128
Figure 5-14: Schematic representation of biodegradation by surface erosion of LDPE
Untreated LDPE
(smooth surface)
Oxidised LDPE
(rough surface)
Fungi-treated LDPE
(surface eroded)
CHAPTER 6
LACCASE AND CO-METABOLISM
Laccase and co-metabolism
130
6.1. Chapter overview
In this chapter, the attempts that were made to identify enzymes that participate
in LDPE biodegradation are described. Also, the oxidation capability of laccase from
Fusarium oxysporum and factors that influence its activity in vitro are described.
Finally, the effect of co-metabolism on LDPE biodegradation and the reasons for it were
detailed.
6.2. Introduction
Biodegradation of oxidised LDPE using enzymes is an important step in
devising an ideal disposal method. It was also shown that biodegradation of LDPE is
possible with cell-free extracts of Fusarium oxysporum. In this chapter, an investigation
of the enzymes present in cell-free extracts that participate in LDPE biodegradation
process is described. Gel filtration chromatography (GFC) was used to examine the
composition of cell-free extract. The GFC method of cell-free extract preparation was
described in Section 3.17.2. An attempt to degrade LDPE using laccase from Fusarium
oxysporum is also described.
6.3. Biochemical analysis of fungal extract
The amount of protein and carbohydrates in concentrated Fusarium oxysporum
extract (10 ml) were analysed. The detailed procedures were given in Section 3.7.6 and
Section 3.13.2. The protein concentration of the extract was measured as 20 µg/L. while
the carbohydrate concentration was calculated as 293 µg/L.
6.4. Gel filtration chromatography of Fusarium oxysporum extracts
Fusarium oxysporum extracts incubated with oxidised LDPE were examined
using GFC and the protein content of individual fractions was determined (as per the
method described in Section 3.13.2). A peak was seen in Fraction 3 (see Figure 6-1).
Laccase and co-metabolism
131
0
0.2
0.4
0.6
0.8
1
1.2
1.4
1 2 3 4 5 6 7 8 9 10
OD
at 2
54 n
m
Fraction number
Figure 6-1: GFC of Fusarium oxysporum extracts
6.5. SDS-PAGE analysis of Fusarium oxysporum extracts
This experiment was carried out to identify the proteins involved in LDPE
biodegradation by Fusarium oxysporum. The cell-free extract was separated by SDS-
PAGE as described in Section 3.17.3.
SDS-PAGE analysis indicated the presence of a major protein band between the
75 kDa and 60 kDa markers, indicating that LDPE degradation requires protein.
However, three additional light bands were observed at the region near the last marker
(around 37 KDa) (not visible in Figure 6-2), which indicates that other proteins are also
participate in biodegradation.
Laccase and co-metabolism
132
Figure 6-2: SDS-PAGE of the concentrated Fusarium oxysporum extract (Lane 1).
M=molecular weight marker
6.6. Identification of fungal enzymes using plate assays
In order to identify enzymes that participate in biodegradation, Fusarium
oxysporum was cultured on modified Czapek’s-Dox agar plates supplemented with
pulverised oxidised LDPE powder and various enzyme substrates. The plates were
cultivated for a week and observed for the presence of positive reactions (as per Section
3.13.1). The presence of laccase is indicated by the development of a blue colour. The
fungal culture was found to exhibit laccase activity (see Figure 6-3).
Laccase and co-metabolism
133
Figure 6-3: Identification of laccase secretion by Fusarium oxysporum
From the above result it can be concluded that Fusarium oxysporum was
induced to produce laccase by the presence of LDPE. Though Fusarium oxysporum has
the capability to produce several cell wall degrading enzymes (see Section 2.21), only
laccase is secreted in the presence of LDPE. This shows the inductive capacity of
LDPE. Laccase production has been shown to be triggered by the presence of polymeric
substances (Lee et al. 2004). It has previously been shown that production of laccase
from Fusarium oxysporum is dependent on several other factors, such as the type of
cultivation, carbon limitation and the nitrogen source (Shraddha et al. 2011). In the
above experiment Fusarium oxysporum was incubated in Czapek’s-Dox media (see
Section 3.6.6), which contains copper, ferrous ion, zinc, nitrates and phosphates. Also, it
has LDPE as the only carbon source. So it can be concluded that all these ions and
nitrogen and carbon sources further encouraged production of laccase by Fusarium
oxysporum.
Laccase and co-metabolism
134
6.7. Biodegradation with laccase
Laccase was purified from fungal extracts by gel filtration (as per Section 3.9.1)
and 100 mmol phosphate buffer was added to each extract to make up the total volume
to 10 mL (Patrick et al. 2009). LDPE pellets were immersed in this buffer. The pH was
set at 7 and the temperature was set at 30 °C. The weight loss of the LDPE pellets and
dissolved carbon dioxide concentrations were measured.
The results indicated that biodegradation did not occur in the presence of laccase
alone. A weight loss of less than 0.02% was observed. No difference in
dissolved carbon dioxide concentration between the control (without laccase) and the
test (with laccase) was observed. These data indicate that laccase have little or no effect
on LDPE weight loss. This indicates that biodegradation of polyethylene is complicated
and requires other factors. This experiment was performed several times with no
detectable biodegradation occurring each time.
6.8. Effect of co-metabolites
These experiments were conducted in order to monitor the effect of co-
metabolites of LDPE in its biodegradation. All these experiments were conducted in
two stages. In the first stage, biodegradation experiments were conducted with
Fusarium oxysporum mycelia. In the next stage, they were conducted with Fusarium
oxysporum cell-free extract. The preparation of cell-free extracts was described in
Section 3.13.
6.8.1. Effect of monosaccharides
Glucose, galactose and fructose were added to the Czapek’s-Dox medium at
0.25, 0.50, 0.75 and 1 % (w/v) and the weight loss of the LDPE pellets was measured
after 45 days. The details of the incubation protocol were described in Section
3.8.1. This experiment was performed in triplicate with both mycelia and cell-free
extracts. The average weight loss for each sugar concentration was determined, and
the results are shown in Figure 6-4.
Laccase and co-metabolism
135
0
2
4
6
8
10
12
14
16
18
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of monosaccharides (% w/v)
GlucoseGalactoseFructose
A
0
2
4
6
8
10
12
14
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of monosaccharides (% w/v)
GlucoseGalactoseFructose
B
Figure 6-4: Effect of monosaccharides on LDPE biodegradation with mycelia (A) and
with cell-free extract (B) of Fusarium oxysporum.
Laccase and co-metabolism
136
Biodegradation consistently increased with increasing concentrations of glucose,
fructose and galactose. However, among these monosaccharides, galactose showed a
lesser effect on weight loss. The addition of galactose inhibited Fusarium oxysporum
growth, but did not prevent biodegradation to occur. All monosaccharides showed
biodegradation in both an active and passive manner. In the active manner, weight loss
is caused by the increase in fungal biomass, which in turn increased the output of
enzymes. Under the passive biodegradation mechanism, the weight loss is caused by
activating enzymes that participate in biodegradation. This implies that biodegradation
is not solely dependent on the growth of Fusarium oxysporum.
6.8.2. Effect of disaccharides
Maltose, lactose and sucrose were added to the Czapek’s-Dox medium at
0.25, 0.50, 0.75 and 1 % (w/v) and the weight loss of the LDPE pellets was measured
after 45 days. This experiment was performed in triplicate, and the average weight loss
for each sugar concentration determined. This experiment was also performed with
mycelia and cell-free extracts of Fusarium oxysporum (see Figure 6-5).
Laccase and co-metabolism
137
0
2
4
6
8
10
12
14
16
18
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of disaccharides (% w/v)
MaltoseLactoseSucrose
A
0
2
4
6
8
10
12
14
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of disaccharides (% w/v)
MaltoseLactoseSucrose
B
Figure 6-5: Effect of disaccharides on LDPE biodegradation with mycelia (A) and with
cell-free extract (B) of Fusarium oxysporum.
Laccase and co-metabolism
138
A gradual increase in weight loss with increasing disaccharide concentration was
observed. At higher concentrations, lactose outperformed maltose and sucrose in
increasing biodegradation. Among these sugars, sucrose had a consistently smaller
effect on biodegradation (with the exception of at 0.25%). Compared to the
monosaccharides, the addition of disaccharides resulted in a greater LDPE weight loss.
6.8.3. Effect of polysaccharides
Starch and cellulose were added to the Czapek’s-Dox medium at 0.25, 0.50, 0.75
and 1 % (w/v) and the weight loss was measured after 45 days. This experiment was
performed in triplicate and the average weight loss for each concentration was
determined. This experiment was also performed with mycelia and with cell-free extract
(see Figure 6-6).
Laccase and co-metabolism
139
0
2
4
6
8
10
12
14
16
18
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of polysaccharides (% w/v)
StarchCellulose
A
0
2
4
6
8
10
12
14
16
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of polysaccharides (% w/v)
StarchCellulose
B
Figure 6-6: Effect of polysaccharides on LDPE biodegradation with mycelia (A) and
with cell-free extract (B) of Fusarium oxysporum.
Laccase and co-metabolism
140
An increased concentration of polysaccharides was associated with greater
weight loss of LDPE. At higher concentrations, cellulose outperformed starch in
facilitating biodegradation.
6.8.4. Effect of methanol, ethanol and propanol
Alcohols were tested for their impact on the biodegradation of oxidised LDPE
(see Figure 6-7). They were added to the Czapek’s-Dox medium at concentrations of
0.25%, 0.50%, 0.75% and 1% v/v.
0
2
4
6
8
10
12
14
16
18
20
22
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of alcohols (% v/v)
EthanolMethanolPropanol
A
Laccase and co-metabolism
141
0
2
4
6
8
10
12
14
16
18
20
Control 0.25 0.5 0.75 1
Aver
age
wei
ght
loss
(%)
Concentration of alcohols (% v/v)
EthanolMethanolPropanol
B
Figure 6-7: Effect of alcohols on LDPE biodegradation with mycelia (A) and with cell-
free extract (B) of Fusarium oxysporum.
The effect of ethanol on LDPE biodegradation was substantial, followed by
propanol and methanol.
6.8.5. Summary of effect of co-metabolites
Addition of co-metabolites of LDPE to the growth medium increased the extent
of fungal biodegradation. Among all these co-metabolites, the polysaccharides showed
a major impact in terms of increased weight loss. Co-metabolites in general increase the
growth of microbes, causing a ‘priming effect’ (Ladygina et al. 2006). The activation of
microorganism growth by easily available substrates is mostly considered to be due to a
priming effect. The priming effect of the above-mentioned carbon sources triggered the
production of enzymes with a non-specific substrate attachment capacity. This increased
the degradation of LDPE. Following the addition of ethanol, it was observed that the
fungi grew on oxidised LDPE and formed a cocoon-like structure that enveloped the
Laccase and co-metabolism
142
LDPE pellets. Among all the co-metabolites, the addition of ethanol resulted in
an immediate growth reaction from the fungi. This is because ethanol is readily soluble
in the growth medium, and hence it is available more rapidly than other carbon sources.
It has been proven that the amount and composition of carbon sources available in
a growth medium greatly influences its microbial activity (Nelson et al. 1994).
The addition of sucrose, glucose and lactose to polyolefin chains has been
demonstrated to significantly increase its biodegradation (Galgali et al. 2002) by
increasing its susceptibility to enzymatic attack. Volke-Seplveda et al. (2002) observed
that ethanol can be used to degrade non-oxidised LDPE and that the addition of ethanol
increases biodegradation. This is due to both the priming effect of sugars and their
capacity to physically direct the enzymes that are participating in biodegradation.
6.9. Effect of laccase
The enzyme laccase that was isolated and separated from Fusarium oxysporum
by GFC (see Section 3.17.2) was assayed for its capacity to oxidise LDPE. The
incubation protocol was described in Section 3.20.5. The effect of oxidation was
measured in terms of the carbonyl index, A1712/A1465 (see Figure 6-8).
Laccase and co-metabolism
143
0
0.5
1
1.5
2
0 1 2 3 4
Car
bony
l ind
ex
Period of incubation in hours
Figure 6-8: The effect of period of incubation with laccase on oxidation, as reflected by
the carbonyl index
From the above results it can be seen that laccase increases the oxidation of
LDPE by increasing the formation of carbonyl groups (C=O) on the LDPE surface. It
has been reported that laccase can assist in the oxidation of the hydrocarbon backbone
of polyethylene (Santo et al. 2012; da Luz et al. 2013; Bhardwaj et al. 2012). During the
photo-oxidation process of LDPE reactive groups are created on its surface. These
reactive groups contain carbonyl and hydroxyl groups (see Section 4.4.1), which allow
further oxidation by laccase via electron transfer. Thus, laccase is able to increase the
carbonyl index of LDPE.
6.9.1. Effect of manganese, copper, iron (II) and zinc chloride on laccase
oxidation
This experiment was conducted to assess the influence of co-enzymes
(manganese, copper, ferrous iron and zinc chloride) on LDPE biodegradation on laccase
enzyme. As it has been demonstrated that metal ions with +2 valency act as co-enzymes
in LDPE biodegradation by Fusarium oxysporum it is important to determine the effect
Laccase and co-metabolism
144
of these ions on LDPE oxidation in the presence of laccase. To do this, photo-oxidised
LDPE pellets were incubated with laccase in phosphate buffer at pH 7 at 30 °C.
Chloride salts were added to the samples at concentrations of 0.25%, 0.50% and 0.75%
v/v and the mixes were then incubated for 24 h (Section 3.21). The results are shown in
Figure 6-9.
0.05
0.15
0.25
0.35
0.45
0.55
1 2 3 4
Car
bony
l ind
ex
Control0.25 mmol0.50 mmol0.75 mmol1 mmol
Figure 6-9: Effect of manganese, copper, ferrous and zinc chloride on oxidation of
LDPE (measured by the carbonyl index)
Ferrous and zinc ions showed little effect on LDPE oxidation by laccase. In
contrast, copper and manganese enhanced laccase-dependent oxidation of LDPE, with
Cu2+ having a greater impact on laccase activity. As laccase contains copper in its
enzymatic structure, addition of this ion to the reaction mix enhances enzymatic activity
(Cañero & Roncero 2008). The copper ions of laccase accept electrons from the reactive
groups of the LDPE and participate in its oxidation. Moreover, it has previously been
shown that laccase activity is enhanced in the presence of copper and manganese ions
(Palmieri et al. 2000; Rogalski et al. 2006). These two ions may also participate in the
reaction mediated by laccase.
Laccase and co-metabolism
145
6.9.2. Effect of co-metabolism and laccase
These experiments were performed to investigate effect of laccase in co-
metabolic degradation of LDPE. As observed in Chapter 5, the biodegradation of LDPE
was enhanced by presence of sugars and alcohols. Addition of these carbon sources
increased the cell density and facilitated the enzymatic attack of LDPE by Fusarium
oxysporum.
6.9.3. Effect of sucrose and ethanol
This experiment was performed to examine the effect of sucrose (disaccharide)
on laccase. First, photo-oxidised LDPE pellets were incubated with laccase in phosphate
buffer at pH 7 at 30 °C. Sucrose and ethanol were added to the reaction mixtures at
concentrations of 0.25, 0.50, 0.75, and 1 % w/v and v/v, respectively. The reaction
mixes were then incubated for 24 h (see Section 3.21.1).
0
0.5
1
1.5
2
2.5
0 1 2 3 4 5 6
Car
bony
l ind
ex
Concentration of sucrose and ethanol in mmol/L
SucroseEthanol
Figure 6-10: Effect of concentration of sucrose on LDPE oxidation (measured by the
carbonyl index)
The carbonyl index of the UV-treated LDPE was found to increase with the
addition of sucrose (see Figure 6-10). Sucrose participates in electron transfer between
Laccase and co-metabolism
146
LDPE reactive groups and laccase. This results in an increase of the carbonyl index of
LDPE. It has been shown that low concentrations of sucrose induce the activity of
laccase secreted by strains of Fusarium oxysporum (Cañero & Roncero 2008).
Likewise, laccase secreted by Pleurotus ostreus was found to be activated by the
addition of sucrose to the basal media (Sivakami et al. 2012). Thus, it can be concluded
that sucrose increases electron transfer between the substrate (oxidised LDPE) and the
enzyme (laccase).
The addition of ethanol to the laccase buffer increased the enzyme’s oxidation
efficiency against UV-treated LDPE. The resulting oxidation efficiency was greater
than that observed when sucrose was used to increase the oxidation capacity of laccase
against UV-treated LDPE. As ethanol possesses higher solubility than sucrose it is
easily available to participate in reaction. It is already well established that ethanol
induces laccase activity, depending on the laccase source. Laccase secreted by Coriolus
versicolor was activated by ethanol at concentrations between 0–2.5 M, and the activity
of laccase secreted by Trametes versicolor was increased by the addition of ethanol to
the fungal culture (Maceiras et al. 2001). Thus, it can be concluded that ethanol also
encourages electron transfer between LDPE and laccase.
6.9.4. Summary
Fusarium oxysporum secretes laccases in the presence of LDPE, and these
laccases appear to play an oxidative role in LDPE biodegradation. However, the
presence of laccase alone was not enough to degrade oxidised LDPE. Biodegradation of
LDPE was not observed in the presence of laccases in vitro, indicating that
biodegradation might not be possible with a single enzyme; but rather, may require a
combination of enzymes.
Co-metabolites (ethanol and sucrose) enhanced the extent of biodegradation by
two major pathways. First, these carbon moities increased the cell density of Fusarium
oxysporum by serving as an immediately available carbon sources. Second, they also
oxidise LDPE by activating the laccases secreted by the fungi. These factors
cumulatively increased the biodegradation by fungi. Figure 6-11 illustrates the pathways
that increase LDPE biodegradation by co-metabolism.
Laccase and co-metabolism
147
Figure 6-11: Cumulative effect of co-metabolism on laccase-induced biodegradation of
LDPE
Co-metabolites (ethanol and sucrose)
Increased cell quantity Increased laccase activity
Increased laccase quantity Increased oxidised LDPE quantity
Increased biodegradation
CHAPTER 7
CONCLUSION AND FUTURE STUDIES
Conclusion and future studies
149
In this chapter, a description is given of the factors that were found to influence
the biodegradation of LDPE. Future directions for research that may lead to methods to
allow the efficient oxidation and rapid biodegradation of LDPE are also suggested.
7.1. Conclusion
Biodegradation of oxidised LDPE is possible if an appropriate strain of fungus
can be isolated. Various genera of fungi show the capacity to degrade oxidised LDPE.
Fusarium is an ideal fungus to use to study the process of biodegradation, as it
possesses a natural capacity to degrade plant polymers. Unlike Mucor, it is easily
recognisable on PDA plates as it forms pale pink colour on this medium. Additionally,
selective media (with malachite green) can be used to isolate Fusarium species to
minimise contamination. Mucor is also a promising candidate for the selection of
strains with biodegradative capacity.
Biodegradation of oxidised LDPE resulted in reduced roughness of the polymer
surface. The fungi isolated preferentially degraded rougher surfaces on LDPE, as these
offer sites for physical interaction between the fungal enzymes and the LDPE and hence
are more rapidly degraded than smoother surfaces. It was noted that depending on the
incubation period and the strains of microbes used, the crystallinity of the
biodegraded LDPE increased. This indicates that creating rougher surfaces is advisable
in order to achieve faster biodegradation rates. Surface roughness can be created in
polymers by techniques such as chemical or heat treatment and using copper-based
salts. The carbonyl groups created by oxidation can be easily biodegraded. These
groups can be in either in an acid (–COOH) or ester (–C=O) form. This property
indicates that LDPE with these functional groups will be degraded more rapidly than
LDPE with other functional groups. LDPE to which Irganox® has been added can be
biodegraded if an appropriate strain of fungus is isolated. The factors that influenced its
biodegradation were similar to that of LDPE without added antioxidant. However, it
shows differences in its fungal compatibility.
The detection of biodegradation, and its extent, can be easily achieved and
compared using a simple staining technique. Methylene blue can be used to check the
extent of biodegradation in a cost-effective method.
Conclusion and future studies
150
The rate of biodegradation is influenced by the addition of salts, co-metabolites
and alcohols. Salts of copper, manganese and ferrous iron accelerate biodegradation by
Fusarium oxysporum. Alcohols (ethanol, propanol and methanol) and sugars (sucrose,
fructose and maltose) also enhance LDPE biodegradation, as do nitrates and phosphates.
Chemical oxidation is more effective than thermal treatment and UV treatment and
rearranges entire crystalline phases of the polymer. However, only surface oxidation of
LDPE pellets can be achieved. In order to prevent this, it is advisable to produce flat
sheets of LDPE to increase its surface area before submitting it to oxidation and
subsequent biodegradation. The summative use of these factors increases the
biodegradation of LDPE.
The necessity for biofilm formation during biodegradation was examined, and it
was concluded that biofilm formation on LDPE can be used as an indication of
biodegradation but that it is not essential to achieve biodegradation. In fact, the
formation of biofilm on LDPE decreased the rate of its biodegradation, probably by
decreasing oxygen availability. It is therefore advisable to produce LDPE that can resist
biofilm formation. Additives that can prevent the accumulation of carbohydrates on the
surface of LDPE can be incorporated into its structure. This will prevent biofilm
formation on the LDPE surface but will not inhibit enzymatic action. LDPE
biodegradation happens by surface erosion of it.
Fungal extracts exhibiting the capacity to degrade LDPE suggest that
biodegradation of LDPE can take place in vitro. However, the effect of extracts is
relatively less than that of intact fungi (3%). GFC and SDS-PAGE analysis of fungal
extract identified the presence of a protein responsible for biodegradation, while PDA-
based enzymatic analysis suggested the presence of laccase enzyme. The laccase
secreted by Fusarium oxysporum played an important role in LDPE biodegradation. Co-
metabolism of LDPE increases its rate of biodegradation. In the presence of co-
metabolites, laccase enhanced LDPE oxidation, resulting in increased biodegradation of
LDPE by fungi.
Conclusion and future studies
151
7.2. Suggestions for future study
Samples from soil, leachate and many other sources were thoroughly screened
for their ability to promote biodegradation. In contrast, marine samples were not well
examined. Microbes isolated from marine environments will possess different metabolic
rates and growth requirements. This variation might lead to the isolation of more
competent strains with higher degradation capabilities. Along with biodegradation,
biofouling can be seen during biodegradation in the marine ecosystem (Flemming
1998), which may enhance the degradation effect.
Table 7-1: Various microbes isolated from marine environment
Polymer Microbe isolated from marine environment
Polyethylene (PE) Rhodococcus ruber (Orr et al. 2004)
Poly (vinylchloride) (PVC) Micrococcus luteus (Patil & Bagde 2012)
Poly (ethylene terephthalate)
(PET)
Alteromonas australica sp. Nov. (Ivanova et al.
2013)
Biodegradation of LDPE needs to be understood on a large scale (e.g., in
landfills and fermenters). The mechanism of oxidation of LDPE by weathering needs to
be understood in order to be able to accelerate it. The effect of commercial additives
such as stabilisers, slip agents and antistatic agents on biodegradation must be
understood. Utilising the genes for laccase and other enzymes to construct genetically
engineered microorganisms to produce these enzymes in a continuous manner will
accelerate biodegradation.
Increasing the efficiency of biodegradation is necessary as it provides an
efficient and effective disposal method for LDPE. Large-scale biodegradation of LDPE
is required to address its massive accumulation in the environment. This work provides
a comprehensive idea of how to accelerate the biodegradation of LDPE at larger scales
using fungi.
Fusarium strains have the capacity to degrade LDPE at a relatively rapid rate.
Methods that can increase the rate of biodegradation were proposed and discussed in
Conclusion and future studies
152
detail in this thesis. A laccase was identified from a strain of Fusarium isolated in this
study, and was shown to have the ability to increase the oxidation of LDPE, especially
in the presence of co-metabolites.
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