Application of White-rot Fungi for the

Biodegradation of Natural Organic Matter

in Wastes

A thesis submitted in the fulfilment of the requirements for

the degree of Master of Engineering

Monn Kwang Lee

Bachelor of Engineering (Chemical Engineering)

School of Civil and Chemical Engineering

RMIT University, Melbourne

October 2005
Declaration
I, Monn Kwang Lee, certify that except where due acknowledgement has been made, the
work is that of the author alone; the work has not been submitted previously, in whole or in
part, to qualify for any other academic award; the content of the thesis is the result of work
which has been carried out since the official commencement date of the approved research
program; and, any editorial work, paid or unpaid, carried out by a third party is
acknowledged.

Monn Kwang Lee

October 2005

ii
Acknowledgements
I would like to first thank Prof. Felicity Roddick and Dr. John Harris for being my
supervisors. I do thank you, especially Prof. Felicity Roddick, for the precious advice, support
and guidance given during the course of the research. Thank you for helping me to find
financial support from RMIT.

I would love to express my deep appreciation to my beloved parents for their support and
encouragement. Thank you so much for being there with me.

Thanks to Farhad Younos for microbiological techniques and laboratory technical assistance.
Also my thanks to Chau Nguyen for instrument assistance and safety advice. Thank you to
Dr. Nichola Porter and my fellow postgraduate students for sharing ideas whenever we have
our CRC meetings.

Lastly I want to acknowledge Australian Water Quality Centre for HPSEC analyses. I am
grateful to those who helped me with my research, directly or indirectly, thank you very
much.

iii
Table of Contents
Declaration ................................................................................................................................ii
Acknowledgements ..................................................................................................................iii
Table of Contents.....................................................................................................................iv
Abbreviations ..........................................................................................................................vii
List of Figures ..........................................................................................................................ix
List of Tables..........................................................................................................................xiii
SUMMARY............................................................................................................................... 1
List of Publications ................................................................................................................... 4
Chapter 1 Introduction ...................................................................................................... 5
Chapter 2 Literature Review ............................................................................................. 7
2.1 Natural Organic Matter............................................................................................... 7
2.1.1 Origin.................................................................................................................. 7
2.1.2 Composition and chemical structure .................................................................. 8
(i) Non-humic substances.......................................................................................... 10
(ii) Humic substances ............................................................................................. 10
2.2 Characterisation of NOM ......................................................................................... 13
2.2.1 Whole water characterisations.......................................................................... 13
(i) TOC, DOC and BOM analyses ............................................................................ 13
(ii) Spectrophotometric analysis............................................................................. 13
(iii) Size characterisation of NOM .......................................................................... 14
2.2.2 Fractionation..................................................................................................... 14
2.3 Impact of NOM on Water Quality and Treatment ................................................... 16
2.4 White-rot Fungi ........................................................................................................ 17
2.4.1 Effect of temperature and pH ........................................................................... 18
2.4.2 Effect of supplements ....................................................................................... 19
2.4.3 Effect of agitation ............................................................................................. 22
2.5 Mechanism of Enzymatic Degradation by White-rot Fungi .................................... 22
2.5.1 Lignin peroxidase ............................................................................................. 23
2.5.2 Manganese-dependent peroxidase.................................................................... 25
2.5.3 Laccase ............................................................................................................. 27
2.6 Potential Applications of White-rot Fungi in Bioremediation ................................. 29
Chapter 3 Materials and Methods .................................................................................. 33
3.1 NOM Samples .......................................................................................................... 33

iv
 3.2 Micro-organisms....................................................................................................... 34
3.3 Medium and Culture Conditions .............................................................................. 34
3.4 Preparation of Inoculum ........................................................................................... 35
3.4.1 Fungal inoculum ............................................................................................... 35
3.4.2 Yeast inoculum................................................................................................. 35
3.5 Supplements ............................................................................................................. 35
3.6 Analytical Methods .................................................................................................. 35
3.6.1 pH ..................................................................................................................... 35
3.6.2 Dissolved organic carbon ................................................................................. 36
3.6.3 Absorbance ....................................................................................................... 36
3.6.4 Determination of absorbance correction factor ................................................ 36
3.6.5 Determination of glucose concentration........................................................... 37
3.6.6 Dry weight of biomass...................................................................................... 37
3.6.7 Enzyme assays.................................................................................................. 37
(i) Laccase activity .................................................................................................... 37
(ii) Lignin peroxidase ............................................................................................. 38
(iii) Manganese-dependent peroxidase.................................................................... 38
3.6.8 High performance size exclusion chromatography .......................................... 39
3.6.9 Fractionation of NOM ...................................................................................... 40
Chapter 4 Decolourisation and Bioremediation of MIEXTM NOM ............................. 42
4.1 Removal of Different Preparations of NOM by P. chrysosporium ATCC 34541 ... 42
4.2 Fractionation of the MIEXTM NOM Preparations .................................................... 45
4.2.1 Comparison of A446 of the NOM fractions....................................................... 46
4.2.2 Comparison of A254 of the NOM fractions....................................................... 47
4.2.3 Comparison of molecular weight distribution of the NOM fractions .............. 48
4.3 Molecular Size Distribution of the NOM after Treatment with P. chrysosporium
ATCC 34541 ........................................................................................................................ 52
4.4 Selection of Medium ................................................................................................ 54
4.5 Selection of Organism .............................................................................................. 57
Chapter 5 Biodegradation of NOM by Trametes versicolor .......................................... 66
5.1 Improving NOM Removal by Altering Culture Conditions..................................... 66
5.1.1 Incubation temperature ..................................................................................... 66
5.1.2 Carbon source level .......................................................................................... 72
5.1.3 Types of inoculum ............................................................................................ 75
5.1.4 Effect of NOM concentration ........................................................................... 77

v
 5.2 Enhancement of Enzyme Production ....................................................................... 80
5.2.1 Determination of cultivation time based on maximum laccase activity........... 81
5.2.2 Effect of supplements ....................................................................................... 82
5.2.3 Effect of temperature and pH on laccase activity............................................. 84
(i) Effect of temperature ............................................................................................ 84
(ii) Effect of pH ...................................................................................................... 85
5.2.4 Effect of agitation ............................................................................................. 86
5.3 Enzymatic Treatment of NOM ................................................................................. 90
5.3.1 Effect of pH ...................................................................................................... 90
5.3.2 Effect of temperature ........................................................................................ 91
5.4 Biodegradation of NOM by T. versicolor ................................................................ 93
Chapter 6 Conclusions and Recommendations ............................................................. 98
6.1 Conclusions .............................................................................................................. 98
6.2 Recommendations .................................................................................................... 99
References ............................................................................................................................. 101
APPENDICES....................................................................................................................... 114
Appendix 1 Correlation between NOM concentration and A446 and A254 ..................... 114
Appendix 2 Absorbance correction factor...................................................................... 115
Appendix 3 Typical standard curve for glucose determination ..................................... 117
Appendix 4 Standard curve for determination of apparent molecular weight (Dalton) in
HPSEC analysis.................................................................................................................. 118

vi
Abbreviations
A254 Absorbance at 254 nm
A446 Absorbance at 446 nm
AOC Assimilable organic carbon
AWQC Australian Water Quality Centre
BDOC Biodegradable dissolved organic carbon
BOM Biodegradable organic matter
CHA Hydrophilic charged
C:N Ratio of carbon to nitrogen
DBP Disinfection by-products
DMP 2, 6-dimethoxyphenol
DNS 3', 5’-dinitrosalicylic acid
DOC Dissolved organic carbon
GAC Granular activated carbon
HAAs Haloacetic acids
HPSEC High performance size exclusion chromatography
Lac Laccase
LAS Linear alkyl benzene sulphonate
LiP Lignin peroxidase
Mi Molecular weight
Σni M i
Mn Number average molecular weight, M n =
Σni

Σn M
2

Mw Weight average molecular weight, M w = i i

Σn i M i
Mw: Mn Polydispersity
MEA Malt extract agar
MIEXTM Magnetic ion exchange
MnP Manganese-dependent peroxidase
ni Number of molecules of weight Mi
NEU Hydrophilic neutral
NOM Natural organic matter
PAC Powdered activated carbon
PAHs Polyaromatic hydrocarbons
PCP Pentachlorophenol

vii
POC Particulate organic carbon
RBBR Remazol Brilliant Blue R
RCF Relative centrifugal force
RH Lignin or phenolic substrate
rpm Revolutions per minute
SEC Size exclusion chromatography
SHA Slightly hydrophobic acids
SSF Solid-state fermentation
A254
SUVA Specific UV absorbance,
DOC
THMs Trihalomethanes
TOC Total organic carbon
Tween 80 Polyoxyethylene sorbitan monooleate
U Unit
VA Veratryl alcohol
VHA Very hydrophobic acids
Yx/s Yield, biomass produced__
glucose consumption

viii
List of Figures
Figure 2.1 General structure of NOM.................................................................................. 9
Figure 2.2 Classification of NOM. .................................................................................... 10
Figure 2.3 Chemical properties of humic substances. ....................................................... 11
Figure 2.4 Model structure of humic acid.......................................................................... 12
Figure 2.5 Model structure of fulvic acid. ......................................................................... 12
Figure 2.6 Veratryl alcohol as electron transfer mediator. ................................................ 19
Figure 2.7 Proposed schemes for the biodegradation of lignin. ........................................ 23
Figure 2.8 Mechanism of Cα-Cβ cleavage by LiP.............................................................. 24
Figure 2.9 Catalytic cycle of lignin peroxidase. ................................................................ 25
Figure 2.10 Catalytic cycle of manganese peroxidase......................................................... 26
Figure 2.11 Oxidation of a free phenolic β-1 substructure by MnP. ................................... 27
Figure 2.12 Proposed degradation of phenolic β-1 model compounds by laccase from C.
versicolor.......................................................................................................................... 28
Figure 3.1 Proposed mechanism for the oxidation of guaiacol by laccase. ....................... 37
Figure 3.2 Proposed mechanism for catalysing reduction of VA by LiP. ......................... 38
Figure 3.3 Proposed mechanism for the oxidation of DMP by MnP................................. 39
Figure 3.4 Schematic diagram of the fractionation unit..................................................... 41
Figure 4.1 History plot showing pH, glucose consumption, A446 and A254 for NOM 3
incubated with P. chrysosporium ATCC 34541 at 36oC and 130 rpm for five days. (A254
represents readings of 1/10 dilution of culture medium).................................................. 43
Figure 4.2 A446 and A254 of NOM 1, NOM 2 and NOM 3, 100 mg C/L initial NOM
concentration, P. chrysosporium ATCC 34541. (A254 represents readings of 1/10 dilution
of culture medium) ........................................................................................................... 44
Figure 4.3 NOM removals (as mg, converted from A446 and A254), glucose consumption
(g/L) and dry weight of biomass generated (mg) for the three NOM preparations on day
5, 100 mg C/L initial NOM concentration, P. chrysosporium ATCC 34541................... 45
Figure 4.4 The proportions of each fraction in the three NOM preparations: (A) NOM 1,
(B) NOM 2 and (C) NOM 3 solutions. (N = 2; i.e., number of times each was
determined)....................................................................................................................... 46
Figure 4.5 A446 and A446/DOC of the fractions of the three NOM preparations................ 47
Figure 4.6 A254 and A254/DOC of the fractions of the three NOM preparations................ 47
Figure 4.7 HPSEC chromatograms for the (A) ‘whole’ NOM, (B) VHA, (C) SHA, (D)
CHA and (E) NEU fractions for all NOM preparations................................................... 50

ix
Figure 4.8 Weight average molecular weight (Mw), number average molecular weight
(Mn) and polydispersity for the three NOM preparations................................................. 51
Figure 4.9 HPSEC chromatograms for the three NOM preparations incubated with P.
chrysosporium in Waksman medium, control-NOM (medium plus NOM), control-P.
chry 34541 (P. chrysosporium ATCC 34541 grown without NOM)............................... 53
Figure 4.10 Reduction in weight average molecular weight (Mw) and number average
molecular weight (Mn) after the treatment of the NOM preparations with P.
chrysosporium ATCC 34541............................................................................................ 54
Figure 4.11 Decolourisation and glucose consumption in the different growth media
containing 100 mg C/L NOM and P. chrysosporium ATCC 34541. ............................... 55
Figure 4.12 Dry weight biomass (mg) and NOM removal in terms of A446 (converted to
mg/L) by P. chrysosporium ATCC 34541 in different culture media after 14 days........ 56
Figure 4.13 Decolourisation of the NOM (100 mg C/L initial concentration) in Waksman
medium by (A) white-rot fungi and (B) Saccharomyces sp............................................. 57
Figure 4.14 Biomass dry weight (mg) and glucose consumptions (g/L) of
Saccharomyces spp. 1-3 and T. versicolor, Waksman medium with 2 g/L initial glucose
content……. ..................................................................................................................... 58
Figure 4.15 Biomass of Saccharomyces sp. 2 (A) incubated in the absence of NOM and (B)
after incubation with 100 mg C/L NOM. ......................................................................... 59
Figure 4.16 HPSEC chromatograms for NOM remaining after treatment with
Saccharomyces sp. 2 in Waksman medium for seven days (control-NOM: culture in the
absence of the yeast; control-Saccharomyces sp. 2: culture in the absence of NOM). .... 59
Figure 4.17 Comparison of NOM removal (as mg, converted from A446 and A254), glucose
consumption and biomass for the three white-rot fungi at initial concentrations of 2 g/L
glucose and 100 mg C/L NOM after five days................................................................. 60
Figure 4.18 Biomass of (A) P. chrysosporium ATCC 34541 and (B) T. versicolor in the
absence of NOM (top), and after five days incubation with 100 mg C/L NOM
(bottom)………………………………………………………………………………….61
Figure 4.19 HPSEC chromatograms for NOM treated with P. chrysosporium ATCC 34541
and T. versicolor ATCC 7731 (control-NOM: culture in the absence of the fungi;
control-P. chry 34541 or control-T. ver 7731: culture in the absence of NOM).............. 61
Figure 4.20 Weight average molecular weight (Mw) and number average molecular weight
(Mn) for the NOM (control) and the NOM remaining after five days treatment with P.
chrysosporium ATCC 34541 or T. versicolor ATCC 7731. ............................................ 62

x
Figure 4.21 Reaction of guaiacol on T. versicolor agar plate colony indicating presence of
the laccase enzyme. .......................................................................................................... 63
Figure 4.22 Activity of the extracellular phenoloxidase enzymes in 3-day cultures of the
three white-rot fungi. ........................................................................................................ 63
Figure 5.1 History plots for T. versicolor cultures containing 100 mg C/L NOM incubated
at 30oC and 36oC, Waksman medium 2 g/L initial glucose. ............................................ 67
Figure 5.2 NOM removals (as mg, converted from A446 and A254) and biomass produced
(mg) in T. versicolor cultures at 30oC and 36oC............................................................... 68
Figure 5.3 Biomass of T. versicolor incubated at (A) 30oC and (B) 36oC in the absence of
(top) and presence of (bottom) NOM after nine days incubation, 100 mg C/L NOM..... 68
Figure 5.4 Activity of the extracellular phenoloxidase enzymes of the T. versicolor
cultures incubated at 30oC and 36oC. ............................................................................... 69
Figure 5.5 HPSEC chromatograms for NOM remaining after treatment with T. versicolor
incubated at 30oC and 36oC; controls represent fungal cultures grown in the absence of
NOM……......................................................................................................................... 71
Figure 5.6 Weight average molecular weight (Mw) and number average molecular weight
(Mn) for the NOM (control) and the NOM remaining after nine days treatment with T.
versicolor ATCC 7731 at 30oC and 36oC......................................................................... 72
Figure 5.7 History plots for T. versicolor cultures containing 100 mg C/L NOM incubated
o
at 30 C, Waksman medium containing 2 g/L and 5 g/L initial glucose........................... 73
Figure 5.8 NOM removals (as mg, converted from A446 and A254) and biomass produced
(mg) in T. versicolor cultures containing 2 g/L and 5 g/L initial glucose........................ 74
Figure 5.9 Activity of the extracellular phenoloxidase enzymes of the T. versicolor
cultures containing 2 g/L and 5 g/L initial glucose. ......................................................... 74
Figure 5.10 History plots for T. versicolor cultures in Waksman medium (2 g/L glucose)
containing 100 mg C/L NOM incubated at 30oC, inoculated with either spore
suspensions or three agar plugs. ....................................................................................... 75
Figure 5.11 A446 and A254 for Waksman medium (2 g/L glucose) containing 100 mg C/L
NOM and three agar plugs without fungus. ..................................................................... 76
Figure 5.12 NOM removals (as mg, converted from A446 and A254) and laccase activity in
T. versicolor cultures, inoculated with either spore suspension or plugs......................... 77
Figure 5.13 NOM removals (as mg, converted from A446 and A254), and glucose
consumption in T. versicolor cultures with different NOM concentrations, plug
inoculum. .......................................................................................................................... 77

xi
Figure 5.14 NOM removals (as mg, converted from A446 and A254) by T. versicolor for
different NOM concentrations.......................................................................................... 78
Figure 5.15 Activity of the extracellular phenoloxidase enzymes of T. versicolor in cultures
containing varying NOM concentrations, plug inoculum. ............................................... 79
Figure 5.16 A446 for 100 mg C/L NOM incubated in Waksman medium containing 4.5 g/L
wheat bran in the absence of fungus................................................................................. 81
Figure 5.17 History plot of T. versicolor cultivated in the presence of 4.5 g/L wheat bran.
The data points correspond to mean values of duplicate assays. ..................................... 82
Figure 5.18 Effect of temperature on Lac activity at pH 4.5 in cultures supplemented with
4.5 g/L wheat bran (+WB) and 4.5 g/L wheat bran plus 0.5% Tween 80 (+WB +Tw80).
The data points correspond to mean values of duplicate assays. ..................................... 84
Figure 5.19 Effect of pH on Lac activities at 50oC in cultures supplemented with 4.5 g/L
wheat bran (+WB) and 4.5 g/L wheat bran plus 0.5% Tween 80 (+WB +Tw80). The data
points correspond to mean values of duplicate assays. .................................................... 85
Figure 5.20 Effects of (A) temperature and (B) pH on Lac activities after dilution for
cultures supplemented with 4.5 g/L wheat bran plus 0.5% Tween 80 (+WB +Tw80). ... 86
Figure 5.21 Glucose consumption (g/L) and Lac activity (U/L) for T. versicolor cultures in
different media and agitation conditions: (A) continuous agitation and (B) agitated every
6 hours for 30 minutes, both at 30oC and 130 rpm........................................................... 88
Figure 5.22 A446 and A254 for cultures in the presence of 500 mg C/L NOM, and 500 mg
C/L NOM plus Tween 80, agitated continuously at 30oC and 130 rpm........................... 89
Figure 5.23 Reduction in A446 and A254 at different pH....................................................... 91
Figure 5.24 Reduction in A446 and A254 at different temperatures....................................... 92
Figure 5.25 History plot for T. versicolor cultures containing 100 mg C/L NOM,
supplemented with 4.5 g/L wheat bran and 0.5% (v/v) Tween 80, Waksman medium
with 5 g/L initial glucose, incubated at 30oC and 130 rpm. ............................................. 93
Figure 5.26 NOM removals (as mg, converted from A446 and A254), and glucose
consumption in T. versicolor cultures with the two supplements and different NOM
concentrations, plug inoculum.......................................................................................... 94
Figure 5.27 A446 for 100, 600 and 700 mg C/L NOM incubated in Waksman medium (5
g/L glucose) containing 4.5 g/L wheat bran in the absence of fungus, as controls.......... 95
Figure 5.28 Activity of the extracellular ligninolytic enzymes of T. versicolor in cultures
containing the two supplements and different NOM concentrations, plug inoculum. ..... 96

xii
List of Tables
Table 2.1 The various forms of NOM in different environments........................................... 7
Table 2.2 Elemental compositions of some examples of different organic matter. ........... 8
Table 2.3 Composition of NOM fractions............................................................................ 15
Table 2.4 Production of ligninolytic enzymes by SSF. ........................................................ 21
Table 3.1 Characterisation of MIEXTM NOM concentrates. ................................................ 33
Table 3.2 Composition of Waksman medium agar slants. ................................................... 34
Table 3.3 Composition of growth media. ............................................................................. 35
Table 4.1 The C:N ratios of the different media used. ......................................................... 55
Table 5.1 Comparative ligninolytic enzyme activities in different culture media. .............. 83
Table 5.2 Different medium contents used for T. versicolor cultures for testing effect of
agitation. ........................................................................................................................... 87
Table 5.3 Na2HPO4-citric acid buffer formulation. .............................................................. 90

xiii
SUMMARY
Natural organic matter (NOM), a complex mixture of organic compounds, influences drinking
water quality and water treatment processes. The presence of NOM is unaesthetic in terms of
colour, taste and odour, and may lead to the production of potentially carcinogenic
disinfection by-products (DBPs), as well as biofilm formation in drinking water distribution
systems. Some NOM removal processes such as coagulation, magnetic ion exchange resin
(MIEXTM) and membrane filtration produce sludge and residuals. These concentrated NOM-
containing sludges from alum precipitation, membrane treatment plants and MIEX
regeneration must therefore be treated prior to disposal.

The white-rot fungi possess a non-specific extracellular oxidative enzyme system composed
of lignin peroxidase (LiP), manganese-dependent peroxidase (MnP) and laccase (Lac) that
allows these organisms to mineralise lignin and a broad range of intractable aromatic
xenobiotics. Rojek (2003) has shown the capability of Phanerochaete chrysosporium ATCC
34541 to remove 40-50% NOM from solution, however, this was found to be mainly due to
adsorption and to be a partially metabolically linked activity. Consequently, the
bioremediation of NOM wastes by selected white-rot fungi was further investigated in the
present study.

The P. chrysosporium seemed to preferentially remove the very hydrophobic acid (VHA)
fraction, and so was most effective for a NOM preparation with a high proportion of
hydrophobic content (and so high in colour and specific UV absorbance (SUVA)). The extent
of NOM decolourisation by P. chrysosporium in three growth media with different C:N ratios
followed the trends: Waksman (C:N = 6) > Fahy (C:N = 76) > Fujita medium (C:N = 114),
such that the lower the C:N ratio, the greater NOM removal. This was consistent with the
findings of Rojek (2003), who used a different NOM preparation and demonstrated that the
removal of NOM increased with decreased C:N ratio (1.58-15.81).

As removals of NOM with P. chrysosporium ATCC 34541 were low, and little
biodegradation occurred, this organism was compared with P. chrysosporium strain ATCC
24725, Trametes versicolor ATCC 7731, and three strains of yeast (Saccharomyces species
arbitrarily denoted 1, 2 and 3). T. versicolor gave the greatest removal (59%) which was
attributed largely to degradation, whereas the NOM removal by the two strains of P.
chrysosporium (37%) and the yeast was predominantly due to adsorption as indicated by the

1
deep brown colouration of the biomass. Saccharomyces sp. 1, 2 and 3 removed 12%, 61% and
23% of the colour, respectively. Although Saccharomyces sp. 2 had similar high colour
reduction to T. versicolor, the specific removal values differed markedly: 0.055 compared to
0.089 mg NOM/mg biomass, respectively. The low level of the ligninolytic enzymes secreted
by both strains of P. chrysosporium corresponded with the low degree of NOM removal by
biodegradation as shown by high performance size exclusion chromatography (HPSEC). The
high NOM removal attained by T. versicolor was attributed to the activities of the ligninolytic
enzymes, especially laccase. The NOM removal was attributed to the breakdown of the high
molecular weight compounds to form a pool of low molecular weight materials, which were
then most likely utilised by the T. versicolor.

Growth of T. versicolor cultures at 36oC caused inhibition or denaturation of the activity of

the phenoloxidase enzymes compared to those grown at 30oC. The low activity of LiP in both
cultures suggested that this enzyme may not play much of a role in NOM removal. The higher
levels of MnP and Lac activities at 30oC were responsible for the greater NOM removal (73%
vs. 59%) and thus the cleavage of aromatic rings, conjugated and Cα-Cβ bonds in phenolic
moieties, as well as catalysing alkyl-aryl cleavage in the NOM structures.

T. versicolor cultured in Waksman medium with higher initial glucose (5 g/L cf. 2 g/L) led to
lower ligninolytic enzyme activities and a lower degree of NOM removal (25% less colour
reduction), probably due to preferential use of glucose over NOM as carbon source. NOM
removal (mg removed) increased linearly with NOM concentration up to 600 mg C/L (62 mg
(A446); 31 mg (A254)), above which removal decreased markedly. This trend coincided with
increasing total ligninolytic enzyme activity, where the level of Lac increased up to 600 mg
C/L NOM although MnP decreased gradually across the range while LiP was only detected
for 100 and 300 mg C/L NOM. Hence, the removal of NOM from solution by T. versicolor
was associated with high oxidative enzyme activity, particularly of laccase.

Laccase was the major extracellular enzyme secreted by T. versicolor and by deduction,
played a major role in NOM removal. The optimum temperature for Lac activity secreted by
T. versicolor cultured in Waksman medium supplemented with 4.5 g/L wheat bran plus 0.5%
Tween 80 was determined to be 50oC. The optimum pH for the Lac activity for guaiacol and
NOM was identified as pH 4.0-4.5. Although the optimum enzyme activity occurred at 50oC,
30oC was recommended for enzymatic removal of NOM as the phenoloxidase enzyme
activity may be denatured if the NOM removal process were considered to run for long period

2
at high temperature. Although agitation led to apparent enzyme denaturation, fermentations
with continuous agitation promoted enzyme activity faster than those with occasional
agitation (agitated every 6 hours for 30 minutes at 130 rpm and 30oC) as it provides better
mass transfer. However, it seemed that continuous agitation had an adverse effect on the
fungal growth and enzyme production over extended fermentation periods.

Addition of 4.5 g/L wheat bran to modified Waksman medium in the absence of NOM led to
high production of Lac activity compared with LiP and MnP activities, showing its great
potential as a laccase inducer. Addition of Tween 80 alone to the cultures led to a small
improvement in Lac activity; however, with the presence of wheat bran it caused marked
increases in LiP, MnP and Lac activities. When NOM was added to cultures of T. versicolor
with the two supplements, it led to markedly reduced Lac activity, but increased LiP and MnP
activities, and no improvement in NOM removal compared with the cultures in the absence of
supplements (12 mg (or 61%) cf. 15 mg (or 73%) for 100 mg C/L after corrected for colour
from and adsorption by wheat bran).

3
List of Publications
Monn K Lee, Roddick, FA and Harris, J 2004, ‘Application of white-rot fungi or yeast for the
biodegradation of natural organic matter in wastes’, The 12th Annual RACI Analytical and
Environmental Division, University of Melbourne, Melbourne, December 5-8, 2004, Poster
Presentation.

Monn K Lee, Solarska, S, Roddick, FA and Harris, J 2004, ‘Application of white-rot fungi for
the biodegradation of natural organic matter’, in Environmental Research Event, The 9th
Annual Environmental Postgraduate Conference, Hobart, Tasmania, November 29-December
2, 2005, Submitted for ERE paper.

4
Chapter 1 Introduction
The importance of maintaining high quality natural surface water sources, on both national
and global scales, is well known as a major requirement for public supplies as well as for
future industrial growth. The presence of natural organic matter (NOM) in raw water reduces
drinking water quality and interferes with water treatment processes. NOM, a complex
mixture of organic carbon compounds, is undesirable for several reasons, including its
contribution to taste and odour of the water, as well as to the formation of disinfection by-
products (DBPs) when NOM reacts with disinfectants such as chlorine. Many
epidemiological studies have suggested that DBPs are carcinogenic and thus present a health
risk to the consumer.

As disinfection remains important in public health protection because of its effectiveness in

maintaining pathogen-free water and providing residual protection to control bacterial
regrowth in the distribution system, its application in drinking water treatment is crucial
(Carraro et al. 2000). Therefore, there has been considerable research focused on the
development of more economic and efficient drinking water treatments with improved NOM
removal prior to chemical disinfection. There are many NOM removal processes available,
such as coagulation, granular activated carbon (GAC) adsorption, ozonation, magnetic ion
exchange resin (MIEXTM), and membrane filtration. However, these processes have been
identified as having high operating cost and producing sludge and residuals, some of which
contain NOM. These wastes can be problematic regarding disposal, as government legislation
concerning the release of wastes has become more stringent.

Bioremediation technology is viewed as an attractive approach to the removal of NOM as this

environmentally friendly method is potentially more cost-effective, and limits by-product
formation and associated mutagenic product generation. This process would be able to
remove biodegradable organic matter (which is responsible for microbial regrowth in the
distribution systems), increase disinfectant stability in the water delivery network, as well as
reduce chlorine demand (Carraro et al. 2000). Furthermore, this biological treatment could be
applied in drinking water treatment as well as for the treatment of concentrated NOM wastes
such as those found in alum precipitation, regenerant wastes from the MIEXTM process, and
retentates from membrane treatment plants.

5
Bioremediation utilises living organisms such as bacteria or fungi, or isolated enzyme
systems, to break down organic pollutants and transform them into harmless products or
valuable by-products (Burton et al. 1999). Previous work has shown that white-rot fungi are
able to degrade lignin and a wide variety of recalcitrant organic pollutants due to their non-
specific extracellular oxidative enzyme system which may include lignin peroxidase (LiP),
manganese-dependent peroxidase (MnP) and laccase (Lac) (Kirk & Chang 1975, Bumpus &
Aust 1987, Lonergan 1992, Barr & Aust 1994, Nyanhongo et al. 2002). Phanerochaete
chrysosporium ATCC 34541 has been shown to remove 40-50% NOM from solution,
however, this was found to be mainly due to adsorption and to be partially metabolically
linked (Rojek et al. 2004). They found that environmental conditions such as carbon and
nitrogen content, pH and NOM concentration played an important role in the removal of
colour by the fungus. In addition, Rojek (2003) found that a combination of yeast
contaminants isolated from a MIEX concentrate with the P. chrysosporium gave NOM
removals of 70-80%.

The research herein builds on the work of Rojek (2003) and further investigates the
bioremediation of NOM wastes. The aim of this research was to evaluate the effectiveness of
selected white-rot fungi and yeast for the biodegradative removal of concentrated NOM from
solution. The effectiveness of the white-rot fungus P. chrysosporium ATCC 34541 for the
biological degradation of different batches of MIEXTM NOM concentrate and the effects of
different characteristics of the NOM on the process and thus the resistance of the NOM to
microbial removal were investigated. The effects of the C:N ratios in three different simple
culture media inoculated with the fungus were studied. Three white-rot fungus and three yeast
strains were tested to identify the most suitable organism for the bioremediation of the
concentrated NOM from solution.

The effects of environmental conditions namely incubation temperature, carbon source level,
type of inoculum and different initial NOM concentrations on the fungal growth and the
production of the extracellular ligninolytic enzymes to enhance NOM removal by Trametes
versicolor were determined. This was followed by the investigations of enhancement of
enzyme production by studying the effects of supplements, temperature, pH and agitation.
The enzymatic treatment of the concentrated NOM in vitro by T. versicolor was further
examined to improve the NOM removal by determining the optimum temperature and pH.
The biological treatment of NOM with the presence of two supplements was investigated to
enhance ligninolytic enzyme induction, and hence improve NOM degradation.

6
Chapter 2 Literature Review
This chapter critically reviews literature relevant to the present research topic: the nature of
natural organic matter (NOM), mechanisms of lignin degradation by white-rot fungi and
applications of their enzymes, as well as a review of the potential applications of white-rot
fungi for the removal of NOM from wastes arising from drinking water treatment.

2.1 Natural Organic Matter

2.1.1 Origin
Natural organic matter (NOM) varies in structure, binding and state of solubility in the soil
and aqueous phase of the terrestrial and aquatic environments (Table 2.1) (Kördel et al. 1997).
The total carbon contents of humic substances in soils (soil organic matter) and marine
environments (marine humus) are 2500 x 1012 kg and 3000 x 1012 kg, respectively (Kördel et
al. 1997).

Table 2.1 The various forms of NOM in different environments.

Terrestrial Environment Aquatic Environment
Solid phase of soils Surface water
Soil water • Freshwater
• Interstitial water • Sea water
• Drain and seepage water Sediment
Groundwater • Solid phase
• Interstitial water

NOM, a complex mixture of organic carbon, can comprise as much as 90% of the total
reduced carbon in aquatic ecosystems (Frazier et al. 2002). NOM can be categorised into
dissolved (DOC) and particulate (POC) organic carbon. Aquatic humic substances contribute
about 40-60% of DOC and comprise the largest fraction of NOM in waters (Kördel et al.
1997). The NOM load in ecosystems is formed from allochthonous and autochthonous
sources (Frazier et al. 2002, Hood et al. 2003). Allochthonous DOC (derived from surface
and subsurface leaching of vegetation and soils in the surrounding catchment) is typically
enriched in fulvic acids and is highly aromatic and coloured, while autochthonous DOC
(derived from algal and bacterial biomass in aquatic systems) is characterised by a lower
fulvic acid content and C:N ratio (McKnight et al. 1994, Hood et al. 2003).

7
2.1.2 Composition and chemical structure
NOM mainly comprises carbon, oxygen, hydrogen and nitrogen. Table 2.2 shows the
elemental compositions of some examples of different organic matter (Weber 2001).

Table 2.2 Elemental compositions of some examples of different organic matter

(Weber 2001).
% Dry ash-free basis
Substances C H O N
Fulvic acids 44-49 3.5-5.0 44-49 2.0-4.0
Humic acids 52-62 3.0-5.5 30-33 3.5-5.0
Proteins 50-55 6.5-7.3 19-24 15.0-19.0
Lignin 62-69 5.0-6.5 26-33 -

Most NOM is comprised of a range of hydrocarbon compounds, from small hydrophilic acids,
proteins and amino acids to larger humic and fulvic acids. The organic components found in
NOM can range from largely aliphatic to highly aromatic (coloured), from highly charged to
uncharged, and from apparent molecular weights around 10,000 down to 200 Dalton. Most
characterisation studies reported that NOM in surface water, on average, showed a significant
charge due to carboxylic acid groups, and some aromatic/hydrophobic character (Croué et al.
2000).

Figure 2.1 shows a general hydrocarbon structure of NOM that displays the diversity of
moieties which appear in NOM (Shevchenko & Bailey 1996). It is not surprising that NOM is
a complex mixture and structure of organic compounds with a great range of attached
functional groups (amide, carboxyl, hydroxyl, ketone and various minor functional groups),
given the wide variety of NOM sources, seasonal variations, and the numerous degradation
and transformation mechanisms which affect NOM (Leenheer & Croué 2003).

8
Figure 2.1 General structure of NOM (Shevchenko & Bailey 1996).

NOM consists of non-humic (hydrophilic) and humic (hydrophobic) substances (Hood et al.
2003). The non-humic hydrophilic fractions are composed predominantly of well-defined
chemical structures such as hydrophilic organic acids and low molecular weight compounds
(carbohydrates, carboxylic acids, amino acids, lipids, proteins etc), which are easily attacked
by micro-organisms (Motheo & Pinhedo 2000).

On the other hand, humic substances, which originate from microbial or chemical conversion
of bacteria, plants and other living organism residues, are naturally occurring heterogeneous
organic substances that are based on N-containing polymers. Humic substances comprise both
aliphatic and aromatic high molecular weight components (Liao et al. 1982). They have a
complex chemical structure with no defined chemical and physical properties, which are
generally refractory to attack by micro-organisms (Motheo & Pinhedo 2000). Furthermore,
they are categorised into three classes based on their solubility characteristics: humin, fulvic
acid and humic acid (Weber 2001) (Figure 2.2).

9
Figure 2.2 Classification of NOM (http://www.humintech.com).

(i) Non-humic substances

Aiken et al. (1992) compared the characteristics of hydrophobic and hydrophilic acids
isolated from different environments. The hydrophilic organic acids had lower carbon and
hydrogen contents, higher oxygen and nitrogen contents, and were lower in molecular weight
than hydrophobic organic acids. Moreover, the hydrophilic acids had a lower concentration of
aromatic carbon and greater hetero-atom, ketone and carboxyl content than the fulvic acid
(Aiken et al. 1992).

Other common types of non-humic substances found are carbohydrates. Carbohydrates

constitute about 5-25% of organic matter in soils. They are divided into three subclasses:
monosaccharides, oligosaccharides and polysaccharides, which are aldehyde and ketone
derivatives of the higher polyhydric alcohols (Weber 2001).

(ii) Humic substances

Humic acid is not soluble in water under acidic conditions (pH< 2). Humic acids are the
major extractable component of soil humic substances and are dark brown to black in colour.
The fraction called fulvic acid is soluble in water under all pH conditions and remains in
solution after removal of humic acid by acidification. Fulvic acid is light yellow to yellow-

10
brown in colour. Humins are soluble neither in bases nor in acids and are black in colour
(Kördel et al. 1997, Weber 2001).

Figure 2.3 Chemical properties of humic substances (http://www.humintech.com).

These three fractions vary considerably in colour, C-, O- and N-contents. The differences
between humic acid and fulvic acid can be described by disparity in molecular weight,
numbers of functional groups (carboxyl, phenolic OH) and extent of polymerisation. The
molecular weight distributions for aquatic fulvic acid and humic acid were reported to be
between 500 and 2000 Dalton and between 2000 and 5000 Dalton, respectively (Howe &
Clark 2002). The postulated relationships are represented in Figure 2.3, in which colour
intensity, degree of polymerisation, carbon and oxygen contents, and acidity vary with
increasing molecular weight. The high molecular weight humic acids have lower oxygen but
higher carbon contents than the low molecular weight fulvic acids. Fulvic acids, which
contain more functional groups of an acidic nature, especially COOH, have significantly
higher acidities than humic acids (900-1400 meq/100 g cf. 400-870 meq/100 g) (Kördel et al.
1997, Weber 2001).

Humic acids are complex aromatic macromolecules with amino acids, amino sugars, peptides
and aliphatic compounds involved in linkages between the aromatic groups. The structure of
humic acid, shown in Figure 2.4, contains quinone structures, phenolic OH groups, nitrogen
and oxygen as bridge units and COOH groups diversely located on aromatic rings (Weber
2001).

11
Another study characterised the general structure of aquatic humic acids and reported that
they may consist of (a) single aromatics rings with mainly three to six substituents as alkyl
side chain, carboxylic acid, ketone or hydroxyl groups, (b) short aliphatic carbon chains, and
(c) polycyclic ring structures including polynuclear aromatics, polycyclic aromatic-aliphatic
and fused rings involving furan and possibly pyridine (Liao et al. 1982).

COOH COOH
OH HC O
COOH H
(HC.OH)4 sugar OH
O O
R CH
H HC O COOH
O
O N O
O O H2
HO
CH C

OH OH O O O COOH
CH

N O

O NH

R CH OH
O
C O peptide
NH

Figure 2.4 Model structure of humic acid (Redrawn from http://www.humintech.com).

Fulvic acids are identified as a heterogeneous mixture of medium molecular weight (400-
2000 Dalton) yellow organic acids (McKnight et al. 1994). They are the most hydrophilic of
humic substances, contain a wide variety of aromatic and aliphatic structures, and both are
extensively substituted with oxygen-containing functional groups, particularly –COOH, –OH
and C=O (Kördel et al. 1997, Weber 2001) (Figure 2.5).

OH COOH CH 2OH
CH 3
HOOC CH 2 CH

C CH

O CH 2 COOH
HOOC CH 2 CHOH

COOH C COOH
OH CH 2

Figure 2.5 Model structure of fulvic acid (Redrawn from http://www.humintech.com).

12
2.2 Characterisation of NOM
Characterisation of NOM can be divided into two categories: studies of (a) whole water,
where DOM is characterised in water and its inorganic constituents, and (b) DOM fractions
isolated from water and inorganic constituents (Leenheer & Croué 2003).

2.2.1 Whole water characterisations

(i) TOC, DOC and BOM analyses

Total organic carbon (TOC) is the common measurement used to represent the NOM
concentration in aquatic systems. Particulate organic matter (POC) is the organic carbon that
is retained on a 0.45-μm-porosity membrane, while dissolved organic matter (DOC) is the
fraction of the TOC smaller than 0.45 μm in diameter (Leenheer & Croué 2003).

According to Leenheer and Croué (2003), most NOM is considered to be refractory to rapid
biodegradation but autochthonous NOM is more biodegradable than allochthonous NOM.
Biodegradable organic matter (BOM) can be measured based on standard protocols such as
biodegradation over a given time and is expressed as biodegradable dissolved organic carbon
(BDOC) or assimilable organic carbon (AOC).

(ii) Spectrophotometric analysis

Absorbance at 446 nm (A446) is generally used as a measurement for colour whereas
absorbance at 254 nm (A254) is an indicator of the presence of UV-absorbing components in
NOM. Leenheer and Croué (2003) reported that the aromatic chromophores present in NOM
molecules (particularly humic substances) absorb both visible and UV light.

DOC is a general indicator of the NOM content in a water sample however it does not give
specific information on the potential for NOM to serve as disinfection by-product (DBP)
precursors. Specific UV absorbance (SUVA), the ratio of A254 to DOC, is used to represent
the enrichment of DOC in DBP precursors and as a measure of the aromaticity of the DOC
(White et al. 2003). High SUVA waters suggest that the samples would likely result in DBP
formation and are generally enriched in hydrophobic NOM, such as humic substances
(Leenheer & Croué 2003). In addition to absorbance, fluorescence is also used to characterise
NOM. Humic-type molecules are considered to be largely responsible for the fluorescence
observed in natural waters (Leenheer & Croué 2003).

13
(iii) Size characterisation of NOM
Sequential ultrafiltration fractionation is used for low-resolution separations while size
exclusion chromatography (SEC) is employed for higher-resolution size separations
(Leenheer & Croué 2003). However, neither sequential ultrafiltration fractionation nor SEC
gave absolute measures of the molecular weight of NOM (Schäfer et al. 2002).

Chin et al. (1994) suggested that number- (Mn) and weight-averaged (Mw) molecular weights
for the humic substances could be determined by employing high performance size exclusion
chromatography (HPSEC). This technique requires relatively small sample volumes and can
be used with many samples without pre-concentration, thus allowing determinations of both
molecular size and weight on whole-water samples. It also gives significant understanding of
the nature of chemical interactions at the molecular level between dissolved organic carbon
and other organic constituents (Chin et al. 1994).

2.2.2 Fractionation
DOM is generally characterised by isolating NOM into distinct fractions using resin sorbents
(Leenheer & Croué 2003). The analytical method of DOC fractionation separates NOM into
humic (hydrophobic acids, bases and neutrals) and non-humic (hydrophilic acids, bases and
neutrals) substances based upon their adsorption on non-ionic and ion-exchange resin
adsorbents (Leenheer 1981, Thurman & Malcolm 1981).

Chow et al. (2004) further developed a rapid fractionation system by employing DAX-8,
XAD-4 and IRA-958 resins to obtain very hydrophobic acids (VHA), slightly hydrophobic
acids (SHA), hydrophilic charged (CHA) and hydrophilic neutral (NEU) compounds. Table
2.3 provides a list of organic compounds classified according to different NOM fractions
(Barber et al. 2001, Swietlik et al. 2004).

14
Table 2.3 Composition of NOM fractions (Barber et al. 2001, Swietlik et al. 2004).
Fraction Organic compound class Reference

Humic substances or hydrophobic fractions (VHA + SHA)

Hydrophobic Soil fulvic acids; C5-C9 aliphatic carboxylic Leenheer (1981)
acid acids; 1- and 2-ring aromatic carboxylic Aiken et al. (1992)
acids; 1- and 2-ring phenols; linear alkyl Marhaba et al. (2000)
benzene sulphonate (LAS); LAS degradation Barber et al. (2001)
products
Hydrophobic Portion of humic substances retained by Leenheer (1981)
base XAD-8 resin at pH 7 which can be eluted by Marhaba et al. (2000)
HCl; 1- and 2-ring aromatics amines except Barber et al. (2001)
pyridine, proteinaceous substances, cationic
surfactants
Hydrophobic Hydrocarbons; >C5 aliphatic alcohols, Leenheer (1981)
neutral amides, esters, ketones, aldehydes; long chain Marhaba et al. (2000)
(>C9) aliphatic carboxylic acids and amines; Barber et al. (2001)
>3-ring aromatic carboxylic acids and amines;
chlorophyll and related pigments; LAS and
optical brighteners

Non-humic substances or hydrophilic fractions (CHA + NEU)

Hydrophilic <C5 aliphatic carboxylic acids; polyfunctional Leenheer (1981)
acid carboxylic acids; mixture of various hydroxy Aiken et al. (1992)
acids; LAS degradation products Marhaba et al. (2000)
Barber et al. (2001)
Hydrophilic Amphoteric proteinaceous materials Leenheer (1981)
base containing aliphatic amino acids, amino Marhaba et al. (2000)
sugars, peptides and proteins; <C9 aliphatic Barber et al. (2001)
amines; pyridine
Hydrophilic <C5 aliphatic amides, alcohols, aldehydes, Leenheer (1981)
neutral esters, ketones; polyfunctional alcohols; Marhaba et al. (2000)
carbohydrates; cyclic amides; polysaccharides Barber et al. (2001)

15
2.3 Impact of NOM on Water Quality and Treatment
A few decades ago research into the nature of NOM in potable water and methods for its
removal was predominantly focused on removing colour and turbidity from public water
supplies. Since then, several other problems associated with NOM have arisen in the
treatment and delivery of drinking water. These include its demand for coagulants and
disinfectants, its tendency to foul membranes and to interfere with the removal of other
contaminants, as well as its potential to transport metals and hydrophobic organic chemicals
and thus causing pipe corrosion in distribution systems (Jacangelo et al. 1995).

The presence of NOM in water is known to have adverse impacts on water treatment
processes including coagulation, membrane filtration, GAC and disinfection. NOM can
reduce the effectiveness of flocculation, which in turn increases the demand for coagulants
due to its reactions with coagulants. Many studies have suggested that NOM is the most
important membrane foulant as some DOC can penetrate and clog microfiltration and
ultrafiltration membrane pores and thus contribute to fouling (Carroll et al. 2000, Howe &
Clark 2002). NOM can also compete with other contaminants such as algal toxins and
pesticides for adsorption sites on activated carbon and consequently block GAC pores. The
fractions of NOM that cannot be successfully removed by conventional drinking water
treatment can react with disinfectants to form DBPs, especially trihalomethanes (THMs) and
haloacetic acids (HAAs), some of which are potentially carcinogenic in addition to being
mutagenic (Carraro et al. 2000, Aoustin et al. 2001). Furthermore, NOM can serve as an
electron donor in metal complexation, sorption of xenobiotics and adsorption on to mineral
phases and on to activated carbon (Frimmel 1998).

Another concern of water utilities is the contribution of NOM as a substrate for bacterial
regrowth (i.e., formation of biofilms) in drinking water distribution systems. This can lead to
undesirable tastes, odour, particles, as well as the increased risk of gastrointestinal illness for
drinking water consumers (Prévost et al. 1998). As a result of the deleterious effects of NOM
on water quality, removal of NOM from drinking water is critical before delivery to the
consumers. However, application of some treatments results in the generation of concentrated
NOM wastes, such as sludges from alum precipitation, membrane treatment plants and MIEX
regeneration, which must be treated prior to disposal.

16
2.4 White-rot Fungi
White-rot fungi belong to the wood-destroying basidiomycetes and are best known as the only
micro-organisms responsible for the mineralisation of all major wood polymers, including
lignin, cellulose and hemicellulose (Crawford & Crawford 1980). Several species have been
investigated for their lignin-degrading capability, such as Phanerochaete chrysosporium,
Coriolus versicolor (now called Trametes versicolor), Pleurotus ostreatus, Phlebia radiata,
Ceriporiopsis subvermispora, Panus tigrinus and Bjerkandera adusta. The mechanisms for
their wood biodegradation ability are dependent on the fungal species and conditions, e.g., T.
versicolor degrades wood by simultaneous attack on both lignin and polysaccharides while C.
subvermispora preferentially degrades lignin (Del Pilar Castillo 1997).

Research on the lignin-degrading enzyme system led to the detection of extracellular

phenoloxidase enzymes in P. chrysosporium (Glenn et al. 1983, Tien & Kirk 1983, Kuwahara
et al. 1984). The ligninolytic system of white-rot fungi is considered to contain a pool of
enzymes, particularly lignin peroxidase (LiP), manganese-dependent peroxidase (MnP) and
laccase (Lac), which are highly effective in oxidising and cleaving wood and lignin (natural
components of the ecosystem), as well as various intractable xenobiotic pollutants structurally
similar to lignin. Applications of the system present many potential advantages, such as the
organisms do not have to adapt to the organic pollutants since the lignin-degrading system is
triggered by the absence of a single nutrient, and the extracellular oxidative enzymes are not
dependent upon the concentration of the pollutants (Del Pilar Castillo 1997); and the system is
not specific and so allows the mineralisation of a broad range of persistent organic pollutants
as opposed to bacterial systems which may require separate enzymes to catalyse the breakage
of each bond type (Lonergan 1992). Moreover, the system is also recognised as a safe,
versatile and economic biological treatment which is able to bio-transform hazardous
compounds including xenobiotics; reduce ammonia, iron and manganese levels; as well as
bio-oxidise assimilable organic carbon (AOC) that allows production of biologically stable
water (Carraro et al. 2000).

White-rot fungi apply oxidative mechanisms for pollutant degradation and the organo-
pollutants are metabolised through cycles of oxidation and subsequent quinone reduction
reactions, leading to intermediates which undergo aromatic ring cleavage (Gold & Alic 1993).
These allow the white-rot fungi to oxidise a wide range of recalcitrant organic compounds, for
instance polyaromatic hydrocarbons (PAHs) (Muncnerova & Augustin 1994), polycyclic

17
aromatic hydrocarbons (Bogan & Lamar 1996), polychlorinated biphenyls (Xu 1996),
polychlorinated dibenzo(p)dioxins, and the pesticides DDT and lindane (Fujita et al. 2002). It
has also been reported that P. chrysosporium ATCC 24725 is able to mineralise di-, tri-, tetra-
and penta-chlorophenol (PCP) (Choi et al. 2002, Shim & Kawamoto 2002).

The control of culture parameters is important for optimum ligninolytic activity.

Understanding the physiological bases of culture requirements will facilitate investigation of
the extracellular ligninolytic enzyme system. It has been reported that factors such as
temperature, pH, supplements (veratryl alcohol (VA), Tween 80 and lignocellulose residues),
as well as agitation conditions affect the secretion of enzymes and thus the performance of
white-rot fungi.

2.4.1 Effect of temperature and pH

Many researchers have investigated the effect of both temperature and pH on white-rot fungi
for the bioremediation of different compounds. The effect of both factors varied when
different substrates were investigated. It was reported that the optimum temperature and pH
for an unidentified basidiomycete wood-rotting fungus used for the decolourisation of cotton
bleaching effluent was found to be 27oC and pH 4-5, respectively (Zhang et al. 1999).
Another study stated that the optimum pH for textile dyestuff decolourisation by C. versicolor
MUCL was pH 4.5 and the fungus was capable of decolourising the dyestuff with lower
efficiencies at pH 6 and 7 (Kapdan et al. 2000). Schliephake et al. (2000) determined the
optimum temperature for a purified laccase from Pycnoporus cinnabarinus CBS 101046
during the degradation of the diazo dye Chicago Sky Blue. They established that the laccase
was stable at 60oC for one hour and remained active in bioreactors at 37oC for 25 days
(Schliephake et al. 2000).

The effect of temperature on textile dye decolourisation by Trametes modesta, investigated by

Nyanhongo et al. (2002), showed that different dyes were decolourised at different rates at
different temperatures. They reported that the rate of dye decolourisation due to laccase
increased with temperature up to 50-60oC, after which it decreased (Nyanhongo et al. 2002).
A catechol polymerisation study catalysed by laccase from T. versicolor ATCC 200801
demonstrated that the optimum pH for the oxidative process was pH 5 and the reaction rate
increased exponentially with temperature up to 45oC, after which the rate tended to plateau
(Aktas & Tanyolac 2003). Rancaño et al. (2003) reported that pH 5 was optimal for the

18
decolourisation of Phenol Red by laccase from T. versicolor CBS 100.29.

Dodor et al. (2004) worked with laccase from T. versicolor and observed that the rate of
oxidation of anthracene and benzo[α]pyrene increased with increasing temperature up to
40oC, after which the rate started to decrease. Another group of researchers found that the
optimum temperature and pH for the enzymatic decolourisation of Remazol Brilliant Blue R
(RBBR) by Funalia trogii ATCC 200800 growing in a solid-state fermentation (SSF) medium
containing wheat bran and soybean waste were 50oC and pH 3.0, respectively (Deveci et al.
2004).

2.4.2 Effect of supplements

There have been many investigations of the potential of different supplements (veratryl
alcohol (VA), Tween 80 and lignocellulose residues) for inducing the production of the
ligninolytic enzymes and thus improving the performance of white-rot fungi. It was reported
that VA (3, 4-dimethoxybenzyl alcohol) is capable of stabilising LiP activity (Tsai 1991) and
acts as a substrate for LiP (Feng et al. 1996). It is also believed that VA works as a redox
mediator to facilitate the oxidation of chemicals (Harvey et al. 1986), has a protective effect
against inactivation of certain LiP isoenzymes by H2O2 (Wariishi & Gold 1989), and has the
ability to enhance the action of LiP on many substrates, including lignin (Hammel et al.
1993). The mediator concept is shown in Figure 2.6 for LiP catalysed oxidation of lignin.

Figure 2.6 Veratryl alcohol as electron transfer mediator (Redrawn from Harvey et al.
1986).

Tween 80 (polyoxyethylene sorbitan monooleate), a non-ionic surfactant, is able to transform

the cell membrane structure and promote permeation of LiP from the cell into the medium
(Asther et al. 1987). Tween 80 can also protect LiP in culture fluids against mechanical

19
inactivation due to agitation (Venkatadri & Irvine 1990). They determined that the presence
of Tween 80 caused a 1.3- to 1.4-fold increase in LiP enzyme activity, even under stationary
conditions. Asther and co-workers (1987) explained that the saturated and unsaturated fatty
acids, released by the hydrolysis of Tween 80 by most fungi, are involved in the activation of
LiP production, either by providing an extracellular energy source for secondary metabolism
or by serving as an inducer. However, the addition of Tween 80 as a culture supplement
caused significant foaming due to aeration. The foaming caused by aeration is still of concern
although Tween 80 has a protective effect against mechanical inactivation of the enzyme
(Shim & Kawamoto 2002).

Recently, many studies have demonstrated the importance of utilising lignocellulose residues
for enhancing the production of the extracellular phenoloxidase enzymes by white-rot fungi
(Couto et al. 2001, Lorenzo et al. 2002, Couto et al. 2004, Kapich et al. 2004, Couto &
Sanromán 2005). Lignocellulose residues are mainly composed of polysaccharides (cellulose
and hemicellulose) and lignin and occur in a wide range of wastes from the agricultural and
forestry industries. Lorenzo et al. (2002) showed that due to their cellulose content barley
bran, grape stalks and grape seeds have significant potential to improve laccase production in
submerged cultures of T. versicolor CBS 100.29. They investigated the potential enzymatic
decolourisation of Phenol Red by the ligninolytic extracellular fluids obtained in the cultures
with the aforementioned supplements and established that the cultures with barley bran and
grape stalks had high decolourisation ability. The presence of the lignocellulose wastes
provides the fungus an environment similar to its natural habitat (wood), and so may assist the
stimulation of the secretion of the lignin-degrading enzymes.

Kapich et al. (2004) illustrated that the addition of solid lignocellulose-containing substrates
such as hemp woody core and wheat straw to liquid medium led to the production of LiP and
MnP in submerged cultures of P. chrysosporium ME-446. They suggested that the
immobilisation of the mycelium on the surface of the substrates possibly provides a greater
surface area and increases mass transfer and so improves the production of the enzymes.
Moreover, the fungal mycelium may penetrate the lignocellulose substrate releasing
additional water-soluble aromatic/phenolic substances, which in turn may increase the
secretion of the ligninolytic enzymes (Kapich et al. 2004). In their review Couto and
Sanroman (2005) noted that the use of organic wastes rich in lignin was ideal for the
production of LiP, while utilisation of organic wastes rich in cellulose induced laccase
production.

20
Table 2.4 shows the production of ligninolytic enzymes by different micro-organisms using
SSF (Couto & Sanromán 2005).

Table 2.4 Production of ligninolytic enzymes by SSF (Couto & Sanromán 2005).
Support Micro-organisms Ligninolytic Reference
enzymes
Bagasse Polyporus BH1, Polyporus BW1 LiP Nigam et al. (1987)
T. versicolor MnP, Lac Pal et al. (1995)
P. ostreatus, P. chrysosporium LiP, MnP, Lac Pradeep and Datta (2002)
Cotton P. chrysosporium, Funalia trogii LiP, MnP, Lac Sik and Unyayar (1998)
Grape C. versicolor, Panus tigrinus, P. LiP Golovleva et al. (1987)
chrysosporium
Olive mill P. tigrinus MnP, Lac Fenice et al. (2003)
wastewater
Wheat, bean Pleurotus eryngii, Pleurotus MnP Martinez et al. (1996)
pulmonarius
Wheat bran P. pulmonarius Lac De Souza et al. (2002)
P. ostreatus, P. chrysosporium LiP, MnP, Lac Shinners-Carnelley et al. (2002)
Fomes sclerodermeus MnP, Lac Papinutti et al. (2003)
Wheat straw C. versicolor, P. tigrinus, P. LiP Golovleva et al. (1987)
chrysosporium
P. radiata LiP, MnP, Lac Vares et al. (1995)
Pleurotus sp. MnP, Lac Lang et al. (1996)
P. chrysosporium LiP, MnP Castillo et al. (1997)
Fujian et al. (2001)
P. ostreatus Lac Baldrian and Gabriel (2002)
P. pulmonarius Lac De Souza et al. (2002)
Wood Bjerkandera sp. strain BOS55 LiP, MnP Mester et al. (1998)
C. subvermispora MnP, Lac Ferraz et al. (2003)
P. ostreatus, P. chrysosporium LiP, MnP, Lac Pradeep and Datta (2002)

21
2.4.3 Effect of agitation
Early research showed that culture agitation suppressed both LiP production and lignin
degradation in P. chrysosporium cultures (Kirk et al. 1978, Faison & Kirk 1985). It has been
reported that agitation had no apparent effect on LiP formation by P. chrysosporium, but
caused inactivation of the secreted enzyme (Venkatadri & Irvine 1990).

However, other investigations illustrated that LiP can be protected in agitated (200 rpm)
submerged cultures of P. chrysosporium by supplementing with Tween 80 (Jäger et al. 1985).
Lignin degradation has been achieved under agitated conditions (136 rpm), as reported by
Reid et al. (1985). Another group of researchers demonstrated that agitation was very
effective in improving (more than doubling) the rate of Orange II decolourisation by mycelial
pellets of an unidentified basidiomycete fungus. This may be due to the improved mass
transfer of oxygen and substrates in agitated cultures (Knapp et al. 1997).

2.5 Mechanism of Enzymatic Degradation by White-rot Fungi

White-rot fungi play a significant role in the recycling of lignin, capable of completely
mineralising lignin to CO2 and H2O due to their non-specific extracellular ligninolytic enzyme
system. Degradation of the complex irregular aromatic structure of lignin polymer by white-
rot fungi enables them to access cellulose and hemicellulose, which they then utilise as carbon
and energy sources (Leatham 1986, Kirk & Farrell 1987).

A general scheme for lignin biodegradation, which involves the oxidative reactions catalysed
by LiP, MnP and Lac, is shown in Figure 2.7. Kirk and Chang (1975) studied the chemical
changes of fungally degraded lignin and showed that it had decreases in phenolic hydroxyl,
aliphatic hydroxyl and methoxyl contents. Furthermore, higher contents of α-carbonyl and
conjugated carboxyl groups in the degraded lignin were obtained. Another observation was
that the degraded lignin contained oxidised side chains and aliphatic residues resulting from
oxidative cleavage of aromatic rings (Kirk & Chang 1975).

The lignin-degrading enzyme system of white-rot fungi has been widely studied. A number of
white-rot fungi and their enzymes have been used successfully in various configurations in
different types of industrial applications. Since a diverse range of recalcitrant organic
pollutants contain a chemical structure similar to lignin, the ligninolytic system and oxidative
mechanism is considered to be involved in the degradation of the pollutants (Gold & Alic

22
1993) as well as humic acids (Blondeau 1989), and so of naturally occurring organic matter
due to the similarity between the structures of lignin and NOM.

fragments from phenylpropane

Ligninolytic enzymes: side chain
LiP, MnP, Lac
Lignin ring cleavage

quinones

aromatic aldehydes and acids

CO2
aldehyde and acid oxidoreductases
aromatic alcohols
quinone oxidoreductases
hydroquinones

dioxygenases
ring cleavage

Figure 2.7 Proposed schemes for the biodegradation of lignin (Redrawn from Leisola
& Garcia 1989).

Brief descriptions of three major extracellular phenoloxidase enzymes secreted by white-rot

fungi, which are involved in the oxidative degradation mechanism, are described below.

2.5.1 Lignin peroxidase

Lignin peroxidase (LiP) is a glycoprotein that contains one mole of iron protoporphyrin IX as
a prosthetic group. It has a series of isoenzymes with molecular weight of 41,000-42,000
(Kirk & Farrell 1987, Gold et al. 1989). The enzyme can be assayed by the oxidation of
veratryl alcohol (VA) to veratraldehyde at 310 nm (Tien & Kirk 1984). Fungi that were
identified as producing LiP include P. chrysosporium, T. versicolor, Pleurotus ostreatus,
Phlebia tremellosus etc.

LiP has a relatively high redox potential and has no substrate specificity. LiP has been shown
to oxidise phenolic and non-phenolic lignin related compounds as well as a wide variety of
model lignin and related compounds (Barr & Aust 1994). Among the oxidation reactions
catalysed by LiP are the cleavage of the Cα-Cβ and aryl Cα bonds, ring cleavages in β-O-4
compounds, aromatic ring opening, demethylation and phenolic oxidation (ten Have &

23
Teunissen 2001). All of these reactions are involved in the same mechanism in which the
oxidised enzyme intermediates LiP (I) and LiP (II) catalyse the initial one-electron oxidation
of the substrate to yield an aryl cation radical, followed by a series of non-enzymatic reactions
to yield the final products (Gold et al. 1989). This is illustrated in Figure 2.8 for the LiP-
catalysed oxidation of a β-1 diarylpropane dimeric model compound, showing the mechanism
of Cα-Cβ cleavage.

CHO

. H+ H2COH
H2COH H2COH
HC .
HC OMe e-1 HC OMe OC2H5

HCOH HCOH OC2H5

+.
OC2H5 OC2H5 OMe

OC2H5 OC2H5
H2COH H2COH

HCOH HC + e-1
H2O

NR
OMe OMe
H2COH
H2COH
C O .
HC O O
O2

O2
OMe
OMe
2

Figure 2.8 Mechanism of Cα-Cβ cleavage by LiP (Redrawn from Gold et al. 1989).

The catalytic cycle of LiP is illustrated in Figure 2.9 (Gold et al. 1989). The native enzyme
reacts with H2O2 forming the two-electron oxidised intermediate, LiP (I), which then oxidises
the lignin substrate (RH) to produce the one-electron oxidised intermediate LiP (II) and a

24
substrate radical (R.). The LiP (II) is then reduced back to the resting enzyme state by
oxidising a second substrate compound while the free radical can undergo a range of
reactions. With excess H2O2 LiP (II) can be converted to an inactive form of the enzyme LiP
(III) (Tien 1987, Gold et al. 1989, Del Pilar Castillo 1997). However, as mentioned in Section
2.4.2, the presence of VA can protect against inactivation of the LiP by the excess H2O2 as
well as mediate the oxidation of lignin as shown in Figure 2.6.

Figure 2.9 Catalytic cycle of lignin peroxidase (Redrawn from Gold et al. 1989).

2.5.2 Manganese-dependent peroxidase

The discovery of LiP initiated the search for other oxidative enzymes and led to the detection
of manganese-dependent peroxidase (MnP). MnP is also a heme peroxidase and a
glycoprotein. Like LiP they have a family of isoenzymes, containing one iron protoporphyrin
IX group per mole of enzyme with molecular weight approximately 46,000 (Kuwahara et al.
1984). MnP has been shown to degrade a wide range of phenols and dyes (Kuwahara et al.
1984) by oxidising Mn (II) to the oxidant Mn (III) (Kirk & Farrell 1987), which diffuses from
the active site of the enzyme and reacts with different phenolic substrates.

25
The catalytic cycle of MnP is illustrated in Figure 2.10 (Gold et al. 1989). It is basically the
same as for LiP, except that Mn (II) is required to complete the cycle. As shown in Figure
2.10, the resting enzyme reacts with H2O2 to form MnP (I), which is then transformed to MnP
(II) by oxidising one equivalent of Mn (II) to yield Mn (III). A second Mn (II) is then
responsible to reduce MnP (II) back to the resting enzyme. Similarly, the MnP (I) can oxidise
phenolic substrates (RH), but at a slower rate. Phenolic compounds are, however, not capable
of efficiently converting MnP (II) back to the resting enzyme. This may be due to the Fe4+ =
O centre in MnP (II) being partially buried and so the site is not available to organic
substrates. Thus, Mn (II) is essential for the completion of the cycle (Wariishi et al. 1988, Del
Pilar Castillo 1997).

Figure 2.10 Catalytic cycle of manganese peroxidase (Redrawn from Gold et al. 1989).

The oxidation of a free phenolic β-1 lignin dimer by MnP is shown in Figure 2.11. The initial
reaction involves a one-electron oxidation of the phenol to produce a phenoxy radical
intermediate. As a result, dehydrogenation would yield the ketone (B), alkyl phenyl cleavage
of the radical intermediate would form products (C) through (E) and finally, Cα-Cβ would
yield the products (F) through (H) (Gold et al. 1989).

26
 OH
O
OMe

+ + OH

OMe OMe
OMe OMe
O
OH H
O

(C) (D) (E)

Alkyl phenyl cleavage

OMe
OMe
OH
OH

Dehydrogenation
OMe
OMe
O Ketone
OH
(B)
HO
HO

OMe
OMe

H O

OMe OMe

OH
+ OH
+
OMe OMe
O
OH OH

(F) (G) (H)

Figure 2.11 Oxidation of a free phenolic β-1 substructure by MnP (Redrawn from Gold
et al. 1989).

2.5.3 Laccase
Laccase (Lac) is an extracellular glycosylated enzyme and belongs to the copper-containing
polyphenol oxidase family. The enzyme is generally larger than LiP and MnP, having a
molecular weight around 60,000 (ten Have & Teunissen 2001). Laccase is capable of
catalysing the oxidation of phenolic and non-phenolic compounds (Bourbonnais & Paice
1990) and is able to decolourise a wide range of synthetic dyes (Rodriguez et al. 1999,
Swamy & Ramsay 1999). It was reported that fungal laccase worked synergistically with

27
MnP in oxidising phenols and phenolic substructures of lignin through decarboxylation and
subsequent demethylation of methoxy groups (Galliano et al. 1991, Schlosser & Höfer 2002).

Cleavage of Cα-Cβ

Figure 2.12 Proposed degradation of phenolic β-1 model compounds by laccase from
C. versicolor (Kawai et al. 1988).

Kawai (1988) proposed that laccase not only catalyses alkyl-aryl cleavage and Cα oxidation,
but also catalyses Cα-Cβ cleavage of phenolic moieties via phenoxy radicals. The proposed
pathway, which proceeds via the formation of oxygen mediated phenoxy radicals of the
phenolic units by laccase, is illustrated in Figure 2.12.

28
2.6 Potential Applications of White-rot Fungi in Bioremediation
As outlined in Chapter 1, there are several available processes for the removal of NOM in
waste and drinking water treatment industries. However, all have advantages and
disadvantages. Conventional water treatments have been identified as having low removal
efficiency and high operating cost, are mostly based on chemical addition and applicable to a
limited concentration range, as well as producing sludge and residuals (Vickers et al. 1995,
Burton et al. 1999). The NOM-containing wastes generated from alum precipitation,
membrane process plant and the anionic exchange MIEXTM process can be problematic
regarding disposal. Conventional coagulation, which requires the addition of a coagulant such
as alum, ferric salts or polyaluminium chlorides, has low NOM removal efficiency (10-50%)
and generates sludge disposal problems (Jacangelo et al. 1995, Vickers et al. 1995). In
addition, the control of coagulant addition and adjustment in pH are necessary, as these must
be adjusted with any change in the raw water (Jacangelo et al. 1995, Vickers et al. 1995).
Some studies on coagulation suggested that lower molecular weight, hydrophilic, uncharged
and fulvic acid-like components still remain in natural waters after the treatment (Chow et al.
1999, Drikas et al. 2003, Chow et al. 2004). This is consistent with Page et al. (2003), who
established that the proportion of polysaccharide-derived compounds (i.e., of hydrophilic
character) generally increased after alum treatment, indicating that these compounds are
refractory to alum coagulation.

Adsorption by granular or powdered activated carbon (GAC or PAC), which is widely used in
the United States, has limitations since its adsorption capacity is limited and may be
exhausted after a short period. Consequently, frequent reactivation or replacement of activated
carbon is necessary (Jacangelo et al. 1995). Ozonation, which is effective in transformation of
refractory NOM to biodegradable dissolved organic carbon (BDOC), has some limitations.
These include high operating costs and the possibility of mutagen (aldehydes and other
ozonation by-products, e.g. ketones, bromates) formation (Gilli et al. 1990, Kirisits et al.
2001). Furthermore, the maximum BDOC production by ozonation is only approximately
30% of the total DOC in raw water even if the ozone dose is increased, due to the fact that
BDOC may also consume ozone (Volk et al. 1993, Wricke et al. 1996, Nishijima et al. 2003).

Membrane filtration has become an established process in water treatment industry. However,
one of the most significant factors limiting the implementation of membrane filtration is
fouling. Colloidal matter may cause fouling by forming a cake at the membrane surface, while

29
dissolved matter, some of which can penetrate pores, causes fouling by forming a surface
cake, penetrating and clogging pores, or adsorbing within membrane pores to reduce the pore
diameter (Carroll et al. 2000, Howe & Clark 2002). These limitations reduce the effectiveness,
lead to frequent cleaning, and so reduce the lifetime of membranes. Furthermore, membrane
filtration can generate sludge, and thus a sludge disposal problem.

Consequently, reduction of treatment costs and development of energy-efficient waste

treatment processes are needed as much sludge, some of which contains NOM, is generated in
NOM removal processes. Biological treatment can be an option for the breakdown of the
NOM, as the treatment would overcome some problems associated with conventional
treatment processes, including chemical usage and sludge management. Bioremediation
technology is viewed as an attractive approach to the removal of NOM as it is ‘natural’ and a
potentially chemical-free process, which should have good public acceptance, and leads to
minimal waste production.

White-rot fungi are recognised for their ability to degrade lignin and an array of persistent
aromatic pollutants due to their non-specific extracellular ligninolytic enzymes and their
ability to adapt to severe environmental constraints (Lonergan 1992). Del Pilar Castillo (1997)
suggested that since the lignin-degrading system of P. chrysosporium has been shown to
remove pollutants such as pesticides, it could be an option for the treatment of contaminated
farmland. The wastewater effluents of textile, paper, printing and dye industries are highly
coloured and contain toxic aromatic amines (Kapdan et al. 2000). Kapden et al. (2000)
investigated the effects of environmental conditions such as pH, carbon source and dyestuff
concentration on the textile dyestuff decolourisation performances of C. versicolor MUCL.
Complete decolourisation was observed for initial dyestuff concentrations lower than 500
mg/L whereas any concentrations beyond 1200 mg/L may have a toxic effect on the fungus.
They established that biodegradation rather than adsorption of dyestuff on the fungus (<20%)
was the major mechanism involved in the removal of dyestuff concentrations up to 1200
mg/L.

Due to the similarity of parts of the structure of NOM with lignin, several researchers have
shown the capability of a number of white-rot fungi to decolourise and degrade humic
substances, among which were P. chrysosporium (Blondeau 1989, Ralph & Catcheside 1994),
Trametes (Coriolus) species (Dehorter & Blondeau 1992, Yanagi et al. 2003), Clitocybula
dusenii (Ziegenhagen & Hofrichter 1998) and Panus tigrinus (Zavarzina et al. 2004).

30
Blondeau (1989) reported that the mineralisation of humic acids by LiP of P. chrysosporium
BKM-F 1767 occurred during secondary metabolism in nitrogen-limited medium, however,
low activity of LiP was produced in the presence of humic acids. A reduction in the high
molecular weight range and no accumulation of low molecular weight species were obtained
after the fungal treatment. Ralph and Catcheside (1994) studied the decolourisation and
depolymerisation of low rank coal (lignin-like polymers) with P. chrysosporium and
established that the degradation of the coal coincided with the presence of extracellular LiP
and MnP. They also suggested that aryl cation radicals and possibly Mn3+ generated by LiP
and MnP, respectively, diffused into the coal matrix and initiated the free radical oxidation
reactions resulting in the fission of carbon bonds and thus depolymerisation, as has been
proposed for lignin mineralisation.

Dehorter and Blondeau (1992) illustrated a relationship between the degradation of humic
acid and extracellular enzyme activity for P. chrysosporium and T. versicolor, showing
increased formation of extracellular LiP and MnP with increasing humic acid concentration,
which was in contrast to the results of Blondeau (1989). They found that T. versicolor was
more effective in degrading humic acids than P. chrysosporium with MnP as a key enzyme
responsible for mineralisation. A study by Yanagi et al. (2003) with Coriolus consors IFO
9078 demonstrated that decolourisation of humic acids with different chemical properties
ranged from 9 to 40%, with higher aromaticity and humification providing higher resistance
to microbial decolourisation.

C. dusenii b11 was another white-rot fungus investigated for its ability to degrade humic acids
obtained from low rank coal. Ziegenhagen and Hofrichter (1998) determined the optimum
conditions for the action of MnP and established that the MnP-catalysed depolymerisation of
humic acids in vitro produced low molecular weight fulvic acids by breaking down covalent
bonds. Recently Zavarzina et al. (2004) studied the changes in the structures of soil- and peat-
derived humic acids after biotransformation by laccase from the white-rot fungus P. tigrinus
8/18. It was reported that the purified laccase was able to polymerise and depolymerise humic
acids, and the transformations were dependent upon the nature and properties of humic acids.

Melanoidin, a highly coloured constituent of the spent wash largely produced in the ethanol
industry, has high environmental pollution potential (Fahy et al. 1997, Dahiya et al. 2001).
The brown melanoidin polymers, formed by the Maillard amino-carbonyl reaction (Wedzicha
& Kaputo 1992), have antioxidant properties and structures similar to wood and NOM. Fahy

31
at al. (1997) and Dahiya et al. (2001) studied a microbial decolourisation process for
melanoidin pigments present in spent wash using P. chrysosporium. Dahiya et al. (2001)
reported that 80% colour removal was achieved and showed that the rate of melanoidin
decolourisation for the high molecular weight fractions was faster than for the low molecular
weight fractions.

Rojek et al. (2004) have shown that Phanerochaete chrysosporium ATCC 34541 was able to
remove 40-50% NOM from solution, although this was shown to be mainly due to adsorption
and to be partially metabolically linked. Following the work of Rojek (2003), further
investigations were conducted to examine the bioremediation of NOM wastes to develop an
improved biological system applicable to wastewater treatment.

32
Chapter 3 Materials and Methods

3.1 NOM Samples

The highly coloured MIEXTM NOM concentrate from Hope Valley Reservoir, located in
South Australia, was utilised as a source of organic matter throughout the experiments. The
NOM concentrate was obtained from the regeneration process of the strong base magnetic ion
exchange (MIEX) resin, a recently developed process for the removal of dissolved organic
carbon (DOC) (Slunjski et al. 2000).

The concentrate was filtered (0.45 μm hydrophilic PVDF, Millipore Millex-HV) and stored at
4oC prior to treatment and analysis. The characteristics of the NOM concentrates used in this
study, which were collected at different times, are tabulated in Table 3.1.

Table 3.1 Characterisation of MIEXTM NOM concentrates.

Description Units MIEXTM NOM Concentrates
Batch NOM 1 NOM 2 NOM 3

Collection Date March, 2001 July, 2003 August, 2003

pH 7.63 7.15 8.30
Absorbance at 446 nm cm-1
0.079 0.024 0.728
(1:100)a
Absorbance at 254 nm cm-1
0.114 0.032 0.582
(1:2000)b
DOC g C/L 6.5 2.0 31.0
SUVAc L.mg-1.m-1 3.5 3.2 3.8
a
Dilution factor of 100
b
Dilution factor of 2000

c A254
SUVA (Specific UV absorbance) =
DOC

33
3.2 Micro-organisms
Phanerochaete chrysosporium strains ATCC 34541 and ATCC 24725, Trametes versicolor
strain ATCC 7731 and Saccharomyces species arbitrarily denoted 1, 2 and 3 (isolated from
NOM concentrate in the RMIT University laboratory) were used in this study. P.
chrysosporium and the yeast strains were maintained by subculturing monthly on Waksman
medium agar slants for 3-4 days whereas T. versicolor was grown on 2% (w/v) malt extract
agar (MEA) for 4-5 days. All organisms were incubated at 30oC and then maintained at 4oC
prior to use.

The composition of the Waksman medium agar slants for fungal maintenance is detailed in
Table 3.2. All chemicals were of AR purity.

Table 3.2 Composition of Waksman medium agar slants.

Medium component Sources Content (g/L)
Agar Southern Biological 25.0
Mycological peptone Oxoid 5.0
D-Glucose Merck 10.0
NH4Cl Ajax Chemicals 2.0
KH2PO4 Asia Pacific Specialty Chemicals Limited 1.0
MgSO4 .7H2O Asia Pacific Specialty Chemicals Limited 0.5

3.3 Medium and Culture Conditions

Three simple growth media, i.e., modified Waksman medium (Booth 1971), Fahy medium
(Fahy et al. 1997) and Fujita medium (Fujita et al. 2002) were employed. Medium was first
adjusted to pH 4.5 before being sterilised by autoclaving at 121oC for 20 minutes. The
composition of the listed media is given in Table 3.3.

Medium was prepared and 200 mL added to 500 mL Erlenmeyer flasks, which were then
autoclaved. The shake flasks were then supplemented with filter sterilised (0.45 μm
hydrophilic PVDF, Millipore Millex-HV) NOM (final concentrations of 100-700 mg C/L)
and inoculated with the fungal or yeast strains (Section 3.4). The cultures were incubated at
30oC or 36oC as indicated for various periods at 130 rpm.

34
Table 3.3 Composition of growth media.
Composition (g/L)
Medium
Glucose NH4Cl NH4NO3 KH2PO4 MgSO4
Waksman 2.0 or 5.0a 0.5 - 1.0 0.5
Fahy 25.0 0.5 - 2.5 1.25
Fujita 10.0 - 0.1 1.0 0.5
a
Glucose content varied according to the experiment.

3.4 Preparation of Inoculum

3.4.1 Fungal inoculum

The fungi were inoculated as either a spore suspension or as plugs, as indicated. Spore
suspensions were prepared by washing agar plates with sterilised water and then filtering
through sterile glass wool. Spore concentration was determined by measuring the absorbance
at 650 nm and calculated on the basis that A650 = 1.0 cm-1 corresponds to 5.0 x 106 spores/mL
(Kirk et al. 1978). Spore suspension (10 mL) was then added to the culture media to attain a
concentration of 1.0-1.5 x 105 spores/mL. Inoculation as plugs involved the addition of three
agar plugs of 1 cm2 each excised from a fungal colony actively growing on a MEA plate with
a sterilised cutter.

3.4.2 Yeast inoculum

Yeast cultures were prepared with a loopful of inoculum from malt extract agar (MEA) plate.

3.5 Supplements
In some experiments, as indicated, two supplements were employed: wheat bran and Tween
80. Wheat bran (4.5 g/L, Kellogg’s) and Tween 80 (0.5% v/v, Merck) were added to the
growth medium prior to sterilisation.

3.6 Analytical Methods

3.6.1 pH
A HACH Sension 156 Multiparameter Meter was used to measure medium pH. Any
adjustment of pH was made with 0.1 M, 0.5 M or 1.0 M NaOH and H2SO4, as appropriate.

35
3.6.2 Dissolved organic carbon
DOC concentration was determined using a total organic carbon (TOC) analyser (Sievers,
Model 820). Samples were filtered (0.45 μm hydrophilic PVDF, Millipore Millex-HV) and
diluted as required with Milli-Q water prior to analysis.

3.6.3 Absorbance
Absorbance measurements were performed with a double beam scanning UV/vis
spectrophotometer (Unicam, Model UV2) fitted with a cell of 1 cm pathlength. The
absorbance of NOM solution was measured at both 446 nm (colour) and 254 nm (UV-
absorbing components). The correlations between NOM concentration for the three different
preparations and absorbance at 446 nm and 254 nm are provided in Appendix 1. Samples
were centrifuged until the solution was clear and were diluted to 1:10 with Milli-Q water prior
to A254 measurements.

3.6.4 Determination of absorbance correction factor

The absorbance of NOM varies with pH, therefore corrections were applied where necessary,
using the absorbance at initial pH as reference. NOM in medium at varying concentrations
(50, 100 and 200 mg C/L) and pH (2-6) for the three different NOM preparations were
prepared in duplicate and the absorbances at 446 nm and 254 nm were measured. As the pH
of the culture fluid was usually in the range 2-6, the change in absorbance (%) was calculated
using the absorbance at pH 6 as initial value, and pH 2 and its corresponding absorbance as
final value. An almost linear relationship between pH versus absorbance was obtained hence
the least squares method was applied to calculate the formula for the relationship between pH
change (difference from initial to final) and the absorbance correction factor (%). Thus the
formula for calculating percent change in absorbance for all NOM concentrations for any
change in pH was established.

All data for 446 nm and 254 nm presented in this study have been corrected for pH. Plots
illustrating the influence of pH on absorbances at 446 nm and 254 nm for the three NOM
preparations and description of the calculation of corrected absorbance can be referred to in
Appendix 2.

36
3.6.5 Determination of glucose concentration
Reducing sugar concentrations of samples were measured by the 3', 5’-dinitrosalicylic acid
(DNS) method (Miller 1959) using D-glucose as a standard. A typical standard curve for
glucose determination is shown in Appendix 3.

3.6.6 Dry weight of biomass

Dry weight of biomass for both fungal and yeast strains was determined at the end of
fermentations by filtering the biomass on pre-weighed dried membrane filters (0.45 μm
Whatman WCN sterile membranes), washing with distilled water and then drying them in an
oven at 90oC to constant weight. The dry weight of the biomass was then calculated by
difference.

3.6.7 Enzyme assays

All enzyme assays were performed at 50oC, unless stated otherwise, as the reaction rates were
slow and so would have been even slower at lower temperatures.

(i) Laccase activity

Laccase (Lac) activity was determined spectrophotometrically as described by Coll et al.
(1993) with guaiacol as a substrate. The oxidation of guaiacol (Sigma, 100%) to the polymer
tetraguaiacone was monitored by increase in absorbance at 465 nm. The activity was
expressed in U/mL where one unit (U) of Lac activity was defined as the amount of the
enzyme that caused an increase of one absorbance unit per minute (Coll et al. 1993). The
reaction (Figure 3.1) was started by addition of guaiacol.

Lac
R O O R

OMe
OH
Tetraguaiacone
Guaiacol (brown component)

where R represents the molecule after having donated H

Figure 3.1 Proposed mechanism for the oxidation of guaiacol by laccase (Niwa 2004).

37
The reaction mixture (3 mL) contained:
1.0 mM guaiacol 300 μL
0.2 M sodium acetate-acetic acid buffer, pH 4.5 1800 μL
Distilled water 600 μL
Extracellular culture fluid 300 μL

(ii) Lignin peroxidase

Lignin peroxidase (LiP) activity was measured according to Tien and Kirk (1988) by
monitoring the oxidation of veratryl alcohol (VA) (Aldrich, 96%) to veratraldehyde at 310
nm. Enzyme activity was calculated from the molar extinction coefficient of ε = 9.30 mM-
1
cm-1 and expressed in units (U) which correspond to 1.0 μmole of veratraldehyde produced
per minute (Tien & Kirk 1988). The reaction (Figure 3.2) was started by addition of hydrogen
peroxide.

CH2OH CH2OH

LiP/H2O2
+.

OMe OMe
OMe OMe
Veratryl alcohol Cation radical

Figure 3.2 Proposed mechanism for catalysing reduction of VA by LiP (Aust 1995).

The assay mixture (3 mL) included:

20.0 mM VA 300 μL
0.2 M sodium acetate-acetic acid buffer, pH 3.0 840 μL
4.0 mM H2O2 300 μL
Extracellular culture fluid 1560 μL

(iii) Manganese-dependent peroxidase

Manganese-dependent peroxidase (MnP) activity was assayed according to Wariishi et al.
(1992) by measuring the oxidation of 2, 6-dimethoxyphenol (DMP) (Aldrich, 99%) to
coerulignone at 469 nm. The activity was expressed in units (U) and determined from the
molar extinction coefficient of ε = 49.6 mM-1cm-1. One unit of MnP activity is defined as the

38
amount of the enzyme catalysing the formation of 1.0 μmole of coerulignone per minute
(Wariishi et al. 1992). The reaction is shown in Figure 3.3.

MnP/ Mn 2+
2 2

OMe OMe OMe OMe

OH O.

DMP

Figure 3.3 Proposed mechanism for the oxidation of DMP by MnP (Wariishi et al.
1992).

The assay mixture (3 mL) contained:

0.5 mM DMP 300 μL
0.2 M sodium acetate-acetic acid buffer, pH 4.5 1800 μL
1.0 mM MnSO4 300 μL
0.5 mM H2O2 300 μL
Extracellular culture fluid 300 μL

3.6.8 High performance size exclusion chromatography

The molecular weight distribution of samples was determined using high performance size
exclusion chromatography (HPSEC) at the Australian Water Quality Centre (AWQC),
Adelaide.

The analysis was conducted using a Waters 2690 Alliance system with a temperature
controlled oven at 30oC and a Shodex KW 802.5 glycol functionalised silica gel column with
a Waters 996 Photo Diode Array detector set at 260 nm. The column was calibrated with
polystyrene sulphonate standards and the apparent molecular weight (Dalton) of NOM was
calculated from the linear regression of the relationship between the retention time (t,
minutes) and the logarithm of molecular weight of the standards (log (Mw)):
log(M w ) = −0.399 × t + 7.205 (Appendix 4).

The weight average molecular weight (Mw) and the number average molecular weight (Mn)
were calculated as per Equation 3.1 and Equation 3.2, respectively.

39
 Σni M i
2

Mw = Equation 3.1
Σni M i

Σni M i
Mn = Equation 3.2
Σn i

where ni is the number of molecules of weight Mi. Polydispersity was calculated as the ratio
Mw: Mn.

3.6.9 Fractionation of NOM

A NOM fractionation system as designed by Chow et al. (2004) was constructed as shown in
Figure 3.4. The system allows fractionation of the NOM into four categories: very
hydrophobic acids (VHA), slightly hydrophobic acids (SHA), hydrophilic charged (CHA) and
hydrophilic neutral (NEU) compounds. The VHA, SHA and CHA fractions were adsorbed by
DAX-8 resin, XAD-4 resin and IRA-958 resin respectively; and the NEU fraction was the
effluent from the IRA-958 column.

Three 20 cm glass columns for DAX-8, XAD-4 and IRA-958 resins respectively were set up
in series as shown in Figure 3.4 after exhaustive cleaning of resins with methanol and Milli-Q
water. Resin-water slurries were added to give bed volumes of approximately 14.7 mL, 14.1
mL and 20.6 mL respectively. Each bed was backwashed with 2-3 L Milli-Q water to classify
the resin particles and to remove air bubbles and debris (Chow et al. 2004).

Before fractionation, samples were filtered through a 0.45 μm hydrophilic PVDF (Millipore
Millex-HV) and acidified to pH 2.0 with concentrated HCl. The pH-adjusted samples were
then passed through the DAX-8 column at the rate of 0.2 bed volumes/min. The first two bed
volumes were discarded before collecting the effluent. A sub sample of 100 mL was stored
for TOC, A446 and A254 assays. The remaining effluent was then passed through the second
(XAD-4) column. The same procedures were followed except that the effluent from the XAD-
4 column was adjusted to pH 8.0 with NaOH solution before pumping through the last (IRA-
958) column. The DOC of each fraction was calculated by the difference between the DOC of
effluents of the columns.

40
Figure 3.4 Schematic diagram of the fractionation unit (Chow et al. 2004).

41
Chapter 4 Decolourisation and Bioremediation of MIEXTM NOM
Following the work of Rojek (2003), preliminary fermentations using P. chrysosporium
ATCC 34541 for the removal of NOM from various batches of MIEX NOM concentrate were
conducted to identify the effect of different characteristics of NOM on the process, and so the
applicability of the process to the treatment of NOM waste arising from drinking water.

Three NOM preparations (refer to Materials and Methods-Section 3.1 where described) were
fractionated before the treatment to determine the natures of the organic compounds present.
A comparison of the changes in colour (A446), UV-absorbing components (A254) and
molecular weight distribution (HPSEC) was carried out after the fermentations to establish the
fractions removed by P. chrysosporium. This was followed by the selection of a simple
medium from three growth media with different C:N ratios. Three strains of white-rot fungi
(P. chrysosporium ATCC 34541 and 24725, and T. versicolor ATCC 7731) and yeast
(Saccharomyces species 1, 2 and 3) were investigated to develop an improved NOM waste
biodegradation system.

4.1 Removal of Different Preparations of NOM by P. chrysosporium ATCC 34541

The three NOM preparations were added to 200 mL (final concentration of 100 mg C/L) of
Waksman medium (2 g/L glucose), inoculated with 10 mL of spore suspension of P.
chrysosporium ATCC 34541, and incubated at 36oC and 130 rpm for five days. The
fermentations for each NOM preparation were performed in duplicate. Data points represent
mean values of duplicates.

The trends of the history plots for the three NOM preparations were similar, and the plot for
NOM 3 (Figure 4.1) is shown as it gave the highest reduction in A446 and A254. The pH
dropped markedly over the first two days of incubation and then plateaued. The change in pH
occurred concurrently with the decrease in glucose content. The drop in pH was probably due
to the accumulation of organic acid as metabolite or by the freeing of hydrogen ions in
substrate transfer by the fungus (Griffin 1994). This may support biosorption as the
bioadsorptive capacity of a fungus increased with decreasing pH for humic acids as reported
by Zhou and Banks (1991). The fungus consumed only approximately half of the glucose (1.0
g/L) provided over the incubation period. This is consistent with the results found by Rojek
(2003), where the fungus consumed only 1-1.2 g/L glucose even though higher initial glucose
concentrations (4 and 10 g/L) were supplied. A446 decreased for the first three days and then

42
plateaued whereas A254 decreased gradually during the whole fermentation. This is contrary to
the findings of Blondeau (1989), where the decolourisation of humic acids by P.
chrysosporium BKM-F 1767 only started after an initial lag phase of four days and continued
up to day 15 and suggested that the lignin-degrading system played a role in the humic acid
decolourisation.

6.00 0.40 pH

5.00 Glucose
consumption (g/L)

0.30 consumption
pH OR Glucose

4.00 A446

Absorbance
A254
3.00 0.20

2.00
0.10
1.00

0.00 0.00
0 1 2 3 4 5
Incubation period (days, 36oC &
130 rpm )

Figure 4.1 History plot showing pH, glucose consumption, A446 and A254 for NOM 3
incubated with P. chrysosporium ATCC 34541 at 36oC and 130 rpm for five days. (A254
represents readings of 1/10 dilution of culture medium)

Figure 4.2 clearly indicates that there was a decrease in absorbance at 446 nm whereas there
was less reduction at 254 nm for all the NOM preparations. The plot did not give clear
information on the extent of reduction in A254 as the culture fluids were diluted in 1/10 and so
removal in terms of total carbon was calculated (Figure 4.3).

The initial and final colour (as A446) of NOM 3 was much higher than for the other two NOM
preparations even though the initial NOM concentrations in terms of total carbon were the
same. The greatest decolourisation for all NOM preparations was obtained on day 3: 37%
(NOM 3), 31% (NOM 1) and 29% (NOM 2). Losses in A254 of 9% for NOM 3 and 3% and
1% for NOM 2 and NOM 1, respectively, occurred after five days (Figure 4.2).

43
 0.40
NOM 1

Absorbance at 446 nm or 254 nm NOM 2

0.30
A254 NOM 3

0.20

0.10
A446

0.00
0 1 2 3 4 5
o
Incubation period (days, 36 C & 130 rpm )

Figure 4.2 A446 and A254 of NOM 1, NOM 2 and NOM 3, 100 mg C/L initial NOM
concentration, P. chrysosporium ATCC 34541. (A254 represents readings of 1/10 dilution
of culture medium)

To elucidate NOM removed in terms of total carbon, NOM removals (converted to mg),
measured at 446 nm and 254 nm, were calculated (Figure 4.3). The extent of NOM removal
followed the trends: NOM 3 > NOM 1 > NOM 2 for A446, and NOM 3 > NOM 2 > NOM 1
for A254. The low reduction in A254 indicates that the removal of conjugated double bonds
(unsaturated aldehydes, phenols, aliphatic and aromatics) by these systems was negligible;
consequently, little if any chemical change to the UV-absorbing components due to fungal
activity or removal by adsorption occurred. The addition of NOM 1 and NOM 2 led to similar
NOM removal, glucose consumption and biomass generation. In contrast, the fungus caused
the removal of more NOM, higher glucose consumption and produced more biomass for
NOM 3.

It was observed that the fungal pellets appeared brownish in colour. This indicated that NOM
molecules were bonded to the fungal mycelium and so adsorption of NOM to the biomass
seemed to play a role in colour removal as found by Rojek (2003). As the removal of colour
was greatest for NOM 3, for which the biomass was greatest, and the fungal pellets were
uniform in colour for all three NOM preparations, the major mechanism for NOM removal
appeared to be via adsorption. This concurs with the previous proposal that NOM removal by
biosorption occurred due to the pH drop (Rojek 2003) as low pH supports the binding of
humic acid components to the fungal cell wall surface (Zhou & Banks 1991). In addition, the
degree of removal of NOM was directly related to the amount of biomass produced due to the
greater number of adsorption sites.

44
 80 10.0 Biomass

NOM removal (mg) or Glucose

A446
8.0
60

consumption (g/L)
A254
Biomass (mg)

6.0 Glucose
40 consumption
4.0

20
2.0

0 0.0
NOM 1 NOM 2 NOM 3
NOM preparation

Figure 4.3 NOM removals (as mg, converted from A446 and A254), glucose
consumption (g/L) and dry weight of biomass generated (mg) for the three NOM
preparations on day 5, 100 mg C/L initial NOM concentration, P. chrysosporium ATCC
34541.

To better understand the components of the three NOM preparations, they were fractionated
according to the method of Chow et al. (2004). This was to provide information on which
types of compounds are removed in the treatment, as was mentioned by Yanagi et al. (2003)
where the extent of decolourisation of humic acids varied with different chemical properties.

4.2 Fractionation of the MIEXTM NOM Preparations

The three NOM (100 mg C/L) preparations were separated into four fractions: VHA, SHA,
CHA and NEU to establish the types of organic compounds present (see Section 3.6.9). The
relative proportions of DOC, A446 and A254 in each fraction were determined. The proportion
of each fraction in Figure 4.4 is an average of duplicate determinations, and the values of the
duplicates varied only by ±1%.

The three preparations were dominated by hydrophobic acids such that VHA > SHA > CHA
> NEU (Figure 4.4). NOM 1 and NOM 2 exhibited almost identical proportions of these
fractions, but those in NOM 3 were markedly different. NOM 3 had the highest hydrophobic
content with 69% VHA and 17% SHA. The hydrophilic neutral fraction was relatively small
for all the preparations. This is because the MIEXTM NOM concentrates were obtained from
the resin regeneration process, and constitute only approximately 80% of the NOM originally
present in the water since the MIEX resin is an anionic exchanger and so cannot remove the
neutral components of NOM (Slunjski et al. 2000).

45
 VHA
2%
1% 2%
SHA
20% 19% 12%
CHA
17% NEU
49% 51%

30% 28% 69%

A B C

Figure 4.4 The proportions of each fraction in the three NOM preparations: (A)
NOM 1, (B) NOM 2 and (C) NOM 3 solutions. (N = 2; i.e., number of times each was
determined)

It has been shown that NOM fractions contain a wide range of compounds and the types of
compounds in each fraction are dependent on the water source. The typical classes of
compounds in each fraction are summarised in Table 2.3 (Swietlik et al. 2004). The
hydrophobic fractions are inclined to possess greater aromaticity than the hydrophilic
fractions; therefore NOM 3 had the highest proportion of conjugated aromatic and high
molecular weight compounds as supported by the HPSEC chromatograms in Figure 4.7(A).

The A446 (Figure 4.5) and A254 (Figure 4.6) for each fraction were measured to determine the
colour and the UV-absorbing components, respectively, in the different NOM preparations, to
give information on specific organic groups, rather than just the total dissolved organic
species. The molecular weight distributions were then compared using HPSEC (Figure 4.7).

4.2.1 Comparison of A446 of the NOM fractions

Colour, as A446, was mainly contributed by the hydrophobic fractions, particularly the VHA
fraction for which NOM 3 had the highest value (Figure 4.5A). The hydrophilic fractions
(CHA and NEU) contributed little colour for all three NOM preparations. Thus the greatest
removal of colour in the fermentation for NOM 3 (Figure 4.3) was due to its higher VHA
content.

To better demonstrate the relative colour of each fraction in the NOM preparations, the A446
values were normalised against the DOC concentration of each fraction (A446/DOC) (Figure
4.5B). It is clear that the specific A446 values for the VHA and SHA fractions of NOM 3 were
higher than for NOM 1 and NOM 2. The decolourisation of NOM must be due to the breaking
of bonds and/or adsorption of the VHA and SHA fractions of NOM as the specific values for

46
the CHA and NEU fractions were relatively low. This also explained the higher colour
removal by P. chrysosporium from NOM 3 as being due to the greater hydrophobic content.

NOM 1
0.25 0.0030
NOM 2
Absorbance at 446 nm

0.20
NOM 3
0.0020

A446 / DOC
0.15

0.10
0.0010
0.05

0.00 0.0000
VHA SHA CHA NEU VHA SHA CHA NEU
A Fraction B Fraction

Figure 4.5 A446 and A446/DOC of the fractions of the three NOM preparations.

4.2.2 Comparison of A254 of the NOM fractions

The VHA fractions were the most intensely UV absorbing, with NOM 3 having the highest
and NOM 1 and NOM 2 preparations displaying similar UV-absorbing contents (Figure
4.6A). The UV-absorbing species in the hydrophilic fractions were mostly in the CHA
fraction; the NEU fractions showed very low A254 values.

NOM 1
3.0 0.05
NOM 2
Absorbance at 254 nm

2.5 0.04
NOM 3
2.0
A254 / DOC

0.03
1.5
0.02
1.0

0.5 0.01

0.0 0.00
VHA SHA CHA NEU VHA SHA CHA NEU

A Fraction Fraction
B

Figure 4.6 A254 and A254/DOC of the fractions of the three NOM preparations.

However when the normalised A254 values were calculated, the VHA fraction in NOM 3 had
relatively lower UV-absorbing content per unit DOC than NOM 1 and NOM 2. In contrast,
the UV-absorbing content of the SHA fraction in NOM 3 was relatively higher. The A254 and
also the normalised A254 of the NEU fractions were low for all three NOM preparations
(Figure 4.6B). The higher total normalised A254 of the hydrophobic fractions (VHA + SHA)

47
for NOM 3 is consistent with the higher SUVA for this preparation and thus indicative of
greater aromaticity. In contrast, the three NOM preparations had low total normalised A254 of
the hydrophilic fractions (CHA + NEU) and so did not contain high aromatic nor conjugated
compounds in the hydrophilic fractions. Therefore, NOM 3 was considered to be composed
largely of aquatic humic substances and a relatively higher content of hydrophobic and
aromatic compounds compared with NOM 1 and 2, which is consistent with the high SUVA
for this preparation (Table 3.1).

As the A254 for all three NOM preparations was largely associated with the VHA and SHA
fractions, reduction in A254 in the presence of P. chrysosporium is considered to correspond to
breaking of the UV-absorbing bonds in or removal of these fractions. To elucidate the
breakdown products, GC-MS can be applied to fully establish bond breakage. Again, the
greater removal of NOM 3 (Figure 4.3) was attributed to the higher VHA and SHA contents,
although some CHA fraction may also be involved.

4.2.3 Comparison of molecular weight distribution of the NOM fractions

The molecular size distributions of the UV-absorbing components of the ‘whole’ NOM and
each NOM fraction for the three NOM preparations were investigated (Figure 4.7). NOM 1
and NOM 2 had very comparable molecular weight distribution with a large peak at ~1500
Dalton and another at 400 Dalton (Figure 4.7A). NOM 3 had peaks at almost similar apparent
molecular weights, with an additional peak at 1000 Dalton, and it exhibited some higher
molecular weight compounds.

The UV-absorbing species in all the NOM preparations were mostly in the hydrophobic
fractions (VHA and SHA). The hydrophobic fractions contained molecules of apparent
molecular weight >2000 Dalton for the three NOM preparations, but this was not so for the
CHA fractions. The molecular weight distribution of each fraction was very similar for NOM
1 and NOM 2; NOM 3 had a very different pattern for all fractions. It should be noted that the
NEU fractions were very small for all and so no trends were apparent.

The VHA fraction gave two peaks for all the NOM preparations: at apparent molecular
weights of ~1500 Dalton and 400 Dalton (Figure 4.7B). The absorbance at the peak of ~1500
Dalton for NOM 3 was lower than for NOM 1 and NOM 2, however, it contained more
compounds with apparent molecular weight >2000 Dalton.

48
The SHA fraction comprised markedly fewer high molecular weight UV-absorbing
compounds than the VHA fraction for the three NOM preparations. NOM 3 possessed two
peaks in the molecular weight range of 1000-2000 Dalton but with lower absorbance
compared with the VHA fraction. There was only one peak in the SHA fraction for NOM 1
and NOM 2, which was at ~1500 Dalton (Figure 4.7C).

The CHA fraction for NOM 3 was relatively lower than for NOM 1 and NOM 2. All NOM
preparations had a peak at ~1500 Dalton and an additional peak at 1000 Dalton for NOM 3
(Figure 4.7D). Their UV-absorbing contents were very small compared to the hydrophobic
fractions, especially in NOM 3.

There was little, if any, UV-absorbing species in the NEU fraction for all the NOM
preparations (Figure 4.7E) as the most UV-absorbing compounds in the hydrophilic fractions
(CHA and NEU) were in the CHA fraction.

The three NOM preparations contained less non-humic (hydrophilic) than humic
(hydrophobic) substances due to the lower contents of the UV-absorbing species in the
hydrophilic fractions, especially in NOM 3, than the hydrophobic fractions. Possible
constituents of the hydrophilic fractions would be hydrophilic base compounds such as
amphoteric proteinaceous materials containing aliphatic amino acids, amino sugars, peptides
and proteins (Table 2.3) as the NOM preparations were highly alkaline (pH 7.2-8.3) (Table
3.1). Polysaccharides, hydrophilic neutral compounds, were unlikely to be present in the
NOM preparations as the NEU fraction was very small (Figure 4.4).

All the NOM preparations most likely contained fulvic acids and humic acids with NOM 3
having the highest humic acid content as it was highly alkaline and dark brown in colour, and
had more high molecular weight compounds (>2000 Dalton). This agrees with the
hypothetical relationships for chemical characteristics of humic substances (Figure 2.3) where
humic acids showed higher colour intensity and molecular weight, and lower acidity than
fulvic acids. In addition, Howe and Clark (2002) reported that the molecular weight
distribution for aquatic humic acid was 2000-5000 Dalton, where NOM 3 showed the highest
content (Figure 4.7A). NOM 3 contained both aliphatic and aromatic high molecular weight
compounds with less for NOM 1 and NOM 2 as NOM 3 had the highest total normalised A254
of the hydrophobic fractions (Figure 4.6B), which is consistent with the highest SUVA value
for this preparation (Table 3.1) and thus indicative of greater aromaticity.

49
 0.400
NOM 1
0.350
NOM 2
0.300

UV Abs @ 260 nm
NOM 3
0.250

0.200

0.150

0.100

0.050

0.000
100 1000 10000 100000

A Apparent m olecular w eight (Dalton)

0.250 0.100

0.200 0.080
UV Abs @ 260 nm
UV Abs @ 260 nm

0.150 0.060

0.100 0.040

0.050 0.020

0.000 0.000
100 1000 10000 100000 100 1000 10000 100000
B C

0.100 0.100

0.080 0.080
UV Abs @ 260 nm

UV Abs @ 260 nm

0.060 0.060

0.040 0.040

0.020 0.020

0.000 0.000
100 1000 10000 100000 100 1000 10000 100000
D Apparent molecular weight (Dalton) E Apparent molecular weight (Dalton)

Figure 4.7 HPSEC chromatograms for the (A) ‘whole’ NOM, (B) VHA, (C) SHA, (D)
CHA and (E) NEU fractions for all NOM preparations.

50
 12000 1.40 Mw

Mn
1.20
Molecular weight (Dalton) 10000
Polydispersity
1.00
8000

Polydispersity
0.80
6000
0.60
4000
0.40

2000 0.20

0 0.00
1 2 3

NOM preparation

Figure 4.8 Weight average molecular weight (Mw), number average molecular weight
(Mn) and polydispersity for the three NOM preparations.

The weight average molecular weight (Mw) and the number average molecular weight (Mn)
were calculated so as to compare the variation in molecular weight and the extent of
polymerisation (Figure 4.8). Mw for NOM 1 and NOM 2 were similar, that of NOM 3 was
higher. Mn for all NOM preparations was comparable. Polydispersity, calculated as the ratio
of Mw: Mn, is a measure of the extent of polymerisation. The polydispersity of NOM 3 was
approximately 20% higher than that NOM 1 and NOM 2. These results were consistent with
the previous observations where NOM 3 had the highest content of high molecular weight
compounds and greatest colour intensity. These findings were also in agreement with the
postulated relationships reported by Weber (2001) (Figure 2.3), where the degree of
polymerisation increases with the intensity of colour and molecular weight.

51
4.3 Molecular Size Distribution of the NOM after Treatment with P. chrysosporium
ATCC 34541
The molecular weight distribution of the UV-absorbing species for the three NOM
preparations after five days treatment with P. chrysosporium (Section 4.1) was determined
using HPSEC (Figure 4.9).

A small shift from higher to lower molecular weight for the NOM remaining after treatment
with P. chrysosporium was observed for all NOM preparations (Figure 4.9). NOM 3 showed a
slightly greater shift to lower molecular weight following the treatment. The removal of the
high molecular weight compounds was accompanied by the accumulation of the low
molecular weight species, presumably due to some breakdown by the fungus, especially for
the NOM 3 preparation. Blondeau (1989) also obtained reduction in the high molecular
weight range but without accumulation of low molecular weight species after the fungal
treatment. However, adsorption also contributed to the removal of the coloured high
molecular weight fractions as the fungal pellets turned brown as found by Rojek (2003).

All the NOM preparations exhibited a decrease in the UV absorbance for the molecular
weight fraction >2000 Dalton. This was largely due to the decrease in the high molecular
weight hydrophobic fractions and was most marked for NOM 3 as only the hydrophobic
fractions exhibited apparent molecular weight >2000 Dalton (Figure 4.7A & B). For the
cultures with NOM 1 and NOM 2 there was almost no shift in the apparent molecular weight
range 1000-2000 Dalton, and there was a similar reduction in absorbance for the peak at
~1500 Dalton. In contrast, there was a slight shift in apparent molecular weight in the range
1000-2000 Dalton and some decreases in absorbance for the culture with NOM 3.

P. chrysosporium seemed to preferentially remove the VHA fraction, and so was most
effective for the NOM preparation with the highest VHA content. However, this could not be
demonstrated by fractionation of the NOM-containing medium after growth of the fungus as it
contained glucose, metabolic products and salts, which would have interfered with the
function of the resins.

52
 0.500
NOM 1
0.450
UV Abs @ 260 nm 0.400
0.350
0.300
0.250
0.200
0.150
0.100
0.050
0.000

0.500
NOM 2
0.450
0.400
UV Abs @ 260 nm

0.350
0.300
0.250
0.200
0.150
0.100
0.050
0.000

0.500
NOM 3
0.450
0.400
UV Abs @ 260 nm

0.350
0.300
0.250
0.200
0.150
0.100
0.050
0.000
100 1000 10000 100000
Apparent molecular weight (Dalton)

control-NOM control-P. chry 34541 + P. chry 34541

Figure 4.9 HPSEC chromatograms for the three NOM preparations incubated with
P. chrysosporium in Waksman medium, control-NOM (medium plus NOM), control-P.
chry 34541 (P. chrysosporium ATCC 34541 grown without NOM).

53
The weight average molecular weight (Mw) and number average molecular weight (Mn) for
UV- absorbing components based on HPSEC were calculated after five days treatment of the
three NOM preparations with P. chrysosporium. The greatest reduction in Mw and Mn was
obtained for the culture containing NOM 3. The original values of Mw of 10,188 (NOM 3),
8626 (NOM 2) and 8514 (NOM 1) were reduced by 3%, 2% and 2%, respectively. There was
a similar trend of reduction in Mn for all the NOM preparations (Figure 4.10).

The NOM 3 preparation was selected for further investigation as this batch showed greatest
NOM removal, had the greatest colour, aromaticity and the highest SUVA, and exhibited the
greatest change in the high molecular weight compounds due to its high proportion of the
hydrophobic fractions and so provided a suitable material to further investigate NOM
removal.

300
Mw
Reduction in molecular weight

250 Mn

200
(Dalton)

150

100

50

0
1 2 3
NOM preparation

Figure 4.10 Reduction in weight average molecular weight (Mw) and number average
molecular weight (Mn) after the treatment of the NOM preparations with P.
chrysosporium ATCC 34541.

4.4 Selection of Medium

The aim of this study was to develop a system for the bioremediation of concentrated NOM in
a simple medium. As removal of NOM from Waksman medium by P. chrysosporium ATCC
34541 was low, two other media were tested to see if NOM removal could be increased with
different C:N ratios.

The effects of three different growth media: Waksman, Fahy and Fujita medium, on NOM
decolourisation by P. chrysosporium were established. Carbon starvation (1.0 g/L glucose)
54
did not trigger a degradation process (Rojek et al. 2004) and so Waksman medium with 2.0
g/L glucose was selected. These media were chosen as they showed a wide range of initial
glucose and nitrogen concentrations, thus giving different ratios. C:N ratio has been
considered as a better predictor of lignin degradation than the absolute levels of carbon and
nitrogen (Reid 1979). The C:N ratios of each medium are stated in Table 4.1.

Table 4.1 The C:N ratios of the different media used.

Composition (g/L)
Medium C:N ratios
Glucose NH4Cl NH4NO3
Waksman 6 2.0 0.5 -
Fahy 76 25.0 0.5 -
Fujita 114 10.0 - 0.1

Growth medium (200 mL) with the addition of NOM 3 preparation (final concentration of
100 mg C/L) was inoculated with a spore suspension of P. chrysosporium. The fungal
cultures were then incubated at 36oC and 130 rpm. All cultures were performed in duplicate.

50 5.0
Waksman
Decolourisation at 446nm (%)

Glucose consumption (g/L)

40 4.0 Fahy
Fujita
30 3.0

20 2.0

10 1.0

0 0.0
0 2 4 6 8 10 12 14 0 2 4 6 8 10 12 14

Incubation period (days) Incubation period (days)

A B

Figure 4.11 Decolourisation and glucose consumption in the different growth media
containing 100 mg C/L NOM and P. chrysosporium ATCC 34541.

Maximum colour removal over the 14-day incubation period occurred on day 10 with 41%,
32% and 29% removal for Waksman, Fahy and Fujita medium, respectively (Figure 4.11A).
Although the extent of decolourisation for Fahy and Fujita medium was lower than for
Waksman medium, they consumed more glucose (2.54 g/L and 3.54 g/L versus 1.14 g/L for
Waksman medium by day 10) (Figure 4.11B). Decolourisation started to decline after day 10,

55
yet the fungus continued to consume glucose, especially in Fahy and Fujita medium. The
decrease in decolourisation may be due to desorption of the NOM from the fungal cell wall.

Incubation of P. chrysosporium in Waksman medium, which contained the lowest glucose

content, generated the highest biomass and achieved the greatest NOM removal. Fahy
medium had slightly higher NOM removal and biomass production compared with Fujita
medium (Figure 4.12). The fungal pellets generated in the three culture media had a brown
appearance indicating involvement of NOM adsorption in the decolourisation of NOM. In a
study by Carliele et al. (2001), it was demonstrated that carbon and nitrogen ratios of 10:1 or
less were optimum for the growth of fungi. This is consistent with the present results where
the lower the C:N ratio, the greater biomass formation and thus higher NOM removal
achieved. Rojek (2003) also demonstrated that the removal of NOM increased with decreased
C:N ratio although a different NOM preparation was used. However, as a different NOM
preparation was employed there was a little lower decolourisation (41% cf. 47%) even though
a little lower C:N ratio was used (6 cf. 7.90) compared with the results of Rojek (2003).

120
Biomass
40
100 NOM removal
Dry weight biomass (mg)

NOM removal (mg/L)

80 30

60
20
40

10
20

0 0
Waksman Fahy Fujita
Culture m edium

Figure 4.12 Dry weight biomass (mg) and NOM removal in terms of A446 (converted to
mg/L) by P. chrysosporium ATCC 34541 in different culture media after 14 days.

The synthesis efficiency (yield), which is dried mycelium weight divided by weight of carbon
source used, for Waksman, Fahy and Fujita medium was 0.36, 0.13 and 0.10, respectively.
Thus, it is more efficient to use the medium with the lowest glucose content (Waksman) as it
was apparently not limiting to growth, and gave the greatest NOM removal both in absolute
terms and per unit of glucose consumed. Thus, Waksman medium with the C:N ratio of 6 was
selected as the growth medium in the following experiments.

56
4.5 Selection of Organism
As removals of NOM with P. chrysosporium ATCC 34541 were low, and little
biodegradation occurred, this organism was compared with P. chrysosporium strain ATCC
24725, another species of white-rot fungus (Trametes versicolor ATCC 7731), and three
strains of yeast (Saccharomyces species 1, 2 and 3), which had been isolated from a bottle of
MIEX concentrate. As the aim of this study was to develop a simple system for NOM
removals from solution within a short time frame, 5 or 7-day fermentations were investigated.

Waksman medium (200 mL) with the addition of the NOM 3 preparation (final concentration
of 100 mg C/L) was inoculated with either 10 mL spore suspension of white-rot fungi or a
loopful of yeast from a MEA culture. The cultures were then incubated at 130 rpm and 36oC
(white-rot fungi) or 30o C (the yeast). The yeast used had been shown to decolourise NOM
solutions in a preliminary experiment conducted in the laboratory at RMIT.
P. chrysosporium ATCC 34541 sp. 1
80 80
P. chrysosporium ATCC 24725 sp. 2
Decolourisation at 446 nm (%)

Decolourisation at 446 nm (%)

T. versicolor ATCC 7731 sp. 3

60 60

40 40
`

20 20

0 0
0 1 2 3 4 5 0 1 2 3 4 5 6 7
A Incubation period (days) B Incubation period (days)

Figure 4.13 Decolourisation of the NOM (100 mg C/L initial concentration) in

Waksman medium by (A) white-rot fungi and (B) Saccharomyces sp.

Cultures with white-rot fungi and the yeast were incubated for five days and seven days
respectively and monitored for decolourisation of NOM (Figure 4.13). Of the white-rot fungi,
T. versicolor achieved the highest colour removal. Decolourisation increased rapidly until it
reached a plateau on day 3, at which 59% colour removal was attained. The two strains of P.
chrysosporium displayed comparable colour reduction (37%), which plateaued after day 2
(Figure 4.13A). Saccharomyces sp. 1 and 3 seemed to have little potential for the removal of
NOM (Figure 4.13B). Saccharomyces sp. 2 gave similar high colour reduction to T.
versicolor, however, the specific removal values differed markedly: 0.055 compared to 0.089
mg NOM/mg biomass, respectively.
57
 250 2.50 Biomass

Glucose consumption (g/L)

Glucose
Biomass dry weignt (mg) 200 2.00 consumption

150 1.50

100 1.00

50 0.50

0 0.00
Species 1 Species 2 Species 3 T. versicolor
ATCC 7731
Organism

Figure 4.14 Biomass dry weight (mg) and glucose consumptions (g/L) of
Saccharomyces spp. 1-3 and T. versicolor, Waksman medium with 2 g/L initial glucose
content.

Saccharomyces sp. 2 and T. versicolor consumed comparable amounts of glucose with 0.031
mg NOM removed per unit glucose consumed. Saccharomyces sp. 1 and 3 gave 0.015 and
0.013 mg NOM removal per unit glucose consumed, respectively. Saccharomyces sp. 2
produced the highest biomass whereas Saccharomyces sp. 1 provided the least biomass
(Figure 4.14).

The colour removal by the Saccharomyces species was attributed predominantly to adsorption
as indicated by the deep brown colouration of the biomass for all species (see Figure 4.15 for
example), whereas the T. versicolor was light brown in colour (Figure 4.18B). The yeast
removed little UV-absorbing NOM as shown in the HPSEC chromatograms in Figure 4.16;
there was only a small reduction in the high molecular weight range and no formation of
lower molecular weight materials. This was consistent with the NOM removal measured as
reduction in A446 (Figure 4.13B) being primarily due to adsorption rather than biodegradation.
This biosorption, combined with the difficulty experienced in getting consistent results with
the yeast, led to no further investigation of the yeast.

58
 A B

Figure 4.15 Biomass of Saccharomyces sp. 2 (A) incubated in the absence of NOM and
(B) after incubation with 100 mg C/L NOM.

control-NOM
0.500
control-Saccharomyces sp. 2
0.450
Saccharomyces sp. 2
0.400
UV Abs @ 260 nm

0.350

0.300

0.250

0.200

0.150

0.100

0.050

0.000
100 1000 10000 100000

Apparent m olecular w eight (Dalton)

Figure 4.16 HPSEC chromatograms for NOM remaining after treatment with
Saccharomyces sp. 2 in Waksman medium for seven days (control-NOM: culture in the
absence of the yeast; control-Saccharomyces sp. 2: culture in the absence of NOM).

Plots of NOM removal (A446 and A254, converted to mg), glucose consumption (g/L) and
biomass generated (mg) for the three white-rot fungi were constructed for comparison (Figure
4.17). T. versicolor attained the highest reduction in colour and UV-absorbing components
whereas the two strains of P. chrysosporium exhibited similar NOM removals. NOM
removals in terms of colour and UV-absorbing species achieved by T. versicolor were
approximately 62% and 80% greater, respectively, than for both strains of P. chrysosporium.
The ratios of A254/A446 removal for the two strains of P. chrysosporium were similar (0.25)
whereas for T. versicolor it was slightly higher (0.29).

59
T. versicolor consumed all glucose supplied, unlike both strains of P. chrysosporium (Figure
4.17). The yields (Yx/s) for P. chrysosporium ATCC 34541 and ATCC 24725 and T.
versicolor were 0.32, 0.29 and 0.35, respectively, indicating that T. versicolor was more
efficient in generating biomass under these conditions. It was observed that the P.
chrysosporium pellets were deep brown (Figure 4.18A) whereas the T. versicolor pellets were
slightly lighter in colour (Figure 4.18B).

16.0 4.0
A446 A254
NOM removal (A 446, mg)

NOM removal (A 254, mg)

12.0 3.0

8.0 2.0

4.0 1.0

0.0 0.0

2.50 150
Glucose
consumption
Biomass
Glucose consumption (g/L)

2.00
Dry weight biomass (mg)

100
1.50

1.00
50

0.50

0.00 0
P. chrysosporium P. chrysosporium T. versicolor
ATCC 34541 ATCC 24725 ATCC 7751

Figure 4.17 Comparison of NOM removal (as mg, converted from A446 and A254),
glucose consumption and biomass for the three white-rot fungi at initial concentrations
of 2 g/L glucose and 100 mg C/L NOM after five days.

60
 A B

Figure 4.18 Biomass of (A) P. chrysosporium ATCC 34541 and (B) T. versicolor in the
absence of NOM (top), and after five days incubation with 100 mg C/L NOM (bottom).

0.300

0.250

0.200
UV Abs @ 260 nm

0.150

0.100

0.050

0.000
100 1000 10000 100000
Apparent m olecular w eight (Dalton)

control-NOM control-P. chry 34541 + P. chry 34541

control-T. ver 7731 + T.ver 7731

Figure 4.19 HPSEC chromatograms for NOM treated with P. chrysosporium ATCC
34541 and T. versicolor ATCC 7731 (control-NOM: culture in the absence of the fungi;
control-P. chry 34541 or control-T. ver 7731: culture in the absence of NOM).

61
HPSEC analysis was performed to determine any changes in the molecular weight
distribution of the UV-absorbing species of the NOM remaining after treatment. A shift from
high molecular weight towards lower molecular weight species was observed for both fungal
species and was most marked for T. versicolor. T. versicolor gave greater degradation of the
high molecular weight compounds. The absorbance of the peak at ~1500 Dalton for both P.
chrysosporium and T. versicolor cultures was reduced by approximately 0.03 cm-1 and 0.10
cm-1, respectively, after NOM treatment. The absorbance of the low molecular weight
compounds for both cultures was increased after the treatment, indicating that biodegradation
of NOM had occurred to form a pool of low molecular weight compounds; this was greater
for the T. versicolor culture. There were two new peaks formed at 700 and 1000 Dalton after
the treatment with T. versicolor, again indicating the greater breakdown of the high molecular
weight NOM, and suggesting a possible mechanism of sequential breakdown of the larger
NOM molecules via molecules of intermediate size (Figure 4.19).

2500
Mw

Mn
Molecular weight (Dalton)

2000

1500

1000

500

0
control-NOM + P. chry + T. ver 7731
34541

Figure 4.20 Weight average molecular weight (Mw) and number average molecular
weight (Mn) for the NOM (control) and the NOM remaining after five days treatment
with P. chrysosporium ATCC 34541 or T. versicolor ATCC 7731.

Mw and Mn were calculated after five days treatment of the NOM with P. chrysosporium and
T. versicolor. The greatest reductions in Mw and Mn were obtained for the T. versicolor
culture, 25% and 30%, respectively. There were lower reductions in Mw and Mn for the P.
chrysosporium culture, approximately 13% (Figure 4.20). This is consistent with the
outcomes of the molecular weight distribution (Figure 4.19) for the NOM treated with T.
versicolor where there was apparent sequential breakdown of the larger NOM molecules via
molecules of intermediate size.

62
It was considered that the extracellular phenoloxidase enzymes (LiP, MnP and Lac) may have
been involved in the biodegradation of NOM by T. versicolor. Consequently, a spot test on a
colony on an agar plate was undertaken.

A reddish brown zone was visible on the T. versicolor colony when 0.02% guaiacol was
added indicating the presence of laccase as laccase catalyses the oxidative polymerisation of
guaiacol to form brown tetraguaiacone (Figure 4.21).

0.02% guaiacol

Figure 4.21 Reaction of guaiacol on T. versicolor agar plate colony indicating presence
of the laccase enzyme.

To investigate this further the activity of LiP, MnP and Lac for the three white-rot fungi was
assayed on day 3 (fermentations in Figure 4.13A) to determine if they were involved in the
removal of the NOM (Figure 4.22).

4.0
LiP
MnP
Enzyme activity (U/L)

3.0 Laccase

2.0

1.0

0.0
P. chrysosporium P. chrysosporium T. versicolor
ATCC 34541 ATCC 24725 ATCC 7731

Figure 4.22 Activity of the extracellular phenoloxidase enzymes in 3-day cultures of

the three white-rot fungi.

63
LiP was the main enzyme secreted by P. chrysosporium, where strains ATCC 34541 and
ATCC 24725 produced 2.4 U/L and 1.5 U/L LiP, respectively; very-low activities of MnP and
Lac were found. Interestingly, Lac activity was detected in the cultures of P.chrysosporium
under the conditions used. Moreover, it is generally accepted that this fungus does not
produce laccase. The detected activity may be attributed to the non-specificity of the substrate
(guaiacol) used for the enzyme assays. The very low level of LiP activity detected for the P.
chrysosporium cultures was most likely due to low levels of dissolved oxygen as the system
requires high dissolved O2 for lignin decomposition (Kirk et al. 1978). There was little MnP
activity, this may have been due to low levels of Mn2+ in the liquid cultures inhibiting MnP
production, as Mn2+ can stimulate MnP production and functions as a substrate for MnP (ten
Have & Teunissen 2001). However, Rojek et al. (2004) reported that addition of Mn2+ (1.2
mg/L final concentration) did not result in greater decolourisation and so MnP may not be
involved in the process or sufficient Mn was already present in the culture medium.

For P. chrysosporium, LiP was the major enzyme that caused biodegradation of the NOM.
This enzyme has the ability to partly depolymerise and cleave Cα-Cβ linkages in side chains.
A study by Dehorter and Blondeau (1992) reported that P. chrysosporium produced 0.4 U/mL
and 1.1 U/mL LiP and MnP, respectively, to achieve high decolourisation (~60%) of 0.05%
soil humic acids after a 5-day treatment. Another study stated that 1.2 U/mL LiP was
produced by P. chrysosporium for pentachlorophenol (PCP) degradation with no mention of
MnP and Lac activities (Shim & Kawamoto 2002). However, the small magnitude of LiP
activity (0.002 U/mL) for the two strains of P. chrysosporium in the present study, compared
with the findings of other researchers, probably led to the low degree of NOM removal by
biodegradation. Thus, this also verified that P. chrysosporium removed the NOM primarily by
adsorption as indicated by the deep brown colouration of the biomass (Figure 4.18A), rather
than enzymatically oxidised via ligninolytic enzymes.

T. versicolor had high Lac activity compared with LiP and MnP activity. The high NOM
removal attained by T. versicolor was attributed to the laccase, although the pellets were light
brown in colour. The HPSEC pattern for T. versicolor suggests that the high Lac activity
allowed it to break down a higher proportion of the high molecular weight compounds than P.
chrysosporium and so form lower molecular weight molecules including two peaks at 700 and
1000 Dalton. Furthermore, this also indicates that laccase was able to break different types of
bonds compared with LiP, such as demethylation of methoxy groups and catalysing Cα-Cβ
cleavage of phenolic moieties via phenoxy radicals (Kawai et al. 1988).

64
From the results, T. versicolor has more potential for breaking down the NOM while P.
chrysosporium removed the NOM primarily by adsorption under the conditions studied.
Consequently, T. versicolor was selected for further investigation. Enhancement of NOM
removal was studied by determining the effects of supplements, pH, temperature and initial
NOM concentration on fungal growth and extracellular enzyme activities, as described in
Chapter 5.

65
Chapter 5 Biodegradation of NOM by Trametes versicolor
Of the organisms investigated in Chapter 4, T. versicolor was the most effective for the
biodegradation of NOM and gave the greatest NOM removal under the conditions used.
Therefore, further investigation was undertaken into the conditions to improve growth and
enhance the extracellular enzyme activities of T. versicolor to enhance NOM removal. This
was followed by investigation of the enzymatic treatment of the concentrated NOM in vitro.

5.1 Improving NOM Removal by Altering Culture Conditions

To improve NOM removal from solution by T. versicolor the impacts of incubation
temperature, carbon source level, type of inoculum and NOM concentration were
investigated.

5.1.1 Incubation temperature

As most studies used 30oC (Mehna et al. 1995, Swamy & Ramsay 1999, Lorenzo et al. 2002,
Rancaño et al. 2003, Dodor et al. 2004) as the incubation temperature for T. versicolor
cultures, a comparison of the effect of the temperatures 30oC and 36oC on NOM removal was
undertaken.

Cultures containing 200 mL Waksman medium (2 g/L glucose) and filter sterilised NOM
(final concentration of 100 mg C/L) were prepared and inoculated with fungal spore
suspensions, and incubated at 30oC or 36oC and 130 rpm for nine days. All fermentations
were performed in duplicate. Data points correspond to the average of replicates.

The pH and glucose consumption trends for the two conditions differed markedly (Figure
5.1). The pH dropped to almost 3 on day 2 for the cultures at 36oC whereas this occurred on
day 4-5 for the cultures at 30oC. The cultures incubated at 36oC consumed all the glucose
within the first three days. At 30oC, glucose consumption was slow until day 5, after which
there was a major increase such that it was exhausted by day 9.

The performance of the fungus in terms of NOM removal measured at both 446 nm and 254
nm was greater at the higher than the lower temperature over the first three days, after which
it plateaued. This plateau coincided with the exhaustion of the glucose, where 50% and 20%
reductions in A446 and A254, respectively, were obtained for the cultures at 36oC. Rapid NOM
66
removal and then cessation coinciding with the rapid and then cessation of glucose
consumption suggests that either NOM removal is linked with glucose consumption/growth
and/or to biosorption at 36oC. In contrast, there were only slight reductions in both A446 and
A254 until day 4 at 30oC, which then increased to give 73% and 55% reductions in both A446
and A254, respectively, by day 9. The slow reductions within the first four days at 30oC were
probably due to initial adsorption or uptake of NOM, which was followed by enzyme
induction and thus breakdown of NOM.

6.0 6.0 pH
0.40 0.40
5.0 5.0 Glucose
consumption
consumption (g/L)

consumption (g/L)
A446
pH OR Glucose

4.0 0.30 pH OR Glucose 4.0 0.30

Absorbance

Absorbance
A254
3.0 3.0
0.20 0.20
2.0 2.0

0.10 0.10
1.0 1.0

0.0 0.00 0.0 0.00

0 2 4 6 8 0 2 4 6 8

Incubation period (days) Incubation period (days)

30oC 36oC

Figure 5.1 History plots for T. versicolor cultures containing 100 mg C/L NOM
incubated at 30oC and 36oC, Waksman medium 2 g/L initial glucose.

The NOM removal efficiency in terms of colour was higher at 36oC than at 30oC, 0.075 cf.
0.062 mg NOM removed/mg biomass (Figure 5.2). The high specific colour removed at 36oC
was at least partially due to adsorption as indicated by the brown colouration of the biomass
(Figure 5.3B) compared to the cultures at 30oC, where the pellets were cream in colour
(Figure 5.3A). This supports the findings in Figure 5.1 where biosorption was partially
involved in the NOM removal at 36oC. The specific removal of UV-absorbing components at
30oC (0.046 mg NOM removed/mg biomass) was markedly better than that at 36oC (0.030 mg
NOM removed/mg biomass). The ratio A254/A446 at 30oC was higher than at 36oC, 0.75 cf.
0.40, suggesting that the fungus removed/cleaved a higher proportion of molecules containing
conjugated bonds at the lower temperature.

Less biomass was produced at 36oC than at 30oC (Figure 5.2), with the yields (Yx/s, biomass
per unit glucose consumed) of 0.33 and 0.60 respectively. This further indicates that 30oC is a

67
more appropriate incubation temperature for T. versicolor. Growth of organisms at elevated
temperatures results in the inhibition of fungal growth and possibly inhibition of or
denaturation of enzymes.

16 300
A446
14 A254
250
12 Biomass
NOM removal (mg)

200
10

Biomass (mg)
8 150

6
100
4
50
2

0 0
30oC 36oC

Incubation tem perature

Figure 5.2 NOM removals (as mg, converted from A446 and A254) and biomass
produced (mg) in T. versicolor cultures at 30oC and 36oC.

A B

Figure 5.3 Biomass of T. versicolor incubated at (A) 30oC and (B) 36oC in the absence
of (top) and presence of (bottom) NOM after nine days incubation, 100 mg C/L NOM.

There are clear differences in the behaviour and performance of the organism at the different
temperatures. The behaviour of T. versicolor at 36oC was very similar to that of P.

68
chrysosporium (Figure 4.1), in that the fast NOM removal rate over the first three days
followed by a plateau indicated that the mechanism of NOM decolourisation was primarily
biosorption, as confirmed by the brown colouration of the fungal pellets. This is consistent
with the findings of Rojek (2003), who reported the same trends for P. chrysosporium ATCC
34541 and established that adsorption contributed 60-70% of the overall removal of NOM.
On the contrary, the initial slow and then higher rate of NOM removal with T. versicolor at
30oC was attributed to the production of oxidative enzymes during secondary metabolism.
The lack of brown colouration of the fungal pellets (Figure 5.3A) supports this premise of
NOM removal by biodegradation rather than adsorption. The trends in reduction in A446 and
A254 can be related to the activities of the extracellular phenoloxidase enzymes (Figure 5.4).

18
LiP
16 MnP
14 Laccase
Enzyme activity (U/L)

12

10

0
30oC 36oC
Incubation tem perature

Figure 5.4 Activity of the extracellular phenoloxidase enzymes of the T. versicolor

cultures incubated at 30oC and 36oC.

The low production of the phenoloxidase enzymes at 36oC was very apparent. The activity of
the LiP, MnP and Lac enzymes was much higher at 30oC, as was the removal of NOM. This,
in combination with the different patterns for pH, A446 and A254 reductions and glucose
consumption indicates that these enzymes played a major role in NOM removal. The
reduction in A254 indicates that conjugated bonds and aromatic rings were enzymatically
broken. However, the low activity of LiP in both cultures suggests that this enzyme plays only
a minor role in NOM removal for T. versicolor under the conditions employed. The ratio of
LiP to Lac activity for both cultures was similar (~0.075), while the ratio of MnP to Lac
activity at 30oC was markedly higher than that at 36oC, 0.358 cf. 0.082, suggesting that MnP
and Lac are the main enzymes involved in the removal of NOM at 30oC under the conditions
studied. The high activities of MnP and Lac obtained in this culture would be responsible for
69
the cleavage of aromatic rings, conjugated bonds and Cα-Cβ bonds in phenolic moieties, as
well as catalysing alkyl-aryl cleavage in the NOM structures. The lignin-degrading enzymes
may also be involved in the breakdown of β-O-4 and β-1 linkages that occur within the lignin
polymer.

Although two studies (Blondeau 1989, Dehorter & Blondeau 1992) reported that LiP and
MnP were associated with humic acid degradation, the resistance of humic substances to
microbial decolourisation is largely related to the differences in their chemical structures and
is microbial species dependent, as suggested by Yanagi et al. (2002), and is dependent upon
the culture conditions. Moreover, Dehorter and Blondeau (1992) established that MnP rather
than LiP was the major enzyme involved in the microbial degradation of different
concentrations of humic acid with T. versicolor. The high levels of MnP and Lac observed for
T. versicolor at 30oC are consistent with the findings of Galliano et al. (1991), who reported
that these enzymes worked synergistically in the degradation of lignin. They reported that
when these two enzymes were isolated and purified from Rigidoporus lignosus, neither was
able to solubilise lignin. However, degradation of lignin occurred when the two enzymes were
added to the reaction medium simultaneously.

These findings support the suggestion that removal of NOM by T. versicolor incubated at
30oC was mainly due to enzymatic breakdown whereas at 36oC it was probably removed by
different mechanisms such as: chemical and physical sorption, metabolically dependent
sorption and accumulation, and biodegradation as reported by Rojek (2003) in relation to the
NOM removal with P. chrysosporium.

The impact of the NOM-degrading enzymes on the molecular weight distribution of the UV-
absorbing species for the NOM remaining after nine days treatment with T. versicolor at 30oC
and 36oC was determined (Figure 5.5).

A shift from high molecular weight towards lower molecular weight species was observed for
both temperatures. It was observed that low molecular weight compounds for the culture at
36oC were formed from the breakdown of the high molecular weight compounds. However T.
versicolor at 30oC was markedly more effective in removing and converting the high
molecular weight UV-absorbing species to lower molecular weight compounds. The fungus at
30oC was able to remove compounds of apparent molecular weight smaller than 1000 Dalton,
and the first peak at 350 Dalton remained similar or reduced slightly, indicating that either the

70
low molecular weight species were produced (from the breakdown of the high molecular
weight molecules) and were removed simultaneously probably by metabolism or, that the low
molecular weight species were not UV-absorbing. As noted previously, the ability of T.
versicolor to degrade NOM was greater at 30oC than at 36oC probably due to its greater
production of NOM-degrading enzymes, especially MnP and Lac, where MnP is presumably
responsible for the removal of low molecular weight species. Thus, MnP and Lac may act
synergistically in the enzymatic breakdown of medium molecular weight (500-2000 Dalton)
fulvic acids and high molecular weight (2000-5000 Dalton) humic acids.

0.300

0.250
UV Abs @ 260 nm

0.200

0.150

0.100

0.050

0.000
100 1000 10000 100000

Apparent m olecular w eight (Dalton)

control-36oC 36oC

NOM 3 30oC
control-30oC

Figure 5.5 HPSEC chromatograms for NOM remaining after treatment with T.
versicolor incubated at 30oC and 36oC; controls represent fungal cultures grown in the
absence of NOM.

The greater reduction in Mw and Mn for the T. versicolor cultures at 30oC further
demonstrates that the biodegradation of NOM at lower temperature was markedly more
effective than at higher temperature (Figure 5.6). A reduction of 40% in both Mw and Mn was
achieved for T. versicolor at 30oC, suggesting that the NOM removal was via enzymic
breakdown, which was probably followed by fungal metabolism.

71
 2500 Mw

Mn
Molecular weight (Dalton) 2000

1500

1000

500

0
control-NOM T. ver 36oC T. ver 30oC

Figure 5.6 Weight average molecular weight (Mw) and number average molecular
weight (Mn) for the NOM (control) and the NOM remaining after nine days treatment
with T. versicolor ATCC 7731 at 30oC and 36oC.

5.1.2 Carbon source level

This experiment was performed as the glucose provided in the preceding experiment was
exhausted after nine days at 30oC. A study by Rojek (2003) suggested that glucose exhaustion
could have an inhibitory effect on NOM decolourisation by P. chrysosporium ATCC 34541.
Thus a higher initial glucose concentration (5 g/L) was supplied to see if this would increase
fungal growth, secretion of ligninolytic enzymes and NOM removal with T. versicolor at
30oC.

Waksman medium (200 mL) containing either 2 or 5 g/L glucose and 100 mg C/L NOM was
inoculated with T. versicolor as spore suspensions and incubated at 30oC and 130 rpm for
nine days. The history plot for the culture with 2 g/L glucose was obtained from Figure 5.1,
the fermentation containing 5 g/L glucose was conducted in duplicate and the average values
obtained.

The pH trends for both cultures at different initial glucose concentrations were comparable,
where the pH dropped to 3 on day 4-5 (Figure 5.7). Although the pH dropped to 3, it did not
seem to be associated with long-term adsorption of NOM to biomass when the phenoloxidase
enzymes were active. The rate of glucose consumption was low initially and increased after
day 5 for 2 g/L initial glucose, whereas it was greater before and then decreased after day 5
for 5 g/L initial glucose cultures.

72
As discussed in Section 5.1.1 microbial breakdown of NOM occurred in the T. versicolor
culture with 2 g/L glucose at 30oC, as the rate of NOM removal (which was slow initially and
then increased markedly after day 4) was correlated with the induction of NOM-degrading
enzymes. With 5 g/L initial glucose content, there were only slight reductions in A446 and A254
for the first three days, greater reductions occurred during day 3-4, and then both absorbances
stayed constant. The sharp absorbance drop occurred at the same time as the pH drop for the
cultures with 5 g/L initial glucose, indicating possible biosorption of the NOM by the fungal
cell walls. This was confirmed by the brown colouration of the biomass. The T. versicolor
preferentially consumed glucose rather than NOM in the culture, as glucose is a more easily
obtained source of carbon than NOM. If the fermentation were left longer, it is possible that
once the glucose was depleted then the NOM would be utilised as a carbon source and so
removed from solution and possibly from the surface of the biomass.

6.0 6.0
pH
0.40 0.40
5.0 5.0 Glucose
consumption
A446
consumption (g/L)
consumption (g/L)

pH OR Glucose
pH OR Glucose

4.0 0.30 4.0 0.30

Absorbance
Absorbance

A254
3.0 3.0
0.20 0.20
2.0 ` 2.0
0.10 0.10
1.0 1.0

0.0 0.00 0.0 0.00

0 2 4 6 8 0 2 4 6 8
Incubation period (days) Incubation period (days)

2 g/L 5 g/L

Figure 5.7 History plots for T. versicolor cultures containing 100 mg C/L NOM
incubated at 30oC, Waksman medium containing 2 g/L and 5 g/L initial glucose.

Increasing the glucose content to 5 g/L resulted in higher biomass production (Figure 5.8).
The yields (Yx/s) for the 2 g/L and 5 g/L glucose cultures were 0.60 and 0.65 respectively.
Although the yield was higher for the culture with the higher initial glucose concentration, the
fungus did not attack the NOM until the glucose was depleted. Higher glucose concentration
led to lower NOM removal efficiencies in terms of A446 and A254, which were 0.026 and 0.012
mg NOM removed/mg biomass respectively. The ratio of A254 to A446 for the culture with
higher glucose concentration was lower, 0.45 cf. 0.75, demonstrating that this culture
removed a higher proportion of coloured than the conjugated/aromatic species.

73
 20 400
A446
A254
NOM removal (mg) 15 300 Biomass

Biomass (mg)
10 200

5 100

0 0
2 g/L 5 g/L

Glucose level

Figure 5.8 NOM removals (as mg, converted from A446 and A254) and biomass
produced (mg) in T. versicolor cultures containing 2 g/L and 5 g/L initial glucose.

Enzyme activities were lower in the presence of the higher glucose concentration (Figure 5.9).
Lac activity was markedly lower, especially in relation to MnP activity, for the higher glucose
concentration. The low levels of these enzymes in the culture resulted in the inability of the
fungus to break down the NOM. These further support the argument that the enzymatic
activity is enhanced upon nutrient starvation (as established in the culture with the lower
initial glucose content), which is known to contribute to the secondary metabolism during
which the white-rot fungi produce the ligninolytic enzymes, and thus resulting to higher
enzymatic removal of NOM.

20.0
LiP
MnP
16.0 Laccase
Enzyme activity (U/L)

12.0

8.0

4.0

0.0
2 g/L 5 g/L

Glucose level

Figure 5.9 Activity of the extracellular phenoloxidase enzymes of the T. versicolor

cultures containing 2 g/L and 5 g/L initial glucose.

74
5.1.3 Types of inoculum
Spore collection for T. versicolor was very tedious, as approximately 20-30 agar plates with
an actively growing fungal colony were needed to achieve a concentration of 1.0-1.5 x 105
spores/mL in each culture. Hence a comparison of NOM removal using inoculation by spores
and by plugs was undertaken as the latter is a much easier method.

Cultures containing 200 mL Waksman medium (2 g/L glucose) and sterilised NOM (final
concentration of 100 mg C/L) were prepared and inoculated with the fungus as either spore
suspensions or as three MEA plugs (each 1 cm2) of T. versicolor and incubated at 30oC and
130 rpm for eight days. All fermentations were performed in duplicate.

The pH, glucose consumption, and A446 and A254 trends for both culture types were similar
(Figure 5.10). With plugs the pH dropped to pH 3.5 cf. pH 3.0 for the spore culture. The
fungus in the plug culture did not consume glucose for the first two days, as there was some
readily assimilable carbon available in the MEA plugs. The major decrease in A446 and A254
for both cultures occurred after day 4, presumably due to the production and action of
ligninolytic enzymes involved in the degradation of NOM. After eight days the biomass was
collected and dried and it appeared that the biomass on the plugs and the biomass pellets
formed were creamy brown in colour, indicating that the removal of NOM was not due to
adsorption to the biomass.

6.00 0.40 6.00 0.40 pH

5.00 5.00 Glucose

0.30 0.30 consumption
consumption (g/L)

consumption (g/L)
pH OR Glucose

pH OR Glucose

4.00 4.00 A446

Absorbance

Absorbance

3.00 0.20 A254

3.00 0.20

2.00 2.00
0.10 0.10
1.00 1.00

0.00 0.00 0.00 0.00

0 2 4 6 8 0 2 4 6 8
Incubation period (days) Incubation period (days)
Plugs Spores

Figure 5.10 History plots for T. versicolor cultures in Waksman medium (2 g/L
glucose) containing 100 mg C/L NOM incubated at 30oC, inoculated with either spore
suspensions or three agar plugs.

75
To determine if there was adsorption of NOM to the agar component of the MEA plugs an
experiment was carried out where three MEA plugs (each 1 cm2) without the fungus were
added to 200 mL Waksman medium (2 g/L glucose) and sterilised NOM (final concentration
of 100 mg C/L). The flask was then incubated at 30oC and 130 rpm for eight days.

The A446 and A254 stayed constant throughout the incubation period (Figure 5.11). This
verified that there was no NOM adsorption to the agar, nor detectable leaching of brown-
coloured material from the plugs.

0.40
A446

A254
0.30
Absorbance

0.20

0.10

0.00
0 2 4 6 8

Control Incubation period (days)

Figure 5.11 A446 and A254 for Waksman medium (2 g/L glucose) containing 100 mg
C/L NOM and three agar plugs without fungus.

The total NOM removed (mg) in terms of both coloured and UV-absorbing species for
cultures with plug inoculation was less, although a higher level of Lac activity was obtained
(Figure 5.12). The ratio of A254 to A446 removal for the plug cultures was lower than for the
spore cultures, 0.56 and 0.71, respectively.

Although the degree of NOM removal (measured as decolourisation) was 8% greater when
the spore suspension inoculum was used, subsequent investigations were conducted using the
plug inoculation method, as it was much simpler. Furthermore, plug inoculation led to higher
enzyme activity than spore inoculation, even though there was a longer lag phase for the
enzyme secretion, and thus provides better potential for the biodegradation of NOM.

76
 14 25
A446

12 A254
20
Laccase

Laccase activity (U/L)

NOM removal (mg)

10

15
8

6
10

4
5
2

0 0
Plugs Spores

Figure 5.12 NOM removals (as mg, converted from A446 and A254) and laccase activity
in T. versicolor cultures, inoculated with either spore suspension or plugs.

5.1.4 Effect of NOM concentration

The effect of initial NOM concentration on NOM degradation was examined in order to
determine the maximum concentration tolerated by the fungus. Cultures containing 200 mL
Waksman medium (2 g/L glucose) and varying NOM concentration (final concentrations of
100, 300, 400, 500, 600 and 700 mg C/L) were prepared and each inoculated with three
fungal plugs and incubated at 30oC and 130 rpm for ten days.

70 2.50 A446

A254
60
2.00 Glucose
Glucose consumption (g/L)

50 consumption
NOM removal (mg)

1.50
40

30
1.00

20
0.50
10

0 0.00
100 300 400 500 600 700
Initial NOM concentration (m g C/L)

Figure 5.13 NOM removals (as mg, converted from A446 and A254), and glucose
consumption in T. versicolor cultures with different NOM concentrations, plug
inoculum.
77
NOM removed, when measured as both A446 and A254, increased with NOM concentration up
to 600 mg C/L and decreased dramatically when the NOM concentration was increased to 700
mg C/L (Figure 5.13). The removal of NOM measured as colour (A446) per unit glucose
consumed increased linearly with NOM concentration up to 600 mg C/L NOM. However, the
removal of UV-absorbing species per unit glucose consumed for 100-300 mg C/L NOM were
approximately the same, increased linearly between 300-600 mg C/L and decreased
considerably at 700 mg C/L NOM.

A linear relationship with good correlation (R2 = 0.9901) was established between initial
NOM concentration (100-600 mg C/L) and NOM removal in terms of A446 (Figure 5.14). This
relationship was in good agreement with the study by Rojek (2003), where the correlation
found for P. chrysosporium in the NOM concentration range of 120-480 mg C/L was
y = 0.095 x − 0.8521 (R2 = 0.998).

70
A446

60 A254

50
NOM removal (mg)

y = 0.0952x + 6.6653
R2 = 0.9901
40

30

20
y = 0.0415x + 3.1796
R2 = 0.9206
10

0
0 200 400 600 800
Initial NOM concentration (m g C/L)

Figure 5.14 NOM removals (as mg, converted from A446 and A254) by T. versicolor for
different NOM concentrations.

A linear correlation between initial NOM content (100-600 mg C/L) and NOM reduction in
terms of A254 was established: y = 0.042 x + 3.180 (R2 = 0.9206) (Figure 5.14). This differs
from the observations of Rojek (2003), who reported that there was a similar extent of UV-
absorbing NOM removed (approximately 15%) in all cultures and thus no relationship
between A254 and initial NOM content was established. However, a different NOM
concentrate and fungus was used in that study, which may account for the different result. It

78
should be noted that both relationships obtained in the present study hold only for the NOM
concentration range of 100-600 mg C/L.

The pattern of decolourisation and removal of UV-absorbing species increasing with initial
NOM concentration can be correlated with the production of extracellular ligninolytic
enzymes at different NOM contents (Figure 5.15). The Lac activity increased with NOM
concentration up to 600 mg C/L, however, both LiP and MnP activities decreased with NOM
content, indeed, there was no LiP activity when the NOM concentration exceeded 300 mg
C/L. The culture with 700 mg C/L initial NOM content gave a markedly lower Lac activity.
Dehorter and Blondeau (1992) illustrated a relationship between humic acid degradation and
extracellular enzyme activity of P. chrysosporium and T. versicolor, and showed that the
production of extracellular LiP and MnP increased with increasing concentrations of humic
acids. The surfactant properties of the humic acids were suggested to be responsible for the
increase in enzyme activities. However, 1% or higher concentrations of humic acids were
found to be inhibitory to growth and enzyme induction. This is in an agreement with the
present findings, where LiP and MnP activities decreased with NOM concentrations from 300
mg C/L (equivalent to 1.0%) to 700 mg C/L (equivalent to 2.3%).

200
LiP
MnP
160
Laccase
Enzyme activity (U/L)

120

80

40

0
100 300 400 500 600 700
Initial NOM concentration (m g C/L)

Figure 5.15 Activity of the extracellular phenoloxidase enzymes of T. versicolor in

cultures containing varying NOM concentrations, plug inoculum.

As illustrated in Figure 5.14, NOM removal increased linearly with NOM concentration up to
600 mg C/L. This is coincident with the secretion of laccase, where the activity increased until
600 mg C/L NOM. The ratio of A254 to A446 for the 700 mg C/L culture was the lowest (0.23),
79
matching the sharp decrease in Lac activity. Thus, the removal of NOM from solution by T.
versicolor was associated with high Lac activity. Again, the laccase in the cultures was
responsible for breakdown of the NOM, probably by the cleavage of aromatic rings,
conjugated and Cα-Cβ bonds in phenolic moieties, as well as catalysing alkyl-aryl cleavage.

5.2 Enhancement of Enzyme Production

According to the findings in Sections 5.1.1 and 5.1.4, it was established that laccase was the
major extracellular enzyme secreted by T. versicolor and by deduction, involved in NOM
removal. Consequently, the next phase of the study was the determination of the cultivation
period at which maximum Lac activity occurred, and the effects of supplements, temperature
and pH, and agitation.

Wheat bran was employed in this study, as it has been used to supply nutrients for white-rot
fungal growth. Many researchers and industries have applied the solid-state fermentation
(SSF) technique for the production of enzymes and spores by using lignocellulose residues
(Ooijkaas et al. 2000, Pandey et al. 2000, Robinson et al. 2001, Couto & Sanromán 2005).
There has been increased interest in SSF for the development of bioprocesses, such as
bioremediation and biodegradation of hazardous compounds, biological detoxification of
agro-industrial residues, biopulping, etc (Pandey et al. 2000) due to the recent improvements
in reactor designs (Couto & Sanromán 2005).

Medium with higher glucose content (5 g/L) was utilised in this study as previous
experiments showed that T. versicolor did not grow well at lower glucose concentration (2
g/L) without the presence of NOM. NOM was not added to the cultures since preliminary
experiments demonstrated that NOM adsorbed to the wheat bran (Figure 5.16). There was a
15% decrease in A446 although no fungus was cultured in the medium. To prevent adsorption
of NOM to wheat bran, T. versicolor was pre-cultured in the wheat bran-containing Waksman
medium in the absence of NOM to increase the production of ligninolytic enzymes.

80
 0.30

Absorbance at 446 nm 0.25

0.20

0.15

0.10

0.05

0.00
0 2 4 6 8 10 12

Incubation period (days)

Figure 5.16 A446 for 100 mg C/L NOM incubated in Waksman medium containing 4.5
g/L wheat bran in the absence of fungus.

Cultures were established in 250 mL Erlenmeyer flasks containing 100 mL of modified

Waksman medium (5 g/L glucose) supplemented with 4.5 g/L wheat bran (Nyanhongo et al.
2002) or 0.5% (v/v) Tween 80 (Couto et al. 2001), or both, and three agar plugs (each 1 cm2)
of T. versicolor. All fermentations were performed in duplicate.

5.2.1 Determination of cultivation time based on maximum laccase activity

The cultivation period at which maximum Lac activity occurred was determined. This time
was then used for subsequent experiments involving enzyme assays and NOM removal.

The fungus was cultured in modified Waksman medium supplemented with 4.5 g/L wheat
bran at 30oC and 130 rpm. Samples (3 mL) were collected periodically and centrifuged prior
to pH, Lac activity and glucose measurements.

Microbial growth became apparent on day 4 and so the measurements were undertaken
thereafter. The pH dropped markedly until day 7, after which it stayed constant. Lac activity
became apparent from day 4 and peaked on day 9, the increase coinciding with the exhaustion
of glucose on day 8 (Figure 5.17). Therefore the cultivation time for maximum Lac activity of
T. versicolor under the conditions used was established as nine days.

81
 70.0 6.00 Laccase
activity
60.0 5.00 pH

Glucose consumption (g/L)

Laccase activity (U/L)

50.0 Glucose
4.00
consumption
40.0

pH/
3.00
30.0
2.00
20.0

10.0 1.00

0.0 0.00
0 2 4 6 8 10 12 14
Incubation period (days)

Figure 5.17 History plot of T. versicolor cultivated in the presence of 4.5 g/L wheat
bran. The data points correspond to mean values of duplicate assays.

5.2.2 Effect of supplements

Two supplements were employed in this study, viz., Tween 80 and wheat bran. Before
inoculation with T. versicolor plugs, the culture medium without added NOM was
supplemented with 0.5% (v/v) Tween 80 or 4.5 g/L wheat bran, or both, and was then
incubated at 30oC and 130 rpm. After nine days (as established in the preceding experiment),
the cultures were filtered through a fine sieve to remove the fungal plugs and pellets and the
remaining wheat bran, and then were centrifuged at 8500 rpm (12,200 RCF) and 20oC for 15
minutes. The supernatant was frozen and then defrosted and the precipitated polysaccharides
were removed by centrifugation (4400 rpm for 30 minutes) (Nyanhongo et al. 2002). The
resulting clear solution was used for enzyme activity assays.

Table 5.1 indicates the activity of the extracellular phenoloxidase enzymes (LiP, MnP and
Lac) of T. versicolor for the different culture conditions. There was little growth apparent in
cultures in the absence of NOM and supplements; hence these displayed very little enzyme
activity. Addition of wheat bran led to high Lac activity compared with LiP and MnP,
demonstrating its great potential as a laccase inducer in submerged cultures of T. versicolor.
This result is consistent with those reported by Lorenzo et al. (2002), who found that
lignocellulosic materials can stimulate the laccase-producing ability of T. versicolor; this
effect was attributed to the cellulose content of the residues. Furthermore, these residues act
as a source of nutrients to the fungus for secondary metabolism, consequently they have been
termed support-substrates. This suggests that wheat bran gives T. versicolor an environment
82
similar to its natural habitat (wood), which would probably stimulate the fungus to secrete
lignin-degrading enzymes.

Table 5.1 Comparative ligninolytic enzyme activities in different culture media.

Enzyme activities (U/L)
Culture broths
LiP MnP Lac
No supplement - - 0.30
+ 0.5% Tween 80 - - 27.0
+ 4.5 g/L wheat bran 0.65 7.84 60.3
+ 4.5 g/L wheat bran & 0.5% Tween 80 1.83 13.8 955

Addition of Tween 80 alone to the cultures led to a small improvement in Lac activity, being
similar to the effect of the presence of 100 mg C/L NOM (Figure 5.15). However, with the
presence of wheat bran it caused 1.8-fold, 0.8-fold and 15-fold increases in LiP, MnP and Lac
activities, respectively. The cultures with the additions of the two supplements would
probably be able to break conjugated, Cβ-C4 and Cα-Cβ bonds and cleavage aromatic rings of
NOM due to the presence of the three-ligninolytic enzymes, particularly laccase. Asther et al.
(1987) suggested that Tween 80, a non-ionic surfactant, was capable of transforming the cell
membrane structure to promote permeation of LiP from the cell into the medium. Another
study reported that Tween 80 can protect LiP in culture broths against inactivation due to
agitation (Venkatadri & Irvine 1990). These observations are consistent with those of Shim
and Kawamoto (2002) where the addition of Tween 80 to a culture of P. chrysosporium
immobilised on Biolace resulted in a slight increase in LiP activity.

In spite of the increases in the ligninolytic enzyme activities for cultures in the presence of the
two supplements, the activities of LiP and MnP were still low compared with those for the
cultures in the presence of 300 and 400 mg C/L NOM, respectively (Figure 5.15). However,
the combination of the two supplements led to markedly greater activity of laccase compared
with the NOM-containing cultures in the range 100-700 mg C/L. Consequently, wheat bran
and Tween 80 can act synergistically in improving the production of laccase, but not MnP and
LiP, compared with cultures in the presence of NOM.

Determinations of the optimal pH and temperature of laccase secreted by the cultures

supplemented with wheat bran and wheat bran plus Tween 80 were investigated as described
below.
83
5.2.3 Effect of temperature and pH on laccase activity
The effect of temperature on Lac activity was established by performing assays at
temperatures ranging from 25oC to 90oC at pH 4.5. The effect of pH was determined by
measuring the activity at pH varying from 2.5 to 7.0 at the determined optimum temperature
established in the preceding experiments.

The assay mixtures were equilibrated at the different temperatures and pH for 2.5 minutes
before introducing the substrate guaiacol. Lac activity was determined by measuring the
initial velocity of reaction.

(i) Effect of temperature

The activity of the enzyme secreted in the presence of Tween 80 was higher over a broader
temperature range (50-70oC compared with 60-70oC), but decreased rapidly at greater than
70oC compared with the enzyme secreted in the absence of the detergent (Figure 5.18). The
culture in the presence of Tween 80 retained 91-100% Lac activity over the temperature range
of 50-70oC, whereas 93-100% Lac activity was maintained over 60-70oC by the culture in the
absence of the detergent.

80.0 1200
+ WB

1000 + WB + Tw 80
Laccase activity (U/L)
Laccase activity (U/L)

60.0
(+ WB + Tw80)

800
(+ WB)

40.0 600

400
20.0
200

0.0 0
20 30 40 50 60 70 80 90

Tem perature (oC)

Figure 5.18 Effect of temperature on Lac activity at pH 4.5 in cultures supplemented

with 4.5 g/L wheat bran (+WB) and 4.5 g/L wheat bran plus 0.5% Tween 80 (+WB
+Tw80). The data points correspond to mean values of duplicate assays.

50oC was chosen as the temperature for enzyme assays as the enzyme may not be able to
tolerate the higher temperature if incubated for extended periods.

84
(ii) Effect of pH
As for temperature, there was a similar broadening of pH range close to maximum activity for
enzyme secreted in the presence of 0.5% Tween 80, the activity at pH ≥ 5 being markedly
greater for the latter (Figure 5.19). The culture in the absence of Tween 80 retained 100% Lac
activity over the pH range of 4-4.5, whereas 92-100% Lac activity was maintained over pH 4-
5 by the culture in the presence of the detergent.

80.0 1200
+ WB
70.0
1000 + WB + Tw 80
60.0
Laccase activity (U/L)

Laccase activity (U/L)

800

(+ WB + Tw80)
50.0
(+ WB)

40.0 600

30.0
400
20.0
200
10.0

0.0 0
2.0 3.0 4.0 5.0 6.0 7.0 8.0

pH

Figure 5.19 Effect of pH on Lac activities at 50oC in cultures supplemented with 4.5
g/L wheat bran (+WB) and 4.5 g/L wheat bran plus 0.5% Tween 80 (+WB +Tw80). The
data points correspond to mean values of duplicate assays.

The culture fluids were tested neat, and gave results as in Figure 5.18 and Figure 5.19. As the
activity of the wheat bran plus Tween 80 preparation was markedly higher, it was diluted in
buffer to approximately the same activity as the wheat bran preparation, and retested (Figure
5.20). Although the dilution was somewhat greater than anticipated on the basis of
calculation, the results for the diluted enzyme preparation confirmed those for the original
culture fluid, i.e., higher Lac activity over broader temperature (50-70oC cf. 60-70oC) and pH
(4.0-5.0 cf. 4.0-4.5) ranges than the wheat bran preparation.

85
 250 250
+ WB + Tw80
(diluted)
Laccase activity (U/L) 200 200

Laccase activity (U/L)

150 150

100 100

50 50

0 0
20 30 40 50 60 70 80 90 2.0 3.0 4.0 5.0 6.0 7.0 8.0
Temperature (oC) pH

Figure 5.20 Effects of (A) temperature and (B) pH on Lac activities after dilution for
cultures supplemented with 4.5 g/L wheat bran plus 0.5% Tween 80 (+WB +Tw80).

The optimum temperature for the Lac activity from the cultures with wheat bran and wheat
bran plus Tween 80 was determined to be 50oC although the highest activities were obtained
at 70oC (WB preparation) and 60oC (WB+Tw80 preparation) as the Lac activities would
deteriorate at higher temperature. The optimum activity of the enzyme from the two cultures
of T. versicolor was identified as pH 4.0-4.5. The results were consistent with the study by
Coll et al. (1993), who stated that the optimum temperature and pH for Lac activity secreted
by T. versicolor was 50oC and pH 4.0-5.0, respectively.

5.2.4 Effect of agitation

There are several forms of growth that may affect fungal performance, namely filaments,
fungal mats and pellets. It was reported that stationary cultures formed fungal mats whereas
agitation led to pellet formation (Swamy & Ramsay 1999). Agitation provides better aeration,
which in turn may result in greater decolourisation, however, some researchers showed that
agitation could suppress fungal phenoloxidase enzyme activity (Kirk et al. 1978, Kirk &
Farrell 1987).

In this experiment, cultures with three plugs (each 1 cm2) of T. versicolor and different
medium contents (Table 5.2) were prepared and two different incubation conditions were
applied: (A) continuous agitation at 130 rpm and 30oC and (B) agitation every 6 hours for 30
minutes at 130 rpm and 30oC. Condition (B) was selected as reducing agitation may prevent
damage to the mycelium and secreted enzymes caused by shear stress and/or changes in the

86
morphology of the fungus (Rancaño et al. 2003). These conditions may provide a trade-off
between providing good aeration and inhibition of fungal ligninolytic activity. All cultures
were run as duplicates.

Table 5.2 Different medium contents used for T. versicolor cultures for testing effect
of agitation.
Designation Culture content
WM Waksman medium (WM, 5.0 g/L glucose)
WM + Tw80 WM + 0.5% Tween 80
WM + WB WM + 4.5 g/L wheat bran
WM + WB + Tw80 WM + 4.5 g/L wheat bran + 0.5% Tween 80
WM + NOM WM + 500 mg C/L NOM
WM + NOM + Tw80 WM + 500 mg C/L NOM + 0.5% Tween 80

Cultures with continuous agitation (Figure 5.21A) consumed glucose faster than those
agitated every 6 hours for 30 minutes (Figure 5.21B), except for the culture without
supplement. The high glucose consumption rates were probably due to the improved mass
transfer of oxygen and substrates in agitated cultures.

The cultures agitated every 6 hours formed fungal mats on the surface of the medium whereas
continuous agitation led to pellet formation. The level of Lac activity increased with the
addition of Tween 80 for both incubation conditions. Tween 80 appeared to have the ability to
protect enzymes in the culture fluids against mechanical inactivation due to agitation, as
reported by Jäger et al. (1985). In addition, it was reported that Tween 80 acts as a LiP
inducer (Asther et al. 1987) and so may also be effective in stimulating other enzymes such as
laccase. The WM and WM + Tw80 cultures which were agitated every 6 hours for 30 minutes
showed increasing Lac activity with time unlike those under continuous agitation, as less
agitation prevented damage to the mycelium and secreted enzymes caused by shear stress
and/or changes in the morphology of the fungus. However, continuous agitation for the two
cultures resulted in low increase rate of the enzyme activity with time after day 9, which can
be attributed to shear stress.

For the cultures in the presence of wheat bran and wheat bran plus Tween 80, enzyme activity
increased with time for occasional agitation whereas decreased sharply, probably due to
denaturation, after day 9 for the continuously agitated cultures. Again, Tween 80 is able to

87
increase secretion of laccase and/or, protect laccase significantly from inactivation under
agitated conditions as the Lac activity remained high in both conditions with the addition of
the detergent in the presence of wheat bran, especially for the cultures with continuous
agitation.

6.00 1000
WM

5.00
800
Glucose consumption (g/L)

WM+ Tw80

Laccase activity (U/L)

4.00 WM+ WB
600

3.00 WM+ WB + Tw80

400
WM+ NOM
2.00

WM+ NOM+ Tw80

200
1.00

0.00 0
0 2 4 6 8 10 12 14 16 0 2 4 6 8 10 12 14 16
A

6.00 200

5.00
Glucose consumption (g/L)

160
Laccase activity (U/L)

4.00
120

3.00
80
2.00

40
1.00

0.00 0
0 2 4 6 8 10 12 14 16 0 2 4 6 8 10 12 14 16

Incubation period (days) Incubation period (days)

B

Figure 5.21 Glucose consumption (g/L) and Lac activity (U/L) for T. versicolor
cultures in different media and agitation conditions: (A) continuous agitation and (B)
agitated every 6 hours for 30 minutes, both at 30oC and 130 rpm.

Lac activity in the cultures provided with NOM and NOM plus Tween 80 increased with time
for both conditions. Addition of NOM enhances Lac activity as observed in Section 5.1.4
(Figure 5.15) where laccase increases with NOM concentration up to 600 mg C/L. However,
there was no increase in laccase in the presence of Tween 80, indeed, there was a decrease, in
contrast with the induction of laccase by the combination of wheat bran and Tween 80. Thus,
the addition of wheat bran or NOM alone, and the combination of wheat bran plus Tween 80
88
can stimulate the laccase-producing ability of T. versicolor, however, addition of Tween 80
alone did not give significant improvement.

Fermentations with continuous agitation promote higher and faster enzyme activity than those
with occasional agitation because it provides better mass transfer. This is consistent with the
findings of Knapp et al. (1997), who demonstrated that agitation was very effective in
improving the rate of Orange II decolourisation by mycelial pellets of an unidentified
basidiomycete fungus. This may be due to the improved mass transfer of oxygen and
substrates in agitated cultures. However, it seemed that continuous agitation had an adverse
effect on the fungal growth and enzyme activity over longer fermentation periods. Thus
discontinuous agitation like condition (B) would be a better alternative if longer contact time
were required.

1.20 25
A254 WM + NOM
1.00 WM + NOM + Tw 80
20
Absorbance at 254 nm
Absorbance at 446 nm

0.80
15
0.60
10
0.40
A446 5
0.20

0.00 0
0 2 4 6 8 10 12 14

Incubation period (days)

Figure 5.22 A446 and A254 for cultures in the presence of 500 mg C/L NOM, and 500
mg C/L NOM plus Tween 80, agitated continuously at 30oC and 130 rpm.

Removal of NOM in terms of A446 and A254 was slightly higher for cultures with NOM alone
than those with NOM plus Tween 80 (Figure 5.22). This is consistent with the laccase results,
where the culture with NOM only has a higher laccase level than the culture with Tween 80,
further evidence for the role of laccase in the breakage of bonds that lead to reduction in A446
and A254. T. versicolor removed 86% and 82% (measured as A446) and 43% and 39%
(measured as A254) of the NOM from the NOM and NOM + Tw80 cultures, respectively.

89
5.3 Enzymatic Treatment of NOM
Filtered extracellular culture fluid obtained from the WB + Tw80 culture at 9 days was
incubated with NOM and NOM removal at varying pH and temperature was investigated. The
reaction mixture contained 4.0 mL of the culture fluid, 3.0 mL of Na2HPO4-citric acid buffer
and 3.0 mL of NOM (final concentration of 300 mg/L). The controls (NOM + buffer + water)
showed insignificant changes in A446 and A254 at different pHs and temperatures.

5.3.1 Effect of pH
The reaction mixtures were prepared with buffers in the range pH 3-7. The mixture was
incubated in a water bath set at 30oC. The Na2HPO4-citric acid buffer formulations are
tabulated in Table 5.3.

Table 5.3 Na2HPO4-citric acid buffer formulation (Dawson 1969).

o
pH at 18 C 0.2M Na2HPO4 (mL) 0.1M citric acid (mL)
2.5 0.855 9.145
3.0 2.055 7.945
4.0 3.855 6.145
5.0 5.150 4.850
6.0 6.315 3.685
7.0 8.235 1.765

There was reduction in A446 and A254 for all pHs except pH 7 for which there was an increase
in A254 (Figure 5.23). The increase in A254 at pH 7 was probably due to a change in the NOM
structure. The NOM removal ability for T. versicolor measured as A446 and A254 improved
with pH up to 4.5, above which it decreased markedly with increasing pH. The ratios of
A254/A446 for pH 3-5 were ~0.77, whereas they were ~0.58 for pH 6 and 7. The different ratios
of A254/A446 suggest that different bonds were broken by laccase at the different pH. The trend
for NOM removal matched the results for optimal pH for Lac activity, pH 4.5 (Figure 5.19);
hence the optimum pH for the reaction was determined as pH 4.5.

The results agree with those of many researchers (Zhang et al. 1999, Kapdan et al. 2000,
Aktas & Tanyolac 2003, Rancaño et al. 2003), in that the optimum pH for laccase was pH 4-
5, even though different substrates were utilised. Kapdan et al. (2000) stated that the dyestuff
decolourisation efficiencies of C. versicolor MUCL were lower at pH 6 and 7 compared with

90
pH 4.5, which is consistent with the current results. However, one research group found that
the optimum pH for the enzymatic decolourisation of Remazol Brilliant Blue R (RBBR) by
Funalia trogii ATCC 200800 growing in a solid-state fermentation (SSF) medium containing
wheat bran and soybean waste was pH 3.0 (Deveci et al. 2004), contrary to the present results.

25
A446
20 A254

15
% Reduction

10

0
3.0 4.0 4.5 5.0 6.0 7.0
-5

-10
pH

Figure 5.23 Reduction in A446 and A254 at different pH.

5.3.2 Effect of temperature

The reaction mixtures were prepared at pH 4.5 since it was the optimum value found in the
preceding experiments. The mixtures were incubated in water baths at different temperatures
in the range 30-80oC.

The optimum temperature for the reaction was established as 50oC (Figure 5.24), i.e., at which
maximum reduction in A446 and A254 occurred. The ratios of A254/A446 for the temperature
range 30-50oC were 0.77-0.79, whereas they were 0.58-0.69 for 60-80oC. As with pH,
different ratios of A254/A446 at the different temperatures were obtained, suggesting that
different bonds are being broken by laccase at the different temperatures. There was a
comparable reduction in A254 at 30-50oC, less reduction at 60oC and an increase at 70-80oC,
the latter probably due to thermal disruption of the NOM structure. No reduction in A446 was
observed for cultures at 70-80oC. These observations were different from when guaiacol was
used as a substrate to investigate the effects of temperature on Lac activity (Figure 5.18)
where high Lac activity occurred at 50-70oC.

91
 30
A446

25 A254

20
% Reduction

15

10

0
30 40 50 60 70 80
-5
Tem perature

Figure 5.24 Reduction in A446 and A254 at different temperatures.

Therefore, the effect of temperature varies when different substrates are used. Schliephake et
al. (2000) reported that the laccase from Pycnoporus cinnabarinus CBS 101046 when used
for the degradation of the diazo dye Chicago Sky Blue was stable at 60oC for one hour only,
but remained active at 37oC for long periods (25 days). Nyanhongo et al. (2002) showed that
different dyes were decolourised at different rates at different temperatures for which Lac
activity increased with temperature up to 50-60oC, after which it decreased. Although the
optimum temperature for laccase when using guaiacol as a substrate was 50oC, it was not so
clear-cut for NOM. Hence, 30oC was chosen as the incubation temperature for the in vitro
studies of Lac activity, as the laccase may be denatured if the NOM removal process were run
for long period at high temperature (30oC cf. 50oC), and it corresponds to the temperature at
which fungal growth (Figure 5.2) and enzyme production (Figure 5.4) were high (30oC cf.
36oC).

92
5.4 Biodegradation of NOM by T. versicolor
As discussed in Section 5.2.2, culture medium without added NOM supplemented with
Tween 80 or wheat bran, or both, produced low LiP and MnP activities but high Lac activity
compared with the cultures in the presence of NOM. This led to a further study to investigate
the potential of NOM as an inducer of the phenoloxidase enzymes in the presence of the two
supplements. Although a preliminary experiment (results shown in Figure 5.16) illustrated
that adsorption of NOM to wheat bran occurred, it was not to a great extent, and so
experiments were conducted to investigate the trade-off between adsorption and enzyme
induction, and hence NOM degradation.

Cultures containing 200 mL Waksman medium (5 g/L glucose) supplemented with 4.5 g/L
wheat bran and 0.5% (v/v) Tween 80, and sterilised NOM (final concentration of 100, 600
and 700 mg C/L) were prepared and each was inoculated with three fungal plugs. The cultures
were incubated at 30oC and 130 rpm for thirteen days. All fermentations were performed in
duplicate.

6.00 0.50 pH

Glucose
5.00
Glucose consumption (g/L)

0.40 consumption
A446
4.00
Absorbance

0.30 A254
pH OR

3.00
0.20
2.00

0.10
1.00

0.00 0.00
0 2 4 6 8 10 12

Incubation period (days)

Figure 5.25 History plot for T. versicolor cultures containing 100 mg C/L NOM,
supplemented with 4.5 g/L wheat bran and 0.5% (v/v) Tween 80, Waksman medium
with 5 g/L initial glucose, incubated at 30oC and 130 rpm.

Unlike the history plot for T. versicolor culture with no added supplements incubated at 30oC
(Figure 5.1), the pH drop occurred over the first five days, after which the pH increased
slightly (Figure 5.25). The pH fall coincided with increase in glucose consumption, but the
A446 and A254 did not vary greatly. The major reductions in the absorbances happened only

93
after day 5. It should be noted that the apparent glucose consumption was greater than that
provided (5 g/L) as the wheat bran also provided some reducing sugar as a carbon source for
the fungus.

The slow decreases in both A446 and A254 within the first five days, similar to the trends for
the cultures with no supplement, suggested that there was no or very little adsorption of NOM
on the fungal pellets or wheat bran despite the pH drop, which can lead to adsorption. There
was probably enzyme induction during the period and thus breakdown of NOM when there
were marked reductions in A446 and A254 from day 5-13. High removals of colour and UV-
absorbing compounds were achieved, 79% and 49% respectively. However, it should be noted
that the initial A446 and A254 for these systems were higher than the cultures with no
supplement due to the added coloured and UV-absorbing materials from the wheat bran.
Hence, the cultures with 100, 600 and 700 mg C/L NOM in the presence of wheat bran and
Tween 80 contained 26, 128 and 148 mg total carbon contents (converted from A446),
respectively. As there was 5.0-5.6 mg of the colour added to the culture fluids due to the
wheat bran, a corrected removal of NOM must be considered.

A446
80 5.00

A254
Glucose consumption (g/L)

4.00
NOM removal (mg)

60 Glucose
consumption
3.00

40
2.00

20
1.00

0 0.00
100 600 700

Initial NOM concentration (m g C/L)

Figure 5.26 NOM removals (as mg, converted from A446 and A254), and glucose
consumption in T. versicolor cultures with the two supplements and different NOM
concentrations, plug inoculum.

The overall apparent reductions in colour for the 100, 600 and 700 mg C/L NOM cultures
were 79%, 61% and 17%, respectively (Figure 5.26). The highest apparent removal of NOM
measured as A446 and A254 was obtained for the 600 mg C/L NOM cultures. The 700 mg C/L

94
NOM cultures had higher and lower apparent reductions in colour and UV-absorbing
molecules, respectively, than the 100 mg C/L NOM cultures. These trends matched the trends
for the NOM cultures without any supplements (Figure 5.13). The NOM removed per unit
glucose consumed followed the same trends, where the 600 mg C/L cultures had the highest.

When taking the colour from the wheat bran into account, the colour reductions for the
cultures with 100, 600 and 700 mg C/L NOM were 73%, 60% and 14%, respectively. No
improvement in NOM removal was obtained for all the cultures with the addition of
supplements, except that the culture containing 600 mg C/L NOM had 11 mg more reduction
in colour.

The adsorption of NOM on the wheat bran must be considered. To calculate the amount of
NOM adsorbed to the wheat bran, adsorption experiments were set up for three different
NOM concentrations (100, 600 and 700 mg C/L) in the absence of T. versicolor (Figure 5.27).

20
100

600
Reduction (% A446)

15
700

10

0
0 2 4 6 8 10 12

Control Incubation period (days)

Figure 5.27 A446 for 100, 600 and 700 mg C/L NOM incubated in Waksman medium (5
g/L glucose) containing 4.5 g/L wheat bran in the absence of fungus, as controls.

There was approximately 16% adsorption of NOM by the wheat bran for all the cultures with
different NOM concentrations. Therefore, the overall decolourisations after colour and
adsorption corrections due to the addition of wheat bran were 61%, 50% and 12% for the
cultures containing 100, 600 and 700 mg C/L NOM, respectively. After all, there was no
improvement in colour removal with the addition of the two supplements for all the cultures
compared with those in the absence of wheat bran and Tween 80.

95
The marked increases in LiP and MnP activities but decrease in Lac activity led to no
improvement in NOM removals for the systems (Figure 5.28). There was no increase in NOM
removal compared with the cultures with no supplements, probably due to the very large drop
in Lac activity. This is further evidence that laccase played a major role in the removal of
NOM.

100
LiP
MnP
80
Laccase
Enzyme activity (U/L)

60

40

20

0
100 600 700

Initial NOM concentration (m g C/L)

Figure 5.28 Activity of the extracellular ligninolytic enzymes of T. versicolor in

cultures containing the two supplements and different NOM concentrations, plug
inoculum.

As established in Section 5.2.2, addition of the wheat bran and Tween 80 combination
enhanced LiP, MnP and Lac production. However, the LiP and MnP activities were low in the
absence of NOM (Table 5.1). By adding NOM to the cultures supplemented with both wheat
bran and the detergent, Lac activity decreased markedly. In contrast, the addition of wheat
bran and Tween 80 to the NOM cultures led to increases in LiP and MnP activities, especially
LiP. The enhancement of LiP and MnP activities was attributed to the protective ability of
Tween 80 against mechanical inactivation of the enzymes (Venkatadri & Irvine 1990), and
subsequently led to a greater NOM removal. The results are consistent with the findings of
Venkatadri and Irvine (1990), who reported that the presence of Tween 80 caused a 1.3- to
1.4-fold increase in LiP enzyme activity. Furthermore, Tween 80 can aid the secretion of LiP
by promoting the permeation of LiP from the cell into the medium (Asther et al. 1987).
Therefore, the combination of NOM, wheat bran and Tween 80 can lead to increases in LiP
and MnP activities, however, they had an adverse effect on laccase although wheat bran and
Tween 80 can induce the production of laccase.

96
The overall apparent removal of NOM for the supplemented system with 100 mg C/L NOM
was 79%, however, the colour reduction was corrected to 73% due the colour from the wheat
bran. The NOM removal after corrected for colour from and adsorption by wheat bran was
markedly reduced to 61%. The addition of wheat bran and Tween 80 to cultures of T.
versicolor did not improve the decolourisation/removal of NOM for all the cultures when
adsorption by and colour from the wheat bran were taken into account.

97
Chapter 6 Conclusions and Recommendations

6.1 Conclusions
Phanerochaete chrysosporium ATCC 34541 had been shown to remove 40-50% NOM from
solution largely by adsorption rather than biodegradation (Rojek 2003). Consequently, the aim
of this research was to develop an improved biological treatment for the bioremediation of
NOM wastes.

When cultured in the presence of NOM, P. chrysosporium caused NOM removal and
decolourisation largely by adsorption. P. chrysosporium seemed to preferentially remove the
VHA fraction of the NOM, and so was most effective for the NOM preparation with the
highest VHA content (i.e., greatest colour, aromaticity and the highest SUVA). P.
chrysosporium cultures grown in media with different C:N ratios achieved the greatest
decolourisation with the lowest the C:N ratio: Waksman (C:N = 6) > Fahy (C:N = 76) > Fujita
medium (C:N = 114). This is in agreement with the findings of Rojek (2003), who also
demonstrated that the removal of NOM increased with decreased C:N ratio (1.58-15.81) for a
different NOM preparation.

Of the organisms tested, the two strains of P. chrysosporium (ATCC 34541 and ATCC
24725) and Saccharomyces spp. 1-3 removed NOM primarily by adsorption as indicated by
the deep brown colouration of the biomass, whereas T. versicolor had the greater removals
(59%) and the NOM removal was largely due to biodegradation.

The higher NOM removal with T. versicolor at 30oC corresponded to the greater production
of the oxidative enzymes than at 36oC (73% vs. 50%), suggesting reduced production of the
phenoloxidase enzyme activity at higher temperature. Increasing initial glucose content from
2 to 5 g/L did not improve NOM removal (48% cf. 73%), and biosorption occurred due to the
uptake of the NOM by the fungal cell walls and low enzyme activities were present.
Increasing NOM concentration led to increasing NOM removal and different activities of the
ligninolytic enzymes. The increase in NOM removal coincided with increasing Lac activity
but decreasing LiP and MnP activities, indicating laccase played a major role in NOM
degradation.

Addition of wheat bran, and wheat bran plus Tween 80, to the cultures caused increases in the
activities of LiP, MnP and Lac, especially laccase, as wheat bran can enhance the production
98
of the extracellular phenoloxidase enzymes (Couto et al. 2001, Lorenzo et al. 2002) whereas
Tween 80 can protect LiP against mechanical inactivation due to agitation as well as act as an
inducer (Asther et al. 1987, Venkatadri & Irvine 1990). Although the optimum activity of the
enzyme from the culture supplemented with 4.5 g/L wheat bran plus 0.5% Tween 80 occurred
at 50oC, 30oC was recommended as the ligninolytic enzyme activity may deteriorate if the
NOM removal process were run for long period at high temperature. The optimum pH for the
enzyme was identified as pH 4.0-4.5.

Although agitation led to apparent enzyme denaturation, fermentations with continuous

agitation promote faster and higher enzyme activity than those with occasional agitation due
to better mass transfer conditions. However, it seemed that continuous agitation had an
adverse effect on the fungal growth and enzyme production over longer fermentation periods
(greater than 9-13 days). Thus, discontinuous agitation would be a better option if extended
fermentation were required.

In the wheat bran and Tween 80 systems, there were two corrections needed to be taken into
account: colour from wheat bran, which contributed to the overall apparent NOM colour, and
adsorption of NOM on to the wheat bran. Addition of NOM to cultures of T. versicolor
supplemented with wheat bran and Tween 80 led to markedly reduced Lac activity, but
increased LiP and MnP activities, and no enhancement in NOM removal compared with the
cultures in the absence of supplements (12 mg (or 61%) cf. 15 mg (or 73%) for 100 mg C/L
after corrected for colour from and adsorption by wheat bran).

6.2 Recommendations
As the biological treatment of NOM wastes was conducted in shake flasks, further
investigations in a bioreactor would be recommended to determine the optimal operational
conditions such as dissolved oxygen levels and agitation. Kirk et al. (1978) suggested that the
lignin-degrading system required high O2 demand and so the impact of dissolved oxygen on
NOM removal may play a role in the biological treatment. Furthermore, Knapp et al. (1997)
demonstrated that agitation was very effective in improving (more than doubling) the rate of
Orange II decolourisation by mycelial pellets of an unidentified basidiomycete fungus due to
the improved mass transfer of oxygen and substrates in agitated cultures. Agitation may
therefore influence the performance of NOM removal and could be optimised via a trade-off
between enhanced mass transfer and ligninolytic enzyme denaturation. The operational

99
conditions could easily be optimised as the control of culture conditions can more easily be
managed in bioreactors.

NOM removal by T. versicolor immobilised in a membrane bioreactor or on other suitable

substrate may be an option to develop a reliable system applicable to drinking or wastewater
treatment. There are two types of configurations of membrane bioreactor systems, submerged
and side-stream (Mallia & Till 2001). Millia and Till (2001) reported that membrane
bioreactor technology is able to produce consistently high effluent quality and to reduce
disinfection as membranes with pore openings generally in the 0.1-0.5 mm range can trap a
significant proportion of pathogenic organisms. Furthermore, membrane bioreactor reduces
sludge production and requires a smaller design footprint. Shim and Kawamoto (2002)
established that immobilisation of mycelia cell culture on a bio-carrier is more effective in
promoting cell growth and LiP production compared to conventional stationary liquid culture.

The research was mainly focussed on the biological treatment of MIEX concentrate, further
investigations could be done on other NOM wastes such as sludges from membrane plants
and alum precipitation.

100
References
Aiken, GR, McKnight, DM, Thorn, KA & Thurman, EM 1992, 'Isolation of hydrophilic
organic acids from water using nonionic macroporous resins', Organic Geochemistry, 18(4),
pp. 567-573.

Aktas, N & Tanyolac, A 2003, 'Reaction conditions for laccase catalyzed polymerization of
catechol', Bioresource Technology, 87(3), pp. 209-214.

Aoustin, E, Schafer, AI, Fane, AG & Waite, TD 2001, 'Ultrafiltration of natural organic
matter', Separation and Purification Technology, 22-23(-), pp. 63-78.

Asther, M, Corrieu, G, Drapron, R & Odier, E 1987, 'Effect of Tween 80 and oleic acid on
ligninase production by Phanerochaete chrysosporium Ina-12', Enzyme and Microbial
Technology, 9(4), pp. 245-249.

Aust, SD 1995, 'Mechanisms of degradation by white rot fungi', Environmental Health

Perspectives, 103(Suppl 5), pp. 59-61.

Baldrian, P & Gabriel, J 2002, 'Variability of laccase activity in the white-rot basidiomycete
Pleurotus ostreatus', Folia Microbiologica, 47(4), pp. 385-390.

Barber, LB, Leenheer, JA, Noyes, TI & Stiles, EA 2001, 'Nature and transformation of
dissolved organic matter in treatment wetlands', Environmental Science & Technology,
35(24), pp. 4805-4816.

Barr, DP & Aust, SD 1994, 'Mechanisms white-rot fungi use to degrade pollutants',
Environmental Science & Technology, 28(2), pp. A78-A87.

Blondeau, R 1989, 'Biodegradation of natural and synthetic humic acids by the white-rot
fungus Phanerochaete chrysosporium', Applied and Environmental Microbiology, 55(5), pp.
1282-1285.

Bogan, BW & Lamar, RT 1996, 'Polycyclic aromatic hydrocarbon-degrading capabilities of

Phanerochaete laevis HHB-1625 and its extracellular ligninolytic enzymes', Applied and
Environmental Microbiology, 62(5), pp. 1597-1603.

Booth, C 1971, 'Fungal culture media', In Methods in Microbiology, 4th edn, Academic Press,
New York.

101
Bourbonnais, R & Paice, MG 1990, 'Oxidation of nonphenolic substrates - An expanded role
for laccase in lignin biodegradation', FEBS Letters, 267(1), pp. 99-102.

Bumpus, JA & Aust, SD 1987, 'Biodegradation of DDT 1,1,1-trichloro-2,2-bis(4-

chlorophenyl)ethane by the white-rot fungus Phanerochaete chrysosporium', Applied and
Environmental Microbiology, 53(9), pp. 2001-2008.

Burton, SG, Boshoff, A, Edwards, W, Jacobs, EP, Leukes, WD, Rose, PD, Russel, AK,
Russel, IM & Ryan, D 1999, 'Developing biotechnological systems for the treatment of
organic pollutants: final report to the Water Research Commission', Water Research
Commission, Pretoria, WRC Report 687/1/98.

Carraro, E, Bugliosi, EH, Meucci, L, Baiocchi, C & Gilli, G 2000, 'Biological drinking water
treatment processes, with special reference to mutagenicity', Water Research, 34(11), pp.
3042-3054.

Carroll, T, King, S, Gray, SR, Bolto, BA & Booker, NA 2000, 'The fouling of microfiltration
membranes by NOM after coagulation treatment', Water Research, 34(11), pp. 2861-2868.

Castillo, MD, Ander, P & Stenstrom, J 1997, 'Lignin and manganese peroxidase activity in
extracts from straw solid substrate fermentations', Biotechnology Techniques, 11(9), pp. 701-
706.

Chin, YP, Aiken, G & Oloughlin, E 1994, 'Molecular-weight, polydispersity, and

spectroscopic properties of aquatic humic substances', Environmental Science & Technology,
28(11), pp. 1853-1858.

Choi, SH, Moon, SH & Gu, MB 2002, 'Biodegradation of chlorophenols using the cell-free
culture broth of Phanerochaete chrysosporium immobilized in polyurethane foam', Journal of
Chemical Technology and Biotechnology, 77(9), pp. 999-1004.

Chow, CWK, Fabris, R & Drikas, M 2004, 'A rapid fractionation technique to characterise
natural organic matter for the optimisation of water treatment processes', Journal of Water
Supply Research and Technology-Aqua, 53(2), pp. 85-92.

Chow, CWK, van Leeuwen, JA, Drikas, M, Fabris, R, Spark, KM & Page, DW 1999, 'The
impact of the character of natural organic matter in conventional treatment with alum', Water
Science and Technology, 40(9), pp. 97-104.

102
Coll, PM, Fernandezabalos, JM, Villanueva, JR, Santamaria, R & Perez, P 1993, 'Purification
and characterization of a phenoloxidase (laccase) from the lignin-degrading Basidiomycete
Pm1 (Cect-2971)', Applied and Environmental Microbiology, 59(8), pp. 2607-2613.

Couto, SR, Dominguez, A & Sanroman, A 2001, 'Utilisation of lignocellulosic wastes for
lignin peroxidase production by semi-solid-state cultures of Phanerochaete chrysosporium',
Biodegradation, 12(5), pp. 283-289.

Couto, SR, Rosales, E, Gundin, M & Sanroman, MA 2004, 'Exploitation of a waste from the
brewing industry for laccase production by two Trametes species', Journal of Food
Engineering, 64(4), pp. 423-428.

Couto, SR & Sanromán, MA 2005, 'Application of solid-state fermentation to ligninolytic

enzyme production', Biochemical Engineering Journal, 22(3), pp. 211-219.

Crawford, DL & Crawford, RL 1980, 'Microbial degradation of lignin', Enzyme and

Microbial Technology, 2(1), pp. 11-22.

Croué, JP, Korshin, G & Benjamin, M 2000, 'Characterisation of natural organic matter in
drinking water', AWWA Research Foundation: Denver, Co., USA, AWWARF Report No.
90780.

Dahiya, J, Singh, D & Nigam, P 2001, 'Decolourisation of synthetic and spentwash

melanoidins using the white-rot fungus Phanerochaete chrysosporium JAG-40', Bioresource
Technology, 78(1), pp. 95-98.

Dawson, RMC 1969, Data for Biochemical Research, 2nd edn, Oxford University Press, New
York, pp. 192-209.

De Souza, CGM, Zilly, A & Peralta, RM 2002, 'Production of laccase as the sole
phenoloxidase by a Brazilian strain of Pleurotus pulmonarius in solid state fermentation',
Journal of Basic Microbiology, 42(2), pp. 83-90.

Dehorter, B & Blondeau, R 1992, 'Extracellular enzyme activities during humic acid
degradation by the white-rot fungi Phanerochaete chrysosporium and Trametes versicolor',
FEMS Microbiology Letters, 94(3), pp. 209-215.

Del Pilar Castillo, M 1997, 'Degradation of pesticides by Phanerochaete chrysosporium in

solid substrate fermentation', C634394, Sveriges Lantbruksuniversitet (Sweden).

103
Deveci, T, Unyayar, A & Mazmanci, MA 2004, 'Production of Remazol Brilliant Blue R
decolourising oxygenase from the culture filtrate of Funalia trogii ATCC 200800', Journal of
Molecular Catalysis B-Enzymatic, 30(1), pp. 25-32.

Dodor, DE, Hwang, HM & Ekunwe, SIN 2004, 'Oxidation of anthracene and benzo[α]pyrene
by immobilized laccase from Trametes versicolor', Enzyme and Microbial Technology, 35(2-
3), pp. 210-217.

Drikas, M, Chow, CWK & Cook, D 2003, 'The impact of recalcitrant organic character on
disinfection stability, trihalomethane formation and bacterial regrowth: An evaluation of
magnetic ion exchange resin (MIEX (R)) and alum coagulation', Journal of Water Supply
Research and Technology-Aqua, 52(7), pp. 475-487.

Fahy, V, FitzGibbon, FJ, McMullan, G, Singh, D & Marchant, R 1997, 'Decolourisation of

molasses spent wash by Phanerochaete chrysosporium', Biotechnology Letters, 19(1), pp. 97-
99.

Faison, BD & Kirk, TK 1985, 'Factors involved in the regulation of a ligninase activity in
Phanerochaete chrysosporium', Applied and Environmental Microbiology, 49(2), pp. 299-
304.

Feng, W, Ozaki, H, Terashima, Y, Imada, T & Ohkouchi, Y 1996, 'Activities of ligninolytic

enzymes of the white rot fungus, Phanerochaete chrysosporium and its recalcitrant substance
degradability', Water Science and Technology, 34(7-8), pp. 69-78.

Fenice, M, Sermanni, GG, Federici, F & D'Annibale, A 2003, 'Submerged and solid state
production of laccase and Mn peroxidase by Panus tigrinus on olive mill wastewater-based
media', Journal of Biotechnology, 100(1), pp. 77-85.

Ferraz, A, Cordova, AM & Machuca, A 2003, 'Wood biodegradation and enzyme production
by Ceriporiopsis subvermispora during solid state fermentation of Eucalyptus grandis',
Enzyme and Microbial Technology, 32(1), pp. 59-65.

Frazier, SW, Nowack, KO, Goins, KM, Cannon, FS, Kaplan, LA & Hatcher, PG 2002,
'Characterization of organic matter from natural waters using tetramethylammonium
hydroxide thermochemolysis GC-MS', Journal of Analytical and Applied Pyrolysis,
00(Article in Press), pp. 1-30.

104
Frimmel, FH 1998, 'Characterization of natural organic matter as major constituents in aquatic
systems', Journal of Contaminant Hydrology, 35(1-3), pp. 201-216.

Fujian, X, Hongzhang, C & Zuohu, L 2001, 'Solid state production of lignin peroxidase (LiP)
and manganese peroxidase (MnP) by Phanerochaete chrysosporium using steam-exploded
straw as substrate', Bioresource Technology, 80(2), pp. 149-151.

Fujita, M, Ike, M, Kusunoki, K, Ueno, T, Serizawa, K & Hirao, T 2002, 'Removal of color
and estrogenic substances by fungal reactor equipped with ultrafiltration unit', In 3rd World
Water Congress of the International Water Association, Melbourne, Australia, 7-12 Apr,
2002, IWA Publishing, London, UK, pp. 353-358.

Galliano, H, Gas, G, Seris, JL & Boudet, AM 1991, 'Lignin degradation by Rigidoporus

lignosus involves synergistic action of 2 oxidizing enzymes - Mn peroxidase and laccase',
Enzyme and Microbial Technology, 13(6), pp. 478-482.

Gilli, G, Scursatone, E, Palin, L, Bono, R, Carraro, E & Meucci, L 1990, 'Water disinfection -
A relationship between ozone and aldehyde production', Ozone-Science & Engineering,
12(3), pp. 231-241.

Glenn, JK, Morgan, MA, Mayfield, MB, Kuwahara, M & Gold, MH 1983, 'An extracellular
H2O2-requiring enzyme preparation involved in lignin biodegradation by the white-rot
Basidiomycete Phanerochaete chrysosporium', Biochemical and Biophysical Research
Communications, 114(3), pp. 1077-1083.

Gold, MH & Alic, M 1993, 'Molecular biology of the lignin-degrading Basidiomycete

Phanerochaete chrysosporium', Microbiological Reviews, 57(3), pp. 605-622.

Gold, MH, Wariishi, H & Valli, K 1989, 'Extracellular peroxidases involved in lignin
degradation by the white-rot Basidiomycete Phanerochaete chrysosporium', Acs Symposium
Series, 389(-), pp. 127-140.

Golovleva, LA, Maltseva, OV, Leontievskii, AA, Miasoedova, NM & Skriabin, GK 1987,
'Ligninase biosynthesis by the fungus Panus Tigrinus during solid state fermentation of
straw', Doklady Akademii Nauk Sssr, 294(4), pp. 992-995.

Griffin, DH 1994, Fungal Physiology, Wiley-Liss edn, New York.

105
Hammel, KE, Jensen, KA, Mozuch, MD, Landucci, LL, Tien, M & Pease, EA 1993,
'Ligninolysis by a purified lignin peroxidase', Journal of Biological Chemistry, 268(17), pp.
12274-12281.

Harvey, PJ, Schoemaker, HE & Palmer, JM 1986, 'Veratryl Alcohol as a mediator and the
role of radical cations in lignin biodegradation by Phanerochaete chrysosporium', FEBS
Letters, 195(1-2), pp. 242-246.

Hood, E, McKnight, DM & Williams, MW 2003, 'Sources and chemical character of

dissolved organic carbon across an alpine/subalpine ecotone, Green Lakes Valley, Colorado
Front Range, United States', Water Resources Research, 39(7), pp. 1-12.

Howe, KJ & Clark, MM 2002, 'Fouling of microfiltration and ultrafiltration membranes by

natural waters', Environmental Science & Technology, 36(16), pp. 3571-3576.

Jacangelo, JG, DeMarco, J, Owen, DM & Randtke, SJ 1995, 'Selected processes for removing
NOM: an overview', Journal AWWA, pp. pp. 64-77.

Jäger, A, Croan, S & Kirk, TK 1985, 'Production of ligninases and degradation of lignin in
agitated submerged cultures of Phanerochaete chrysosporium', Applied and Environmental
Microbiology, 50(5), pp. 1274-1278.

Kapdan, IK, Kargia, F, McMullan, G & Marchant, R 2000, 'Effect of environmental

conditions on biological decolorization of textile dyestuff by C. versicolor', Enzyme and
Microbial Technology, 26(5-6), pp. 381-387.

Kapich, AN, Prior, BA, Botha, A, Galkin, S, Lundell, T & Hatakka, A 2004, 'Effect of
lignocellulose-containing substrates on production of ligninolytic peroxidases in submerged
cultures of Phanerochaete chrysosporium ME-446', Enzyme and Microbial Technology,
34(2), pp. 187-195.

Kawai, S, Umezawa, T & Higuchi, T 1988, 'Degradation mechanisms of phenolic β-1 lignin
substructure model compounds by laccase of Coriolus versicolor', Archives of Biochemistry
and Biophysics, 262(1), pp. 99-110.

Kirisits, MJ, Snoeyink, VL, Inan, H, Chee-Sanford, JC, Raskin, L & Brown, JC 2001, 'Water
quality factors affecting bromate reduction in biologically active carbon filters', Water
Research, 35(4), pp. 891-900.

106
Kirk, TK & Chang, HM 1975, 'Decomposition of lignin by white-rot fungi. 2.
Characterization of heavily degraded lignins from decayed spruce', Holzforschung, 29(2), pp.
56-64.

Kirk, TK & Farrell, RL 1987, 'Enzymatic combustion - the microbial degradation of lignin',
Annual Review of Microbiology, 41(-), pp. 465-505.

Kirk, TK, Schultz, E, Connors, WJ, Lorenz, LF & Zeikus, JG 1978, 'Influence of culture
parameters on lignin metabolism by Phanerochaete chrysosporium', Archives of
Microbiology, 117(3), pp. 277-285.

Knapp, JS, Zhang, FM & Tapley, KN 1997, 'Decolourisation of Orange II by a wood-rotting

fungus', Journal of Chemical Technology and Biotechnology, 69(3), pp. 289-296.

Kördel, W, Dassenakis, M, Lintelmann, J & Padberg, S 1997, 'The importance of natural

organic material for environmental processes in waters and soils', Pure and Applied
Chemistry, 69(7), pp. 1571-1600.

Kuwahara, M, Glenn, JK, Morgan, MA & Gold, MH 1984, 'Separation and characterization
of 2 extracellular H2O2-dependent oxidases from ligninolytic cultures of Phanerochaete
chrysosporium', FEBS Letters, 169(2), pp. 247-250.

Lang, E, Nerud, F, Novotna, E, Zadrazil, F & Martens, R 1996, 'Production of ligninolytic

exoenzymes and C-14-pyrene mineralization by Pleurotus sp. in lignocellulose substrate',
Folia Microbiologica, 41(6), pp. 489-493.

Leatham, GF 1986, 'The ligninolytic activities of Lentinus edodes and Phanerochaete

chrysosporium', Applied Microbiology and Biotechnology, 24(1), pp. 51-58.

Leenheer, JA 1981, 'Comprehensive approach to preparative isolation and fractionation of

dissolved organic carbon from natural waters and wastewaters', Environmental Science &
Technology, 15(5), pp. 578-587.

Leenheer, JA & Croué, JP 2003, 'Characterizing aquatic dissolved organic matter',

Environmental Science & Technology, 37(1), pp. 18A-26A.

Leisola, MSA & Garcia, S 1989, 'The mechanisms of lignin degradation', In Enzyme systems
for lignocellulose degradation, Elsevier Science Publishers, Essex, UK, 89-99.

107
Liao, W, Christman, RF, Johnson, JD, Millington, DS & Hass, JR 1982, 'Structural
characterization of aquatic humic material', Environmental Science & Technology, 16(7), pp.
403-410.

Lonergan, GT 1992, 'White-rot fungi - An environmental panacea?' Australasian

Biotechnology, 2(4), pp. 214-217.

Lorenzo, M, Moldes, D, Rodriguez Couto, S & Sanroman, A 2002, 'Improving laccase

production by employing different lignocellulosic wastes in submerged cultures of Trametes
versicolor', Bioresource Technology, 82(2), pp. 109-113.

Mallia, H & Till, S 2001, 'Membrane bioreactor: wastewater treatment applications to achieve
high quality effluent', In 64th Annual Victorian Water Industry Engineers and Operators'
Conference, All Seasons International Hotel-Bendigo, Australia, September 5-6, Water
Industry Operators Association (WIOA).

Marhaba, TF, Van, D & Lippincott, RL 2000, 'Changes in NOM fractionation through
treatment: A comparison of ozonation and chlorination', Ozone-Science & Engineering, 22(3),
pp. 249-266.

Martinez, MJ, RuizDuenas, FJ, Guillen, F & Martinez, AT 1996, 'Purification and catalytic
properties of two manganese peroxidase isoenzymes from Pleurotus eryngii', European
Journal of Biochemistry, 237(2), pp. 424-432.

McKnight, DM, Andrews, ED, Spaulding, SA & Aiken, GR 1994, 'Aquatic fulvic acids in
algal-rich Antarctic ponds', Limnology and Oceanography, 39(8), pp. 1972-1979.

Mehna, A, Bajpai, P & Bajpai, PK 1995, 'Studies on decolorization of effluent from a small
pulp mill utilizing agriresidues with Trametes versicolor', Enzyme and Microbial Technology,
17(1), pp. 18-22.

Mester, T, Sierra-Alvarez, R & Field, JA 1998, 'Peroxidase and aryl metabolite production by
the white rot fungus Bjerkandera sp. strain BOS55 during solid state fermentation of
lignocellulosic substrates', Holzforschung, 52(4), pp. 351-358.

Miller, GL 1959, 'Use of dinitrosalicylic acid reagent for determination of reducing sugar',
Analytical Chemistry, 31(3), pp. 426-428.

108
Motheo, AJ & Pinhedo, L 2000, 'Electrochemical degradation of humic acid', The Science of
The Total Environment, 256(1), pp. 67-76.

Muncnerova, D & Augustin, J 1994, 'Fungal metabolism and detoxification of polycyclic

aromatic hydrocarbons: A review', Bioresource Technology, 48(2), pp. 97-106.

Nigam, P, Pandey, A & Prabhu, KA 1987, 'Cellulase and ligninase production by

Basidiomycete culture in solid state fermentation', Biological Wastes, 20(1), pp. 1-9.

Nishijima, W, Fahmi, Mukaidani, T & Okada, M 2003, 'DOC removal by multi-stage

ozonation-biological treatment', Water Research, 37(1), pp. 150-154.

Niwa, M 2004, 'Control of hazardous bacteria in acidic beverages by using a guaiacol

detection kit (peroxidase method)', Japan Food Science, 2004(2), pp. 23-28.

Nyanhongo, GS, Gomes, J, Gubitz, GM, Zvauya, R, Read, J & Steiner, W 2002,
'Decolorization of textile dyes by laccases from a newly isolated strain of Trametes modesta',
Water Research, 36(6), pp. 1449-1456.

Ooijkaas, LP, Weber, FJ, Buitelaar, RM, Tramper, J & Rinzema, A 2000, 'Defined media and
inert supports: their potential as solid-state fermentation production systems', Trends in
Biotechnology, 18(8), pp. 356-360.

Pal, M, Calvo, AM, Terron, MC & Gonzalez, AE 1995, 'Solid state fermentation of sugarcane
bagasse with Flammulina velutipes and Trametes versicolor', World Journal of Microbiology
& Biotechnology, 11(5), pp. 541-545.

Pandey, A, Soccol, CR & Mitchell, D 2000, 'New developments in solid state fermentation: I-
bioprocesses and products', Process Biochemistry, 35(10), pp. 1153-1169.

Papinutti, VL, Diorio, LA & Forchiassin, F 2003, 'Production of laccase and manganese
peroxidase by Fomes sclerodermeus grown on wheat bran', Journal of Industrial
Microbiology & Biotechnology, 30(3), pp. 157-160.

Pradeep, V & Datta, M 2002, 'Production of ligninolytic enzymes for decolourisation by

cocultivation of white-rot fungi Pleurotus ostreatus and Phanerochaete chrysosporium under
solid state fermentation', Applied Biochemistry and Biotechnology, 102-103(-), pp. 109-118.

109
Prévost, M, Rompre, A, Coallier, J, Servais, P, Laurent, P, Clement, B & Lafrance, P 1998,
'Suspended bacterial, biomass and activity in full-scale drinking water distribution systems:
Impact of water treatment', Water Research, 32(5), pp. 1393-1406.

Ralph, JP & Catcheside, DEA 1994, 'Decolorization and depolymerization of solubilized low-
rank coal by the white-rot Basidiomycete Phanerochaete chrysosporium', Applied
Microbiology and Biotechnology, 42(4), pp. 536-542.

Rancaño, G, Lorenzo, M, Molares, N, Rodriguez Couto, S & Sanroman, MA 2003,

'Production of laccase by Trametes versicolor in an airlift fermentor', Process Biochemistry,
39(4), pp. 467-473.

Reid, ID 1979, 'Influence of nutrient balance on lignin degradation by the white rot fungus
Phanerochaete chrysosporium', Canadian Journal of Botany-Revue Canadienne De
Botanique, 57(19), pp. 2050-2058.

Robinson, T, McMullan, G, Marchant, R & Nigam, P 2001, 'Remediation of dyes in textile

effluent: a critical review on current treatment technologies with a proposed alternative',
Bioresource Technology, 77(3), pp. 247-255.

Rodriguez, E, Pickard, MA & Vazquez-Duhalt, R 1999, 'Industrial dye decolorization by

laccases from ligninolytic fungi', Current Microbiology, 38(1), pp. 27-32.

Rojek, K 2003, 'Decolourisation of aquatic NOM with the white-rot fungus Phanerochaete
chrysosporium', Master of Engineering thesis, School of Civil and Chemical Engineering,
RMIT University.

Rojek, K, Roddick, FA & Parkinson, A 2004, 'Decolorisation of natural organic matter by

Phanerochaete chrysosporium: the effect of environmental conditions', Water Science and
Technology: Water Supply, 4(4), pp. 175-182.

Schäfer, AI, Mauch, R, Waite, TD & Fane, AG 2002, 'Charge effects in the fractionation of
natural organics using ultrafiltration', Environmental Science & Technology, 36(12), pp. 2572-
2580.

Schliephake, K, Mainwaring, DE, Lonergan, GT, Jones, IK & Baker, WL 2000,

'Transformation and degradation of the disazo dye Chicago Sky Blue by a purified laccase
from Pycnoporus cinnabarinus', Enzyme and Microbial Technology, 27(1-2), pp. 100-107.

110
Schlosser, D & Höfer, C 2002, 'Laccase-catalyzed oxidation of Mn2+ in the presence of
natural Mn3+ chelators as a novel source of extracellular H2O2 production and its impact on
manganese peroxidase', Applied and Environmental Microbiology, 68(7), pp. 3514-3521.

Shevchenko, SM & Bailey, GW 1996, 'Life after death: Lignin-humic relationships

reexamined', Critical Reviews in Environmental Science and Technology, 26(2), pp. 95-153.

Shim, S-S & Kawamoto, K 2002, 'Enzyme production activity of Phanerochaete

chrysosporium and degradation of pentachlorophenol in a bioreactor', Water Research,
36(18), pp. 4445-4454.

Shinners-Carnelley, TC, Szpacenko, A, Tewari, JP & Palcic, MM 2002, 'Enzymatic activity

of Cyathus olla during solid state fermentation of canola roots', Phytoprotection, 83(1), pp.
31-40.

Sik, S & Unyayar, A 1998, 'Phanerochaete chrysosporium and Funalia trogii for the
degradation of cotton stalk and their laccase, peroxidase, ligninase and cellulase enzyme
activities under semi solid state conditions', Turkish Journal of Biology, 22(3), pp. 287-298.

Slunjski, M, Cadde, K, O'Leary, B & Tattersall, J 2000, 'MIEX® resin water treatment
process', In Aquatech Amsterdam, Amsterdam, the Netherlands, 26-29 September, 2000.

Swamy, J & Ramsay, JA 1999, 'Effects of glucose and NH4+ concentrations on sequential dye
decoloration by Trametes versicolor', Enzyme and Microbial Technology, 25(3-5), pp. 278-
284.

Swietlik, J, Dabrowska, A, Raczyk-Stanislawiak, U & Nawrocki, J 2004, 'Reactivity of

natural organic matter fractions with chlorine dioxide and ozone', Water Research, 38(3), pp.
547-558.

ten Have, R & Teunissen, PJM 2001, 'Oxidative mechanisms involved in lignin degradation
by white-rot fungi', Chemical Reviews, 101(11), pp. 3397-3413.

Thurman, EM & Malcolm, RL 1981, 'Preparative isolation of aquatic humic substances',

Environmental Science & Technology, 15(4), pp. 463-466.

Tien, M 1987, 'Properties of ligninase from Phanerochaete chrysosporium and their possible
applications', Crc Critical Reviews in Microbiology, 15(2), pp. 141-168.

111
Tien, M & Kirk, TK 1983, 'Lignin-degrading enzyme from the Hymenomycete
Phanerochaete chrysosporium burds', Science, 221(4611), pp. 661-662.

Tien, M & Kirk, TK 1984, 'Lignin-degrading enzyme from Phanerochaete chrysosporium -

Purification, characterization, and catalytic properties of a unique H2O2-requiring oxygenase',
Proceedings of the National Academy of Sciences of the United States of America-Biological
Sciences, 81(8), pp. 2280-2284.

Tien, M & Kirk, TK 1988, 'Lignin peroxidase of Phanerochaete chrysosporium', Methods in

Enzymology, 161(-), pp. 238-249.

Tsai, TS 1991, 'Biotreatment of red water - A hazardous waste stream from explosive
manufacture - with fungal systems', Hazardous Waste & Hazardous Materials, 8(3), pp. 231-
244.

Vares, T, Kalsi, M & Hatakka, A 1995, 'Lignin peroxidases, manganese peroxidases, and
other ligninolytic enzymes produced by Phlebia radiata during solid state fermentation of
wheat straw', Applied and Environmental Microbiology, 61(10), pp. 3515-3520.

Venkatadri, R & Irvine, RL 1990, 'Effect of agitation on ligninase activity and ligninase
production by Phanerochaete chrysosporium', Applied and Environmental Microbiology,
56(9), pp. 2684-2691.

Vickers, JC, Thompson, MA & Kelkar, UG 1995, 'The use of membrane filtration in
conjunction with coagulation processes for improved NOM removal', Desalination, 102(1-3),
pp. 57-61.

Volk, C, Renner, C, Roche, P, Paillard, H & Joret, JC 1993, 'Effects of ozone on the
production of biodegradable dissolved organic carbon (BDOC) during water treatment',
Ozone-Science & Engineering, 15(5), pp. 389-404.

Wariishi, H, Akileswaran, L & Gold, MH 1988, 'Manganese peroxidase from the

basidiomycete Phanerochaete chrysosporium - Spectral characterization of the oxidized states
and the catalytic cycle', Biochemistry, 27(14), pp. 5365-5370.

Wariishi, H & Gold, MH 1989, 'Lignin peroxidase compound III - Formation, inactivation,
and conversion to the native enzyme', FEBS Letters, 243(2), pp. 165-168.

112
Wariishi, H, Valli, K & Gold, MH 1992, 'Manganese(II) oxidation by manganese peroxidase
from the Basidiomycete Phanerochaete chrysosporium - Kinetic mechanism and role of
chelators', Journal of Biological Chemistry, 267(33), pp. 23688-23695.

Weber, J 2001, Definition of soil organic matter, Humintech, viewed 7 June 2005
<http://www.humintech.com/001/articles/article_definition_of_soil_organic_matter.html>.

Wedzicha, BL & Kaputo, MT 1992, 'Melanoidins from glucose and glycine - composition,
characteristics and reactivity towards sulfite ion', Food Chemistry, 43(5), pp. 359-367.

White, DM, Garland, DS, Narr, J & Woolard, CR 2003, 'Natural organic matter and DBP
formation potential in Alaskan water supplies', Water Research, 37(4), pp. 939-947.

Wricke, B, Petzoldt, H, Heiser, H & Bornmann, K 1996, 'NOM removal by biofiltration after
ozonation - results of a pilot plant test', In Advances in slow sand as alternative biological
filtration, John Wiley & Sons, New York.

Xu, F 1996, 'Oxidation of phenols, anilines, and benzenethiols by fungal laccases: Correlation
between activity and redox potentials as well as halide inhibition', Biochemistry, 35(23), pp.
7608-7614.

Yanagi, Y, Hamaguchi, S, Tamaki, H, Suzuki, T, Otsuka, H & Fujitake, N 2003, 'Relation of

chemical properties of soil humic acids to decolorization by white rot fungus-Coriolus
Consors', Soil Science and Plant Nutrition, 49(2), pp. 201-206.

Zavarzina, AG, Leontievsky, AA, Golovleva, LA & Trofimov, SY 2004, 'Biotransformation

of soil humic acids by blue laccase of Panus tigrinus 8/18: an in vitro study', Soil Biology &
Biochemistry, 36(2), pp. 359-369.

Zhang, F-m, Knapp, JS & Tapley, KN 1999, 'Decolourisation of cotton bleaching effluent
with wood rotting fungus', Water Research, 33(4), pp. 919-928.

Zhou, JL & Banks, CJ 1991, 'The adsorption of humic acid fractions by fungal biomass',
Environmental Technology, 12(6), pp. 519-530.

Ziegenhagen, D & Hofrichter, M 1998, 'Degradation of humic acids by manganese peroxidase

from the white-rot fungus Clitocybula dusenii', Journal of Basic Microbiology, 38(4), pp.
289-299.

113
APPENDICES

Appendix 1 Correlation between NOM concentration and A446 and A254

0.15 5.00 A 446

NOM 1 A 254
4.00
Absorbance at 446 nm

Absorbance at 254 nm
0.10
A 446 = 0.0012x + 0.0009 3.00
R2 = 0.9995

2.00
0.05

A 254 = 0.0363x - 0.0252 1.00

R2 = 0.9992

0.00 0.00

0.20 5.00

NOM 2
4.00
0.15
Absorbance at 446 nm

Absorbance at 254 nm

A 446 = 0.0016x + 0.0021

R2 = 0.9979 3.00

0.10

2.00

0.05
1.00
A 254 = 0.0384x - 0.0833
R2 = 0.9983

0.00 0.00

0.25 5.00
NOM 3
0.20 4.00
Absorbance at 446 nm

A 446 = 0.0021x - 0.0022

Absorbance at 254 nm

R2 = 0.9997
0.15 3.00

0.10 2.00

0.05 1.00
A 254 = 0.0381x - 0.0771
R2 = 0.9999
0.00 0.00
0 20 40 60 80 100

NOM conce ntr ation (m g C/L)

Figure A1 The correlation between NOM concentrations and A446 and A254 for the
three NOM preparations.

114
Appendix 2 Absorbance correction factor

25
NOM 1
20
Correction Factor (%)

15
CF = 5.5983x + 1.0684
R2 = 0.9862
10

20

NOM 2
Correction Factor (%)

15

10
CF = 3.423x + 0.1878
R2 = 0.9823
5

25
NOM 3
20
Correction Factor (%)

15
CF = 5.2819x + 0.9718
R2 = 0.9871
10

0
0 1 2 3 4 5
pH Difference (pH0 -pHi )

Figure A2 pH impact on the absorbance at 446 nm and 254 nm.

115
The experiments were conducted twice and the samples for each were analysed three times.
The mean value values were plotted.

The description of the calculation of corrected absorbance based on the plots on previous page
(Figure A2) is explained in Table A2.

Table A2 Calculation of corrected absorbance.

Y-axis X-axis Corrected absorbance,
Aci
Aci − Ai pH difference Ai × CF
CF (%) = × 100 Aci = Ai +
Ai = pH 0 − pH i 100

CF = Correction factor (%) pH0 = Initial pH Aci = Corrected absorbance

Ai = Measured absorbance pHi = pH on day i on day i

on day i
Aci = Corrected absorbance
on day i

116
Appendix 3 Typical standard curve for glucose determination

0.80
Absorbance at 570 nm

0.60
y = 0.346x - 0.005
R2 = 1.000
0.40

0.20

0.00
0.0 0.5 1.0 1.5 2.0
Glucose concentration (g/L)

Figure A3 Typical standard curve for glucose determination by DNS method.

117
Appendix 4 Standard curve for determination of apparent molecular weight (Dalton)
in HPSEC analysis

8.0

6.0
log(M w)

4.0
log (Mw) = -0.399(t) + 7.205
R2 = 1.000
2.0

0.0
0 2 4 6 8 10 12
Retention tim e (m inutes)

Figure A4 Standard curve for HPSEC using polystyrene sulphonate standards.

118