microbial degradation of polyester...
TRANSCRIPT
Microbial degradation of polyester polyurethane
By
Zia Ullah Shah
DEPARTMENT OF BIOTECHNOLOGY
FACULTY OF BIOLOGICAL SCIENCES
QUAID-I-AZAM UNIVERSITY
ISLAMABAD
2012
A thesis submitted in partial fulfillment of the requirements for
the
Degree of
DOCTOR OF PHILOSOPHY (Ph.D.)
IN
BIOTECHNOLOGY
By
Zia Ullah Shah
DEPARTMENT OF BIOTECHNOLOGY
FACULTY OF BIOLOGICAL SCIENCES
QUAID-I-AZAM UNIVERSITY
ISLAMABAD
2012
DEDICATED TO
MY Parents and my Aunty
DECLARATION
The material and information contained in this thesis is my original work.
I have not previously presented any part of this work elsewhere for any
other degree.
Zia Ullah Shah
CERTIFICATE
This thesis, submitted by Zia Ullah Shah, is accepted in its present form
by Department of Biotechnonology, Faculty of Biological Sciences,
Quaid-i-Azam University, Islamabad as satisfying the thesis requirements
for the degree of
Doctor of Philosophy (Ph.D.)
EXTERNAL EXAMINER:
_________________________
EXTERNAL EXAMINER:
_________________________
INTERNAL EXAMINER:
_________________________
CHAIRMAN:
_________________________
Dated:………………
TABLE OF CONTENTS
_____________________________________________________________________
Acknowledgement i
List of Tables iii
List of Figures iv
List of Abbreviations ix
Abstract xi
Introduction 1
Aims and Objectives 8
Literature Review
2.1. Importance of Plastics in Our Daily Life: Polyurethane Share 9
2.2. Introduction of Polyurethane 9
2.3. Environmental Concerns about Plastics Waste: Polyurethane 13
2.4. Plastics Waste Management: Polyurethane 14
2.4.1. Biodegradation of Polyurethane 15
2.4.1.1. Fungal Degradation 18
2.4.1.2. Bacterial Degradation 19
2.4.1.3. Polyurethane Degrading Enzymes 21
2.5. Polymer Degradation Analysis 23
2.5.1. Laboratory Tests 23
2.5.1.1. Visual Observations 25
2.5.1.2. Changes in Mechanical Properties and Molar Mass 25
2.5.1.3. Weight Loss: Determination of Residual Polymer 26
2.5.1.4. CO2 evolution/O2 Consumption 26
2.5.1.5. Determination of Biogas 28
2.5.1.6. Radio Labeling 28
2.5.1.7. Clear-zone Formation 29
2.5.3. Simulation Tests 29
2.5.3.1. Field Tests 29
2.6. Degradation products of Polyurethane 30
Materials and Methods
3.1. Isolation of Pu Degrading Microorganisms 33
3.1.1. Materials 33
3.1.2. Preparation of thin PU films 33
3.1.3. Media used in degradation experiments and 33
esterase activity analysis
3.1.4. Soil sample for isolation of PU degrading microorganisms 33
3.1.5. Isolation of PU degrading fungus 34
3.1.6. Isolation of polyurethane degrading bacterial strains 34
3.2. Identification of the Isolated Microorganisms 35
3.2.1. Identification of fungal isolate 35
3.2.2. Identification of bacterial isolates 35
3.2.2.1. Morphological characterization 35
3.2.2.2.Biochemical characterization 36
3.2.2.3. 16S rRNA gene sequence analysis 37
3.2.2.3.1. DNA extraction 37
3.2.2.3.2. Amplification of 16S rRNA gene 37
3.3. Polyurethane degradation assay 38
3.3.1. Scanning Electron Microscopy of PU film pieces 38
3.3.2. Fourier Transform Infrared Spectroscopy (FT-IR) analysis 39
3.3.3. Gel Permeation Chromatography (GPC) analysis 39
3.3.4. CO2 evolution Test (sturm test) 39
3.4. Esterase Activity Assay 41
3.4.1. Optimization of polyurethaneesterase production from 44
Bacillus subtilis MZA-75
3.4.1.1. Effect of temperature 44
3.4.1.2. Effect of pH 44
3.4.1.3. Effect of time of incubation 44
3.4.1.4. Effect of nitrogen source 45
3.5. Analysis of ester hydrolysis products by GC-MS 45
3.5.1. Growth of the isolated organisms with PU as sole carbon source 45
3.5.2. Extraction of samples from PU culture medium for metabolites 45
3.5.3. Analysis of the extracts on GC-MS 45
3.6. Utilization of 1, 4-butandiol and adipic acid as carbon source 46
3.7. Biofilm quantification by crystal violet staining/SEM observation 46
3.8. Production and purification of cell bound polyurethane esterase 46
3.8.1. Electrophoresis 47
3.8.2. Protein concentration determination 47
3.8.3. Substrate specificity of purified cell bound esterase from MZA-75 47
3.8.4. Degradation activity of purified esterase against PU films 48
3.8. Statistical analysis of results 48
Results
4.1. Isolation of polyurethane degrading microorganisms 49
4.1.1. Isolation of polyurethane degrading Fungus 49
4.1.2. Isolation of PU degrading bacterial strains 49
4.1.3. Growth of bacterial isolates on PU as carbon source 49
4.2. Identification of polyurethane degrading strains 53
4.2.1. Identification of PU degrading fungus 53
4.2.2. Identification of PU degrading bacterial strains 53
4.2.2.1. Biochemical analysis of MZA-75 and MZA-85 53
4.2.2.2. 16S rRNA gene sequence analysis of MZA-75 53
and MZA-85
4.3. Analysis of polyurethane degradation 58
4.3.1. Polyurethane degradation by Aspergillus tubingensis 58
4.3.1.1. SEM analysis 58
4.3.1.2. FTIR analysis 58
4.3.1.3. CO2 evolution test (sturm test) 61
4.3.2. Polyurethane degradation by bacterial strains MZA-75 61
and MZA-85
4.3.2.1. SEM analysis 61
4.3.2.2. FTIR analysis 61
4.3.2.3. GPC analysis 62
4.3.2.4. CO2 evolution test 62
4.4. Analysis of degradation products by GC-MS 71
4.4.1. Detection of ester hydrolysis products 71
4.4.2. Growth of MZA-75 and MZA-85 on 1,4-butanediol 71
and adipic acid
4.5. Analysis of Biofilm Formation by MZA-85 on the Surface of PU 72
4.6. Polyester PU degrading enzyme (esterase) from bacterial strains 79
4.6.1. Esterase assays for MZA-85 79
4.6.2. Esterase assays for MZA-75 79
4.6.3. Optimization of culture conditions for esterase 79
production from MZA-75
4.7. Purification of cell bound esterase from Bacillus subtilis MZA-75 80
4.7.1. Substrate Specificity of Purified PUesterase 80
4.7.2. Degradation activity of purified PU esterase against PU film 80
Discussion 91
Conclusion 100
Future Prospects 101
References 102
___________________________________________________________
i
ACKNOWLEDGEMENTS
All praises to Almighty Allah The Most Merciful and Beneficent, Whose
Blessings always enable me to pursue my goals. Praise to our Holy Prophet Hazrat
Muhammad (PBUH) whose teachings enable me to recognize our Creator Allah
Almighty.
I have this opportunity to express my gratitude and sincere appreciation to Dr.
Aamer Ali Shah for his guaidance and motivation. Throughout my stay in Q. A. U., I
found in him a great teacher and a true role model. His friendly advices and dedicated
personality motivated me to work harder to accomplish this task. I am also deeply
indebted to Dr. Fariha Hasan who looked after this project in the absence of Dr. Aamer
Ali Shah.
I am grateful to Prof. Dr. Asghari Bano, Dean Faculty of Biological Sciences
Quaid-I-Azam University Islamabad for providing access to facilities that ensure
successful completion of this work.
I would also like to appreciate Chairman Biotechnology Prof. Dr. Zabta Khan
Shinwari for his supportive attitude during my research work.
It is a matter of immense pleasure for me to express my sincerest feelings of
gratefulness to all teachers of Department of Microbiology especially Prof. Dr. Abdul
Hameed and Dr. Safia Ahmed for their cooperative attitude and humble guidance
throughout my research work.
I am extremely grateful to Dr. Lee Krumholz for supervising my work at the
Department of Botany and Microbiology, University of Oklahoma, Norman, Oklahoma,
USA. I would like to say thanks to Dr. Deniz F. Aktas for helping me with GC-MS. It
would not have been possible without her help.
It would be injustice not to mention my gratitude for assistance and motivation
that I received from my friends Dr. Samiullah Khan, Dr. Masroor Hussain, Dr. Fazal Ur
ii
Rehman Kakar, Dr. Syed Aun Muhammad, Muhammad Khurram Afzal and Fawad Ali
Bangash.
I am thankful to Higher Education Commission (HEC) Pakistan for providing me
monitory support in the shape of HEC indigenous and IRSIP scholarships.
No word of gratitude seems sufficient to describe the efforts my family made to
make me achieve this goal. Huge credit goes to my parents, my aunty, my brothers and
sister, my wife Sumera Farid and my son Muhammad Zarrar Shah whose innocent face
motivated and energized me to complete this task.
Zia Ullah Shah
iii
LIST OF TABLES
Table
No.
Title Page No.
4.1 Identification of polyurethane degrading bacterial strains 55
4.2 Sturm test for detection of CO2 produced as a result of degradation of
PU by MZA-75.
70
4.3 Sturm test for detection of CO2 produced as a result of degradation of
PU by MZA-85.
70
iv
LIST OF FIGURES
Fig
No.
Title Page
No.
1.1 Condensation reaction between polyisocyanate and polyol that produces
polyurethane.
2
2.1 Urethane linkage. 11
2.2 Polyurethane monomer. R is the hydrocarbon chain from alcohol group, while
R2 represents the hydrocarbon chain from diisocyanate group
11
2.3 Property matrix showing wide range of polyurethane applications (adopted
from media.wiley.com).
12
2.4 Plastic-microbe interaction; Microorganisms excrete extracellular enzymes,
which degrade large molecular weight plastics into short degradation
intermediates (water soluble). These intermediates are assimilated by the cells
and are converted into CO2 and H2O or other final products (adopted from
Muller, 2003).
16
2.5 Proposed biodegradation pathway (adapted from Gautam et al., 2007) 31
3.1 Work flow sheet for project ―Microbial degradation of polyester polyurethane‖ 32
3.2 Modified Sturm test. Arrows represent direction of air flow. Red bottles
containing 3M KOH constitute air pretreatment chamber (removes CO2 present
in the air). Black bottles are Control and Test having culture without and with
PU respectively and Blue bottles are CO2 absorption chamber having 1M KOH
(traps CO2 produced as a result of microbial metabolism).
40
3.3 Plot of Abs410 values against respective concentrations of p-Nitrophenol, used
for determination of milli Moles of p-Nitrophenol produced for determination
of specific esterase activity.
43
v
3.4 Standard curve of BSA used for determination of concentration of total
proteins.
43
4.1 Polyurethane film with fungus growing on MSM-agar plate. Photograph taken
after seven days of incubation at 30oC.
50
4.2 PU degradation by fungi. PU film treated with Aspergillus tubingensis for 4
weeks (B) shows signs of degradation as compared to untreated PU film (A).
50
4.3 Growth and colony morphology of the isolated fungal strain on malt extract
agar.
51
4.4 Growth of MZA-75 in MSM supplemented with PU as a sole source of carbon.
Test: culture with PU and Control: culture without PU or any other carbon
source.
52
4.5 Growth of MZA-85 in the presence of PU as sole source of carbon in liquid
MSM. Test represents growth of MZA-85 in the presence of PU, while Control
represents the growth in absence of any carbon source.
54
4.6 Pylogenetic tree of MZA-75 showing that it has maximum resemblance with
Bacillus subtilis.
56
4.7 Phylogenetic tree of MZA-85 showing that it has maximum resemblance with
Pseudomonas aeruginosa.
57
4.8 SEM of PU film treated with Aspergillus tubingensis for 1 month on MSM-
agar. Mycelial mass adhering to the surface of the film can be seen (B).
Damage caused by fungal mycelia can be seen (C). No mycelial mass seen on
untreated control (A).
59
4.9 The peak at 1696 cm-1
representing urethane linkage is absent in the treated PU
sample. Peaks representing amine linkage (C-N) i.e. at 1164.4 cm-1
and 1136.3
cm-1
are absent in treated PU film. The area 600-700 cm-1
representing C-H
deformation, No peaks was observed in this region.
60
vi
4.10 Cells of Pseudomonas aeruginosa MZA-85 adhering to the surface of treated
PU films with cracks radiating from the point of adherence (B). No such
changes could be seen in the untreated (abiotic) control.
64
4.11 SEM of PU films treated with Bacillus subtilis MZA-75, Small hair like
modifications can be seen on the surface (B) higher magnification reveals these
as cracks on the surface of PU film (C). No such cracks can be seen on the
surface of untreated (Abiotic) control (A).
65
4.12 FTIR spectrum of PU film treated with Bacillus subtilis MZA-75, reveals
disappearance of peak at 1725 cm-1
(B) which is present in the untreated
(abiotic) control (A).
66
4.13 FT-IR spectrum of MZA-85 treated PU film demonstrating polyester portion of
the PU as target for microbial degradation.
67
4.14 Gel permeation chromatography shows increase in the polydispersity index of
PU film treated with Bacillus subtilis MZA-75 for 4 weeks (B). changes in
number average molecular weight (Mn) and weight average molecular weight
(Mw) can also be seen when compared to untreated to control (A).
68
4.15 Gel permeation chromatography shows increase in the polydispersity index of
PU film treated with Pseudomonas aeruginosa MZA-85 for 4 weeks (B).
changes in number average molecular weight (Mn) and weight average
molecular weight (Mw) can also be seen when compared to untreated control
(A).
69
4.16 GC chromatogram overlay of ethyl acetate extract of Bacillus subtilis MZA-75
culture with PU (Red), culture without PU i.e biotic control (Blue), MSM
supplemented with PU without MZA-75 culture i.e. abiotic control (Black). 1,
4-butanediol and adipic acid peaks at retention time 13.10 minutes and 18.2
minutes respectively can be seen only in culture with PU (Black line). M/Z of
1,4-butanediol standard (B) M/Z of 1,4-butanediol extracted (C) M/Z of adipic
73
vii
acid standard (D) M/Z of adipic acid extracted (E).
4.17 GC chromatogram overlay of ethyl acetate extract of Pseudomonas aeruginosa
MZA-85 culture with PU (Black), culture without PU i.e biotic control (green),
MSM supplemented with PU without MZA-85 culture i.e. abiotic control
(Red). 1, 4-butanediol and adipic acid peaks at retention time 13.28 minutes and
18.5 minutes respectively can be seen only in culture with PU (Black line). M/Z
of 1,4-butanediol standard (B) M/Z of 1,4-butanediol extracted (C) M/Z of
adipic acid standard (D) M/Z of adipic acid extracted (E).
74
4.18 Gradual rise in growth of Pseudomonas aeruginosa MZA-85 utilizing 1,4-
butanediol (circle), Adipic acid (square) and control i.e. MSM without any
carbon source (triangle).
75
4.19 Gradual rise in growth of Bacillus subtilis MZA-75 utilizing 1,4-butanediol
(circle), Adipic acid (square) and control i.e. MSM without any carbon source
(triangle).
76
4.20 SEM of PU film treated with P.aeruginosa MZA-85 for one week (B) showing
the presense of biofilm the surface, while no biofilm is visible on the surface of
untreated control (A).
77
4.21 Biofilm quantification by crystal violet staining. Abs750 was taken as measure of
biofilm quantity.
78
4.22 Induction of membrane associated esterase by PU supplementation of the MSM
as depicted by plot of membrane associated esterase activity conducted for
MZA-85 after every 4 days. (triangle) represents membrane associated esterase
activity in culture with PU, while (circle) represent membrane associated
esterase activity in culture without PU.
81
4.23 Induction of extracellular esterase by PU supplementation of the MSM as
depicted by plot of extracellular esterase activity conducted for MZA-75 after
every 4 days. (triangle) represents extracellular esterase activity in culture with
82
viii
PU, while (circle) represent extracellular esterase activity in culture without PU.
4.24 Induction of cell bound esterase by PU supplementation of MSM as depicted by
plot of membrane associated esterase activity against time of incubation
conducted for MZA-75 after every 4 days (triangle) represents cell bound
esterase activity in culture with PU, while (circle) represent cell bound esterase
activity in culture without PU.
83
4.25 Optimization of temperature of incubation for esterase production from Bacillus
subtilis MZA-75; best results can be seen with 37 oC i.e. 0.493 mM/min/mg at
day 21st.
84
4.26 pH optimization for PU esterase production from Bacillus subtilis MZA-75;
best results can be seen on pH7 i.e. 0.593 mM/min/mg day 21st.
85
4.27 Analysis of PU esterase production from Bacillus subtilis MZA-75 in the
presence and absence of yeast extract (as Nitrogen source); best results can be
seen in the presence of nitrogen source i.e. 0.753 mM/min/mg at day 28th.
86
4.28 Optimization of time of incubation for production of PU esterase by Bacillus
subtilis MZA-75; best results can be seen on day 21st i.e. 0.543 mM/min/mg.
87
4.29 Esterase activity of different fractions collected from Sephadex G-75 elution of
cell bound esterase of MZA-75. Fractions 12-16 showing maximum activity.
Primary vertical exis represents enzyme activity (mM/min), while secondary
vertical axis represents Abs280 for rough estimation of total protein contents.
88
4.30 SDS-PAGE of purified PU esterase purified from MZA-75. Lane 1: protein
marker; Lane 2&3: PU esterase approximately 50 KDa.
89
4.31 Specific esterase activity of purified esterase against para Nitrophenyl acyl
ester with fatty acids of different carbon numbers. Best activity was observed
against para Nitro Phenyl butyrate i.e. 2.786 mM/min.
90
ix
LIST OF ABBREVIATIONS
AFM Atomic force microscopy
ANOVA Analysis of Variance
ASTM American Society for Testing and Materials
ATR-FTIR Attenuated total Reflectance-Fourier transformed infra-red
BLAST Basic Local Alignment Search Tool
BPU Biodegradable polyurethane
BSTFA N,O-bis (trimethylsilyl) trifluoro acetamide
CAGR Compound annual growth rate
CE Cholestrol esterase
CFCs Chlorofluorocarbons
DEAE Diethyl amino ethyl
dNTP Deoxy nucleoside triphosphate
EDTA Ethylene diamine tetra acetate
x
FTIR Fourier Transforms Infra-Red Spectroscopy
GC-MS Gas chromatography-Mass spectrometry
GPC Gel Permeation Chromatography
GPC Gel Permeation Chromatography
HCFCs Hydro chlorofluorocarbons
HPLC High Performance Liquid Chromatography
ISO/DIS International Organization for Standardization /Draft
International Standard
KDa Kilo Dalton
MEA Malt extract agar
MEGA Molecular Evolutionary Genetic Analysis
MM Millimeter
Mn Number average molecular weight
MSM Mineral Salt Medium
MSW Muncipal solid waste
MW Molecular weight
Mw Weight average molecular weight
NCBI National Center for Biotechnology Information
NIST National Institute of Standards and Technology
OD600 Optical density as measured from Absorbance at 600nm
PCL Polycaprolactone
xi
PCL-PU Polycaprolactone based polyurethane
PCR Polymerase Chain Reaction
PCU Polycarbonate Urethane
PE Polyethylene
PEU Polyether Urethane
PGA Poly glycolide
PLA Poly lactide
PMSF phenylmethylsulfonyl fluoride
pNPA Para nitrophenyl acetate
PSI Pounds per square inch
PU Polyurethane
PUs Polyurethanes
P-VALUE Probability Value
RPM Revolution Per Minute
RPU Rigid polyurethane foam
SBD Surface binding domain
SDA Sabouraud dextrose agar
SDS-PAGE Sodium Dodecyl Sulphate-Polyacrylamide Gel Electrophoresis
SEM Scanning Electron Microscopy
SMM Surface modifying molecules
THF Tetra hydro furan
xii
ABSTRACT
In this report, Polyurethane (PU) degrading microorganisms (fungi and bacteria) were
isolated from soil through enrichment. The isolated fungal strain was identified by
examination of colony morphology i.e. color, size and colony diameter and shape, color,
size and structure of conidia, hyphae, conidiophores and conidial head as Aspergillus
tubingensis. PU films incubated for one month on MSM-Agar plates inoculated with A.
tubingensis demonstrated visible signs of degradation in terms of changes in color and
flexibility. Thick mycelial growth and adherence of fungal biomass with surface of PU
was confirmed by scanning electron microscopy (SEM). Fourier transformed infra-red
spectroscopy (FTIR) spectrum of the treated PU film, when compared to that of untreated
control revealed changes in important functionalities. Two bacterial strains isolated from
the same soil were identified as Bacillus subtilis MZA-75 and Pseudomonas aeruginosa
MZA-85 by colony morphology, microscopy, biochemical characterization and 16S
rRNA gene sequence analysis. The degradation of PU film pieces exposed to both strain
MZA-75 and MZA-85 was investigated by SEM, FTIR and gel permeation
chromatography (GPC). SEM micrographs of PU film pieces, treated with strains MZA-
75 and MZA-85, showed alterations in the morphological features of surface. FTIR
spectrum demonstrated rise in organic acid functional groups and fall in ester
functionality. GPC results revealed increase in polydispersity, which shows that long
chains of polyurethane polymer are cleaved into shorter chains by microbial action.
Increase in cell growth and CO2 concentration detected through Sturm Test, in
comparison to control further elaborate the degradative capability of strains MZA-75 and
MZA-85. MZA-85 was found capable of producing cell associated esterase measured on
the basis of p-Nitrophenyl acetate (pNPA) hydrolysis assay. Time course study for cell
associated esterase in the presence and absence of PU in MSM broth revealed that this
enzyme is induced by the presence of PU in the medium. Crystal violet staining and SEM
results shows that MZA-85 forms biofilm on the surface of PU.
In case of MZA-75 increase in both cell bound and extracellular esterases was
observed in the presence of PUR films in MSM as compared to control when analyzed
through p-Nitrophenyl acetate (pNPA) hydrolysis assay. PUesterase was purified from
xiii
the MZA-75 by using Sephadex G-75 column chromatography. The purified enzyme
gave single band on SDS-PAGE corresponding to molecular weight 51 KDa. Substrate
specificity analysis was done using p-Nitrophenyl acyl esters of varying carbon numbers.
Maximum esterolytic activity was observed in case of p-Nitrophenyl butyrate (C4).
Analysis of the cell free supernatant by GC-MS, revealed that 1, 4-butanediol and
adipic acid monomers were produced as result of degradation of PU by both MZA-75 and
MZA-85 and both the strains were capable of utilizing these intermediates as carbon
source.
Both MZA-75 and MZA-85 are subject to further studies to understand their
interaction with PU completely, which may be helpful in PU bioremediation and
biochemical monomer recycling from PU wastes.
14
INTRODUCTION
With rising trend of urbanization and industrial development, large quantities of
different industrial products are being produced to improve living standards, which resulted
in both quantitative increase and qualitative diversification of solid wastes being generated.
In particular industry and general living have seen great transformations with the
development of polymeric material. However these materials when disposed of, lead to
pollution of water, soil and atmosphere, which has urged a global interest in the development
of safe disposal methods. Moreover, recycling of waste materials is getting increasingly
appealing due to the current cost of energy and resource. Therefore, it is highly desirable to
work out techniques for the processing of polymeric materials in stable and environmentally
friendly manner. The global demand for thermosetting resin is 20% of the total plastic
requirements worldwide and polyurethane (PU) contributes 50% to the total demand of
thermosetting resin (Im et al., 2008).
The term polyurethane (PU) is used for a polymer produced by condensation reaction
between polyisocyanates and polyols. This polymer is characterized by an intra-molecular
urethane bonds (carbamate ester bond, -NHCOO-) as shown in figure 1.1 (Nakajima-Kambe
et al., 1999). Three basic components are required for the synthesis of PUs: a diisocyanate, a
polyhydroxy alcohol and a chain extender (various low-molecular weight pre-polymer
blocks). Due to presence of terminal hydroxyl groups alternating blocks, referred to as
―segments‖ form in the PU chain. Blocks comprising of isocyanate and chain extender
constitute rigid crystalline phase and are called ―hard segments‖. Blocks producing non-
crystalline or low crystallinity phase are termed as ―soft segments‖ (Young and Lovell. 1994;
Barbari. 1995). Usually, properties like hardness, tensile strength, impact resistance, modulus
and stiffness are dependent on the nature of hard segment, while the soft segment determines
properties like water absorption, softness, elasticity and elongation. The degree of tensile
strength and elasticity can be changed by modifications in the above mentioned segments of
the polymer. Therefore production of versatile PU polymers by bringing about modifications
in the molecular structures of soft and hard segments (Zhang et al., 2003).
15
Fig. 1.1: Condensation reaction between polyisocyanate and polyol that produces
polyurethane (adopted from Nakajima-Kambe et al., 1999)
16
Based on types of polyols i.e. polyether polyol and polyester polyol, two types of PU
can be synthesized. PU synthesized from polyether polyol is referred to as polyether PU,
while PU synthesized from polyester polyol is called polyester PU. Although most of the
PUs used nowadays are polyether type but polyester PU gained importance recently because
of its superior biodegradability and good material properties. It is considered as environment
friendly from the view point of waste treatment.
Because of their versatile physical and mechanical properties, PUs have widely been
employed in different industrial applications. Some of the biodegradable PUs have been used
as biomaterial application (Lee et al., 2001, Moon et al., 2003) for manufacturing of medical
devices such as vascular grafts, values, artificial heart diaphragms, connecting modules for
cardiac pacemakers, neurological lead insulation and catheters (Labow et al., 2001). It has a
great potential in different environmental applications and can be used for the prevention and
control of pollution. High density PU foam can be used as barrier to check direct contact
between the hazardous pollutants and the environment. The PU foam suitability for these
applications depends on the aging behavior, degradation due to environmental and
mechanical factors and structural integrity. Numerous factors like moisture, sunlight, heat,
temperature variations, chemical pollutants, radioactivity and microorganisms etc. cause
aging and degradation of PU barrier (Erlandsson et al., 1997). The mechanisms responsible
for biotic and abiotic degradation of PU can occur simultaneously or subsequently.
Polyurethane waste is currently managed by landfilling and recycling. In developed
countries most of the disposed of PU ends up in landfills (Helsinki university report 2004).
Recycling of PU is done mechanically by regrinding, flexible foam bonding, adhesive
pressing, and compression molding of the PU wastes or chemically by feed stock recovery
(recovery of the monomers) employing techniques like glycolysis, hydrolysis, pyrolysis and
hydrogenation (www.polyurethane.org, 2007).
Biodegradation may offer an advanced answer to the problem of PU wastes (Gautam
et al., 2007). The attack of microorganisms on non-water soluble materials of polymeric
nature is called biodegradation of polymers (Muller, 2003). During this complex process the
carbon based polymeric structure is mineralized into carbon dioxide and biomass. The first
step of biodegradation is catalyzed by extracellular enzyme (enzymes which are secreted by
17
microorganisms in the surrounding medium). In case the polymeric substrate is polyester the
expected site of attack for enzymatic hydrolysis is ester bond, which degrades the
hydrophobic polymer into oligomers and monomers. This initial fragmentation is required to
decrease the molecular weight and increase water solubility of the polymer so that they can
pass through the cell membrane for subsequent metabolism by microorganisms (Tokiwa and
Suzuki 1974, 1977; Herzog et al., 2006; Muller, 2006). As a result of this initial
depolymerization environmental concentrations of oligomers and monomers are built up,
which allow their passage from one environmental component to another (Degli Innocenti,
2005). Microorganisms and their enzymes can also be used for bio chemical monomer
recycling because of their ability to catalyze the hydrolysis of specific substrate as compared
to chemical hydrolysis methods which generate mixture of monomers when mixed wastes
containing different polyesters are treated (Kim and Rhee, 2003, Shah et al., 2008; Shimao,
2001; Suyama et al., 1998; Tokiwa and Calabia, 2008)
Environmental degradation of PU depends upon the type of PU, its chemical and
structural compositions. Several parameters like conditions during processing, chemistry,
additives, morphology, degree of crystallinity and ratio of soft to hard segment etc., are
responsible for the variation in degradability of PU (Erlandsson et al., 1997; Albertson et al.,
1987; Labow et al., 2005; Kim and Kim, 1998; Marten et al., 2000).
Biodegradation of polyester based polyurethanes is easier as compared to polyether
polyurethanes and the presence of at least three methylene groups of unbranched carbon
chains in a row between urethane linkages is necessary for significant enzymatic attack to
occur (Darby and Kaplan, 1968). Increase in polyester chain length leads to decrease in
enzymatic and hydrolytic degradation of the polymer. Different fungal species present in soil
microflora for example Curvularia sengalensis, Aureobasidium pullulans, Fusarium solanii,
and Cladosporium sp. are reported to degrade PU (Crabbe et al., 1994). Some bacterial
species like Pseudomonas fluorescens, Bacillus subtilis, P. chlororaphis and Comomonas
acidovorans are also known for their polyurethanolytic capability (Nakajima-Kambe et al.,
1995, 1997; Howard et al., 1998; Ruiz et al., 1999; Howard et al., 1999; Rowe and Howard,
2002). For PU wastes management we need bacteria efficient in PU-utilizing ability, which
can be explored in various ecological niches by testing their ability in pure culture or
consortia. Polyester PU is more readily degraded by fungi than polyether PU (Darby and
18
Kaplan, 1968). A number of polyester PU degrading fungi have been isolated and
characterized (Morton and Surman, 1994; Huang et al., 1981; Pathirana and Seal, 1984;
Crabbe et al., 1994). The polyester PU degradation has been attributed to hydrolytic enzymes
(esterases) causing hydrolysis of ester linkages. In case of bacteria the same mechanism that
is ester bond hydrolysis is considered important for the degradation of PU. Although data
regarding bacterial degradation of PU is limited and degradation pathways and enzymes are
not fully documented, however it is assumed that degradation of PU may occur due to
utilization of the polymer by microorganisms as carbon and /or nitrogen source (Akutsu et
al., 1998) or due to co-metabolism which takes place in the presence of other nutrients.
Polyurethane being a versatile polymer, a coordinated effort on the part of both
environmental microbiologists and organic chemists is needed, to elucidate the complete
mechanism of biodegradation of this polymer.
Polyurethanase enzymes isolated and characterized so far are of two types; cell
membrane bound PU-esterase also called as cell associated PU-esterase and soluble PU-
esterase also known as extracellular PU-esterase (Akutsu et al., 1998: Allen et al., 1999;
Vega et al., 1999; Ruiz et al., 1999). These types of enzymes appear to have separate
functions in the deterioration of PU. The cell associated PU-esterase would allow contact of
microbial cells with the insoluble PU while, the extracellular PU-esterase would adhere to the
polymer surface and proceed the process of hydrolysis. The proposed mechanism of PU
degradation by Pud A is that PU is degraded by this enzyme in two steps: hydrophobic
adsorption of the enzyme on the polymer surface and then breakdown of ester bonds of PU.
The PU esterase was thought to possess a surface binding domain (SBD) that adheres to the
surface of PU hydrophobically and a catalytic domain that proceeds hydrolysis. The presence
of SBD in PU-esterase was shown to be necessary for the degradation of PU (Akutsu et al.,
1998).
The hydrolytic degradation of the polyester PU can be increased by the addition of
hydrophilic monomers into the polymer chain which helps in water attack. A reasonable
number of PUs and their composites have been synthesized from natural resources and are
termed as biobased PUs (Dwa‘nIsa et al., 2005). Sugar-based [m,n]-polyurethanes have been
synthesized using selectively protected derivatives of hexoses (Garçon et al., 2001;
19
Hashimoto et al., 2005) aldaro (Hashimoto et al., 1995) and aldonolactones, (Yamanaka and
Hashimoto et al., 2002) alditols,(Paz et al., 2007; Paz et al., 2008; Marin and Munoz-Guerra,
2008; Marin and Munoz-Guerra, 2009) or anhydroalditols, (Marin and Munoz-Guerra, 2009;
Braun et al., 1992) as diol monomers. Such diols have been polymerized with diisocyanate
monomers to afford the corresponding [m,n]-polyurethanes.
Conventional polymers like polyethylene and polypropylene are non-biodegradable and
cause substantial environmental problems. These polymers can be replaced by biodegradable
PU (BPU) in near future to reduce environmental burden (Moon et al., 2003). Polyurethanes
prepared from aliphatic polyesters of natural origin will likely constitute a group of
biodegradable polymers of great economic significance (Edlund et al., 2003). BPU are
expected to have widespread applications in the fields of medicine and environment. BPU is
usually produced by using easily hydrolysable soft segments into the polymer chain, such as
poly(3-caprolactone) (PCL), poly(alkylene adipate), poly(lactide) (PLA), and poly(glycolide)
(PGA) into PU. In particular, PCL-based PU (PCL-PU) has been the most widely studied
because of the high modulus and ultimate tensile stress of PCL soft segment (Pena et al.,
2006).
Biodegradation is monitored by using analytical tools like visual observations
(changes in the surface of polymer films such as appearance of holes and cracks, altertions in
color or development of biofilms on polymer surface), SEM, changes in material properties
and molecular weight (Erlandsson et al., 1997), measurement of weight loss and formation of
clear zone on agar plates with polymer dispersed (Nishida and Tokiwa, 1993; Abou-Zeid,
2001), changes in molar mass and mechanical properties, CO2 evolution/O2 consumption
(Puchner et al., 1995) through sturm test. Sturm test give direct indication of polymer
backbone mineralization to end products i.e CO2 in aerobic conditions. Radiolabeling helps
in cases where slowly degrading polymeric substances are monitored for degradation in an
environment having other sources of carbon as well (Albertsson, 1978). For development of
biodegradation techniques for plastic wastes it is important to understand their metabolism by
existing microorganisms along with search for new potential degraders of microbial origin.
Alongside biodegradation studies, PU synthesis also needs some changes like
replacement of phosgene or even isocyanates in the synthesis of PU. The biodegradation
20
studies of isocyanates especially aryl isocyanates also need to be carried out because the
currently available data on the biodegradation of polyester polyurethane only indicates the
enzyme catalyzed hydrolysis of ester linkages leaving aside the fate of isocyanates released.
Although polyurethanes are recalcitrant in nature but the availability of innumerable
microbial resources with versatile catalytic capabilities bring hope to environmental
biotechnologists to deal with solid waste through the development of practically feasible
processes of bioremediation.
The current study focused on isolation and identification of microbial strains from
local soil microflora with polyester PU degradation capability, understanding the role of
microbial enzymes and identification of the products of degradation. Since polyester segment
of PU can be hydrolyzed to polyols and organic acids, microorganisms with substantial
esterolytic capability can not only be used to get rid of polyester based polymers but may
also be useful in recycling of the monomers constituting the polyester, like polyhydroxy
alcohols and organic acids. The ability of the isolated microorganisms to mineralize the
polyester based PU was also preliminarily evaluated to investigate the extent of degradation.
21
Aim and Objectives
1. Isolation of PU degrading microorganisms from local soil micro-flora;
2. Identification of the isolated strains using morphological, biochemical and genetic
characteristics;
3. To study the ability of the isolated strains to degrade PU by bringing about changes in
the chemical structure, surface feature and molecular weight of the exposed polymer
through FTIR, SEM, and GPC.
4. To study the role of esterase in the degradation of polyester based polyurethane;
5. Analysis of PU degradation products by gas chromatography mass-spectrometry
(GC-MS).
9
LITERATURE REVIEW
2.1. Importance of Plastics in Our Daily Life: Polyurethane
Plastics are man-made molecules of polymeric nature (Scott, 1999). They are
extensively used because they are relatively economical and have excellent material
properties. They can be easily manufactured and molded. Plastic do not easily degrade in
the environment because of their stable nature, and their accumulation cause significant
environmental pollution. In particular plastics of synthetic origin are recognized as an
environmental problem. According to statistics issued by US EPA a massive amount of
municipal solid waste (MSW) weighing approximately 236 million tons was produced in
the United States during 2003. The share of plastics in the total amount of MSW
generated was 11.3%. Only a minor portion of this plastic waste consisting generally of
bottles used for soft drinks and other purposes was recycled and the rest of the amount
needed disposal (US EPA, 2005). Polyurethane has a considerable share and is
considered as the 5th
major contributor to the colossal amount of plastic consumed every
year. In the United States alone, PU production increased from 45,000 tons in 1960 to
2,722,000 tons in 2004 (Uhlig, 1999; Howard, 2012).
2.2. Introduction of Polyurethane
Polyurethanes a group of polymers in which each monomers contain a urethane
functional group, were first synthesized by Dr. Otto Bayer in 1937. Urethanes are
carbamic acid derivatives existing in their esters form. Figure 2.1 represents generalized
structure of urethane linkage. Different urethanes can be produced by bringing variations
in the R group and introducing substitutions at the amide hydrogen. Apart from urethane
groups, other moieties like urea, ether and ester or an aromatic can also be included in the
structure of polyurethanes to reduce the number of urethane moieties in the polymer
chain. The urethane linkage is produced through the reaction of an isocyanate with an
alcohol (Dombrow, 1957; Saunders and Frisch, 1964; Kaplan et al., 1968). During
10
reaction the hydrogen atom from the hydroxyl group is transferred to the nitrogen of the
isocyanate. Presence of heteroatoms like oxygen and nitrogen in the polymer chain is the
major advantage of polyurethanes.
Figure 2.2 represents polyurethane monomer, produced by reaction between
polyhydroxy alcohol and polyisocyante. R represents hydroxylated hydrocarbon and R2
represents a hydrocarbon chain associated with diisocyanate functional group and n is the
number of these urethane monomers. Because of their high reactivity towards active
hydrogen containing compounds, diisocyanates are used in polyurethane production
reactions (Dombrow, 1957). Polyhydroxyl compounds can be used for industrial
applications. Similarly at the amide linkage polyfunctional nitrogen compounds can be
used. Different types of PU can be synthesized by varying polyhydroxyl and
polyfunctional nitrogen compounds e.g. polyether and polyester polyurethane in which
polyether and polyester resins containing hydroxyl groups are used respectively
(Dombrow, 1957; Urbanski et al., 1977).
Variety of PU can be produced by changing the number substitutions and space
between and within branch chains. PU produced can be linear, branched, flexible or rigid.
Fibers are produced using linear PU, while flexible PU is employed in the manufacture of
binding agents and coatings. Flexible and rigid foamed plastics which constitute the
major portion of PU produced can also be found in different forms in the industry
(Urbanski et al., 1977; Saunders and Frisch, 1964; Barbari, 1995). PU chain consists of
alternating blocks, called segments of rigid crystalline phase consisting of the isocyanate
and chain extender known as the hard segment and amorphous rubbery phase consisting
of polyester or polyether diols are recognized as the soft segment. These block polymers
are commercially known as segmented PUs. The tensile strength and elasticity of the
polymer can be changed by varying the composition of these segments (Barbari, 1995;
Young and Lovell, 1994). Since PUs with broad range of material properties can be
synthesized, it is used for wide range of purposes like in liquid coatings and paints,
adhesives, sealants, flexible and rigid foams and elastomers. Figure 2.3 represents various
applications of the versatile polymer polyurethanes in different areas.
11
Fig. 2.1: Urethane linkage. R represents hydrocarbon chain from alcohol
Fig. 2.2: Polyurethane monomer. R is the hydrocarbon chain from alcohol group, while
R2 represents the hydrocarbon chain from diisocyanate group
12
Fig. 2.3: Property matrix showing wide range of polyurethane applications (adopted from
media.wiley.com)
13
2.3. Environmental Concerns about Plastics Waste: Polyurethane
Plastics represent a huge group of material with global annual production that has
doubled in quantity during 15 years (245 million tons in 2008) (Lithner, 2011). Plastics
wastes are ubiquitous in the environment with marine environment being the major
recipient of plastic contamination (Lithner, 2011). The estimated amount of plastic that
goes into sea annually is one million ton. Plastic debris is known to harm and/or kill
many marine species. This threat to the survival of different marine species is a matter of
great concern because many of these species are already endangered due to other forms
of human activities. Plastic litter when ingested by marine animals entangles in their guts
and harms them. Other threats of lesser importance are the usage of plastic wastes by
‗‗invader‘‘ species and the absorption of polychlorinated biphenyls from ingested
plastics. Less obvious forms, for example plastic in pellets or scrubbers are harmful also
(Derraik, 2002).
UV radiations, air and water currents can lead to formation of plastic particles less
than 5mm in size, also referred to as microplastics. These microplastics are present both
on land and in sea and can be more dangerous than larger size plastics. In sea these
microplastics may easily be adsorbed by sea plants or ingested by sea animals, which
then act as carrier for these microplastics (Barnes et al., 2009; Andrady, 2011).
Microplastics may also be produced when larger size plastics are ingested by animals
with strong digestive system (Franeker, 2011). These microplastics accumulate in the
environment and are considerable source of health and environmental hazards.
Incorrectly managed landfills can lead to escape of plastics or hazardous
chemical leachates like biphenyls, phthalates and to the environment. These chemical
may cause endocrine, circulatory, respiratory, reproductive and neural symptoms
(Talsness et al., 2009; Meeker et al., 2009; Oehlmann et al., 2009; Hengstler et al., 2011).
The unofficial plastic disposal methods used specially in the developing countries like
burning of plastic insulated wires to retrieve metal can also cause release of toxic
chemical to the environment.
14
Terrestrial wildlife is also affected by plastic litter. Plastic material entangles in
the digestive system and block digestion , especially when it is porous or in the form of
bags (Whitney et al., 1993; Jayasekara et al., 2005). Polyurethane contributes
significantly to the plastic wastes disposed of every year. Isocyanates which are part of
chemical composition of PUs is highly toxic affect not only worker during manufacturing
but also of people living in areas surrounding the manufacturing facilities of PU. Apart
from isocyanates toxic metal catalysts are used during PU manufacturing, which may
release from PU into the environment. Hormonal process may be disrupted in animals
exposed to these chemicals (http://www.environmentalhealth news.org).
2.4. Plastics Waste Management: Polyurethane
Widespread use of polyurethane due to its numerous applications has brought
with it a huge amount of polyurethane waste which has to be disposed of safely at a
suitable place in such a way that is environment friendly (Zia et al., 2007). One way to
get rid of polyurethane wastes is to degrade them for energy and feedstock recovery.
Reactions that lead to bond breakage and chemical changes in chemical structure of
polymer are referred to as polymer degradation reactions. These reactions may be carried
out by physical, chemical or biological agents (Pospisil and Nespurek, 1997). Based on
the nature of the agent causing polymer degradation, it has been classified as thermal
degradation, photo-oxidative degradation, ozone-induced degradation, catalytic
degradation, mechano-chemical degradation and biodegradation (Grassie and Scott,
1985). These phenomenon happen in the environment naturally or are conducted for
waste removal, feedstock recovery and energy recovery. For the later purpose, these
methods are either expensive or environment non friendly. Biodegradation provides an
attractive opportunity to deal with solid wastes of polymeric origin because of its
inexpensive and environment friendly nature.
15
2.4.1. Biodegradation of Polyurethane
Different agencies defined biodegradation and biodegradable plastics similarly.
Japanese society of biodegradable plastics (1994) defines biodegradable plastics as
follows:
“Polymeric materials, which are changed into lower molecular weight
compounds where at least one step in the degradation process is through
metabolism in the presence of naturally occurring organisms”.
Heterotrophic bacteria and fungi can use plastics as a potential nutrient source.
Characteristics like molecular weight, physical forms and crystallinity influence
biodegradability of plastics (Gu et al., 2000). Generally biodegradation of polymers is
inversely related to the molecular weight of polymers i.e. the higher the molecular
weight of polymers the lower its biodegradation and vice versa. The reason behind
decline in biodegradation with rise in molecular weight is the decrease in solubility of
polymers with increase in molecular weight. Polymers must be broken down to low
molecular weight oligomers and monomers so that microorganism can assimilate it for
further degradation and mineralization. The biodegradation of polymeric materials occur
in several steps all catalyzed by enzymes (Fig. 2.4). The most important type of
enzymatic cleavage reaction is hydrolysis (Schink et al., 1992). Glycosidic, peptide and
ester linkages are hydrolyzed by nucleophilic attack on carbonyl carbon atom.
Biodegradation of polymeric material is affected by:
1. Presence of enzymes and microorganisms:
The degradative enzymes may either be induced by the presence of the polymer or they
may be produced constitutively by the organism.
2. Biotic availability of the polymer:
The crystallinity of the polymer determines the accessibility of the cleavable bonds to
the enzymes. The products formed as a result of early degradative reactions should be
similar to compounds present in nature and biodegradable.
16
Fig. 2.4: Plastic-microbe interaction; Microorganisms excrete extracellular enzymes,
which degrade large molecular weight plastics into short degradation intermediates
(water soluble). These intermediates are assimilated by the cells and are converted into
CO2, H2O and other metabolic products (adopted from Muller, 2003).
17
3. Abiotic factors:
Conditions like the presence or absence of oxygen, appropriate temperature and pH
and availability of nutrients to microorganisms also influence biodegradation of the
polymer (Gu and Gu, 2005).
Polymer degrading microorganisms usually secrete hydrolases into the
surrounding milieu, to convert the inassimilable polymer into water soluble low
molecular weight compounds, which are ingested by the cells for mineralization (Gu et
al., 2000). Although urethane and urea groups are hydrolysable, their accessibility to the
hydrolyzing enzymes is difficult and degradation beyond polymer surface may never
happen, polyether PU although less biodegradable, has demonstrated higher release of
radiolabeled products consistently in comparison to the untreated control, from soft
segment when treated enzymatically. The ester linkage may be protected by hydrogen
bonding and secondary structures inside hard segment (Santerre et al., 1994). Increase in
the size of hard segment increases integration of carbonyl groups into secondary
structures, which results in reduced polymer chain mobility and reduced access of
enzymes to the carbonyl groups for degradation (Santerre and Labrow, 1997). PU
constructed from longer repeating units and hydrophilic groups have lesser chances of
packing into highly crystalline regions as normal PU does, and is more accessible to
enzymes responsible for biodegradation (Huang and Roby, 1986). PU containing
polycaprolactone diols have been found biodegradable; moreover it has been found that
increasing the chain length of the polyester increases the extent of degradation.
Research is in progress to understand the effect of chemical additives to the
structure of PU on its biodegradation. Sulfur curing has been found to give some fungal
inertness to PU, however fungal growth still occurs on PU, even after addition of
fungicide to the sulfur cured and peroxide cured PU (Kanavel et al., 1966). In medical
field, PU shows resistance to macromolecular oxidation, hydrolysis and calcification
(Marchant, 1992). Polyurethane elastomers are preferred over other elastomer due to
their better elasticity, tear resistance and toughness. It also resist oxidation and
hydrolysis (Dombrow, 1957; Saunders and Frisch, 1964; Ulrich, 1983). Surface
18
modifying molecules (SMM) such as those giving fluorinated and phosphonated end
groups to the base PU can decrease biodegradation, however some SMM and PU
formulations are non-compatible and increase biodegradation (Tang et al., 1997;
Baumgartner et al., 1997).
2.4.1.1. Fungal Degradation
Microorganisms degrade PUs by using hydrolase enzymes like esterases,
proteases and ureases (Evans and Levisohn, 1968; Hole, 1972; Flilip, 1978; Griffin,
1980). According to some reports PUs can be degraded by fungi (Darby and Kaplan,
1968; Kaplan et al., 1968; Ossefort and Testroet, 1966). These reports reveal that
polyester polyurethanes are relatively more readily degraded by fungi than polyether
type, which is moderately resistant to degradation. Enzymes demonstrating esterase and
polyurethane hydrolase activities have been isolated from fungi Chaetomium globosum
and Aspergillus terreus by using liquid PU in the growth medium as substrate
(Boubendir, 1993). Geomyces pannorum and Phoma sp. has been isolated from PU films
buried in acidic and neutral soil. Both the fungal isolates were found capable of
degrading Impranil DLN (Cosgrove et al., 2007). In a follow up study it has been found
that biostimulation of soil microcosm i.e addition of yeast extract alone or a combination
of yeast extract and Impranil DLN improved degradation of PU to 62% in comparison to
that in the control soil with no additives, it was also found associated with 45% increase
in potentially degrading microorganisms colonizing PU. Bioaugmentation studies
involving soil inoculation with wheat biomass (fungal growth on the surface of sterile
wheat) of the isolated fungi revealed increase in biodegradation. Addition of wheat
colonized with Penicillium viridicatum, Nectria haematococca, P. ochrochloron, or an
unidentified Mucormycotina sp. has been found to increase the degradation of buried PU
(Cosgrove et al., 2010). These results suggested that both biostimulation and
bioaugmentation were operating together to increase PU degradation. Bioaugmentation
was also found to bring about change in the local fungal populations and increased the
number of indigenous PU degrading fungi, suggesting increased biodegradation. These
results demonstrate that both bioaugmenatation and biostimulation may be very useful for
bioremediation of PU.
19
Two endophytic fungi i.e. Pestalotiopsis microspora isolates were found to grow
on PU as carbon source in both aerobic and anaerobic conditions. A serine hydrolase was
found responsible for this polyurethanolytic activity of the isolates (Russell et al., 2011).
Amaral et al., (2012) studied the degradation of two lignin based rigid polyurethane
foams (RPU) and compared to the degradation of commercially available polyether PU,
Lubrenol, when both types were treated with A. niger in liquid media and soil, under
same conditions. Results as observed from surface morphology analyzed by optical
microscopy and scanning electron microscopy suggest both lignin based RPU were more
prone to microbial degradation than commercial samples.
Polyurethane contaminated sands have been found to harbor PU degrader fungal
organisms. The most common enzyme according to a report responsible for their
polyurethanolytic activity was urease, however protease, esterase and laccase activities
have also been found in these isolates (Loredo-Treviño et al., 2011). Several PU
degrading isolates including Fusarium solani, Alternaria solani, Spicaria spp., A.
fumigatus, A. terreus, and A. flavus have been isolated from soil, wall paints, pieces of
plastic debris and plastic shields of the street lights and studied for their ability to degrade
PU by different methods including (1) direct plating, (2) clear zone in a 2-layered agar
media, and (3) liquid shaking culture (Ibrahim et al., 2011).
These findings suggest that PU degrading fungi have widespread distribution in
the environment. Continuous endeavor in search of PU degraders of fungal origin may
ultimately be helpful in bioremediation of PU solid wastes.
2.4.1.2. Bacterial Degradation
Various bacterial species have been isolated for their ability to degrade PUs. In
another study 16 bacterial strains were tested for their polyester PU degradation ability.
Seven of the isolated microorganisms were found to degrade PU when yeast extract was
added to the medium. A Pseudomonas aeruginosa and a Corynebacterium sp. could
degrade PU in basal medium. However, none of the isolates demonstrated any growth on
PU alone. Changes in physical properties like tensile strength and elongation were found
20
associated with the treated PU samples. Infra-red spectrophotometric analysis revealed
hydrolysis of ester linkage as a result of microbial treatment in the exposed PU samples.
It was also found that esterase production in the isolated organisms was inhibited by the
addition of glucose to the basal medium; however the addition of PU could not increase it
(Kay et al., 1991; 1993). In another study Acinetobacter calcoaceticus, Arthrobacter
globiformis, P. aeruginosa and P. putida were found capable of utilizing military aircraft
paint as a soul source of carbon and energy. These species showed esterase activity even
in the absence of PU, suggesting constitutive nature of these enzymes (Halim et al.,
1996).
The analysis of Bacillus sp. PU system was done by employing techniques like
electrophoretic mobility, electrical impedance, and dynamic light diffraction
measurements. The results obtained revealed that Bacillus cells bind to PU, form flocs
which is followed by degradation of the exposed PU particles. It was proposed that in
Bacillus PU system, two populations of cells exist, one which is coated with PU and the
other which is free. The coated cells degrade PU into soluble metabolites, which are
consumed by the uncoated cells as a nutrient source (Blake and Howard, 1998; Rowe and
Howard, 2002).
At least four Pseudomonads have been reported to degrade polyester
polyurethane. Comamonas acidovorans has been found growing on colloidal polyester-
polyurethane. Various polyester PUs was synthesized in the form of solid cubes by
varying polyester segments. Using different polyester segments polyester PUs was
prepared in the form of solid cubes, which were tested for degradation. Complete
degradation was observed when they were used as the only carbon source within a week.
However when they were used as sole carbon and nitrogen source, only 48% of the mass
degraded. Gas Chromatographic analysis of the PU degradation products showed most of
them were derived from the polyester segment of the treated PU sample. Main
metabolites observed were diethylene glycol, trimethylolpropane, and dimethyladipic
acid (Nakajima-Kambe et al., 1995; 1997). In agreement with these findings, Gautam et
al. (2007a) examined the biodegradation of polyester-polyurethane foam by P.
21
chlororaphis ATCC 55729. Both ammonia and diethylene glycol rose in concentration
with time, which was accompanied by rise in bacterial growth and fall in the mass of PU.
Pseudomonas fluorescens was found to degrade and utilize polyester polyurethane as a
sole carbon and energy source. Polyurethane utilization by P. fluorescens followed
simple Michaelis–Menten kinetics. The Ks and μmax values were 0.9 mg ml−1
and 1.61
doublings h−1
, respectively (Howard and Blake, 1998). Mukherjee et al., 2011 isolated
polyester polyurethane degrading P. aeruginosa from soil. High performance thin layer
chromatography results showed that the isolated strain is capable of degrading 32% of the
exposed polymer within 10 days. Maximum degradation was observed during log phase
of growth. Polyurethanolytic ability of the organism was attributed to extracellular
esterase production.
Oceguera-Cervantes (2007) isolated two Alicycliphilus sp. strains BQ1 and BQ8
for their ability to grow on N-methylpyrollidone (NMP) based PU foam. Their ability to
utilize NMP and degrade the exposed polymer was confirmed from IR spectrum and
SEM of the exposed polymer film. A soil bacterium identified as Acinetobacter gerneri
P7, was isolated for its PU degrading ability. This organism was found to grow on PU
and its potential to degrade PU was characterized by SEM observations. An esterase
catalyzing PU degradation was also purified and characterized from this strain (Howard
et al., 2012).
2.4.1.3. Polyurethane Degrading Enzymes
Enzymes can easily bind to water soluble substrates and hydrolyze them.
However water insoluble substrates like PU is difficult to bind with enzymes. In order to
overcome this, enzymes that catalyze the hydrolysis of water insoluble substrates bear
some characteristics that help them bind to the surface of water insoluble substrates (Van
Tilbeurgh et al. 1986; Fukui et al. 1988; Hansen 1992). A polyurethane esterase PUD A
is observed to catalyze polyurethane degradation in two steps i.e. hydrophobic adsorption
on the surface of PU through its surface binding domain (SBD) and then hydrolysis of the
substrate (PU) through its catalytic domain (Akutsu et al. 1998). So far, two types of PU
hydrolyzing enzymes have been isolated and characterized:
22
1. Membrane associated PU esterase and
2. Extracellular PU esterase
Membrane bound enzyme is responsible for the access of cells to the surface of
PU, which is followed by binding of the extracellular enzymes and hydrolytic action.
Data published so far suggest that degradation of PU usually occurs due to esterolytic
degradation of the polyester chains in the PU, which is mediated by esterase (Howard and
Hillard. 1999; Allen et al., 1999; Vega et al., 1999; Nakajima-kambe et al., 1995).
Interestingly, different bacterial species demonstrate ability to hydrolyze PU
enzymatically that is inhibited in the presence of serine hydrolase inhibitors. The role of
esterase and/or protease in the biodegradation of Impranil DLNTM
can be inferred from
this data. (Howard and Hillard. 1999). Comomonas acidovorans strain TB-35 possesses
two esterolytic enzymes, an extracellular esterase and a membrane-bound esterase. The
membrane-bound esterase was observed to have the ability to degrade the majority of
polyester PUs. Purification and characterization of this protein revealed that it has a
molecular mass of 62 kDa, it is heat stable up to 65oC and it loses its polyurethane
degrading activity in the presence of N,N-bis(3-d-gluconamidopropyl) deoxycholamide
(deoxy-BIGCHAP), while retaining its esterolytic activity against para Nitrophenyl
acetate (pNPA). These observations indicated that this enzyme degrades PU in a two-step
reaction: hydrophobic adsorption to the PU surface and hydrolysis of the ester bond of
PU (Akutsu et al., 1998).
P. chlororaphis was found to secrete extracellular polyurethanase enzymes.
Three active protein bands were obtained when these polyurethanolytic proteins were
analyzed on non-denaturing polyacrylamide gel electrophoresis having Rf values of 0.25,
0.417 and 0.917. One of these proteins was purified and demonstrated esterase activity.
This enzyme was inhibited by phenylmethylsulfonyl fluoride (PMSF) and had a
molecular weight of 27000 Da (Howard et al., 1999). PU degrading enzymes have also
been reported from P. fluorescens with molecular size 29,000 daltons and inhibited by
PMSF (Howard and Blake, 1998). Mukherjee et al., 2011 suggested the role of
extracellular esterase in the degradation of PUs, however the enzyme was not purified.
23
A 66 kDa polyurethane esterase was purified from Acinetobacter gerneri. This
enzyme was inhibited by phenyl methylsulfonyl fluoride (PMSF) and ethylene diamine
tetra acetate (EDTA). When tested against different p-Nitrophenyl substrates, it showed
best activity against p-Nitrophenyl propanate (Howard et al., 2012). Most of these
enzymes are serine hydrolases and are either extracellular or memberane bound.
Proteolytic enzymes, papain and urease can degrade segmented, cross-linked
polyester PU of biometric origin. Even though cross linking was negatively correlated
with the extent of degradation (Kaplan et al. 1968), papain (molecular weight 20.7 kDa)
has been proposed to diffuse into the bulk of the polymer and breaks the structural
integrity. Urease activity, because of its size (molecular weight 473 kDa), has been found
limited to the PU surface. It has been proposed that papain degrades the polymer by
hydrolyzing the urethane and urea linkages and produces free amines and hydroxyl
groups (Phua et al., 1987). Mammalian enzymes like human neutrophil elastase,
cholesterol esterase and porcine pancreatic elastase can also degrade PUs specifically
polyester based PU (Labrow et al., 1996; Santerre et al. 1993, 1994; Santerre and Labrow
1997).
2.5. Polymer degradation Analysis
A general problem during degradation analysis of plastics in the environment is to select
a suitable type of test and draw conclusions on the basis of the data acquired. Tests
applied in polymer degradation analysis are of three types:
1. Laboratory tests
2. Simulation tests
3. Field tests
2.5.1. Laboratory Tests
Laboratory tests have the best reproducibility. In Laboratory tests a consortium
for example from waste water or pure culture (single microbial strain) is inoculated in
24
defined medium supplemented with polymer as nutrient source. Such tests are performed
under conditions already optimized for previously screened microorganisms, due to
which polymer degradation rates observed are higher than it would have been in nature.
These tests are of value when polymer degradation mechanisms are to be studied
(Marten, 2000). Degradation tests can be performed in more reproducible and controlled
manner by using systems employing extracellular depolymerases only. Although this
method does not correlate biodegradation to microbial metabolism but it is useful in
performing systematic investigations like establishing relationship between chemical
structure of the polymer and its biodegradability (Tokiwa and Suzuki, 1977; Vikman et
al., 1995; Walter et al., 1995; Marten, 2000). Apart from reproducibility, minimization of
test duration and substantial reduction in the material required for the test are very
important when extended systematic investigations are performed, or when
biodegradation testing is used as a tool for the development of industrial materials.
Although degradation tests performed in compost or soil may require up to one year for
completion, and experiments involving specially screened microorganisms may take a
few weeks, enzymatic degradation can be done within days or even hours. New methods
are being tried by employing polymer nanoparticles for enzymatic degradation tests.
These methods are expected to reduce the duration of enzymatic degradation tests for
polyesters to a few seconds by increasing the surface area of polymer available to
enzymes (Gan et al., 1999; Welzel et al., 2002). Due to the principal disagreement
between the available analysis techniques in different tests and their applicability to
degradation studies under practical conditions, it is necessary to use combination of tests
in order to completely understand the biodegradation behavior of a plastic within a
certain environment.
The analytical methods used for monitoring the process of degradation depend on
the environment used for the test and aim of the study. Some of the various methods used
for the analysis of biodegradability are visual observations, measurement of changes in
weight (weight loss), molecular weight and mechanical properties, comparison of CO2
evolved in the presence and absence of polymer in the culture, formation of clear zone on
polymer dispersed agar plate, radiolabelling and analysis of biogas produced.
25
2.5.1.1. Visual Observations
Visible changes in polymer indicative of degradation like surface roughening,
color changes, appearance of cracks and/or hole and formation of biofilms can be
evaluated in almost all the tests. Although these changes provide any evidence of the
process of biodegradation in terms of microbial metabolism, but it can be used as a first
sign of degradative activity of microorganisms. To study the mechanism of degradation
more advanced techniques like scanning electron microscopy (SEM) or atomic force
microscopy (AFM) are used to obtain observations (Ikada, 1999). These techniques
provide important information like the appearance of crystalline spherolites after initial
degradation suggests that the amorphous fraction of the polymer is preferentially
degraded and the crystalline portions which are not readily degradable have been etched
out of the material. Such observations have been recorded from AFM micrographs of
PHB films degraded enzymatically to understand the mechanism of surface degradation
(Kikkawa et al., 2002).
2.5.1.2. Changes in Mechanical Properties and Molar Mass
Like visual observations, changes in mechanical properties of the polymer do not
prove that the polymer has been metabolized by microorganisms. However if these
changes are accompanied by even slight changes in molecular weight of the polymer
they are considered as evidence of degradation because the tensile strength of the
polymer is significantly affected by changes in molar mass of the polymer. Changes in
molar mass are considered as a direct evidence of degradation (Erlandsson et al., 1997).
Usually the pattern of enzymatic degradation differs from that of abiotic degradation.
Since enzymatic degradation is more restricted to the surface of exposed polymer and the
inner part of the material is not usually affected, the mechanical properties changes only
if there is substantial degradation. Abiotic degradation occurs throughout the bulk of
polymeric material including the inner part due to which it affects the mechanical
properties of the polymer significantly. Due to these reasons these methods are used in
cases where degradation is initiated by abiotic factors e.g., chemical hydrolysis of poly
(lactic acid) or oxidation of modified polyethylenes (Breslin, 1993; Tsuji and Suzuyoshi,
2002).
26
2.5.1.3. Weight Loss Measurements: Determination of Residual Polymer
The loss of mass of polymer films or bars is generally used in analysis of
degradation but again it is of no value for confirmation of biodegradability. Two main
problems with this type of analysis are; correct cleaning of the polymer sample and
recovery of extensively degraded material from culture medium. In case of excessive
degradation, recovery is facilitated by placing the samples in small nets. The degradation
characteristics of plastic samples can be quantified by sieving analysis of the matrix
surrounding the plastics in a better way. In case of polymer samples with very small
particle size weight loss can be measured by adequate extraction or separation of the
polymer from biomass, or oil or compost. Structural analysis of the residual polymeric
material combined with that of low molecular weight intermediates provide detailed
information about the process of degradation, especially when the test is conducted in
defined medium (Witt et al., 2001).
2.5.1.4. CO2 evolution/O2 Consumption
In aerobic environment microbes produce carbon dioxide by oxidation of carbon.
Therefore determination oxygen consumption (respirometric analysis) (Puchner et al.,
1995; Hoffmann et al., 1997) or carbon dioxide production (Sturm test) are indicators of
the degradation of polymers. These methods are often used in laboratory tests for
measuring biodegradation. The accuracy of sturm test depends on the presence of carbon
sources other than the polymer in the culture. For high accuracy, background respiration
is minimized by using low amount of other carbon sources and the test is conducted in
synthetic mineral salt medium. This type of test has long been used for evaluation of the
degradability of diverse substances and chemicals in water, and has now been adapted to
applications in non-water soluble polymeric materials. Mainly modified analytical
methods are used for the determination of CO2. Besides conventional methods like using
Ba (OH)2 for trapping of CO2 the concentrations of oxygen in air stream can also be
monitored by using paramagnetic oxygen detectors. Although systems designed for
continuous measurements in an automatic manner are advantageous, they have their own
disadvantages as well. Stability of the detector signals is necessary and in case of slow
degradation processes the detection system should be able to monitor small changes in
27
CO2 or O2 concentrations which increase the chances of systematic errors. Alternatives to
the above approach for example trapping CO2 in an alkaline solution of pH 11.5 may be
useful (Pagga et al., 2001). The problem of CO2 detection can also be resolved by using
closed non continuously aerated systems and monitor CO2 by using both sampling
technique and infrared-gas analyzer or titration sytem (Calmon et al., 2000; Muller,
1999). In similar tests degradation reactions are carried out in small closed bottles and
increase in CO2 or decrease in O2 is analyzed in the head space (Itavaara and Vikman,
1995; Solaro et al., 1998; Richterich et al., 1998). These closed bottles tests are simple
and not very sensitive to leakages, but the problem is low capacity of the bottles to hold
material and inoculums. Although these tests based on analysis of evolved CO2 were
originally designed to be used in aqueous systems, however they have been adapted for
use in solid matrices such as composts (Pagga et al., 1995). The standardized form of this
method is now called controlled composting test. Since instead of using biowaste as a
matrix mature compost is used in controlled composting test, it is not considered as a
simulation of composting process. Due to high contents of easily degradable carbon,
biowaste generates large amount of background CO2 and reduce accuracy of the test so
mature compost is used instead of biowaste. Detection of CO2 during polymer
degradability analysis in soil is more complicated than in normal compost the degradation
rate is slower and test durations are longer (up to 2 years). The amount of CO2 evolved is
also lower than that of background CO2 evolved from natural carbon sources already
present in soil. The problem of background CO2 evolution can be tackled by using inert,
porous matrix, free of CO2. This matrix is first impregnated with a synthetic medium
followed by inoculation with mixed microbial culture. This method can be used to
simulate compost conditions (degradation at 60 oC) but for soil conditions this has not
been optimized yet (Bellina et al., 1999; 2000).
28
2.5.1.5. Determination of Biogas
As compared to CO2 generation in aerobic environment, in case of anaerobic
microorganisms a mixture of methane and CO2 (known as biogas) is produced as an
extracellular product of anaerobic catabolism. The amount of biogas produced and its
composition depend on the chemical composition of material and can be calculated
theoretically by using Buswell equation (Buswell and Muller, 1952). Plastic degradation
in the absence of oxygen is usually monitored through biogas production (Gartiser et al.,
1998; Reischwitz et al., 1998; Abou-Zeid, 2001), and standards used for evaluation of
anaerobic biodegradation are also based on such measurements (ISO/DIS 15985, ASTM
D 5210, ASTM D 5511). Techniques like manometry or water displacement are used for
measurement of gas volume while the composition of the gas is analyzed by gas
chromatography (Budwill et al., 1996). Background evolution of biogas from anaerobic
sludges is also a problem like CO2 evolution and it makes the measurement of the amount
of biogas evolved from degradation of polymer difficult. For slow degradation processes
the accuracy of the test is particularly affected by this problem. Abou-Zeid (2001) diluted
anaerobic sludges with synthetic medium in an attempt to reduce the evolution of
background biogas.
2.5.1.6. Radio Labeling
Many problems related to background CO2 evolution can be resolved if the
polymer is radiolabelled at its carbon with 14
C. For example, 14
CO2 in very low
concentrations can be detected even in the presence of background CO2 evolved from
biowaste. Therefore radiolabelling is helpful while investigating the degradation of slow
degrading materials in an environment containing other carbon sources than plastics as
well (Albertsson, 1978; Tuomela et al., 2001). Difficulty in producing radiolabelled
plastics and risk involved with handling radioactive materials during experimental
procedures are the main disadvantages.
29
2.5.1.7. Clear-zone Formation
Clear-zone test is a very simple and semi-quantitative method. Polymer in the
form of very fine particles is dispersed within synthetic agar medium which is poured
into plates to prepare opaque agar plates. These plates when inoculated with
microorganisms, the appearance of a clear zone around the colony is indicative of the
ability of the organism to depolymerize the polymer. This technique is generally used for
screening of polymer degrading microorganisms but measurement of growth in the zone
can be used as a semi quantitative measure of degradation (Nishida and Tokiwa, 1993;
Abou-Zeid, 2001; Augusta et al., 1993).
2.5.3. Simulation Tests
For measurement of biodegradation different simulation tests can be used in
laboratory as an alternative to field degradation analysis. These tests are performed by
carrying out degradation in soil, compost or seawater in a controlled bioreactor inside
laboratory. These tests aim at providing environment similar to field test with
controllable external parameters like temperature, pH and humidity. These tests have
another advantage of availability of better analytical tools than those used for field tests
(For example for analysis of intermediates and residues, determination of carbon di oxide
evolution or oxygen consumption). Soil burial test, controlled composting test and test
simulating landfills or aqueous aquarium tests are examples of simulation tests (Pantke
and Seal, 1990; McCarlin et al, 1990; Smith et al., 1990; McCarthy et al., 1992; Puchner
et al., 1995; Pagga et al., 1995; Tosin et al., 1996; Degli-Innocenti et al., 1998; Ohtaki et
al., 1998; Tuominen et al., 2002). Sometimes, the tests can be accelerated by addition of
nutrients to the medium which increase microbial activity and speed up degradation.
2.5.3.1. Field Tests
Although field tests like placing plastic samples in a river or lake, soil burial or a
full scale composting analysis using biodegradable plastics ideally represent
biodegradation under conditions present in the natural environment. However there are
some grave drawbacks associated with these tests. One major problem is that conditions
like humidity, pH and temperature are difficult to control in the environment; secondly
30
the analytical tools for monitoring of the process of biodegradation in the field are very
limited. Most of the times the only possibility is to conclude on the basis of visual
observations of the polymer sample or to measure the extent of degradation by weight
loss analysis. Measurement of degradation based on the later approach is difficult when
the polymer breaks into pieces and recovery of all the pieces from soil, water or compost
is necessary. The complex and chemically undefined environment complicates the
analysis of intermediates and residues. Since physical disintegration of plastics alone is
not considered as biodegradation according to most of the definitions, field tests alone
can‘t be proof of polymer biodegradability (Touminen et al., 2002)
2.6. Degradation Products of Polyurethane
Few reports have been presented about the degradation products of polyurethane.
Some possible degradation products have been detected by Nakajima-Kambe et al., 1997.
Metabolites like adipic acid, diethylene glycol and trimethylolpropane are produced by
the degradation of polyester polyurethane (PU) by Comamonas acidovorans strain TB35.
Results revealed by GC-MS analysis showed that hydrolytic breakdown of ester bonds
caused the release of these metabolites from polyester segments of the PU. Alkaline
treatment of the culture broth detected a previously undetected product, identified as 2,4-
diaminotoluene by GC-MS analysis. It was considered that polyisocyanate segments of
the PU were degraded to the unknown metabolite which is water soluble compound. Two
different esterase enzymes were produced by strain TB-35, one which is secreted to the
extracellular liquid media and the other one which is cell surface-bound. Only the cell-
surface-bound esterase catalyzed the degradation of the polyester PU. Gautam et al.,
(2007a) reveals that esterases can catalyze the degradation of low-molecular weight
(MW) polyester PU. However, in case of high-MW PUs, it is yet to be determined
whether the direct hydrolysis of urethane bond is possible or degradation of high-MW
polymer molecules to low-MW compounds is a pre-condition for urethane linkage
vulnerability. A possible pathway of PU degradation by the cell-bound esterase of strain
TB-35 is shown schematically in Fig. 2.5 (Nakajima-Kambe at al., 1997).
31
Fig. 2.5: Proposed biodegradation pathway (adapted from Gautam et al. 2007b)
32
Materials and Methods
Fig 3.1: Work flow sheet for project ―Microbial degradation of polyester polyurethane‖
33
3.1. Isolation of Polyurethane Degrading Microorganisms
3.1.1. Materials
Polyurethane {poly[4,4‘-methylene-bis(phenyl isocyanate)-alt-1,4-butanediol/
poly(butylene adipate)]} was obtained in the form of pellets from Sigma-Aldrich, GmbH,
Germany. Tetrahydrofuran (THF) was purchased from Panreac Quimica, SA. The rest of
the reagents used were commercial products of the highest grade available.
3.1.2. Preparation of thin polyurethane films
1g of polyurethane pellets were taken in 100 ml of THF in 250 ml conical flask
and sonicated for 30 minutes for complete dissolution. This mixture was poured into 4
clean glass petri dishes in equal amounts to prepare thin films of polyurethane. The THF
was allowed to evaporate slowly by placing the covered petri dishes in desiccator for 48
hours.
3.1.3. Media used in degradation experiments and esterase activity analysis
For degradation experiments and esterase activity analysis in liquid medium,
mineral salt medium (MSM) [g/l: K2HPO4 0.5, KH2PO4 0.04, NaCl 0.1, CaCl2·2H2O
0.002,(NH4)2SO4 0.2, MgSO4·7H2O 0.02, FeSO4 0.001, pH adjusted to 7.0] was used.
Inoculums for such experiments were developed in MSM enriched with 0.5% w/v
peptones. For isolation experiments the bacterial inocula were developed in nutrient
broth, while nutrient agar and sabouraud dextrose agars were used for purification of
bacterial and fungal isolates. Media were sterilization by autoclaving at 121 oC at 15 PSI
pressure for 15 minutes. The pH of the medium was adjusted before to sterilization using
0.1 M NaOH or HCl to the desired value.
3.1.4. Soil sample for isolation of polyurethane degrading microorganisms
For collection of soil samples waste dumping area of Islamabad Pakistan was
chosen. Samples were collected in sterilized polyethylene bags. Large particles (pebbles,
wood pieces and plastic and paper pieces) from the soil sample were removed by sieving
34
through sieving machine (Retsch Model No. 56792), pore size 2 mm. Three potential PU
degrading microorganisms (two bacterial and one fungal) were isolated in three different
experiments from the same soil sample.
3.1.5. Isolation of PU degrading fungus
The soil was transferred to sterilized pots and placed in dark at room temperature.
Five polyurethane films of the same thickness were buried (4-6 inches deep) vertically in
the pots containing soil. At the time of burial 200 ml of MSM containing glucose was
added to the pot, so as to meet the nutrient requirements of the microorganisms. The
films were dug out, one each month, washed with autoclaved distilled water and placed
on Sabouraud Dextrose agar (SDA) plates aseptically. It was incubated at 30ºC for seven
days, then washed with sterilized distilled water and shifted to fresh SDA plates. The
process was repeated again and then it was shifted to MSM agar plates, incubated at 30oC
for 30 days. Fungal colonies grown on the surface of films were isolated and inoculated
on SDA plates. Pictures were taken and the polyurethane (PU) films were carefully
washed and stored for ATR-FTIR and SEM.
3.1.6. Isolation of polyurethane degrading bacterial strains
One gram of soil with uniform texture was incubated in nutrient broth for 24
hours at 37oC. This culture (10 ml) was inoculated in fresh MSM (90 ml), buffered at
pH7, containing PU (250 mg) films and incubated at 37oC in shaker incubator at 150 rpm
for one week. Treated PU pieces were shifted to fresh MSM along with 10 ml of the
culture after a week incubation and allowed to incubate in shaker incubator under the
same conditions for another week. This procedure was repeated four times. Viable cell
count was done each time prior to shifting of polyurethane films to fresh MSM. Isolates
were selected for further studies based on their ability to utilize polyurethane as a sole
carbon source.
35
3.2. Identification of the isolated microorganisms
3.2.1. Identification of fungal isolate
The fungal strain, isolated from soil, attached on the surface of polyurethane films
was purified on agar plates and was identified on the basis of colony morphology, color,
size and colony diameter. The fungal strain was maintained on malt extract agar (MEA).
The composition of MEA is [g/l: malt extracts 30.0; peptone 5.0; agar 15.0; pH 5.4±0.2].
The microscopic examination of the shape, color, size and structure of conidia, hyphae,
conidiophores and conidial head was done under the supervision of Dr. Kishwar Nazir,
Pir Mehar Ali Shah Arid Agriculture University, Rawalpindi, Pakistan.
3.2.2. Identification of bacterial isolates
The identification of bacterial strain was done through
Morphological characters
Biochemical characters and
16S rRNA gene sequence analysis
3.2.2.1. Morphological Characterization
Buchanan and Gibbons (1974) was followed for study of morphological
parameters of bacterial isolates. Parameters like colony morphology i.e. size, margins,
elevation and pigmentation, gram‘s staining and spore staining was studied. Bacterial
motility was also evaluated. For morphology studies the isolated colony of purified
organisms grown on nutrient agar plates under aseptic conditions was used. Gram‘s
staining was performed by preparing a smear of the selected colony. The smear was dried
and heat fixed. Heat fixed smear was treated with crystal violet for 60 Seconds before
washing with distilled water, then treated with gram‘s iodine for one minute. It was
washed with distilled water again and then decolorized with ethanol (95%) before
washing with distilled water again. Finally the slide was counter stained with safranin for
1 minute and then visualized under compound microscope.
36
For spore staining, smear of the isolated organism was prepared. After drying and
heat fixing, it was drenched in malachite green and heated for 3 minutes on hot plate. It
was washed with distilled water after cooling. In the end it was counter stained with
safranin for 60-90 seconds and then washed with distilled water. Stained cells were
observed under compound microscope.
Hanging drop technique was used to check bacterial motility. Petroleum jelly was
applied in the form of a ring along the margin of concavity of depression slide. A drop of
normal saline containing the cells of the isolate was placed in the center of a cover slip.
Depression slide was placed over the cover slip in a manner that concavity of the slide
covered the culture. The slide was turned upside down so that the drop adhering to the
cover slip started hanging into the depression. Then it was examined under compound
microscope for bacterial motility.
3.2.2.2. Biochemical characterization
The following biochemical tests were performed for the detection of identification
of bacterial isolate MZA-75 and MZA-85
Casein hydrolysis Citrate utilization
Startch hydrolysis Triple sugar iron
Lipid hydrolysis Methyl red
Gelatin liquefaction Vogus-Proskauer
Carbohydrate metabolism SIM (Indole & motility)
Glucose Oxidase
Fructose Catalase
Sucrose
Lactose
Raffinose
Mannose
Sorbitol
Urease test
37
Nitrate reduction
3.2.2.3. 16S rRNA gene sequence analysis
3.2.2.3.1. DNA Extraction
Extraction of DNA from bacterial cultures was done with the help of Wizard
genomic Kit (Promega, Madison, USA) as per manufacturer‘s instructions. Bacterial
culture (2 to 3 days old) was suspended in phosphate buffer solution (1 ml) in 1.5 ml
micro-centrifuge tube. This cell suspension was centrifuged at 13000-16000 X g for 2
minutes and supernatant was discarded. The cells were resuspended in 600 µl of lytic
solution by gentle pipetting. For cell lysis 5 minutes of incubation at 80 oC was carried
out. RNAse solution (3µl) was added after cooling the mixture to room temperature.
After 2-5 inversions, the cell lysate was incubated at 37 oC for approximately 1 hour.
After cooling the mixture to 25 oC protein precipitation buffer (200 µl) was added,
vortexed to mix it with cell lysate. Incubation of mixture on ice was carried out for 5
minutes before centrifugation at 13000 x g for 3 minutes. DNA was observed in the form
of visible mass in supernatant. After shifting of supernatant to separate tube DNA was
precipitated by addition of 600 µl of ethanol (70% v/v) followed by centrifugation at
13000 x g for 2 minutes. Ethanol was removed carefully and the DNA pellet was allowed
to air dry for 10 minutes. Finally TE buffer (pH 7) was added to the tube and mixed by
gentle tapping and stored at 4 oC. Concentration of DNA in the sample was determined
using Nanodrop 1000 (Thermo Scientific, Rockford, USA) as per standard procedure.
3.2.2.3.2. Amplification of 16S rRNA gene
Bacterial primers 27F‘ and 1494R‘ were used for PCR amplification of 16S rRNA gene.
20 μl PCR reaction mixture consisted of template DNA (1 μl), 10x PCR buffer (2 μl),
deoxynucleoside triphosphate (dNTP) mix (2 μl), forward and reverse primer (2 μl each),
Ex taq DNA polymerase (Takara Shuzo, Otsu) (0.5 μl) and distilled water (10.5 μl). At
first, template DNA was denatured by incubating the reaction mixture at 96oC for 4 min.
Then 35 amplification cycles were performed at 94oC for 45 sec, 55
oC for 60 sec, and 72
oC for 60 sec. Reaction mixture was further incubated for 7 minutes at 72
oC. DNA
fragments amplified were about 1,400 bps in the case of bacteria. A positive control
38
(E.coli genomic DNA) and a negative control (without template DNA) were included in
the PCR. The PCR product was purified from unincorporated PCR primers and dNTPs by
using Montage PCR Clean up kit (Millipore). The purified PCR products of
approximately 1,400 bps were sequenced by using 2 primers, 518F‘ and 800R‘.
Sequencing was performed by using Big Dye terminator cycle sequencing kit v.3.1
(Applied BioSystems, USA). Sequencing products were resolved on an Applied
Biosystems model 3730XL automated DNA sequencing system (Applied BioSystems,
USA) at the Macrogen, Inc., Seoul, Korea.
All sequences related to MZA-75 and MZA-85 was downloaded from NCBI
GenBank and aligned by Mafft v6.903B sequence alignment program. NJ analysis was
carried out using Maximum Composite-likelihood model. Bootstrap 1000 replicate
were used to the significance of generated tree on MEGA 4.
3.3. Polyurethane degradation assay
The capability of the isolated strains to degrade PU was analyzed by tracking
changes in surface morphology, modifications in chemical bonds as well as molecular
size of the PU with SEM, FTIR spectroscopy and GPC, respectively. Small pieces of PU
film (approx. 2x2cm) were sterilized by autoclaving at 121oC for 15 minutes in 250 ml
conical flask containing 100ml MSM. It was inoculated with the cells of MZA-75
fallowed by incubation at 37oC in shaker incubator at 100rpm for 28 days. Another
culture under the same conditions but without PU films was used as biotic control. For
abiotic control PU films were put in the same medium and were left un-inoculated. The
experiment was done in triplicate. The growth in test vessel was compared with that of
control by measuring the absorbance at 600nm. The experiment was terminated at the
end of fourth week and PU films were recovered for analysis through SEM, FT-IR and
GPC. CO2 evolution test was employed to confirm the ability of the isolated strains to
mineralize the polymer.
39
3.3.1. Scanning Electron Microscopy of PU film pieces
Changes in the surface features of PU films, as a result of microbial treatment,
was tracked by analysis of the recovered PU films through SEM (JSM 5910, Jeol, Japan).
Samples were thoroughly washed with sterile distilled water and then mounted on the
copper stubs with gold paint. To enhance conductivity of the samples, they were gold
coated in vacuum by evaporation. Electron micrographs of the treated PU samples were
compared with those of abiotic control.
3.3.2. Fourier Transform Infrared Spectroscopy (FT-IR) Analysis
FT-IR was employed for detection of changes in the functional groups in the
chemical structure of PU film as a result of incubation with the isolated microorganisms.
After pasting the polymer pieces on FTIR sample plate, a spectrum was taken in single at
500 to 4000 wave-numbers cm-1
for each sample and compared with that of abiotic
control.
3.3.3. Gel Permeation Chromatography (GPC) analysis
1% (w/v) solution of PU films in THF was prepared and analyzed by Agilent
PLGel 5µm 50A, 300x7.5mm GPC column. Flow rate was maintained at 1 ml/min.
Refractive index detector was used for detection. Calibration curve of polystyrene
standard was used for calculation of polydispersity and relative molecular weight.
3.3.4. CO2 evolution Test (sturm test)
CO2 evolved as a result of degradation of PU by strain MZA-75 and MZA-85 was
trapped and compared to the amount evolved in case of biotic control in Sturm test under
similar conditions. The culture bottle (Test bottle) containing 300 ml of MSM was added
approximately 500mg of PU film pieces. Both culture bottles i.e. test and control were
inoculated with the isolated microorganism (overnight grown culture) up to 0.07 Abs600,
No PU was present in the control bottle. Filter sterilized air was pretreated to remove
dissolved CO2 by bubbling it through a series of two bottles, each containing KOH (3M)
(pretreatment chamber). Magnetic stirrer was used for continuous stirring of the test and
control bottles throughout the duration of the test. The test was performed at room
40
Fig 3.2: Modified Sturm test. Arrows represent direction of air flow. Red bottles
containing 3M KOH constitute air pretreatment chamber (removes CO2 present in the
air). Black bottles are Control and Test having culture without and with PU respectively
and Blue bottles are CO2 absorption chamber having 1M KOH (traps CO2 produced as a
result of microbial metabolism).
41
temperature (30°C) for 4 weeks. After completion, the difference in CFU/ml(colony
forming units/ml) and the quantity of carbon dioxide produced in test was compared with
that of control. CO2 evolved as a result of PU degradation was trapped in the absorption
bottles consisting of two bottles each containing KOH (1M). Absorbed CO2 was
precipitated as barium carbonate by titrating aliquots from the absorption chamber
against barium chloride solution (0.1M). Amount of CO2 released was calculated
stoichimetrically from the weight of barium carbonate precipitates produced by addition
of BaCl2. Difference in the amount of precipitates in the test and control was determined
(Muller et al. 1992).
3.4. Esterase activity assay
Strain MZA-75 and MZA-85 were inoculated in 9 ml of liquid MSM with 0.5%
peptone and incubated at 37oC in shaker incubator for 16 hrs. The culture was centrifuged
at 8000 rpm for 10 minutes at 4oC and cell pellet was separated from supernatant. The
pellet was suspended in Tris HCl buffer (pH7) after washing with the same twice. The
method of kanwar et al. (2005) was used for determination of esterase activity in both
supernatant and cells, using pNPA as substrate for esterase. The details of assay are as
follows:
Stock solution of pNPA was prepared in isopropanol. 50 µl of the sample was
added to Tris buffer (0.05M, pH 7) to make the final volume of 3 ml. The reaction
mixture was incubated at 37oC for 20 minutes in a water bath. The reaction was stopped
by adding 1ml of chilled acetone:ethanol mixture (1:1, kept at -20oC overnight). Control
containing no enzyme sample was also incubated with each assay. The absorbance at 410
nm (A410) was measured for both Test and Control. The A410 for control was subtracted
from that of test. The concentration of p-nitrophenol produced was determined from
previously prepared standard curve of p-nitrophenol. All the assays were performed in
quadruplicates (Two biological replicates and two technical replicates for each biological
replicate) and mean values were recorded.
Unit of activity:
The unit of activity is defined as the amount of enzyme that hydrolyzes 1 µM
substrate in 1 minute.
42
Standard curve for p-nitrophenol (pNP):
p-Nitrophenol solutions of different concentrations i.e.; 1mM, 2mM, 3mM up to
9mM were used and absorbance was checked for each concentration on UV-Visible
spectrophotometer. The absorbance results were plotted against respective concentrations
to obtain p-nitro phenol concentration versus absorbance standard curve.
To investigate about the cell associated esterases, 0.2% of N,N-Bis(3-D-
gluconamidopropyl) deoxycholamide (deoxy-BIGCHAP, Dojin Chem. Co., Japan), a
surfactant, was added to the culture broth, and was mixed for 1hr by shaking and then
centrifuged. The cell free supernatant was assayed for esterase activity.
Protein estimation
For the estimation of proteins the method of Lowry et al., 1951 was used and
BSA (bovine serum albumin) was taken as a standard. Four solutions were prepared.
Solution A
Ingredients g/100ml
Na2CO3 1.0
NaOH(0.1) 0.4
NaK tartarate 1.0
Distilled water 100ml
Na2CO3 was dissolved in distilled water then NaK tartarate and finally NaOH.
Solution B
CuSO4.5H2O 0.5g
Distilled water 100ml
Solution C
Solution A 50%
Solution B 50%
Solution C was freshly prepared.
Solution D
Folin‘s phenol in ratio of 1:1 with distilled water.
43
Fig 3.3: Plot of Abs410 values against respective concentrations of p-Nitrophenol, used for
determination of milli Moles of p-Nitrophenol produced for determination of specific
esterase activity.
Fig 3.4: Standard curve of BSA used for determination of concentration of total proteins.
44
Procedure
Different dilutions of BSA (bovine serum albumin) were made from the stock
solution of concentration 1μg/ml. the volume was made up to 1ml by adding distilled
water except blank. 1ml solution C was added, shaken kept for 10min. Folin phenol was
diluted to 1:1 and was added, 0.1ml in each tube. It was shaken and kept for 30 min at
room temperature. Samples were treated in the same way and using spectrophotometer,
A650 was taken. BSA standard curve was used for calculation of unknown protein
quantities.
3.4.1. Optimization of polyurethane esterase production from Bacillus subtilis MZA-
75
Production of cell bound polyurethane esterase from Bacillus subtilis MZA-75
was optimized. Effect of different physical and chemical parameters such temperature,
pH, incubation time and nitrogen source was studied on the production of polyurethane
esterase. 300 ml of MSM was taken in 500 ml Erlenmeyer flasks and inoculated by cells
of MZA-75 (freshly separated from overnight culture of 8ml MSM with 0.5% peptone).
The flasks were incubated at 37oC and 150 rpm for 4 weeks. Samples were drawn weekly
and esterase assay was performed for cell bound esterases.
3.4.1.1. Effect of temperature
The production of polyurethane esterase was carried out at 30, 37, 40, 45 and
50°C, at 150 rpm and pH 7.0.
3.4.1.2. Effect of pH
Effect of pH on the production of polyurethane esterase was determined by
carrying out the growth and enzyme activity calculations at pH 5.0, 6.0, 7.0, 8.0 and 9.0
at 37°C and 150 rpm.
3.4.1.3. Effect of time of incubation
Optimum period of incubation for the production of polyurethane esterase from
was determined by polyurethane esterase assay on weekly basis.
3.4.1.4. Effect of nitrogen source
45
Effect of nitrogen source on polyurethane esterase production was studied in the
presence and absence of yeast extract (1% w/v solution) at pH, temperature, agitation
speed as 7.0, 37 oC and 150 rpm.
3.5. Analysis of Ester Hydrolysis Products by GC-MS
3.5.1. Growth of the isolated organisms with PU as sole carbon source
300 ml of MSM was dispensed in 1 liter of Erlenmeyer flask. 500mg of
polyurethane film pieces were put in the flask. The flask was tightly plugged and
sterilized by autoclaving at 121oC for 15 minutes. After autoclaving, the flask was
inoculated with the isolated bacterial strain (performed for both MZA-75 and MZA-85
separately). Both biotic and abiotic controls were set up in the similar fashion. The
experiment was run in triplicate. 50 ml of sample was taken at zero time and then all
three sets were shifted to shaker incubator at 37oC and 100 rpm. All sets were sampled
after every seven days.
3.5.2. Extraction of samples from polyurethane culture medium for metabolites
Samples were centrifuged at 8000 rpm for 10 minutes at 4oC. The supernatant was
preserved at -20oC while the pellet was discarded. The samples were melted and acidified
to pH 2. They were allowed to stand for 2 hrs and then extracted with three volumes (40,
20, 20 ml) of ethyl acetate. The extracts were pooled and dried by passing through
anhydrous sodium sulfate, concentrated by rotary evaporation, and reduced further under
a stream of N2 to a volume of 50 μl.
3.5.3. Analysis of the extracts on GC-MS
The extracts were derivatized with N,O-bis (trimethylsilyl) trifluoro acetamide
(BSTFA) (Pierce Chemical Co., Rockford, IL) prior to analyses of the resulting
compounds on an Agilent 6890 model gas chromatograph (GC) coupled with an Agilent
model 5973 mass spectrometer (MS). Derivatized components were separated on HP-
5ms capillary column (30mx0.25mm inner diameter x 0.25micro meter film, Agilent)
using temperature programming as follows: The initial temperature of 80oC was kept for
5 minutes and then ramped it up at the rate of 10oC per minute up to a maximum of
46
230oC. The identification of methyl esters of butanediol and adipic acid was done by
comparison of the GC-MS profiles to authentic standards purchased from Sigma-Aldrich
(St Louis, MO) or the National Institute of Standards and Technology (NIST) Mass
Spectral Library, version 2.0a.
3.6. Utilization of 1,4-butanediol and adipic acid as carbon source
5mM concentration of 1, 4-butanediol and adipic acid was taken in 5ml of MSM
in separate test tubes. Tubes were inoculated with strain MZA-75 and incubated at 37oC.
The experiment was set up in triplicate. Abs600 was recorded at 24hrs interval to
demonstrate the ability of strain MZA-75 to utilize 1, 4-butanediol and adipic acid as a
source of carbon and energy.
3.7. Biofilm quantification by crystal violet staining/SEM observation
The ability of P.aeruginosa MZA-85 to form biofilms on surface of PU films was
quantified by the method of O‘Toole et al. (1998). P.aeruginosa MZA-85 was grown
overnight in MSM with 0.5% peptone at 37°C, was centrifuged, pellet was washed with
sterile Trish HCl (pH 7) and suspended in the same. 1ml of this culture (OD600=0.973)
was inoculated in 49 ml sterile MSM (containing PU films) in 100 ml conical flask and
incubated at 37 oC. PU film (2x2cm) was recovered after 4 days interval. The recovered
PU films were gently washed with sterile Tris HCl (pH7) twice, dried and stained with
crystal violet for 15 minutes. After staining they were rinsed gently with Tris HCl three
times. Bound crystal violet was solubilized in 5ml ethanol-acetone (80:20 vol/vol).
Absorbance750 was measured using UV-visible spectrophotometer.
The recovered PU films after the end of experiment were also observed by SEM
to visualize the biofilm.
3.8. Production and purification of cell bound polyurethane esterase
Strain MZA-75 was inoculated in production medium (MSM supplemented with
1% wt/vol PU) and incubated it for 3 weeks. The culture was centrifuged at 10,000xg for
15 min at 4oC. Supernatant was discarded while 0.2% of N, N-bis(3-D-
gluconamidopropyl) deoxycholamide (deoxy-BIGCHAP, Dojin Chem. Co., Japan), a
47
surfactant, was added to the pellet suspended in Tris HCl buffer pH7, and was mixed for
1 h by shaking and then centrifuged. (NH4)2SO4 was added to the supernatant to provide
50% saturation. The precipitated proteins were collected by centrifugation (10,000xg, 15
min) and suspended in a 100 mM phosphate buffer at pH 7.0. Dialysis was performed in
a 100 mM phosphate buffer at pH 7.0 to remove the ammonium sulfate from the protein
extract. Crude protein from cell-free filtrate of Bacillus subtilis strain MZA-75 grown in
1-liter production medium was purified to 100% purity by gel filtration chromatography
using a Sephadex G-75 column (GE Healthcare Life Sciences). All of the fractions were
tested for polyurethanase activity by esterase assay. Rough estimation of protein
concentration was done by taking absorbance at 280 nm in each fraction during
purification.
3.8.1. Electrophoresis
Sodium dodecyl sulfate (SDS)-PAGE was performed as described by Laemmli
(1970) with a 15% (wt/v) polyacrylamide resolving gel. Proteins were denatured by the
addition of 2-mercaptoethanol and heating to 100oC for 5 min. Proteins were visualized
by silver staining.
3.8.2. Protein concentration determination
Protein concentrations were measured by the method of Lowry et al., 1951
(method described in section 3.4).
3.8.3. Substrate specificity of purified cell bound esterase from MZA-75
The effect of acyl chain length on esterolytic activity of the purified PU esterase
was checked by using various p-NP acyl esters such as, acetate (C2), butyrate (C4),
caproate (C6), caprylate (C8), caprate (C10), palmitate (C16) and stearate (C18) according to
the method of Eggert et al. (2000). The reaction was performed with 100 mM potassium
phosphate buffer (pH 7.0) at 37°C.
48
3.8.4. Degradation activity of purified esterase against PU films
PU film were treated with 70% ethanol under UV lamp for 10 min for
sterilization. A piece of PU film (50 mg) was added to a test tube containing 10 ml of 100
mM phosphate buffer. After addition of PU esterase, it was incubated in a reciprocal
shaker incubator at 30°C (125 oscillations/min). Un-inoculated medium containing
similar PU film was taken as a negative control. Plastic degradation was monitored by
measuring the weight of the film before and after incubation. Experiment was performed
in triplicate.
3.9. Statistical analysis of results
The experiments were done in triplicate. Student‘s t-test and two way anova
analysis were done Graph-pad prism version 5.01. P value of 0.05 was set as a level of
significance. The data are expressed as mean ± standard errors.
49
RESULTS
4.1. Isolation of Polyurethane Degrading Microorganisms
4.1.1. Isolation of polyurethane degrading fungus
The soil buried films were recovered and plated on Sabouraud dextrose agar
(SDA) plates. PU films with fungal growth were washed with sterile normal saline and
shifted to MSM agar plates. Growth of fungi accompanied by substantial physical
changes was considered as an indication of the capability of the isolated fungal strain to
utilize PU as substrate (Fig. 4.1 & 4.2).
4.1.2. Isolation of PU degrading bacterial strains
Two bacterial strains MZA-75 and MZA-85 were selected in two different
experiments for their ability to survive and grow in the presence of PU as a sole carbon
source after screening by enrichment. After purification the ability of both MZA-75 and
MZA-85 to grow on PU was confirmed by taking absorbance at 600 nm for each one of
them, using PU as a source of carbon.
4.1.3. Growth of bacterial isolates on PU as carbon source
Abs600 values (corresponding to number of cells) in test culture of MZA-75
(containing PU) rose steadily from 0.05 (P<0.05) at the time of inoculation to a
maximum of 0.215 (P<0.001) on day 12, while no significant change was observed in the
biotic control (culture without PU) (Fig. 4.3). Strain MZA-85 exhibited much slower
growth and Abs600 value reached to the maximum i.e 0.204 (P<0.001) on 22nd
day of
inoculation. No growth was observed in the biotic control of MZA-85 (Fig. 4.4).
50
Fig. 4.1: Polyurethane film with fungus growing on MSM-agar plate. Photograph taken
after seven days of incubation at 30oC.
Fig. 4.2: PU degradation by fungi. PU film treated with Aspergillus tubingensis for 4
weeks (B) shows signs of degradation as compared to untreated PU film (A).
51
Fig. 4.3: Growth of MZA-75 in MSM supplemented with PU as a sole source of carbon.
Test: culture with PU and Control: culture without PU or any other carbon source.
52
Fig. 4.4: Growth of MZA-85 in the presence of PU as sole source of carbon in liquid
MSM. Test represents growth of MZA-85 in the presence of PU, while control represents
the growth in absence of any carbon source.
53
4.2. Identification of Polyurethane Degrading Strains
4.2.1. Identification of PU degrading fungus
The isolated fungal strain was identified on the basis of colony morphology,
structure of conidiophores and conidial head and pigmentation characters as Aspergillus
tubingensis. The isolated fungi, when inoculated on sabouraud dextrose agar grew into
white mycelia initially, which darkened gradually to black colour with the development
of spores on mycelia (Fig. 4.5).
4.2.2. Identification of polyurethane degrading bacterial strains
4.2.2.1. Biochemical analysis of MZA-75 and MZA-85
Both the strains MZA-75 and MZA-85 were identified through standard
morphological and biochemical tests, the results are presented in Table 4.1.
4.2.2.2. 16S rRNA gene sequence analysis of MZA-75and MZA-85
16S rRNA gene sequence of 1.481kb nucleotide length was sequenced from strain
MZA-75. The sequence was aligned with reference sequences obtained from NCBI
GeneBank. The phylogenetic analysis of the 16S rRNA sequence showed that the strain
MZA-75 belonged to genus Bacillus and have similarities (99% max identity) with
several strains of Bacillus subtilis on NCBI, but on the basis of maximum score the
closest related organism is Bacillus subtilis JBE0016 (FJ982665)(Fig. 4.6). The
nucleotide sequence reported here can be obtained from NCBI nucleotide sequence
database under accession number HM101166. In case of MZA-85 the 1.463 kb
nucleotides of 16S rRNA obtained after amplification, when aligned with reference
strains on NCBI GeneBank, revealed that MZA-85 belonged to genus Pseudomonas and
have 100% similarity with Pseudomonas aeruginosa IL1-(DQ989211)(Fig. 4.7) type
strain. The nucleotide sequence reported here for MZA-85 can be obtained from NCBI
nucleotide sequence database under accession number HQ023428.
54
Fig. 4.5: Growth and colony morphology of the isolated fungal strain on sabouraud
dextrose agar.
55
Table 4.1: Identification of polyurethane degrading bacterial strains
Characteristics MZA-75 MZA-85
Colony characteristics
Shape Round Round
Size Large Large
Colour White/pale Pale
Surface Dull, granular,Wrinkled Convex
Margin Irregular Undulate
Morphology
Straight rod + +
Cocci - -
Gram stain + -
Cell arrangement Short chains, single Short chains, single
Spore C C
Motility + +
Granulation + +
Biochemical tests
Casein hydrolysis + +
Startch hydrolysis + +
Lipid hydrolysis + +
Gelatin liquefaction + +
Carbohydrate
Fermentation
Glucose A/- -/-
Fructose A/- -/-
Sucrose A/- -/-
Lactose A/- -/-
Raffinose A/- -/-
Mannose A/- -/-
Sorbitol A/- -/-
Urease - -
Nitrate Reduction + +
Citrate + +
TSI Y/Y -
MR - -
VP + -
SIM + +
Oxidase + +
Catalase + +
Identified
Microorganisms
Bacillus sp. Pseudomonas sp.
C: center; A: acid; Y: yellow; R: red
56
Fig. 4.6: Pylogenetic tree of MZA-75 showing that it has maximum resemblance with
Bacillus subtilis.
57
Fig. 4.7: Phylogenetic tree of MZA-85 showing that it has maximum resemblance with
Pseudomonas aeruginosa IL1.
58
4.3. Analysis of polyurethane degradation
4.3.1. Polyurethane degradation by Aspergillus tubingensis
Polyurethane was treated with Aspergillus tubingensis in both solid media
(Nutrient Agar) and MSM Agar and degradation was analyzed by SEM and FTIR.
4.3.1.1. SEM analysis
Polyurethane film treated with A. tubingensis on MSM agar for one month,
when analyzed through SEM, adherence of fungal mycelia on the surface of film was
observed. The damaged surface of the treated PU film could also be observed in the
SEM. In some areas fungal mycelia traversing the cracks on the film were also noted
(Fig. 4.8). PU films recovered from liquid MSM did not show any mycelial adherence,
nor was there any convincing signs of degradation on the surface of exposed PU films
recovered from MSM broth inoculated with spore suspension. No growth was observed
when A. tubingensis was inoculated as spore suspension in MSM supplemented with PU.
4.3.1.2. FTIR analysis
Attenuated total reflectance-FTIR analysis of the fungus treated PU film on
MSM agar plate shows few changes in the spectra as compared to control. The peak in
sample at wavelength 3271.9 cm-1
was at wavelength 3325.4 cm-1
in control, moreover
the sample peak was broader than control. Another peak at wavelength 2919.3 cm-1
in
sample was present at wavelength 2954.2 cm-1
in control. The disappearance of sharp
peak at wavelength 1725.8 cm-1
was observed, which was present in control. The
appearance of characteristic peak at 1632.0 cm-1
was present in sample spectrum which
was absent in control. No changes in the functional groups of the PU films treated with
Aspergillus tubingensis in liquid MSM was observed (Fig. 4.9).
59
Fig. 4.8: SEM of PU film treated with Aspergillus tubingensis for 1 month on MSM-agar.
Mycelial mass adhering to the surface of the film can be seen (B). Damage caused by
fungal mycelia can be seen (C). No mycelial mass seen on untreated control (A).
60
Fig. 4.9: The peak at 1696 cm-1
representing urethane linkage in the control PU film (A)
is absent in the treated PU sample (B). Peaks representing amine linkage (C-N) i.e. at
1164.4 cm-1
and 1136.3 cm-1
are absent in the treated PU film. The area 600-700 cm-1
representing C-H deformation, No peaks was observed in this region.
61
4.3.1.3. CO2 evolution test (sturm test)
No significant difference in the amount of CO2 released in the test (culture
with PU) and control (culture without PU) was observed. No fungal adherence was seen
to the surface of PU. Since spore suspension was used for inoculation in the culture, they
did not grow during the test period of one month.
4.3.2. Polyurethane degradation by bacterial strains MZA-75 and MZA-85
Polyurethane films treated with the isolated bacterial strains MZA-75 and MZA-
85 were analyzed for changes in the surface morphology through SEM, chemistry
through FTIR and molecular weight through Gel Permeation Chromatography (GPC).
4.3.2.1. SEM analysis
After incubating polyurethane film pieces with the culture of P. aeruginosa MZA-
85 for one month, alteration in the physical characters of the treated PU was analyzed.
The treated PU films gradually changed their color from being transparent to brown.
SEM of PU film pieces was performed to confirm degradative changes in the exposed
films. Figure 4.10B shows strain MZA-85 cells adhering to the surface of PU film with
cracks radiating from the point of adherence. None of these changes can be seen in case
of control (Fig. 4.10A). MZA-85 cell attachment and accompanying surface changes
indicates towards microbial degradation of the exposed polyurethane film. No adherence
of MZA-75 cells was seen with the surface of PU films treated for one month, however
widespread cracks appeared on the surface of treated PU films, which were not present in
the untreaed control (Fig. 4.11).
4.3.2.2. FTIR analysis
ATR-FTIR spectrum of the PU film treated with MZA-75 and MZA-85 revealed
similar changes in the different functionalities. The peak at 1725 cm-1
representing ester
linkage appears in the untreated PU film sample and was found absent in the treated one
showing ester hydrolysis. The carboxylic acid peak in the treated PU sample was
overlapped by amide carboxyl peak at 1685 cm-1
. NH peak in the untreated sample is
present at 3325 cm-1
whereas in treated sample the peak at 3325 cm-1
broadened due to
62
free hydroxyl group. C-O peak for ester group is present at 1135 cm-1
whereas in the
treated sample this is absent indicating toward ester hydrolysis and production of free
hydroxyl groups as a result of treatment with MZA-85 (Fig. 4.12 & 4.13).
4.3.2.3. GPC analysis
GPC chromatogram of the PU film treated with Bacillus subtilis MZA-75 and
Pseudomonas aeruginosa MZA-85, in liquid MSM for 4 weeks reveals changes in
polydispersity index, weight average molecular weight and number average molecular
weight as compared to that of untreated control. The polydispersity index increased from
1.369 to 1.679 in case of strain MZA-75 as compared to untreated control. The weight
average molecular weight (Mw) decreased from 48762 in case of untreated control, to
48152 after treatment. The number average molecular weight (Mn) was 35616 before
treatment and dropped to 28667 after treatment. These results show that microbial
treatment resulted in the cleavage of long chain polyester polyurethane molecules to
fragments of relatively smaller molecular weight (Fig. 4.14).
Incase of MZA-85 increase in polydispersity index i.e. from 1.369 to 1.760 was
observed in the GPC chromatogram of treated PU film as compared to untreated control.
The weight average molecular weight (Mw) of the untreated control was 48762, which
decreased to 42589 after treatment. The number average molecular weight (Mn) was
35616 before treatment and dropped to 24198 after treatment. Melting point of the
untreated control which was 50161 dropped to 49684 in the treated PU samples. These
changes show that microbial treatment resulted in the cleavage of long chain polyester
polyurethane molecules to fragments of relatively smaller molecular weight (Fig. 4.15).
4.3.2.4. CO2 evolution test
The results indicate high amount of CO2 released i.e. 7.62 gram/liter in MZA-75 culture
supplemented with PU film pieces (Test) as compared to the culture (3.5 gram/liter)
without any carbon source (control). MZA-75 also showed better growth in the presence
of PU and CFU/ml increased from 10x106 to 6.6x10
11, which is higher than the rise
observed in control i.e. 11x106 to 2.6x10
9 CFU/ml. In case of MZA-85 similar results
63
were recorded for both CFU/ml and CO2 evolution. The CFU/ml increased from 6.3x105
to 8.31010
in Test while in case of control it increased from 8x105 to 4x10
7. The amount
of CO2 evolved from Test is 9.54 gram/liter as compared to 5 gram/liter in case of
control.
64
Fig. 4.10: Cells of Pseudomonas aeruginosa MZA-85 adhering to the surface of treated
PU films with cracks radiating from the point of adherence (B). No such changes could
be seen in the untreated (abiotic) control (A).
65
Fig. 4.11: SEM of PU films treated with Bacillus subtilis MZA-75, Small hair like
modifications can be seen on the surface (B) higher magnification reveals these as cracks
on the surface of PU film (C). No such cracks can be seen on the surface of untreated
(Abiotic) control (A).
66
Fig 4.12: FTIR spectrum of PU film treated with Bacillus subtilis MZA-75, reveals
disappearance of peak at 1725 cm-1
(B) which is present in the untreated (abiotic) control
(A).Another peak at 1368.6 increased in intensity.
67
Fig 4.13: FT-IR spectrum of MZA-85 treated PU film (B) demonstrating polyester
portion of the PU as target for microbial degradation. The peak at 1725cm-1
which is
present in untreated control (A) disappeared in the treated sample. The intensity of
another peak at 1368.6 increased.
68
Fig 4.14: Gel permeation chromatography shows increase in the polydispersity index of
PU film treated with Bacillus subtilis MZA-75 for 4 weeks (B). Changes in number
average molecular weight (Mn) and weight average molecular weight (Mw) can also be
seen when compared to untreated to control (A).
69
Fig 4.15: Gel permeation chromatography shows increase in the polydispersity index of
PU film treated with Pseudomonas aeruginosa MZA-85 for 4 weeks (B). Changes in
number average molecular weight (Mn) and weight average molecular weight (Mw) can
also be seen when compared to untreated control (A).
70
Table 4.2: Sturm test for detection CO2 produced as a result of degradation of PU by
MZA-75.
CFU/ml
at day 1
St. dev. CFU/ml after
4 weeks
St. dev. CO2
evolved
St. dev.
Control 1.1x107 1 2.6 x 10
9
1.52 3.5g/litre 0.5
Test 1.0x107 1.73 6.6 x 10
11 1.52 7.62g/litre 0.56
Table 4.3: Sturm test for detection CO2 produced as a result of degradation of PU by
MZA-85.
CFU/ml*
at day 1
St. dev* CFU/ml after
4 weeks
St. dev CO2
evolved
St. dev.
Control 8x105 1.732 4 x 10
7 1 5.0g/litre 0.453
Test 6 x 105 1 8.3 x 10
10 0.577 9.54g/litre 0.713
71
4.4. Analysis of degradation products by GC-MS
4.4.1. Detection of ester hydrolysis products
Polyurethane films were exposed to MZA-75 and MZA-85 in liquid MSM. The
MSM culture was sampled on weekly basis, extracted with ethyl acetate, concentrated
under continuous stream of nitrogen and derivatized with N,O-bis (trimethylsilyl)
trifluoro acetamide (BSTFA). The analysis of these derivatized samples showed the
presence of methyl esters of 1,4-butanediol and adipic acid in both the cultures. The
analysis of cell free supernatant of strain MZA-75 by GC-MS demonstrated two new
peaks at retention times 13.1 and 18.19 min, which correspond to 1,4-butanediol and
adipic acid respectively, after comparison with the standard chromatograms. Similar
results were obtained with MZA-85 and both 1,4-butanediol and adipic acid were
observed in the cell free supernatant of MZA-85 supplemented with PU. Neither any
metabolite related to the basic PU structure nor the above mentioned metabolites were
detected in the biotic and abiotic controls. Figure 4.16 and 4.17 show GC-MS based
detection and identification of monomers released as a result of esterolytic breakdown of
PU by the action of MZA-75 and MZA-85 respectively.
4.4.2. Growth of MZA-75 and MZA-85 on 1,4-butanediol and adipic acid
Based on Abs600 data taken after every 24 hours, strain MZA-75 and MZA-85
were found capable of using both 1,4-butandiol and adipic acid for their growth, as
indicated by a gradual increase in optical density (OD) at 37oC within 48 hrs. Absorbance
600 for P.aeruginosa MZA-85 culture in MSM with Adipic acid as a sole source of
carbon increased from 0.053 to 0.64, in case of 1, 4-butanediol the absorbance increased
from 0.06 to 0.4 after 48 hours of incubation whereas no growth was observed in case of
control with no carbon source (Fig. 4.18). Strain MZA-75 also showed ability to utilize
the ester hydrolysis products of polyester PU by growing significantly faster in the
presence of 1,4-butandiol and adipic acid, as indicated by a gradual increase in optical
density (OD600) at 37oC within 48 hrs. Abs600 increased from 0.062 to 0.480 in 48 hours
in the presence of 1,4-butanediol, while in case of adipic acid the Abs600 values increased
72
from 0.062 to 0.331 in 48 hours. No growth was observed in case of control with no
carbon source (Fig. 4.19).
4.5. Analysis of biofilm formation by MZA-85 on the surface of
polyurethane
Crystal violet staining of the PU films treated with MZA-85, showed that MZA-
85 forms biofilm strongly adhered to the surface of films. Treated PU film pieces were
also visualized through SEM for biofilms and compared to untreated PU films (control)
(Fig. 4.20). Increase in biofilm quantity was recorded through measurement of Abs750
after crystal violet staining (Fig. 4.21). Maximum Absorbance was obtained after 48
hours in case of test i.e.
73
Fig 4.16: GC chromatogram overlay of ethyl acetate extract of Bacillus subtilis MZA-75
culture with PU (Red), culture without PU i.e biotic control (Blue), MSM supplemented
with PU without MZA-75 culture i.e. abiotic control (Black). 1, 4-butanediol and adipic
acid peaks at retention time 13.10 minutes and 18.2 minutes respectively can be seen only
in culture with PU (Red line). M/Z of 1,4-butanediol standard (B) M/Z of 1,4-butanediol
extracted (C) M/Z of adipic acid standard (D) M/Z of adipic acid extracted (E).
74
Fig 4.17: GC chromatogram overlay of ethyl acetate extract of Pseudomonas aeruginosa
MZA-85 culture with PU (Black), culture without PU i.e biotic control (green), MSM
supplemented with PU without MZA-85 culture i.e. abiotic control (Red). 1, 4-butanediol
and adipic acid peaks at retention time 13.28 minutes and 18.5 minutes respectively can
be seen only in culture with PU (Black line). M/Z of 1,4-butanediol standard (B) M/Z of
1,4-butanediol extracted (C) M/Z of adipic acid standard (D) M/Z of adipic acid extracted
(E).
75
Fig 4.18: Gradual rise in growth of Pseudomonas aeruginosa MZA-85 utilizing 1,4-
butanediol (circle), adipic acid (square) and control i.e. MSM without any carbon source
(triangle).
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0 10 20 30 40 50 60
Ab
sorb
ance
60
0n
m
Time (Hours)
Butandiol Adipic acid MSM
76
Fig 4.19: Gradual rise in growth of Bacillus subtilis MZA-75 utilizing 1,4-butanediol
(circle), adipic acid (square) and control i.e. MSM without any carbon source (triangle).
A
0
0.1
0.2
0.3
0.4
0.5
0.6
0 10 20 30 40 50 60
Ab
sorb
ance
60
0n
m
Time (Hours)
Butanediol Adipic acid Control
77
Fig 4.20: SEM of PU film treated with P.aeruginosa MZA-85 for one week (B) showing
the presense of biofilm on the surface, while no biofilm is visible on the surface of
untreated control (A).
B
78
Fig: 4.21: Quantification of Biofilm made by MZA-85 on PU film by crystal violet
staining. Abs750 was taken as measure of biofilm quantity.
79
4.6. Polyester polyurethane degrading enzymes (esterase) from bacterial
strains
4.6.1. Esterase assays for MZA-85
No esterase activity was detected in the cell free supernatant, however an increase
in cell associated esterase activity was found during day 1 to day 12, i.e. 0.5023-2.22
mM/min/mg (P<0.01), then gradually decreased to 1.135 mM/min/mg (P<0.05) till day
24, while no marked increase in activity was found in the absence of PU (biotic control)
(Fig. 4.22). The cell associated enzyme was found to be cell membrane bound because
enzyme activity was recorded in the cell free supernatant when they were treated with
0.2% deoxy BIGGCHAP for two hours. This PU associated esterase induction in MZA-
85 was attributed to the isolate‘s ability to hydrolyze ester linkage present in this
polymer.
4.6.2. Esterase assays for MZA-75
Both supernatant and cell suspension were found to be active when tested for
microbial esterases. Extracellular esterase activity increased steadily from 0.182
mM/Min/mg on day one to 0.329 mM/Min/mg (P<0.001) on day 16 and then reached its
maximum i.e. 0.494 mM/Min/mg (P<0.001) on day 24 when PU was used as carbon
source, while no increase in esterase activity was observed without any carbon source
(Fig. 4.23). Cell associated esterase activity dropped from 0.519 mM/Min/mg (P<0.05)
on day one to 0.336 mM/Min/mg (P<0.001) on day 4 and then reached its maximum i.e.
1.210mM/Min/mg (P<0.001) on day 20 in the presence PU as source of carbon and
energy. While no rise in esterase activity was observed in the absence of PU (Fig. 4.24).
4.6.3. Optimization of culture conditions for esterase production from MZA-75
Maximum esterase activity was observed at 37oC on day 21 (Fig. 4.25). Optimum
pH was found to be 7, but strain MZA-75 also demonstrated some esterase activity at
slightly acidic pH, i.e. pH 6 and 5, but little activity was observed when pH was increased
above neutral (Fig. 4.26). Enzyme production was observed to be influenced positively
by the addition of yeast extract in the medium and maximum enzyme activity in the
80
absence of yeast extract that is 0.593 (P<0.05) on day 21 is significantly lower than
esterase activity in the presence of yeast extract which was 0.761 mM/min/mg (P<0.01)
(Fig. 4.27). Maximum enzyme activity (0.543 mM/min/mg) (P<0.001) was observed
after 3rd
week of incubation (21 Days), the enzyme activity slightly decreased after this
period (Fig. 4.28).
4.7. Purification of cell bound esterase from Bacillus subtilis MZA-75
Proteins precipitated by (NH4)2SO4 precipitation, were purified by elution through
Sephadex G-75 column in phosphate buffer at pH 7.0. The eluted fractions were collected
in volume of 2ml each and assayed for esterase activity using p-nitrophenyl acetate assay.
Fractions 12-16 showed high esterase activity and were merged together (Fig. 29). SDS-
PAGE analysis revealed single band corresponding to approximately 50 kDa (Fig. 4.30).
4.7.1. Substrate specificity of purified cell bound esterase
The purified enzyme was active against different p-nitrophenyl (pNP) acyl esters,
Purified enzyme demonstrated significant ester cleavage rate upto p-nitrophenyl caproate
(C6) and then gradually started decreasing from C8-C18. Best enzyme activity was
recorded with p-nitrophenyl butyrate (C4) (Fig. 4.31).
4.7.2. Degradation activity of purified PU esterase against PU film
About 50% of the PU film (initial weight = 50 mg) was degraded by PU esterase from
MZA-75 within 7 days. PU degradation in buffered control (without enzymes) was not
observed (< 0.0001 g) after 7-days of incubation.
81
Fig 4.22: Induction of membrane associated esterase by PU supplementation of the MSM
as depicted by plot of membrane associated esterase activity conducted for MZA-85 after
every 4 days. (triangle) represents membrane associated esterase activity in culture with
PU, while (circle) represent membrane associated esterase activity in culture without PU.
82
Fig 4.23: Induction of extracellular esterase by PU supplementation of the MSM as
depicted by plot of extracellular esterase activity conducted for MZA-75 after every 4
days. (triangle) represents extracellular esterase activity in culture with PU, while (circle)
represent extracellular esterase activity in culture without PU.
83
Fig 4.24: Induction of cell bound esterase by PU supplementation of MSM as depicted by
plot of membrane associated esterase activity against time of incubation conducted for
MZA-75 after every 4 days (triangle) represents cell bound esterase activity in culture
with PU, while (circle) represent cell bound esterase activity in culture without PU.
84
Fig 4.25: Optimization of temperature of incubation for esterase production from Bacillus
subtilis MZA-75 at pH 7; best results can be seen with 37 oC i.e. 0.493 mM/min/mg at
day 21st.
85
Fig 4.26: pH optimization for PU esterase production from Bacillus subtilis MZA-75 at
37 oC; best results can be seen on pH7 i.e. 0.593 mM/min/mg day 21
st.
86
Fig 4.27: Analysis of PU esterase production from Bacillus subtilis MZA-75 in the
presence and absence of yeast extract (as Nitrogen source) at pH 7 and temperature 37
oC; best results can be seen in the presence of nitrogen source i.e. 0.753 mM/min/mg at
day 28th.
87
Fig. 4.28: Optimization of time of incubation for production of PU esterase by Bacillus
subtilis MZA-75 at pH 7 and temperature 37 oC; best results can be seen on day 21
st i.e.
0.543 mM/min/mg.
88
Fig. 4.29: Esterase activity of different fractions collected from Sephadex G-75 elution of
cell bound esterase of MZA-75. Fractions 12-16 showing maximum activity. Primary
vertical exis represents enzyme activity (mM/min), while secondary vertical axis
represents Abs280 for rough estimation of total protein contents.
89
Fig. 4.30: SDS-PAGE of purified PU esterase purified from MZA-75. Lane 1: protein
marker; Lane 2&3: PU esterase approximately 50 KDa,
170 130
34
26
14
43
55
72
96
90
Fig 4.31: Specific esterase activity of purified esterase against para Nitro Phenyl acyl
ester with fatty acids of different carbon numbers. Best activity was observed against
para Nitro Phenyl butyrate.
0
0.5
1
1.5
2
2.5
3
3.5
C2 C4 C6 C8 C10 C16 C18
mM
/min
/mg
pNP acyl esters with different Carbon numbers
91
DISCUSSION
Polyurethanes have found widespread acceptance for a variety of applications.
The shortage of raw materials for synthesis of plastics necessitates the recycling of
disposed of plastics. It is difficult to recycle waste plastics completely because of the
incompatibility of most of the plastics and separation of polymeric materials of different
chemical nature from one another. Therefore, different hydrolytic processes are being
worked out for the consumption of waste either for energy recovery or extraction of raw
material for the production of valuable chemicals (Schnabel, 1981). Polyurethanes are
considered to be comparatively susceptible to microbial degradation (Morton and
Surman, 1994). Three types of polyurethane degradations have been identified in
literature fungal biodegradation, bacterial biodegradation and degradation by
polyurethanase enzymes (Howard, 2002).
This study was designed to isolate microorganisms capable of degrading water
insoluble polyurethane from soil. The involvement of bacterial esterases in the
esterolytic breakdown of the polyester polyurethane has also been studied. Soil samples
were screened for the isolation of PU degrading microorganisms, presuming that the soil
will be a rich source of various organisms. Both fungal and bacterial strains with
polyester polyurethane degrading capability have already been isolated from soil. Four
species of fungi Curvularia senegalensis, Fusarium solani, Aureobasidium pullulans
and Cladosporium sp, were obtained from soil and found to degrade ester-based
polyurethane (Crabbe et al., 1994). Shah et al. (2008) used a consortium of bacterial
strains isolated from soil for degradation of polyester polyurethane. A polyester
polyurethane diol degrading P. aeruginosa have also been isolated from soil.
Extracellular esterases have been found to be involved in this polyurethanolytic activity
of the isolated P. aeruginosa. PU degrading fungal strains have been isolated from sands
contaminated with PU (Loredo-Treveno et al. 2011).
In this study we isolated a fungal strain identified as Aspergillus tubingenesis for
its ability to utilize PU as a sole source of carbon. The isolated strain grew to sporulation
92
when inoculated on MSM agar with PU film overlaid as a sole source of carbon. After
four weeks of incubation, visible signs of degradation were observed on the treated PU
films. The exposed area was very fragile and the film disintegrated on detachment from
the surface of MSM agar plate.
Several fungi were found associated with soil buried PU foam when recovered.
These strains could utilize polyester PU as a sole source of carbon. These include
Trichoderma, Emericella, Fusarium, Aspergillus, Penicilliam and Gliocladium (Bentham
et al., 1987). Geomyces Pannorum has been found to be very effective with regard to its
polyester PU degrading capacity (Barratt et al., 2003). Other soil fungi with notable PU
degrading potential are Nectria, Plectosphaerella, Phoma, Neonectria and Alternaria.
Aspergillus niger has also been found to cause visible deterioration in 30 days (Russell et
al., 2011).
SEM and FTIR results also confirm the adherence of fungal mycelia on the
surface of film and its degradative effects. However no fungal growth or mycelial
adherence to the surface of PU films were observed when fungal spores were inoculated
and incubated in MSM supplemented with PU films in shake flask experiment. No
evidence of production of extracellular esterases was found during shake flask
fermentation. It is hypothesized that the isolated fungal strain needs physical contact with
the PU films to utilize it as a nutrient source, shaking in submerged culture might have
inhibited its ability to adhere to the PU film and thus their growth. In a study by
Bonhomme et al. (2003) SEM micrographs confirmed that fungi colonize the polymer
surface and leave the surface pitted and eroded after removal. The polymer after
microbial attack was physically weak and only a mild pressure was sufficient to
disintegrate it. Color changes like whitening and appearance of small holes on the
polymer surface recovered after 32 years burial, have been reported by Otake et al.
(1995). Wang et al. (2011) prepared rapidly degradable, non-toxic PU material and
confirmed its susceptibility to varying pH and enzymes by SEM and measurement of
weight loss. SEM results demonstrated the emergence of pits on the surface of treated PU
films. In another study two fungal strains i.e. Penicillium chrysogenum and Aspergillus
93
brasiliances were found to degrade PU composites in static cultures on sabouraud
dextrose agar (Macocinschi et al. 2011).
Notable changes in the FTIR spectrum of PU films treated with Aspergillus
tubingensis were the disappearance of peaks in the region 1100-1200 cm-1
(representing
stretching vibration of C-N and C-O) which were present in untreated control. The peak
at 1700 (representing stretching of amide carbonyl) also disappeared after treatment.
These results show that the isolated fungal strain can target urethane linkages (the hard
segment) of the PU used for these experiments. The two spectra (those of treated and
untreated PU films) are totally different with regard to the region between 500 and 800
cm-1
, showing that hydrocarbon portion of the polymer is also subject to degradation
under the test conditions. Although enzyme studies when A. tubingensis was grown in
MSM supplemented with PU were not convincing, combination of urethane bond
degrading enzymes and oxidoreductases might be involved in this polyurethanolytic
process.
Two bacterial strains were also isolated on the basis of their ability to grow solely
on PU as carbon source. Microscopy, biochemical and genetic analysis identified these
strains as Bacillus subtilis MZA-75 and Pseudomonas aeruginosa MZA-85. The ability
of both these species to degrade water dispersible Impranil DLN has already been
reported (Mukherjee et al. 2011; Rowe and Howard 2002). Howard and Hilliard (1999)
reported the growth of P. chlororaphis on Impranil DLN (water dispersible
polyurethane), and purified polyurethane hydrolyzing esterase from it.
Polyester PU films when treated with the isolated organisms MZA-85 and MZA-
75 and analyzed for any physical changes through scanning electron microscopy (SEM)
revealed surface modification indicating degradation. MZA-85 treated PU films
demonstrated bacterial cell adherence with cracks radiating from the point of adherence,
which indicates that in case of P. aeruginosa MZA-85 the degradative activity might be
dependent on physical contact of the cells with the film. In case of Bacillus subtilis MZA-
75 the treated PU films were observed with cracks, accompanied by little cell adherence.
94
Shah et al. (2008) analyzed PU films by SEM after one month treatment with a
consortium of bacterial isolates in Sturm test and observed modifications like appearance
of pits, cracks and black spots on the PU surface after treatment. Sarkar and adhikari
(2007) used SEM for observing modifications of polyester PU surface treated with P.
aeruginosa and reported gradual increase in the appearance of pits, cracks and valleys on
the surface of treated PU films from day 15 to day 180.
The infrared spectroscopy can be used to corroborate the biodegradation of
polymeric materials, mainly by monitoring the variation of characteristic bands. In
general the structural changes are minimal and a reliable interpretation depends on a right
analysis of a set of spectral data (Kloss et al., 2009). FT-IR analysis revealed that P.
aeruginosa strain MZA-85 degrades the polyester portion of the polyurethane. FT-IR
peaks at 1725 cm-1
and 1135 cm-1
representing ester linkage disappeared in the treated
polyurethane sample. Peak at 3325 cm-1
representing NH in the untreated PU film
broadened in the treated samples due to appearance of OH groups of alcohols. The peak
representing ester linkage decreased in intensity and a new peak representing carboxylic
acids appeared in treated samples. Similar results were obtained from FT-IR analysis of
the films treated with Bacillus subtilis MZA-75. Hydrolysis of ester functionality into
carboxylic acids is evident from a peak at 1725 cm-1
in the FT-IR spectrum of the treated
polyurethane films. This data supports the idea of involvement of microbial esterases in
the degradation of polyester polyurethane. Oprea and Doroftei (2011) reported that the
structural changes in the PU as a result of biodegradation, can easily be monitored by FT-
IR spectral analysis. He analyzed biodegradation of polyurethane acrylate with acrylated
epoxidized soybean oil blend elastomers by Chaetomium globosum through IR and
reported changes in the IR profile of the polymer exposed to C. globosum. The changes
observed indicated that both ester and urethane groups were attacked by the fungus.
Oprea (2010) also attributed the broadening of peak at 3270-3335 cm-1
to overlapping of
NH peak by OH. In our previous study we employed FT-IR for the analysis of chemical
changes in the PU treated with microbial consortium (Shah et al., 2008).
95
Gel permeation chromatography (GPC), a technique used for the determination of
molecular weight distribution of polymers, plays an important role in studying
biodegradable materials by giving an insight into the rate at which a material might
degrade, and revealing the presence of degraded polymer chains in a sample (Saunders
and MacCreath, 2010). The degradation of PU by P. aeruginosa strain MZA-85 was
analyzed through GPC. An increase in polydispersity accompanied by decrease in both
number average molecular weight (Mn) and weight average molecular weight (Mw) of
the polymer was observed in GPC results. It indicates that strain MZA-85 brought about
change in molecular weight by breaking the long chain polymer into shorter subunits.
Rek et al. (1986) studied the effect of UV irradiation on the photooxidative degradation
of PU pre-polymer synthesized from diphenylmethane-4,40-diisocyanate (MDI) and
polyester diol oligomers by GPC and observed that photo-oxidative decomposition is
followed by a decrease in molecular mass together with an increase in polydispersity.
Christenson et al. (2006) studied the effect of cholesterol esterase (CE) on the
degradation of commercial poly(ether urethane) (PEU) and poly(carbonate urethane)
(PCU) and observed no significant difference in molecular weight or polydispersity as
compared to the untreated control. In case of MZA-75 GPC analysis revealed an increase
in polydispersity index and decrease in both weight average molecular weight (Mw) and
number average molecular weight of the treated polyurethane films, which is an evidence
of changes in the molecular weight distribution of PU or biodegradation as a result of
treatment with Bacillus subtilis strain MZA-75. These results also indicate that the
degradation carried out by Bacillus subtilis MZA-75 is not limited to the surface of the
films.
The involvement of esterase in the degradation of PU was confirmed by detection
of ester hydrolysis products i.e. 1,4-butanediol and adipic acid by GC-MS analysis. GC-
MS chromatograms of ethyl acetate extract of strain MZA-85 and MZA-75 culture,
having PU as a sole source of carbon reveal 1,4-butandiol and adipic acid peaks, which
were not present in either biotic or abiotic control. Nakajima-Kambe et al. (1997)
investigated PU degradation metabolites, when exposed to C. acidovorans strain TB35
by GC-MS and reported the detection of diethylene glycol, trimethylolpropane, adipic
96
acid and 2,4- diaminotoluene. These compounds were derived from both the polyester
and polyisocyanate portions of the polyurethane. P. aeruginosa strain MZA-85 and
Bacillus subtilis MZA-75 not only depolymerize PU by hydrolysis of ester linkage but
also utilizes the intermediates as carbon source. The mineralization was also confirmed
by comparison of CO2 evolved in the presence and absence of polyurethane through
modified Sturm test. Yabannavar and Bartha (1994) used CO2 evolution test to evaluate
biodegradability of photosensitized polyethylene (PE), starch-PE, extensively plasticized
polyvinyl chloride (PVC), and polypropylene (PP) and found that plastics with additives
evolves more CO2 because of easy degradation of the additive. In our previous report, we
mentioned 4 fold high evolution of CO2 in Fusarium sp. AF-4 culture supplemented with
PE than in biotic control with no carbon source (Shah et al., 2009).
As far as the mechanism of degradation of PU is concerned, the strain MZA-85
easily degraded the aliphatic segment as compared to aromatic segment, resulted in the
formation of monomers 1,4- butanediol and adipic acid, as indicated by the GC-MS
results. We speculate that the esterases from strain MZA-85 specifically adsorb to the
ester bonds in the aliphatic or soft segment leads to depolymerization of polymer chain. It
is the point of initiation of degradation of the polymer chain. Later on the degradation of
aromatic or hard segments may occur after shredding the polymer chain by esterases, but
unfortunately no metabolite corresponding to the polyisocyanate portion could be
detected in the cell free supernatant. The growth of strain MZA-85 in the presence of
ester hydrolysis products revealed that it utilized the products as carbon source and
mineralized to CO2 and H2O.
Solid polymeric material like polyethylene are usually insoluble and difficult for
microorganisms to utilize, this is overcome by formation of microbial biofilm on the
surface of the polymer for efficient enzymatic activities (Gilan et al., 2004). Microbial
biofilms are multicellular microbial communities, adhered to solid surfaces and are
considered as potent degrading agents present in nature. Microbial communities in the
form of biofilm offer more resistance to antimicrobials. Study of different microbial
populations in most of the natural and artificial habitats revealed that most of them
97
produce biofilms which enhance their metabolic activity (Atkinson and Fowler, 1974;
Kirchman and Mitchell, 1982). The ability of P.aeruginosa MZA-85 to degrade polyester
PU by binding to the surface of the film accompanied by the fact that only membrane
associated esterases were induced as a result of PU supplementation indicated towards
the possibility of biofilm formation of P. aeruginosa MZA-85. This was confirmed by
crystal violet staining of the biofilm and scanning electron microscopy visualization.
Crystal violet staining and SEM visualization results, indicated biofilm forming capacity
of the P. aeruginosa MZA-85. No biofilm was observed in case of Bacillus subtilis
MZA-75.
Time course study for esterases from strain Pseudomonas aeruginosa MZA-85
revealed a gradual rise in cell bound esterase activity with time in the presence of PU,
while no change was observed in extracellular esterase activity in the presence or absence
of PU. Ohkawa et al. (1979) found an esterase associated with the outer membrane of P.
aeruginosa having affinity for long chain acyl esters in particular. Akutsu et al., (1998)
and Wilhelm et al. (1999) purified PU esterase from the outer membrane of C.
acidovorans strain TB35 and P. aeruginosa PAO1, respectively. Mukherjee et al., (2011)
also reported the role of extracellular esterases from P. aeruginosa in degradation of
polyurethane diol, but no induction of extracellular esterases was observed in this study.
The isolated Bacillus subtilis strain MZA-75 showed rise in both extracellular and
cell associated esterase activity when grown in the presence of polyurethane in minimal
salt medium. Conditions for production of cell associated esterase by MZA-75 were
optimized by weekly analysis of esterase activity under various conditions of
temperature, pH and in the presence or absence of yeast extract as nitrogen source. Time
of incubation needed to produce optimum enzyme activity was also determined. The cell
associated esterase was salted out by ammonium sulphate after extraction from cell by
French pressure cell homogenizer, and purified by sephadex gel G-75, a single protein
band corresponding to approximately to 51kDa on SDS-PAGE. Pure enzyme when tested
against different para nitrophenyl acyl esters, demonstrated best activity against p-
Nitrophenyl butyrate. The activity decreased by further increasing the length of acyl
98
chain. Lipases are differentiated from esterases on the basis of their ability to catalyze the
hydrolysis of acylglycerols with acyl chain lengths of >10 carbon atoms, but esterases
catalyze the hydrolysis of glycerolesters with acyl chain lengths of <10 carbon atoms
(Rhee et al., 2005). Esterases usually catalyze hydrolysis of triglycerides with fatty acids
shorter than C6 (Bornscheuer, 2002). Therefore, our results suggested that this is a type
of esterase. Purified enzyme degraded 50% of the PU film, when the later was treated
with it in 100 mM phosphate buffer, which confirms the effectiveness of this enzyme as
polyurethane degraders. In case of water insoluble solid substrates like plastics, it is very
difficult for enzymes to bind as compared to water-soluble substrates. To overcome this
problem plastic degrading enzymes possess some special properties that enhance their
ability to adsorb to solid surface. Ohtaki et al. reported that a hydrophobic protein adheres
to the PBSA surface at first and then drags a PBSA-degrading enzyme on to the surface
of PBSA (Ohtaki et al., 2006).
Kumar et al., 2012 recorded both intracellular and extracellular esterase activity
in Bacillus sp. Wang et al., 2010 purified an intracellular esterase from Bacillus cereus
and determined its molecular weight to be 43 kDa by SDS-PAGE. Riefler and Higerd
(1976) obtained intracellular esterase from Bacillus subtilis by sonic disruption, and
purified it by differential chemical and heating precipitation, DEAE-cellulose
chromatography, and Bio-Rad P-150 gel filtration chromatography, with an overall yield
of 59%. The purified enzyme hydrolyzed both aliphatic and aromatic acetate esters at
substrate concentrations of 0.25 M but did not hydrolyze amino acid esters. The exact
mechanism of how both extracellular and intracellular esterases collaborate to degrade
PU, it is however postulated that extracellular esterases produce oligomers by
hydrolyzing the polymer and then leave those for intracellular esterases to metabolize.
Generally, biodegradation occurs in two steps in which during the first step, chain
cleavage occurs and polymers are converted into their corresponding oligomers and
monomers which is also called as depolymerization. This is followed by mineralization in
which monomers and oligomers formed are sufficiently smaller in size and are
transported to the cytoplasm of cells of the microorganisms and get completely
mineralized. This is the process in which various byproducts such as Carbon dioxide,
99
water, methane and other inorganic substances are formed depending on whether the
process is aerobic or anaerobic. The ability of aerobic microorganisms to mineralize
polymeric substances can be traced by entrapment of CO2 evolved as a result of
mineralization by using modified sturm test. Sturm test (Sturm, 1973) has commonly
been employed for evaluation of the biodegradability of polymer materials (Calmon et
al., 2000). Various modifications of this test have been used for the measurement of
carbon dioxide evolution during degradation of biodegradable polymers (Muller et al.,
1992), and the aliphatic and aromatic compounds (Kim et al., 2001).
It was observed during current study that the amount of CO2 produced in case of
both Pseudomonas aeruginosa MZA-85 and Bacillus subtilis MZA-75, when polyester
polyurethane is used in the medium as carbon source, is greater than the amount of CO2
produced in the absence of polyurethane. This provides evidence that both Pseudomonas
aeruginosa MZA-85 and Bacillus subtilis MZA-75 are mineralizing the polymer. The
colony forming units/ml count for both the isolates increased significantly during test
period. Previously we observed similar results when we employed modified sturm test to
evaluate polyurethane mineralization capability of the consortium isolated from soil
(Shah et al., 2008). The ability of the isolated strains to utilize the degradation products
detected during GC-MS, i.e. 1,4-butanediol and adipic acid as sole source of carbon was
analyzed by growth analysis of both MZA-85 and MZA-75 on MSM supplemented with
these metabolites. Both these isolates demonstrated convincing capacity to utilize both
these metabolites.
The role of both extracellular and intracellular esterases of Bacillus subtilis can
further be studied. The results show that the purified enzyme can be a handy addition to
the collection of polyurethanolytic esterases reported so far. Techniques like protein
modeling, site directed mutagenesis and genetic manupulations can be used to enhance its
quantity and activity and may potentially be used for recycling of the monomers
constituting polyesters or polyester based polyurethanes.
100
Conclusions
Waste dumping sites especially those where plastics are dumped may be a rich source of
microorganism capable of degrading plastics including polyurethane.
Bacillus subtilis MZA-75 and Pseudomonas aeruginosa MZA-85 can degrade polyester
polyurethane mainly by targeting the polyester segment.
Sturm test results with MZA-75 and MZA-85 reveal that the degradation products are
mineralized to CO2 and H2O.
Aspergillus tubingensis cannot utilize PU as a substrate in liquid broth in shake flask
fermentation.
Pseudomonas aeruginosa MZA-85 cannot produce extracellular esterase to degrade
polyester polyurethane and rely on its cell bound esterases for this purpose.
Polyurethane supplementation can induce both extracellular and cell bound esterases in
Bacillus subtilis MZA-75.
Cell bound esterase from MZA-75 is a true esterase of 51 KDa and have maximum
esterolytic activity against p-nitrophenyl butyrate.
101
Future prospects
Plastic contaminated sites can be exploited for other potential polyurethane degraders.
After the degradation of polyester portion (soft segment), the fate of hard segment can be
studied.
Field bioremediation tests can be conducted to determine the effectiveness of MZA-75
and MZA-85 in soil.
Cell bound esterase from MZA-85 and extracellular esterase from MZA-75 can be
purified and characterized.
The mechanism by which both extracellular and cell bound esterase in MZA-75 share PU
degradation can be studied.
The expression studies of esterases from MZA-75 and MZA-85 can be conducted in
more efficient expression systems and their utility in biochemical monomerization be
studied.
102
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Appendix
Table 1: Growth of Bacillus subtilis MZA-75 on PU as sole carbon source. T1-T3 represents replicates of Test (PU+MSM+MZA-75),
while C1-C3 represent abiotic control without any carbon source. Enzyme activity is expressed in U/mg. T.avg means average specific
esterase activity in test, and C.avg means average specific esterase activity in control.
Days T1 T2 T3 C1 C2 C3 T.avg C.avg St.dev.T St.dev.C
0 0.045 0.056 0.05 0.053 0.066 0.052 0.050 0.057 0.00550 0.0078
3 0.081 0.062 0.091 0.081 0.062 0.066 0.078 0.069 0.01473 0.0100
6 0.133 0.098 0.099 0.065 0.074 0.065 0.11 0.068 0.01992 0.0051
9 0.151 0.154 0.169 0.078 0.085 0.092 0.158 0.085 0.00964 0.007
12 0.211 0.235 0.2 0.083 0.074 0.069 0.2153 0.075 0.01789 0.007
16 0.198 0.231 0.168 0.088 0.093 0.084 0.199 0.088 0.03151 0.004
19 0.204 0.223 0.234 0.063 0.078 0.075 0.220 0.072 0.01517 0.007
22 0.186 0.191 0.235 0.099 0.063 0.084 0.204 0.082 0.02696 0.0180
120
Table 2: Growth of Pseudomonas aeruginosa on PU as sole carbon source. T1-T3 represents replicates of Test (PU+MSM+MZA-85),
while C1-C3 represent abiotic control without any carbon source. Enzyme activity is expressed in U/mg. Tavg means average specific
esterase activity in test, and Cavg means average specific esterase activity in control
Days T1 T2 T3 C1 C2 C3 T.avg C.avg St.dev.T St.dev.C
0 0.033 0.056 0.041 0.053 0.061 0.042 0.043333 0.052 0.0116 0.009
3 0.063 0.076 0.077 0.071 0.062 0.045 0.072 0.0593 0.0078 0.013
6 0.091 0.13 0.099 0.0732 0.074 0.046 0.106 0.0644 0.0205 0.015
9 0.124 0.154 0.115 0.051 0.032 0.053 0.131 0.0453 0.0204 0.011
12 0.153 0.175 0.188 0.071 0.063 0.066 0.172 0.0666 0.0176 0.004
16 0.178 0.188 0.164 0.051 0.083 0.064 0.176 0.066 0.0120 0.016
19 0.195 0.2 0.186 0.073 0.078 0.049 0.193 0.0666 0.00709 0.015
22 0.186 0.191 0.235 0.065 0.021 0.063 0.204 0.0496 0.0269 0.024
121
Table 3: Time course of specific activity for extracellular esterases from Bacillus subtilis MZA-75. Four replicates (T1-T4) represent
test cultures (MSM+PU+MZA-75), while (C1-C4) represent abiotic control (MSM+MZA-75). Enzyme activity is expressed in U/mg.
Tavg means average specific esterase activity in test, and Cavg means average specific esterase activity in control.
Days T1 T2 T3 T4 C1 C2 C3 C4 T.avg C.avg St.dev.T St.dev.C
0 0.24437 0.1701 0.22 0.0959 0.2431 0.2492 0.1015 0.2677 0.1825 0.2154 0.0655 0.077
4 0.2433 0.2522 0.234 0.221 0.21879 0.2305 0.2655 0.283 0.2377 0.2494 0.0133 0.03
8 0.3206 0.2754 0.298 0.3116 0.21 0.2229 0.1671 0.2614 0.3014 0.2154 0.0196 0.039
12 0.30867 0.3554 0.299 0.2853 0.21206 0.2202 0.2324 0.2284 0.3122 0.2233 0.0304 0.009
16 0.30484 0.3343 0.303 0.3774 0.30712 0.3271 0.1923 0.2971 0.3298 0.2809 0.0349 0.06
20 0.4574 0.4018 0.473 0.5197 0.27684 0.0923 0.2716 0.2821 0.463 0.2307 0.0486 0.092
24 0.48572 0.5299 0.458 0.5046 0.11224 0.1497 0.1995 0.1206 0.4947 0.1455 0.0302 0.039
122
Table 4: Time course of specific activity for cell bound esterases from Bacillus subtilis MZA-75. Four replicates (T1-T4)
represent test cultures (MSM+PU+MZA-75), while (C1-C4) represent abiotic control (MSM+MZA-75). Enzyme activity is
expressed in U/mg. Tavg means average specific esterase activity in test, and Cavg means average specific esterase activity in
control
Days T1 T2 T3 T4 C1 C2 C3 C4 Tavg Cavg St.dev.T St.dev.C
0 0.65906 0.597 0.426 0.3954 0.33747 0.6801 0.1713 0.6594 0.5195 0.4621 0.1285 0.249
4 0.34069 0.3558 0.336 0.3155 0.23449 0.0673 0.2019 0.1978 0.3369 0.1754 0.0167 0.074
8 0.64787 0.7058 0.708 0.6637 0.3336 0.1859 0.3063 0.3336 0.6815 0.2899 0.0304 0.07
12 0.91207 0.9313 0.937 0.956 0.34959 0.3443 0.2966 0.143 0.934 0.2834 0.0181 0.097
16 0.94055 0.9685 0.974 0.9965 0.28429 0.345 0.2401 0.4002 0.9698 0.3174 0.023 0.07
20 1.18354 1.2035 1.236 1.2185 0.24342 0.2338 0.2097 0.229 1.2104 0.229 0.0223 0.014
24 1.15755 1.1295 1.151 1.1832 0.15557 0.2074 0.1084 0.231 1.1552 0.1756 0.0221 0.055
123
Table 5:Time course of specific activity for cell bound esterases from Pseudomonas aeruginosa MZA-85. Four replicates (T1-T4)
represent test cultures (MSM+PU+MZA-85), while (C1-C4) represent abiotic control (MSM+MZA-85). Enzyme activity is expressed
in U/mg. Tavg means average specific esterase activity in test, and Cavg means average specific esterase activity in control
Days T1 T2 T3 T4 C1 C2 C3 C4 Tavg Cavg St.dev.T St.dev.C
0 0.489 0.513 0.5 0.506 0.552 0.586 0.584 0.579 0.502 0.5752
5
0.01016 0.0157
4 0.201 0.227 0.219 0.225 0.216 0.212 0.223 0.221 0.218 0.218 0.01183 0.0049
8 0.894 0.966 1.301 1.003 0.369 0.342 0.367 0.359 1.041 0.3592 0.17914 0.0122
12 2.312 2.151 2.201 2.22 0.732 0.73 0.713 0.721 2.221 0.724 0.06728 0.008
16 1.96 1.85 2.19 2.05 1.001 0.8 1.02 0.859 2.0125 0.92 0.14384 0.1075
20 1.754 1.635 1.664 1.55 0.596 0.562 0.571 0.563 1.65075 0.573 0.08413 0.0158
24 1.098 1.234 1.146 1.062 0.613 0.611 0.641 0.611 1.135 0.619 0.07443 0.0146
124
Table 6: Effect of pH on production of cell bound esterase from MZA-75.
Day pH5 (U/mg) Avg pH6 (U/mg) Avg pH7 (U/mg) Avg pH8 (U/mg) Avg pH9 (U/mg) Avg
1 0.153 0.133 0.152 0.146 0.16 0.143 0.153 0.152 0.148 0.142 0.127 0.139 0.169 0.153 0.14 0.154 0.158 0.127 0.132 0.139
7 0.22 0.182 0.192 0.198 0.275 0.312 0.286 0.291 0.296 0.324 0.316 0.312 0.185 0.153 0.163 0.167 0.222 0.246 0.225 0.231
14 0.195 0.181 0.179 0.185 0.311 0.361 0.315 0.329 0.366 0.34 0.35 0.352 0.234 0.193 0.212 0.213 0.194 0.175 0.186 0.185
21 0.218 0.241 0.249 0.236 0.51 0.481 0.497 0.496 0.611 0.588 0.58 0.593 0.244 0.21 0.212 0.222 0.211 0.21 0.188 0.203
125
Table 7: Effect of temperature on production of cell bound esterase from MZA-75.
Day (U/mg) Avg
(U/mg) Avg
(U/mg) Avg
(U/mg) Avg
(U/mg) Avg
1 0.152 0.164 0.15 0.15 0.17 0.149 0.15 0.159 0.164 0.151 0.171 0.16 0.151 0.135 0.143 0.143 0.166 0.149 0.153 0.156
7 0.222 0.196 0.17 0.19 0.1 0.21 0.18 0.19 0.15 0.19 0.18 0.1 0.14 0.14 0.16 0.16 0.11 0.14 0.14 0.13
14 0.38 0.32 0.35 0.35 0.4 0.46 0.43 0.45 0.28 0.25 0.55 0.3 0.15 0.18 0.17 0.17 0.07 0.01 0.05 0.045
21 0.38 0.35 0.35 0.36 0.5 0.48 0.48 0.49 0.34 0.33 0.32 0.3 0.13 0.18 0.17 0.16 0.01 0.04 0.04 0.032
28 0.325 0.31 0.33 0.32 0.35 0.381 0.35 0.361 0.143 0.178 0.154 0.15 0.17 0.14 0.155 0.155 0.0001 0.001 0.0003 0.0004
126
Table 8: Effect of yeast extract on production of cell bound esterase from MZA-75. Enzyme
activity is expressed in U/mg.
Days Yeast extract present (U/mg) Avg Yeast extract absent (U/mg) Avg
1 0.15 0.138 0.15 0.146 0.0532 0.0002 0.0003 0.0179
7 0.518 0.499 0.519 0.512 0.382 0.343 0.37 0.365
14 0.675 0.66 0.622 0.6523 0.439 0.404 0.421 0.4213
21 0.765 0.738 0.756 0.753 0.591 0.610 0.577 0.5926
28 0.584 0.549 0.559 0.564 0.553 0.521 0.519 0.531
127
Table 9: Effect of time of incubation on enzyme production from MZA-75. Enzyme activity is
expressed in U/mg.
Days T1 T2 T3 T4 Avg St.dev.
0 0.045 0.054 0.056 0.041 0.049 0.007164728
7 0.315 0.277 0.294 0.271 0.28925 0.019737865
14 0.422 0.418 0.391 0.4 0.40775 0.014705441
21 0.512 0.531 0.513 0.479 0.50875 0.021669872
28 0.498 0.447 0.469 0.473 0.47175 0.020902552
128
Table 10: Protein concentration and specific enzyme activity measured in different fraction
eluted through Sepahdex G-75, during enzyme purification
Protein conc.
mg/ml
Specific Activity
(U/mg)
0.1001 0.054
0.1112 0.018
0.1136 0.033
0.1236 0.1371
0.2453 0.128
0.2536 0.186
0.1141 0.1076
0.2355 0.1187
0.3523 0.1027
0.1078 0.1153
0.2488 0.1159
0.3654 0.5025
0.3188 0.491
0.3256 0.5122
0.3739 0.6192
0.3664 0.5265
0.1581 0.1292
0.1928 0.1271
0.2329 0.1266
0.0923 0.1531
0.0873 0.0981
0.0875 0.0912
0.0546 0.0631
0.0546 0.0642
129
Table 11: Biofilm quantity as depicted from Abs750, after crystal violet staining.
Days Abs750
1 0.070666667
4 0.1595
8 0.232
130
Table 12: Activity of purified enzymes, against p-nitrophenyl esters of varying acyl chain.
Enzyme activity is expressed in U/ml.
C4 2.699 2.879 2.782 2.786666667
C6 2.654 2.642 2.666 2.654
C8 1.305 1.467 1.2 1.324
C10 0.511 0.536 0.519 0.522
C16 0.227 0.199 0.213 0.213
C18 0.188 0.167 0.167 0.174