Microbial Degradation
Activities
Project Proposal
Poh Yong Rui
Ong Kim Yao
Hwa Chong Institution (High School)
Mentor: Mrs Har Hui Peng
1
Introduction
In the 20th century, the demands of society have caused a great rise in the usage of plastic.
Plastics are defined as polymers which are mouldable upon heating, to take on a desired
shape and then retain that shape (Shah, 2007).
The use of plastics has become a significant part of today’s economy. Various kinds of plastic
have been used extensively for manufacturing a wide range of products. Polyethylene has
been used to make plastic bags, food packaging films and toys; polystyrene for making
disposable cups, packaging material and laboratory ware; and polypropylene for making
bottle caps, straws, and car seats.
Plastics are non-polar and inert in nature, hence they remain almost unaffected once disposed .
As plastics do not biodegrade they pose detrimental consequences on the environment, and
thus the extensive usage of plastics has drawn much attention. Plastic sheets and bags are
impermeable to water and air and, if buried in the ground will cause infertility in soil, disrupt
degradation of other substances, deplete underground water resource and pose danger to the
ecosystem. In the sea, plastic rubbish from ropes and nets chokes and entangles marine life.
Plastic waste cannot be incinerated as they release air pollutants which cause various health
problems. The burning of polyethylene and polystyrene produces toxic irritant products,
leading to human disorders and their classification as human carcinogens (Shah, 2007). There
is thus an increasing need to dispose plastic waste in an environmentally friendly manner.
Despite the inert nature of polymer plastics, past research has shown that biodegradation of
plastic is not impossible or improbable. Degradation is termed as the physical or chemical
change in polymer as a result of environmental factors, such as light, heat, moisture, chemical
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conditions or biological activity. Past research has shown that some bacterial and fungus are
able to degrade plastic, the process is catalysed by exposure to light, UV radiation, and heat.
This project aims to investigate the optimal conditions for photodegradation (physical
degradation of polymer into shorter chains) and thermal degradation (chemical degradation of
polymer by rupture of bonds into radical sites). Their efficiency under different aerobic
activity will also be investigated, so that the most efficient breakdown of plastic may occur.
This is coupled with the biodegradation of HDPE plastics by Pseudomonas putida and
Sphingomonas macrogoltabidus. Such information would not only allow for efficient
biodegradation of polymer plastic, but also, plastic can now be produced from monomers
vulnerable to the degredation of microorganisms, allowing them to be biodegradable.
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Literature Review
High-density polyethylene (HDPE) is a synthetic polymer which consists of chains of
ethylene (Nanda et al., 2010). The light weight and durability of HDPE plastics has resulted
in its widespread use. However, its rapid use has resulted in landfills to be filled with HDPE.
Coupled with the low biodegradability of HDPE, rapid growth of landfills has posed a threat
to the ecosystem. Delgi-Innocenti et al. (2001) showed that polymer which may appear safe
for degradation may instead produce toxic chemicals on degrading (as cited in Nanda et al.,
2010, p. 57). This has caused a pressing need to find alternatives to accelerate the
biodegradation process of HDPE. Much research has shown that microbes can help to
degrade plastics. Microbes can attach to the hydrophilic surfaces of HDPE, and grow with the
plastic polymer as the carbon source (Arutchelvi et al., 2008). They will then secrete
extracellular enzymes, resulting in the cleavage of carbon-carbon bonds of the HDPE
(Premraj & Doble, 2005), thereby releasing oligomers and monomers which are utilised by
the microorganisms as energy (Nanda et al., 2010).
Cornware, also known as starch-polyethylene plastics, are polyethylene mixed with other
biodegradable substances such as corn, yam and starch (Olive Green Marketing, n.d.). These
plastics are more environmentally friendly than HDPE due to its use of starch, which is
biodegradable. Furthermore, the use of starch in making plastic has decreased the time taken
for biodegradation to occur (Borghei et al., 2010). Bastioli (2001) showed that such plastic
reduced the greenhouse effect (Borghei et al., 2010) In addition to this eco-friendly
advantages, the use of starch allows the use polyethylene to be maximised as well as provide
an alternative to our limited resources due to the “consistent growth of plants” (Kaur &
Gautam, 2010). A research done by Borghei et al. (2010) showed that low-density
polyethylene (LDPE) with potato starch can biodegrade to a greater extent than LDPE, and to
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an even greater extent by Pseudomonas aeruginosa. This was because starch became the only
source of carbon for the bacteria, and the breakdown of starch caused a fracture in the
polyethylene structure, thereby resulting in biodegradation.
Providing pre-treatment to the plastic such as UV and thermal radiation before inoculating
them in bacteria culture have been observed to accelerate chemical degradation through the
absorption of energy. This is because the plastic polymer absorbs the radiant energy which
will cause a cleavage (breaking of bonds) within the bonds of the polymer plastic, resulting in
the formation of free radicals from the plastic (Gijsman et al., 1999). These radicals cannot
recombine easily (Albano et al., 2005), thereby speeding up the biodegradation process. A
research done by Albano et al. (2005) showed a decrease in tenacity of HDPE plastic with
increasing gamma irradiation, further suggesting that pre-treatment will enhance the
breakdown of plastic.
Thermal exposure has also been shown to catalyse the breakdown of plastics. Thermal
exposure allows the plastic to reach a higher temperature than its surroundings (Andrady,
1999) and this rise in temperature results in an acceleration of light-induced degradation. It is
thus suggested that pre exposure to heat will cause accelerated breakdown of polymer plastic
due to the synergistic effect with UV pre- treatment. Also, thermal exposure is interconnected
to humidity which also affects breakdown. High humidity is known to accelerate the rates of
biodegradation of polymer plastic as absorbed water leads to increased accessibility of
atmospheric oxygen. As such, exposure to heat as pre- treatment is likely to show synergistic
biodegradation under aerobic conditions.
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Plastic biodegradation can occur under 2 conditions: aerobic and anaerobic. Under aerobic
conditions, enzymes secreted by the bacteria catalyses the breakdown of polymer plastic into
oligomers, which diffuse into the bacteria (Arutchelvi et al., 2008). These oligomers are then
converted into energy or assimilated. The final products are heat, carbon dioxide, water and
biomass. A research done by Huang et al. (2004) suggested that the breakdown of PEGs
(polyethylene glycols) under aerobic conditions was more efficient due to “the employment
of efficient microbes”. This research suggests that HDPE, which is polyethylene as well,
undergoes biodegradation more efficiently with oxygen. Under anaerobic conditions,
polymer plastics are first broken down into monomers, catalysed by bacterial enzymes. The
monomers are then converted into carbon dioxide and organic acids containing C and H.
Methane is then produced, largely a result of the reduction of carbon dioxide with hydrogen,
together with the breakdown of organic acid bonds to form carbon dioxide and methane
(Verma, 2002). The final products are thus carbon dioxide, water and methane (Arutchelvi et
al., 2008). While Huang et al. (2004) suggested that PEGs break down more efficiently under
aerobic conditions, Rees (1980) showed that aerobic biodegradation might cause the
temperature to rise above 70°C (as cited in Lanini et al., 2001, p. 68). This might in turn
hinder the development of the bacteria, as well as contribute to global warming.
While much research has been done on biodegradation of plastic in liquid medium, Sierra et
al. (2003) suggested that biodegradation of polychlorinated biphenyls (PCBs) would occur
faster under soil conditions. The presence of natural compounds such as flavonoids and lignin
would help to replace conditions that enhance PCB degradation such as biphenyl addition,
which poses adverse health effect. Though PCBs are not plastic, this gives a general idea of
how soil conditions can accelerate the biodegradation of plastic, by providing organic
compounds. Furthermore, research has suggested that loamy soil would result in higher
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biodegradation rate than sandy soil, due to different growth conditions for microbes, and
environmental differences between different soil, such as temperature and moisture (Mostafa
et al., 2010).
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Rationale
While past researches have identified the use of bacteria to enhance the biodegradation of
HDPE, little have been done to find out the effect of soil contents on biodegradation of
plastics. Studies by Sierra et al. (2003) suggested that the presence of compounds such as
lignin would help to enhance biodegradation as well. Therefore, this project investigates the
effect of soil bacteria on the biodegradation of plastic under various conditions.
UV radiation during the pre-treatment process contributes to the biodegradation process since
it releases free radicals from plastic, thereby increasing the surface area for bacterial enzymes
to act on. Therefore, this project seeks to study the optimum exposure time for UV irradiation
on HDPE that accelerates the biodegradation process.
Objectives
The objectives of this project are:
1. To study the effects of different environmental conditions on the biodegradation of
HDPE by the selected microbes.
2. To study the effects of varying exposure time of HDPE to UV radiation on its
biodegrability by the selected microbes.
3. To determine the optimum conditions to achieve maximum biodegradation of HDPE
by the selected microbes.
Hypothesis
The following factors affect the rate of biodegradation of HDPE plastic:

Types of microbe culture

Environmental conditions for biodegradation
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
Exposure time to UV radiation

Types of plastic
Variables
Independent Variables

Types of bacterial culture

Exposure time to UV
radiation


Dependent Variables
Constant Variables



Change in dry mass
Amount of
of HDPE samples
bacterial culture
Amount of dissolved
used
Environmental conditions
O2 gas present in
for soil degradation
tube

Amount of culture
medium for
bacteria/fungus
Types of plastic

Amount of HDPE
plastic used
Other possible independent variables (can be experimented on if time allows)

Wavelength of UV radiation

Temperature and time of thermal radiation
Apparatus

Sterile test containers

Inoculating loop

Alcohol burner

Incubator

Tweezer

Oven (capable of 150°C)
9

UV lamp (for 365 nm UV radiation)

Measuring scale

Autoclave

Vernier dissolved O2 probe

Datalogger

Spatula

Thermometer (to measure small changes in temperature of at least 0.25°C)

Spectrophotometer

Rotary shaker
Materials

Bacteria (as suggested from AOS side)
o P. putida bacterial strain

Obtained from ATCC (Catalog Number: 4359)
o S. macrogoltabidus bacterial strain

Obtained from ATCC (Catalog Number: 51380)

Nutrient broth

Loamy soil (obtained from Far East Flora Singapore)

Wire mesh

Ethanol

Aluminium foil

Paper towels

HDPE plastic (from grocery bags, with the symbol “2”)

Deionised water

Bleach
10

M63 minimal media

Cornware

Petri dishes

LB agar powder
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Outline of method
The flow chart below summarises our method:
Preparing bacterial
culture
Preparing soil
conditions
Pre-treatment of plastic
Treatment of plastic by microbes
Change in dry
mass of HDPE
samples
Amount of carbon
dioxide gas present in
flask
Temperature of
contents of
flask
Cell density of
bacterial
culture
A. Preparing bacterial culture
1. 17 test containers will be prepared. They will be labelled as shown below:
Liquid medium
+115°C, 48h
+115°C, 48h
Ctrl + UV, H
Ctrl + UV, H
Sphingomonas macrogoltabidus
Ctrl + UV, H
Ctrl – H
UV – 120h
UV – 96h
UV – 72h
Ctrl + H – UV
UV – 120h
UV – 96h
UV – 72h
Pseudomonas putida
UV – 72h 96h 120h
Control
Legend: H: heat treatment, UV: UV treatment
Soil conditions
Ctrl
Sphingomonas macrogoltabidus
Best
Ctrl
Best
Ctrl
Best
Pseudomonas putida
Control
For soil conditions, the best conditions concluded from plastic biodegradation
in liquid medium will be used, as shown in the above diagram.
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Each set-up above will be done in triplicates. This can be simplified by the diagram
below:
Liquid medium
Add microbes individually
Control (x3)
Treatment
Control
UV radiation
(72h, 96h and
120h)
This set-up will
contain bacterial
culture and heattreated HDPE
plastic which is
not UVirradiated.
3 set-ups will be prepared.
All 3 will not contain
bacterial culture. The 3 setups will contain heattreated HDPE plastics
which are UV-irradiated for
72h, 96h and 120h
respectively.
Soil conditions
By using the best conditions
concluded from plastic degradation in
liquid medium, degradation under soil
conditions will be carried out.
The controls will be

Thermal radiation with microbes

Thermal radiation without microbes

UV radiation without microbes
2. Bacterial culture (Pseudomonas putida and Sphingomonas macrogoltabidus) will be
prepared by inoculating each bacteria in 10ml of NB broth. P. putida culture will be
inoculated overnight, while S. macrogoltabidus culture will be inoculated for 48h.
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The cell density of the bacteria culture will be adjusted to approximately 4 × 108
cells/ml, as suggested by Farrell & Quilty (2002). The corresponding cell densities are
0.44 AU for P. putida and 0.46 AU for S. macrogoltabidus (according to Section H).
3. For liquid medium, the test containers will be prepared according to Huang et al.
(2004), with some modifications. 2ml of bacterial culture (prepared in Section A Step
2) will be inoculated in a centrifuge tube with 13.5ml of M63 minimal media and
4.5ml of NB. The NB will allow the bacteria to grow and adapt to its environment
before plastic degradation starts. The control tube will contain 15ml of M63 minimal
media and 5ml of NB.
B. Preparing soil conditions
1. The soil samples will be prepared according to Mostafa et al. (2010) and Orhan et al.
(2004), with some modifications. Loamy soil will be purchased from Far East Flora
Singapore, and sieved with a mesh to remove gravel.
2. The soil will be autoclaved to remove any microbes present in the soil.
3. The test containers will be filled with 20cm3 of loamy soil, and the soil will be
adjusted to 50% of its maximum water holding capacity. The maximum water holding
capacity of the soil will be determined by adding deionised water to a centrifuge tube
filled with 50cm3 of autoclaved soil, until the tube is filled to the brim. The volume of
water added is the maximum water holding capacity of the soil.
4. 2ml of bacterial culture (prepared in Section A Step 2) will be added to the soil each.
The control set-up will contain the soil only.
5. The containers are kept at room temperature and 70% humidity.
C. Pre-treatment of HDPE
1. HDPE plastics will be cut from a grocery bag for UV radiation. UV radiation will be
carried out according to Johnson et al. (1993) and Morancho et al. (2006), with some
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modifications. Each HDPE sample will be exposed to UV radiation with a wavelength
of 365 nm using a UV lamp. The exposure time will be varied as 72h, 96h and 120h
(since Morancho et al. (2006) suggested an exposure time of 98.7h). The control
plastic samples will not be exposed to any UV radiation.
2. The samples will then be exposed to heat treatment at 115°C for 48 hours in an oven.
3. The plastics will be cut into small pieces of plastics, with masses between 0.015g and
0.025g. The mass of each plastic sample will be recorded. The plastics will be placed
in test containers, and the containers will be labelled with the type of bacteria and UV
irradiation time, as shown in Section A Step 1.
D. Treatment of HDPE with bacteria
1. The treatment process will be carried out according to Nanda et al. (2010), with some
modifications. HDPE samples will be disinfected with ethanol before they are placed
into the liquid medium in the respective test containers prepared in Section A Step 3
(1 HDPE sample each). The containers are then loosely capped and kept at 37°C.
2. Separate HDPE samples will be placed in the soil in the respective containers
prepared in Section B Step 4 at 2.5cm depth, and kept at room temperature (1 HDPE
sample each).
3. The set-ups containing liquid medium will be placed on a slow rotary shaker to ensure
that the HDPE samples are evenly exposed to the bacterial culture.
4. The M63 media will be changed every week as suggested by Pometto et al. (1992).
The HDPE sample will be removed, and the culture will be allowed to stand for 1h.
5ml of liquid medium will then be removed using a pipette. New M63 media will then
be added to replace the amount of culture removed. (refer to Section F). This process
will remove waste materials and dead cells from the medium.
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5. The HDPE samples will be exposed to the bacterial culture until a significant change
in the mass of plastic is recorded. Readings will be taken every 7 days.
E. Dry mass of plastic samples after treatment
1. At every reading (every 7 days), the HDPE samples will be removed from the tube
using a tweezer.
2. The HDPE samples will be rinsed with deionised water. This will help to remove any
bacterial biofilm and cell mass attached to the samples.
3. Bleach will be added to the waste materials before disposal.
4. The samples will then be dried and weighed. To find the percentage change in mass,
the difference between the final mass and initial mass (after pre-treatment) of the
samples will be divided by the initial mass. This can be summarised by the below
formula:
Percentage change in mass 
Initial mass (measured after pre-treatment)  Final mass
100%
Initial mass
5. A graph will then be plotted to show the percentage change in mass of HDPE samples
over time. A faster rate of decrease in HDPE mass would indicate a faster rate of
biodegradation.
6. The HDPE samples will then be disinfected using ethanol, to reduce the chances of
contamination by other microbes, before being returned to the tube.
F. Amount of dissolved oxygen present in the tube
1. The amount of dissolved oxygen will be monitored every 7 days.
2. Dissolved oxygen concentration will be measured from the liquid medium using a
carbon dioxide probe and a datalogger.
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3. Oxygen is taken in during the biodegradation process and through respiration. Thus, a
decrease in oxygen concentration will suggest microbe activity in the containers.
4. Similarly, a graph will be plotted to show the change in amount of oxygen over time.
G. Cornware
1. The entire experiment will be repeated with cornware in place of HDPE plastics.
H. Plotting the standard curve
1. A standard curve will be plotted before the experiment for each bacteria to determine
the turbidity of bacteria culture at different cell densities. It will be carried out
according to Reynolds & Farinha (2005), with some modifications.
2. Bacterial culture samples will be prepared by inoculating 1 loopful of bacteria into
10ml of NB.
3. 10-fold serial dilutions will be carried out until 10-8.
4. 0.1ml and 1ml of bacterial culture for each dilution factor will be added to a petri dish
respectively. The petri dishes will be labelled with the dilution factor and volume of
culture added.
5. NB agar will be prepared by adding deionised water to NB agar powder. The agar will
be autoclaved, poured into the petri dishes and allowed to solidify. Directions for
preparing agar plates will be followed.
6. The agar plates will then be inverted and incubated at 37°C for 24h.
7. At the end of the incubation, the number of colonies on each plate will be counted.
Plates with too many colonies and cannot be counted, and plates with less than 30
colonies will not be counted.
8. The cell density of the bacterial culture of each dilution factor will be calculated using
the following formula (assuming each colony represents 1 bacteria cell):
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Cell density (no. of bacteria cells/ml ) 
Number of colonies
Dilution factor  Amount of bacterial culture plated (ml )
9. Following that, 1ml of bacterial culture for each dilution factor (prepared in Section J
Step 3) will be pipetted into a cuvette and its optical density will be read at an
absorbance of 550nm. 1ml of NB will be used as the blank.
10. A graph will be plot showing the optical density of each sample against the cell
density of the sample.
11. The cell density of the bacterial culture will be adjusted to 4 × 107 cells/ml according
to the graph plotted (refer to Section A Step 2).
Safety precautions
During the experimentation, latex gloves will be worn when handling microorganisms. All
microbial cultures and vessels used to contain them will be decontaminated by autoclaving
before disposal.
Perpex goggles will be worn when working with UV irradiation, and plastic pieces will be
irradiated in an opaque box.
Timeline
Period
Activity
Write proposal
October – November 2011
Confirm methods and proposal
November 2011
Experimentation
March – July 2012
Analyse results
May – July 2012
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Write research paper
July 2012
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