ACCELERATING THE BIODEGRADATION OF POLY(LACTIC ACID) AT MESOPHILIC
TEMPERATURES
By
Pooja C Mayekar
A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Packaging—Doctor of Philosophy
2024
ABSTRACT
As plastic production and consumption increase globally, so does the amount of plastic
waste to be disposed of and managed. Most worldwide plastic waste ends in landfills, open
dumps, or leaks into the environment. Hence, new alternatives, such as developing biobased
and biodegradable polymers, have been proposed to tackle the ever-growing plastic waste
problem and help reduce the amount of plastic coupled with organic waste reaching landfills or
incineration facilities. It is essential to understand the degradation behavior of these novel
polymers to guarantee their ultimate biodegradation together with organic waste. Among them,
poly(lactic acid) (PLA), a popular biobased and compostable polymer produced from renewable
sources has garnered much interest due to its low environmental footprint and ability to replace
conventional polymers and be disposed of in industrial compost environments. Although PLA is
industrially compostable when subjected to a suitable set of conditions (i.e., aerobic thermophilic
conditions for an extended period), its acceptance in industrial composting facilities is affected
adversely due to longer timeframes to degrade than the readily biodegradable organic fraction
of waste. So, PLA’s requirement to be fully exposed to thermophilic conditions for prolonged
periods to biodegrade has restricted its adoption and hindered its acceptance in industrial
composting facilities. This dissertation proposes three different approaches to improve PLA
biodegradation under mesophilic conditions to open new avenues of biodegradation, such as
home/backyard composting and guaranteeing industrial composting biodegradation at similar
times as that of readily biodegradable materials.
For the first approach, a reactive blend of PLA with thermoplastic starch (TPS) was
produced and evaluated for biodegradation under mesophilic (37°C) and thermophilic (58°C)
conditions. Films were tested for biodegradation by analysis of evolved CO2 for 180 days in
simulated composting conditions in an in-house built direct measurement respirometer (DMR).
The films' average molecular weight (Mn) and crystallinity (Xc) were tracked throughout the test
duration, and the kinetic degradation rate was calculated. The introduction of TPS positively
affected accelerating PLA hydrolysis during the lag phase in both mesophilic and thermophilic
conditions due to increased chain mobility, resulting in faster degradation of PLA at both
biodegradation conditions.
The second approach involved the introduction of biostimulants in compost to target
different stages of biodegradation and enhance the enzymatic activity of microorganisms.
PLA and biostimulants, Fe3O4 nano-powder, skim milk, gelatin, and ethyl lactate were
introduced into the compost media at 37°C. The CO2 evolution, Mn, and Xc of PLA, PLA added
with single biostimulants, and PLA added with a combination of biostimulants were
evaluated to investigate the degradation of PLA. To attain Mn values of ≲10 kDa for PLA,
PLA added with skim milk experienced a biodegradation acceleration of 15%, 25% with
gelatin, and 22% with ethyl lactate. Fe3O4 enhanced the biodegradation rate by 17% whereas
the combination of gelatin and Fe3O4 resulted in a substantial increase of biodegradation rate
of around 30%.
The last approach explores the use of enzymatic pretreatments. PLA films were
pretreated with proteinase K enzyme at 37°C for 7 and 10 days and at 58°C for 2 and 5 days.
These films were later introduced in inoculated vermiculite at 37°C and 58°C in the DMR to
investigate the effect of pretreatment by simulating home and industrial composting settings.
The results showed a higher CO2 evolution and visible degradation for PLA films pretreated
with proteinase K compared to the untreated control PLA films.
This dissertation presented three innovative methods to speed up PLA film
biodegradation in composting. It provides potential solutions to remove the barrier for
degrading PLA in home and industrial composting conditions and to help address the plastic
pollution challenge by effectively degrading biodegradable polymers with organic waste.
Copyright by
POOJA C MAYEKAR
2024
ACKNOWLEDGEMENTS
When it comes to crafting a thesis, the acknowledgment section stands out as one of
the most daunting tasks. It brings to mind a quote from my advisor, Dr. Rafael Auras, who
often says, "It's never about the destination, but it's about the journey," and indeed, what an
extraordinary journey it has been.
I want to kick off my acknowledgments by expressing my deepest appreciation to Dr.
Rafael Auras for his unwavering support, guidance, and belief in me throughout this
endeavor. Not only has he been an exceptional advisor, but he has also served as a
remarkable role model, a guide, and a mentor who always showed faith in me, guiding me
towards the right path. From granting us the autonomy to work at our own pace to caring
about our well-being as graduate students by checking if we've had enough rest, he has gone
above and beyond the call of duty.
I also extend my heartfelt gratitude to my committee members, Dr. Maria Rubino, Dr.
Eva Almenar, and Dr. Ramani Narayan (College of Engineering, Michigan State University),
for their invaluable support and insightful feedback during my research journey.
No research endeavor can succeed without access to proper laboratory equipment and
a competent lab manager. Therefore, I would like to acknowledge Mr. Aaron Walworth
(School of Packaging, Michigan State University) for graciously allowing me to utilize the lab
resources. He is always ready to help and teaches things with a smile no matter how many
times it is needed.
Additionally, I am immensely grateful to my RAA research group members:
particularly Dr. Anibal Bher, and Dr. Wanwarang Limsukon for being my friends and
colleagues and for their unwavering guidance and support.
I would also like to express my gratitude to my SOP friends for their constant
encouragement, transforming into a family away from home. A special mention goes to my
v
partner Dr. HenryManoj Dsouza for his caring nature, encouragement, for creating a safe
space for me, and for his endless support through the most difficult journey. I am truly
grateful to my best friend Kajal Rajwal for donning so many hats – sister, soulmate, unpaid
therapist, and more. I would like to extend my heartfelt appreciation to Dr. Kaustubh Rane
for all the love, support, and companionship, for making me laugh and helping me find my
spark back. You three inspire me every day to keep going and always have my back. I am
profoundly grateful for all the moments we have shared and thank you for being my rock and
for believing in me. A special mention to my oldest friends, Dhruv Ghiya & Amey Talekar for
endlessly listening to me go on about my research and keeping it light and fun when I needed
it.
Finally, none of this would have been possible without the love and support of my
family, my father Chandrashekhar Mayekar, my mother Chhaya Mayekar, my brother
Prasad Mayekar, and my fur baby my loyal companion Rhea (I miss you very dearly). I am
deeply thankful to the Universe for granting me the strength and blessing me with the
opportunity to be a Spartan! and aiming for the impossible.
vi
TABLE OF CONTENTS
LIST OF ABBREVIATIONS .................................................................................................. viii
CHAPTER 1: INTRODUCTION ............................................................................................... 1
REFERENCES ............................................................................................................... 6
CHAPTER 2: LITERATURE REVIEW ..................................................................................... 8
REFERENCES ........................................................................................................... 153
CHAPTER 3: BREAKING IT DOWN: HOW THERMOPLASTIC STARCH ENHANCES
POLY(LACTIC ACID) BIODEGRADATION IN COMPOST – A COMPARATIVE
ANALYSIS OF REACTIVE BLENDS .................................................................................. 205
REFERENCES ........................................................................................................... 227
APPENDIX 3A: MATERIAL PROCESSING ............................................................ 234
APPENDIX 3B: PHYSICOCHEMICAL CHARACTERISTICS ................................ 236
APPENDIX 3C: MORPHOLOGICAL CHARACTERIZATION OF PLA-g-TPS
REACTIVE BLEND ................................................................................................... 237
APPENDIX 3D: ROUGHNESS AND CONTACT ANGLE MEASUREMENTS ...... 238
APPENDIX 3E: APPLICATION OF THE TIME-TEMPERATURE SUPERPOSITION
PRINCIPLE FOR PREDICTING HYDROLYTIC DEGRADATION IN MESOPHILIC
CONDITIONS ............................................................................................................ 240
APPENDIX 3F: CRYSTALLINITY MEASUREMENTS .......................................... 242
CHAPTER 4: ACCELERATING BIODEGRADATION: ENHANCING POLY(LACTIC
ACID) BREAKDOWN AT MESOPHILIC ENVIRONMENTAL CONDITIONS WITH
BIOSTIMULANTS ................................................................................................................ 244
REFERENCES ........................................................................................................... 266
CHAPTER 5: SPEEDING IT UP: DUAL EFFECTS OF BIOSTIMULANTS AND IRON ON
BIODEGRADATION OF POLY(LACTIC ACID) AT MESOPHILIC CONDITIONS .......... 272
REFERENCES ........................................................................................................... 291
APPENDIX 5A: PHYSICOCHEMICAL CHARACTERISTICS ................................ 297
APPENDIX 5B: CO2 EVOLUTION AND MINERALIZATION OF PLA IN THE
PRESENCE OF SKIM MILK, GELATIN, AND ETHYL LACTATE ....................... 298
APPENDIX 5C: CO2 EVOLUTION AND MINERALIZATION OF PLA IN THE
PRESENCE OF FE3O4 NANOPOWDER, SKIM MILK + FE, AND ETHYL
LACTATE + FE .......................................................................................................... 304
CHAPTER 6: ENHANCING BIODEGRADATION OF POLY(LACTIC ACID) IN
MESOPHILIC AND THERMOPHILIC ENVIRONMENTAL CONDITIONS: THE ROLE
OF PROTEINASE K AS A PRETREATMENT .................................................................... 308
REFERENCES ........................................................................................................... 320
APPENDIX 6A: PHYSICOCHEMICAL CHARACTERISTICS ................................ 322
CHAPTER 7: CONCLUSIONS AND RECOMMENDATIONS FOR FUTURE WORK....... 323
vii
BA
BOD
BT
CO2
EFP
EPS
LIST OF ABBREVIATIONS
bonds of 1,4-butanediol of adipic acid
biochemical oxygen demand
bonds of 1,4-butanediol and terephthalic acid
carbon dioxide
environmental footprint
extracellular polymeric substances
GHG
greenhouse gas
MSW
municipal solid waste
Xc
Mno
Mn
Mw
degree of crystallinity
initial number-average molecular weight
number-average molecular weight
weight-average molecular weight
MWD
molecular weight distribution
OH
hydroxyl group
PBAT
poly(butylene adipate-co-terephthalate)
PBS
poly(butylene succinate)
PBSA
poly(butylene succinate-co-adipate)
PBSe
poly(butylene sebacate)
PBSeT
poly(butylene sebacate terephthalate)
PBST
poly(butylene succinate terephthalate)
PBT
poly(butylene terephthalate)
PDLA
poly(D-lactide)
PDLLA
poly(D, L-lactide)
PE
poly(ethylene)
viii
PEA
PES
PET
PGA
PHA
PHB
PHU
PLA
poly(ethylene adipate)
poly(ethylene succinate)
poly(ethylene terephthalate)
poly(glycolic acid)
poly(hydroxyalkanoates)
poly(hydroxybutyrate)
polyhydroxyurethane
poly(lactic acid)
PLLA
poly(L, L-lactide)
PS
PU
polystyrene
polyurethane
PVOH
poly(vinyl alcohol)
SEC
SUP
TCA
THF
TPS
UV
VOC
size exclusion chromatography
single-use plastic
tricarboxylic acid
tetrahydrofuran
thermoplastic starch
ultraviolet radiation
volatile organic compounds
ix
CHAPTER 1: INTRODUCTION
1.1
Background and motivation
The last two decades have seen a significant revolution in the plastic industry
concerning the fate of polymers in the environment post-consumer use [1]. As the global
production of plastic increases, so does the attention towards its disposal and waste
management scenario [2]. The current problem of plastic waste finding its way to oceans and
remote locations, such as the alps, arctic has revealed the lack of waste management systems
and policies targeting their responsible use and consumption [3–5]. The growing concern and
environmental awareness of the problem of white pollution have been a driving force to
develop green, biodegradable, recyclable, or compostable polymers with a low environmental
footprint [6,7]. Replacing the fossil-based polymers with biodegradable polymers such as
poly(lactic acid)
(PLA), thermoplastic starch
(TPS), polycaprolactone
(PCL), and
polyhydroxyalkanoate (PHA) is perceived as one of the many solutions to tackle the plastic
pollution problem [8,9]. Implementing these polymers can expand their commercial market
and help divert organic waste from reaching landfills.
PLA is a bio-based polymer that can be synthesized from the fermentation of dextrose
obtained from crops such as corn, sugarcane, and cassava [10,11]. PLA is compostable in
nature and can be biodegraded under the right conditions to produce carbon dioxide, water,
and biomass, which is cycled back and non-toxic to the environment. In addition, PLA
displays a moderate barrier to gas and flavor, is easily processed using extrusion, has high
clarity and good stiffness equivalent to traditional polymers such as polystyrene [12].
PLA has found its applications in food packaging such as containers and trays [13–
16]. It can be collected with the organic fraction of municipal solid waste (MSW). The organic
collection of MSW can be treated in industrial composting treatment facilities. These organic
composting treatment facilities were created keeping in mind the considerable volume of
1
organic waste that comes from food waste streams. However, they were not designed to
process bioplastics such as PLA, whose degradation rate is lower than readily biodegradable
feedstocks such as food waste.
Since PLA undergoes chemical hydrolytic degradation at temperatures higher than
its glass transition temperature (Tg ~ 58°C), which has been reported as the primary
controlling degradation mechanism, this brings a considerable change in the mechanical
properties and reduction in molecular weight to a point where it can be easily assimilated by
microorganisms present in the given environment [13]. PLA shows a more extended
biodegradation lag phase at lower temperatures and degrades at a slower rate than other
readily biodegradable materials such as starch and cellulose. This dependence of PLA on
degrading at elevated temperatures reduces its disposal scenario to industrial composting
and limits degradation on home or backyard composting [14,15].
Furthermore, since industrial composting facilities handle large volumes and the
turnaround time can be faster than the current standard requirements, PLA may not be
wholly degraded at the end of the composting process [16]. Any variation in the process
related to the frequency of turning the compost pile, uniform mixing to aerate the piles,
maintaining optimal temperature range, and avoiding the formation of pockets of anaerobic
conditions through the compost heaps can easily derail the biodegradation of PLA, thereby
leaving half degraded or at times whole packages of the same by the end of the composting
cycle [17]. This defeats the purpose of compostable polymers and produces low-quality,
contaminated, and unmarketable compost (soil conditioner and fertilizer). Overall, this
negative effect has discouraged composting facilities from accepting biodegradable polymers
like PLA [15,16,18].
Since biodegradation of PLA is a complex phenomenon and involves the action of
extracellular enzymes released by the microbes at the same time, researchers have studied
2
and advocated that apart from chemical hydrolysis, the action of the hydrolases class of
enzymes plays an essential role in the breakdown of the high molecular weight PLA by
severing the backbone chain [19–23]. Since PLA is marketed as a compostable polymer,
inspecting different paths for improving PLA degradation in composting and ambient
conditions is essential. So, enhancing the degradation of PLA under lower temperature
conditions such as mesophilic range can open new avenues to dispose of PLA in-
home/backyard composting and provide some assistance towards guaranteeing the
degradation at industrial composting facilities.
Biostimulation, which is the addition of selective compounds in the degradation
environment, is one way of inducing enzymatic degradation and accelerating the degradation
of PLA. Researchers have studied the prospective of biostimulation by introducing different
groups of nutrients and enhancing the degradation activity. Modifying the PLA structure
through a process such as blending PLA with other biodegradable polymers to tailor its
properties such as hydrophobicity, tensile strength, and elongation at the break while still
retaining/ improving the biodegradability can be another option to advance its slow
degradation rate. Finally, bioaugmentation is another favorable option that involves
introducing specific microbial strains known to degrade PLA in the current environment and
boosting its degradation rate. These three methods, modification of PLA structure (blending),
biostimulation, and bioaugmentation, can open the venue to make PLA easily compostable
in industrial facilities and provide an opportunity for home composting.
1.2 Overall goal and objectives
This dissertation aims to elucidate the degradation process of PLA at mesophilic
temperatures and investigate different pathways to accelerate the aerobic biodegradation of
PLA in simulated composting conditions. Three objectives have been outlined to accomplish
this goal:
3
1) To modify PLA structure by blending PLA with other biobased biodegradable
polymers to increase its biodegradation.
2) To assess and understand the effect of adding biostimulants to enhance the
biodegradation of PLA by altering different steps involved in the biodegradation process.
3) To investigate the effect of pretreating PLA with proteinase K enzyme prior to
introducing the films in a compost environment.
1.3 Dissertation overview
This dissertation is organized as follows.
The current chapter, Chapter 1, details the background and motivation for this
dissertation study, including the main goal and the specific objectives outlined to accomplish
this goal.
Chapter 2 lays out an extensive literature review regarding the main polymer
classification, abiotic and biotic polymer degradation mechanism, mesophilic environments,
polymer properties affecting the biodegradation rate, different assessment techniques to
measure biodegradation, and polymers that can be blended with PLA to improve its
properties and biodegradation. The chapter also covers the main class of enzymes active in
PLA biodegradation and concludes by highlighting the biodegradation pathway of PLA and
other biodegradable polymers.
Chapter 3 investigates the effect of reactively blending thermoplastic starch with PLA
and compares biodegradation at two different simulated composting settings: industrial
(thermophilic temperature range ~ 58°C) and home/backyard (mesophilic temperature range
~ 37°C). The study explores how the addition of thermoplastic starch improves PLA
biodegradation.
Chapter 4 investigates the biostimulation technique to improve PLA biodegradation
at mesophilic conditions. Different biodegradable biostimulants were added to the compost
4
environment to stimulate the native microbial population and target different steps involved
in the biodegradation process.
Chapter 5 explores the effect of a combination of biostimulants on PLA biodegradation
when introduced in the compost environment. The combination of biostimulants targets two
different steps (i.e., adding compounds to trigger enzyme generation and electron
donor/acceptor compounds) in the PLA biodegradation process.
Chapter 6 presents the results of the bioaugmentation technique wherein PLA films
are pretreated with proteinase K enzyme and subjected to industrial and home composting
simulated degradation setting. A comparative analysis of PLA degradation at two different
temperatures, 37°C and 58°C, for two different pretreatment time intervals was investigated.
Chapter 7 summarizes all the works in this dissertation and concludes with future
work recommendations.
5
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7
CHAPTER 2: LITERATURE REVIEW
2.1
Abstract
Finding alternatives to diminish plastic pollution has become one of the main
challenges of modern life. A few alternatives have gained potential for a shift towards a more
circular and sustainable relationship with plastics. Biodegradable polymers, derived from
both bio- and fossil-based sources, have emerged as one feasible alternative to overcome
inconveniences associated with the use and disposal of non-biodegradable polymers.
Biodegradation of biodegradable polymers depends on the environments, abiotic and biotic
factors, and the polymer bulk and surface properties, resulting in a plethora of parameters
that create a complex process whereby biodegradation times and rates can vary immensely.
The intent of this review is to provide background and a comprehensive, systematic, and
critical assessment of the factors affecting the biodegradation of biodegradable polymers,
with special focus on the mesophilic range. To accomplish this goal, the literature on
biodegradable polymers since 1990 was assessed to create a holistic understanding of the
main proxies responsible for biodegradation such as abiotic and biotic mechanisms of
degradation, environments, factors, microbial populations, and polymer properties. Insights
gained and remarks for potential future research are provided with focus on mesophilic
aerobic environments and the main biodegradable polymers produced.
2.2
Introduction
Plastics are pervasive and have become an indispensable part of our everyday life.
The nature of plastics and their easy processability, durability, low cost, and availability
favor their use, opening up an array of opportunities in market segments such as consumer
goods, food and medical packaging, plastic films and pots for the agriculture sector,
construction, and automotive parts [1,2]. Between 1950 and 2020, global plastic production
8
reached an accumulated amount of c. 9,5001 million metric tons [1,3]. With annual production
of c. 370 million metric tons in 2020, estimates for 2030 are c. 6002 million metric tons. The
global value of the plastic industry in 2020 was around USD 580 billion [4,5], and is expected
to reach USD 8003 billion by 2030. Flexible and rigid single-use plastic (SUP) represents c.
40% of the total global plastic production [1].
The benefits of plastics and SUPs are numerous, and include lower water and energy
consumption during production and use than glass and metal, which in turn reduces the cost
of resources used [6]. However, the ability of plastics to persist, even in harsh environments,
has led to white pollution (i.e., leakage and accumulation of plastics in the environment).
SUPs have been blamed as one of the main offenders for white pollution and are a growing
concern for our modern society since increasing amounts end up in landfills as a portion of
municipal solid waste (MSW), as litter on land and in drainage systems worldwide, and
ultimately leaking into rivers and oceans [7–9]. At present, c. 8 million metric tons of plastic
end up in our oceans annually, in addition to the 150 million metric tons that are already
circulating in marine environments since the dawn of the plastic era [10,11]. A recent
prediction, if business continues as usual without mitigation measures, is that c. 90 million
metric tons of plastic waste will reach the world’s aquatic environments by 2030 [12].Plastics
ending up in the environment mostly start as macromolecular structures formed as films,
bottles, trays, etc. (macroplastics) and then break down into smaller fragments called
microplastics, which are formed due to mechanical abrasion, radiation light, and heat, and
1 Estimations were obtained from references [1,3]. Results are based on production estimated
from reference [3] until 2015 and the addition of production for the 2016–2020 period from reference
[1].
2 Estimation was obtained based on a linear projection growth rate from 2006–2018 from each
global region from references [1,3] and extrapolated to 2030.
3 Estimation obtained based on a linear global projection growth rate from 2016–2020 from
references [4,5] and extrapolated to 2030.
9
can even be reduced to nanoplastics. Microplastics are a concern due to the ability to
concentrate pollutants and become a channel for bioaccumulation, while nanoplastics are
also a health concern since they can potentially translocate in cell membranes of living
organisms and become a source of transporting toxic chemicals [13–15].
Most of the plastic waste in the ocean comes from land-based sources, such as
agricultural soils, open dumps and industries, or mismanaged plastic waste from land litter
and incomplete collection, and then ending up in sludge, sewage, and polluted streams,
finding its way through river pathways and leading to global marine pollution [11,16,17].
Apart from rivers [16], wind and snow have also been identified as responsible for
transporting airborne plastic debris to locations perceived uninhabitable and remote such as
the polar regions and the French and Swiss Alps [18,19]. So, plastic pollution has called
attention worldwide in the form of a global crisis leading to ecological imbalance [7,20].
A consumer paradigm shift about plastics has occurred due to the growing amount of
unmanaged disposal of flexible SUPs, pushing industries to embrace the long-term circular
economy of plastics [21–23]. As part of this circular economy, new challenges have been
highlighted such as novel policies targeting responsible consumption, a push for worldwide
waste management infrastructure creation to recover plastics, and the development,
production and use of highly recyclable or biodegradable plastics with lower environmental
footprint (EFP) [24–26]. Novel policies targeting responsible consumption have been
developed, such as the 2030 Agenda for Sustainable Development by the United Nations
establishing the seventeen Sustainable Development Goals (SDGs) to achieve a better and
more sustainable future for all [27]. Specifically, Goal 12 stipulates sustainable consumption
and production, which has been adopted by countries around the world to create novel policies
about use of materials such as plastics [28]. For example, various U.S. states have established
“extended producer responsibility” for packaging and have banned plastics bags [29–31].
10
Furthermore, bans or extra fees for some SUPs are already effective in the European Union
and countries in Asia such as China and Indonesia [32–34], and are in development in New
Zealand and Australia [35].
The need for worldwide waste management infrastructure has been noted: in 2016,
the world generated c. 2 billion metric tons of MSW, and is expected to generate c. 2.6 billion
metric tons of waste by 2030 if no measures are taken to curb the growing generation of waste
[36]. High and upper-middle income economies generated 66% of the global MSW in 2016,
representing c. 1.34 billion metric tons, and are expected to generate c. 1.6 billion metric tons
by 2030; lower-middle and low-income economies generated 34% of global MSW in 2016,
representing c. 0.7 billion metric tons, and are expected to generate 1.0 billion metric tons by
2030 [36]. Inadequate or scarce waste management solutions are prevalent in lower-middle
and low-income economies. In 2016, high and upper-middle income economies had MSW
collection rates of 96% and 82%, respectively, in urban areas; whereas lower-middle and low-
income economies had collection rates of 51% and 39%, respectively, in urban areas [36].
Therefore, as economies move from low and lower-middle to upper-middle and high-income,
MSW generation will increase, and waste management systems and collection must improve
accordingly. Concentrated efforts are being directed to improve material recovery facilities
around the world, with special emphasis on the lower-middle and low-income economies
[37,38]. However, without MSW collection, white pollution will not be deterred.
To tackle the problem of plastic pollution, cradle approaches to deal with the
production of highly recyclable and biodegradable polymers with lower EFP are increasingly
being considered [39,40]. Replacing fossil-based plastics with bio-based plastics is one
strategy shown to reduce the greenhouse gas (GHG) emissions produced by plastics [41–43].
For example, substitution of c. 66% of the world’s fossil-based plastics with bio-based plastics
has been estimated to reduce GHG emissions by 241 to 316 Mt CO2-eq. per year [44]. To date,
11
mechanical recycling has been the main recovery method for polymers such as poly(ethylene
terephthalate) (PET) and poly(ethylene) (PE). However, novel chemical upcycling methods
for polymers are being explored, which entail the use of rejected plastic (waste) and
converting it to a high-value resource in the form of fuels, chemicals and novel polymers, thus
avoiding the buildup of plastic and reducing the use of fossil fuels [45]. Most of these new
recovery methods are also being explored for flexible SUPs [46].
Among the cradle approaches to deal with littering and recovery of plastics, the
production of biodegradable polymers is a promising solution, primarily since they can be
recovered by traditional waste management options, including mechanical and chemical
recycling and energy recovery, and the additional recovery route through aerobic industrial
and home composting or anaerobic digestion. If enough volume of isotropic biodegradable
polymers is collected and treated, they can be recycled. However, if volumes collected are low
and/or the materials are contaminated, they can be routed towards biodegradation recovery
scenarios such as industrial and home composting, with soil biodegradation being a special
route for agriculture films.
The efficacy of biodegradation is conditioned by drastically different environmental
conditions, such as heat, humidity, and acidic or alkaline media, and by the polymer
characteristics such as chemical structure and physical properties.
Previous reviews on the biodegradation of polymers in some of these environments
and conditions have focused on biodegradable polymers in general [47–49], biodegradable
polyesters [50,51], and mechanisms of degradation [52,53]. Furthermore, recently works have
reviewed and identified gaps and research needs in this area [54,55]. This comprehensive
review expands on these previous works, and aims to provide an overview of the mechanisms,
environments, and factors affecting the biodegradation of biodegradable polymers, giving
special attention to the mesophilic degradation range (20 - 45℃). The specific goals are:
12
1) to provide a transdisciplinary background on the aspects affecting the
biodegradation of biodegradable polymers,
2) to describe the different methods used for assessing biodegradation at mesophilic
temperatures, and
3) to provide insights on the degradation pathway followed by polymers susceptible to
biodegradation.
2.3
Bio- and fossil-based biodegradable polymer classification
Figure 2.1 provides a general classification of polymers according to their feedstock
source and their ability to experience biodegradation. The first group of polymers are bio-
based in nature and non-biodegradable, such as bio-based poly(ethylene) (Bio-PE) and bio-
based poly(ethylene terephthalate) (Bio-PET). The second group of polymers are bio-based
and biodegradable, such as poly(lactic acid) (PLA), poly(hydroxyalkanoate)s (PHAs),
cellulose, and starch. The third group includes polymers that are derived from fossil-based
sources, but also present biodegradable characteristics, such as poly(butylene adipate-co-
terephthalate) (PBAT) and poly(caprolactone) (PCL). The fourth and final group corresponds
to the conventional group of polymers that are derived from fossil-based sources and are non-
biodegradable, such as low- and high-density PE (LDPE, and HDPE) and PET. This
classification is very general since the characteristics of the material, the environment, and
the rate of biodegradation for polymers vary widely among these groups.
As shown in Figure 2.1, biodegradability of a polymer in regular environmental
conditions is not strictly related to the source of the polymer, and factors such as its chemical
structure and physical properties are essential [56]. Some bio-based polymers, such as bio-
PE and bio-PET, are more difficult to degrade than their fossil-based counterparts (i.e., PE
and PET); whereas some fossil-based polymers, such as fossil-poly(butylene succinate) (PBS)
13
and PBAT, are as easily biodegradable as some bio-based polymers, such as bio-PBS and
PHAs, when assessed under standard conditions [57,58].
Figure 2.1 Classification of polymers considering their bio-based or fossil-based feedstock
and ability to be biodegradable or non-biodegradable in environments as compost, soil, and
aquatic; and carbon cycle of bio-based and fossil-based polymers; adapted from [59,60].
Considering the carbon used to produce polymers, the main benefits of biodegradable
polymers can be obtained when the polymers are produced from renewable resources since
they can replenish the carbon cycle (i.e., the times needed to create them and to convert them
to biomass are equivalent) (Figure 2.1). Fossil-based polymers can also be considered
renewable like the bio-based polymers, but the main difference between both is the amount
of time needed to convert to biomass and then back to their original form. Biodegradable
polymers produced from bio-based resources take far less time to be converted to biomass
whereas the fossil-based polymers take millions of years to achieve the same. The longer time
frames are due to the imbalance between the rate of consumption and the replenishment
14
rate, which further leads to mass imbalance in the carbon cycle. There is no additional carbon
footprint associated with renewable-carbon feedstock used to produce biodegradable
polymers, such as starch-heavy crops not intended for human consumption (e.g., starches
from field/feed corn), due to quite similar time frames for consumption and conversion to
biomass [43,60,61].
2.4
Abiotic and biotic polymer degradation mechanisms
Polymer degradation is defined as an irreversible change of the chemical structure,
physical properties, and visual appearance due to the chemical cleavage of the polymer’s
constitutive macromolecules by one or more mechanisms acting concurrently [49]. More than
one mechanism can simultaneously take place due to the action of external factors, and one
mechanism can be more dominant than others at any time [48]. External factors associated
with the environment, such as heat, humidity, radiation, and acidic or alkaline conditions,
could modify the degradation process and its rate. The degradation process can alter polymer
properties such as mechanical, optical, electrical, discoloration, phase separation or
delamination, erosion, cracking and crazing [49]. The four main abiotic mechanisms
associated with polymer degradation are mechanical, thermal (or thermo-oxidative), photo
(photo-oxidative), and hydrolytic (chemical) degradation, some of which can be assisted by
catalysis. In addition, ozone degradation (chemical) is considered a mechanism of
degradation for polymers but is less common (Figure 2.2). Mechanical and thermal
degradation can occur at the early stage of polymer processing. Photodegradation is observed
when polymers are exposed to radiation like ultraviolet (UV) light or gamma rays. Abiotic
hydrolytic (chemical) degradation is considered the most significant mechanism for
biodegradable polymers such as polyesters. These abiotic mechanisms of degradation can be
combined, accelerating the final biotic enzymatic process, for example, increasing the
15
exposed surface area for interaction with the microbial population [48]. We will be discussing
hydrolytic and biotic degradation in detail below.
Figure 2.2 Main abiotic and biotic mechanisms of polymer degradation.
2.4.1 Hydrolytic degradation
Chemical hydrolytic degradation or abiotic hydrolysis is one of the main abiotic
degradation mechanisms for biodegradable polymers, especially for aliphatic and
aliphatic/aromatic polyesters. In this review, we refer to this mechanism also as chemical
hydrolysis. With the uptake of water, susceptible chemical bonds in polymers can undergo
chain scission, resulting in a reduction of Mw, loss of mass and mechanical properties, and
increased surface area of the polymer, thereby increasing the available sites for attack by
enzymatic activity, the biotic step initiated by microorganisms [48,52,62].
Chemical hydrolysis proceeds via two mechanisms when considering the macro
structure: bulk or surface erosion. Depending on the conditions, these mechanisms can occur
independently or combined. Bulk erosion is the dominant mechanism when the water
diffusion is higher than the hydrolysis reaction rate, and surface erosion is dominant when
16
the water diffusion into the polymer matrix or bulk is lower than the hydrolysis reaction rate
[48,50,63].
When bulk erosion is the dominant mechanism, the Mw of the polymer bulk is reduced
so that the polymer loses its mechanical properties in a short period of time. Due to the Mw
reduction and higher mobility of shorter polymeric chain segments, crystallinity may change.
Loss of mass and change/s in geometric shape take more time. The byproducts of bulk erosion
are first accumulated; when the polymer chains are short enough, and reaching n-mers size,
they can start to diffuse out. When the polymer undergoes surface erosion, the loss of mass
is mostly from the surface while the bulk remains intact. As degradation advances, the loss
of mass happens faster at the surface and the polymer gets smaller in size. When compared
with bulk erosion, the mechanical properties and Mw are preserved for an extended period,
and release of byproducts from the surface occurs from the beginning [63].
The kinetic rate of the chemical hydrolysis – surface or bulk dominant – depends on
and can be affected by several factors associated with the polymer itself and the environment.
The roles of some factors are discussed in the next sections and additional information can
be found elsewhere [48,50,63,64].
In terms of environmental factors, increases in temperature and moisture intensify
the rate of chemical hydrolysis [65]. Polymer chain mobility increases as the temperature
increases. Hence, the susceptibility of hydrolysable bonds to undergo chain scission increases.
The chemical potential of water on the surrounding media plays a significant role in
hydrolysis of polymers [66]. Hydrolysis in acidic or alkaline conditions can occur through
different mechanisms so the byproducts of the reactions can differ [67]. Finally, catalysts can
modify the rate of the hydrolytic process [68,69].
In terms of polymer factors, hydrophilic polymers are more susceptible to hydrolytic
degradation than are hydrophobic polymers [48]. Hydrolysis depends on the presence of
17
hydrolysable covalent bonds, such as esters, ethers, anhydrides, carbamide (urea), and ester
amide (urethane), which increase the rate of chemical hydrolysis [52,70]. Table 2.1 compares
the half-lives of hydrolysable bonds in various polymers and shows that poly(anhydride)s are
subjected to rapid hydrolysis due to the presence of hydrolysable bonds of very low half-life.
By contrast, polyamides are very resistant to hydrolysis due to the resistance of the amide
bonds to hydrolysis. The kinetics of the hydrolysable bond half-life presented in Table 2.1
can increase or decrease due to the influence of neighboring groups.
The presence of amorphous regions increases the chemical hydrolysis rate due to easy
diffusion of water into the polymer matrix, compared to semi- and crystalline polymers
showing well-organized structure where diffusion is limited, even at temperature higher than
Tg [71–73]. So, for polymers with lower or similar values of Tg than the mesophilic range, the
diffusion likely is mostly controlled by the amorphous region where chemical hydrolysis is
dominant.
Table 2.1 Half-lives of hydrolysable bonds (in water at pH 7 and 25 ºC). Adapted from [74].
Polymer
Chemical structure
Half-life*
Poly(anhydrides)
Poly(ketal)
Poly(ortho esters)
0.1 hours
3 hours
4 hours
18
RCOOCOnOCORR''R'nOOORn
Table 2.1 (cont’d)
Poly(acetal)
Poly(ester)
Poly(urea)
Polycarbonate
Polyurethane
Polyamides
0.8 years
3.3 years
33 years
42,000 years
42,000 years
83,000 years
*Half-life: time required for 50% hydrolysis in water at pH = 7 and 25 °C for the low Mw (methyl, ethyl)
model compounds.
The macro structure properties, such as size and shape of the polymer, are factors
that condition whether the dominant mechanism will be either surface or bulk erosion. In
this way, a material can go from surface to bulk erosion when its thickness is reduced to a
value lower than a critical value, called critical sample thickness (Lcrit) [64,75].
Polyesters that contain ester groups are degraded mainly by chemical hydrolysis.
Bulk degradation is predominant for aliphatic polyesters, such as poly(glycolic acid) (PGA),
PLA, PCL, and PBS. The main stages of the hydrolytic degradation of polyesters undergoing
bulk erosion can be summarized as 1) diffusion of water in the polymer matrix (amorphous
regions); 2) water reacting with random ester linkage to produce shorter chains; 3)
19
OCORR'HnCCOCH3HOnNHNHRNHNHR'OOnOCOORnRNCOHOnNCOORnH
autocatalysis due to the presence of acid chain ends in the medium; and 4) release of water-
soluble oligomers and monomers creating a void core and subsequent reduction in Mw [48,76].
The duration of the chemical hydrolysis process depends mainly on the initial Mw,
crystallinity, temperature, and pH [53].
PLA is an example for chemical hydrolysable polymer degradation that does not
require enzymes to catalyze the hydrolytic degradation. In this sense, the media to which the
material is exposed and factors such as temperature, pH, and moisture play major roles in
delaying or speeding up the hydrolytic degradation rate. In an industrial composting process
(~58 ℃ and ~60% RH), PLA can absorb water and undergo chemical hydrolytic degradation.
However, at lower temperatures, such as in agricultural soil environments (~25 ℃ and ~45%
RH), the rate of chemical hydrolysis is low, increasing the time for the enzymatic hydrolysis
process to start. One of the main differences between bulk and surface erosion mechanisms
can be recognized in the diffusion of the degradation byproducts. During bulk degradation of
polyesters, these hydrolysis-formed oligomers and monomer byproducts, such as carboxylic
acid and hydroxyl groups, are trapped and accumulated inside the bulk, leading to an
autocatalytic degradation that tends to accelerate the degradation kinetics [50,77].
Burkersroda et al. [75] reported that hydrolytic degradation of PLA, evaluated at 37 °C,
follows a bulk erosion mechanism for thicknesses between 0.5 and 2 mm, a core-accelerated
erosion for thicknesses between 2 mm and 74 mm, and surface erosion for thicknesses greater
than 74 mm. Hoüglund et al. [78] reported that the hydrolysis of 100% PLLA increased upon
the addition of a low percentage of d-Lactide units, due to reduction of the polymer order
structure, showing the effect of tacticity and optical purity on the hydrolytic degradation of
PLA. PLA hydrolysis is delayed in comparison to PGA hydrolysis due to the presence of the
methyl group in PLA that blocks the attack of water to interact with the hydrolysable bonds
20
[48,63]. A review of the hydrolysis of PLA at mesophilic and thermophilic conditions has been
reported by Tsuji [64].
PBAT, due to the presence of an aromatic group in its polyester chain experiences a
lower hydrolytic degradation rate than polyester with only aliphatic units as PLA and PGA
[79]. The presence of the aromatic group reduces chain flexibility, provides less susceptible
bonds, and creates an steric interference effect to the access of the susceptible ester bonds
[80]. The soft aliphatic domain bonds consisting of 1,4-butanediol and adipic acid monomers
(BA) are more susceptible to hydrolysis than the hard aromatic bonds of 1,4-butanediol and
terephthalic acid monomers (BT). In this sense, PBAT displays good biodegradability when
the aromatic moiety concentration is kept below 55 mol% [81]. Kijchavengkul et al. [82] also
demonstrated that the increase of crosslinking on PBAT has a detrimental effect, not only on
chemical hydrolysis but also in enzymatic hydrolysis.
Polymers that undergo surface erosion are desirable when designing medical devices
and for drug release, since the retention of mechanical properties and capacity for a controlled
release of drugs can be achieved by mass loss without compromising the Mw. Polymers that
can mostly undergo surface erosion are polyanhydrides, some poly(ortho esters), and some
polycarbonates [83–85].
2.4.2 Biotic enzymatic degradation
Biotic enzymatic degradation is the mechanism of degradation where microorganisms
break down organic substances through an enzymatic process. The four main stages of biotic
degradation are shown in Figure 2.3. The main outcome of biotic degradation is reduction of
the polymer to small molecules that are utilized by the microorganisms as a source of carbon
and energy, resulting in final products like CO2 and H2O in aerobic conditions or CO2, H2O
and CH4 in anaerobic conditions. Microorganisms like bacteria and fungi are actively
involved in the biodegradation process. These microorganisms have their own optimal growth
21
conditions; for this reason, biotic degradation is a complex process where several factors
associated with the polymer, microorganisms, and the environment come into play [65].
Abiotic chemical hydrolysis degradation, and biotic enzymatic degradation are the two
main processes for the cleavage of polymer bonds and degradation [63]. The biodegradation
process involves several steps, some of them are abiotic, and some are biotic and act
synergistically to decompose the organic matter. Some of the abiotic mechanisms described
above, such as photo, hydrolytic, or even mechanical degradation, can enhance the biotic
degradation process by increasing the surface area for biofilm formation or by reducing the
Mw [53]. However, the dominant mechanism in the biotic degradation process is related to
biotic agents.
The first stage of the biodegradation process, as shown in Figure 2.3, is biofilm
formation. In the second stage, depolymerization, microorganisms secrete extracellular
enzymes. These agents can cleave the molecules by random or end chain scission, reducing
the Mw and resulting in the generation of oligomers, dimers, and monomers. The
susceptibility of a polymer to microbial attack depends on enzyme availability, availability of
sites in the polymer for enzyme attack, enzyme specificity for the polymer, and the presence
of cofactors [53]. In the third stage, bioassimilation, the molecules transported from the
depolymerization stage are assimilated into the microorganism’s metabolism and are used to
produce energy, new biomass, and primary and secondary metabolites. In the final stage,
mineralization, simple molecules such as CO2, CH4, H2O, N2, and different salts from
intracellular metabolites, which are completely oxidized, are released to the environment
[86]. These non-toxic components are redistributed through the carbon, nitrogen, and sulfur
cycles in nature. Some simple and complex metabolites, such as organic acids, terpenes,
aldehydes, and antibiotics, can reach the extracellular surroundings by excretion [52].
22
Figure 2.3 Main stages of the biotic degradation process: (1) biofilm formation, (2)
depolymerization, (3) bioassimilation, and (4) mineralization. Adapted from [87].
The literature has also described these processes as biodeterioration. Biodeterioration
is commonly described as the undesirable degradation of materials by microorganisms, and
it is considered the mechanism responsible for causing several irreparable damages. In this
review, to avoid any confusion regarding the stages of the biodegradation, we will not use
this term to describe any particular process. The term biodeterioration is mostly used to
describe a combined mechanism when microorganisms are in contact with a polymer surface
and the polymer experiences fouling, degradation of leaching components (as additives),
biocorrosion (enzymes and radicals attack additives and polymer backbone), hydration and
penetration in the bulk polymer, and changes in color of the polymer matrix [88,89].
2.4.3 Biofilm formation
Biofilm structure and formation have been identified as the dominant phase of life for
microorganisms on Earth. Studies have shown that microorganisms, in general, live in
aggregates or mixed species rather than as single cells in pure cultures [90,91]. When a
biodegradation process occurs, biofilm formation is considered the first step and a necessary
23
step in the process. However, the formation of a biofilm on a surface does not necessarily
imply that biodegradation will occur [89].
In biofilm formation, a microbial community is established on a surface. These
surfaces, such as metals, sediments, or polymers, can exist in different forms, have different
properties, and different compositions. Biofilms are considered highly sophisticated and
complex synergistic structures originated by the selective attachment of phylogenetically and
functionally diverse communities of bacteria, fungi, protozoans, or algae [92]. The
organization of microorganisms on a surface is specific for the material and dependent on
that material’s surface properties and the environmental conditions. Biofilms can be
developed in solid/liquid and solid/air interfaces [89]. The first step of biofilm formation for
bacteria is the microorganism’s initial attachment to the surface via the cell pole or the
flagellum (within minutes after the first contact with the substrate) (Figure 2.4a). The
initial attachment is a reversible step. The second step of biofilm formation is the
microorganisms’ irreversible attachment to the surface using a glue-like substance and tail-
like structures. The attached microorganisms start producing slimy extracellular polymeric
substances (EPS), formed by proteins, polysaccharides, nucleic acids, lipids, and humic
substances, and develop clusters of cells in contact with each other and with the substrate.
EPS production allows the microbial community to develop a complex structure highly
influenced by the environmental factors, and is the main factor responsible for the adhesion
to surfaces and for the integrity of the biofilm [93]. During this second step, the growth of
microbial communities can occur in a matter of hours. Biofilm maturation occurs in the third
step, when cell clusters embedded in the EPS become mature and layered. A high level of
biofilm maturation is achieved as cell clusters and microcolonies reach their maximum
average thickness in the fourth step. In the final step, as the maturation of the colonies
progresses, the complex structures weaken, detach from the substrate, release, and
24
propagate. This variable sized group of cells can now attach to a different zone of the surface,
or another previously optimally developed biofilm. The detachment step is characterized by
cells evacuating from the interior of the clusters, forming void spaces [90,94].
In the case of fungi population, the development of filamentous fungi biofilm has been
proposed (Figure 2.4b) [95]. The first step, similar for bacteria biofilm, implies the deposition
and adsorption of spores and/or hyphal fragments. The second step implies the development
of a fungal EPS for active attachment to the surface. In the third step a microcolony is formed
with branching of a monolayer hyphal and extension of the EPS for better adherence of the
microcolony to the surface of the substrate. In the fourth step a colony is formed, a hyphal
compacted network is developed, and the maturation of the colony occurs. Finally, in the fifth
step the dispersal or release of new cells takes place. These new cells can start a new cycle.
A detailed discussion of the biofilm formation mechanism can be found elsewhere [96–98].
25
Figure 2.4 a) bacteria biofilm formation, steps: (1) attachment of microorganisms to the
surface using a specialized glue-like substance and tail-like structures, (2) colonization, (3)
growth, (4) maturation, and (5) detachment; b) Fungi biofilm formation, steps: (1)
attachment of microorganisms to the surface using a specialized glue-like substance and
tail-like structures, (2) colonization, (3) growth, (4) maturation, and (5) detachment.
Adapted from [91,95,99].
Table 2.2 Functions of extracellular polymeric substances (EPS) in biofilm formation.
Reproduced with permission from [93].
Function
Adhesion
Relevance for biofilms
Allows the initial steps in the colonization of abiotic and
biotic surfaces by planktonic cells, and the long-term
attachment of whole biofilms to surfaces
Aggregation of bacterial
cells
Enables bridging between cells, the temporary
immobilization of bacterial populations, the development of
high cell densities and cell-cell recognition
26
1235412345ab
Table 2.2 (cont’d)
Cohesion of biofilms
Forms a hydrated polymer network, mediating the
mechanical stability of biofilms, determines biofilm
architecture, and allows cell-cell communication
Retention of water
Maintains a highly hydrated microenvironment for biofilm
microorganisms, leading to their tolerance of desiccation in
water-deficient environments
Sorption of organic
compounds
Allows the accumulation of nutrients from the environment
and the sorption of xenobiotics
Sorption of inorganic ions
Promotes polysaccharides gel formation, ion exchange,
mineral formation, and the accumulation of toxic metal ions
Enzymatic activity
Enables the digestion of exogenous macromolecules for
nutrient acquisition and the degradation of structural EPS,
allowing the release of cells from biofilms
Nutrient source
Electron donor or
acceptor
Provides a source of carbon, nitrogen, and phosphorus
containing compounds for utilization by the biofilm
community
Allows redox activity in the biofilm matrix
Export of cell components Releases cellular material as a result of metabolic turnover
Sink for excess energy
Stores excess carbon under unbalanced carbon-to-nitrogen
ratios
Binding of enzymes
Results in the accumulation, retention, and stabilization of
enzymes through their interaction with polysaccharides
Many of the identified extracellular enzymes, such as oxidoreductases and hydrolases,
are responsible for the degradation of biopolymer substrates. Extracellular enzymes can
break down substrates available on the EPS or from the surface to which the biofilm is
attached; the resulting low molecular mass products are used as a source of carbon and
energy by the microorganisms. Hence, enzymes capable of degrading substrates turn the EPS
into an external digestive system for the microorganism [93]. Extracellular enzymes are key
for the breakdown of water-soluble substrates (e.g., polysaccharides, proteins, and nucleic
acids) or water-insoluble substrates (e.g., cellulose, lipids, and bio- or fossil-based polymers
chains), leading to depolymerization [93,100].
27
2.4.4 Depolymerization
The enzymatic activity that occurs after biofilm formation is the main contributor to
the depolymerization step. Enzymatic activity can occur via a hydrolytic or oxidative route
(Figure 2.5), involving either random or end chain scission [53,65].
The oxidative mechanism is called enzymatic oxidation. In the case of non-
hydrolysable polymers, due to the absence of hydrolysable groups, redox reactions are the
most effective way to break the backbone made of C-C bonds. However, extracellular enzymes
must have redox potentials high enough to allow the electron extraction from non-reactive C-
H or C-C bonds. A high redox potential requirement could be an important obstacle for
ultimate polymer biodegradation [101].
Enzymatic hydrolysis mirrors abiotic chemical hydrolysis. As chemical hydrolysis
progresses, the Mw is reduced and consequently the polymer becomes available for enzymatic
hydrolysis, which starts dominating the depolymerization step. For hydrolysable polymers,
with ester, carbonate or amide groups, the hydrolytic enzymatic degradation by extracellular
hydrolases has been reported and is presented and discussed in further section in detail.
Within the major enzyme classes (Table 2.3), hydrolases (EC 3) and oxidoreductases
(EC 1) are the main groups of enzymes linked to depolymerization.
28
Figure 2.5 Enzymatic hydrolysis or oxidation routes for depolymerization. Adapted from
[101].
Table 2.3 International Union of Biochemistry and Molecular Biology (IUBMB) classification
of enzymes by function of the reactions they catalyze. Adapted from [102].
EC Number
Enzyme class
Reaction
1
2
3
4
5
6
Oxidoreductases
Oxidation-reduction
Transferases
Chemical group transfers
Hydrolases
Hydrolytic bond cleavages
Lyases
Nonhydrolytic bond cleavages
Isomerases
Changes in arrangements of atoms in
molecules
Ligases
Joining together of two or more molecules
The lower activation energy needed for enzymatic hydrolysis of ester linkages, such
as those in aliphatic and semi-aromatic polyesters, appears to facilitate the depolymerization
29
of polyesters in comparison to polyolefins, where non-hydrolysable linkages are present.
However, large differences have been reported in the rates of biodegradation for polyesters
as a function of their morphology and chemical structure. For example, the aromatic
polyester PBT is considered non-biodegradable; however, the copolymer obtained from
terephthalic acid and adipic acid, PBAT, is biodegradable. Besides the presence of the
aromatic ring in both structures as well as hydrolysable bonds, the presence of the adipic acid
component in PBAT improves the flexibility of the polymer structure, making it more
susceptible to attack by extracellular enzymes [51].
Enzymes are macromolecules made up mostly of proteins, which are complex chemical
structures, with high Mw and hydrophilic groups acting as biocatalysts that accelerate the
depolymerization reaction rates by lowering the activation energy of the reaction [103]. The
simplest enzymes consist entirely of amino acids while conjugated enzymes contain a non-
protein component, a cofactor (or co-enzyme) along with a protein component.
Extracellular enzymes are released when optimal conditions are present between the
polymer surface and the attached biofilm. Enzymes bind to a substrate by their active site
and transform the substrate into a product. Figure 2.6 shows the steps of this process. First,
an enzyme binds to its substrate and positions it properly in its active site to catalyze the
reaction. In the second step, the enzyme-substrate complex is formed. In the third step, the
enzyme-substrate complex aligns reactive groups in the substrate and places strain on
specific bonds, reducing the activation energy required for making the reaction to occur. In
the fourth step, the cleaved products are released. Finally, in the fifth step, the enzyme is
ready to begin the catalytic cycle again.
The main factors influencing the susceptibility of a polymer towards microbial attack
by extracellular enzymes are:
30
Enzyme availability. Availability is determined by the type of microorganisms and the
environment.
Available sites on the polymer for enzyme attack. Extracellular enzymes are classified
as exo- and endo-enzymes. Exo-enzymes are responsible for chain end scission, while endo-
enzymes are responsible for random chain scission [104].
Enzyme specificity. Enzymes are known as catalysts of biochemical reactions with
high substrate specificity. This means that an enzyme catalyzes a special reaction with high
efficiency. Therefore, many different reactions catalyzed by different enzymes can run in
parallel simultaneously. The specificity is a function of the three-dimensional structure of
the enzyme [104].
Presence of cofactors. Cofactors are additional chemical groups incorporated to the
structure of the active site of the enzyme to facilitate a biochemical reaction. Cofactors can
be metal ions (e.g., calcium, magnesium, potassium, sodium, or zinc) or co-enzymes (organic
cofactors). A common function of cofactors is to provide a geometric place for the substrate to
bound to the enzyme by maintaining the stability and activity of the enzyme at the active
site [105].
31
Figure 2.6 Catalytic cycle of an enzyme. Adapted from [86].
The priority of extracellular enzymes is obtaining carbon to ensure a supply of
resources. Additionally, the microbial community is able to shift enzyme production between
groups of substrate-specific enzymes and non-specific enzymes, to match substrate
requirements. In other words, enzymes are selectively produced to increase the supply of the
most limiting element and to target the most available substrates [106]. From an energy
point of view, enzyme production is energy intensive. For this reason, microorganisms
produce enzymes at the expense of growth and metabolism when nutrients are scarce.
Furthermore, when available nutrients are scarce, microorganisms can produce adaptive
32
enzymes to obtain resources from complex sources [107]. On the other hand, when assimilable
nutrients are available and abundant, the production of constitutive enzymes may be
decreased [108].
For polymer degradation, depolymerases are the extracellular enzymes secreted by
microorganisms that cleave complex polymeric substrates into oligomers, dimers, and
monomers. The hydrolytic cleavage can be by exo-attack or endo-attack. Exo-attacks occur at
the end of the polymer chain, and the byproducts are oligomers or monomers that can be
assimilated by the cell. Endo-attacks occur randomly along the polymer chain, reducing the
Mw; hence, products are not assimilable without further depolymerization [109,110]. An
important characteristic of extracellular enzymes is that they are too large to penetrate
deeper into the polymer material. For this reason, enzymes can only act on the polymer
surface, making depolymerization by enzymatic activity a surface erosion process [104].
Increasing the surface area can increase the rate of depolymerization by extracellular
enzymes [111]. Fragments small enough to go through the membrane cells as monomers are
transported inside the cell and transformed to obtain energy for the growth process by the
action of intracellular enzymes. Usually these are oxidative enzymes, and the process is
called bioassimilation or assimilation.
2.4.5 Bioassimilation
Bioassimilation is related to the acquisition or uptake of substances for the microbial
metabolic process. Compounds small enough to pass the semi-permeable membrane after the
depolymerization step can be potentially processed by the metabolism of the microorganism
and finally mineralized to CO2 (dissimilation) or be used for biosynthesis of new products
through metabolic pathways (assimilation) (Figure 2.7). In general, the periplasmic space –
the cell membrane – is where the cleavage takes place, and from where oligomers can be
33
transported across the cytoplasmic membrane for further oxidation in the -oxidation cycle.
Oligomers can be internalized with the aid of surfactants produced by microorganisms during
biofilm formation and be used as carbon and energy sources by the action of intracellular
enzymes. The presence of water for the transport of components is a critical factor during the
bioassimilation step [86].
2.4.6 Mineralization
Mineralization, or ultimate biodegradation, refers to the degradation of polymer
fragments to the mineralized components and biomass, plus CO2 and H2O in aerobic
conditions or CO2, H2O, and CH4 in anaerobic conditions (Figure 2.7). Depending on the
polymer composition, other compounds also can be released, including sulphide, sulphate or
sulphite, ammonia, nitrite or nitrate, phosphate or phosphite, chloride, and fluoride. By
measuring the mineralization levels (i.e., CO2 released or evolved), biodegradation rates and
% mineralization can be quantified. Bioassimilated monomers are part of the catabolism
cycle. During this step, organic compounds, such as carbohydrates and proteins, are used as
metabolites of the tricarboxylic (TCA) cycle or Krebs cycle by aerobic and anaerobic respiring
microorganisms to produce energy [93,112]. Insights on external factors affecting
mineralization of biodegradable polymers can be found elsewhere [113].
Figure 2.7 Microbial bioassimilation and mineralization during the polymer biodegradation
process. Adapted from [114].
34
2.5
Biodegradation environments
There are many feasible waste management recovery processes for polymers, ranging
from recycling, biodegradation included as defined by the U.S. Environmental Protection
Agency (EPA), waste-to-energy conversion, and landfill, each with trade-off environmental
impacts. Littering or leakage to the environment must not be considered waste management
processes. Each of these waste management environments present specific conditions that
can tailor the degradation rate of polymers. So, evaluation of degradation of a polymer in
different environments may reveal different rates due to the influence of external abiotic and
biotic factors [115–117].
Heat (temperature) and other factors in the environment, such as aeration,
acidic/alkaline media, and water content, play crucial roles in how degradation of polymers
takes place. Temperature is one fundamental parameter affecting the rate of degradation
and has been studied in detail [76,118–120]. Besides affecting the abiotic mechanisms of
degradation, temperature drives chemical and microbial changes through the different
phases of biodegradation. Microbial activity is temperature dependent since each
microorganism’s optimal population and growth is driven by different temperature regimes.
Marine environments and some lakes and rivers are associated with temperatures in the
psychrophilic range (0–20 °C); however, depending on the geographical region, rivers and
lakes can also reach the mesophilic range (20–45 °C). Soil and home composting
environments are mostly in the mesophilic range, whereas composting under industrial
conditions is mostly conducted in the thermophilic range (45–60 °C). Table 2.4 presents a
summary of the temperature ranges and the main environments where biodegradation
occurs.
35
Table 2.4 Typical temperature ranges and conditions for different environments where
polymers can be subjected to aerobic biodegradation.
Environment
General description
Management
Temperature
range, °C
20–30
Soil/Agricultural
soils
20–45
Home composting
Large scale. Soil structure
(texture, porosity),
moisture, aeration,
radiation
Small scale. C/N ratio,
moisture, aeration, heat, pH
Uncontrolled
Controlled
Controlled
45–60
Industrial
composting
Medium scale. C/N ratio,
moisture, aeration, pH
2.5.1 Soil environments
Soils provide diverse environments where the biodegradation of polymers can take
place. Soil is a typical disposal scenario for biodegradable and non-biodegradable polymers
employed as agricultural mulch films [121,122]. For several decades, non-biodegradable
fossil-based polymers, such as PE, have been employed as mulch films for crops. However, in
the last 15 years, bio-based and fossil-based biodegradable polymers have gained market
momentum since their use can avoid the removal of the plastic film after harvest and reduces
the leakage of plastic debris [123,124]. The study of plastic use in agriculture is known as
“plasticulture” and includes products such as drip irrigation tape, greenhouse covers, hoop-
house covers, silage bags, row covers, hay bale wraps, plastic trays and pots, and mulch films
[125,126].
Soil is a diverse and typical habitat for microorganisms [127,128], and biodegradation
usually takes place in the mesophilic range of temperature. Biodegradation of polymers in
soil is affected by both biotic and abiotic components such as microorganisms, solid particles
(i.e., organic matter and inorganic minerals), water, and air. Solid, liquid, and gaseous phases
make up the soil environment, along with different organisms. The liquid and gaseous
36
phases, water and air, vary with the climatic conditions and human activity, whereas the
solid phase generally is resistant to these activities [127]. The main parameters used to
classify a soil are based on its granularity and porosity due to the amount of clay, sand, and
silt (Figure 2.8). The texture and structure of soils is determined by the relative proportions
of clay, sand, and silt and their relative sizes [129,130]. Silty soils can possess high water
retention capacity, but clay soils possess the highest water retention capacity. Hence, fungal
populations can be supported by a dry, sandy, well-ventilated soil, whereas an insufficiently
aerated clay soil provides habitat for facultative bacteria [131].
Figure 2.8 USDA soil texture classes determined according to the relative proportions of
sand, clay, and silt. Adapted from [129,132].
The chemical and biological properties of soils are characterized by acidic/alkaline
media, cation exchange capacity, organic carbon concentration, and soil respiration [133].
These properties control the formation and activity of the microbial diversity, and the
37
combination of the mentioned factors create habitats where only certain microorganisms
can grow [70]. The distribution of the different particles creates pores of different sizes that
can retain water or surrounding living organic material. The soil connectivity determines the
circulation of nutrients, soluble organic compounds, and water and is ultimately tied to the
pore geometry and network [134]. Thus, the size of the pores is a factor that determines and
helps to explain the spatial separation of living organisms [127].
The biodegradation process occurring in soil environments should consider the surface
layer and underground matrix [135]. The surface layer of the soil is highly affected by abiotic
factors. On the other hand, the underground matrix is associated with the microbial
population and factors for its optimal activity [70]. The factors playing major roles in the
biodegradation process in soil are soil texture and structure, water content, organic matter,
pH, temperature, O2, and sunlight [131].
Water content and retention is a function of the texture and structure of the soil, as
discussed above. While a dry soil encourages the formation of fungal populations, a wet soil
promotes the genesis of bacterial populations [131]. Fungi spread through the soil using
hyphae, which are thin filaments forming the mycorrhizal network. Under dry conditions,
while in search of water and nutrients, the hyphae spread and take different routes. The
fungi continue enlarging this network and bridge the gaps between different small pockets
of water and nutrients, thus enabling survival and growth in soil, where the moisture content
may be low [136].
Microorganisms can adapt to specific ranges of pH values. Thus, the soil pH is a factor
that can limit the growth of microorganisms. Alkaline to neutral pH favors bacterial growth
whereas acid pH favors fungal development [131]. The pH influences the availability of
nutrients and concentrations of trace metals such as zinc, iron, calcium, magnesium, and
phosphorus. Fungi take in these molecules across their membranes by creating a proton
38
gradient; this proton gradient affects the ability to take up the nutrients when exposed to
extreme pH conditions [137]. In acidic media, certain nutrients, such as phosphorus, become
less available and other nutrients like magnesium and aluminum can become more toxic,
thus creating a hostile environment for helpful soil bacteria.
The O2 content of the medium determines whether the microbial population expressed
is aerobic or anaerobic. Soil temperature governs the physical, chemical, and biological
processes in the soil. Changes in soil respiration rate, due to the fluctuation in temperatures
also affect the bioactivity. Microbial activity is inhibited or reduced drastically with lowering
temperatures [125]. Radiation, mostly from UV light, can inhibit the growth of microbial
populations, depending on the intensity of the radiation. The optimal conditions of
temperature, organic matter, aeration and O2, and water content are in the first 30 cm of the
soil layer [129,135].
Agricultural soils can be considered as a particular type of soil environment, and have
been extensively studied in the plasticulture field [123,126]. One of the most studied
applications has been polymeric mulch films, which undergo several steps in biodegradation.
This process involves a period of intense photodegradation when the mulch film starts
crosslinking and eroding, followed by an intensive period of biodegradation [125,138–140].
Figure 2.9 shows a typical life cycle of polymeric mulch films in agriculture soils.
39
Figure 2.9 Biodegradable mulch film cycle, starting from raising the bed and applying
herbicide in spring, to harvesting and the disposal of the films in late fall, and the associated
degradation processes. [141] (Copyright 2008. Reproduced with permission from Elsevier
Science Ltd.).
2.5.2 Home and industrial composting
Home composting could become a common disposal scenario for polymers and polymer
blends used for packaging (food and beverages) [142]. Home composting is garnering interest
since it can be very instrumental in diverting the household organic fraction from going to
landfill [143]. Additionally, as consumers are becoming more aware of plastic pollution, home
composting has also become important as a potential methodology to reduce organic waste
and contaminated packages that cannot be efficiently recovered or diverted through the MSW
40
management system. The US Composting Council describes home composting as the natural
aerobic decomposition of organic wastes or materials, usually in small-scale composters by
“slow-stack” treatment methods where temperatures are in the psychrophilic (0–20 °C) to
mesophilic (20–45 °C) range [144]. As per the US Composting Council, home composting can
also be labeled as “backyard” or “composting at home”. However, the terminology varies in
different geographical regions worldwide since “composting at home” may imply composting
in designed vessels inside the apartment or house [145,146], and “backyard” composting may
refer to uncontrolled composting units outside the house subject to the environmental
conditions.
The typical matrix for home composting includes biowastes, which are generated in
the kitchen, and garden waste such as weeds and leaves (Figure 2.10). Many key factors,
such as temperature, pH, moisture, composter efficiency, substrate, C/N ratio, and microbial
populations, affect the home and industrial composting process [147,148]. Home composting
is a far less controlled process in comparison to industrial composting, since the process takes
place in the backyard and hence on a smaller scale. Usually, it never reaches the high
temperatures in the thermophilic range for long periods of time, as seen in industrial
composting. The small size of the installation, accompanied with difficulties in reaching an
optimum control of factors, result in home composting requiring a longer time to achieve a
mature compost [131]. The material volumes that can be handled and the abundance of
microorganisms are lower for home composting settings. In addition, seasonal changes can
influence “backyard composting” depending on the geographical location, and hence lower
and more variable temperatures are inevitable.
Regardless of whether the composting is done at home, in a community backyard, or
in an industrial facility, the composting process must ensure a succession of microbial
communities (i.e., mesophiles–thermophiles–mesophiles) and the corresponding temperature
41
regimes to operate. These factors are necessary to guarantee the safety and quality of the
compost process and the final product.
One advantage of home composting is that it can be helpful in rural and suburban
areas where collection of organics is limited or there is no infrastructure for industrial
composting [143,149,150].
Figure 2.10 Home composting representation. [60] (Copyright 2008. Reproduced/adapted
with permission from Wiley & Sons, Ltd.).
Industrial composting is a process designed to handle large volumes of yard, food, and
manure waste [60,148,151]. By employing better aeration, moisture control, and higher
temperatures the biodegradation in industrial composting is accelerated significantly in
comparison to natural and home composting processes. The industrial composting process
requires a proper system in place for collection of wastes and a good infrastructure (e.g.,
windrow, aerated static piles, and in-vessel composting) [152]. Biodegradation in industrial
composting takes place mostly in the thermophilic temperature range. Figure 2.11 shows a
representation of an industrial composting process.
42
Figure 2.11 Industrial composting process. [60] (Copyright 2008. Reproduced with
permission from Wiley & Sons, Ltd.).
The industrial composting process follows four main stages. The first stage is the
mesophilic stage (20–45°C), where microorganisms decompose the simplest organic,
degradable substances into CO2 and water in an exothermic reaction. The high amount of
substrate ensures high microbial activity, which leads to the generation of large quantities
of metabolic heat energy that causes the temperature to rise swiftly. The second stage is the
thermophilic stage (45–60 °C) where bacteria and fungi mesophiles become less active and
are replaced by thermophiles. As the temperature rises above 55 °C, microorganisms such as
pathogens are destroyed. For safety reasons, several certifying entities require that the
43
temperature must reach above a certain temperature, such as 55 °C, and remain at that level
for a set period of time, such as 15 days, to ensure that the resulting compost is pathogen free
[153]. Temperatures in some industrial compost facilities during the early stages commonly
reach values of c. 70 °C [154]. Such high temperatures expedite the disintegration process of
high energy carbohydrates and structurally complex molecules. As the disintegration process
comes to an end, there is no longer any supply of these high energy compounds, and the third
stage kicks into action where the mesophiles take over once again. The third stage is a
transition stage from high to low temperature. The final stage, also called “curing” or
maturation, can take several months to result in stabilized compost [131]. The total
composting time varies in systems used worldwide, from two to more than six months; thus,
certified compostable packages can encounter difficulties to fully disintegrate in some
operations [155].
2.5.3 Aquatic environment
Natural aquatic environments (i.e., oceans, rivers, and lakes) unfortunately are
environments where discarded polymers from activities such as fishing and shoreline
recreation are commonly found [11,16,156]; however, these are not formal waste
management scenarios and must not be considered as such. The natural aquatic environment
is a non-desired end-of-life scenario due to the creation of white pollution and a lack of proper
conditions for biodegradation and control of the process due to its complexity [157].
Biodegradation in the aquatic environment can happen in lakes, rivers, and oceans as well
as in reservoirs, and wastewater facility treatments (aerobic or anaerobic); however, our
discussion is focused on the natural aquatic environments.
Geographical considerations of the aquatic environment play an important role in
understanding the presence and flow of plastics. Lakes are generally low-flow environments
and act as a point for accumulation for plastics and microplastics [158]. Rivers are considered
44
the essential route for transporting plastics to the ocean. Considering their proximity to
urban and industrial areas, rivers become an easy access point to the marine environment
with respect to plastic pollution. Plastics are extensively carried out during floods in cities
with poor waste management systems [12]. For the marine environment (seawater), three
main habitats can be considered when addressing the degradation of plastics (Figure 2.12):
the pelagic zone, an illuminated and aerated column of water; the littoral zone, which is the
beach sediment periodically covered by water due to waves or tide; and the sublittoral zone,
which is the seabed interface up to 200 meters in depth that is aerated and photosynthetically
active. The physical and chemical properties of seawater, including the essential nutrients
for living organisms, vary with the depth, latitude, and proximity to land. Because of this
variation, the microbial populations within seawater also vary. Furthermore, the
degradation process of the plastics entering this environment can be altered by agitation and
turbulence caused by ocean currents, salinity, temperature gradients, and solar radiation
among others [159,160]. Biodegradation of polymers in aquatic environments is described in
terms of scarce evolution for synthetic biodegradable polymers. However, high efficiency has
been reported for natural polymers as cellulose, starch, and PHAs regardless of the low
temperatures reached (Table 2.9). The “plastisphere,” the development of biofilms on the
surface of polymers present in water, has been extensively studied to elucidate the main
components and behavior of microorganisms during the colonization and depolymerization of
polymers in aquatic environments [161–163].
45
Figure 2.12 Main habitats to consider when addressing the degradation of polymers in the
marine environment. The pelagic zone, an illuminated and aerated column of water; the
littoral zone, which is the beach sediment periodically covered by water due to waves or tide;
and the sublittoral zone, which is the seabed interface up to 200 meters in depth that is
aerated and photosynthetically active. Adapted from [164].
2.6
Factors and properties that affect degradation rate
The rate of polymer degradation is affected by the degradation mechanisms, the
environments, and the polymer properties. This framework creates a complex interplay
governing what is reflected in the rate and efficiency of the whole degradation process.
In the next subsections, we selectively provide a discussion of factors important for
mesophilic biodegradation and correlate these factors to the information already provided
about mechanisms and environments. Detailed discussions of these factors are also provided
in selected reviews [48,50,53].
2.7
Environmental factors
Factors that can affect the degradation rate of a polymer are related to the
environment where the degradation process takes place, and include thermal energy (heat),
acidic/alkaline media, moisture, aeration, and microbial populations. Some of these factors
are more relevant or critical than others and are important during the abiotic and biotic
degradation stages affecting both the polymer’s properties and the microbial activity.
2.7.1 Heat
The amount of thermal energy, identified as the temperature of the system, is one of
the main factors affecting the rate of abiotic and biotic degradation mechanisms and varies
46
neriticpelagicepipelagiclittoralsublittoralphotic200 moceanic
with the environment (Table 2.4). In this section, we are not expanding on the thermal
degradation mechanism; instead, we briefly discuss temperature as a factor that can modify
the rate of other mechanisms such as chemical hydrolysis and microbial activity. At an early
stage in the degradation process, mechanisms such as chemical hydrolysis can be dominant
and the temperature plays a crucial role on the rate [64,165]. For example, for PLA, the
chemical hydrolytic degradation is dependent on the temperature since a large initial
reduction of the Mw is needed before microorganisms can assimilate the byproducts [64].
Higher temperatures activate chain mobility, increasing free volume and polymer
rearrangements. If the temperature is higher than the Tg of the polymer, mobility and
reaction are accelerated, increasing the rate of polymer degradation (Table 2.5).
Furthermore, the presence and potential growth of different microorganisms depends on the
environment temperature, and a change in temperature regulates both presence and activity
[86,131]. Biodegradation rate is a function of temperature, mostly described by the Arrhenius
equation above and below Tg [120].
2.7.2 Moisture
The presence of water plays a crucial role in the degradation of hydrolysable chemical
bonds, such as in polyesters, since they are susceptible to chain scission reactions [166,167].
Furthermore, microorganisms need water for transport of nutrients through the cell
membrane and for growth. The amount of water in the different environments, such as soil
and home or
industrial composting, can create different surroundings
for the
microorganisms. Low levels of moisture can lead to dry environments with low biological
activity [168]. High values of moisture will lead to loss of the porosity of the matrix (soil or
compost), turning the process into one with anaerobic conditions [169]. Pore spaces are
essential for the normal air flow and aerobic regimen; the optimal humidity range for
microbial activity is a function of the percentage of pore space needed that does not obstruct
47
the air flow required for the microbial activity [170]. For example, for the composting process,
an optimal range of moisture content is 45 to 65% [148].
2.7.3 Acidic and alkaline media
Acidic or alkaline media can modify the rate of reactions and the mechanism of
hydrolytic degradation [64]. For example, for PLA in acidic conditions the hydrolysis proceeds
via a chain-end scission, while in alkaline solution the hydrolysis takes place via backbiting
[64]. In the case of PCL films evaluated at extreme pH values (1 and 13) at 37 °C, different
behavior was observed for reduction of Mw and crystallinity, suggesting a surface erosion
process in alkaline media and bulk erosion in acidic media [171]. During the biodegradation
process, pH values close to neutral are highly favorable for the growth of microbial
populations. In soil environments, a pH range close to alkaline-neutral values is favorable
for bacteria populations, whereas fungi are more tolerant to acidic and alkaline media;
fluctuations of pH are considered a harmful situation for living organisms [131,172].
2.7.4 Light and UV radiation
If sufficient energy is absorbed by light and UV radiation, polymers can be subjected
to photodegradation, experiencing changes in their chemical structure and physical
properties. Light and UV radiation is important in agricultural soils and aquatic
environments. So, photodegradation can be the precursor of the degradation process before
microorganisms can use byproducts [82,140].
2.7.5 C/N ratio
Microorganisms need carbon as a source of energy and nitrogen for synthesis of amino
acids, proteins and nucleic acids [148]. The C/N ratio is a key parameter in environments
such as compost and soil. Optimal values for the C/N ratio in compost and soil are in the
range of 15:1 to 30:1. During the active aerobic phase of breakdown, microorganisms use
around 30 parts of carbon for each part of nitrogen, due to the high energy requirement. If
48
carbon levels are higher, microorganisms need to undergo several life cycles to oxidize the
excess carbon, slowing down the biodegradation process. If carbon levels are low,
microorganisms do not have sufficient energy source to survive [148].
2.7.6 Oxygen flow and porosity
Aeration and porosity are key factors for the normal activity of the microbial
population in soil and compost environments. To maintain aerobic conditions the porosity
should allow O2 concentrations of around 5%. Porosity is highly correlated with the air flow
within a matrix. Low porosity hinders air flow, whereas high porosity can lead to excessive
aeration and low water retention capacity. The shape, size, and structure of particles of the
matrix (soil or compost) affects its texture. Therefore, a tight packing arrangement reduces
the porosity and the compressed matrix impacts the air flow [148].
2.8
Polymer properties
The factors affecting degradation associated with the bulk polymer matrix can be
categorized as chemical structure and physical properties such as morphology, crystallinity,
constitutional unit, flexibility, crosslinking, Mw, tacticity, density, shape, and polarity. The
surface properties affecting degradation are related mostly to hydrophobic/hydrophilic ratio,
roughness, surface energy, and available surface area.
2.8.1 Bulk properties
Chain flexibility. A polymer chain that is highly flexible is more accessible to attack
by microorganisms. Longer aliphatic chains can exhibit high biodegradation rates. However,
aromatic rings can act as obstacles, providing steric hindrance to the enzyme attacking the
ester bonds, thereby lowering the rate of biodegradation [70]. During the depolymerization
step, enzyme binding is favored by high flexibility of the polymer chains. In this sense, it is
aptly recognized that microorganisms are more likely to start the biodegradation process in
the amorphous region of the polymer [48,49]. Polymers with Tg values in or below the
49
mesophilic range, such as PCL, PBS, PBAT, PHAs, and PGA, will be more flexible in favoring
chemical and enzymatic hydrolysis in the mesophilic range (see Table 2.6). Flexibility and
mobility are enhanced by copolymerization, blending, or by increasing the temperature, and
are reduced by crystalline domains [80,173].
Table 2.6 Polymer structure and thermal properties (Tg,Tm) of the biodegradable polymers
discussed in this work.
Polymer
Structure
Tg
(°C)
Tm
(°C)
PGA
PLA
PCL
PBAT
PBS
PBSA
PBST
35–40
220–
230
55–65
170–
200
-60
58–63
-30
106
-28 to
-32
112–
114
-43 to
-45
95
-20 to
-30
~179
50
OOnOOnOOnOOOOOOOOmnOOOnOOOOOOOOOxyOOOOOOOOmn
Table 2.6 (cont’d)
PBA
PES
PEA
PHB
PHV
PHBV
PU
PVOH
PBSe
51
-61 to
-64
41–61
-9 to -
17
96–
105
-46 to
-50
48
4
180
-10
100–
200
-8 to -
1
180
-63
-
-
-
-62
65
OOOOnOOOOnOOOOHOOnOOCH3nOOCH2-CH3nOOOOmnNCOORnHOHnHOOOHO
Table 2.6 (cont’d)
PBSeT
~ -43
25 to
91
Chemical structure (functional units and functional groups). Chemical structure is an
inherent property of a material and determines whether the polymer is prone to undergo
biodegradation. The chemical structure depicts the spatial arrangement of chemical bonds
and atoms in the molecule influencing the molecular geometry and governs how the
molecules are packed together allowing the formation of crystalline or amorphous regions.
The presence of bulky groups in the main chain, such as aromatic rings, restricts the free
movement of the polymer molecule, reducing chain flexibility such as in PBT. However, when
the linear copolymer of adipic acid and 1,4-butanediol is added to the main chain of PBT to
obtain PBAT, polymer flexibility improves and susceptible hydrolysable bonds are
introduced, so the polymer is more flexible and prone to biodegradation [81]. Modifications
such as inclusion of functional groups by copolymerization in the main chain of initially non-
biodegradable chemical structures can make a polymer more prone to biodegradation [56].
The addition of functional groups also can impart a hydrophilic nature to a hydrophobic
polymer thus improving its likelihood of undergoing biodegradation [109].
Chain structure configuration (side chains and crosslinking). The length of side chains
influences the degradation process. For example, Li et al. [174] concluded that the enzymatic
degradation of PHA was dependent on the length of side chain in the PHA structure.
Crosslinking can occur and play a significant role in polymer mass transfer properties and
chain flexibility hindering biodegradation. Kijchavengkul et al. demonstrated that increasing
the amount of crosslinking reduces the biodegradation of PBAT [175].
52
OOOOOOOnm
Crystallinity. Crystallinity can increase the stiffness and density of a polymer [48]. A
high crystalline fraction decreases the abiotic and biotic degradation rates. The amorphous
region is more susceptible to chemical hydrolysis due to the ease of water diffusion. A
characteristic of the crystalline region is its low mass transfer to gases and vapors, decreasing
the rate of the hydrolytic degradation [176,177]. Extracellular enzymes mainly attack the
amorphous region of the polymer structure [104,178]. Biodegradable polymers are, in
general, semicrystalline polymers with a crystalline and amorphous region.
Molecular weight (Mw). To obtain polymers with usable thermal, mechanical, and
barrier properties a high Mw is required. However, microorganisms assimilate polymers when
selected thresholds of low Mw fractions of the polymer are reached. The higher the Mw value
of the polymer residue, the harder it is for microorganisms to assimilate the chain segments
and assimilate that to their cell, which reduces the rate of the biodegradation. So, a critical
threshold low Mw value must be reached to kick off the degradation by enzymatic attack [70].
Generally, this Mw is attainable by a precursor degradation mechanism such as
photodegradation or chemical hydrolysis, as with polyesters. In the case of PLA, the polymer
first undergoes primarily chemical hydrolysis, accelerated under industrial composting
conditions, until reaching a Mw ≤10 kDa, and then enzymatic activity becomes the dominant
degradation mechanism, with a high mineralization rate [113].
Density and porosity. Denser and more compact polymers have lower chances to
experience water diffusion. For polyesters, chemical hydrolysis is generally the initial trigger
mechanism of degradation, mostly through a bulk erosion process, so water diffusivity of the
polymer plays a crucial role. One way to modify the diffusion or the hydrophilicity of a
polymer matrix is by blending different polymers. So, biodegradable blends and copolymers
53
can be used to tailor some of these bulk properties. Blends of PLA and TPS have shown higher
biodegradation rates [179].
2.8.2 Surface properties
Hydrophobic/hydrophilic ratio, surface roughness, surface energy, and surface/volume
ratio are the more relevant factors during the degradation process. Chemical hydrolysis is
highly affected by the hydrophobic/hydrophilic ratio of the polymer surface. Furthermore,
enzyme activity, biofilm formation, and colonization are also linked to surface properties.
Hydrophobic/hydrophilic ratio. In the case of isotropic polymers, surface and bulk
water sensitivity plays a major role in the degradation process. Hydrophobic surfaces will not
allow water to be adsorbed and will delay water uptake, so that any degradation mechanism
triggered by water diffusion will be delayed. Table 2.7 shows that polymers with hydrophobic
surface and high-water diffusion, such as the polyester PLA, mostly degrade under a bulk
degradation process [63]. So, by tailoring the surface and bulk hydrophobicity and the water
diffusion of the polymer matrix, the overall chemical hydrolysis can be controlled, as shown
for PLA [166]. In terms of enzymatic activity, a hydrophobic/hydrophilic balance allows the
presence of necessary water for optimal microbial activity [162]. Some studies have
demonstrated that biofilms develop faster on hydrophobic nonpolar surfaces [180]. However,
Tsuji et al. reported an alkaline treatment to increase the hydrophilicity of PLLA and PCL
to improve enzymatic attack. The effect was important for PLLA films, where enzymatic
attack by Proteinase K was higher on hydrophilic surfaces [181,182]; however, the attack by
lipases on PCL films remained unchanged [182]. The fact that lipases need a hydrophobic
surface to be active could be an important conditioning of the scarce activity on PCL films.
Furthermore, the exposure to hydrophobic surfaces has been reported to be a relevant signal
for the production of extracellular enzyme cutinases by fungi to act on the surface of
polyesters such as PCL, PBS, and PBSA, among others [183]. Tribedi et al. reported the effect
54
of cell hydrophobicity when comparing enzymatic esterase activity of two strains of
Pseudomonas on the surface of PES. The strain with higher hydrophobicity also showed
higher microbial activity, which is indicative that the interaction and hydrophobic balance
between the microorganism and polymer surface is also relevant for microbial and enzymatic
activity [184].
Table 2.7 Water diffusion and surface property as related to the main degradation process.
Water diffusion
Surface
Degradation process
Example
Low
High
Hydrophilic
Surface
PHAs
Hydrophilic
Bulk/surface
High
Hydrophobic
Bulk
Low
Hydrophobic
Surface (depending on
the ratio of
hydrophobic depletion
and water diffusion)
Starch, TPS,
Cellulose
PLA, PCL,
PBS, PCL
PLA with chain
extender
Surface roughness. Surface roughness is a measure of the finely spaced micro-
irregularities on the surface texture and depicts the irregularities on the polymer surface.
Some researchers have used surface roughness as an indicator of surface biodegradation
[125,185]. The types of microbes able to colonize a surface and the formation of biofilms
depend on the surface roughness. Increased roughness favors bacterial adhesion because of
the greater area of contact between the polymeric material and the bacterial cells [186]. A
rough surface offers micro- and nano-irregularities in the range of 0.5 to 2 m, which appear
as voids and can provide sites for microorganisms to attach and eventually access the polymer
chains, increasing the rate of biodegradation [187,188].
Surface area. The shape (e.g., film, pellet, powder, and fiber) and size (macro, micro,
and nano) of the polymer play important roles during the degradation process [53]. For
example, thicker polyester samples take more time to biodegrade [75]. The surface area
55
available has a high effect on the rate of biodegradation: as the surface to volume ratio
increases with time, so does the speed at which biodegradation occurs. Pits and cracks
continue to increase as time proceeds, and gradually the sample shape and size change,
enabling access to the inside of the matrix [189]. Extracellular enzymes are highly active on
the surface of a polymer since they are relatively large to penetrate the bulk. Hence,
increasing the surface area available for enzymatic attack translates into an increase in the
kinetics of the biodegradation process. Herzog et al. [111] showed that the enzymatic
degradation of a polyester by Candida cylindracea at 40 °C was more effective on
nanoparticles (100 nm diameter) than on thick films (110 μm thickness) of the same
polyester. The effect of morphology on water biodegradation of PHBV was evaluated by
Komiyama et al. [190]. Samples evaluated in powder form showed the faster biodegradation
due to the larger surface area available for biofilm formation in comparison to film, undrawn
fiber, and fivefold drawn fiber.
2.9
Biodegradation assessment
The misuse of the terms “biodegradable” or “biodegradation” has given rise to inflated
and unsubstantiated claims. Claims about general biodegradable products that are used to
deceive consumers into believing that products are environmentally friendly have been
coined as “greenwashing.” It is essential to avoid such false claims, guarantee transparency
to consumers, and stop the unqualified use of vague terms. Certification for biodegradation,
per se, does not exist worldwide. Some polymer and paper materials are certified for
biodegrading in specific environments such as home and industrial composting, soil, and
water [142]. Standards and methods have been developed to aid certification, to avoid
confusion, and to define the environment and conditions in which the samples can be
biodegraded [191].
56
2.10 Standards for evaluation of biodegradation at mesophilic conditions
Several organizations are associated with developing
the standards
for
biodegradability of materials in different environments for different countries and world
regions [191]. Various reviews and reports have provided the standards available for
biodegradation in soils [192], aquatic environments [193], or home and industrial composting
[194,195]. In this review we specifically summarize, in Table 2.8, the different standards
used to assess biodegradability under aerobic conditions for mesophilic temperatures and
tracking evolution of CO2 and O2 demand and cite published works that reported the use of
these standards. Furthermore, standards with specifications of materials to be evaluated and
for certification are described in ASTM 6400, ISO 17088, and EN13432. The environmental
conditions in which the biodegradation takes place are an important aspect since the
biodegradability of a material differs from one environment to another. Development of
standards for assessing biodegradation in different environments is essential [196,197].
57
Table 2.8 Standards for assessing aerobic biodegradation of polymers at mesophilic conditions in different environments, and
selected studies that used the standards to conduct their biodegradation tests.
Parameter
evaluated
Biodegradation
requirement
Temperature
range
Time
frame
Environment
Standard
Name
Selected
published
works
Measure
CO2
evolved
> 60% for
reference
material (end of
test)
Natural aqueous
medium
(inoculum from
activated sludge,
compost, or soil)
20–25 °C
(± 1 °C)
6
months
[198–200]
Measure
O2
demand
> 60% for
reference
material (end of
test)
Natural aqueous
medium
(inoculum from
activated sludge,
compost, or soil)
20–25 °C
(± 1 °C)
6
months
[115,201–
207]
ISO
14852:2018
ISO
14851:2019
Determination of
the ultimate
aerobic
biodegradability
of plastic
materials in an
aqueous medium
— Method by
analysis of
evolved carbon
dioxide
Determination of
the ultimate
aerobic
biodegradability
of plastic
materials in an
aqueous medium
— Method by
measuring the
oxygen demand
in a closed
respirometer
58
Measure
O2
demand,
CO2
evolved
> 60% for
reference
material (plateau
phase or end of
test)
Soil
20–28 °C
(preferably
25 °C, ± 2 °C)
6
months
[115,208–
210]
Measure
CO2
evolved
> 60% for
reference
material after
180 days
Seawater /
sandy
sediment
interface
15–25 °C
(don’t exceed
28 °C, ± 2 °C)
≤ 24
months.
[211]
Table 2.8 (cont’d)
ISO
17556:2019
ISO
19679:2019
Plastics —
Determination of
the ultimate
aerobic
biodegradability of
plastic materials
in soil by
measuring the
oxygen demand in
a respirometer or
the amount of
carbon dioxide
evolved
Plastics —
Determination of
aerobic
biodegradation of
non-floating
plastic materials
in a
seawater/sediment
interface —
Method by
analysis of evolved
carbon dioxide
59
Measure
O2
demand
> 60% for reference
material (after 180
days)
Seawater / sandy
sediment interface
15–25 °C (don’t
exceed 28 °C, ±
2 °C)
≤ 24
months.
Measure
CO2
evolved
≥ 90% for reference
material (within 2
years)
Marine
15–25 °C (don’t
exceed 28 °C, ±
2 °C)
24
months.
Table 2.8 (cont’d)
ISO
18830:2016
ISO
22403:2020
Plastics —
Determination of
aerobic
biodegradation of
non-floating
plastic materials
in a
seawater/sandy
sediment
interface —
Method by
measuring the
oxygen demand
in closed
respirometer
Plastics —
Assessment of
the intrinsic
biodegradability
of materials
exposed to
marine inocula
under mesophilic
aerobic
laboratory
conditions —
Test methods and
requirements
60
Measure
CO2
evolved
> 60% for reference
material (after 180 days)
Marine
sediment
15–25 °C (don’t
exceed 28 °C, ±
2 °C)
≤ 24
months.
Measure
CO2
evolved
Sea water
15–25 ºC
≤ 24
months
Table 2.8 (cont’d)
ISO
22404:2019
ISO 23977-
1:2020
Plastics —
Determination of
the aerobic
biodegradation of
non-floating
materials exposed
to marine
sediment —
Method by
analysis of
evolved carbon
dioxide
Plastics —
Determination of
the aerobic
biodegradation of
plastic materials
exposed to
seawater — Part
1: Method by
analysis of
evolved carbon
dioxide
61
Table 2.8 (cont’d)
ISO 23977-
2:2020
EN
17033:2018
ASTM
D5988-18
Plastics —
Determination of
the aerobic
biodegradation of
plastic materials
exposed to
seawater — Part
2: Method by
measuring the
oxygen demand
in closed
respirometer
Plastics –
Biodegradable
mulch films for
use in agriculture
and horticulture
– Requirements
and test methods
Standard Test
Method for
Determining
Aerobic
Biodegradation of
Plastic Materials
in Soil
Measure
O2
demand
Sea water
15–25 ºC
≤ 24
months
Measure
CO2
evolved
> 90%
conversion
Agriculture
soil
24
months
20–28 °C
(25 °C
preferred,
± 2 °C)
[120]
Soil and
mature
compost
25 ± 2 °C
6
months
[120,189,218–
227,208,228,229,209,212–
217]
Measure
CO2
evolved
> 70% for
reference
material
after 180
days
(starch or
cellulose)
62
Table 2.8 (cont’d)
ASTM
D6691-
17
ASTM
D7991-
15
Standard Test
Method for
Determining
Aerobic
Biodegradation of
Plastic Materials
in the Marine
Environment by a
Defined Microbial
Consortium or
Natural Sea
Water Inoculum
Standard Test
Method
for Determining
Aerobic
Biodegradation of
Plastics Buried in
Sandy Marine
Sediment under
Controlled
Laboratory
Conditions
Measure
CO2
evolved
> 70% for
reference
material
Marine (seashore and open
ocean). Synthetic seawater
with pre-grown population of
at least 10 aerobic marine
micro-organisms. Natural
seawater with inorganic
nutrients
30 ± 2 °C
10–90
days
[115,230–
232]
Measure
CO2
evolved
> 60% for
reference
material
(after 180
days)
Marine (tidal zone, sandy
sediment + seawater)
24
months
[230,233]
15–25 °C
(do not
exceed
28 °C, ±
2 °C)
63
Table 2.8 (cont’d)
ASTM
D5929-18
AS 5810-
2010
NF U52-
001:2005
Standard Test
Method
for Determining
Biodegradability
of Materials
Exposed to
Source-Separated
Organic
Municipal Solid
Waste Mesophilic
Composting
Conditions by
Respirometry
Biodegradable
plastics—
Biodegradable
plastics suitable
for home
composting
Biodegradable
materials for use
in agriculture
and horticulture
– Mulching
products –
Requirements
and test methods
Total O2 uptake > 80g
Volatile fatty acids >
2g/kg (invalid test)
Municipal solid
waste
inoculated with
compost
40 ±
2 °C
45 days
Measure
O2
uptake,
Measure
CO2
evolved
Measure
CO2
evolved
≥ 90% (dry weight)
degradation of test
sample.
Organic waste,
kitchen waste
25 ±
5 °C (<
30 °C)
12 months
[234]
Measure
CO2
evolved
60% for reference
(cellulose) in soil, 90%
for cellulose in
compost or water
media
Soil, compost,
and water
28 ± 5
ºC
12 months in
soil, 6 months in
compost, 6
months in water
64
2.11 Methods for biodegradation assessment
Different methodologies, both quantitative and qualitative, are used to determine the
biodegradation process. When used in combination, the different methodologies help to
recognize if there is any disagreement among the achieved results. Also, supporting
quantitative methodologies, such as CO2 evolution and Mw reduction, with qualitative
methodologies, such as scanning electron microscopy (SEM), visual observation, and
spectroscopy, is helpful in corroborating the biodegradation of the material under study. The
main methodologies to assess and report the degree of biodegradation in aerobic conditions
have been summarized in several reviews [49,195,235–237]. The oldest and most common
methodology is the gravimetric reduction in weight or mass loss of the material under
biodegradation. Significant deterioration in mechanical properties has also been reported as
a degree of biodegradation. Macro visualization, mass loss, and deterioration of mechanical
properties are methods for the approximate assessment of biodegradation. These methods
are more related to physical degradation of the material and not to the biological process
conducted by a population of microorganisms. In general, are more useful for gaining
insights during the early step of polymer biodegradation as during abiotic degradation or
during biofilm formation on the surface of the polymer.
For enzymatic activity clear zone formation, turbidimetric assays, and techniques
that monitor the release of soluble products into the supernatant solution as Total Organic
Carbon (TOC), and spectroscopy combined with chromatography have been reported.
Nowadays the use of microbalance with dissipation monitoring measurements constitutes
an additional analytical technique to evaluate the evolution of the enzymatic hydrolysis of
hydrolysable polymers.
For tracking CO2 evolution and mineralization, respirometric methods has been
developed and are supported by standards for assessing the conversion of the C present in
65
the polymer to CO2. Furthermore, standards also describe for specific environments the
measure of biochemical oxygen demand (BOD) instead of CO2. Radio labeling and tracking
of C has been reported as an adequately technique to complement with respirometric
methods.
Associated with each of the main evaluation methodologies are several techniques
used to quantify the degree of biodegradation. Table 2.9 lists published studies conducted
to measure biodegradation using techniques to measure CO2 and/or O2 under aerobic
conditions at mesophilic temperatures. This section provides a brief description of the
methodologies used and the published studies using those methodologies in the mesophilic
range.
66
Table 2.9 Biotic degradation of polymers at mesophilic conditions measuring CO2 evolution or O2 demand. Polymer details as
shape, initial molecular weight (Mw), and initial crystallinity (Xc); environment in which the biodegradation study is conducted,
testing temperature, the extent of biodegradation with the time frame, and the corresponding selected studies are mentioned.
Published
Parameter Polymer (shape, initial
studies
[219]
[237]
Main result (test
duration)
-
-
Mw, initial Xc)
Cellulose (powder)
Cellulose (paper mulch)
Temperature,
°C
15, 20, 28
27
Environment
CO2
CO2
CO2
PBS (dumbbell, 21.2 kDa,
57.6%)
CO2
PCL (powder, 100 kDa)
CO2
PLA (films, 100–200 kDa),
starch (powder)
Soil
Soil in laboratory
conditions
Soil compost in
laboratory
conditions
Compost in
laboratory
composting
conditions
Soil in laboratory
conditions
CO2
CO2
PLA (sheets, 170 and 180
kDa)
PLLA (film, 100 kDa, 30–
35%)
Soil inoculated in
laboratory
conditions
Aquatic laboratory
conditions
CO2
PLA (films, 163 kDa)
Soil in laboratory
conditions
30
67
25 ± 2
65% CO2 evolution (180 days)
[226]
40
20% mineralization (180
days)
28, 40
30
25, 37
PLA (100kDa): 10–40%
mineralization (28 ºC, 180
days), PLA (200 kDa): 30–
95% mineralization (40 ºC,
180 days)
5–40% mineralization (60
days)
PLA (25 ºC): 10%
mineralization (180 days),
PLA (37 ºC): 12%
mineralization (180 days)
10–25% mineralization (150
days)
[246]
[247]
[222]
[248]
[236]
Table 2.9 (cont’d)
CO2
CO2
CO2
CO2
PHB (powder and film),
PCL (powder), starch
(powder)
Soil in laboratory
conditions
22 ± 3
PHBV (powder, -, 68.9%),
cellulose (powder)
PHBV (film), cellulose
(powder)
PHB (film), PBSe (film),
PBSeT (film)
Marine in
laboratory
conditions
Soil in laboratory
conditions
Marine in
laboratory
conditions
25
28
25
CO2
PLLA (powder and film, 5,
11, 34, 256 kDa, 0, 18,
42%)
Compost
30, 37
[225]
[241]
[224]
[218]
[249]
PHB powder: 91%
mineralization (90 days),
PCL powder: 102%
mineralization (270 days),
PHB films: 26%
mineralization (210 days)
PHBV: 90% mineralization
(450 days)
PHBV: 90% mineralization
(120 days)
PHB: 70% mineralization
(360 days) and 95%
mineralization (200 days),
PBSe: 95% mineralization
(365 and 200 days), PBSeT:
85% mineralization (360
days) and 90%
mineralization (200 days)
PLA (5 kDa): 70%
mineralization (40 days),
PLA (11 kDa): 55%
mineralization (40 days),
PLA (34 kDa): 35%
mineralization (40 days),
PLA (256 kDa): 20%
mineralization (40 days)
68
Table 2.9 (cont’d)
CO2
PHA, PBS, cellulose
(powder)
Soil in laboratory
conditions
25, 37
CO2
CO2
CO2
CO2
CO2
CO2
CO2
CO2
PU (films)
PBAT (films, -, 9%)
PBSe (powder), cellulose
(powder)
PHB (film), PBSe (film),
PBSeT (film), cellulose
(powder)
Soil/Sturm test
Soil
Soil
Soil
30
25
28
25
Cellulose (powder)
UV irradiated PLA
(powder, 198 kDa)
Soil
Inoculated
sterilized compost,
Sturm test
25 ± 2
37
PHB (powder, 470 kDa))
Sturm test
27
PLA (film, -, 20.8), PHBV
(film, -, 72.6), cellulose
Soil
23–25
CO2
PHA (films), PHB (films)
Soil
23 ± 4
69
PHA (25 ºC): 95%
mineralization (150 days),
PHA (37 ºC): 90%
mineralization (180 days),
PBS (25 ºC): 90%
mineralization (200 days),
PBS (37 ºC): 75%
mineralization (180 days)
10 g CO2 evolution (30 days)
5% mineralization (100 days)
55–90% mineralization (140
days)
PHB: 95% mineralization
(360 days), PBSe: 90%
mineralization (360 days),
PBSeT: 90% mineralization
(360 days)
-
PLA (compost): 35–45%
mineralization (40 days),
PLA (Sturm test): 10–20%
mineralization (40 days)
10–80% mineralization (28
days)
PLA: 5% mineralization (190
days), PHBV: 25%
mineralization (190 days)
PHA: 0.2 mM/mg CO2 (90
days), PHB: 0.3 mM/mg CO2
(90 days)
[217]
[250]
[251]
[195]
[216]
[234]
[73]
[252]
[227]
[221]
Table 2.9 (cont’d)
CO2
PHB (film, 175–225 kDa,
48–52%) PHBV (films,
400–300 kDa, 48–52%)
with 1% nucleating agent
CO2
PHBV (films, 455 kDa,
47%), cellulose (powder)
CO2
CO2
CO2
CO2
CO2
PHA (film), PLA (bag,
bottle)
PHBV (film, 500–600 kDa,
14–58%), cellulose, starch
PHA (film), cellulose
(paper)
PLA with chain extender
(films sheets, 449 kDa,
0.9%), PBAT (films sheet,
44 kDa, 15.2%), cellulose
(powder)
PLA (sheets), PHB
(sheets), PBS (sheets), TPS
(sheets), PCL (sheets),
cellulose (powder)
Microorganisms
from marine
environment in
simulated
laboratory
conditions
Marine (foreshore
sand, sand &
seawater,
seawater) in
laboratory
conditions
Marine
30
25
30
Soil
Soil
Soil in laboratory
conditions
25
20 ± 2
28
Soil, home
composting*,
marine pelagic,
and fresh water
25 ± 2, 28 ± 2,
30 ± 1, and 21
± 1
70
PHB: 80–95% mineralization
(115 days), PHBV: 90–100%
mineralization (115 days)
[239]
PHBV (foreshore sand): 90%
mineralization (250 days)
[253]
[254]
[223]
[235]
[220]
[128]
PHA: 38–45% mineralization
(180 days), PLA (bag): 4.5%
mineralization (180 days),
PLA (bottle): 3.1%
mineralization (180 days)
PHBV: 90% mineralization
(250 weeks)
PHA: 70% mineralization
(660 days)
PLA: 10% mineralization
(180 days), PBAT: 20%
mineralization (180 days)
PLA (soil): negligible (141
days), PLA (home
composting): <20%
mineralization (365 days),
PLA (marine water): <10%
relative biodegradation
Table 2.9 (cont’d)
PU (films, 48.7 kDa)
PBAT (films)
Sturm test
Soil
35, 30
30
CO2
CO2
CO2
CO2
CO2
CO2
CO2
CO2
CO2
CO2
PU (films)
PBSA (films)
PLA (sheets)
Cellulose (foil)
PBS (sheets, 90 kDa,
58.9%), PEA (sheets, 88
kDa, 40.6%)
PBSA (films), cellulose
(powder)
PHA (films), PVOH (films)
PCL, PHBV, PBSA,
PVOH, PEA, starch,
cellulose
Sturm test
Sturm test
Sterilized soil,
non-sterilized soil,
non-sterilized
inoculated soil in
laboratory
conditions
Respirometer
Sturm test
(activated sludge)
Compost
Sea water
Aqueous solution
CO2
PLA 3001D (films, -, 7.7%),
cellulose (powder)
CO2
PBAT (films, 56–38 kDa)
Aqueous mineral
solution (including
wastewater)
Soil incubation
71
7.6–8.6 g/l CO2
15% mineralization (120
days)
4.46 g/l CO2
78% mineralization (40 days)
PLA inoculated: 20%
mineralization (60 days)
[255–257]
[258]
[259]
[260]
[228]
-
PBS: 18% mineralization (40
days), PEA: 12%
mineralization (50 days)
70% mineralization (55 days)
PHA: 100% mineralization
(100 days), PVOH: 85%
mineralization (100 days)
PCL: 26% mineralization,
PHBV: 53% mineralization,
PBSA: 3% mineralization,
PVOH: 5% mineralization,
PEA: 36% mineralization (2
weeks)
5% mineralization (115 days)
7–15% mineralization (6
weeks)
[215]
[261]
[242]
[240]
[205,206]
[207]
[149]
35
37
30
20
25
25
30
30
30
25
Table 2.9 (cont’d)
CO2
CO2
CO2
CO2
O2
O2
O2
PU (foam)
PU (foam), cellulose
(paper)
PU (foam)
Non-isocyanate
polyurethane (NIPU)
polyhydroxyurethane
(PHU) (film)
PCL (powder), cellulose
(powder)
PHB (film, 735 kDa, 65%),
PHBV (film 484 kDa, 46%),
PCL (films, 187 kDa, 63%),
PES (film, 87 kDa, 61%),
PEA (film, 144 kDa, 74%),
PBS (film, 79 kDa, 63%),
PBA (film, 81 kDa, 70%),
PBSe (films, 31.5 kDa,
68%)
PHB (film, 735 kDa, 65%),
PHBV (film 484 kDa, 46%),
PCL (films, 187 kDa, 63%),
PES (film, 87 kDa, 61%),
PEA (film, 144 kDa, 74%),
PBS (film, 79 kDa, 63%),
PBA (film, 81 kDa, 70%)
Soil
Soil
Sewage
water/modified
Sturm test
Soil
21 ± 2
27 ± 1
22 ± 2
20–28
43% mineralization (192
days)
10% mineralization (320
days)
32–45.6% mineralization (60
days)
40% mineralization (120
days)
[229]
[230]
[262]
[231]
Aqueous
environment
Freshwater (river)
25
25
Freshwater (lake)
25
30–35% BOD (150 days)
[208]
[263,264]
PHB: 75 ± 16% BOD, PHBV:
76 ± 2% BOD, PCL: 75 ± 8%
BOD, PES: 83 ± 2% BOD,
PEA: 70 ± 3% BOD, PBS: 3 ±
1% BOD, PBA: 20 ± 4% BOD,
PBSe: 6 ± 3% BOD (28 days)
[263]
PHB: 52 ± 7% BOD, PHBV:
71 ± 1% BOD, PCL: 77 ± 1%
BOD, PES: 77 ± 1% BOD,
PEA: 68 ± 8% BOD, PBS: 12
± 8% BOD, PBA: 80 ± 13%
BOD (28 days)
72
Table 2.9 (cont’d)
O2
O2
PHB, PHBV, PCL, PES,
PEA, PBS, PBA
Seawater (bay)
25
PHB, PHBV, PCL, PES,
PEA, PBS, PBA
Seawater (ocean)
25
O2
Cellulose (filter paper)
O2
O2
PLA (film), PBAT (film),
PCL (film and powder),
cellulose (powder)
PLA (films, fibers), PHA
(films)
Seawater (pelagic,
eulittoral,
sublittoral,
supralittoral, deep
sea, buried under
sediments)
Inoculum from
activated sludge
11–26
30 ± 2
Soil
30, 40
73
[263]
[263]
[213]
[209]
[265]
PHB: 27 ± 10% BOD, PHBV:
84 ± 2% BOD, PCL: 79 ± 2%
BOD, PES: 1 ± 1% BOD,
PEA: 65 ± 3% BOD, PBS: 1 ±
1% BOD, PBA: 20 ± 2% BOD
(28 days)
PHB: 14 ± 10% BOD, PHBV:
78 ± 5% BOD, PCL: 43 ± 14%
BOD, PES: 3 ± 2% BOD,
PEA: 46 ± 13% BOD, PBS: 2
± 0% BOD, PBA: 10 ± 5%
BOD (28 days)
-
PLA: 3.7% BOD, PBAT:
15.1% BOD, PCL (film):
34.8% BOD, PCL (powder):
37.7% BOD (28 days)
PLA (films, 30 ºC, 20 days):
9.8–10.3% BOD, PLA (films,
40 ºC, 10 days): 11.8–17.9%
BOD, PLA (fiber, 30 ºC, 20
days): 9% BOD, PLA (fiber,
40 ºC, 10 days): 16% BOD,
PHA (films, 30 ºC, 20 days):
26.3% BOD, PHA (films, 40
ºC, 12 days): 49.5% BOD
Table 2.9 (cont’d)
O2
O2
O2
PBS (sheets), cellulose
(powder)
PHBV (powder, 376 kDa,
58.5%), cellulose (powder)
PLA (film)
O2
PCL (film), PLA (film)
Inoculum from
activated sludge
Aqueous
conditions
Lake water,
compost, soil in
laboratory
conditions
Compost,
activated sludge,
river water, sea
water
O2
O2
O2
O2
PBAT (film, 16 kDa)
Mineral medium
PHBV (powder, film,
undrawn fiber, fivefold-
drawn fiber, 250 kDa)
PCL (powder), cellulose
(powder)
PLA (powder), PCL
(powder)
Freshwater,
seawater
Activated sludge
Aqueous
conditions
74
25
20
20
20
25
25
25
30
PBS: 31% BOD (80 days)
[212]
PHBV: 80 % BOD (28 days)
[214]
PLA (lake water): ~5
mgO2/dm3 water, PLA
(compost): ~25 mgO2/kg
compost, PLA (soil): ~100
mgO2/kg soil (28 days)
PCL (compost): 140
mgO2/dm3, PLA (compost):
125 mgO2/dm3, PCL
(activated sludge): 120
mgO2/dm3, PLA (activated
sludge): 115 mgO2/dm3, PCL
(river water): 10 mgO2/dm3,
PLA (river water): 8
mgO2/dm3, PCL (sea water):
5 mgO2/dm3, PLA (sea
water): 5 mgO2/dm3 (7 days)
10% BOD (22 days), 45%
BOD (45 days)
Powder: 18% BOD, film: 18%
BOD, undrawn fiber: 18%
BOD, fivefold-drawn fiber:
8% BOD (28 days)
PCL: 20–100% (100 days)
PLA: 35% (40 days), PCL:
100% (days)
[266]
[267]
[268,269]
[196]
[210]
[211]
Table 2.9 (cont’d)
O2
PLA (film, particle), PBAT
(film, particle), PBS (film,
particle), PBSA (film,
particle), PCL (film,
particle), PHB (particle)
Seawater in
laboratory
conditions
27
[270]
PLA: 0.3 % BOD, PBAT: 1–
1.4% BOD, PBS: 0.1–1.3%
BOD, PBSA: 0.4–29.2 %
BOD, PCL: 14.5–40.9% BOD,
PHB: 44–60.4% BOD (4
weeks)
BOD: biochemical oxygen demand; *using ISO 14855; **relative to the reference material
75
2.12 Mass loss and mechanical properties deterioration
Measurement of mass loss is the most commonly used method to indicate the extent
of degradation and is indicated as mass loss measured from the samples retrieved during
the degradation test [195]. Mass loss is used mostly to designate the degradation occurring
on the polymer surface and is contingent on the disintegration phenomena. Many
researchers have reported the use of mass loss determination to indicate that the material
has undergone degradation. Furthermore, deterioration of mechanical properties (assessed
on films, sheets, or dumbbell specimens) indicative of degradation by the action of abiotic
mechanisms has been reported along with mass loss.
Quartz crystal microbalance with dissipation (QCM-D). In terms of enzymatic
degradation, microbalance weight loss technique in the nanogram scale has been reported
during enzymatic hydrolysis of aliphatic and aromatic polyesters as PCL and PBAT. This is
a unique approach to monitor the dynamic of the enzymatic hydrolysis and has showed high
sensitivity [262–266].
2.13 Macro and micro visual analysis of the polymer surface
Macro visual analysis is the second most commonly used technique after mass loss.
Macro visual changes of the polymer do not necessarily indicate biodegradation, but these
changes are usually the first evidence of microbial colonization and biofilm formation.
Micro visual inspection using microscopic techniques like SEM, transmission
electron microscopy (TEM) or atomic force microscopy (AFM) can impart more knowledge
regarding the biodegradation process at the early stage, specifically biofilm formation and
the structure of the sample [49]. The topographical changes occurring in the polymer are
usually seen as the formation of holes, cracks, cavities (material erosion), discoloration, or
surface roughness [267]. Kijchavengkul et al. studied the surface evolution during
76
biodegradation of PBAT films and demonstrated the consequences of the degradation by
using SEM methodology among
other techniques [268]. For PBAT samples with c. 30% or less crosslinking, biofilm formation
was observed. Large number of microbes consumed PBAT samples, creating pits in the film
surface. For samples with more than 30% crosslinking, no cavities were observed on the
PBAT film surface, indicating that increased crosslinking results in reduced biodegradation
[268]. Shah et al. reported changes in surface morphology, such as pit formation and erosion,
due to the biodegradation of PHBV films in a basal salt medium after two weeks of
immersion [269]. Techniques as TEM and AFM were extensively used for identification of
chemical and enzymatic degradation of polyesters [270–273].
2.13.1 Chromatography
Size exclusion chromatography (or gel permeation chromatography). It is used to
study the reduction of Mw. Reduction and distribution in Mw are a preferred parameter that
provides evidence of the biodegradation process. When accompanied with mineralization,
the Mw reduction can provide more insights into understanding the process.
High performance liquid chromatography is widely used for qualitative and
quantitative analysis of soluble compounds derived from enzymatic activity that are
released into solution.
Lu et al. examined biodegradability of PPC/starch composites in soil at room
temperature; the study of Mw change for unburied, 40 and 180 days along with weight loss
and other qualitative techniques like FTIR, SEM, and photographs helped the researchers
conclude that PPC was the last component to biodegrade post microbial colonization and
starch degradation [274]. Reduction of Mw by chemical hydrolysis has been reported for
aliphatic and aromatic polyesters as PLA, PCL, PHB, and PBAT, among others [64,275,276].
77
2.13.2 Spectroscopy
A qualitative way to assert biodegradation is by identifying the chemical changes in
the polymer structure [195]. These changes could translate into the formation of low Mw
compounds resulting from the polymer degradation. Changes in the molecular structure can
be identified by various spectroscopic analysis methods.
Nuclear magnetic resonance (NMR): The nuclei of any given type (C, H, N, P, or O)
resonate at different energies. The information from the NMR signal (position and pattern)
gives critical information about the nuclei environment and presence [79,277]. The use of
NMR has been reported for the degradation of different polymers in different environments.
Kijchavengkul et al. studied the biodegradation of PBAT in compost and tracked the
evolution of the BT and BA dimers using 1H NMR and showed that the soft aliphatic portion
and the amorphous region are more susceptible to hydrolysis and biodegradation than the
rigid aromatic portion and the crystalline region [79].
Fourier-transform infrared spectroscopy (FTIR): FTIR analysis of any given material
provides a specific fingerprint spectrum for that material, and the appearance and
disappearance of peaks associated with the functional groups can help explain the changes
happening in the material structure [79,195]. Mass spectroscopy is an analytical technique
widely used for identification of products during enzymatic degradation of polymers. In
general, it is used along with techniques as liquid chromatography.
Weng et al. studied the biodegradation of PHB/PLA blends buried in soil at different
depths at c. 20 °C; the FTIR spectra showed that the peaks in the 4000 to 3000 cm-1 region
were broad in nature due to the formation of -OH and -COOH groups after degradation [278].
Furthermore, Mbarki et al., conducted both the FTIR and NMR analysis on PDLA samples
immersed in the soil/liquid culture at 37 °C and found no significant difference in the
chemical structure before and after immersion (45 days for FTIR and 28 days NMR); the
78
conclusion derived was that the biodegradation phenomena was only surface and not bulk
[279].
2.13.3 Plate (clear zone formation) and turbidimetry assays
Plate tests were originally designed to gauge the resistance of plastics to degradation
via microorganisms. However, in addition to testing resistance they are now used to see if
the polymer can support the growth of microorganisms through biofilm formation. The
polymeric material is dispersed in a petri dish containing a mineral salts agar medium that
serves as the sole carbon source. The polymer in the surface, suspended in the medium, is
then inoculated with microorganisms and held for a predetermined amount of time at a
constant temperature to allow the microorganisms to grow. The formation of a halo or clear
zone around the microorganism colony marks an end for this test, since the clear zone
indicates that the microorganism can at least depolymerize the polymeric material. The test
is also used in screening, isolating, and identifying the potential degrading microorganisms
for any given polymer [280,281]. Urbanek et al. isolated, screened, and assessed the
degrading capability of Antarctic soil microorganisms on PCL, PBS, and PBSA at low
temperature with the help of the clear zone formation technique [282].
2.13.4 Respirometric tests for CO2 evolution and biochemical O2 demand
These tests involve measuring the consumption of O2 or formation of CO2 under
aerobic conditions. The CO2 evolved can be measured by three different techniques [60]: in
cumulative measurement respirometry (CMR) the evolved CO2 (trapped in basic solution
such sodium hydroxide, barium hydroxide) is quantified by titration method [283]; in
gravimetric measurement respirometry (GMR) the evolved CO2 is trapped in absorption
columns and the weight increase is used to quantify the amount of CO2 [283]; and in direct
measurement respirometry (DMR) the evolved CO2 is quantified by means of an inline non-
dispersive infrared gas analyzer or gas chromatograph [113]. Kale et al. compared the use
79
of CMR, GMR, and DMR to assess the biodegradation of PLA under simulated composting
conditions, and found similar evolution of biodegradation [283]. They concluded that the
biodegradation process is further dependent on various factors, including shape, size,
thickness, and sample/compost ratio, among others. The advantages and disadvantages
associated with these techniques are explained in detail elsewhere [268].
Techniques measuring O2 consumption, reported as BOD, are assessed in specific
aquatic environments as sewage sludge and wastewater. However, standards have been also
developed for assessing O2 consumption in soil environments (Table 2.8).
2.13.5 Radiolabeling
An understudied approach for assessing the degree of biodegradation is the use of
radiolabeled carbon. This is one of the absolute tests to determine biodegradation and
involves tracking carbon from biodegradable polymers into CO2 and biomass. The approach
is based on labeling the carbon atoms in the polymer backbone with carbon isotopes: 13C
(stable in nature) and 14C (radioactive) [284,285].
Early works conducted by Albertsson et al. showed that the technique of
radiolabeling polymers using 14C were useful not only for detecting the biotic stage of the
biodegradation process but also the abiotic stage [286–289]. In this sense, PE films produced
using a 14C marker showed 14CO2 evolution of the carbonyl oxidized byproducts when the
films were exposed to soil [286].
Studies in the area of biodegradable polymers addressed in this review are
insufficient. Zumstein et al. employed the use of 13C labeled polymer along with isotope-
specific analytical methods (i.e., cavity ring-down spectroscopy) to track the biodegradation
of PBAT in soil [140]. This technique allowed for tracking of the basic biodegradation steps
by distinguishing the labelled PBAT CO2 from the CO2 evolved due to the mineralization of
organic matter in the soil.
80
In summary, two or more methods are commonly employed together to determine
biodegradation. The change in Mw, weight loss, and surface analysis are used widely but
these alone do not guarantee biodegradation, and at most hint towards disintegration of the
material under study. The evolution of CO2 and radiolabeling represents the complete
assessment of the breakdown of the material into biomass and need to be employed on a
more regular basis for biodegradation studies. Though the respirometry method gives the
mineralization value, radiolabeling is far more advanced by showing the actual integration
of polymer carbon into the microbial biomass.
With respect to standards, there is no international standard specifying how home
composting should be conducted for effective biodegradation of biodegradable polymers.
Also, many standards for determining polymer biodegradation in aquatic environments, as
listed in Table 2.8, mention temperatures (laboratory simulated settings conditions) that
are much higher than the actual conditions encountered in real-world environments. In
general, adaptations of the international standards to specific conditions are implemented
to assess and report, for example, biodegradation of polymers in home composting or at
mesophilic temperatures.
2.14 Microorganisms and enzymes able to biodegrade polymers
The ability to degrade biodegradable polymers is widely distributed among bacteria,
fungi, and actinomycetes, and there is much variation in ability. Table 2.10 lists
extracellular enzymes and/or microorganisms able to biodegrade polymers in different
mesophilic environments, as reported in the published literature.
2.14.1 Microbial Population
Some microorganisms can digest several polymer structures
in different
environments, and degradation rate efficiency can differ. A high portion of the published
works have reported the digestion activity of a specific microorganism in the highly
81
controlled conditions of incubated or culture media; under these conditions, the polymer
substrate is mostly the only source of nutrient for the microorganism. In contrast, in less
restricted environments, such as soil, home composting, industrial composting, or aquatic
environments, the complexity of the biological activity process increases, and several sources
of substrates and microorganisms may be available. In natural soil or aquatic environments,
an active population of microorganisms with different requirements, in terms of nutrients
and optimal growth conditions, are competing or working cooperatively for the resources
available.
The presence of microorganisms and the formation of a biofilm, due to the
colonization
of the polymer surface, creates an effect that sometimes can alter the abiotic degradation of
the polymer. For example, when PCL biodegradation was evaluated under low stirring, the
impediment of biofilm formation resulted in a higher weight loss [298].
82
Table 2.10 Enzymes and/or microorganisms with activity for degrading biodegradable polymers when tested in mesophilic
conditions. Different parameters such as the enzymes released, microbial species used, the environment from which the
microorganisms were isolated / testing media, polymer studied, the temperature and pH for conducting the biodegradation study,
optimal conditions for the microorganisms, and studies reporting them are mentioned.
Environment Polymer
Microorganism*
Reference
Enzymes*
T
(ºC),
pH
Optimal
conditions of T
(ºC) and pH
Alcalase (3.4.21.62)
Bacillus licheniformis (B)
Buffer
solution
PLA
40,
8.0
60, 9.5
[290]
Amidase
(3.5.14)/esterase
(55 kDa)
Rhodococcus equi strain TB-60
Soil/culture
PU
30, 7
45, 5.5
[291]
Carboxyl esterase
(3.1.1.1)
Alcanivorax borkumensis (B),
Rhodopseudomonas palustris (B)
Culture
Carboxyl esterase
Alcanivorax borkumensis (B)
Culture
-
Culture
Aspergillus oryzae RIB40 (F)
Culture
Chymotrypsin
(3.4.21.1)
Cutinase (3.1.1.74)
(21.6 kDa)
Cutinase
Alternaria brassicicola (F),
Aspergillus fumigatus (F), Aspergillus
oryzae (F), Humicola insolens (F),
Fusarium solani (F)
Culture
PCL
PCL,
PDLLA,
PBSA
PES,
PHBV,
PDLLA
PLLA,
PEA
PBS,
PBSA,
PLA
30–37, 9.5–10
[292]
55–60, 9.5–10
[292]
-, -
[293]
35–55, 9.0
[183]
-, -
[294]
30,
8.0
30,
8.0
37,
7.0
37,
8.0
40,
3, 5,
8
Cutinase
Fusarium solani (F)
PBAT
30, -
-, -
[140]
Buffer
solution
83
Table 2.10 (cont’d)
Cutinase (21
kDa)
Cryptococcus magnus (F)
Larval midgut of stag beetle
(Aegus laevicollis)/culture
PBS, PBSA,
PCL, PDLLA,
PLLA
30,
7.4
40,
7.5
[295]
Cutinase
Fusarium solani (F)
Buffer solution
PCL
Cutinase (20
kDa)
Fusarium sp. FS1301 (F)
Soil/liquid culture
PBS, PCL
Cutinase (19.7
kDa)
Paraphoma-related fungal
strain B47-9 (F)
Barely phyllophane/liquid
culture
PBAT, PBS,
PBSA, PCL,
PDLLA
Cutinase
Pichia pastoris (F)
Buffer solution
PBS
Cutinase
-
Cutinase (20.3
kDa)
Pseudozyma
antarctica JCM 10317 (Y)
Cutinase
Fusarium solani (F),
Fusarium moniliforme (F)
Cutinase
Bacillus sp. KY0701
Culture
Culture
Culture
Culture
Cutinase
Aspergillus oryzae (F)
Buffer solution
PBS, PBA
PBS, PBSA,
PCL, PLLA,
PDLLA
PCL
PCL
PCL
-, -
[296]
37,
7.2
30, - 50,
8.0
30,
7.2
45,
7.2
[297]
[298]
-, -
[299]
-, -
[300]
40,
9.5
9–
10
50,
7
-, -
[301,302]
[303]
[304]
[305]
37,
7.4
37,
7.4
30
22
30,
7
40,
8
Cutinase
Pseudozyma jejuensis
OL71 (F)
Cutinase-like
enzyme (22 kDa)
Cryptococcus flavus GB-1
(Y)
Leaves of Citrus unshiu/culture
PCL
30, -
-, -
[306]
Culture
PBSA
30,
6.8
45,
7.8
[307]
84
Table 2.10 (cont’d)
Cutinase-like
enzyme
Close related
to Cutinase
Cryptococcus sp. Strain S-2 (F)
Liquid culture
Pseudomonas pachastrellae JCM12285T (B)
Marine, coastal
seawater/culture
PBS,
PLA,
PCL
PCL
30, -
[308]
37,
7.0
30, -
-, -
[309]
Elastase
Esterase
(3.1.1.1)
Esterase
Esterase
Esterase
-
Culture
PLA
37, 7.0
-, -
[293]
Aspergillus sp. strain S45 (F)
Bacillus sp. AF8 (B), Pseudomonas sp. AF9 (B),
Micrococcus sp. 10 (B), Arthrobacter sp. AF11
(B), Corynebacterium sp. AF12 (B)
Solid waste
dumpsite/liquid
culture
PU
30, 7.0
-, -
[240]
Soil/culture
PU
30–35
-, -
[250]
Hog liver
Buffer solution
PGA
37, 7.5
-, -
[310]
Bacillus subtilis (B)
Buffer solution
Esterase
Aspergillus tubingensis (F)
Esterase
Bacillus licheniformis (B)
Esterase
Esterase
Alicycliphilus sp. (B)
Leptothrix sp. TB-71 (B)
PCL,
PLA
PU
37, -
-, -
[258]
(30, 37,
40), (5
– 9)
[311]
37,
7.0
PLLA
32, 7.4
-, -
[312]
Soil/solid and liquid
culture
Compost/liquid
culture
Culture
PU
37, 7
-, -
[313]
Soil, fresh
water/culture
PBSA,
PES,
PCL
30, -
-, -
[314]
85
Table 2.10 (cont’d)
Esterase (62
kDa)
Comamonas acidovorans strain TB-
35 (B)
Soil/liquid culture
PU
30, 7.2
Esterase (28
kDa)
Esterase (42
kDa)
Esterase
Esterase
Esterase
Esterase
Esterase
Curvularia senegalensis (F)
Soil/liquid culture
PU
(21–25), 30,
35, 45, (4.0–
8.0)
Comamonas acidovorans (B)
Culture
PU
30, 5–8
-, -
[319]
Penicillium verrucosum (F),
Aspergillus ustus (F)
Pseudomonas aeruginosa MZA-85
(B), Bacillus subtilis MZA-75 (B)
Pseudomonas aeruginosa strain S3
(B)
Compost soil/culture
PLA
30, 5.6
-, -
[320]
Soil/liquid culture
PU
37, 7.0
-, -
[246–
248]
Culture
PLA
30–37, 8
37, 8
[321]
Pseudomonas (B)
Porcine liver
Soil/Culture
Buffer solution
45,
6.5
[315–
317]
-, 7–8
[318]
PES
PLA
PES,
PCL
PLA
PBS,
PBSA
PCL
30, -
40, 8.0
30, 7.0
37, 7.2
30, 7.0
-, -
40,
8.0
40–
45, -
- ,-
-, -
[184]
[290]
[322]
[323]
[324]
30, -
-, -
[308]
Close related to
esterase
Bacillus pumilus strain KT1012 (B)
Soil, water/culture
Lipase (3.1.1.3)
Rhizopus delemar (F)
Buffer solution
Lipase
Lipase
Acidovorax delafieldii Strain BS-3
(B)
Soil/solid and
emulsified substrate
Rhizopus oryzae (F), Burkholderia
sp. (B)
Liquid culture
86
Table 2.10 (cont’d)
Lipase
Lipase (36
kDa)
Lipase
Lipase
Lipase
Lipase
Lipase
Lipase
Lipase
Lipase (23
kDa)
Candida rugosa (F)
Buffer solution
PCL, PLA
37, -
-, -
Aspergillus niger MTCC 2594 (F)
Liquid culture
PCL, PLA
30, 7
37, 7.0
Aspergillus oryzae (F)
Buffer solution
Aspergillus tubingensis (F)
Soil/solid and
liquid culture
PCL
PU
37, 7.0
-, -
(30, 37,
40), (5–9)
37, 5.0
[258]
[325]
[326]
[311]
Burkholderia cepacia PBSA-1 (B),
Pseudomonas aeruginosa PBSA-2 (B)
Soil/culture
PBSA
27, 37
-,
[251]
Candida cylindracea (F)
Buffer solution
PLA
40, 8.0
40, 8.0
[290]
Candida antarctica (F)
Buffer solution
PCL, PBS
45, 7.2
-, -
[296,327,328]
Candida rugosa (F)
Liquid culture
PU
Chromobacterium viscosum (B),
Rhizopus orizae (F), Rhizopus niveus (F)
Culture
PCL, PBS,
PBSA
(20–50),
(4–9)
35, 7.0
[329]
37, 7.0
-, -
[330]
Cryptococcus sp. MTCC 5455 (F)
Liquid culture
PBAT
25, -
-, -
[331]
Lipase
Cryptococcus sp. MTCC 5455 (F)
Buffer solution
PU
30, 7.0
[332]
37,
(7.0–
8.0)
87
Lipase (25
kDa)
Lipase
Lipase
Lipase
Lipase (22
kDa)
Lipase
Lipase (34
kDa)
Lipase
Lipase
Lipase
Lipase
Lipase
Table 2.10 (cont’d)
Lipase
Lactobacillus plantarum (B)
Culture
Penicillium sp. strain 14-3 (F)
Soil/liquid culture
Pseudomonas (B)
Buffer solution
PCL
PEA
PLLA, PCL,
PDLLA
37, 8.0
-, -
30, 6.0
45,
4.5
[333]
[334]
37, 7.0
-, -
[335,336]
Pseudomonas cepacia (B)
Buffer solution
PCL
37, 7,0
Pseudomonas cepacia (B), Rhizopus
delemar (F)
Buffer solution
PCL, PPS
30, 7.2
Lipase
Pseudomonas fluorescens (B)
Buffer solution
PCL
37, 7.4
Cryptococcus sp. (Y)
Buffer solution
PBS, PBSA
30, 7
Fusarium solani (F)
Culture
PCL
22, 6.8
-, -
Pseudomonas sp. strain DS04-T (B)
Activated Sludge/liquid
medium
Rhizopus oryzae (F)
Solution
Rhizopus arrhizus (F)
Pseudomonas (B)
Rhizopus oryzae (F)
Rhizopus delemar (F)
Buffer solution
Buffer solution
Buffer solution
Buffer solution
PLLA, PCL,
PHB
PBS, PLLA,
PBA
PCL
PCL
PBAT
PU
88
-, -
-, -
-, -
-, -
[201]
[337]
[338]
[339]
[340]
[341]
37, 8
50,
8.5
40, 5
40, 7
[263]
30, 7
25, 37,
7
30, -
37, -
-, -
-, -
-, -
-, -
[182]
[342]
[140]
[343]
Table 2.10 (cont’d)
Lipase
Lipase
Pseudomonas (B)
Buffer solution
PCL
37, 7
Achromobacter sp (B), Candida
cylindracea (F), Rhizopus arrhizus (F),
Rhizopus delemar (F), Geotrichum
candidum (F)
Buffer solution
PEA, PCL
Lipase
Bacillus sp. (B)
Soil/culture
buffer solution
PBAT
Lipase
Lipase
Lipase
Pseudomonas sp. (B)
Buffer solution
PEA
Stenotrophomonas sp. YCJ1
Soil/culture
PBAT
Candida Antarctica (F)
Buffer solution
PBAT
-, -
-, -
[344]
[345]
-, -
[346]
-, -
[347]
[348]
37,
7.5
-, -
[349]
37,
7.0
30–
37,
7.4
37,
7.0
30,
7.2
45,
7.2
PBAT hydrolase (close
related to cutinase)
(18.9 kDa)
PBAT hydrolase (close
related to Lipase)
PBAT hydrolase (close
related to lipase)
Rhodococcus fascians NKCM 2511 (B)
Soil/liquid
culture
Rhodococcus fascians (B)
Liquid culture
PBAT, PCL,
PBSA, PES,
PBS (low
activity)
PBAT, PCL,
PBSA, PES,
PBS
25, -
-, -
[259]
30, 7
-, -
[350]
Bacillus pumilus (B) (NKCM3101,
NCKM3201, NCKM3202, KT1012),
Brevibacillus choshinensis PBATH (B)
Soil/liquid
culture
PBAT (low
activity), PBSA,
PBS, PES, PCL
30,
7.0
-, -
[81]
PLA depolymerase
(related to lipase)
Paenibacillus amylolyticus Strain TB-13
(B)
Soil/culture
PBS, PBSA,
PDLLA, PCL,
PES
37, 8
45–
55,
10.0
[351]
89
Table 2.10 (cont’d)
PBAT hydrolase
Isaria fumosorosea strain
NKCM1712 (F)
Soil/culture
PBAT, PBA, PBS, PBSA,
PES, PHB, PCL
-, -
[260]
25–
45,
7.0
PBS-degrading enzyme
(44.7 kDa)
Aspergillus sp. XH0501-a
(F)
Soil/culture
PBSA
30
40, 8.6
[352]
PCL depolymerase
(63.5 kDa) (esterase)
Brevundimonas sp. strain
MRL-AN1 (B)
Liquid culture
PCL, data not shown for
PLA, PES, PHB, and
PHBV
37, 7
30, 6–
8
[353]
PCL depolymerase
Penicillium oxalicum
strain DSYD05-1 (F)
Soil/liquid culture
PCL, PHB, PBS
PCL depolymerase
Alcaligenes faecalis TS22
(B)
Culture
PCL depolymerase
Paecilomyces lilacinus
strain D218 (F)
Soil/solid culture
PCL
PCL
PLA depolymerase (58
kDa)
Pseudomonas tamsuii
TKU015 (B)
PLLA degrading
enzyme
Actinomadura
keratinilytica T16-1 (B)
Soil/culture
PLLA
Culture
PLLA
45, 7
PHA depolymerase
(3.1.1.76)
Alcaligenes faecalis (B)
Buffer solution
PHB, PHBV, PHA
PHA depolymerase (48
kDa)
Pseudomonas stutzeri
YM1414 (B)
Fresh
water/buffer
solution
PHB
PHA depolymerase
Ralstonia pickettii T1 (B)
Buffer solution
PHB, PHBV
37,
7.4
37,
7.4
37,
7.5
90
30,
6.8
-, -
[354]
30, -
-, -
[355]
30,
5.2
30,
7.0
[356]
30,
3.5–
4.5
60, 10
[357]
45, 6–
8
[358]
-, -
[359]
55, 9.5
[360]
-, -
[174]
Table 2.10 (cont’d)
PHA depolymerase
Ralstonia pikettii T1 (B), Acidovorax sp.
TP4 (B)
Buffer
solution
PHA
PHA depolymerase
PHA depolymerase
(50 kDa)
PHA depolymerases
(33.8 and 59.4 kDa)
PHA depolymerase
(intracellular)
PHB depolymerase
(3.1.1.75)
PHB depolymerase
PHB depolymerase
(57 kDa)
PHB depolymerase
(49 kDa)
PHB depolymerase
(42.7)
Comamonas sp. DSM 6781 (B),
Pseudomonas lemoignei LMG 2207 (B),
Pseudomonas fluorescens GK13 DSM 7139
(B)
Comamonas testosteroni (B)
Pseudomona mendocina DS04-T (B)
Liquid
culture
PHB, PHV,
PHBV
Buffer
solution
Mineral
medium
PHB,
PHBV
PHB,
PHBV
37,
38,
7.5,
8.0
30,
7.2
37,
7.4
37, -
-, -
[361]
-, -
[362]
[363]
[364]
-, 9.5–
10
50, 8
and
8.5
Pseudomonas putida LS46 (B)
Culture
PHB, PCL,
PES
30, 7
-, -
[365]
Alcaligenes faecalis (B)
Culture
PHB
Alcaligenes faecalis (B), Pseudomonas
stutzeri (B), Comamonas acidovorans (B)
Aspergillus fumigatus (F)
Buffer
solution
Buffer
solution
Comamonas testosteroni strain ATSU (B)
Soil/culture
PHB, PEA,
PES
PHB,
PHBV,
PEA, PES
PHB,
PHBV
37,
7.4
37,
7.4
45,
8.0
37,
7.4
-, -
[366]
-, -
[367]
70, 8
[368,369]
70,
8.5
[370]
Aureobacterium saperdae (B)
Buffer
solution
PHB
37, 7
45, 8
[371]
91
Table 2.10 (cont’d)
PHB depolymerase
(57 kDa)
PHB depolymerase
(50–48 kDa)
Aspergillus fumigatus 76T-3
Emericellopsis minima W2 (F) Wastewater/liquid culture
PHB,
PES, PBS
45, -
PHB,
PHBV
30,
8.0
PHB depolymerase
(40 kDa)
Microbacterium paraoxydans
RZS6 (B)
Dumping yard/culture
PHB
30, -
Penicillium sp. DS9701-D2 (F)
Activated sludge/culture
PHB
28–
30,
6.8
55,
6.4
55,
9.0
30,
7
30,
5
[372]
[373]
[374]
[375]
Sewage sludge/culture
PHBV
30, 8
-, -
[376]
Streptoverticillium
kashmirense AF1 (A)
Acidovorax sp. strain TP4 (B)
Arthrobacter sp. strain W6 (B)
Soil/culture broth
Pond water, river water,
farm soil/culture
Fusarium solani Thom (F)
Wastewater/culture
PHB
25, 8
PHB depolymerase
(62.3 kDa)
Bacillus megaterium N-18-25-
9 (B)
Penicillium sp. (F)
Culture
Culture
Marinobacter sp. NK-1 (B)
Culture
PHB depolymerase Nocardiopsis aegyptia sp. nov.
DSM 44442T (B)
Marine seashore
sediments/culture
PHB,
PHBV
30, 7
-, -
[384]
92
PHB
PHB,
PHBV
30,
8.5
30, 7
PHB
PHB
PHB
30–
37, 9
40, 4–
6
37,
7.4
-, -
[377]
50,
8.5
55,
7
65,
9
50,
5
[378]
[379]
[380]
[381]
-, 8
[382,383]
PHB depolymerase
(46.8 kDa)
PHB depolymerase
PHB depolymerase
(50 kDa)
PHB depolymerase
(47 kDa)
PHB depolymerase
(85 kDa)
PHB depolymerase
(44.8 kDa)
PHB depolymerase
(61.8–70 kDa)
Table 2.10 (cont’d)
PHB depolymerase
(33 kDa)
PHB depolymerase
(36 kDa)
Penicillium funiculosum (F)
Culture
PHB
Penicillium simplicissimum LAR13 (F)
Soil/culture
PHB
PHB depolymerase
Paecilomyces lilacinus D218 (F)
Soil/liquid culture
PHB depolymerase
PHB depolymerase
(48 kDa)
Pseudomonas fluorescens (B),
Pseudomonas aeruginosa (B),
Pseudomonas putida (B)
Contaminated
soil/culture
Comamonas acidovorans YM1609 (B)
Freshwater/culture
PHB depolymerase
Pseudomonas stutzeri (B)
Sea water/Buffer
solution
PHB depolymerases
(44, 46 kDa)
PHB depolymerase
(49 kDa)
PHB depolymerase
(40 kDa)
PHB depolymerase
(53 kDa)
Agrobacterium sp. K-03 (B)
Culture
Streptomyces exfoliatus K10 (B)
Culture
Pseudomonas pickettii (B)
Culture
PHB
-, 6.5
[385]
45, 5.0
[386]
50, 6.5–
7.5
[356]
-, -
[105]
-, -
[387]
-, 7–7.5
[388]
30,
7.5
25,
30,
37, -
30,
6.0
30,
7.9
37,
7.4
30–
45,
7.4
30, 8
45, 7,9
and 8.1
[389]
25–
37, 8
40, 8.5–
9
[390]
40, 5.5
[391]
37,
7.4
PHB,
PCL
PHB,
PHBV
PHB,
PHBV
PHB
PHB,
PHBV
PHB
Comamonas sp. (B)
Solid culture
PHB
37, 8
-, -
[392]
93
Table 2.10 (cont’d)
PHB depolymerase (65
kDa)
Alcaligenes faecalis AE122
(B)
PHB depolymerase (95.5
kDa)
Alcaligenes faecalis AE122
(B)
Seawater/culture
PHB
37,
-, -
[393]
Seawater/culture
PHB
30, 6.8–7.5 55,
9
[394]
PHB depolymerase (40
kDa)
PHB depolymerase (48
kDa)
Aspergillus fumigatus (F)
Culture
PHB
30–32, 8
-, -
[395]
Alcaligenes faecalis T1 (B)
Activated
sludge/culture
PHB
30, 7.5
[396]
-,
7.5
PHB depolymerase
Ralstonia pikettii (B)
Culture
PHB depolymerase (45
kDa)
Paecilomyces lilacinus F4-5
(F)
PHB depolymerase (52.2
kDa)
Diaphorobacter sp. PCA039
(B)
PHB depolymerase (63.7
kDa)
Aspergillus fumigatus 202
(F)
Soil/culture
Culture
Soil/culture
PHB,
PHBV
PHB,
PHBV
PHB,
PHBV
PHB
20, 7.5
-, -
[270]
27–37, 7
30, -
30, 37, 45,
7
[397]
[398]
[399]
[400]
50,
7
45,
8
45,
7
50,
5
PHB depolymerase (20
kDa)
Penicillium expansum (F)
Wastewater/culture
PHB
30, 5
PHB depolymerase
Streptomyces sp. SNG9 (B) Marine/liquid culture
PHB,
PHBV
30, 7
-, -
[401]
94
Table 2.10 (cont’d)
PHB depolymerase
(45 kDa)
PHB depolymerase
(37 kDa)
PHB depolymerase
(48 kDa)
PHB depolymerase
PHBV depolymerase
(36, 68, 72, 90 kDa)
PHBV depolymerase
(43.4 kDa)
PHBV depolymerase
(51 kDa)
PHV depolymerase
(43.6 kDa)
Polyurethanase –
lipase (28 kDa)
Polyurethanase
esterase (27 kDa)
Bacillus (B), Clostridium (B),
Streptomyces (B), Alcaligenes (B),
Comamonas (B), Pseudomonas (B),
Zoogloea (B)
Soil, lake water,
activated sludge,
air/liquid culture
PHB,
PHV,
PHBV
4–58,
4.8–
10.6
29–
35,
9.4
[402]
Penicillium funiculosum (F)
Culture
PHB
30, 5
-, 6
[403]
Paecilomyces lilacinus D218
Buffer solution
Aspergillus clavatus strain
NKCM1003 (F)
Soil/culture
PHB,
PHBV
30,
6.8
45, 7
[404]
30, -
-, -
[405]
PES,
PHB,
PCL, PBS
Aspergillus sp. NA-25 (F)
Soil/solid culture
PHBV
Acidovorax sp. HB01
Activated sludge/
PHBV,
PHB,
PCL
Streptomyces sp. strain AF-111 (B) Sewage sludge/culture
PHBV
30,
7.0
37,
6.8
30–
37,
45,
7.0
[406]
50, 7
[407]
[408]
35–
55,
7–8
Pseudomonas lemoignei (B)
Liquid culture
PHB,
PHV
37, 8
-, -
[409,410]
Bacillus subtilis (B)
Soil/liquid culture
PU
30, 7
-, -
[411]
Pseudomonas chlororaphis (B)
Liquid culture
PU
30,
7.2
-, 7–
8
[412]
95
Pseudomonas chlororaphis (B)
Yeast
extract salts
medium
PU
30, -
[413]
-, 8.5
and
7
Table 2.10 (cont’d)
Polyurethanase
esterase/protease (63 kDa),
Polyurethanase esterase
(31 kDa)
Polyurethanase protease
(29 kDa)
Pseudomonas fluorescens (B)
Polyurethanase lipase
Pseudomonas protegens strain Pf-5 (B)
Polyurethanase (66 kDa)
Acinetobacter gerneri P7 (B)
Polyurethanase – protease
Alternaria solani Ss1-3 (F)
Liquid
culture
Liquid
culture
Liquid
culture
PU
30, 7.2
[414]
25,
5.0
PU
27, 7.4
-, -
[415]
PU
30, 7.0
[416]
[417]
37,
8.0
30,
7.0
Soil/liquid
culture
PU
(20–
35),
(4.0–
8.0)
Polyurethanase – esterase
and amidase
Polyurethanase serine
hydrolase family (21 kDa)
Alicycliphilus sp. BQ8 (B)
Pseudomonas chlororaphis (B), Pestalotiopsis
microspora (E2712A, 3317B) (F),
Lasiodiplodia sp. E2611A (F), Bionectria sp.
strain E2910B (F), Aspergillus niger (F),
Pleosporales sp. E2812A (F)
Protease (3.4.21)
Amycolatopsis orientalis (A)
Protease
Bacillus licheniformis (B)
Liquid
culture
Soil/liquid
culture
PU
37, 7.0
-, -
[418]
PU
30, -
-, -
[419]
Liquid
culture
Buffer
solution
PLLA 30–40,
-, -
[420]
7.0
PLA
37, -
-, -
[258]
96
Table 2.10 (cont’d)
Protease
PLA degrading
enzyme close related
to Protease (40–42
kDa)
Protease, esterase,
and lipase
Protease, PLA
degrading enzyme
Tritirachium album (F), Lentzea
waywayandensis (A),
Amycolatopsis orientalis (A)
Culture
PLLA
Amycolatopsis sp. strain 41 (A)
Soil/liquid culture
PLLA
Amycolatopsis sp. strain
SCM_MK2-4 (A)
Soil/liquid, solid
culture
Stenotrophomonas pavanii CH1
(B), Pseudomonas geniculata
WS3 (B)
Soil, wastewater
sludge/liquid
culture
PLA, PCL
PLA
Proteinase K
(3.4.21.64)
Proteinase K
Proteinase K
Proteinase K
Proteinase K
-
-
Buffer solution
PLLA
Buffer solution
Amorphous PLLA
(not crystalline
PLLA)
Tritirachium album
Liquid culture
PLA
-
-
Culture
PLLA, PES, PEA,
PBS, PBSA, PCL
Culture
PLLA
-, -
[421]
[422]
37–
45,
6.0
-, -
[423]
30,
7.5
30,
8.0
-, -
[424]
[182]
-, -
[335]
-, -
-, -
[308]
[293]
-, -
[181,425]
30,
7
37,
7.0
30,
7.0
30, -
37,
8.6
37,
8.6
30, -
37,
7.0
37,
8.6.
97
Table 2.10 (cont’d)
Proteinase K
Tritirachium album
Buffer solution
(PVAase)-Cu3(PO4)2
Bacillus niacini (B)
Culture
PVOH oxidase (1.1.3.30)
Sphingomonas sp. (B)
PVOH oxidase
Sphingopyxis sp. PVA3 (B)
Activated
sludge/culture
Activated
sludge/culture
PLA
PVOH
PVOH
PVOH
Pseudomonas (B)
Buffer solution
PVOH
PVOH degrading enzyme
(30 kDa)
PVOH degrading enzyme
37, -
-, -
[258]
30, 7
[426]
-, -
[427]
-, -
[428]
40, 7–9
[429]
30,
8.0
25,
7.5
30,
7.2
27,
7.3
Streptomyces venezuelae
GY1
Culture
PVOH
30, 8
-, -
[430]
PVOH degrading enzyme
Penicillium sp. WSH0-21
(F)
Activated
sludge/culture
PVOH
30, 7
-, -
[431]
PVOH degrading enzyme
(67 kDa)
Alcaligenes faecalis KK314 River water/culture
PVOH
Serine enzyme (3.4.21)
(24 kDa)
Amycolatopsis sp. strain
K104-1 (A)
Soil/liquid medium
PLLA
Subtilisin (3.4.21.62)
-
Culture
PLA, PEA, PBS,
PBSA, PCL
-, -
[432]
55–60,
9.5
[433]
-, -
[293]
30,
7.2
37,
7.0
37,
7.0
98
Table 2.10 (cont’d)
Trypsin (3.4.21.4)
-
Culture
PLA, PEA
37, 7.0
-, -
[293]
Aliphatic-aromatic co-polyester
degrading enzyme (27–31 kDa)
Roseateles depolymerans
TB-87 (B)
Soil, fresh
water/culture
Esterase and protease activity
Paenibacillus
amylolyticus TB-13 (B)
Soil/culture
PBS, PBSA,
PCL, PBST,
PES
PLA, PBSA,
PBS, PCL, PES
20–40,
6–11
35,
7
[434,435]
30, -
-, -
[436]
Esterase and amidase
PU esterase (48 kDa)
Lipase, manganese peroxidase,
laccase
Fungal peroxidase (1.11.1.7),
Laccase (1.10.3.2)
Esterase deacetylase (3.5.1.)
-
Buffer solution
Culture
PU
PU
37, 7
37, -
Liquid medium
PVOH
28, -
Pseudomonas fluorescens
(B)
Penicillium
brevicompactum OVR-5
(F)
[437]
[438]
[439]
-, -
-, -
30,
7
Aspergillus sp. (F)
Buffer solution
PU
30, 7
-, -
[440]
Comamonas sp. strain
NyZ500
Activated
sludge/culture
PVOH
37, -
-, -
[441]
-
Pseudomonas aeruginosa
(B)
Culture
PU
37, -
-, -
[442]
99
Table 2.10 (cont’d)
-
-
-
-
Nocardioides OK12
Aspergillus flavus (F)
Culture
Culture
PHB,
PHBV
PU
30, -
28, 6–
6.5
Aspergillus versicolor (F)
Culture
PBSA
30, 7.2
Pseudomonas chlororaphis ATCC 55729 (B)
Culture
PU
(foam)
29, -
PHB
28, 37, -
- Aspergillus fumigatus (F), Paecilomyces farinosus (F), Fusarium
solani (F), Penicillium simplicissimum (F), Penicillium
minioluteum (F), Penicillium pinophilum (F), Penicillium
funiculosum (F)
Activated sludge
soil/farm soil
-
-
-
-
Pseudonocardia sp. RM423 (A)
Culture
PLA
30, 7
Fusarium solani (F), Candida ethanolica (F)
Compost, Soil
PU
25, 45
Enterobacter sp. IBP-VN1 (B), Bacillus sp. IBP-VN2 (B),
Gracilibacillus sp. IBP-VN3 (B), Enterobacter sp. IBP-VN4 (B),
Enterobacter sp. IBP-VN5 (B), Enterobacter sp. IBP-VN6 (B)
Seawater/culture
Acidovorax delafieldii (B7-7, B7-21, B7-28) (B), Streptomyces
acidiscabies A2–21 (A), Streptomyces griseus A2–10 (A),
Fusarium oxysporium F1–3 (F), Paecilomyces lilacinus F4–5 (F),
Paecilomyces farinosus F4–7 (F)
Natural
Soil/incubated
artificial soil
PHB,
PHBV
27.1–
30.4,
7.0–7.5
PHBV
30, -
100
[443]
[444]
[445]
[446]
[243]
[226]
[447]
[448]
[449]
-,
-
-,
-
-,
-
-,
-
-,
-
-,
-
-,
-
-,
-
-,
-
Table 2.10 (cont’d)
-
-
-
-
-
-
-
-
-
-
-
-
-
Pseudomonas aeruginosa (B)
Soil/liquid culture
PDLA
37, -
Fusarium solani WF-6 (F)
Soil/culture
PBS
30, -
Flammulina velutipes (F)
Culture
PVOH
28, -
Aspergillus flavus (F), Aspergillus oryzae (F),
Aspergillus parasiticus (F), Aspergillus
racemosus spp. (F)
Soil/culture
PHB,
PHBV
28 – 30,
6 – 7
[279]
[450]
[451]
[452]
-,
-
-,
-
-,
-
-,
-
Azospirillum brasilense BCRC 12270 (B)
Liquid culture
PBSA
30, 7.0
Aspergillus fumigatus (F)
Compost/culture media
PCL
23, 25,
30, 37,
5.5
-, -
-, -
[453]
[178,454
]
Aspergillus fumigatus (F) strain NKCM1706
Soil/culture
Leptothrix sp. TB-71 (B)
Culture nutrient
broth
PBS, PBSA, PES,
PHB, PCL
30, 7
30, -
[455]
PBST, PBAT
30, -
-, -
[456]
Burkholderia cepacia (B)
Culture
PLLA
35, 7
Bacillus pumilus strain 1-A (B)
Soil/Culture
PBSA, PBS, PCL
30, 7.0
Bacillus sp. JY14 (B)
Marine/culture
PHB, PHBV
Pseudomonas sp. (B)
Marine
water/culture
PCL
30, -
25, -
-, -
-, -
-, -
-, -
[457]
[458]
[459]
[460]
Actinomadura AF-555 (A)
Soil/culture
PHBV
37, -
-, -
[269]
101
Table 2.10 (cont’d)
-
-
-
-
-
-
-
-
Trichoderma viride (F)
Soil/liquid
culture
PLA
28, -
-, -
[461]
Chryseobacterium S1 (B), Sphingobacterium S2 (B),
Pseudomonas aeruginosa (S3, S4) (B)
Compost/liquid
culture
PLA
30, 7.2
-, -
[462]
Amycolatopsis sp. (SST, SNC, SO1.2, SO1.1) (A)
Amycolatopsis sp. (A)
Amycolatopsis sp strain 3118 (A)
Amycolatopsis sp. strain HT-32 (A)
Amycolatopsis sp. strain KT-s-9 (A)
Soil/basal
medium
Culture
Soil/liquid
medium
Soil/liquid
culture
Soil/liquid
medium
PLLA
30, 7
-, -
[463]
PLLA,
PCL,
PHB
PLLA
30, 7.3
-, -
[464]
43,
7.0
[465]
(30,
37, 43,
48),
7.0
PLLA
30, 7.0
-, -
[466]
PLLA
30, -
-, -
[467]
Acidovorax facilis (B), Varivorax paradoxus (B), Pseudomonas
syringae (B), Comamonas testosteroni (B), Cytophaga jhonsonae
(B), Bacillus megaterium (B), Bacillus polymyxia (B),
Streptomyces spp. (B), Aspergillus fumigatus (F), Paecilomyces
marquandii (F), Penicillium daleae (F), Penicillium
simplicissimum (F), Penicillium ochrochloron (F), Penicillium
adametzii (F), Penicillium chermisimun (F), Penicillium
restrictum (F), Acremonium sp. (F)
Soil/incubated
PHB,
PHBV
-, -
[172]
(15,
28,
40),
(3.5,
3.9,
6.3,
6.5,
7.1)
102
Table 2.10 (cont’d)
-
-
-
-
Acinetobacter calcoaceticus, Arthrobacter artocyaneus, Bacillus
aerophilus, Bacillus megaterium, Bacillus sp., Brevibacillus agri,
Brevibacillus invocatus, Chromobacterium violaceum, Cupriavidus
gilardii, Mycobacterium fortuitum, Ochrobactrum anthropi,
Staphylococcus arlettae, Staphylococcus haemoliticus, Staphylococcus
pasteuri, Pseudomonas acephalitica, Rodococcus equi, Bacillus cereus,
Bacillus megaterium, Bacillus mycoides, B. agri, Gordonia
terrari, Microbacterium paraoxydans, Burkholderia sp, Streptomyces,
Mycobacterium spp, Nocardiopsis, Gongronella butleri, Penicillium,
Acremonium recifei, Paecilomyces lilacinus, Trichoderma
pseudokoningii,
Amycolatopsis thailandensis strain CMU-PLA07T (A)
Bacillus pumilus B12 (B)
Kibdelosporangium aridum (B)
- Lentzea (B), Saccharothrix (A), Amycolaptosis (B), Kibdelosporangium
Culture
PLLA
30, 7
(B), Streptoalloteichus (B)
-
-
-
Pseudonocardia alni AS4.1531T (A)
Soil
PLA
30, -
Saccharothrix waywayandensis (A)
Culture
PLLA
30, 7
Tritirachium album ATCC 22563 (F)
Liquid culture
with gelatin
PLLA
30, -
103
Soil
PHB,
PHBV
(26–
31), -
-,
-
[468]
Soil/liquid
culture
Soil/minimal
salt medium
agar
Solid/liquid
culture
PLLA
30, -
PLA
30, -
PLLA
30,
6.6–
7.8
[469]
[470]
[471]
[472]
[473]
[474]
[475]
-,
-
-,
-
-,
-
-,
-
-,
-
-,
-
-,
-
Table 2.10 (cont’d)
-
-
-
-
-
-
-
-
-
Parengyodontium (F), Aspergillus (F), Penicillium (F),
Fusarium (F)
Soil/agar medium
PLLA, PCL
Stenotrophomonas maltophilia LB 2-3 (B)
Mortierella sp. (F), Doratomyces microsporus (F),
Fusarium solani (F), Fennellomyces sp. (F), Aspergillus
fumigatus (F), Verticillium sp. (F), Lecanicillium
saksenae (F), Cladosporium sp. (F), Trichoderma sp. (F)
Compost/sturm
test
PLLA exposed
to UV
irradiation
Compost, soil
PLLA
Bordetella petrii PLA-3 (B)
Compost
PLLA
-, -
[476]
25,
7.0,
6.0
37, 7
-, -
[242]
-, -
[477]
-, -
[239]
25,
7.2
30,
37,
7.0
Flammulina velutipes (F)
Quartz
sand/culture
PVOH
28, -
-, -
[451]
Bacillus cereus RA 23 (B)
Oil sludge/culture
PVOH
Bacillus sp. (B), Curtobacterium sp. (B)
Eutypella sp. BJ (F)
Sewage
sludge/culture
Soil
compost/culture
PVOH
Geomyces pannorum (F), Phoma sp. (F)
Soil/solid culture
PU
PVOH
30, -
-, -
[480]
28,
7
-, -
[478]
[479]
30,
7.0
35,
8.0
-, -
[481]
-, -
[282]
< 25,
5.5,
6.7
(14,
20,
28), -
- Geomyces sp. B10I (F), Fusarium sp. B3’M (F), Sclerotinia
sp. B11IV (F)
Antarctic
soil/liquid culture
PCL, PBS
104
2.15 Extracellular enzymes
Figure 2.13 depicts the main extracellular enzymes reported for depolymerization
of aliphatic and aliphatic/aromatic polyesters, PUs derived from ester, where the ester bond
cleavage is considered as the rate-determining step, [104]; and PVOH. These enzymes belong
to the esterase (EC: 3.1) and peptidase (EC: 3.4) groups of the main group hydrolases (EC:
3); and oxidoreductases (EC 1).
Enzymes like cutinases, esterases, lipases, and PHA/PHB depolymerases are the
main extracellular enzymes for enzymatic hydrolysis of the ester group and belong to the
/ hydrolase family that are structurally similar but with diverse functionality
[173,482,483]. The natural activity of the esterase group of enzymes is the hydrolysis of
lipids. Proteases are the main group for enzymatic degradation of the peptidase group. For
polyurethanes, various esterases, proteases, amidases (EC: 3.5.1.4), and ureases (EC:
3.5.1.5) also have been reported to induce enzymatic degradation. In this case, esterases are
involved in ester scission, and ureases are more inclined to scission of urethane bonds and
are more resistant to chemical and enzymatic hydrolysis [101]. In the case of PVOH, and
also some PU, an oxidative pathway prior to the hydrolytic enzymatic degradation has been
reported, and the main extracellular enzymes are the oxidoreductases (EC: 1).
105
Figure 2.13 Classification of the main extracellular enzymes reported for enzymatic activity of aliphatic and aliphatic/aromatic
polyesters, PUs derived from esters, and PVOH. The numbers in parentheses are the enzyme codes according to the Enzyme
Commission (EC) nomenclature [102].
106
2.15.1 Carboxylesterases
In general, carboxylesterases (3.1.1.1) are reported as esterases, creating some
confusion in the literature since the main group classification (3.1) is esterases. Since the
natural function of esterases is the hydrolysis of lipids, for polymer attack they need a
hydrophobic surface to be activated for scission of ester bonds. Carboxylesterases, in general,
act hydrolyzing short-chains (C < 10) and present a lid domain that covers the active site.
The lid domain (present also in lipases), when binding to the hydrophobic substrate, opens
the active site to promote the catalysis. The lid domain structure is important since some
differences can determine the specificity of the enzymes toward some substrates. Disulfide
bonds is not present in carboxylesterases [484]. Hajighasemi et al. reported the action of
carboxylesterases from Alcanivorax borkumensis and Rhodopseudomonas palustris on PLA
and other polyesters; the enzymatic endo and exo activity resulted in the production of
oligomers, dimers, and monomers of PDLLA but did not show activity for PDLA or PLLA
[292].
2.15.2 Lipases
Lipases (3.1.1.3) are water-soluble extracellular enzymes reported to show enzymatic
hydrolysis activity for several biodegradable polymers such as PLA (PLLA and PDLA), PCL,
PBS, PBSA, PBAT, PBA, PEA, and PU esters. The typical structure of lipases is a protein
structure covered by a lid-like structure. Similar to carboxylesterases, lipases need a
hydrophobic surface to be activated since its natural function is the hydrolysis of lipids;
increased lipase activity is observed when a hydrophobic substrate starts to form an
emulsion due to its contact with a hydrophilic aqueous medium [482,485]. However, the
difference respect to carboxylesterases is that lipases prefer to break down long chains (C >
10). Lipases are unable to hydrolyze ester bonds in intermediates that become water soluble
[111,173]. However, Rizzarelli and Impallomeni reported some ability of lipases to hydrolyze
107
dissolved esters in water solution [486]. Such findings indicate that the nature of the
polymer could be more important than the stereo chemistry in the vicinity of the ester bond
for substrate preference by enzymes with lid-like structures, such as lipases. Also, some
works have reported that lipases act preferentially by random chain scission, showing an
endo-type behavior where Mw reduction is highly affected in comparison to end chain scission
[487].
In the case of lipases, the active site is found in a deep cavity of the protein structure.
This is shielded by a lid-like α-helical structure that is reoriented when in contact with the
substrate. The degree of freedom of polymer chains to move is a key factor in controlling the
hydrolytic depolymerization of polyesters. This mobility ensures that the polymer chain can
fold itself and fit in the active site of the lipase enzyme to carry out the depolymerization
[482]. Hence, polyesters must be mobile enough to reach the active site of the lipase, making
thermal and conformational properties other key factors for depolymerization since exposure
temperature controls the mobility of polymer chains [173]. In general, lipases require a
hydrophobic surface to reach full hydrolytic activity. For this reason, lipases are not likely
to be observed developing high enzymatic activity for the PHAs family of aliphatic
polyesters. The molecular weight of lipases from bacteria has been reported in the range of
20 to 77 kDa [482], thus their size allows activity on the surface of polymers.
2.15.3 Cutinases
Cutinases (3.1.1.74) are hydrolytic enzymes considered the smallest members of the
/ hydrolase superfamily (20–25 kDa) [488,489]. Enzymatic activity for cutinases has been
reported for several biodegradable polymers. Cutinases are mainly produced and released
by fungal pathogens and are able to degrade the polyester cutin, a natural crosslinked lipid
polymer composed of n-C17 and n-C18 hydroxy and epoxy fatty acids, present in plant cell
108
walls and insoluble in water. However, some bacteria can also produce cutinases [490].
Cutinases are able to show enzymatic activity without needing interfacial activation like
lipases and are capable of being active in both soluble and emulsified substrates [490],
primarily due to the absence of the hydrophobic lid that covers the active site. Furthermore,
the active site of cutinases is considered large enough to locate and catalyze even high Mw
polyesters [491]. Cutinases act mostly against aliphatic polyesters; however, results for
aliphatic-aromatic polyesters are scarce. A comparative study of five extracellular cutinases
released by five species of microorganisms found that some cutinases are more stable and
have higher activity towards polymer substrates than others; the higher stability and
efficiency were related to additional disulfide bond formation [294]. In general, the presence
of covalent disulfide bonds and neutral charge in the crowning area of the active site provides
extra stability to the tertiary structure by linking regions of proteins. The presence of
disulfide bonds in a cutinase was also reported by Liu et al. [305], and together with a
favored catalytic triad resulted in improved activity, enhanced thermostability, and higher
activity towards PCL. A cutinase (21.6 kDa) from the fungus Aspergillus oryzae was able to
degrade PBS and PBSA and also showed low activity for PLA [183]. Furthermore, PCL was
reported to be an optimal substrate for cutinases [303].
A study on the effect of pH on the surface charge of the area around the active site of
cutinases reported that the active site becomes more positive as pH decreases from alkaline
to acidic values, resulting in lower activity towards polymers such as PCL [294].
Electrostatic surface potentials generated by charged residues affect the enzyme/substrate
interaction, transition stage stabilization, and efficiency during the product release stage
[294]. Similar results were reported from the interaction of cutinase and PBS; the release of
acidic monomers from PBS affected the pH and the activity of the cutinase, lowering the
degradation rate of the PBS films [296,299]. The presence of the cofactors Ca2+, Na+, and K+
109
increased the activity of cutinases towards polymers such as PCL, PBS, and PBSA; however,
the cofactors Mg2+ or Zn2+ did not show a significant effect or significantly inhibited the
activity of the enzymes [295,297,298].
2.15.4 PHA, PHB depolymerases
PHA and PHB depolymerases
(3.1.1.75 and 3.1.1.76) are produced by
microorganisms and accumulate within the cells as an intracellular carbon and energy
storage. Thus, they can undergo enzymatic degradation by functioning intra or/and
extracellular. PHB depolymerases (3.1.1.75) show activity against short-chain length PHAs
as PHB, PHV, and PHBV, while others (3.1.1.76) show more depolymerization of medium-
chain length PHAs [489]. The primary structure of PHA depolymerases is formed by two
functionally domains, a catalytic domain and a substrate binding domain, and is activated
by the presence of Ca+2 and Mg+2 or inhibited by Cu+2, Fe+2, Mn+2, and Hg+2 [489]. Inhibition
of enzymatic hydrolysis due to the presence of detergents highlights the likely presence of a
hydrophobic region near the active site of PHA depolymerases [489]. Furthermore, PHA
depolymerases are reported to have exo and endo behavior since they were able to release
monomers and oligomers [489]. The extended presence of hyphae due to the colonization of
fungi on PHBV surface has been reported as evidence of enzymatic degradation by
extracellular PHAs depolymerases released by fungus [449]. In terms of the structure, some
PHA depolymerases are reported to belong to the serine esterases group due to the presence
of lipase boxes [386]. Even though PHA depolymerases are specific for PHAs, enzymatic
activity has been reported also for other polyesters.
The presence of additional carbon sources may reduce the enzymatic activity against
polymers. For example, the reduction of PHA depolymerase produced by Aspergillus sp.
showed a repression apparently influenced by the type of carbon source added to the media,
which was indicative of a regulated behavior as a function of the available carbon source
110
[452]. This finding is in accordance with the hypothesis that when abundant labile nutrients
are present, the decomposition of more recalcitrant compounds is inhibited [107]
2.15.5 Peptidases (Proteinase K and Protease)
Peptidases (3.4), a group of enzymes acting on peptide bonds, are also commonly
called proteases, generating some confusion in the literature. Peptidases or proteases
hydrolyze peptide bonds that link amino acids in a protein. For example, Proteinase K
(3.4.21.64) and proteases (3.4.21.112) belongs to the serine endo peptidases (3.4.21), enzymes
that preferentially catalyze bond scission in the middle of the substrate chain. Also, it has
been reported that enzymes belonging to the serine endo peptidases are able to hydrolyze
polyesters like PLA. Proteinase K and proteases have been identified for major enzymatic
activity on PLA. Lim et al. [293] reported the ability of Proteinase K to depolymerize PES,
PEA, PBS, PBSA, and PCL but at lower levels of enzymatic activity than on PLA. More
specifically, Proteinase K showed a higher enzymatic activity for PLLA (amorphous
preferentially) than for PDLA and PDLLA.
The activity of these classes of enzymes towards PLA is still not fully understood in
the sense that these enzymes are more prone to attack the scission of peptide bonds. Tokiwa
and Jarerat [492] concluded that enzymes showing activity on PLA belong to the peptidases
or protease-type group, and these enzymes are able to recognize the repeated L-lactic acid
unit of PLA as the natural homologue L-alanine unit of silk fibroin, a natural protein present
in silk. Later work by Lim et al. [293] reported the enzymatic ability of serine proteases on
PLA, PHB, PES, PEA, PBS, PBSA, and PCL; in particular, alpha-chymotrypsin, a
mammalian enzyme, showed preferential activity for PLA. In studies on PLA
biodegradability, the incorporation of agents to the media, such as silk fibroin or gelatin, as
a nitrogen source to induce production of protease, has resulted in increased enzymatic
activity for PLA since proteases are more prone to interact with peptide bonds [424].
111
2.15.6 Amidases and ureases
In addition to esterases and proteases, PUs derived from esters can be enzymatically
degraded by amidases (3.5.1.4) and ureases (3.5.1.5). Amidases attack the amide groups,
proteases can attack amide and urethane bonds, esterases attack the ester bonds, and
ureases catalyze the hydrolysis attack of the urea groups. However, the information in terms
of amidases and ureases showing enzymatic activity towards PUs derived from esters in
mesophilic environments is scarce [493–496].
2.15.7 Oxidoreductases PU and PVOH-oxidases
Oxidoreductases (1) have shown activity for PVOH and PU within the groups EC 1.1,
EC 1.10, and EC 1.11. More specifically 1.1.3 (with O2 as electron acceptor), polyvinyl-alcohol
oxidase (1.1.3.30) or dehydrogenase have shown enzymatic activity on PVOH; these are
enzymes that can act extra or intracellular. Laccase (EC 1.10.3.2) has been reported to show
enzymatic activity against both PVOH and PU.
2.16 Biosurfactants and synthetic surfactants
Biosurfactants are amphipathic molecules with the capacity of reducing surface and
interfacial tension between liquids, solids, and gases. They contain both hydrophilic and
hydrophobic moieties that can improve the interaction between phases of different degree of
polarity and hydrogen bonding [497,498]. Microorganisms are capable of synthesize and
release biosurfactants such as glycolipids and phospholipids to emulsify the substrate and
stimulate other functions as extracellular enzymatic activity [499].
Hydrophobins are a type of amphipathic surfactant secreted by
fungi
microorganisms, that besides other functions, they attach to the surface of biodegradable
polymers and stimulate the hydrolysis by extracellular enzymes. They present a dual
behavior with hydrophobic and hydrophilic parts (amphipathic proteins), and are adsorbed
to the surface of the polymer, condensing, and stimulating its enzymatic hydrolysis by
112
recruiting extracellular enzymes [500,501]. Hydrophobins are important to support the
growth of fungal aerial structures (hyphae) and conidiospores by playing an important role
for fungal adhesion to hydrophobic surfaces, development of a protective surface coating,
and reduction of water tension [500,501].
Synthetic commercial surfactants are widely used during studies of extracellular
enzymatic activity on polymers and are classified in function of the nature of their polar
grouping. It has been reported the use of ionic and nonionic surfactants. Some common
synthetic nonionic surfactants are added to culture studies for emulsification, like
commercials ones as polysorbate 80 and polyoxyethylene type, to increase microbial activity
on the polymer surface by increasing the hydrophilicity of the surface [109].
Interaction between surfactants and enzymes is still a subject of exploration.
Holmberg mentioned that probably nonionic surfactant are more benign than ionic
surfactants. The way in what ionic surfactants interact with enzymes can introduce
significant changes in the conformational structure of the enzyme [502]. Detailed discussion
about bio and commercial surfactants can be found elsewhere [497,498,502,503].
2.17 Polymers susceptible to biodegradation
The main group of biodegradable polymers susceptible to biodegradation in the
mesophilic range are the aliphatic and aliphatic-aromatic polyesters; besides that, the soft
segment of PUs derived from esters and PVOH are also considered biodegradable to some
extent. Commercialized cellulose and starch-derived polymers, which are bio-based and
naturally biodegradable, are also important to consider when discussing biodegradable
polymers. Figure 2.14 shows the tentative pathways of the most common bio- or fossil-
based polymers reported to undergo depolymerization by specific microorganisms. These
polymers are reviewed individually in this section.
113
Figure 2.14 Main tentative biodegradation pathways for biodegradable polymers in aerobic conditions.
114
2.17.1 Cellulose
Cellulose is a linear homopolymer of D-glucose units joined by −14 glycosidic
linkages, with a degree of polymerization ranging from several hundreds to over 10,000 [504].
Each glucose molecule is upside down in relation to the neighboring glucose molecule so that
the repeating unit is cellobiose, consisting of two glucose molecules linked by a −14
glycosidic bond. The fibrils of cellulose can have crystalline and amorphous regions.
Depending on the origin and treatment, the crystallinity of cellulose can vary from fully
amorphous to fully crystalline. Higher crystallinity makes cellulose resistant to chemical
attacks. In the secondary wall of plant cells, cellulose forms several sheets organized as
parallel microfibrils. These microfibrils are embedded in the matrix of hemicellulose and
lignin. Pure cellulose is available in several forms, such as cotton and filter paper. Before
1950, cellulose-based polymers were one of the most important groups of polymers. Cellulose
nitrate, the oldest plastic, was produced by replacing nitrates on all three hydroxyl groups of
the cellulose glucose units. Several other cellulose ether and ester thermoplastics, such as
cellulose acetate and cellulose butyrate, have been produced through the years [505]. The
main cellulose ethers are: methylcellulose (MC) – non-thermoplastic, water-soluble with high
O2 barrier, generally used as filler and thickener agent; carboxymethyl cellulose (CMC) –
hydrophilic, non-thermoplastic, generally used as a viscosity modifier and thickener;
hydroxypropyl cellulose (HPC) – thermoplastic, with water barrier and grease resistance,
generally used for coatings, and as binder and thickener; and hydroxypropyl methyl cellulose
(HPMC) – non-thermoplastic, non-heat-sealable, generally used for coating purposes.
Cellulose esters are thermoplastic, and they are produced by the reaction of organic or
inorganic acid substituting the hydroxyls of the glucose unit. The prominent cellulose esters
include cellulose acetate (CA), which is thermoplastic, used for molding and extrusion, and
115
can exist in several forms such as cellulose acetate butyrate (CAB), cellulose acetate
propionate (CAP), and cellulose triacetate (CTA) [505].
Cellulolytic and non-cellulolytic mixed populations of microorganisms are present
where cellulosic waste is present. These microorganisms interact synergistically to complete
the biodegradation of cellulose, which is ultimately converted to CO2 and H2O in aerobic
environments through the pathways shown in Figure 2.14, and to CO2, CH4, and H2O in
anaerobic environments.
Cellulose, as well as starch, is enzymatically hydrolyzed to glucose by extracellular
enzymes, which are produced by bacteria and fungi (Figure 2.15). Natural polymers, such
as cellulose and starch, are mostly attacked for enzymatic hydrolysis by cellulases and /
amylases. In addition, oxidoreductases have been identified that can act prior to hydrolytic
enzymes on cellulose [506].
Figure 2.15 Pathway for enzymatic degradation, bioassimilation and mineralization of the
natural polymers, cellulose and starch.
After glucose is produced, glycolysis converts the glucose to pyruvic acid, which acts
as the precursor for the TCA cycle. Glucose, together with the adenosine triphosphate (ATP),
which is the molecule providing the energy source in the cell plus NAD+, and inorganic
phosphate, breaks down into two pyruvates. In the pyruvic acid cycle, Figure 2.16, three
main steps take place. First, a carbonyl group is removed from pyruvic acid, releasing CO2 to
the surrounding media and resulting in a two-carbon hydroxyethyl group bound to the
116
enzyme pyruvate dehydrogenase. Second, the hydroxyethyl group is oxidized to an acetyl
group and the electrons are picked up by the NAD+ (nicotinamide adenine dinucleotide),
forming NADH. This electron will later be used by the cell to create energy through the ATP
process. And third, the enzyme bound to the acetyl group is transferred to CoA, producing a
molecule of acetyl CoA. This molecule is then further converted through the TCA cycle [86].
Figure 2.16 Pyruvic acid to Acetyl-CoA reaction pathway. Adapted from [86].
Cellulose biodegradation occurs primarily by cellulolytic microorganisms belonging to
the bacteria and fungi. Aerobic biodegradation of cellulose occurs mostly by cellulolytic
bacteria; several species
identified
in
the genera Cellulomona, Pseudomona,
Thermomonospora, and Microbispora have been shown to biodegrade cellulose [507].
Cellulose undergoes biodegradation in several environments. Amorphous forms of
cellulose are used as positive controls for biodegradation studies due to their negligible
chemical hydrolysis and rapid enzymatic hydrolysis rates and assimilation by
microorganisms. In thermophilic and mesophilic environments, such as industrial
composting or soil biodegradation, cellulose is widely used as a positive control, as stated in
ASTM and ISO standards (Table 2.8). In marine environments, mineralization of cellulose
powder was reported to reach ~95% after 450 days of testing at 25 °C [233], indicative of its
high biodegradability in aquatic environments. Anunciado et al. [217] used cellulose in the
form of a mulch paper, instead of powder, as a positive control in soil and composting
117
conditions; after 365 days of testing mineralization values were in the range of 50 to 80% for
samples in soil at 27 °C.
2.17.2 Starch
Low-cost starch, mainly obtained from crops not intended for human consumption, is
a bio-based material that can be blended with other polymers to produce novel bio-based and
biodegradable blends. Starch consists of two main molecules making up the constitutional
unit: amylose (linear) and amylopectin (branched). Starches with high amylose content have
been used to produce suitable blends and to improve the thermal, mechanical, and gas barrier
properties of the resulting blends [508–512]. The Tg of pure starch is above its decomposition
temperature, meaning the material does not soften and flow. To make it processable, starch
needs to be combined with plasticizers such as glycerol, poly(ethylene glycol) (PEG), or
sorbitol to obtain thermoplastic starch (TPS). The starch granules are plasticized by using
plasticizers under heating, which provides a viscous melt that can then be processed using
traditional methods such as extrusion foaming and injection molding [513]. TPS is highly
hydrophilic, resulting in leaching of plasticizer during storage and poor dimensional stability
and mechanical properties with time [514]. However, TPS can be used to blend with other
bio-based polymers, improving O2 barrier and elongation at break due to the presence of
glycerol [515–522]. Since the properties of TPS by itself are not sufficient for producing
polymeric structures for some applications, the possibility of blending TPS with other
polymers to improve its mechanical and water barrier properties has opened a wide field for
the development of novel TPS blends, with reactive functionalization as one of the suitable
methods to enhance the compatibilization of TPS [512].
Since starch is sensitive to water, starch or the portion of blends containing TPS will
mostly hydrolyze by enzymatic hydrolysis to glucose. The main extracellular enzymes
118
involved during enzymatic degradation of starches are -amylases (Figure 2.14). The
general pathways for biodegradation and bioassimilation/mineralization of starch are shown
in Figure 2.15 and Figure 2.16, respectively.
Starch, TPS, or TPS blends with other biodegradable polymers have shown high
production of CO2, which is indicative of the high biodegradability of TPS even in mesophilic
environments. Ho and Pometto [118] reported values of mineralization of ~70% for starch at
28 at 40 °C in a soil environment under laboratory conditions after 180 days of testing. The
main characteristic was rapid initial degradation at 40 °C, with a negligible abiotic phase of
degradation, reaching the plateau stage at around day 60; lower activity was observed at 28
°C, reaching the plateau stage at around day 100.
2.17.3 Poly(glycolic acid) – PGA
PGA, the simplest aliphatic polyester, is a biodegradable and biocompatible
thermoplastic, extensively used for many decades in the medical field for implants [523]. PGA
can be synthesized using many mechanisms. Direct polycondensation polymerization of
glycolic acid results in low Mw PGA (Mw < 50 kDa). Ring opening polymerization of glycolic
acid results in high Mw PGA (Mw >50 kDa). Solid-state polycondensation is used to increase
the Mw by increasing the polymer chain lengths in the absence of heat and O2, by constant
removal of byproducts using inert gas or under vacuum [524,525].
PGA has a Tg in the range of 35 to 40 °C and Tm between 220 to 230 °C (Table 2.5)
[526]. PGA displays good gas barrier properties due to its crystalline and stereochemistry
structure [523]. PGA is also resistant to most organic solvents. In addition, the high density
of c. 1.53 g/cm3 awards PGA good mechanical properties compared with other biodegradable
polymers; however, the high cost associated with the PGA production process has hampered
its entry into the consumer market as compared to other biodegradable polymers [523]. In
119
general, PGA is blended with other polymers to improve their properties. For example, when
PGA is blended with PLA, the result is better mechanical properties and improved flexural
modulus of the PLA/PGA blend [527]. Due to its high O2 and H2O barrier properties, PGA
can be used in packaging of products sensitive to O2 [528]. PGA is widely used in biomedical
applications such as sutures, drug delivery, and tissue engineering [529].
PGA degradation starts by abiotic degradation, and chemical hydrolysis is by a non-
specific chain scission of the ester backbone, with bulk erosion as the dominant mechanism
(Figure 2.14) [50,523]. Therefore, water diffusion activated by temperature plays a crucial
role in the initial hydrolysis of the ester backbone. The absence of asymmetrical methyl
groups turns PGA more hydrophilic than PLA, increasing its bulk chemical hydrolysis rate.
Currently, there is limited published information on PGA depolymerization by enzymatic
activity in the mesophilic range in open environments. Since PGA has been used mainly for
biomedical applications, most of the biodegradation data is from in vivo studies at 37 °C.
Extracellular enzymes like esterases have been reported to have enzymatic activity on PGA
sutures [310]. After the initial chemical and enzymatic hydrolysis, PGA is degraded into
small oligomers and glycolic acid, which can be bioassimilated and oxidized to become a
substrate for the TCA cycle, as shown in Figure 2.17.
Figure 2.17 Biodegradation pathway for PGA in aerobic conditions.
In terms of CO2 evolution and mineralization studies, biodegradation of PGA in a
marine environment at around 30 °C, which is high for marine environments, showed a
longer lag phase than for cellulose, but 75% mineralization was reached at 28 days for both
PGA and cellulose [523]. At thermophilic conditions in a simulated industrial composting
120
environment at 58 °C, PGA showed lower mineralization than cellulose; 70% mineralization
was reached at around day 40 for cellulose and at around day 70 for PGA [523].
2.17.4 Poly(lactic acid) – PLA
PLA, a biodegradable aliphatic polyester, is a widely used alternative for conventional
fossil-based plastics. In addition to PLA being biocompatible, biodegradable (compostable),
its production from renewable resources results in energy savings and lower greenhouse gas
(GHG) emissions [530]. The building block for PLA is lactic acid or lactide, which is derived
from the fermentation of glucose obtained from varied sources such as corn and sugar cane.
Lactic acid has two enantiomers: L-lactic and D-lactic acid [531]. Lactide can be produced in
three stereochemical configurations: L, L-lactide; L, D-lactide, and D, D-lactide. High Mw PLA
is obtained by ring opening polymerization of the different lactides and polycondensation of
low Mw lactic acid [532,533]. PLA presents acceptable thermal, mechanical and barrier
properties and its main applications include food and medical product packaging, medical
devices, fibers, textiles, plasticulture, and automotive parts [532]. The ratio of L-lactic and D-
lactic acid in a final PLA formulation plays a crucial role in its final properties and
degradation rate [532–534].
The hydrolysable ester bonds in the backbone of PLA structure (Table 2.5) makes it
susceptible to chemical hydrolysis. Several mechanistic, phenomenological, and probabilistic
models have been developed for PLA and can be extended to other aliphatic polyesters,
explaining how diffusion and geometric properties can modify the pathways and incentivize
one or the other mechanism [75]. The mechanism proceeds in different stages, starting with
water diffusion into the material, followed by the degradation of amorphous regions. After
degradation of the amorphous regions, the random chain scission and cleavage of ester bonds
results in the release of soluble oligomers and monomers [62], which can be used as
substrates for bioassimilation (Figure 2.18). The hydrolysis rate of PLA, as well as other
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polyesters, is highly dependent on temperature (below or above Tg), pH, and several other
properties of the polymer such as Mw and crystallinity, as reviewed elsewhere [64]. In the
absence of other factors accelerating other mechanisms, chemical hydrolysis is the most
important mechanism in the mesophilic range for the abiotic process.
The degradation activity of PLA by microorganisms has been monitored by different
methods and correlated to different biodegradation stages. The crystal structure change or
biofilm formation for PLA degraded in a compost environment was observed using SEM [239].
Weight loss indicating depolymerization of PLA was measured by size exclusion
chromatography [424], the degree of biofragmentation of PLA fibers was monitored by X-ray
diffraction (XRD) [290], and the generation of lactic acid was detected using an enzymatic
bioanalysis kit [475].
The biotic degradation stage implies enzymatic activity and microbial assimilation.
The enzymatic degradation of PLA involves interaction of the polymer with a reagent, such
as water, in the hydrolysis reaction. Hydrolases, such as proteases and esterases, catalyze
the hydrolysis reactions. When lactic acid becomes available for bioassimilation, it is
transported through the semipermeable membrane and is oxidized to pyruvic acid through a
dehydrogenization reaction, which then follows the pyruvic acid pathway (Figure 2.18), as
previously described.
Figure 2.18 Biodegradation pathway for PLA in aerobic conditions.
Various bacteria, fungi and actinomycetes strains have been identified as having some
ability to degrade PLA in different forms such as pellet, film, powder, and sheet. These
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microorganisms were isolated from different environments, such as soil, compost, wastewater
sludge, by enrichment culture media, while some were procured from research facilities. The
extracellular enzymes secreted by these microorganisms have been reported to preferentially
degrade the amorphous regions of PLA, since the backbone chains are highly disordered and
have higher mobility as compared to the crystalline region. This flexibility and mobility aids
in the binding of the backbone chain to ensure a fit into the active site of the enzyme [323].
The extracellular enzymatic activity efficiency is dependent on the type of PLA (PLLA, PDLA,
or PDLLA) as well as the temperature, crystallinity, and Mw of the PLA [239,535].
The enzymatic degradation of PLA involves the hydrolases, with esterases (3.1) and
peptidases (3.4) as the main groups of enzymes. Carboxyl esterases ABO2449 and RPA1511
(3.1.1.1) have been reported to hydrolyze PLLA and PDLA with the highest activity for
ABO2449 at 30 to 37 °C and for RPA1511 at 55 to 60 °C [292]. The analysis of the hydrolysis
suggested that, like other hydrolases (e.g., nucleases and proteases) that are active in
depolymerizing polymeric substrates, these enzymes can exhibit both exo and endo-esterase
types of cleavage [292]. Several esterases (3.1) able to degrade PLA such as lipases, cutinases,
and carboxyl esterases. Peptidases (3.4) have been reported to be able to degrade PLA in
culture media. For example, Proteinase K (3.4.21.64) has been shown to be efficient during
scission of polymer chains, favoring the hydrolysis of the amorphous region of PLLA [536].
The enzymatic degradation of PLA revealed the preferential activity of proteases for PLLA
and for PDLA of lipase/cutinase/esterase type. The enzymatic activity of lipase on PLLA was
affected by the addition of Na+ and K+ that increased the activity. However, Zn+2, Mg+2, Cu+2,
and Fe+2 showed inhibition of the enzymatic activity [341]. Furthermore, the presence of
anionic surfactant showed a significant inhibition of Proteinase K activity towards PLLA
[293]. However, the presence of the same anionic surfactant showed a dual behavior during
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enzymatic activity of / hydrolases on PDLLA, facilitating the binding of carboxylesterases
on PDLLA and also reducing the hydrolytic activity by lipase-like esterases [292,537].
Nonionic surfactant also reduced the enzymatic activity towards PDLLA by lipase-like
esterases [537]. Several studies have reported the importance of factors as stereochemistry,
crystallinity, and hydrophilicity on the enzymatic degradation of PLLA, PDLA, and PDLLA
mostly by action of Proteinase K [538–543].
Some biodegradation studies for PLA in the mesophilic range and different
environments have reported high values of mineralization while other studies have reported
low values of CO2 evolution or mineralization. Ho and Pometto reported values of
mineralization for three types of PLA films in soil environments under laboratory conditions;
after 180 days of testing, mineralization values ranged from 10 to 45% at 28 °C and from 30
to 90% at 40 °C, depending on the film [118]. The effect of temperature also can be observed
in the work of Muniyasamy et al., where mineralization values for PLA films in soil
environment at c. 25 °C were a negligible 5% after 190 days of testing [225]. Biodegradation
studies by Kim et al. in compost showed different mineralization values for PLA with
different Mw and crystallinities after 40 days of testing [239]. The dependence of enzymatic
activity on the initial Mw of PLA was evident, with mineralization values of c. 70% for low Mw
PLA (5 kDa) and c. 30% for higher Mw PLA (34 kDa). The same study reported a similar trend
for different levels of crystallinities at 30 °C, with reduced biodegradation rates for PLA with
high crystallinity [239].
In aquatic environments, biodegradation of PLLA granules resulted in mineralization
value of c. 10% after 180 days of testing at 25 and 37 °C [238]. After 50 days of testing,
mineralization values had reached a similar plateau at both temperatures. Lower
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mineralization values are indicative of limited chemical and enzymatic hydrolysis in aquatic
conditions for PLA.
When PLA (initial Mw c. 188 kDa) powder was exposed to UV irradiation and studied
by using the Sturm test and in compost for 40 days, the highest mineralization values were
found in both the Sturm test (c. 20%) and compost (c. 45%) for samples treated for 8 hours;
longer UV irradiation treatment times resulted in decreased mineralization values after 40
days of testing [242]. The authors stated that a Norrish reaction was not identified as the
main effect for reduced biodegradation with longer UV irradiation time, leaving the presence
of crosslinking as the most probable one.
Biodegradation of PLA sheets after 180 days of testing in soil at 28 °C resulted in c.
10% of mineralization [218]. Lower values obtained for PLA sheets in comparison to powder
samples indicates the effect of shape and size as important factors decreasing the chemical
hydrolysis and the mineralization rate.
Biostimulation and bioaugmentation of soil environments to improve PLA
biodegradation under mesophilic conditions was studied by Satti et al. [216]. After 150 days
of testing, improved results, with respect to natural biodegradation of PLA, were obtained
for biostimulated soil with lactate and bioaugmented soil with previously isolated PLA-
degrading bacteria strains. Techniques such as biostimulation and bioaugmentation to
improve biotic conditions are increasingly considered as feasible alternatives to increase the
biodegradation rate of polymers. In this sense, UV irradiated PLA sheets in soil, inoculated
with Pseudomonas geniculata WS3 at 30 °C, showed maximum biodegradation values of c.
35% after 60 days of testing. However, in the case of soil non inoculated, the biodegradation
was just about 15% after 60 days [220].
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2.17.5 Polycaprolactone – PCL
PCL is a synthetic, aliphatic biodegradable polymer, semicrystalline in nature, and is
obtained by the ring opening polymerization of caprolactone [544]. PCL has a Tg of around
−60 °C and a Tm of 60 °C. Since the Tg is so low, PCL shows high molecular chain mobility
due to its rubbery state [545]. Also, this low Tg provides good flexibility and malleability to
PCL [544]. PCL is non-hazardous and biocompatible, so the polymer is often used in
biomedical applications such as tissue engineering, drug delivery, and in the construction of
scaffolds and sutures [546]. PCL displays excellent rheological and viscoelastic properties.
Aside from the many listed advantages, the mechanical properties are less suited for rigid
applications. The inferior mechanical advantage coupled with improved degradation rate
warrants the use of fillers and incorporation of different polymers to attain the necessary
mechanical properties. PCL is usually a raw material for production of polyurethanes, as
polyol polyester-type [495,547].
The main abiotic degradation mechanism for PCL in the mesophilic range is chemical
hydrolytic degradation through bulk erosion [548]. Furthermore, PCL can photodegrade
when exposed to radiation via Norrish II reactions [549]. UV treatment also has been
effective in increasing the degradation rate of PCL films, making it easier for microorganisms
to attack during the biodegradation phase [550]. Due to its relatively low Tm (60 °C), PCL can
undergo thermal degradation at conditions similar to the thermophilic conditions of the
industrial composting process. A short abiotic lag phase was reported for PCL in home
composting conditions, showing a biodegradation trend similar to readily biodegradable
materials like cellulose or starch [115]. However, the initial Mw of the used PCL was low (Mw
c. 50 kDa) in comparison to others polyesters evaluated such as PLA and PHAs [551].
A comparison of the hydrolysis mechanisms for PCL in water and phosphate buffer
solutions revealed that, in general, enzymatic hydrolysis was faster than abiotic chemical
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hydrolysis in terms of mass loss, and that enzymatic hydrolysis is a surface erosion process
whereas abiotic chemical hydrolysis is a bulk erosion process [338]. However, in more real-
world conditions the enzymatic hydrolysis also could be affecting the chemical hydrolysis.
For example, a comparison test for PCL abiotic degradation at 30 °C by Aspergillus fumigatus
showed a different pattern; samples studied for chemical hydrolysis remained without
surface changes, while samples in culture media showed an erosion pattern indicative of
surface enzymatic degradation [178].
Carboxyl esterase (3.1.1.1), lipases (3.1.1.3), and cutinases (3.1.1.74) are able to
degrade PCL. Also, low enzymatic activity by peptidases such as Proteinase K (3.4.21.64) has
been observed. Cutinases from fungal phytopathogens are indicated as PCL depolymerases
showing enzymatic activity [100]. Based on earlier works studying and identifying aerobic
microorganisms able to biodegrade PCL, it has been reported that the natural polymers cutin
and suberin are enzymatically degraded by lipases; since these materials are considered as
natural analogous to PCL, enzymatic activity of PCL by lipases was potentially considered.
Nishida et al. demonstrated that lipases are highly active in the degradation of PCL [552].
This finding was also indicative of potential microbial populations for PCL biodegradation
being extensive in natural environments such as soil, home composting, and water. When
enzymatic degradation of PCL by lipases and Proteinase K available in those environments
was studied, lipase activity was reported but none for Proteinase K; the authors associated
this result to the preferential specificity of lipases for ester bonds on hydrophobic substrates
as in PCL [335].
Temperature and pH are key factors also identified in playing a main role in the
degradation of PCL. The high stability of cutinases able to degrade PCL was associated with
stabilization of the enzymes by neutral surfaces and additional disulfide bond formation
[294]. Baker et al. compared cutinases for PCL degradation and showed that enzyme activity,
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stability, and efficiency was affected by the type of microorganism that releases the
extracellular enzyme and by temperature; the authors reported a similar residual activity for
the enzymes at 25 °C, but reduced residual activity at 45 °C for some of the cutinases [294].
Li et al. reported the presence of 6-hydroxy-hexanoic acid instead of PCL oligomers
during the enzymatic degradation of PCL by Penicillium oxalicum, indicative of an exo-type
chain-end scission by the enzyme [354].
PCL degrading microorganisms and extracellular enzymes have been reported also in
marine environments, showing relatively good activity in comparison to other aliphatic
polyesters as PLA [553].
When 6-hydroxycaproic acid becomes available for bioassimilation after chemical and
enzymatic depolymerization of PCL (Figure 2.14), it is transported through the
semipermeable membrane, and then is converted to acetyl-CoA by -oxidation of fatty acids,
becoming available for the TCA cycle (Figure 2.19).
Figure 2.19 Tentative biodegradation pathway for PCL in aerobic conditions. Adapted from
[544].
A few studies have demonstrated PCL biodegradation with production of CO2 and
mineralization at mesophilic conditions like soil or home composting. Ohtaki et al. reported
a low mineralization value of c. 15% for PCL powder (Mw c. 100 kDa) in compost after 8 days
of testing [138]. Modelli et al. reported ~102% mineralization for PCL in a soil environment
in laboratory conditions at 22 °C after 270 days of testing [223]. Narancic et al. tested the
biodegradation of PCL sheets in home composting and in marine (30 °C) and fresh water
(21°C) environments [115]. In home composting, the PCL reached mineralization values of c.
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90% after 180 days of testing, with a negligible abiotic degradation stage; however, PCL failed
the marine (56 days) and freshwater tests, with mineralization values of c. 80 and 50%,
respectively.
2.17.6 Poly(alkylene dicarboxylate)s
Poly(alkylene dicarboxylate)s are a family of biodegradable aliphatic polyesters
derived from dicarboxylic acids and dihydroxy compounds [50]. This family includes PBA,
PBS, PBSA, PBST, PBSe, PBSeT, PEA, PES, among others. Their general structures are
presented in Table 2.5, and a general description of the main polymers of the family is
provided here.
PBA is a biodegradable polyester that can be synthesized via polycondensation of
adipic acid with 1,4-butanediol in the presence of a catalyst. Due to its low Tm (41–61 °C),
PBA is generally copolymerized to obtain polyesters with improved mechanical properties.
Potential applications for PBA are mainly in the medical area [50].
PBS, a biodegradable, linear, semi-crystalline, thermoplastic aliphatic polyester, is a
result of the condensation polymerization of succinic acid (SA) and 1,4-butanediol (BDO).
PBS can be 100% bio-based (bio-based SA and BDO), partially bio-based (bio-based SA and
petrochemical BDO) or fossil-based (petrochemical SA and BDO), depending on the
production route used [554]. SA is derived from maleic anhydride, which can be produced by
the oxidation of butane or benzene or from the fermentation of carbohydrate sources such as
glucose and starch [555]. BDO, on the other hand, can be derived via three routes: using
petrochemicals, hydrogenation of SA, or fermentation of sugars [556]. PBS provides easy
processability and mechanical properties comparable to LDPE and PP. The fact that PBS is
flexible and not rigid and brittle like other biodegradable polymers (e.g., PLA, PBAT, and
PHB) makes it a more viable and a cost-effective option for common applications [557]. The
properties of PBS can be further fine-tuned for designated applications by blending with
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other polymers. For example, PBS/PLA blends have improved toughness and elongation at
break with the help of random copolymers of poly(butylene succinate-co-lactic acid) as
compared with neat blends [558]. PBS applications vary from food packaging, agriculture
mulch films, hygiene products, fishing nets, plant pots, and coffee capsules, among others
[559].
PBSA is obtained when adipic acid is added when synthesizing PBS. The addition of
adipic acid decreases the crystallinity and increases the degradation rate [50]. In comparison
to PBS, PBSA has a lower Tm of c. 95 °C (Table 2.5) and higher flexibility in terms of
mechanical properties [489].
PBST is an aliphatic/aromatic polyester synthesized by direct esterification and
polycondensation using titanium tetraisopropoxide as a catalyst. PBST has a potential
development, and works about PBST biodegradation are limited [50].
PEA, an aliphatic polyester, is produced by the polycondensation of ethylene glycol
and adipic acid or by the polycondensation of dimethyl adipate and ethylene glycol [560]. PEA
has a Tg of c. −50 °C and Tm of c. 48 °C (Table 2.4). The polymer displays good flexibility due
to the low Tg, but at the same time demonstrates low mechanical strength [561]. PEA is
usually blended with other polymers. When blended with PLA, PEA helps in reducing the
brittleness, improving the thermal stability, and has also been shown to increase the
elongation at break compared with neat PLA [562]. PEA has application as a plasticizer,
when low migration, good plasticity and better mechanical properties are desired for the
copolymer blend [563].
PES is synthesized by ring opening polymerization of succinic anhydride with
ethylene oxide or by polycondensation of succinic acid and ethylene glycol (Table 2.5). PES
is highly permeable to O2 [50].
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In terms of abiotic degradation, the density of the ester bond is a factor affecting the
chemical hydrolysis rate of poly(alkylene dicarboxylates). As reported for PES, a smaller
ester bond reduces the hydrophilicity, affecting the overall process of hydrolysis [50]. For
PBS, abiotic degradation generally occurs through chemical hydrolysis, with bulk erosion as
the predominant mechanism [48]. In general, PBS is copolymerized with the aim of
increasing the hydrolysis rate. For example, the addition of more hydrophilic components,
such as PEG, has been reported to increase abiotic hydrolysis; however, adipic acid is the
most common component added to obtain PBSA [50]. Hayase et al. reported a higher
degradation of PBSA than PBS in the presence of Bacillus pumilus, which was attributed to
the faster degradation of the adipate units [458].
Extracellular enzymes with activity for poly(alkylene dicarboxylate)s have been
reported, mostly for PBA, PBS, PBSA, PEA, and PES (Table 2.10). In general, lipase activity
data is scarce for PBA, PBS, and PBSA; however, the activity of cutinases is well reported for
PBS and PBSA (Table 2.10). Cutinases are indicated as being more active for polyesters with
chain lengths less than 10 C [489].
Fungi and bacteria have been shown to depolymerize PBS. For example, Ishii et al.
[455] reported succinic acid and 1,4-butanediol as hydrolysis products due to the action of
Aspergillus fumigatus strain NKCM1706. Li et al. [352] reported the action of an exo
extracellular enzyme on PBS as an exo attack, since products identified by mass spectrometry
were succinic acid and butylene succinate monomers rather than PBS oligomers.
Furthermore, the presence of butylene succinate monomers and not 1,4-butanediol showed
that the enzyme cut the polymer chain from the carboxyl end [352]. In the case of PBSA, 1,4-
butanediol, succinic acid, and adipic acid were detected by HPLC during depolymerization by
Leptothrix sp. TB-71 [314]. Enzymatic activity of Rhizopus delemar lipases against PEA
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produced, besides ethylene glycol and adipic acid, PEA oligomers; indicative of an endo attack
of the lipase extracellular enzyme [345].
When SA and BDO become available for bioassimilation (Figure 2.20) after chemical
and enzymatic hydrolysis, they are transported through the semipermeable membrane and
converted to succinyl-CoA, becoming available for the TCA cycle (Figure 2.20).
Figure 2.20 Tentative biodegradation pathway for PBS in aerobic conditions. Adapted from
[564].
Some studies reported mineralization (or CO2 production) for PBS, showing limited
biodegradation in the mesophilic range, including home composting, soil, marine and
freshwater environments. Narancic et al. tested the biodegradation of PBS sheets in soil,
home composting, and in marine and freshwater environments, and found mineralization
values lower than 20% after 365 days of testing in soil and home composting, whereas values
were c. 20 and 5% in marine and freshwater after 56 days of testing, respectively [115].
However, other group evaluated PBS in powder form in soil environments at 25 and 37 °C,
and mineralization values reached c. 85 and 80%, respectively, after 180 days of testing [210].
PBSe and PBSeT (films) were assessed in a marine environment at 25 °C under
laboratory conditions, and similar mineralization values of c. 90% were obtained after 360
days of testing with stirring and without stirring the media containing the samples [211].
PBSe and PBSeT (films) in soil at 25 °C reached mineralization values of c. 90% after 360
days of testing [209]. Furthermore, when PBSe (powder) was evaluated in soil at 28 °C,
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mineralization improved for samples with higher available surface area; at day 140, the
mineralization values were c. 55% for samples with 33 cm2 surface area, and 80 to 95% for
samples with 89 to 1650 cm2 surface area [189].
PBSA films was evaluated in compost at 25 ºC with values of mineralization of ~70%
after 55 days of testing, with an abiotic phase duration of ~1 week [234].
2.17.7 Polyhydroxyalkanoates – PHAs
PHAs comprise a family of naturally occurring biodegradable aliphatic polyesters
produced due to the fermentation of carbohydrate sources, such as sugar and lipids, by the
action of a broad range of microorganisms [531,565]. PHAs are synthesized and stored as an
intracellular energy resource for later metabolism under conditions of scarcity. PHAs can be
classified according to the length of the side chain. The most common are short-chain-length
PHAs, with 3 to 5 carbon atoms in their monomeric structure [50]. Poly(3-hydroxybutyrate)
(PHB), poly(hydroxyvalerate) (PHV), and the copolymer poly(hydroxy-butyrate-co-valerate)
(PHBV) are the most common, and PHB is abundantly manufactured. PHAs can be derived
from renewable and non-renewable sources [566], and have excellent barrier and good
thermo-mechanical properties [567]. However, drawbacks for PHAs in conventional thermal
processing include a narrow processing window and high production costs. To improve the
processability and ensure large-scale production, PHAs are often blended with other
polymers. PHAs have been commonly used for cutlery, trays, food packaging, and cosmetics,
and in the development of medical devices, surgical sutures, implants and tissue engineering,
among others [568].
The most common PHAs undergo abiotic degradation by chemical hydrolysis scission
of the ester bonds (Figure 2.14). A discussion is still open in the field whether PHAs go
through a bulk or surface erosion process regardless of the thickness [48,50]. However, some
studies have reported reduction of Mw, mass loss, and mechanical properties deterioration
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during degradation, indicative of surface erosion as the dominant mechanism [569,570].
Specifically, for the copolymer PHBV, a surface erosion process for both enzymatic and
chemical hydrolysis was reported [571].
After depolymerization of PHB (as an example PHA), 3-hydroxybutyric acid is
bioassimilated and, through a redox reaction, converted to acetyl-CoA, which feeds the TCA
cycle (Figure 2.21).
Figure 2.21 Tentative biodegradation pathway for PHB in aerobic conditions.
Degradation of short-chain-length PHAs by enzymatic activity from bacteria and
fungi has been extensively reported [48,50,489,572]. In general, an increase in side-chain
length decreases the hydrolytic rate of the PHAs [359].
PHAs can be metabolized by intra or extracellular depolymerases depending on its
location. In this sense, in vivo granules are amorphous PHAs that can be metabolized by
intracellular enzymes. Denatured PHAs, after cell lysis, becomes semi-crystalline and can be
depolymerized by microorganisms that release extracellular depolymerases [489,573].
PHB depolymerases (3.1.1.75) and PHA depolymerases (3.1.1.76) are the main
enzymes reported as able to degrade PHB and other PHAs. A bacterial PHB depolymerase
has been shown to have two functions during the hydrolysis of PHB films, which takes place
via adsorption and hydrolysis, binding, and catalytic domains [366]. Investigations revealed
that the binding domain of the enzyme is non-specific for binding to the surface of PHA films;
however, the active site in a catalytic domain is specific for the hydrolysis of the PHB
molecule [366].
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The activity of extracellular PHB depolymerases in enzymatic depolymerization
occurs initially on the surfaces of the polymer after biofilm formation, and the rate is
dependent on the Mw, crystallinity, and microbial community [574]. PHA depolymerases can
be described as serine hydrolases with protein progression formed by four regions. First is
the signal sequence, second is a catalytic area which contains the lipase box, third is a
substrate-binding domain where the adsorption of the polymer substrate takes place, and
eventually a domain which connects the catalytic area to the substrate securing areas [100].
Stimulation activity for PHB depolymerase was observed in presence of Na+, K+, Ca2+,
and Mg2+ [105,380]. However, Fe+2, Hg+2, Mn+2, and Cu+2 were reported as inhibitory of
enzymatic PHB activity [399,489].
Nishida et al. reported the effect of crystallinity and amorphous fraction on microbial
degradation [575]; showing that increased crystallinity repressed microbial activity.
PHB, PHV, and PHBV have been reported to be biodegradable in several mesophilic
environments such as soils, composts, and natural waters (Table 2.9). A large microbial
population has been identified as associated with biodegradation of PHB and the copolymer
PHBV in mesophilic conditions [50]. Biodegradation in soil of PHBV films was reported as
the combined action of fungi, bacteria and actinomycetes; however, the fungi population was
identified as the dominant one due to the ability to increase the surface growth of hyphae
[48,449]. Copolymer composition, crystallinity, microstructure, and surface morphology are
factors reported to play an important role during biodegradation of PHBV in soil [221].
During degradation of PHBV in seawater, an increase in surface roughness was observed,
which was reported as both surface erosion by chemical hydrolysis and enzymatic activity
[48,576].
During biodegradation of PHBV (films) evaluated in soil at 25 °C, mineralization
values of c. 25% were achieved after 190 days of testing [225]. Higher mineralization values
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were reported for PHBV in marine environments. Thellen et al. [231] reported high
mineralization for high Mw PHBV films with different contents of valerate in a marine
environment at 30 °C; after 100 days of testing, mineralization values reached c. 100% for
PHBV with 5 and 8% of valerate, and c. 90% for PHBV with 12% valerate. A recent study by
Meereboer et al. [233] evaluated PHBV in powder form in a simulated marine environment
at 25 °C; mineralization values were higher than 50% at day 190, and values reached c. 90%
at the end of the test (450 days).
Mineralization of PHB in a soil environment was reported after 360 days of testing,
with a degree of biodegradation of c. 95% at 25 °C [209]. High mineralization values were
also reported by Narancic et al. [115]. After 136 days of testing in soil at 25 °C, mineralization
values higher than 100%, showing priming effect, were reported; however, when evaluated
in home composting, low mineralization values (less than 20%) were reported for PHB at 28
°C [115].
On the other hand, PHB in marine environment showed c. 70% of mineralization after
360 days of testing and c. 95% in the same test after 200 days, but with a stirring system; the
difference in CO2 evolution was attributed to the shortage of O2 in the system without stirring
[211]. Thellen et al. also reported high mineralization values (in the range of 80 to 90%) for
high Mw and high crystallinity content PHB films in a simulated marine environment at 30
°C; this work was indicative of the high degradability of polymers from the PHA family even
though the % crystallization and initial Mw was high for both PHB and PHBV [231]. Narancic
et al. reported mineralization values of c. 90% for PHB sheets at day 56 of testing in a marine
environment at 30 °C, whereas in a freshwater environment at 21 °C the values were c. 90%
for PHB [115].
PHA biodegradation has been evaluated in soil. Gómez et al. reported mineralization
values of c. 70% for PHA (injection molding samples) in soil after 650 days at 20 °C [215]. A
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more recent study reported mineralization values of c. 90% for PHA (powder) in soil after 150
days at both 25 and 37 °C [210].
2.17.8 Poly(butylene adipate-co-terephthalate) – PBAT
PBAT is a co-polyester synthesized from 1,4-butanediol (BDO), adipic acid (AA), and
dimethyl terephthalate by a polycondensation reaction. Adipic acid and BDO polymerize to
produce their own polyester and water. Dimethyl terephthalate and BDO react to form their
own polyester and methanol. The resulting polyester then reacts with the polyester of AA
and BDO using tetrabutoxytitanium as a catalyst for the transesterification. The reactions
are carried out at temperatures higher than 190 °C, under high vacuum, and usually require
long times [577]. PBAT has a Tg of c. 30 °C and Tm of c. 106 °C. PBAT has good toughness
and ductility, biodegradability, and is flexible. However, higher production costs, coupled
with lower mechanical and heat resistance in comparison to common fossil-based plastics,
has hindered PBAT development and acceptance in the consumer market [577]. These
shortcomings can be overcome by blending PBAT with other biodegradable polymers. For
example, blends of PBAT and PLA demonstrated higher yield stress, modulus, and
rheological properties than those of neat PBAT [578]. PBAT is widely used for agricultural
mulch films, and also for packaging applications including trash bags, shopping bags,
wrapping films, and disposable food containers [579].
The reported abiotic mechanisms of degradation associated with PBAT are primarily
mechanical, photodegradation, thermal, and hydrolysis. Mechanical degradation is, in
general, associated with the entire spectra of biodegradable polymers; in the case of PBAT,
erosion is a common situation due to its main application in agricultural mulch films.
Photodegradation has been reported as the main abiotic mechanism of degradation for PBAT
agricultural mulch films. A crosslinking effect as a result of exposure to sunlight has been
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reported; the effect is known to delay the biodegradation rate of PBAT by decreasing the
chain mobility of the polymer [141,175,276].
The addition of aliphatic acids to aromatic polyesters improves the water uptake and
hydrolysis of these polymers. For example, by adding adipic acid to poly(butylene
terephthalate), PBAT is obtained and it can undergo faster hydrolytic degradation. However,
PBAT still offers more resistance to chemical hydrolysis than aliphatic polyesters, such as
PLA, due to the steric hindrance of the large aromatic ring repeating units.
PBAT is depolymerized through chemical and enzymatic hydrolysis into adipic acid,
1,4-butanediol, and terephthalic acid (Figure 2.14). Then each compound is bioassimilated
or undergoes a redox reaction to feed the TCA cycle, as shown in Figure 2.22. The adipic
acid pathway is through adipyl-CoA, and 1,4-butanediol is converted to succinic acid and to
succinyl-CoA. Several bioassimilation pathways have been reported for terephthalic acid.
The most probable, in the case of PBAT, seems to be the transport of terephthalic acid
through the cell membrane, followed by degradation to protocatechuic acid and then through
the pyruvic acid pathway to Acetyl-CoA to enter the TCA cycle [564].
138
Figure 2.22 Tentative biodegradation pathway for PBAT in aerobic conditions. Adapted from
[564,580].
In terms of biodegradation and enzymatic activity at mesophilic conditions, PBAT has
been reported to be degraded by cutinases [298], lipases [331], and PBAT hydrolases [260].
As for chemical hydrolysis, enzymatic activity is affected due to the presence of aromatic
groups that make enzyme accessibility more difficult for scission of the ester bonds that are
close to these groups [50,51]. The presence of the aromatic ring has been associated with a
decrease of the enzymatic activity by creating a steric impediment to access the active site of
the enzymes. Butanediol-terephthalate bonds have been reported to be hydrolyzed at a lower
rate in comparison to adipate-butanediol bonds [81].
Low values of enzymatic activity reported for the actinobacteria Rhodococcus fascians
in comparison to a mesophilic PBAT-degrading fungus show that both the type of enzyme
139
and the microorganism producing the enzyme play major main roles in activity [259,260].
These results could be associated with the favorable conditions offered to microorganism
populations in soil environments in the mesophilic range. The enzymatic hydrolysis of PBAT
by a fungal strain generated terephthalic acid, adipic acid, and 1,4-butanediol, as identified
by mass spectroscopy [260]. Furthermore, enzymatic activity of PBAT hydrolase by Bacillus
pumilus on PBAT showed degradation products as adipate, 1,4-butanediol, and terephthalate
[81].
Crosslinking due to exposure to UV-radiation treatment has also been shown to
decrease the enzymatic activity against PBAT, due to the reduced flexibility of the polymer
chains after crosslinking [581].
PBAT has been reported to be degraded in soil environments or soil in laboratory
conditions (Table 2.9). In general, rates of biodegradation at mesophilic conditions are low.
Biodegradation studies of PBAT showing CO2 and mineralization in simulated and controlled
media in the mesophilic range are limited [140]. Studies in more controlled environments
like culture and/or buffer media are more commonly focused on identification of microbial
activity and/or enzymatic activity towards PBAT. However, identification of extracellular
enzymes able to degrade PBAT is relatively limited in comparison to those for common
aliphatic polyesters. Most of the environments assessed for PBAT degradation are
agricultural soils. CO2 production from PBAT in soil environment media has been reported,
with mineralization values of c. 10% after six weeks [140].
A novel approach by Zumstein et al. [140] demonstrated the mineralization of PBAT
13C to 13CO2, with higher values of mineralization for 13C derived from depolymerization of
the adipate structure, and lower values of mineralization associated with depolymerization
of the aromatic terephthalate fraction. This finding is indicative of the increased complexity
of aromatic polyesters towards depolymerization and assimilation. On the other hand, the
140
presence of the aromatic component in the co-polyester was shown to improve the overall rate
of biodegradation, even in the mesophilic range like in the soil environment evaluated [140].
PBAT films with 1% of a chain extender had low mineralization values of c. 20% after
180 days in soil at 28°C [218]. The effect of the chain extender on delaying Mw reduction and
biodegradation was evident.
An interesting outcome of biodegradation studies for PBAT in a soil environment is
that the degradation products have been shown to be harmless to the microbial population
[582]. Although biodegradation can be a longer process in the mesophilic environment, and
the formation of PBAT degradation products does not affect the quality and health of the soil
and its microbial population, development of some microorganisms over others can be
modified [583].
2.17.9 Poly(urethanes)s – PUs
PUs are synthetic plastics, insoluble in water, and produced by the condensation
reaction of polyols and polyisocyanate having urethane bonds [49,584]. Polyisocyanates and
polyols react with a chain extender to give polyurethane polymers with alternate soft and
rigid segments. Polyol forms the soft segment and can be obtained from polyester or polyether
polyols; whereas the rigid segment is derived from the isocyanate and chain extender, and
has restricted mobility compared with the soft polyol segment (Figure 2.23) [495]. The rigid
segment is considered the crystalline region and the soft segment the non-crystalline or
amorphous region of PUs [493,585]. Depending on the polyol used, the resulting PU can be
identified as polyester PU or polyether PU. The resulting properties and degradation
behavior are dependent upon the selection and chemistry of the soft segment [586].
141
Figure 2.23 Soft and hard segment of the poly(urethane)s structure. Adapted from [495].
Poly(urethane)s are used in the medical, construction, and automotive fields, among
others. Typically, products that contain PUs include furniture, paints, fibers, flexible foams,
rigid foams, coatings, adhesives, synthetic skins, sutures, and tissue scaffolds [494,587,588].
Poly(urethane) elastomers (thermoplastic) are used in the medical field due to their high
elasticity and toughness compared with other elastomers [584,587]. The good mechanical,
thermal, and electrical properties of PUs allow these polymers to offer good adhesion for
coatings, tensile strength, and abrasion resistance for several uses [587]. Poly(urethane)
foam are a typically example of thermoset PUs [494].
Early studies demonstrated that PUs with long repeating units and hydrolytic groups
were susceptible to biodegradation [589]. This review concentrates on polyester PUs. The
ester bond of polyester PUs is susceptible to hydrolytic degradation and can be catalyzed with
the help of extracellular enzymes. The extracellular enzymes for PU degradation have a
hydrophobic area, which assists in attaching onto the polymer surface [585,587]. Microbial
attack of PUs can occur by action of extracellular hydrolases such as ureases (3.5.1.5),
amidases (3.5.1.4), proteases, and esterases (Figure 2.14). The cleavage site and the product
of the breakdown is dependent on the type of the enzyme acting during depolymerization
(Figure 2.24). Adipic acid and diethylene glycol were reported as degradation products by
the action of extracellular enzymes on polyester PUs; however, no identification of the
142
isocyanate hard segment byproducts was reported [315,316]. Later work by Shah et al.
reported the probable presence of a hydrolyzed portion of the hard segment, detected by FTIR
spectrum, when polyester PU was attacked by both Bacillus subtilis MZA-75 and
Pseudomonas aeruginosa MZA-85 [246]. Furthermore, the mixing of esterase and amidase
has been reported to hydrolyze the hard segment via the urethane bonds [437].
A bacterial esterase was identified to degrade ester PUs by acting in a two-step
reaction: first, a hydrophobic adsorption of the enzyme on the surface of the PU; and second,
the hydrolysis of the ester bonds of the PU [316]. Studies of enzymatic activity have shown
that the rate of biodegradation decreases with decreasing ester content, indicating the impact
of the esterase activity as relevant for PU depolymerization [584]. Fungal communities have
been identified to degrade PU to some extent [481].
Figure 2.24 Sites of scission for urethane bonds in extracellular enzyme function. Adapted
from [585].
A tentative route for metabolism of the soft segment (PEG) is presented in Figure
2.25 for PU derived from ester.
143
Figure 2.25 Tentative metabolism pathway for poly(urethane)s derived from esters. Adapted
from [590].
Biodegradation studies showing CO2 evolution or mineralization in mesophilic
environments for PUs are limited. Most of the reported studies on PUs are for enzymatic
activity of both fungi and bacteria. More investigations of the abiotic degradation process of
PUs prior to the biotic degradation stage, such as hydrolysis or photodegradation, in the
mesophilic range would help determine whether PUs derived from esters are fully
biodegradable in soil, home composting, industrial composting, or water environments. The
biodegradation of polyester-PUs studied under mesophilic composting conditions resulted in
mineralization between 5 and 43% after 45 days of testing, and this wide range was
attributed to the different chemical structures of the PUs [591]. A high content of the hard
segment led to decreased biodegradation rates and mineralization, whereas biodegradation
increased as the amount of diol carbon chains of the polyol (soft segment) increased. The hard
segment composition in PUs was presented as a more dominant effect than the crystallinity
or surface properties during PU biodegradation in composting [591]. The presence of aromatic
144
diisocyanates decreased the rate of biodegradation in comparison to PUs with aliphatic
diisocyanates [591].
Biodegradation of PU films during the Sturm test showed high CO2 evolution at 30 °C
for 28 days in comparison to the control [240]. Also, a Sturm test revealed the production of
CO2 during the enzymatic hydrolysis of PU films by Bacillus subtilis MZA-75 and
Pseudomonas aeruginosa MZA-85, hydrolyzing the ester portion in 1,4-butanediol and adipic
acid products [246–248]. This result indicated that Bacillus subtilis was able to hydrolyze
and assimilate the intermediates as carbon sources with final mineralization. An interesting
outcome of the reported enzymes attacking ester PUs is the evidence of the presence of
membrane-bound enzymes, besides extracellular enzymes. For an esterase not secreted to
the culture medium, its high hydrophobicity was reported as the most probably cause for its
membrane-bound characteristic [547].
A new approach is the development of non-isocyanate PUs (NIPU). NIPU are a
promising and more sustainable alternative for traditional PUs [592,593]. However, studies
in this area looking at degradation and biodegradation are still limited. Production and
biodegradation assessment of polyhydroxyurethane, a NIUP based on cyclic carbonate and
polyamine, was reported by Ghasemlou et al. [229]. Mineralization values for film samples
reached c. 40% after 120 days of testing in soil conditions.
2.17.10
Poly(vinyl alcohol) – PVOH
PVOH is a synthetic, water-soluble polymer produced by partial or complete
hydrolysis of polyvinyl acetate. Unlike other polymers, PVOH is not synthesized from the
polymerization of its monomer (vinyl alcohol), due to the unstable nature of the high density
of hydroxyl groups in the monomer. Polyvinyl acetate is first synthesized by the
polymerization of vinyl acetate and then subjected to saponification wherein the ester groups
of vinyl acetate are replaced by hydroxyl groups in the presence of caustic soda [594].
145
Different grades and properties of PVOH are available, depending on the degree of hydrolysis
and the variation in initial length of the vinyl acetate polymer. PVOH is odorless and non-
toxic in nature, has excellent resistance to aroma and gases, is resistant to solvents and oil,
has good optical and adhesive properties, and film forming capacity [595]. In terms of
disadvantages, PVOH is expensive, and mechanical properties are highly conditioned by the
presence of water or humidity so it needs to be blended with other polymers to achieve more
desirable properties [596]. Due to its good adhesion to other hydrophilic surfaces, PVOH is
used widely in emulsifiers, binders, and hydrogels for a broad range of industries, including
textile, paper sizing, fabrics, and packaging films as a protective film for laundry and dish
detergents. The applications are not limited and extend to the biomedical, cosmetic, and food
packaging industries [597,598].
The degree of solubility of PVOH in water can be tailored, depending on the amount
of OH groups and remaining acetate bonds.
Besides the abiotic mechanism of biodegradation, PVOH could be considered as
partially biodegradable since the number of microorganisms and enzymes identified to
biodegrade it is rather scarce in comparison to polyesters.
Biodegradation of PVOH has been reported to start from random chain scission where
the action of oxidative enzymes catalyzes the break of the carbon backbone. Mostly
dehydrogenases or oxidases are responsible for the carbon-carbon bond scission. Hydrolases
or aldolases have been reported as responsible for the chain scission of the hydroxyl group
(Figure 2.14). Furthermore, a two-step process has been proposed for the enzymatic
degradation of PVOH: the first step, by action of PVOH oxidases, involves the oxidation of
hydroxyl groups to form diketone or monoketone structures; and the second step involves
hydrolysis of the carbonyl structure formed by oxidized PVOH hydrolases [599].
146
Since PVOH is a water-soluble polymer, its biodegradation has been studied mostly
in aqueous media. The identified microorganisms and enzymes able to biodegrade PVOH are
associated mainly with contaminated environments, such as waste sludge, which are
common end-of-life scenarios for PVOH.
Abiotic degradation of PVOH by UV/chlorine oxidation via generation of active free
radicals has been investigated; in acidic media the efficiency was higher due to the higher
ratio of [HOCl]/[OCl-] [600]. The abiotic degradation of PVOH by photocatalytic oxidation or
radiation and ozone also has been reported [478,601].
Published works have identified that microorganisms able to biodegrade PVOH are
mostly from the genus Pseudomonas [602]. Also, many PVOH degradation pathways have
been proposed for different bacteria such as Alcaligenes and Pseudomonas species [603].
These routes include scission of the polymer chain by an extracellular oxidase
(dehydrogenase), followed by aldolase and hydrolase reactions, releasing compounds such as
acetic acid and hydroxyl fatty acids that can be incorporated into the -oxidation and TCA
cycle, respectively [100]. In Figure 2.26 is presented a tentative metabolization route for
PVOH [439].
As stated, scarce mineralization was reported for PVOH films in water conditions,
with ~ 10% after 100 days of testing at c. 30 ºC [232].
147
Figure 2.26 Proposed pathway for metabolism of PVOH. Adapted from [109,603].
2.18 Final insights and remarks
Addressing plastic pollution has become one of the main concerns of our modern
society. The impacts of plastic waste in terms of global climate change, health, and social
effects, circular economy, sustainable use of resources and production, and improved waste
management systems have garnered the attention of industry, government, and NGO
stakeholders and the society in general. The development of new plastic waste pacts and
commitments to curb the use of virgin plastic are ongoing globally, with targets for 2025 [21].
However, the damage to ecosystems has already created deleterious impacts, which will
require forward-thinking actions to remediate, to mitigate, and to avoid permanent damage
[9].
The development of biodegradable polymers derived from both bio- and fossil-based
resources has transcended from the lab scale to commercial applications in the last two
decades, and these polymers have become an option for packaging and consumer goods
148
applications to mitigate the impact of plastic waste. However, as with any material created
by society, biodegradable polymers also must reach a waste management end-of-life to avoid
a rebound effect on creating additional pollution.
The degradation process for biodegradable polymers starts by the action of external
abiotic and biotic factors. The main abiotic mechanisms of degradation associated with
mesophilic environments are photodegradation, mechanical degradation, and chemical
hydrolysis. Photodegradation in the presence of O2 introduces modifications during the
degradation of biodegradable polymers in specific environments such as agriculture soils,
inducing a dual effect: chain scission that contributes to the degradation and also crosslinking
that act to delay the process. The initial deterioration of the polymer structure enhances the
mechanical degradation, generating micro and nano plastics but not guaranteeing
biodegradation. Chemical hydrolysis is the crucial mechanism for the large majority of
biodegradable polymers since most of them contain ester bonds that are prone to water
attack.
The formation of biofilms, the stage prior to the release of extracellular enzymes,
affects the whole dynamic of the degradation process, as discussed. Since biofilms create an
extra layer on the polymer surface and potentially affect water diffusion during chemical
hydrolysis, a better understanding of biofilm formation and its effect on water diffusion and
bulk erosion are needed.
Extracellular enzymes act at the surface level of polymers, making enzymatic activity
a surface erosion process. As presented in this review, the main groups of enzymes reportedly
able to break chemical bonds in polymers belong to the esterase group (amidases, cutinases,
esterases,
lipases, and PHA depolymerases), proteases (specific for PLLA), and
oxidoreductases (for PVOH and PU). Recent advances in the identification of protein
sequences and residues, structural domains, mechanisms of substrate binding, kinetic
149
analysis, and the presence and effects of cofactors have provided a better understanding of
enzymatic activity on biodegradable polymers. However, a better understanding of
bioassimilation and mineralization is still needed at the biochemical level of monomer
compounds produced from the chemical and enzymatic hydrolysis of biodegradable polymers.
In terms of polymer properties, the key bulk properties affecting biodegradation in the
mesophilic range are stereochemistry, crystallinity, and Mw, which are tailored for each
application. The amorphous region offers the optimal conditions for chemical hydrolysis due
to the easy diffusion of water and also for exo and endo enzymatic attack by extracellular
enzymes. Microorganisms start the assimilation process when low Mw compounds such as
dimers and monomers are released. Since biofilm formation, microbial colonization, and
enzymatic activity are surface related processes, the key surface properties of polymers
impacting biofilm formation and colonization are hydrophobic/hydrophilic balance,
roughness, and surface energy parameters.
This review has summarized the enzymes and microorganisms (e.g., bacteria, fungi,
and actinomycetes) isolated from several environments and showing activity towards
aliphatic, aliphatic-aromatic polyesters, PUs derived from esters, and PVOH. Usually the
identification of microorganisms and/or enzymes involves techniques, such as culturing,
where the polymer is the solely source of carbon for the biotic process. These studies provide
unique insights on enzymatic activity and pathways of degradation. However, natural
environments introduce far more complexities to the degradation process, creating a dynamic
that undoubtedly affects the rate of degradation. Microbial consortia have demonstrated an
increased efficiency for elimination of toxic metabolites in comparison to pure cultures.
Studies showed that some microorganisms are directly involved in the degradation process,
while other microorganisms showed activity towards eliminating toxic metabolites excreted
by the first ones. However, besides symbiotic, mutual, and synergistic interactions, efficiency
150
differs among microbial consortia. Complex tracking of microorganism population dynamics
during biodegradation should provide better insights on the real pathways of degradation
and assimilation of these polymers in actual environments. Research in the areas of
biostimulation (addition of specific nutrients to the soil, compost, or water environments to
stimulate the activity of naturally occurring microorganism populations), bioaugmentation
(addition of specific microorganisms to increase the biodegradation rate of an indigenous
microbial population), and engineering of enzymes (modification of enzymes to reach specific
reactions) are needed to address the complexities associated with microbial consortia
involved in the biodegradation process, extracellular enzymes, and biocatalytic cascades of
enzymes.
Standards, methodologies, and techniques have been developed to assess the
degradation of polymers in the environments as discussed. Some are more focused on
evaluating the degradation of mechanical properties and the mass loss due to various factors
of abiotic mechanisms. However, the assessment of CO2 or O2 and ultimate mineralization
must be the definitive assessment to determine the extent of biodegradability in a specific
environment. The use of complementary techniques, such as carbon tracking and Mw
reduction, constitutes important tracking parameters that must be incorporated when
evaluating biodegradation to the mineralization level.
Many works reported the CO2 or mineralization for the biodegradable polymers
available in the market in soil, home composting, and aquatic environments at mesophilic
conditions using several standards. From the aliphatic polyesters group (e.g., PGA, PEA,
PLA, PCL, and PBS), chemical hydrolysis has been reported to be the main controlling step.
Other aliphatic polyesters, such as PCL, PBS, and PBSA, were consistently reported to
biodegrade in soil conditions. For the aliphatic-aromatic polyesters (PBAT and PBST),
reports of PBAT biodegradation in soil and marine environments are limited. The
151
degradation of the natural polyesters PHB and PHBV in aquatic environments was reported
extensively, showing the high level of biodegradability at mild conditions. For the PUs, the
presence of the soft segment offers availability for enzymatic attack and biodegradation with
mineralization at some extent; however, the bioassimilation pathway of the hard segment
has not been well identified and/or described.
New, innovative methods to tailor the biodegradation of biodegradable polymers
through creation of novel polymers with tailored biodegradation [604], biostimulation,
bioaugmentation, and addition of natural enzymes [605–607] or modified enzymes [608] are
opening new routes to accelerate the biodegradation process. However, these new methods
must be connected to standards to fully track the biodegradation process and the end
products in the environment, so that further insights on biodegradation pathways of
polymers can be elucidated.
152
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CHAPTER 3: BREAKING IT DOWN: HOW THERMOPLASTIC STARCH
ENHANCES POLY(LACTIC ACID) BIODEGRADATION IN COMPOST - A
COMPARATIVE ANALYSIS OF REACTIVE BLENDS
3.1
Abstract
Poly(lactic acid) (PLA) is a sustainable, bio-based, and industrial compostable polymer
with a recalcitrant abiotic degradation phase limiting its organic recovery to well-managed
industrial composting facilities. We present a methodology to biodegrade PLA in industrial
and home composting settings fully. Thermoplastic starch (TPS) and PLA were reactively
blended by adding a chemical modifier and peroxide radicals to obtain a PLA-g-TPS blend by
twin screw extrusion and later processed into films by cast extrusion. Biodegradation of the
films was investigated using a direct measurement respirometer (DMR) for 180 days by
tracking the CO2 evolution in compost media at 58 and 37 °C, and the molecular weight (Mn)
reduction was measured by size exclusion chromatography. The hydrophilic nature of TPS
and its role as a nutrient source accelerated the degradation of PLA in both abiotic and biotic
phases of the composting process. The kinetic curve of Mn reduction showed the positive effect
of TPS on accelerating PLA hydrolysis during the lag phase in both mesophilic and
thermophilic conditions due to increased chain mobility. This work unlocks the capability of
PLA-based films to be successfully composted in industrial and home composting without
compromising its desired properties for applications in everyday life.
3.2
Introduction
Poly(lactic acid) (PLA), a bio-based polymer derived from renewable resources with
end-of-life scenarios such as recyclability and industrial compostability, is considered a green
and sustainable polymer and an alternative polymer to displace the ever-growing fossil-based
plastic pollution [1]. PLA has been used in medical applications, but its use has mainly grown
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in food packaging applications [2,3]. PLA and its products are labeled and marketed as
industrial compostable [4,5], so it can be collected with the organic fraction of municipal solid
waste (OFMSW) routed to industrial composting. Marketing bio-based and compostable
plastics, such as PLA, fuels the assumption that they can be discarded with the organic
fraction and treated and recovered in the same manner [6,7]. However, some challenges arise
when these polymers cannot fully disintegrate in the same time frame as readily
biodegradable organic fraction wastes, such as food, starch, and cellulose, leaving fragments
at the end of the compost cycle. Mostly, this situation occurs due to the slow abiotic
degradation controlling phase, such as in the case of PLA, associated with inherent
properties, such as high initial molecular weight (Mn) and crystallinity (Xc). Although
composting of the organic fraction is accomplished in industrial composting facilities, the
infrastructures have not been precisely designed to keep bio-based and compostable plastics
in mind. Therefore, some composters are not keen on accepting these products [8–10].
Standards are available to delineate what conditions polymers should meet to be
certified as compostable in industrial composting operations [11]. Conditions for home and
industrial composting are different since temperatures and humidities encountered in these
two environments differ drastically. Since the material volume, reached temperature, and
abundance of microorganisms are lower in home composting, it is very challenging to
maintain the necessary conditions for several days.
Regardless of whether the composting is done at home, in a community backyard, or
in an industrial facility, the process must ensure a succession of microbial communities (i.e.,
mesophiles–thermophiles–mesophiles) and the corresponding temperature regimes to
operate. These factors ensure and guarantee the safety and quality of the composting process
and the final product. Furthermore, this process and high temperatures are crucial to
degrade a polymer, such as PLA with a high glass transition temperature (Tg c. 58 ℃) and
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whose initial breakdown is based on being able to traverse a rapid abiotic stage to reduce the
weight average molecular weight number (Mn) low enough (≲10 kDa) so that the
microorganisms can use PLA oligomers as a food source [12]. Furthermore, the industrial
composting process is highly controlled. Efficient control of temperature, moisture, and
airflow is exercised to achieve complete waste biodegradation. Any deviation from the
required conditions in terms of turning frequency of the piles, consistent mixing to maintain
adequate aeration and pH, and reduction of the composting process can lead to the formation
of anaerobic pockets or increase the time needed for PLA to initially degrade through the
abiotic reduction process and delay the complete breakdown of PLA, thereby leaving behind
partially degraded or whole packages at the end of the composting cycle. Additionally,
polymer fragments at the compost pile surface could take longer to degrade since they are
not exposed to the same conditions. This is especially relevant for plastics that can constantly
resurface during pile turning due to their lower density. So, incomplete biodegradation of
PLA as well as other compostable plastics can impact the sale of compost by failing to abide
by the agricultural compost requirements. To avoid this problem, some composting facilities
are apprehensive about incorporating PLA with organic wastes [9]. Methods able to
accelerate the biodegradation of PLA in industrial composting facilities and possibly expand
its biodegradation efficiency in home/backyard composting are highly sought.
One potential way to improve PLA properties and its degradation rate in ambient
conditions is by blending it with different biodegradable polymers to improve specific
properties such as toughness, flexibility, and ductility and promote its complete
biodegradation post-use [1,13–15]. One such polymer that can be used to blend with PLA to
improve its biodegradability is starch, which is low-cost, renewable, non-toxic, readily
available, and 100% bio-based. Thermoplastic starch (TPS) is a plasticized polymer used to
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produce blends with different polymers. Starch undergoes rapid biodegradation by enzymatic
hydrolysis to produce glucose, which is further assimilated to produce CO2 and H2O in
compost or soil environments [16]. To process TPS efficiently and to blend and derive good
interfacial adhesion between hydrophilic starch and hydrophobic PLA, compatibilizers are
generally used [17–19]. Blending PLA and TPS helps improve the oxygen and water vapor
barrier properties [20] and the elongation at break of the resulting matrix [21]. Furthermore,
the hydrophilic feature of TPS can enhance PLA’s sensitivity to humid environments and act
as an excellent initial nutrient source for the microbes when introduced into the compost
environment [22–24].
PLA is environmentally benign, to begin with, and the addition of starch derived from
non-human crop consumption creates a safe polymer. Biodegradation of PLA-g-TPS
ultimately will not leave behind traces of microplastics since degradation of the PLA evolves
to lactic acid traces and degradation of the starch evolves to glucose, two naturally occurring
and benign monomers that can be acted upon and utilized by the microorganisms. Adding
TPS can further help accelerate the low abiotic degradation rate of PLA at mesophilic
conditions, which otherwise takes longer. PLA-starch blends can open avenues for
home/backyard composting and address the growing concern about microplastics [25–28] and
their exo-toxicological effects [29,30] since their formation can be avoided.
In recent years, several studies have been conducted focusing on the biodegradation
of PLA in soil environments [31–34] by introducing starch in the PLA matrix, where the
biodegradation was determined by visual analysis, weight loss, or change in mechanical
properties but not CO2 evolution or Mn reduction. Biodegradation studies have also been
conducted where additives such as wood flour [24], coir [35], and montmorillonite [36] have
been blended with PLA, in addition to starch. Still, the additives failed to enhance properties,
and the resulting biodegradation was attributed to the presence of starch. However, a
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detailed assessment of PLA and TPS mineralization and Mn evolution is still lacking, and
there is a need to understand the role of starch in accelerating the biodegradation of PLA in
compost. In this work, we investigated the role of starch in expediting the biodegradation of
PLA under thermophilic and mesophilic conditions in a compost environment, and we
monitored CO2 evolution and quantified the changes in Mn throughout the test duration so
as to replicate real-life scenarios encountered under home composting and industrial
composting conditions.
3.3 Materials and methods
3.3.1 Materials
PLA resin, Ingeo™ 2003D with L-lactic content of 96%, was provided by NatureWorks
LLC (Minnetonka, MN, USA). Cassava starch was procured from Aldema LLC (Cooperativa
Agricola e Industrial San Alberto Ltda., Puerto Rico, Misiones, Argentina), with an amylose
content of 25 ± 6% wt/wt and moisture content of approximately 12%. Glycerol (>99.5%),
maleic anhydride (MA), dicumyl peroxide (DCP), and cellulose (~20 µm particle size) were
purchased from Sigma-Aldrich (Milwaukee, WI, USA). Except for the PLA resin, all materials
were processed in the condition they were received in. PLA pellets were dried at 50 °C
overnight at 67 kPa to avoid hydrolytic degradation while processing.
3.3.2 Preparation of PLA and thermoplastic starch (TPS) masterbatches and
films
Cassava starch and glycerol (70/30% wt.) were mixed and held for 12 h before
processing into the TPS masterbatch. The temperature profile for the TPS masterbatch was
set at 25/100/105/110/115/120/120/120/115/115 °C for each zone from feed to the die, with a
screw speed of 120 rpm. A ZSK-30 co-rotating twin screw extruder (Century, Traverse City,
MI, USA) [37] was used to process the various masterbatches; the screw length was 1,260
mm and the diameter was 30 mm, with an L/D ratio of 42:1. PLA, MA, and DCP were mixed
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prior to processing; PLA-g-MA (PLA with 2% wt. MA and 0.65% wt. DCP, based on PLA
weight) was processed in the twin screw extruder. The temperature profile for the PLA-g-MA
masterbatch was set at 140/150/160/160/160/170/170/170/170/160 °C for each zone from feed
to the die, with a screw speed of 120 rpm, feed rate of 70 g/min, and residence time of
approximately 3 min. The extrudate from the die was fed through a water bath and pelletized
using a BT 25 pelletizer (Scheer Bay Co., Bay City, MI, USA). The PLA-g-MA masterbatch
was kept in an oven at 50 °C for 3 h to remove any traces of moisture and was later
transferred to the freezer and stored at −15 °C until processed into films. The same procedure
was followed to produce a PLA control masterbatch. To obtain PLA-g-TPS, PLA (56% wt.),
TPS (30% wt.), and PLA-g-MA (14% wt.) were processed together to produce the final reactive
blend. The
temperature profile
for
the PLA-g-TPS masterbatch was set at
140/150/160/160/160/170/170/170/170/160 °C for each zone from feed to the die, with a screw
speed of 120 rpm, feed rate of 70 g/min, and residence time of approximately 3 min, which
was dried in a vacuum oven at 50 °C for 3 h to remove any residual traces of moisture. Figure
S1 of the supporting information depicts the processing of PLA-g-MA, PLA and TPS
masterbatches and the temperature profile for the twin screw extruder.
The masterbatch was introduced in a RCP-0625 microextruder (Randcastle
Extrusion Systems, Inc., Cedar Grove, NJ, USA) to produce cast films; the screw diameter
was 1.5875 cm, with an L/D ratio of 24/1 and volume of 34 cm3, and the temperature profile
was set at 140/150/160/160/160/170/168 °C from feed to die. The screw speed, nip roller, and
winding roller were set at 30 rpm, 50 rpm, and 12 rpm, respectively [38]. The same procedure
was used to process PLA films. Figure S2 shows the processing of PLA-g-MA, PLA and TPS
masterbatches into PLA-g-TPS film using the cast film extruder.
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3.3.3 Biodegradation test in simulated compost conditions
The biodegradability of PLA and PLA-g-TPS films was investigated by analysis of
evolved CO2 in a simulated composting environment using an in-house built direct
measurement respirometer (DMR) system [11,39]. The biodegradation tests were run at 37
± 2 °C and 58 ± 2 °C to simulate mesophilic and thermophilic environmental conditions,
respectively. The DMR chamber was maintained at specific temperatures, and a Li-COR® LI-
820 non-dispersive infrared gas analyzer (Lincoln, NE, USA) was used to measure the
evolving CO2 concentration from each DMR bioreactor. The relative humidity was
maintained at 50% ± 5% and the optimal airflow rate was set at 40 ± 2 sccm [40,41].
Compost was procured from the Michigan State University composting facility (East
Lansing, MI, USA). The compost was filtered using a 10-mm screen to remove any larger
lumps of material and then was conditioned at 37 ± 2 °C and 58 ± 2 °C for one week before
the respective test use. The procured compost was characterized for its physicochemical
parameters (presented in Table A3.1, Appendix 3B). When running the biodegradation test,
deionized water was added to the compost to regulate and sustain the moisture content at
around 50%. Vermiculite, an inert, inorganic hydrous phyllosilicate matrix, obtained from
Sun Gro® Horticulture Distribution Inc. (Bellevue, WA, USA), was mixed 1:4 (wt/wt) with
compost. Vermiculite does not affect the biodegradation test and helps provide better
aeration, improving the accessible space, thus improving the microbial activity [42].
Each bioreactor was loaded with 400 g of compost which was then mixed with 8 g of
film samples (cut into 1-cm2 pieces) or 8 g of cellulose. Experiments with film samples in
compost media were run in triplicate. Cellulose, a positive control reference, was used in the
test due to its high biodegradation nature and as requested by ASTM D5338-15(2021) [11].
Three bioreactors were run with compost only (blank) and without any film samples or
211
cellulose to identify the background CO2 evolution. CO2-free air (≲30 ppm CO2) was passed
through each bioreactor, and the evolved CO2 was measured for a set period. After completing
measurements for each bioreactor, the whole system was purged using CO2-free air to ensure
no residual CO2 was disturbing the baseline test [41]. The mineralization formula shown in
equation (3.1) was used to calculate the amount of carbon that was converted to CO2:
𝑀𝑖𝑛𝑒𝑟𝑎𝑙𝑖𝑧𝑎𝑡𝑖𝑜𝑛 % =
(𝐶𝑂2)𝑡 − (𝐶𝑂2)𝑏
𝑀𝑡 𝑥 𝐶𝑡 𝑥
44
12
𝑥 100
(3.1)
where (CO2)t is the average cumulative mass of CO2 evolved from the bioreactor
containing the sample, (CO2)b is the average cumulative mass of CO2 evolved from the blank,
Mt represents the total mass of the sample in the bioreactor, Ct is the total carbon content of
the sample and is derived from CHN analysis, 44 is the molecular weight of CO2, and 12 is
the atomic weight of carbon. The equation numerator represents the actual CO2 evolved from
the sample after accounting for the background compost activity, and the denominator
depicts the maximum theoretical CO2 the sample produces, i.e., when 100% of the sample
carbon is converted to CO2.
3.3.4 Hydrolysis experiment
A hydrolysis test method adapted from ASTM D4754-18 [43] was run for PLA films
at 58 ± 2 °C and 37 ± 2 °C to understand the hydrolytic degradation. The hydrolysis cell
consisted of a stainless-steel wire, glass beads, and a glass vial with cap. PLA films were cut
into small discs of 2-cm diameter, and ten such discs were strung into a stainless-steel wire
and separated by glass beads. The vial was filled with 35 mL of HPLC-grade water (J.T.
Baker, Center Valley, PA, USA). The water was preconditioned, and the hydrolysis cell was
stored at the same temperature. Triplicates of the PLA films were retrieved at predetermined
time intervals and dried before running size exclusion chromatography to assess the Mn
reduction.
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3.3.5 Size exclusion chromatography
The film samples were collected at specific intervals to determine the Mn, weight
average molecular weight (Mw), and molecular weight distribution (MWD). Samples were
weighed and dissolved in tetrahydrofuran (THF) solvent in a ratio of 2:1. THF solvent was
used as a mobile phase. A size exclusion chromatography (SEC) unit from Waters Corp.
(Milford, MA, USA) was supplied with an isocratic pump (Waters® 1515), an autosampler
(Waters® 717), a series of Styragel® columns (HR4, HR3, HR2), and a refractive index detector
(Waters® 2414). The detector was maintained at 35 °C, and a 1 mL/min flow rate was applied
for the mobile THF solvent. The Mark-Houwink constants of K = 0.000174 mL/g and α =
0.736 were used to determine the Mn and Mw of the PLA fraction in the samples. A detailed
description of the technique can be found elsewhere [44]. The data was analyzed using the
Waters BreezeTM 2 software.
3.3.6 Elemental analysis
The carbon, hydrogen, and nitrogen content of the PLA and PLA-g-TPS films was
calculated using a PerkinElmer Series II CHNS/O Elemental Analyzer (PerkinElmer Inc.,
Shelton, CT, USA). Approximately 2 mg of sample was weighed in a small tin capsule and
tested. Samples were tested in triplicates. The values are presented in Table 3.1.
Table 3.1 Percent of carbon, hydrogen, and nitrogen content by weight of cellulose and film
samples.
% Carbon
% Hydrogen
% Nitrogen
Material
Cellulose
PLA
42.50 ± 0.34
49.72 ± 0.19
PLA-g-TPS
48.20 ± 0.05
6.53 ± 0.05
5.72 ± 0.04
6.00 ± 0.04
0.04 ± 0.01
0.11 ± 0.07
0.05 ± 0.01
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3.3.7 Statistical analysis
The statistical analysis was conducted using MINITABTM software (Minitab Inc.,
State College Park, PA, USA). The statistical significance at p < 0.05 was evaluated using
one-way ANOVA and Tukey-Kramer test. Data is reported as means ± standard deviation.
3.4 Results and discussion
PLA films have a recalcitrant abiotic degradation stage, limiting its deployment to
well-managed industrial composting facilities. Facilities that rotate compost in a very short
period (c. ≲2 months) and do not follow a practice of allowing a full cycle of mesophilic,
thermophilic, mesophilic, and mature phases are not able to fully biodegrade PLA, leaving
residual at the end of the period [8]. Previously, we developed PLA-g-TPS reactive blend films
with suitable thermal, mechanical, and barrier properties for several applications, including
packaging and agriculture, to be recovered through composting [37,45]. Figure A3.3 b,
Appendix 3C shows the SEM images for PLA-g-TPS film showing uniform dispersion of the
TPS phase in the PLA matrix by reactive blending as opposed to physical blending. Here, we
present the biodegradation of these PLA-g-TPS films in simulated composting at
thermophilic and mesophilic conditions using a DMR unit and report the Mw reduction and
thermal properties.
3.4.1 CO2 evolution and mineralization of PLA and PLA-g-TPS films
PLA-g-TPS films were evaluated alongside PLA in simulated composting conditions
at 58 ± 2 °C and 37 ± 2 °C to understand the effect of reactive blending TPS on PLA
degradation. Figure 3.1a and b present the CO2 evolution and % mineralization of the blank
(compost only), cellulose, PLA films, and PLA-g-TPS films in compost at 58 ± 2 °C. The blank
showed maximum CO2 evolution of 28.3 g at day 180. The positive control cellulose achieved
a 40.8 g for CO2 evolution and showed maximum mineralization of c. 100%. No lag phase
214
was observed for cellulose, as it is a readily available food source for microorganisms and
easily biodegradable. Cellulose is degraded by the action of a battery of enzymes that work
simultaneously and synergistically. Cellulases catalyze the hydrolysis of β-1,4-linkages in the
cellulose [46]. The action of exoglucanases and endoglucanases on the ends and at random
internal sections of cellulose’s amorphous region produces varying lengths of cello-
oligosaccharides, which are then hydrolyzed by glucosidases to produce glucose [47]. Glucose
is finally converted to CO2 through a series of further cycles. Previous research has shown
fungi, a few bacteria species, and actinomycetes in compost and the soil environment produce
cellulase and are completely involved in the degradation of cellulose [48–52].
The biodegradation curve for PLA showed a lag phase of around 20 days, attributed
to the initial abiotic hydrolysis phase. Ester bonds of PLA are cleaved during the hydrolytic
abiotic degradation phase due to its susceptibility to water. As hydrolytic degradation
proceeds, PLA chains are broken into smaller fragments, releasing low Mw oligomer
populations. These lactic acid oligomers are available for microbial assimilation, releasing
CO2 and water, which can be observed during the biotic degradation phase. A maximum CO2
evolution of 39 g and mineralization of c. 77% was observed for PLA over the test duration of
180 days at 58 °C.
PLA-g-TPS films showed a CO2 evolution of 42.8 g and a maximum mineralization of
100% over the test duration of 180 days. PLA-g-TPS did not show any lag phase compared to
the 20-day phase for PLA. This finding can be ascribed to the presence of TPS acting as an
initial food supply for the microorganisms until the PLA undergoes hydrolysis,
fragmentation, and is depolymerized into small fragments available for further assimilation.
Starch, like cellulose, is a natural hydrophilic, biodegradable polymer and can be used up
almost immediately as the microorganisms get acclimatized to the compost environment. The
presence of hydroxyl groups in TPS contributes to and enhances the biodegradation of PLA
215
by further enabling the disintegration process of PLA-g-TPS films. Several research studies
have shown that introduction of starch increases the water absorption characteristics of a
matrix. Maran et al. [53] demonstrated improved water absorption due to the presence of
starch in specimens buried in soil for degradation studies. Also, the presence of starch
increases the hygroscopic characteristics of films, which favors water absorption, thereby
providing suitable conditions for fast hydrolysis and microbial invasion and colonization
[34,54,55]. The presence of amylopectin branched hydroxylated chains improves the water
penetration in PLA-g-TPS film matrices [33,56]. This sensitivity of starch to water helps in
the enzymatic hydrolysis to glucose.
Figure 3.1 CO2 evolution and % mineralization of PLA and PLA-g-TPS films in compost at
58 °C (a & b) and 37 °C (c & d). The shade in the background for each material represents
the standard error between replicates.
216
Figure 3.1c and d depict the cumulative CO2 evolution and % mineralization for
cellulose, PLA films, and PLA-g-TPS films at 37 °C to evaluate the biodegradation behavior
in mesophilic conditions. Cellulose achieved mineralization of c. 86% in 50 days and evolved
around 37 g of CO2 by the end of 180 days. There was no lag phase for cellulose, as also
observed at 58 °C.
Bioreactors containing compost and PLA produced c. 22 g of CO2 in compost around
180 days. In contrast, blank bioreactors evolved c. 26 g of CO2, which is translated to negative
mineralization in the case of PLA. In a blank bioreactor, the microbes can easily utilize the
organic matter in the compost, whereas in a bioreactor containing PLA, the presence of PLA
leads to a reduction in the working efficiency, which translates to reduced carbon conversion
as a collective. The negative mineralization values are generated as an artifact and indicate
that the blank bioreactors produce more CO2 than the bioreactors containing PLA samples.
This could mean that PLA offers a physical hydrophobic barrier to water and air, and the
microbes have difficulty accessing the carbon source as nutrients. It could also indicate that
there is non-uniform mixing of the compost and PLA samples in the bioreactors, which could
additionally hinder the microbes. Any change in the optimal conditions for the biological
activity, either due to an insufficient supply of water (dryness of the compost), excess of water
(agglomeration), or obstruction in the airflow passage, can create an unfavorable
environment for microbial development. The negative mineralization values in the case of
PLA do not necessarily imply the absence of hydrolytic degradation or enzymatic activity due
to the action of extracellular enzymes secreted by the microbes but more like inhibition of the
microbial activity due to the hydrophobic layer barrier created by the presence of the sample.
The lower values of CO2 evolution in the case of PLA indicate that the samples are still
undergoing hydrolysis and are yet to be reduced to a point (≲10 kDa) needed to activate the
217
biodegradation stage where the microorganisms can start assimilating low Mw PLA, such as
oligomers, dimers, and monomers for their biochemical processes. The hydrolytic degradation
progresses very slowly when PLA is exposed to temperatures lower than Tg (c. 60 °C) due to
scarce chain mobility of the amorphous fraction and low water diffusion in the crystalline
fraction for effective bulk erosion [12,57,58]. PLA segments have little to no mobility and are
not flexible below Tg, preventing diffusion or attack by water. Since the initial and rate-
limiting step in PLA degradation is chemical hydrolysis, the lack of chain scissions, perhaps
accompanied by simultaneous low surface colonization by microorganisms due to the
hydrophobic surface [56], could have further prevented penetration by water molecules and
slowed down the hydrolysis rate. Further attestation to this rationale is the lower CO2
evolution values obtained for PLA (22 g) compared to the blank (26 g). The above reasons, as
well as the lower chemical hydrolysis rate of PLA, can account for lower CO2 evolution as
compared to the blank.
In the case of PLA-g-TPS, c. 58% mineralization was observed by the end of the test
(180 d) with an upward mineralization trend, since starch, like cellulose, is a readily available
nutrient for the microbial species in the compost [59]. No lag phase was seen in the case of
PLA-g-TPS films, which evolved 34.2 g of CO2 by the end of the test. Ho and Pometto [22]
documented mineralization of c. 70% for starch alone at 28 °C after 98 days and a higher
mineralization rate with increasing temperature. The mineralization for PLA-g-TPS films at
mesophilic conditions corroborates the research presented earlier [23,60,61]. The
hydroxylated chains in TPS impart hydrophilicity to PLA, pave the way for water dispersion
and hydrolysis in the blend matrix and support microbial development and growth [33,62].
The hydrophilic characteristics further stimulate microbial colonization and biofilm
formation on the surface of PLA-g-TPS films. This is further corroborated by changes in the
218
roughness and contact angle for PLA and PLA-g-TPS film due to the addition of TPS, as seen
in Figure A3.4 and Figure A3.5, Appendix 3D.
Starch is enzymatically hydrolyzed by the action of extracellular enzymes, namely
/-amylase belonging to the glycoside hydrolase family. Glucose produced as a result is then
converted to pyruvic acid by glycolysis. Pyruvic acid is a precursor for the tricarboxylic acid
cycle for energy production [63]. Once the microorganisms use starch as their food source,
fragmented PLA films (holes and cracks) are left behind. These structural discrepancies in
the form of macroscopic fractures aid in the biodegradation process. Starch biodegradation
and PLA chemical hydrolysis processes complement each other, resulting in improved
biodegradation of a starch-PLA mixture compared to PLA alone. Usually, at higher
temperatures, such as industrial composting conditions, these complementary processes
happen concurrently, and hence an improved degradation is seen for PLA. Figure 3.1b and
d show that adding TPS to PLA not only accelerates the disintegration of PLA-g-TPS films
in industrial composting but also opens a venue to make PLA blends biodegradable in home
composting conditions if key parameters such as aeration and moisture are controlled.
3.4.2 Molecular weight reduction rate
The Mn was measured, and the reduction rate calculated during the abiotic hydrolysis
stage was used to elucidate if the TPS fraction accelerated the hydrolysis of PLA besides also
enhancing the biotic degradation. PLA and PLA-g-TPS films were retrieved at specific time
intervals from the sampling bioreactors to track the reduction of Mw.
Figure 3.2 presents the MWD over time for PLA films and the PLA fraction in PLA-
g-TPS films in thermophilic and mesophilic conditions. For 58 ± 2 °C (Figure 3.2a & b), both
films show shifting of the peaks depicting a decrease in Mn and broadening of the peaks,
indicating an increase in dispersity (Đ) associated with hydrolytic degradation of the PLA
219
chains. The shifting of peaks in the abiotic phase reveals that the chemical hydrolysis is due
to random chain scission occurring in the bulk of the polymer and is not restricted to the
surface [64,65]. The change in the peaks from monomodal to multimodal after day 15 is a
characteristic feature of reconfiguration and crystallization of the newly crystalline region
resulting from the regrouping of newly formed short polymer chains formed at Mn ≲10kDa.
The amorphous region is subjected to hydrolysis, while the crystalline region remains stable
[65]. Slight differences can be observed at day 20 on the MWD but may not be sufficient to
impact the samples. Although the portion of PLA in PLA-g-TPS overall MWD follows the
same path, no lag phase was observed in Figure 3.1b, supporting a more rapid disintegration
of the samples.
For 37 ± 2 °C (Figure 3.2c & d), the PLA MWD shift occurred very slowly, as shown
in Figure 3.2c, especially in the case of PLA. The peak amplitude remained the same, even
at day 180, and a minimal shift of Mn is observed, indicating that chemical hydrolysis
happened very slowly. Whereas for PLA-g-TPS films (Figure 3.2d), shifting and significant
peak reduction are observed after day 90, indicating the reduction of the Mn and initial
change in crystallinity (Figure A3.6 and Table A3.2, Appendix 3F). As biodegradation of
the TPS component of the PLA-g-TPS film proceeded, so did hydrolysis of the PLA matrix,
shown by the broadening of the peaks after day 60 and the peak reduction. This was
accompanied by a stable increase in % mineralization, as shown in Figure 3.1d.
220
Figure 3.2 Molecular weight distribution of PLA and PLA-g-TPS films in compost at 58 °C
(a & b) and 37 °C (c & d), respectively.
Figure 3.3 shows the reduction in Mn of PLA and PLA-g-TPS films in compost at 58
± 2 °C and 37 ± 2 °C. In the thermophilic test, the samples could be retrieved only until day
20. For the mesophilic test, the samples were collected until day 180. Figure 3.3 shows a
more rapid reduction of the Mn at 58 ± 2 °C than at 37 ± 2 °C due to the higher hydrolysis
rate at elevated temperatures. Furthermore, a larger deviation of the Mn of PLA-g-TPS
seemed to occur at 180 days.
221
Figure 3.3 Normalized Mn reduction as a function of time for PLA and PLA-g-TPS films in
compost at 58 °C and 37 °C. The experimental data were fitted using a first-order reaction of
the form Mn /Mno = e(−kt), where Mno is the initial Mn, k is the rate constant, and t is the time.
The data points indicate the original experimental triplicate values at specific times.
Hydrolysis experiments for PLA films were concurrently performed at 58 ± 2 °C and
37 ± 2 °C to decouple, simulate, and understand the abiotic phase (lag phase of biodegradation
curve) dominated by chemical hydrolysis, as also reported elsewhere [44]. Hydrolysis in solid
state (compost) and water are not exactly the same. We have previously conducted these
experiments in sterilized vermiculite to show the difference; however, in this case, hydrolysis
in water should be a sufficient proxy to model the abiotic stage. Additionally, the hydrolysis
data obtained at 58 ± 2 °C was also used to predict the lifetime of PLA at 37 ± 2 °C using the
master curve technique reported by Limsukon et al., who studied the hydrolytic degradation
of PLA films over a series of temperatures (40 to 95 °C) and constructed a master curve to
222
predict the lifetime of PLA films at any temperature [57]. More details about this
methodology can be found in section S5 of the Supporting Information. For this work, the fit
obtained from the accelerated degradation testing at 58 ± 2 °C was used to predict the
hydrolytic degradation at 37 ± 2 °C, as seen in Figure 3.4. To achieve the same reduction in
Mn, c. 10 kDa or similar disintegration values for PLA, approximately 730 days were required
at 37 ± 2 °C as compared to 25 days at 58 ± 2 °C.
Figure 3.4 Hydrolysis of PLA at 58 ± 2 °C (a) and master curve prediction for hydrolytic
degradation at 37 ± 2 °C (b). a) represents the actual experimental data obtained at 58 ± 2
°C. The data points indicate the original experimental triplicate values at specific times.
As shown in Figure 3.3 and Figure 3.4, PLA degradation follows a first-order
reaction, where chemical hydrolysis is the dominant phase at high and lower temperatures
and controls the degradation rate. So, the degradation rate of PLA and PLA in PLA-g-TPS in
compost at 37 ± 2 °C was modeled to a first-order reaction. Table 3.2 details the rate constant
k (d-1) for Mn reduction of PLA and PLA-g-TPS films for both conditions. No significant
difference in Mn reduction among PLA and PLA-g-TPS films was observed in thermophilic
223
conditions at 58 ± 2 °C. At the higher temperature, PLA is in the rubbery state, enabling easy
diffusion of water and, thereby, more rapid hydrolysis. The presence of starch definitely aids
in the initial disintegration and biodegradation of PLA [23], but at the higher temperature,
the process of chemical hydrolysis is dominant and governs the reduction in Mn to
differentiate any advantage associated with the presence of starch on the abiotic degradation
of PLA.
Table 3.2 Rate constant (k) for PLA and PLA-g-TPS films evaluated in compost media at 58
± 2 °C and 37 ± 2 °C.
Film
k (d−1) at 37 ± 2 °C
k (d−1) at 58 ± 2 °C
PLA
0.0885 ± 0.0010a
0.0045 ± 0.0001a
PLA-g-TPS
0.0900 ± 0.0167a
0.0056 ± 0.0002b
Values with different letters in a column are statistically different (α = 0.05 Tukey-Kramer Test).
The chemical hydrolysis process proceeds at a lower rate at 37 ± 2 °C than at 58 ± 2
°C. Conversely, there is a significant effect on the reduction in Mn of the PLA-g-TPS films
due to the presence of starch in the PLA matrix at 37 ± 2 °C, suggesting that the addition of
starch leads to enhanced degradation at mesophilic conditions. The presence of starch
increases the hydrophilicity of the PLA-g-TPS films, further promoting water absorption and
microbial colonization and leading to changes in the structural integrity. This results in the
accelerated degradation of PLA in PLA-g-TPS films as compared to neat PLA with a different
(k) rate constant at 37 ± 2 °C.
Lv et al. conducted a biodegradation study in outdoor soil conditions to evaluate and
account for the role starch plays in PLA degradation, and found higher degradation of PLA
in PLA/starch composites as compared to pure PLA; the authors reported the theoretical and
experimental values of PLA degraded in the presence of starch through weight loss and
component content analysis determination [31,66]. Yu et al. also demonstrated higher
biodegradability, using the mass loss technique, for PLA/starch blends as compared to PLA
224
in soil conditions of 44 °C and high humidity due to the presence of acetylated starch [67].
Palai et al. recorded Mw reduction, depletion in mechanical properties, along with scanning
electron microscopy morphological analysis, and reported higher degradation for PLA/starch
blown films as compared to PLA and PLA/PBSA (poly butylene succinate-co-adipate) blend
in a soil environment at 30 ± 2 °C due to the presence of TPS [68].
In this work, we demonstrated that reactive blending of starch with PLA presents an
array of options for PLA to be easily accessed by water and microorganisms, thereby creating
avenues for accelerated disintegration and rapid mineralization of PLA-g-TPS films at
thermophilic conditions, which can increase the acceptance of PLA by industrial composting
facilities. The blending of TPS and PLA also improves PLA biodegradation in mesophilic
environments, thereby opening an opportunity for creation of PLA-based films that can be
home composted. Overall, these findings can also help tackle the ever-growing problem of
microplastics left behind after disintegration and incomplete biodegradation of plastics.
3.5
Conclusion
The biodegradation performance of a polyester-based reactive blend was evaluated in
simulated composting conditions in both mesophilic and thermophilic conditions. PLA-g-TPS
films showed the highest mineralization trend at 58 ± 2 °C by day 180, indicating that the
presence of starch significantly affects the final biodegradation of PLA by reducing the initial
lag phase and accelerating Mn reduction. Under mesophilic conditions (37 ± 2 °C), PLA-g-TPS
showed improved degradation compared to PLA due to plasticized starch, achieving more
than 57% mineralization. The fact that the amount of starch was around 30% of the blend
indicates that PLA is being degraded and TPS influences Mn reduction of PLA as well. The
presence of starch imparts hydrophilicity and surface roughness to PLA-g-TPS composites
and creates favorable conditions for enhanced microbial activity. Furthermore, starch
225
accelerates the biodegradation of pure PLA by acting as a food reservoir for the growth of
microorganisms while PLA undergoes chemical and enzymatic hydrolysis. Starch can be used
to improve PLA degradation in industrial composting conditions by reducing the lag phase of
disintegration of PLA and, even at mild conditions as in home composting settings, by
inducing biodegradation during the initial days.
3.6
Acknowledgments
P.M. thanks the undergraduate and graduate students at the School of Packaging,
Michigan State University, for their help in running the biodegradation tests reported in this
manuscript. P.M. also acknowledges the School of Packaging at Michigan State University
for partial support; R.A. acknowledges the USDA National Institute of Food and Agriculture
and Michigan State University AgBioResearch, Hatch project number MICL02665, for
partial support of the study.
226
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233
APPENDIX 3A: MATERIAL PROCESSING
Maleic anhydride (MA) and dicumyl peroxide (DCP) along with PLA was used to
produce PLA-g-MA masterbatch. Similarly, glycerol and starch were processed together to
produce a thermoplastic starch (TPS) masterbatch. PLA pellets were processed to produce a
PLA masterbatch. The different zones in the co-rotating twin screw extruder namely the
feeding-melting zone, large kneading/mixing zone, conveying zone, short kneading zone, and
conveying zone provide the necessary shear in the form of mechanical and thermal energy to
produce TPS [69]. Figures A3.1 and A3.2 show the processing of PLA, PLA-g-MA, and TPS
masterbatches and PLA-g-TPS films respectively.
Figure A3.1 Processing of PLA-g-MA, TPS, and PLA masterbatches in a twin screw extruder
and temperature profile from feeder to the die of the twin screw extruder.
234
The masterbatches produced from the twin screw extruder were blended in specific
proportion (56% PLA, 14% PLA-g-MA, and 30% TPS) in a cast film extruder to produce PLA-
g-TPS films.
Figure A3.2 Processing of PLA-g-MA, TPS and PLA masterbatches into PLA-g-TPS film by
cast film extrusion.
235
APPENDIX 3B: PHYSICOCHEMICAL CHARACTERISTICS
Some compost was collected and sent to the Soil and Plant Nutrient Laboratory at
Michigan State University (East Lansing, MI, USA) to evaluate its physicochemical
parameters (dry solids, volatile solids, and C/N ratio) as previously described elsewhere [12].
The physicochemical parameters are reported below in Table A3.1.
Table A3.1 Physicochemical parameters and total nutrient analysis of compost used in the
biodegradation test.
Parameter
Dry solids, %
Volatile solids, %
pH
C/N ratio
Carbon, %
Nitrogen, %
Phosphorus, %
Potassium, %
Calcium, %
Magnesium, %
Sodium, %
Sulfur, %
Iron, ppm
Zinc, ppm
Manganese, ppm
Copper, ppm
Boron, ppm
Aluminum, ppm
Compost
42.5
41.7
8.0
10.1
24.2
2.42
1.21
3.15
5.07
2.82
0.58
0.58
9878
480
413
107
41
6751
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APPENDIX 3C: MORPHOLOGICAL CHARACTERIZATION OF PLA-g-TPS
REACTIVE BLEND
The PLA-TPS physical blend and PLA-g-TPS reactive blend were characterized by
SEM to observe the morphological changes [70]. Figure A3.3 a shows the non-uniform
dispersion of the TPS phase in the PLA matrix for PLA-TPS physical blend while Figure
A3.3 b shows good compatibilization and distribution of TPS domain in the PLA matrix.
Figure A3.3 SEM images for (a) PLA-TPS physical blend and (b) PLA-g-TPS reactive blend.
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APPENDIX 3D: ROUGHNESS AND CONTACT ANGLE MEASUREMENTS
The roughness of the PLA film and PLA-g-TPS films in Figure A3.4 a and b
respectively was conducted using a CypherTM atomic force microscope (Oxford Instruments
Asylum Research, Inc., Santa Barbara, CA, USA) in the contact mode. Roughness
parameters, calculated as the root mean square (Rq) and average roughness (Ra), were
determined for each type of film and were calculated from the Htr mode image. Images were
obtained in the Dfr mode [71]. The film area for the determination of roughness was 900 μm2.
Figure A3.4 Surface roughness of (a) PLA and (b) PLA-g-TPS films as measured by atomic
force microscopy. Roughness parameters, calculated as the root mean square (Rq) and
average roughness (Ra), were determined for each type of film and were calculated from the
Htr mode image. Within columns, values followed by a different letter are significantly
different at p ≤ 0.05 (Tukey’s test).
The contact angle of the films was measured by the sessile drop technique using an
in-house built goniometer equipped with a diffuse light source and a digital camera. Film
samples of 7 cm length and 2 cm wide were attached to microscope slides. Next, a drop of
HPLC-grade water (3 μL) was deposited on each film surface, and the contact angle was
determined by using the tangent method as seen in Figure A3.5 a and b [72].
238
abFilms Rq, nm Ra, nm PLA 9.0 ± 2.1a 6.3 ± 1.3a PLA-g-TPS 183.5 ± 0.3b 143.6 ± 5.4b
Figure A3.5 Water contact angle measurement on (a) PLA and (b) PLA-g-TPS films.
239
ab75.3º ±2.568.1º ±1.6
APPENDIX 3E: APPLICATION OF THE TIME-TEMPERATURE SUPERPOSITION
PRINCIPLE FOR PREDICTING HYDROLYTIC DEGRADATION IN MESOPHILIC
CONDITIONS
In a previous work, Limsukon et al. [73] studied the hydrolytic degradation of PLA
film at temperatures from 40 to 95 °C and presented the lifetime prediction methods at the
lower temperature of interest using the time-temperature superposition principle. The
Williams-Landel-Ferry (WLF) equation was used to fit the hydrolysis of PLA film over the
wide range of experiment temperatures crossing the Tg. The WLF equation is given as
follows:
logαT= −
C1(T − Ti)
C2+(T − Ti)
(A3.1)
C1 and C2 can be expressed as:
C1 =
E/R
2.303 (Ti − Ts)
and C2 = (Ti − Ts)
(A3.2)
where T and Ti are experimental temperature and the temperature of interest. Ts is
the temperature at which the conformation entropy induced by segmental motion approaches
zero (is about 50 K below the Tg of PLA immersed in water which is 51.6 °C or 324.75K [74]).
E is the pseudo activation energy over a temperature range from below to above Tg. R is the
gas constant (8.314 J/mol∙K). αT is the factor in shifting the data obtained at temperature T
to overlap one at Ti.
To predict the hydrolysis of PLA in the mesophilic condition, αT value was calculated
by substituting C1 and C2 into S1 to shift the experimental data curves of 58 °C and construct
the master curve at 37 °C using E of the hydrolysis of neat PLA film measured within the
temperature range crossing the Tg reported as 2720 J/mol ; αT can be determined as 0.03205.
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Finally, the master curve of the hydrolysis at 37 °C can be constructed by multiplying the
hydrolysis time range by 1/αT and plotting as a function of the prediction line of hydrolysis at
58 °C. A detailed description of the calculation can be found elsewhere [73,75].
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APPENDIX 3F: CRYSTALLINITY MEASUREMENTS
The films retrieved from the sampling bioreactors were analyzed for the evolution of
crystallinity throughout the test for PLA and PLA-g-TPS films at both 37 °C and 58 °C,
respectively. The crystallinity (Xc) of the samples was tracked using a DSC Q100 (TA
Instruments, New Castle, DE, USA). Samples weighing between 5 and 10 mg were sealed in
aluminum pans and subjected to a cycle ranging from −5 °C to 210 °C. The samples were
cooled to −5 °C and then heated to 210 °C at a ramp rate of 10 °C/min under a nitrogen
atmosphere where the purge flow was maintained at 70 mL/min. The data obtained was
analyzed using the software Thermal Universal Analysis 2000, V4.5 (TA Instruments). The
% Xc was calculated using the equation S3:
Xc(%) =
(∆Hm − ∆Hc)
∆Hm
° (1 −
%wtfiller
100
)
x 100
(A3.3)
where ∆Hm is the heat of fusion, ∆Hc is the enthalpy of cold crystallization, ∆Hm
° is the
enthalpy fusion for 100% pure crystalline PLA (93 J/g), and wtfiller is the weight fraction of
TPS (30%).
Table A3.2 Crystallinity evolution for PLA and PLA-g-TPS films at 58°C and 37°C.
Material
58 ± 2 °C
37 ± 2 °C
Day 0
Day 15
Day 0
Day 90
PLA
3.1 ± 0.6
35.5 ± 0.2
28.2 ± 3.5
32.3 ± 1.8
PLA-g-TPS
7.0 ± 0.6
39.8*
7.0 ± 0.6
18.5 ± 2.2
* Only one measurement is noted for the sample due to the fragile state of the film.
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Figure A3.6 DSC thermograms showing the evolution of crystallinity of a) PLA and b) PLA-
g-TPS films in compost at 37 °C.
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CHAPTER 4: ACCELERATING BIODEGRADATION: ENHANCING POLY(LACTIC
ACID) BREAKDOWN AT MESOPHILIC ENVIRONMENTAL CONDITIONS WITH
BIOSTIMULANTS
4.1
Abstract
Poly(lactic acid) – PLA – has garnered interest due to its low environmental footprint
and ability to replace conventional polymers and be disposed of in industrial composting
environments. Although PLA is compostable when subjected to a suitable set of conditions
(i.e., aerobic thermophilic conditions for an extended period), its broader acceptance in
industrial composting facilities has been affected adversely due to longer degradation
timeframes than the readily biodegradable organic waste fraction. PLA must be fully exposed
to thermophilic conditions for prolonged periods to biodegrade, which has restricted its
adoption and hindered its acceptance in industrial composting facilities, also negating its
home composting potential. Thus, enhancing PLA biodegradation is crucial to expand its
acceptance. PLA's biodegradability was investigated in a compost matrix under mesophilic
conditions at 37°C for 180 days by biostimulating the compost environment with skim milk,
gelatin, and ethyl lactate to enhance the different stages of PLA biodegradation. The evolved
CO2, number average molecular weight, and crystallinity evolution were tracked. To achieve
Mn ≲10 kDa for PLA, biodegradation rate was accelerated by 15% by adding skim milk, 25%
by adding gelatin, and 22% by adding ethyl lactate. This work shows potential techniques to
help biodegrade PLA in compost by adding biostimulants.
4.2
Introduction
Poly(lactic acid) (PLA) is a well-known biodegradable thermoplastic polymer and has
garnered a lot of interest due to its ability to exhibit reasonable properties as compared to
fossil-based conventional polymers [1]. In addition, PLA has turned out to be a key player
244
leading the global bioplastic segment as per the market research conducted by nova-Institute
which estimates an increase in bioplastics production from 2.18 million tonnes in 2023 to 7.43
million tonnes in 2028 [2]. Due to its biodegradable nature instead of contributing to plastic
waste in landfills, PLA can be rerouted to composting facilities [3]. PLA is known to undergo
chemical hydrolytic degradation during composting, significantly reducing its molecular
weight [4]. This reduction in molecular weight precedes the assimilation by microorganisms
and final mineralization to CO2. However, it is difficult to degrade PLA at lower temperatures
(e.g., backyard or home composting environments – mostly mesophilic conditions) because of
its dependence on high temperatures (≳60°C) to undergo chemical hydrolysis. Additionally,
PLA degradation in natural environments (usually associated with lower temperatures) is
slow due to the sparse distribution of PLA-degrading microbes compared with other more
aggressive thermophilic degradable environments such as industrial composting [5,6]. A
detailed discussion about the compostability and biodegradation of PLA can be found
elsewhere [7,8].
Biostimulation and bioaugmentation
techniques are
included under
the
bioremediation domain and are used to eradicate hazardous pollutants/waste materials that
may be toxic to the environment. These techniques use enhanced settings or biological
workers/microorganisms to eliminate pollutants. They also are designed to make up for the
lack of factors that can speed up the removal process of pollutants in an eco-friendly
approach. Biostimulation is an approach that addresses limiting factors such as nutrients (in
the form of enzyme inducers),[9,10] electron donor or acceptor compounds,[11,12] nitrogen-
supplying compounds,[13,14] and compounds that can activate the biochemical processes by
providing necessary resources or chemicals to stimulate the environment [15,16]. The
biostimulation technique has been used extensively to revitalize the native microbial
245
communities to boost the degradation of heavy and toxic metals,[11,17,18] diesel oil,[19,20]
chlorinated hydrocarbons,[21] and petroleum hydrocarbons,[22,23] since these compounds
are hard to biodegrade by native microorganism populations.
PLA biodegradation proceeds much faster at temperatures higher than its glass
transition temperature (Tg) ~ 58°C, commonly reached in industrial composting conditions.
Chemical hydrolysis, the dominant mechanism responsible for the significant reduction in
molecular weight of PLA, takes place quickly at higher temperatures [24]. The resulting low
molecular weight PLA is then available for further assimilation by the microorganisms.
Under thermophilic conditions (i.e., 45–60°C), the chemical hydrolysis of PLA
proceeds quickly, efficiently, and in a suitable time frame compared to mesophilic conditions
(i.e., 20–45°C). Accelerating PLA degradation, particularly hydrolytic degradation in
compost, is exceptionally challenging because of the limitation of the solid complex matrix to
be at high temperatures to activate the biodegradation process. Several research studies have
focused on including metal compounds in PLA to catalyze its degradation [25–29]. But most
of these metal compounds are included in PLA as blends/nanocomposites at lower
concentrations, significantly altering physical properties such as number and weight average
molecular weight, melting temperature, and Tg [30–33]. These final properties do not
accurately represent the post-consumer PLA discarded packages ending up in compost, soil,
or landfill scenarios.
Apart from the chemical hydrolysis, the biodegradation of PLA also entails enzymatic
degradation, the release of extracellular enzymes by the microbes, which simultaneously
break down PLA into low molecular weight compounds. The extracellular depolymerases
released by the microorganisms in the case of PLA degradation belong mainly to the
hydrolase class of enzymes, primarily proteases, and some also belong to lipase and cutinase
[34–36]. The serine proteases produced by microbes disintegrate the ester bonds of PLA [37–
246
39]. Several studies have used the biostimulation technique to expedite the biodegradation
of bio-based polymers. But most of these studies were conducted with PLA as the sole carbon
source, in liquid media with specific microbial strains, and usually at higher temperatures
[40–43]. So, there is a gap between these experiments conducted in highly controlled
incubated culture media and the real-world unchecked environment in solid media such as
soil, industrial composting, and home composting. The biological activity in such natural
environments is highly complex due to the diversity of microorganisms competing or working
together to consume organic resources.
This study aimed to evaluate the addition of different biostimulant compounds, which
can be added externally in the compost media (solid) to accelerate the biodegradation of PLA
in mesophilic conditions. The compounds selected as biostimulants were screened through
multiple factors and were expected to meet various criteria: not toxic towards the
microorganisms present, not antibacterial, able to disintegrate, consumable by the
microorganisms, biodegradable in a suitable timeframe, cost-effective, and readily available
or accessible.
Hydrolases (EC 3) are the hydrolytic enzymes responsible for degrading hydrolyzable
polymers such as esters, carbonate, and amide groups. The ester bonds in the backbone of
polyesters act as hydrolyzable linkages and are acted upon by esterases (EC 3.1) and
proteases (EC 3.4), which are released by the PLA-degrading microbes. To induce the activity
of these enzymes by microbes, skim milk and gelatin have been reported as suitable
compounds [44–47]. So, skim milk and gelatin can incite proteolytic activity for enzymatic
degradation, resulting in the cleavage of PLA polymer chains.
To stimulate the lactate-utilizing microbial species in compost, ethyl lactate can be
used. Ethyl lactate belongs to the lactate esters family. It has been used as a food source for
dechlorinators to remediate heavy metals in soil under anaerobic conditions [48–50]. It is
247
considered a green solvent with numerous advantages such as favorable low toxicity and low
cost, and is readily available. It is environmentally benign, highly biodegradable, and has a
strong solvency power [51,52]. Ethyl lactate undergoes hydrolysis to produce ethanol and
lactate, where both can act as electron donor compounds for reductive degradation. Lactate
has been shown to act as an electron donor compound in the case of anaerobic degradation
[53,54]. Hydrolysis of ethyl lactate is a slow process ensuring a constant, long-term supply of
hydrogen as the electron donor for the microbial redox process. The use of ethyl lactate as a
biostimulant for polymer degradation has not yet been reported.
Thus, this study aimed to investigate the effectiveness of biostimulating the compost
environment with compounds that may enhance enzymatic degradation (skim milk and
gelatin), and with an electron donor compound (ethyl lactate) to improve the aerobic
biodegradation of PLA in compost at mesophilic conditions under which it is more difficult to
degrade PLA. The difference in the CO2 evolution and changes in the number average
molecular weight (Mn) and crystallinity (Xc) of PLA degradation with and without
biostimulants was tracked to account for the activity of biostimulants in compost.
4.3 Materials and methods
4.3.1 Materials
Commercial PLA resin, Ingeo™ 2003D, with l-lactic acid content of 96%, with weight
average molecular weight (Mw of 2.23 ± 0.04 x 105 Da) and number average molecular weight
(Mn of 1.14 ± 0.07 x 105 Da) was obtained from NatureWorks LLC (Minnetonka, MN, USA).
Skim milk powder was procured from the local grocery store. Gelatin of the brand McCormick
& Co. (Hunt Valley, MD, USA) was purchased online. Ethyl lactate was procured from Sigma
Aldrich (St. Louis, MO, USA). Cellulose of ∼20 μm particle size was procured from from
Sigma-Aldrich (Milwaukee, WI, USA).
248
4.3.2 Characterization of PLA and the biostimulants
The carbon, hydrogen, and nitrogen contents of PLA resin, skim milk, gelatin, and
ethyl lactate were determined using a CHNS/O Elemental Analyzer, PerkinElmer 2400
Series II (Shelton, CT, USA), and are presented in Table 4.1. The methodology was provided
in our previous work [55].
Table 4.1 Percent of carbon, hydrogen, and nitrogen content by weight of cellulose and film
samples.
Material
% Carbona
% Hydrogena
% Nitrogena
PLA
49.72 ± 0.19
5.72 ± 0.04
0.11 ± 0.07
Skim milk
41.28 ± 0.13
6.33 ± 0.03
5.75 ± 0.06
Gelatin
44.80 ± 0.30
7.00 ± 0.10
16.4 ± 0.10
Ethyl lactate
31.93 ± 0.98
5.46 ± 0.34
0.05 ± 0.03
a: Percentage by weight.
4.3.3 Biodegradation in compost
The biodegradation of PLA and the effect of biostimulant addition on the degradation
of PLA were evaluated under aerobic mesophilic conditions using an in-house direct
measurement respirometric (DMR) system for analysis of evolved CO2 under simulated
composting conditions. The test was adapted to ASTM D53338-15 and ISO 14855-12, as
currently there are no standards in place for evaluating the aerobic biodegradation of plastic
materials under controlled composting conditions at mesophilic temperatures [56–58]. The
system was equipped with a non-dispersive infrared gas analyzer (NDIR) Li-COR® LI-820
(Lincoln, NE, USA), which measures the CO2 concentration. The temperature and relative
humidity (RH) of the chamber was maintained at 37 ± 2°C and 50% ± 5% RH, and the airflow
249
rate was controlled at 40 ± 2 cm3/min. Detailed information about the DMR chamber can be
found elsewhere [24].
Manure compost was obtained from the Michigan State University (MSU) Composting
Facility (East Lansing, MI, USA). The compost was sieved using a 10-mm screen to remove
any large debris or chunks that might be present, and post screening was conditioned at 37
± 2°C. Deionized water was added to adjust the moisture content to 50%. Vermiculite, a
hydrous phyllosilicate inorganic material of premium grade, was obtained from Sun Gro
Horticulture Distribution Inc. (Bellevue, WA, USA). The vermiculite was mixed 1:4 with
compost (dry weight basis). Samples of the compost were analyzed by the Soil and Plant
Laboratory at MSU (East Lansing, MI, USA) for physicochemical parameters such as dry
solids, volatile solids, pH, and C/N ratio, as presented in Table 4.2.
The bioreactors were packed with 400 g of compost, and then 8 g of PLA sample and
8 g of the selected biostimulant were added, and all the samples were tested in triplicate.
Table 4.2 Physicochemical parameters and total nutrient analysis of compost used in the
biodegradation test.
Parameter
Compost
Parameter
Compost
Dry solids, %
42.5
Magnesium, %
2.82
Volatile
solids, %
pH
C/N ratio
Carbon, %
Nitrogen, %
Phosphorus, %
Potassium, %
Calcium, %
41.7
8.0
10.1
24.2
2.42
1.21
3.15
5.07
Sodium, %
Sulfur, %
Iron, ppm
Zinc, ppm
Manganese, ppm
Copper, ppm
Boron, ppm
0.58
0.58
9878
480
413
107
41
Aluminum, ppm
6751
250
4.3.4 Biostimulation
Ethyl lactate, skim milk, or gelatin were added at 8 g to separate bioreactors and
mixed thoroughly with the compost matrix containing the 8 g of PLA pellets to ensure
uniform distribution. The bioreactor used to study the biodegradation of PLA samples is
shown in Figure 4.1.
Figure 4.1 Bioreactor used to study the biodegradation of samples.
Moisture content was adjusted, and the optimal conditions were maintained by
injecting the required amount of deionized water weekly into all bioreactors. CO2-free air
(<30 ppm) was supplied to each bioreactor and the CO2 evolved was measured for a specific
time. The measurement system was purged after every measurement to get rid of any traces
of CO2 from the previous bioreactor and to ensure that the baseline was achieved.
Mineralization, which is defined as the total amount of carbon converted to CO2 molecules,
was calculated according to equation (4.1)
𝑀𝑖𝑛𝑒𝑟𝑎𝑙𝑖𝑧𝑎𝑡𝑖𝑜𝑛 % =
(𝐶𝑂2)𝑡 − (𝐶𝑂2)𝑏
𝑀𝑡 𝑥 𝐶𝑡 𝑥
44
12
𝑥 100
(4.1)
251
The numerator expresses the CO2 evolved from the sample, which is calculated by
subtracting the average cumulative mass of CO2 evolved from the blank (CO2)b from the
average cumulative mass of CO2 evolved from the bioreactor containing the sample (CO2)t.
The denominator represents the theoretical amount of CO2 that can be produced by the
sample and is calculated as a product of the total mass of the sample in the bioreactor (Mt),
and the total carbon content of the sample as derived from CHN analysis (Ct). 44 is the
molecular weight of CO2, and 12 is the atomic weight of carbon.
4.3.5 Hydrolysis experiment
A hydrolysis test method adapted from ASTM D4754-18 [59] was run for PLA films
at 58 ± 2 °C and 37 ± 2 °C to understand the hydrolytic degradation. The hydrolysis cell
consisted of a stainless-steel wire, glass beads, and a glass vial with cap. PLA films were cut
into small discs of 2-cm diameter, and ten such discs were strung into a stainless-steel wire
and separated by glass beads. The vial was filled with 35 mL of HPLC-grade water (J.T.
Baker, Center Valley, PA, USA). The water was preconditioned, and the hydrolysis cell was
stored at the same temperature. Triplicates of the PLA films were retrieved at predetermined
time intervals and dried before running size exclusion chromatography to assess the Mn
reduction.
4.3.6 Size exclusion chromatography
A size exclusion chromatography (SEC) instrument was used to quantify the weight
average (Mw), number average (Mn) molecular weight and molecular weight distribution
(MWD) of the PLA treated with and without biostimulants. SEC system from Waters Corp.
(Milford, MA, USA) was furnished with an autosampler (Waters® 717), a refractive index
detector (Waters® 2414), an isocratic pump (Waters® 1515), and a series of Styragel®
columns (HR4, HR3, HR2). The flow rate and temperature was maintained at 1mL/min and
35°C respectively. The Mark-Houwink constants of K = 0.000174 dL/g and α = 0.736 were
252
used for PLA dissolved in tetrahydrofuran solvent. Waters BreezeTM2 software was used to
examine the data obtained.
4.3.7 Differential scanning calorimetry
A differential scanning calorimeter (DSC), model Q100 (TA Instruments, New Castle,
DE, USA), was used to determine the glass transition temperature (Tg), melting temperature
(Tm), crystallization temperature (Tc), and crystallinity (Xc) for the PLA samples retrieved
from biostimulated compost. Samples weighing between 5–10 mg were sealed in aluminum
pans and subjected to a heating cycle to understand the evolution of crystallinity. The
samples were cooled down to −5°C, and then heated to 210°C at a ramp rate of 10°C/min. The
cooling was achieved using a nitrogen cooling system that maintained the purge flow rate at
70 mL/min. Thermograms were analyzed using the Thermal Universal Analysis 2000
software, V4.5 (TA Instruments). The degree of crystallinity was calculated using equation
(4.2)
𝜒𝑐 % =
𝛥𝐻𝑚 − 𝛥𝐻𝑐
0
𝛥𝐻𝑚
𝑥 100
(4.2)
where 𝛥𝐻𝑚 is the heat of fusion, 𝛥𝐻𝑐 is cold crystallization enthalpy, and 𝛥𝐻𝑚
0 is the
heat of fusion for 100% crystalline pure PLA (93 J/g).
4.3.8 Statistical analysis
All the statistical analyses was conducted using MINITABTM 18 software (Minitab
Inc., State College Park, PA, USA). One-way ANOVA and Tukey’s test were used to evaluate
statistical significance at p < 0.05. All the values are reported as mean ± standard deviation.
4.4 Results and discussion
The biodegradation of PLA in biostimulated compost with skim milk, gelatin, or ethyl
lactate under mesophilic conditions (37°C) was evaluated for 180 days to understand the
effect of these compounds on stimulating home composting operations and understanding the
253
potential effects in industrial composting. To decouple the effects of these compounds, the
CO2 evolution, Mn, and Xc of the samples were tracked.
4.4.1 Effect of skim milk on cellulose and PLA degradation
Figure 4.2a and b present the CO2 evolution and % mineralization of the blank
(compost only), cellulose, skim milk, PLA, cellulose+skim milk (Cell+skm), and PLA+skim
milk (PLA+skm) in compost at 37°C. Cellulose evolved around 36.8 g of CO2 and reached a
mineralization of 87.7%, whereas skim milk showed around 39.2 g of CO2 evolution and
attained over 100% of its carbon conversion over 180 days. The priming effect is observed in
the case of skim milk, due to over-degradation of the indigenous carbon present in the
compost. Since skim milk and cellulose are readily biodegradable and can be easily utilized
as a carbon source by the microorganisms present in the compost, no lag phase was observed.
Skim milk was added to the compost to induce the protease activity of the microbes present,
since serine protease (3.4.21.112), belonging to the peptidases, is in the class of extracellular
enzymes able to hydrolyze the peptide bonds linking to the amino acids in the protein
structure.
Skim milk comprises lactose, casein, and whey protein, making it a good precursor for
the protease enzymatic activity by the microorganisms in the compost. Since cellulose and
skim milk constitute excellent carbon sources when both are present in the compost, they are
expected to show a much higher CO2 evolution. However, the CO2 evolution of the bioreactor
containing both cellulose and skim milk was around 49.1 g, which is close to the CO2 evolution
of cellulose and skim milk separately and shows 96.6% biodegradation.
To better understand the interaction and isolate the degradation behavior of cellulose
in the presence of skim milk (Cellskm), we estimated the mineralization value wherein the
background signal from the bioreactor containing skim milk is subtracted. As seen in Figure
4.2b, the mineralization % depicting the degradation behavior of cellulose reaches a
254
maximum value of 95.9%. The value is higher than for cellulose alone, as skim milk and
cellulose are easily accessible to microorganisms.
PLA pellets in compost showed around 22.5 g of CO2 evolution, whereas the blank
produced around 26.1 g of CO2, implying that no carbon from PLA was degraded. The
negative mineralization values indicate that the blank bioreactors produced more CO2 than
the bioreactors containing PLA samples. In a blank bioreactor, the microbes are easily able
to assimilate the organic matter in the compost, whereas the presence of PLA in a bioreactor
reduces the working efficiency, which in turns shows diminished CO2 production. PLA offers
an initial physical hydrophobic barrier to water, making it difficult for the microorganisms
to utilize it as a carbon source and establish a macro colony able to break down PLA. We did
not see any mineralization in PLA due to the low temperature of 37°C, which is opposite to
the degradation of PLA in thermophilic temperatures [1,24].
255
Figure 4.2 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
skim milk, cellulose + skim milk (Cell+skm), PLA + skim milk (PLA+skm) in compost at
37°C. (c) represents the normalized Mn reduction as a function of time for PLA in control
compost and compost biostimulated by skim milk. The experimental data was fitted using a
first-order reaction of the form Mn /Mno = e(−kt), where Mno is the initial Mn, k is the rate
constant, and t is the time. The inset shows the k-fitted values; values with different
lowercase letters are statistically different (α = 0.05 Tukey-Kramer Test). (d) and (e) depict DSC
thermograms for PLA and PLA in compost biostimulated by skim milk, respectively. (f) and
(g) show the MWD of PLA in compost and compost biostimulated with skim milk,
respectively.
Chemical hydrolysis is the initial and primary driving mechanism in the
biodegradation of PLA. It dramatically reduces molecular weight and aids in PLA
assimilation by microorganisms [60–62]. The Tg of PLA is much closer to the industrial
256
composting conditions, which reduces the rigidity of its polymer chains, creating more free
volume and rearranging the polymer chains for easy diffusion of water into the matrix.[63,64]
The polymer hydrophilicity is also elevated at higher temperatures, further promoting
microbial attachment besides the chemical hydrolysis [65,66]. Apart from this, the initial Mn
of PLA plays a crucial role in the biodegradation of PLA [67]. The bioreactor containing PLA
and skim milk (PLA+skm) shows CO2 evolution of 42.1 g and maximum mineralization of
60.1% by the end of the test duration. This result indicates improved mineralization
compared to PLA alone, where no mineralization was observed. To isolate the degradation
behavior of PLA in the presence of skim milk, mineralization of PLAskm (subtracting blank +
skim milk) was calculated, as described earlier, and is shown in Figure 4.2b. We observed
positive mineralization, indicating PLA's enzymatic degradation due to the presence of skim
milk. To further corroborate the biostimulation activity of skim milk, samples of PLA were
retrieved from the bioreactor at specific intervals, and their Mn was evaluated. The reduction
in Mn of PLA was tracked until the end of the test, as seen in Figure 4.2c. For the skim milk
treatment, the kinetic reduction rate of biostimulated PLA was higher than for PLA. There
was a significant difference in the degradation rates for PLA alone and PLA treated with
skim milk, as indicated by the k values shown in Figure 4.2c inset (PLA k = 0.0045 ± 0.0001
and PLA+skim milk k = 0.0053 ± 0.0003). This increase in k can be interpreted as a final
reduction of around 75 days when PLA biostimulated with skim milk reaches Mn ≲10kDa at
420 days, where microorganism assimilation of PLA n-mers accelerates biodegradation [24].
On the other hand, PLA needs at least 494 days to reach the same Mn conditions –an effective
15% reduction of time.
The PLA samples were also evaluated to understand crystallinity degradation and
evolution at 37°C. The Xc increased from 28.3% to 31.4% for PLA and from 28.3% to 39.5%
257
for PLA samples biostimulated with skim milk, as seen in Figure4.2d and e, and the values
are tabulated in Table 4.3 respectively. The significant difference between the two indicates
the more rapid degradation of PLA samples occurring in the presence of skim milk since the
lowest Mn can reconfigure and recrystallize even though the test was conducted below but
close to Tg. Figure 4.2f and g present the MWD for PLA in compost and compost
biostimulated with skim milk, respectively. The peak amplitude for PLA in Figure 4.2f
remains approximately the same throughout the test duration. No broadening of the peak
and the negligible shift indicate that the chemical hydrolysis proceeded slowly at a mesophilic
temperature of 37°C. Whereas for PLA biostimulated with skim milk, a significant shift and
broadening of the peak are observed in Figure 4.2g, depicting the reduction in Mn,
rearrangement of the PLA molecular chains and change in Xc. The presence of skim milk
initiates protease activity and concurrent chemical hydrolysis, produces a significant peak
shift.
Table 4.3 Crystallinity evolution for PLA and PLA biostimulated with skim milk.
Material
Day 0
Day 180
PLA
28.2 ± 3.5a
31.4 ± 1.8a
PLA + skim
milk
28.2 ± 3.5a
39.5 ± 1.5b
Values with different letters within columns are statistically different (α = 0.05 Tukey-Kramer test).
4.4.2 Effect of gelatin on cellulose and PLA degradation
Gelatin was used as another candidate to stimulate the protease activity of microbial
strains in compost. Gelatin has been reported as a precursor for protease activity
[10,34,40,43]. Figure 4.3a and b show the CO2 evolved and % mineralization of the blank,
cellulose, gelatin, PLA, cellulose+gelatin (Cell+gel), and PLA+gelatin (PLA+gel) in compost
at 37°C. Gelatin shows around 41.5 g of CO2 evolution and mineralization of 116% in 180
days. To further understand the biostimulation of compost by gelatin, it was combined with
258
cellulose. The CO2 evolution in this case (Cell+gel) was 58.5 g, which is as expected and higher
compared to the individual values for cellulose and gelatin, and mineralization of 130.4%. To
better understand the interaction and account for the degradation behavior of cellulose in the
presence of gelatin (Cellgel), the mineralization value was estimated at 180.5% by subtracting
the background signal from the gelatin bioreactor. This higher value depicts that gelatin does
not affect cellulose degradation, and both are used up as carbon sources by the
microorganisms. Additionally, a significant priming effect is observed in the case of bioreactor
containing cellulose biostimulated with gelatin. To understand the influence of gelatin on
PLA degradation, PLA was introduced in the gelatin-amended compost. The bioreactor
containing both PLA and gelatin (PLA+gel) shows CO2 evolution of around 41.8 g and
maximum mineralization of 56.4% by the end of the test (Figure 4.3a). Improved
mineralization is observed as opposed to no CO2 evolution for PLA alone without any
biostimulation of compost. The effect of gelatin on PLA degradation is calculated and
subtracted by plotting the mineralization of PLAgel (subtracting blank + gelatin), as
mentioned earlier. The positive mineralization indicates that gelatin's protease activity helps
in PLA's enzymatic degradation (Figure 4.3b). This is validated by the significant difference
observed in the kinetic rate of degradation for PLA in compost, with and without any
biostimulation by gelatin, as seen in Figure 4.3c inset (PLA k = 0.0045 ± 0.0001 and
PLA+gelatin k = 0.0060 ± 0.0002). This change in k indicates a final reduction of around 124
days when PLA biostimulated with gelatin reaches an Mn ≲10kDa at 371 days compared to
PLA, which needs at least 494 days to reach the same Mn conditions – an effective 25%
reduction of time. The significant difference in the evolution of Xc from 28.3% to 31.4% for
PLA, and from 28.3% to 41.9% for PLA samples biostimulated with gelatin, as seen in Figure
259
4.3d and e, and as seen in Table 4.4 respectively, further shows the improvement in the
enzymatic degradation of PLA due to the presence of gelatin.
Figure 4.3 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
gelatin, cellulose + gelatin (Cell+gel), PLA + gelatin (PLA+gel) in compost at 37°C. (c)
represents the normalized Mn reduction as a function of time for PLA in control compost and
compost biostimulated by gelatin. The experimental data was fitted using a first-order
reaction of the form Mn /Mno = e(−kt), where Mno is the initial Mn, k is the rate constant, and t
is the time. The inset shows the k-fitted values; values with different lowercase letters are
statistically different (α = 0.05 Tukey-Kramer Test). (d) and (e) depict DSC thermograms for PLA and
PLA in compost biostimulated by gelatin, respectively. (f) and (g) show the MWD of PLA in
compost and compost biostimulated with gelatin, respectively.
260
Table 4.4 Crystallinity evolution for PLA and PLA biostimulated with gelatin.
Material
Day 0
Day 180
PLA
28.2 ± 3.5a
31.4 ± 1.8a
PLA + gelatin 28.2 ± 3.5a
41.9 ± 1.9b
Note: Values with different letters within columns are statistically different (α = 0.05 Tukey-Kramer
test).
The broadening and change in the amplitude of the MWD peaks through the test
duration of 180 days, as seen in Figure 4.3g for PLA in compost stimulated with gelatin
compared to Figure 4.3f for PLA alone, provide compelling evidence and confirm enhanced
degradation for PLA with gelatin.
4.4.3 Effect of ethyl lactate on cellulose and PLA degradation
Ethyl lactate was used to stimulate the lactate-utilizing microbial species in the
compost to improve PLA degradation at 37°C. Figure 4.4a and b show the CO2 evolved and
% mineralization of the blank, cellulose, ethyl lactate, PLA, cellulose+ ethyl lactate (Cell+el),
and PLA+ethyl lactate (PLA+el) in compost at 37°C. Ethyl lactate evolves around 51.5 g of
CO2, the corresponding mineralization reaches around 270%, and no lag phase is observed.
As noted earlier with skim milk and gelatin, a priming effect is observed with ethyl lactate.
The CO2 evolution of the bioreactor containing both cellulose and ethyl lactate (Cell+el) was
only around 66.2 g and there was a corresponding mineralization of 182.4%, as both are
readily available to be used as carbon sources by the microbial population. The higher value
can be attributed to the lactate-utilizing microbial community, increasing the microbial
population. The cellulose degradation in the presence of ethyl lactate (Cellel) was decoupled
by subtracting the CO2 evolution from ethyl lactate – maximum mineralization of 116.7% was
observed.
261
To understand how ethyl lactate affects the degradation of PLA, PLA was introduced
into the bioreactor biostimulated by ethyl lactate. PLA in the presence of ethyl lactate
(PLA+el) evolved around 51.6 g of CO2, followed a similar trend as ethyl lactate over 180 days,
Figure 4.4 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
ethyl lactate, cellulose + ethyl lactate (Cell-gel), PLA + ethyl lactate (PLA-gel) in compost at
37°C. (c) represents the normalized Mn reduction as a function of time for PLA in control
compost and compost biostimulated by ethyl lactate. The experimental data was fitted using
a first-order reaction of the form Mn /Mno = e(−kt), where Mno is the initial Mn, k is the rate
constant, and t is the time. The inset shows the k-fitted values; values with different
lowercase letters are statistically different (α = 0.05 Tukey-Kramer Test). (d) and (e) depict DSC
thermograms for PLA and PLA in compost biostimulated by ethyl lactate, respectively. (f)
and (g) show the MWD of PLA in compost and compost biostimulated with ethyl lactate,
respectively.
262
and showed maximum mineralization of 105%. A positive mineralization behavior is seen for
PLA when the effect of ethyl lactate is accounted for (PLAel), indicating the lactate-
stimulating activity of ethyl lactate, but the biostimulation is lower than with skim milk and
gelatin.
This finding is substantiated by a significant difference in the kinetic rate of
degradation for PLA with and without the biostimulation effect of ethyl lactate, as indicated
in Figure 4.4c inset (PLA k = 0.0045 ± 0.0001 and PLA-ethyl lactate k = 0.0058 ± 0.0002).
This change in k indicates a final reduction of around 111 days when PLA biostimulated with
ethyl lactate reaches an Mn ≲10kDa at 384 days compared to PLA, which needs at least 494
days to reach the same Mn conditions – an effective 22% reduction of time. The PLA samples
were also evaluated to understand crystallinity degradation and evolution at 37°C. The Xc
increased from 28.3% to 31.4% for PLA, and from 28.3% to 38.4% for PLA samples
biostimulated with ethyl lactate, as seen in Figure 4.4d and e, respectively. The significant
difference between the two indicates the degradation of samples as seen from Table 4.5
occurring in the presence of ethyl lactate. Figure 4.4f and g present the MWD for PLA in
compost and compost biostimulated with ethyl lactate, respectively. The peak amplitude for
PLA in Figure 4.4f remains approximately the same throughout the test duration. No
broadening of the peak, accompanied by a negligible shift, indicates that the chemical
hydrolysis proceeded slowly at a mesophilic temperature of 37°C. Whereas for PLA
biostimulated with ethyl lactate, a significant shift and broadening of the peak are observed
in Figure 4.4g, depicting the reduction in Mn as previously shown for skim milk and gelatin.
263
Table 4.5 Crystallinity evolution for PLA and PLA biostimulated with ethyl lactate.
Material
Day 0
Day 180
PLA
28.2 ± 3.5a
31.4 ± 1.8a
PLA + ethyl
lactate
28.2 ± 3.5a
38.4 ± 2.1b
Values with different letters within columns are statistically different (α = 0.05 Tukey-Kramer test).
The addition of biostimulants in the compost media accelerated the enzymatic
degradation of PLA in mesophilic environments, which could be scaled to home composting
settings. The biostimulants tested in the study are easily available and can be sourced locally,
thus encouraging the practice of home composting, especially for contaminated PLA-based
food packages post use. These practices can be applied and carried out by consumers in the
backyard, and by university campuses diverting cafeteria and dining hall food waste from
ending up in landfill. Additionally, the improvement in PLA degradation because of the
presence of skim milk, gelatin, and ethyl lactate can further improve acceptance of PLA in
thermophilic industrial composting environments, as these biostimulants can help improve
the turnover time as compared to organic fraction such as food waste. In future work, we will
be focusing on analyzing the effect of adding biostimulants periodically on accelerating PLA
degradation. Understanding how the periodic addition of biostimulant impacts the
mechanism of PLA’s enzymatic degradation can further open avenues to investigate
incorporation of enzymes directly to effectively biodegrade PLA.
4.5
Conclusion
The study investigated the biodegradation behavior of PLA using the biostimulation
treatment under an aerobic simulated mesophilic setting. Different compounds – skim milk,
gelatin, and ethyl lactate – were used to study the biostimulation effect by targeting different
steps in PLA biodegradation. As expected, without any treatment, a very long abiotic lag
phase was observed for PLA at mesophilic conditions (37°C), indicating and reaffirming a
264
very slow hydrolysis phase. This is depicted as negative mineralization over the test duration
of 180 days. Improved CO2 evolution was observed for PLA when the compost was
biostimulated with the addition of skim milk, gelatin, or ethyl lactate. This finding was
corroborated by tracking crystallinity evolution and molecular weight changes. PLA
biodegradation was accelerated to reach Mn ≲10kDa (where the biotic phase is taken over to
consume the PLA n-mers) by 15% by adding skim milk, 25% by adding gelatin, and 22% by
adding ethyl lactate. This work opens the door to help biodegrade PLA in compost conditions
by adding selected biostimulants.
4.6
Acknowledgements
The authors thank Anibal Bher for helping with the test, the data acquisition, and
discussion about biodegradation. P.M. acknowledges the School of Packaging at Michigan
State University; R.A. acknowledges the USDA National Institute of Food and Agriculture
and Michigan State University AgBioResearch, Hatch project number MICL02665, for
partial study support.
265
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271
CHAPTER 5: SPEEDING IT UP: DUAL EFFECTS OF BIOSTIMULANTS AND
IRON ON BIODEGRADATION OF POLY(LACTIC ACID) AT MESOPHILIC
CONDITIONS
5.1
Abstract
Plastic pollution presents a growing concern, and various solutions have been
proposed to address it. One such solution involves the development of new plastics that match
the properties of traditional polymers while exhibiting enhanced biodegradability when
disposed of in a suitable environment. Poly(lactic acid) (PLA) is a biobased, compostable
polymer known for its low environmental impact and ability to break down into harmless
components within a specified timeframe. However, its degradation in industrial composting
facilities poses challenges, and it cannot degrade in home composting. In this study, we
investigated the biodegradability of PLA within a biostimulated compost matrix at
mesophilic conditions (37°C) over 180 days. The compost environment was enhanced with
Fe3O4 nanopowder, skim milk, gelatin, and ethyl lactate, individually and in combination, to
target different stages of the PLA biodegradation process. We monitored key indicators, CO2
evolution, number average molecular weight, and crystallinity, to assess the impact of the
various biostimulants and iron. The results demonstrated that the most effective treatment
for degrading PLA at mesophilic conditions was adding gelatin and Fe3O4. Gelatin
accelerated PLA biodegradation by 25%, Fe3O4 by 17%, and a combination of gelatin and
Fe3O4 by c. 30%. The effect of skim milk and ethyl lactate is also reported. This research
introduces novel pathways to enhance PLA biodegradation in home composting scenarios,
offering promising solutions to address the plastic pollution challenge.
272
5.2
Introduction
Poly(lactic acid) – PLA – is a biobased, biodegradable polymer that is an eco-friendly
substitute for fossil-based polymers for a circular and sustainable economy. PLA is derived
from natural resources and quickly degrades and breaks down in suitable waste management
environments, such as industrial composting. PLA is a versatile biobased polymer because it
has properties comparable to conventional polymers, is cost-effective, and can provide an
additional disposal scenario, namely composting, at the end of a contaminated package life
cycle [1]. These benefits, combined with the growing consumer awareness of plastic littering
and white pollution, have propelled PLA to the forefront as the face of the green, biobased
plastic movement [2].
Although PLA is industrially compostable, its practical and rapid biodegradation
depends on reaching temperatures in the thermophilic range, 45 to 60°C, to undergo
chemical hydrolysis and significantly reduce its molecular weight in a shorter period so that
microorganisms present in the compost can use PLA oligomers as a food source. This
constraint makes it difficult to degrade PLA at lower and ambient temperatures (i.e.,
mesophilic range, 20 to 45°C) [3]. Chemical hydrolysis is a crucial step in the PLA
degradation mechanism, involving the breakdown of high into low molecular-weight polymer
chains, such as oligomers, dimers, and monomers, which are easily assimilated by
microorganisms [4]. However, at the lower temperatures commonly encountered in the home
or backyard composting environments (herein referred to as backyard composting), chemical
hydrolysis proceeds at a prolonged pace [5]. Boosting the hydrolytic degradation of PLA,
particularly in backyard composting, is extremely difficult due to the limitation of high
temperature essential to activate the biodegradation process [3].
Several research studies have included nanocomposites and metal compounds within
the PLA matrix at lower concentrations to enhance its depolymerization and degradation [6–
273
8]. Including these metal oxides and nanoparticles significantly alters physical properties,
such as number average molecular weight (Mn), melting temperature (Tm), and glass
transition temperature (Tg) [9–11] of the resulting PLA or PLA blends, and may not be
practical for food contact or single-use PLA discarded packages.
Enzymatic hydrolysis involving the release of extracellular enzymes is also essential
for PLA degradation when low molecular weight PLA chains are available. PLA’s degrading
enzymes include the hydrolase class of enzymes, primarily proteases [3]. The serine proteases
released by the microorganisms in response to the presence of amino acid compounds in
compost have also been shown to cleave the ester bonds in PLA [12,13]. So, the introduction
of enzymes to compost may assist in the biodegradation of PLA.
Adding various components besides enzymes, such as nutrients, electron
donor/acceptor compounds, or compounds essential to trigger the biochemical processes of
microorganisms in the given environment, is termed biostimulation [14–16]. Biostimulation
techniques to enhance PLA biodegradation have been reported [17]. However, most of these
studies were conducted in restricted settings where PLA was the only carbon source, in liquid media
with specific microbial strains, and usually at higher temperatures, which do not replicate the conditions
encountered during backyard composting [18,19].
In a previous study, we assessed the change in PLA biodegradation at a mesophilic
temperature of 37°C in a solid composting matrix by adding biostimulants to the compost.
The compounds selected as biostimulants in that study were screened through multiple
criteria: anticipated to have no toxicity towards the microorganisms present, be able to
degrade, be biodegradable in a suitable timeframe, be consumed, be cost-effective, and be
readily available. The main goal was to introduce compounds enhancing the biotic
degradation stage. Skim milk and gelatin were selected to trigger proteolytic activity. Ethyl
lactate, belonging to the lactate esters family, was used to stimulate the lactate-utilizing
274
microbial species in backyard compost. All these compounds effectively enhanced the
biodegradation of PLA in simulated backyard composting by at least 15%, as determined by
the accelerated reduction of Mn [20].
In this study, we focused on evaluating the effect of adding a metal compound to
catalyze the chemical hydrolysis of PLA and combine that with the previous demonstrated
biotic enhancement. Metals can be added externally to the compost media rather than within
PLA for humification purposes, as previously reported [21,22] and screening through the
earlier criteria. Table 5.1 presents the permissible limits of metal compounds as derived
from regulatory standards for heavy metals in agricultural soils (mg/kg) [23], since the
resulting amended compost may be applied to agricultural land, lawns, or home gardens. The
primary standards include EEA 2007 [24], TMS 2007 [25], BPI 2021 [26], GB 15168-2018
[27], OMOE 2011 [28], and NZME 2012 [29].
Table 5.1 Critical limits of heavy metals in agricultural soils (mg/kg), adapted from [23].
Country
As
Cd
20
20
3
3
Cr
50
Cu
Hg
100
1
Ni
60
Pb
Zn
300
200
250
150
0.8
100
200
500
20 -
40
0.3 -
0.6
150-
300
50-
200
0.5-
3.4
60-
190
70-
240
200-
300
Australia
Canada
China
Germany
50
Tanzania
1
5
1
500
200
100
200
5
2
200
1000
600
100
200
150
Netherlands
76
13
180
190
36
100
530
720
New
Zealand
17
3
290
> 104
200
N/A
160
N/A
UK
43
1.8
N/A
N/A
26
230
N/A
N/A
US
6.5
1.5
105
50
0.4
31
75
250
275
Considering the criteria mentioned before and the information from Table 5.1 only a
few metals could be selected for further consideration due to limitations placed by permissible
limits. The selected compounds were further scrutinized (Table 5.2) for their antibacterial
properties, as reported in the literature, to evaluate their use directly in compost.
Table 5.2 List of elements screened for their antimicrobial properties.
Element
Antibacterial nature
References
Zn
Fe
Cu
Ni
Ti
Cl
Ag
Yes
No
Yes
Yes
Yes
Yes
Yes
[30,31]
[30,32,33]
[30]
[30]
[34,35]
[30,36]
[36,37]
Table 5.2 indicates that iron was an acceptable compound that could be used to target
chemical hydrolysis. It was demonstrated that introduction of a Lewis acid, FeCl3, can speed
up the hydrolysis of PLA in an alkali solution [38]. However, due to chlorine's antimicrobial
behavior, it was impossible to introduce FeCl3 into compost. Alternatively, iron oxides, such
as FeO, FeO2, Fe3O4, or Fe2O3, could be the primary option and are in a form that is not toxic
to the microbes in the compost [32,33]. Several of these iron forms are present in soil
worldwide [39].
Thus, this study aimed to investigate the effectiveness of biostimulating the compost
environment with compounds that may be able to enhance the chemical hydrolysis (Fe3O4),
the enzymatic degradation (skim milk and gelatin), and electroconductivity (ethyl lactate as
electron donor compound) during the aerobic biodegradation of PLA in compost at mesophilic
conditions. Differences in CO2 evolution, changes in Mn, and the crystallinity (Xc) of PLA
276
degradation with and without biostimulants were monitored to account for the activity of
biostimulants in compost.
5.3 Materials and methods
5.3.1 Materials
PLA Ingeo™ 2003D resin, with L-lactic acid content of 96%, was obtained from
NatureWorks LLC (Minnetonka, MN, USA). Iron oxide nano-powder (Fe3O4) was obtained
from US Research Nanomaterials, Inc. (Houston, TX, USA). Skim milk powder was procured
from a local store (Walmart, Lansing, MI, USA). Gelatin of the brand McCormick & Co. (Hunt
Valley, MD, USA) was purchased on Amazon LLC. Ethyl lactate was procured from Sigma
Aldrich™ (St. Louis, MO, USA).
5.3.2 Characterization of PLA and the biostimulants
The carbon, hydrogen, and nitrogen compositions of PLA resin, skim milk, gelatin,
and ethyl lactate were determined using elemental analysis, (CHNS/O Elemental Analyzer,
PerkinElmer 2400 Series II) (Shelton, CT, USA), and are presented in Table 5.3.
Table 5.3 Carbon, hydrogen, and nitrogen content (percentage by weight) of the tested
materials.
Material
% Carbona
% Hydrogena
% Nitrogena
PLA
49.72 ± 0.19
5.72 ± 0.04
0.11 ± 0.07
Skim milk
41.28 ± 0.13
6.33 ± 0.03
5.75 ± 0.06
Gelatin
44.80 ± 0.30
7.00 ± 0.10
16.4 ± 0.10
Ethyl lactate
31.93 ± 0.98
5.46 ± 0.34
0.05 ± 0.03
[a] Percentage by weight.
277
5.3.3 Biodegradation in compost
The biodegradation of PLA and the effectiveness of introducing biostimulants in
compost on the degradation of PLA were evaluated under aerobic mesophilic conditions using
a direct measurement respirometric (DMR) system [40–42]. The system included a non-
dispersive infrared gas analyzer (NDIR) (Li-COR® LI-820, Lincoln, NE, USA) which
measures the CO2 concentration. The DMR system chamber was maintained at temperature
of 37 ± 2 °C and relative humidity (RH) of 50% ± 5%. A flow rate of CO2-free air (concentration
<30 ppm to establish a low baseline) was controlled at 40 ± 2 cm3/min. Detailed information
about the DMR equipment can be found in other source [43].
The mature compost obtained from MSU composting facility was sifted using a 10-
mm screen to get rid of any huge debris or chunks present, and then conditioned at 37°C until
use. Deionized water was used to adjust the moisture content of compost to 50%. Saturated
vermiculite (Sun Gro Horticulture Distribution Inc., Bellevue, WA, USA), was mixed with
compost in 1:4 parts (dry weight). Samples of the resulting compost-vermiculite mixture were
later sent to the Soil and Plant Laboratory at MSU for determining the physicochemical
parameters. Data regarding the nutrient analysis is presented in Table A5.1 (Appendix 5A)
[44]. The bioreactors were packed with 400 g of compost, 8 g of PLA sample and the selected
biostimulant, and all the samples were tested in triplicate. Blank (only compost) and positive
control (cellulose) were also tested.
Biostimulation: Fe3O4 nanopowder at 17 g was mixed with the compost matrix in the
bioreactor containing PLA to target the hydrolysis step. Ethyl lactate, skim milk, and gelatin
were added at 8 g individually per bioreactor and mixed with the compost matrix thoroughly
with the PLA pellets in it to ensure uniform distribution.
278
5.3.4 Size exclusion chromatography
The Mn and molecular weight distribution (MWD) of PLA for the control and each
treatment with the biostimulants were measured using SEC (Waters Corp., Milford, MA,
USA) as described elsewhere [4]. PLA samples weighing approximately 10 mg were retrieved
at predetermined time intervals and dissolved in 5 mL of tetrahydrofuran (THF) solvent. A
temperature of 35°C and a 1 mL/min flow rate were maintained during testing. The Mark-
Houwink constants of K = 0.000174 dL/g and α = 0.736 were used to determine Mn ,Mw and
MWD of the PLA samples. Data analysis was carried out using Waters BreezeTM2 software.
5.3.5 Differential scanning calorimetry
A DSC model Q100 (TA Instruments, New Castle, DE, USA), was used to determine
the Tg, Tm, crystallization temperature (Tc), and crystallinity (Xc) for the PLA samples
retrieved from the regular and biostimulated compost. PLA samples weighing between 5 - 10
mg were packed in aluminum pans and cooled down to −5°C and then subjected to a heating
cycle to reach 210°C at a ramp rate of 10°C/min . This helped to evaluate the evolution of Xc.
The cooling was achieved using a nitrogen cooling system that maintained the purge flow
rate at 70 mL/min. The degree of crystallinity was estimated using equation (5.1):
𝜒𝑐 % =
𝛥𝐻𝑚 − 𝛥𝐻𝑐
0
𝛥𝐻𝑚
𝑥 100
(5.1)
where 𝛥𝐻𝑚 is the heat of fusion, 𝛥𝐻𝑐 is cold crystallization enthalpy, and 𝛥𝐻𝑚
0 is the
heat of fusion for 100% crystalline pure PLA (93 J/g) [45].
5.4 Results and discussion
The CO2 evolution of PLA samples in compost and biostimulated with Fe3O4
nanopowder and the combination of gelatin, skim milk, and ethyl lactate without and with
Fe3O4 nanopowder was tracked over a test duration of 180 days at mesophilic conditions
279
(37°C). Samples were retrieved at specific times to evaluate the Mn and Xc evolution and
determine the kinetic degradation rate.
5.4.1 Effect of Fe3O4 on cellulose and PLA degradation
Figure 5.1a and b shows the CO2 evolution and mineralization, respectively, of
cellulose, cellulose in compost biostimulated with Fe3O4 nanopowder (hereafter referred to as
cellulose + Fe), PLA, and PLA in compost biostimulated with Fe3O4 nanopowder (hereafter
referred to as PLA + Fe) at 37°C. Control compost (blank) evolved around 26.1 g of CO2, and
compost biostimulated with Fe3O4 nanopowder (hereafter referred to as blank + Fe) evolved
around 27.2 g of CO2. The minor difference can be attributed to the difference in weight of
the compost introduced in the bioreactor, and the levels are not significantly different (p >
0.05). Cellulose in compost evolved around 36.8 g of CO2, and reached a mineralization of
87.7%, whereas cellulose + Fe evolved around 44.6 g of CO2, depicting a mineralization of
137.7%. The primary reason for the priming effect (>100% mineralization) observed in the
case of cellulose + Fe may be attributed to the over-deterioration of the endemic carbon
present in the compost [4]. PLA in compost showed around 22.5 g of CO2 evolution, whereas
blank produced around 26.1 g of CO2, implying that no carbon from PLA was degraded. The
negative mineralization values indicate more CO2 production in the blank bioreactors than
in the PLA bioreactors. PLA offers a physical hydrophobic barrier to water, making it difficult
for the microorganisms to utilize it as a carbon source at the beginning of the test and until
day 180 due to the low contribution of chemical hydrolysis at mesophilic temperatures.
Overall, we did not see any mineralization in PLA due to the low temperature of 37°C, which
is insufficient to activate the biotic stage. These values are very low compared to the
degradation of PLA at thermophilic temperatures [4,44] but are similar to earlier reported
values [5,46].
280
In the case of PLA and PLA + Fe, we observed the difference in the CO2 evolution
right from the start of the test. PLA + Fe evolved around 25.9 g of CO2 compared with 22.5 g
of CO2 evolution for PLA. Similar to the case of cellulose + Fe, the reason for the difference
in the CO2 evolution for PLA can be attributed to the presence of Fe3O4. Fe3O4 promotes
microbial activity in soil and enhances the nitrification potential [47]. Fe3O4 is also known to
induce changes by enhancing enzymatic activity and microbial growth [32]. This
characteristic can be corroborated by the kinetic degradation rate (k), as seen in Figure 5.1c
inset (PLA k = 0.0045 ± 0.0001 d-1 and PLA+ Fe k = 0.0052 ± 0.0002 d-1). The significant
difference can be credited to the presence of Fe3O4. Figure 5.1d and e present the molecular
weight distribution (MWD) for PLA in compost and compost biostimulated with Fe3O4. The
peak amplitude for PLA in Figure 5.1dremained approximately the same throughout the test
duration. No broadening of the peak and negligible shift indicates that the chemical
hydrolysis proceeded slowly at a mesophilic temperature of 37°C. In contrast, for PLA + Fe,
a significant shift to low Mw and broadening of the peak are observed in Figure 5.1e, depicting
the reduction in Mn as shown in Figure 5.1c.
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Figure 5.1 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
blank + Fe3O4 (Blank + Fe), cellulose + Fe3O4 (Cellulose + Fe), PLA + Fe3O4 (PLA + Fe) in
compost at 37°C. (c) represents the normalized Mn reduction as a function of time for PLA in
control compost and compost biostimulated with Fe3O4. The experimental data was fitted
using a first-order reaction of the form Mn /Mno = e(−kt), where Mno is the initial Mn, k is the
rate constant, and t is the time. The inset shows the k-fitted values; values with different
lowercase letters are statistically different (α = 0.05, Tukey-Kramer test). (d) and (e) show
the MWD of PLA in compost and compost biostimulated with Fe3O4, respectively.
5.4.2 Mn reduction for PLA with biostimulants in compost
PLA samples were retrieved separately from the control compost and compost
biostimulated with skim milk, gelatin, and ethyl lactate. Figure 5.2 shows the reduction in
Mn of PLA and biostimulated PLA tracked until the end of the test (180 days). A significant
difference was observed in the kinetic reduction rates of PLA with biostimulant treatment
compared with no biostimulation treatment. The CO2 evolution values for PLA, PLA + skim
milk, PLA + gelatin, and PLA + ethyl lactate are provided in Appendix 5B.
282
Skim milk was added to the compost to induce protease activity by the microbes [48–
50]. Serine protease (3.4.21.112) belongs to the peptidases and is the class of extracellular
enzymes able to hydrolyze the peptide bonds linked to amino acids in the protein structure.
Skim milk is composed of different proteins, such as lactose, casein, and whey protein,
making it a good precursor for enzymatic activity, as mentioned earlier. Other researchers
have previously used skim milk to demonstrate extracellular protease synthesis. The
microorganisms present in compost secrete protease to hydrolyze the milk protein. This
protease is used by microorganisms capable of PLA degradation to depolymerize PLA [51].
This increase in k can be deduced as a final reduction of around 75 days on the biodegradation
time when PLA in compost biostimulated with skim milk reaches Mn ≲10kDa at 420 days
(Figure 5.2). Microorganisms assimilate the PLA n-mers at this stage, accelerating the
biodegradation stage [44]. On the other hand, PLA needs at least 494 days to reach the same
Mn – an effective 15% reduction of time.
Gelatin is composed of protein and amino acids, and is a precursor for protease activity
[19,52–54]. The addition of gelatin to compost produced an acceleration of PLA with an
enhancement of k translated to a final reduction of around 124 days when PLA is
biostimulated with gelatin, reaching an Mn ≲10kDa at 371 days compared with at least 494
days for PLA alone – an effective 25% reduction of time (Figure 5.2 ).
Ethyl lactate, on the other hand, was used to stimulate the lactate utilizing microbes
in the compost. Ethyl lactate undergoes hydrolysis to produce ethanol and lactate, where
both can act as a constant long-term supply of hydrogen sources as electron donor compounds
for reductive degradation and microbial redox process. Lactate has been shown to act as an
electron donor compound in the case of anaerobic degradation [55,56]. Lactate has previously
been used for anaerobic degradation, trichloroethane dichlorination, and sulfate reduction
283
[57,58]. PLA + ethyl lactate resulted in a change in k, which can be translated to around 111
days when PLA is biostimulated with ethyl lactate, reaching an Mn ≲10kDa at 384 days
compared to PLA alone – an effective 22% reduction of time (Figure 5.2).
A detailed discussion of the effect of gelatin, skim milk, and ethyl lactate on the
biodegradation of PLA is provided in our previous work [20].
Figure 5.2 Normalized Mn reduction as a function of time for PLA in control compost and
compost biostimulated with skim milk, gelatin, and ethyl lactate. The experimental data was
fitted using a first-order reaction of the form Mn /Mno = e(−kt), where Mno is the initial Mn, k is
the rate constant, and t is the time. The inset shows the k-fitted values; values with different
lowercase letters are statistically different (α = 0.05, Tukey-Kramer test).
5.4.3 Effect of Fe3O4 on cellulose and PLA degradation with gelatin as a
biostimulant
Since gelatin resulted in the most significant Mn reduction for PLA, it was selected to
discuss the effect of a combination of biostimulants (i.e., Fe3O4 nanopowder and gelatin). The
data for Fe3O4 nanopowder, skim milk, and ethyl lactate are provided in Appendix 5B. When
284
Fe3O4 nanopowder was introduced in compost amended with gelatin, the gelatin acted as a
precursor for the protease enzyme secretion by the microbes present in the compost and the
Fe3O4 nanopowder provided the metal to catalyze the hydrolysis.
Figure 5.3a and b show the CO2 evolution and mineralization of cellulose + Fe,
gelatin + Fe, PLA + Fe, cellulose + gelatin + Fe (hereafter referred to as cell + gel + Fe), and
PLA + gel + Fe in compost at 37°C. Gelatin + Fe resulted in around 60.1 g of CO2 evolution
and mineralization of 247.8% in 180 days. Gelatin was combined with Fe to target the
chemical hydrolysis and enzymatic degradation steps. The CO2 evolution in this case
(Cell+gel+Fe 1) was 64.6 g, which is as expected and higher compared with the individual
values for cellulose + Fe and gelatin + Fe and a mineralization of 97.7%. To better understand
the interaction and to account for the degradation behavior of cellulose in the presence of
gelatin (Cell+gel+Fe 2), the mineralization value was estimated at 80.5% by subtracting the
background signal from the bioreactor containing gelatin + Fe. This higher value indicates
that cellulose degradation was not affected by gelatin, and both are used up by the
microorganisms as carbon sources.
285
Figure 5.3 Cumulative CO2 evolution (a) and mineralization (b) of blank + Fe, cellulose +
Fe, PLA + Fe, gelatin + Fe, cellulose + gelatin + Fe (Cell+gel+Fe), PLA + gelatin + Fe
(PLA+gel+Fe) in compost at 37°C. (c) represents the normalized Mn reduction as a function
of time for PLA in control compost and compost biostimulated with gelatin, and gelatin + Fe.
The experimental data was fitted using a first-order reaction of the form Mn /Mno = e(−kt), where
Mno is the initial Mn, k is the rate constant, and t is the time. The inset shows the k-fitted
values; values with different lowercase letters are statistically different (α = 0.05, Tukey-
Kramer test). (d) and (e) depict DSC thermograms for PLA + Fe and PLA + Fe in compost
biostimulated with gelatin. (f) and (g) show the MWD of PLA + Fe in compost and compost
biostimulated with gelatin.
To understand the influence of gelatin + Fe on PLA degradation, PLA was introduced
in the compost amended with gelatin + Fe. The bioreactor containing both PLA and gelatin
+ Fe (PLA+gel+Fe 1) generated CO2 evolution of around 40.5 g and maximum mineralization
286
of 56.5% by the end of the test. Improved mineralization was observed as opposed to no CO2
evolution for PLA alone without any biostimulation of the compost. The effect of gelatin + Fe
on PLA degradation is calculated by plotting the mineralization of PLA+gel+Fe 2
(subtracting gelatin + Fe), as mentioned earlier. Negative mineralization does not necessarily
indicate the absence of gelatin's protease activity in PLA's enzymatic degradation. This
finding is validated by the significant difference observed in the kinetic rate of degradation
for PLA in compost, with and without any biostimulation with gelatin, as seen in Figure 5.3c
inset (PLA+Fe k = 0.0052 ± 0.0001 and PLA+Gel+Fe k = 0.0076 ± 0.0002). The significant
difference in the evolution of Xc from 28.3% to 31.4% for PLA, and from 28.3% to 40.9% for
PLA biostimulated with gelatin + Fe, as seen in Figure 5.3d and e , respectively, further
shows the improvement in the enzymatic degradation of PLA due to the presence of gelatin
+ Fe. Gelatin acts as a precursor for the microbes to release protease enzyme, aiding in the
enzymatic degradation of PLA. The broadening and change in the amplitude of the MWD
peaks through the test duration of 180 days, as seen in Figure 5.3g for PLA in compost
biostimulated with gelatin + Fe compared to PLA + Fe alone in Figure 5.3g , show compelling
evidence for enhanced degradation for PLA in the presence of gelatin and Fe. Iron is an
essential micronutrient, necessary for life-sustaining processes, and plays a critical role in
cell growth of microbes [59]. Iron also functions as a cofactor, promoting and increasing
enzymatic activity. Iron plays important roles in various biological processes such as
respiration, oxido-reduction mechanism, nitrogen fixation, tricarboxylic acid cycle, and
electron transport [33]. S. He et al. demonstrated that a soil matrix amended by Fe
nanoparticles shifted the microbial community composition and stimulated the metabolic
activity of the bacterial community present by enhancing their growth rate [33]. The soil
nitrification potential of the Fe-amended soil was improved by 10% to 19% compared to
287
control soil, indicating that adding Fe aided in the increase of biomass capacity and
eventually enhanced and boosted carbon cycling.
Zhang et al. further showed that the addition of Fe nanoparticles promoted the
degradation of organic matter and amplified the dehydrogenase and urease activities,
significantly improving the overall microbial activity and nitrogen mineralization [47]. Thus,
adding Fe3O4 nanopowder improves the microbial metabolic activity, nitrification potential,
and microbial population. When supplemented with the enzymatic activity associated with
gelatin, these changes improve the degradation of PLA in compost compared with that of
control PLA with no biostimulants present. Y. He et al. showed enhanced enzymatic and
nitrification activity for organic matter degradation in a food-waste composting system due
to compost amendment with Fe-carbon particles [60]. In addition, the bacterial and fungal
communities exhibited significant improvement in the composting process due to the
presence of iron, which can explain the improved PLA degradation found in the presence of
Fe3O4 nanopowder.
Overall, the changes obtained in k indicate a final reduction of around 148 days when
PLA stimulated with gelatin reaches an Mn ≲10kDa at 346 days compared to at last 494 days
for PLA – an effective 25% reduction of time. Similarly, time reductions of 17% or 30% were
observed when PLA + Fe3O4 or gelatin and Fe3O4 nanopowder, respectively, were included in
the compost.
As organic waste disposal is becoming more stringent worldwide and landfill disposal
bans are increasing, several food industries are being impacted and need to find alternative
end-of-life scenarios. The gelatin industry produces a large amount of sludge, resulting in a
tremendous amount of waste generated, which includes collagen fibers, bone residues, and
other inorganic materials. This gelatin sludge usually ends up in landfill or waste
288
management treatment without any pretreatment and creates several problems such as
water contamination, greenhouse gas emission, and health risks for local habitats [61]. So,
this waste could be diverted from landfills and used to produce mature compost along with
the organic fraction of municipal solid waste. Gelatin sludge is high in nitrogen and organic
matter content and can act as a valuable plant nutrient [61]. The nutritional value of the
compost generated from treating gelatin waste and Fe3O4 nanopowder, considering the
benefits mentioned earlier, complement each other to improve soil fertility once the compost
is applied to agricultural land. The selection and combination of specific compounds can open
a new route to accelerate the degradation of compostable polymers in industrial and home
composting operations.
5.5
Conclusion
We investigated the role of different compounds—skim milk, gelatin, and ethyl lactate
in combination with Fe3O4 nanopowder—on PLA degradation at 37°C by biostimulating the
compost media. The different compounds were selected to target different stages of the
biodegradation process. Without any biostimulant compounds, PLA continued to undergo a
long abiotic lag phase affirming a slow hydrolysis phase, which was seen as a negative
mineralization for the test duration of 180 days, whereas a boost in CO2 evolution for PLA
was observed in compost amended by Fe3O4 nanopowder in combination with gelatin, skim
milk, or ethyl lactate. This observation was verified by the molecular weight change and
crystallinity evolution. PLA biodegradation was accelerated by 30% to reach the biotic phase
of Mn ≲10kDa by the addition of gelatin and Fe3O4 nanopowder. The addition of biostimulants
opens new avenues to improve PLA biodegradation in home composting conditions.
289
5.6
Acknowledgments
P.M. acknowledges the School of Packaging at Michigan State University; R.A.
acknowledges the USDA National Institute of Food and Agriculture and Michigan State
University AgBioResearch, Hatch project number MICL02665, for partial study support.
290
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296
APPENDIX 5A: PHYSICOCHEMICAL CHARACTERISTICS
Some compost was collected and sent to the Soil and Plant Nutrient Laboratory at
Michigan State University (East Lansing, MI, USA) to evaluate its physicochemical
parameters (dry solids, volatile solids, and C/N ratio) as previously described elsewhere [44].
The physicochemical parameters are reported below in Table A5.1.
Table A5.1 Physicochemical parameters and total nutrient analysis of compost used in the
biodegradation test.
Parameter
Dry solids, %
Volatile solids, %
pH
C/N ratio
Carbon, %
Nitrogen, %
Phosphorus, %
Potassium, %
Calcium, %
Magnesium, %
Sodium, %
Sulfur, %
Iron, ppm
Zinc, ppm
Manganese, ppm
Copper, ppm
Boron, ppm
Aluminum, ppm
Compost
42.5
41.7
8.0
10.1
24.2
2.42
1.21
3.15
5.07
2.82
0.58
0.58
9878
480
413
107
41
6751
297
APPENDIX 5B: CO2 EVOLUTION AND MINERALIZATION OF PLA IN THE
PRESENCE OF SKIM MILK, GELATIN, AND ETHYL LACTATE
Figure A5.1 a and b shows the CO2 evolution and % mineralization of cellulose, skim
milk, PLA, cellulose + skim milk, and PLA + skim milk. Cellulose reached mineralization of
87.7% whereas skim milk attained over 100 % of its carbon conversion over a period of 180
days. Since skim milk and cellulose are readily biodegradable and can be easily utilized as a
carbon source by the microorganisms present in the compost, no lag phase was observed.
Skim milk was added to the compost with the goal of inducing the protease activity of the
microbes present. PLA shows similar CO2 evolution when compared to blank (compost only).
This indicates that PLA is still undergoing chemical hydrolysis and is yet to breakdown to
Mn of 10k Da, where it can be assimilated by microorganisms.
Due to the compost amendment with skim milk, PLA (PLA + skm1) shows a
mineralization of approximately 60%. In order to account for the effect of skim milk, a
separate mineralization plot (PLA + skm2) is derived. Around 35% mineralization is observed
for the same depicting that the enzymatic degradation of PLA is enhanced due to the presence
of skim milk. This is corroborated by the molecular weight analysis (PLA k = 0.0045 ± 0.0001
and PLA-skim milk k = 0.0053 ± 0.0003).
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Figure A5.1 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
skim milk, cellulose + skim milk (Cell+skm), PLA + skim milk (PLA+skm) in compost at
37°C.
299
Figure A5.2 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
gelatin, cellulose + gelatin (Cell+gel), PLA + gelatin (PLA+gel) in compost at 37°C.
300
Figure A5.2 a and b shows the CO2 evolution and % mineralization of cellulose,
gelatin, PLA, cellulose + gelatin, and PLA + gelatin. Cellulose reached mineralization of
87.7% whereas gelatin attained over 100 % of its carbon conversion over a period of 180 days.
Since gelatin and cellulose are readily biodegradable and can be easily utilized as a carbon
source by the microorganisms present in the compost, no lag phase was observed.
Gelatin is composed of protein which the microorganisms in compost use for their
biochemical process. The microorganisms secrete protease enzyme to digest gelatin which is
the same mechanism when PLA is introduced in gelatin amended compost. Due to the
compost amendment with skim milk, PLA (PLA + gel1) shows a mineralization of
approximately 60%. In order to account for the effect of gelatin, a separate mineralization
plot (PLA + gel2) is derived. Though there seems to be negative mineralization, the molecular
weight analysis [62] shows that gelatin helps in the enzymatic degradation of PLA. This
coupled with the chemical hydrolysis of PLA [5], produces a significant difference with
respect to the kinetic rate of degradation (PLA k = 0.0045 ± 0.0001 and PLA-gelatin k = 0.0060
± 0.0002) [62].
301
Figure A5.3 Cumulative CO2 evolution (a) and mineralization (b) of blank, cellulose, PLA,
ethyl lactate, cellulose + ethyl lactate (Cell+el), PLA + ethyl lactate (PLA+el) in compost at
37°C.
302
Figure A5.3 a and b shows the CO2 evolution and % mineralization of cellulose, ethyl
lactate, PLA, cellulose + ethyl lactate, and PLA + ethyl lactate. Cellulose reached
mineralization of 87.7% whereas ethyl lactate attained over 100 % of its carbon conversion
over a period of 180 days. Since ethyl lactate and cellulose are readily biodegradable and can
be easily utilized as a carbon source by the microorganisms present in the compost, no lag
phase was observed. Ethyl lactate was used to stimulate the lactate utilizing microbial
community present in the compost.
Ethyl lactate evolves around 51.5 g of CO2, and the corresponding mineralization
reaches around 270%. The CO2 evolution of the amended compost containing both cellulose
and ethyl lactate (Cell-el 1) was only around 66.2 g and there was a corresponding
mineralization of 182.4%. PLA in the presence of ethyl lactate (PLA-el 1) evolved around 51.6
g of CO2, followed a similar trend as ethyl lactate over 180 days, and showed maximum
mineralization of 105%. A positive mineralization behavior is seen for PLA when the effect
of ethyl lactate is accounted for (PLA-el 2), indicating the lactate-stimulating activity of ethyl
lactate. This is further confirmed by kinetic rates for PLA alone and PLA in compost amended
with ethyl lactate (PLA k = 0.0045 ± 0.0001 and PLA-ethyl lactate k = 0.0058 ± 0.0002) [62].
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APPENDIX 5C: CO2 EVOLUTION AND MINERALIZATION OF PLA IN THE
PRESENCE OF FE3O4 NANOPOWDER, SKIM MILK + FE, AND ETHYL
LACTATE + FE
Figure A5.4 Cumulative CO2 evolution (a) and Mineralization (b) of blank + Fe, cellulose +
Fe, PLA + Fe, skim milk + Fe , cellulose + skim milk + Fe (Cell+skm+Fe), PLA + skim milk
+ Fe (PLA+skm+Fe) in compost at 37°C. (c) represents the normalized Mn reduction as a
function of time for PLA in control compost and compost biostimulated by gelatin, and gelatin
+ Fe. The experimental data was fitted using a first-order reaction of the form Mn /Mno =
e(−kt), where Mno is the initial Mn, k is the rate constant, and t is the time. The inset shows
the k-fitted values. Values in the column with different lowercase letters are statistically
different (α = 0.05 Tukey-Kramer Test). (d) and (e) depict DSC thermograms for PLA + Fe
and PLA + Fe in compost biostimulated by skim milk. (f) and (g) shows the MWD of PLA +
Fe in compost and compost biostimulated with skim milk.
Figure A5.4 a and b shows the CO2 evolution and mineralization of cellulose + Fe,
skim milk + Fe, PLA + Fe, cell + skm + Fe, and PLA + skm + Fe in compost at 37°C. Skm +
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Fe shows around 60.1 g of CO2 evolution and mineralization of 247.8 % in 180 days. Skim
milk was combined with Fe to target chemical hydrolysis and enzymatic degradation steps.
The CO2 evolution in this case (Cell+skm+Fe) sees a higher production of 54.5 g, which is as
expected and higher compared to the individual values for cellulose + Fe (44.5 g).
Cell+skm+Fe 1 shows a mineralization of 121.6% whereas Cell+skm+Fe 2 after accounting
for skm+Fe shows a mineralization of 99.4% indicating that the presence of skim milk in no
way affects the degradation of cellulose.
To understand the influence of skim milk + Fe on PLA degradation, PLA was
introduced in the compost amended with skim milk and Fe. The bioreactor containing both
PLA and skim milk + Fe ( PLA+skm+Fe 1) shows CO2 evolution of around 36.4 g and
maximum mineralization of 35.3 % by the end of the test. Improved mineralization is
observed as opposed to no CO2 evolution for PLA alone without any biostimulation of compost.
The effect of skim milk + Fe on PLA degradation is calculated by plotting the mineralization
of PLA+skm+Fe 2 (subtracting skim milk + Fe). The negative mineralization does not
necessarily indicate the absence of skim milk’s protease activity in PLA's enzymatic
degradation. The significant difference in the evolution of Xc from 28.3% to 31.4% for PLA,
and from 28.3% to 37.9% for PLA samples biostimulated with skim milk + Fe, as seen in
Figure S4 d and e, respectively further shows the improvement in the enzymatic degradation
of PLA due to the presence of skim milk + Fe. Skim milk acts as a precursor for protease
activity and the broadening and change in the intensity of peaks as seen in Figure A5.4 f
and g enforces that the addition of skim milk and Fe does enhance PLA degradation.
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Figure A5.5 Cumulative CO2 evolution (a) and Mineralization (b) of blank + Fe, cellulose +
Fe, PLA + Fe, ethyl lactate + Fe , cellulose + ethyl lactate + Fe (Cell+el+Fe), PLA + ethyl
lactate + Fe (PLA+el+Fe) in compost at 37°C. (c) represents the normalized Mn reduction as
a function of time for PLA in control compost and compost biostimulated by gelatin, and
gelatin + Fe. The experimental data was fitted using a first-order reaction of the form Mn
/Mno = e(−kt), where Mno is the initial Mn, k is the rate constant, and t is the time. The inset
shows the k-fitted values. Values in the column with different lowercase letters are
statistically different (α = 0.05 Tukey-Kramer Test). (d) and (e) depict DSC thermograms for
PLA + Fe and PLA + Fe in compost biostimulated by ethyl lactate. (f) and (g) shows the MWD
of PLA + Fe in compost and compost biostimulated with ethyl lactate.
Figure A5.5 a and b shows the CO2 evolution and mineralization of cellulose + Fe,
ethyl lactate + Fe, PLA + Fe, cell + el + Fe, and PLA + el + Fe in compost at 37°C. El + Fe
shows around 54.5 g of CO2 evolution and mineralization of 288.1 % in 180 days. Ethyl lactate
was combined with Fe to target chemical hydrolysis and lactate utilizing microbes in compost.
The CO2 evolution in this case (Cell+el+Fe) sees a higher production of 62.2 g, which is as
306
expected and higher compared to the individual values for cellulose + Fe (44.5 g). Cell+el+Fe
1 shows a mineralization of 156.7% whereas Cell+el+Fe 2 after accounting for el+Fe shows a
mineralization of 75.1% indicating that the presence of ethyl lactate does not affect the
degradation of cellulose.
To understand the influence of ethyl lactate + Fe on PLA degradation, PLA was
introduced in the compost amended with ethyl lactate and Fe. The bioreactor containing both
PLA and ethyl lactate + Fe ( PLA+el+Fe 1) shows CO2 evolution of around 46.5 g and
maximum mineralization of 88.8 % by the end of the test. Enhanced mineralization is
observed as opposed to no CO2 evolution for PLA alone without any biostimulation of compost.
The effect of ethyl lactate + Fe on PLA degradation is calculated by plotting the
mineralization of PLA+el+Fe 2 (subtracting ethyl lactate + Fe). The negative mineralization
does not necessarily indicate the absence of ethyl lactate’s lactate stimulating microbial
activity in PLA's enzymatic degradation. The significant difference in the evolution of Xc from
28.3% to 31.4% for PLA, and from 28.3% to 39.9% for PLA samples biostimulated with ethyl
lactate + Fe, as seen in Figure S5 d and e, respectively further shows the improvement in the
enzymatic degradation of PLA due to the presence of ethyl lactate + Fe. As seen in Figure S5
f and g ethyl lactate stimulates the lactate utilizing microbial community in the compost
which aids in the degradation of PLA.
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CHAPTER 6: ENHANCING BIODEGRADATION OF POLY(LACTIC ACID) IN
MESOPHILIC AND THERMOPHILIC ENVIRONMENTAL CONDITIONS: THE
ROLE OF PROTEINASE K AS A PRETREATMENT
6.1
Abstract
Plastic pollution poses a significant environmental challenge, prompting a shift
towards biodegradable and biobased polymers such as poly(lactic acid) (PLA) as sustainable
alternatives. Despite PLA's potential, a slow abiotic process hinders its degradation in
composting environments, influenced by factors like high number molecular weight and
crystallinity. Enzymatic degradation, particularly by PLA depolymerase like proteinase K,
offers a promising solution to accelerate the degradation of PLA during composting if used as
pretreatment. This study aimed to investigate the enzymatic degradation of PLA films
pretreated with proteinase K and degraded in simulated home and industrial composting
conditions. PLA films were pretreated with proteinase K at 37°C and 58°C at different times.
PLA films treated in underwater conditions at 37°C were used as control. The accelerated
CO2 evolution and mineralization of the pretreated PLA films highlight the effectiveness of
proteinase K pretreatment. This study shows the potential of enzymatic pretreatment to
enhance PLA biodegradability at mesophilic and thermophilic temperatures, offering
insights into sustainable waste management strategies to reduce the biodegradation rate of
PLA films and packaging.
6.2
Introduction
Plastic pollution in the environment is a looming crisis. Given the crucial role that
plastics play in creating single-use plastic packaging and the generation of white pollution,
using biodegradable polymers in day-to-day life to combat mismanaged waste is seen as a
welcoming solution. So, the commercial plastic industry is adopting biobased and
308
biodegradable plastics as an environmentally
friendly alternative to
fossil-based
conventional polymers. Poly(lactic acid) - PLA - is one such polymer and is considered a major
substitute for conventional fossil polymers because of its designed biodegradable nature
when exposed to the proper set of composting conditions. So, the use of PLA has increased
rapidly to support the growth of the circular sustainable bioeconomy, accompanied by the
ongoing development in its production process.
Although PLA and its products are labeled as industrial compostable, it is essential
to note that the time frame for the degradation process is longer as compared to the readily
biodegradable organic waste fraction such as food, starch, and cellulose when PLA is collected
together as a part of municipal solid waste and directed to an industrial composting facility
[1]. This scenario arises because of the slow abiotic phase, which determines its degradation
rate and is further influenced by molecular weight (Mn) and crystallinity (Xc). PLA is
subjected to higher temperatures of 58°C-70°C in industrial composting facilities, which
makes it able to degrade. However, its degradation is recalcitrant in soil and home
composting environments where temperatures are much lower in the mesophilic range.
In environments such as soil and home composting, chemical hydrolysis proceeds at a
prolonged rate due to the lower temperatures encountered, and biodegradation is mainly
governed by biotic enzymatic degradation [2]. Abiotic degradation proceeds by reducing the
Mn to values lower enough that the biotic enzymatic degradation can take over and accelerate
the breakdown process. Biotic enzymatic degradation involves the breakdown of polymer
chains into small molecules that the microorganisms can quickly assimilate. The
biodegradation at such mesophilic conditions relies heavily on the depolymerization stage,
wherein the microorganisms release the enzymes. For hydrolyzable polymers such as PLA,
the main enzymes belong to the hydrolases (EC 3) class of extracellular enzymes reported for
depolymerizing aliphatic polyesters. Depending on the substrate specificity, the PLA
309
depolymerases are categorized as protease and lipase types. Both classes of PLA
depolymerases employ the serine hydrolase catalytic mechanisms, yet their different
stereochemistry at the catalytic sites provides a structural basis for different specificities for
substrates [3].
Proteinase K (3.4.21.64) and proteases (3.4.21.112) belong to the serine endo
peptidases (3.4.21), which catalyze bond scission in the middle of the substrate chain and are
also known to hydrolyze polyesters such as PLA. Proteinase K and proteases show enzymatic
activity specifically for PLA containing L-lactate compared to PDLA and PDLLA. These
enzymes can recognize PLA's repeating L-lactic acid unit as structurally homologous to the
proteins composed of L-aminoacids [4]. Williams [5] first reported the hydrolysis of PLA by
proteinase K sourced from Tritirachium album. Since then, the degradation of PLA by
different enzymes has been studied substantially [2]. The commercially available proteinase
K has been used extensively to study PLA enzymatic degradation.
Unlike mechanical and chemical recycling, enzymatic degradation consumes less
energy, needs fewer chemicals, or generates less harmful compounds threatening the
environment. So, enzymatic degradation can assist in further developing the degradation of
PLA at lower temperatures. This study aimed to evaluate the enzymatic degradation of PLA
by pretreating PLA with proteinase K enzyme in a buffer solution at 37°C and 58°C for
different time intervals before introducing it into a simulated compost environment at 37°C
to replicate home composting and at 58°C to simulate industrial composting. The CO2
evolution of the samples was recorded, and the changes in Mn and Xc throughout the test
duration were measured to understand how the pretreatment with proteinase K modified the
biodegradation of PLA in the different composting conditions.
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6.3 Materials and methods
6.3.1 Materials
Crystalline PLA films ranging 25 microns in thickness were obtained from EarthFirst
(Columbus, OH, USA). The crystalline PLA films were heated at 180°C, for 10 mins and fast
quenched in dry ice for 10 minutes to remove any residual crystallinity. Proteinase K from
Tritirachium album (29.3 kDa) from Syd labs (Hopkinton, MA) was procured and used as
received for enzymatic degradation. HPLC grade Tetrahydrofuran (THF) was obtained from
(Sigma-Aldrich, St. Louis, MO, USA). All the chemicals were used as received.
6.3.2 Preparation of buffer and enzyme solution
Buffer powder (MSU, East Lansing, MI) was mixed in distilled water to prepare the
buffer solution. NaOH pellets were mixed in the distilled water and added to the buffer
solution until the pH reached 8.5. The proteinase K enzyme was added to the distilled water
to get an aqueous solution of proteinase K (500 μg/mL). Based on the previous reports, the
concentration was calculated and set to 500 μg/mL [6,7].
6.3.3 Enzymatic degradation by externally adding Proteinase K
The enzymatic degradation of PLA films was performed by externally adding the
proteinase K solution and buffer solution to the PLA films ( 1 cm x 1 cm squares). The reaction
solution was incubated at 37°C for 7 and 10 days and at 58°C for 2 and 5 days with constant
stirring to determine the effect of exposing PLA films to Proteinase K at different intervals.
6.3.4 Hydrolysis experiment
A hydrolysis test was also run as a control to understand the hydrolytic degradation
of the PLA films. The test procedure was adapted from ASTM D4754-18 [8]. The PLA films
were cut into 1 cm x 1 cm and introduced into the beaker with HPLC-grade water (VWR,
Radnor, PA, USA) at 37°C for 10 days. At the end of the test, the samples were dried and
stored for further analysis.
311
6.3.5 Biodegradation in vermiculite
The biodegradation of PLA and the effectiveness of pretreating PLA films with
proteinase K in inoculated vermiculite on the degradation of PLA were evaluated under
aerobic simulated mesophilic and thermophilic conditions using two direct measurement
respirometric (DMR) systems [9–11]. Shortly, the system included a non-dispersive infrared
gas analyzer (NDIR) (Li-COR® LI-820, Lincoln, NE, USA) which measures the CO2
concentration. The DMR system chamber was maintained at a temperature of 37 ± 2 °C and
58 ± 2 °C and provided with air at a relative humidity (RH) of 50% ± 5%. A flow rate of CO2-
free air (concentration <30 ppm to establish a low baseline) was controlled at 40 ± 2 cm3/min.
Additional detailed information about the DMR equipment can be found in another source
[12].
Mature compost obtained from the MSU composting facility was sifted using a 10-mm
screen to get rid of any considerable debris or chunks present and then conditioned at 37 ± 2
°C and 58 ± 2 °C until use. This compost extract was then amalgamated with a mineral
solution in a 1:1 ratio, resulting in the inoculum solution. Detailed information regarding the
preparation of the mineral solution can be found elsewhere [13]. Inoculated vermiculite
provides the benefit of very low CO2 evolution from the blank used as the baseline. Deionized
water was used to adjust the moisture content of the vermiculite during testing to 50%. The
resulting inoculated vermiculite mixture was sent to the Soil and Plant Testing Laboratory
at the University of Missouri (Columbia, MO, USA) to determine the physicochemical
parameters of the media. Data regarding the solid analysis is presented in Table A6.1,
Appendix 6A. The bioreactors were packed with 400 g of inoculated vermiculite, and 8 g of
all the samples were tested in triplicate and positive control (cellulose). Blank (only compost)
was used as the baseline.
312
6.3.6 Size exclusion chromatography (SEC)
As described elsewhere, the Mn and molecular weight distribution (MWD) of PLA for
the control and each biostimulant treatment were measured using SEC (Waters Corp.,
Milford, MA, USA) [14]. Shortly, PLA samples weighing approximately 10 mg were retrieved
at predetermined intervals and dissolved in 5 mL of THF. A temperature of 35°C and a 1
mL/min flow rate were maintained during testing. The Mark-Houwink constants of K =
0.000174 dL/g and α = 0.736 were used to determine the absolute Mn, Mw, and MWD of the
PLA samples. Data analysis was carried out using Waters BreezeTM2 software from Waters.
6.3.7 Differential scanning calorimetry
A DSC model Q100 (TA Instruments, New Castle, DE, USA), was used to determine
the glass transition temperature (Tg), (Tm), crystallization temperature (Tc), and crystallinity
(Xc) for the PLA samples retrieved from the regular and biostimulated compost. PLA samples
weighing 5 - 10 mg were packed in aluminum pans, cooled to −5°C, and then subjected to a
heating cycle to reach 210°C at a ramp rate of 10°C/min. This information was used to
evaluate the evolution of Xc. The cooling was achieved using a nitrogen cooling system that
maintained the purge flow rate at 70 mL/min. The degree of crystallinity was estimated using
equation (6.1):
𝜒𝑐 % =
𝛥𝐻𝑚 − 𝛥𝐻𝑐
0
𝛥𝐻𝑚
𝑥 100
(6.1)
where 𝛥𝐻𝑚 is the heat of fusion, 𝛥𝐻𝑐 is cold crystallization enthalpy and 𝛥𝐻𝑚
0 is the heat of
fusion for 100% crystalline pure PLA (93 J/g) [15].
6.3.8 Statistical analysis
The statistical analysis was conducted using MINITABTM software (Minitab Inc.,
State College Park, PA, USA). The statistical significance at p < 0.05 was evaluated using
one-way ANOVA and Tukey-Kramer test. Data is reported as means ± standard deviation.
313
6.4 Results and discussion
The CO2 evolution of PLA control (PLA), PLA hydrolyzed films at 37°C for 10 days
(PLA 37 hydro 10 D), and enzymatically pretreated PLA samples over 7 and 10 days at 37°C
(PLA-37 proteinase K 7D and PLA-37 proteinase K 10D) and PLA samples over 2 and 5 days at 58°C
(PLA-58 proteinase K 2D and PLA-58 proteinase K 5D) were tracked at simulated mesophilic conditions
(37°C) and thermophilic (58°C) conditions using inoculated vermiculite as media,
respectively.
6.4.1 Mn reduction and crystallinity evolution for PLA and PLA pretreated films
The Mn and Xc of PLA control, PLA hydrolyzed films at 37°C for 10 days, PLA pretreated
with proteinase K at 37°C for 7 and 10 days, and PLA pretreated with proteinase K at 58°C
for 2 and 5 days was determined to understand the effect of proteinase K pretreatment. Table
6.1 presents the values for the same before introducing the samples in the DMR chamber for
further biodegradation testing.
Table 6.1 Initial characterization of PLA, PLA hydrolyzed films, and PLA films treated with
proteinase K at 37°C and 58°C.
Films
Treatment
time (days)
PLA Control
PLA
hydrolysis
PLA
proteinase
K
0
10
7
10
2
5
37 ± 2 °C
58 ± 2 °C
Molecular
weight
(Mn, kDa)
Crystallinity
(Xc)
Molecular
weight
(Mn, kDa)
Crystallinity
(Xc)
79 ± 3.0a
0
79 ± 3.0a
0
75 ± 2.8a
2.5± 2.0a
75 ± 2.8a
2.5± 2.0a
38 ± 1.0b
7.1 ± 1.5b
17 ± 0.9c
12 ± 1.3c
-
-
-
-
-
-
-
-
61 ± 3.0b
23 ± 3.5b
47 ± 2.9c
43 ± 3.0c
A significant difference was seen for Mn for PLA control, PLA hydrolyzed films, and
PLA films pretreated with proteinase K at both 37°C and 58°C. Apart from PLA and PLA
314
hydrolyzed films, again, a significant difference was seen for Xc for PLA proteinase K
pretreated films. This indicates that proteinase K enzymatically hydrolyzes PLA and reduces
it to a lower Mn. The values differ for the different pretreatment time intervals, as seen in
Table 6.1 for 37°C and 58°C. The difference can be associated with the optimal activity of
proteinase K at 37°C. The PLA hydrolyzed films do not show any difference in Mn and Xc
compared to PLA control films because of the slower chemical hydrolysis at 37°C.
6.4.2 CO2 evolution and mineralization of PLA and PLA pretreated films in
inoculated vermiculite
Figure 6.1 a and b present the CO2 evolution and % mineralization of the blank (vermiculite
only), cellulose, PLA control films, PLA hydrolyzed films at 37°C for 10 days, and PLA
pretreated with proteinase K enzyme for 7 days and 10 days, respectively in inoculated
vermiculite at 37 ± 2 °C. Blank showed CO2 evolution of 2.4 g by the end of day 100.
315
Figure 6.1 CO2 evolution and % mineralization of PLA, PLA 37hydro 10D, PLA-37 proteinase K
7D,and PLA-37 proteinase K 10D films in compost 37 °C (a & b) and PLA, PLA 37hydro 10D, PLA-
58 proteinase K 2D, PLA-58 proteinase K 5D films in compost at 58 °C (c & d). The shade in the
background for each material represents the standard error between replicates.
Cellulose showed CO2 evolution of 14.3 g and mineralization of c. 94% by day 100. PLA
films subjected to hydrolysis at 37°C for 10 days showed a very low CO2 evolution of 2.5 g,
whereas PLA control films showed only 2.7 g. A lag phase is observed for both PLA control
and PLA hydrolyzed films, which could mean that PLA offers a physical hydrophobic barrier
to water and air before hydrolysis, and the microbes have difficulty accessing the carbon
source as nutrients. The absence of mineralization does not necessarily imply the absence of
hydrolytic degradation or enzymatic activity due to the action of extracellular enzymes
secreted by the microbes but more like inhibition of the microbial activity due to the
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hydrophobic layer barrier created by the presence of the high Mw PLA. The lower values for
both indicate that PLA is still undergoing hydrolysis since PLA is exposed to temperatures
lower than its Tg (c. 60 °C). PLA segments have little to no mobility and are not flexible below
Tg, preventing diffusion or attack by water. Since the initial and rate-limiting step in PLA
degradation is chemical hydrolysis, the lack of chain scission accompanied by the hydrophobic
barrier provided by PLA can explain the lack of mineralization both for PLA control and PLA
hydrolyzed films.
For PLA films pretreated with proteinase K enzyme for 7 days, CO2 evolution of 6.6 g
and mineralization of c. 28% was seen. In contrast, for PLA films pretreated with proteinase
K enzyme for 10 days, 7.7 g of CO2 was evolved, and c. 35% mineralization was observed by
day 100. No lag phase was observed for proteinase K pretreated films, indicating that the
pretreatment with proteinase K reduces PLA to Mn of ~ 10,000 Da, wherein the biotic
degradation stage kicks in, and the microorganisms can utilize PLA for their biochemical
processes. Figure 6.1c and d present the CO2 evolution and % mineralization of the blank
(vermiculite only), cellulose, PLA control films, PLA hydrolyzed films at 37°C for 10 days,
and PLA pretreated with proteinase K enzyme for 2 days and 5 days, respectively in
inoculated vermiculite at 58 ± 2 °C. The blank showed a maximum CO2 evolution of 2.6 g at
day 60. The positive control, cellulose showed CO2 evolution of 13.1 g and mineralization of
c. 84% at day 60. No lag phase is observed since cellulose is readily biodegradable and a food
source for microorganisms. Cellulose is degraded by the action of a battery of enzymes that
work simultaneously and synergistically. Cellulases catalyze the hydrolysis of β-1,4-linkages
in the cellulose [1,16]. The action of exoglucanases and endoglucanases on the ends and at
random internal sections of cellulose’s amorphous region produces varying lengths of cello-
oligosaccharides, which are then hydrolyzed by glucosidases to produce glucose [17]. Glucose
317
is finally converted to CO2 through a series of further cycles. Previous research has shown
fungi, a few bacteria species, and actinomycetes in compost and the soil environment produce
cellulase, which are involved in the degradation of cellulose [18–22].
PLA control films show a lag phase of around 35 days because of the initial abiotic
hydrolysis phase, wherein PLA ester bonds are broken down mainly due to the hydrolysis by
water. The high Mn PLA chains are cleaved to produce low Mn oligomers used by the
microorganisms in the inoculated vermiculite, releasing CO2 and water, which can be
observed during the biotic degradation phase. A CO2 evolution of 7.0 g and mineralization of
c. 28.5% was observed for PLA at 58 °C. PLA films subjected to hydrolysis at 37°C for 10 days
(Figure 6.1b) showed a CO2 evolution of 5.7 g and mineralization of c. 20.5%. PLA hydrolyzed
films followed a similar trend to PLA control films; however, the mineralization is slightly
lower, maybe due to the higher initial crystallinity of PLA 37 hydro 10D films. PLA films
pretreated with proteinase K enzyme at 37°C for 10 days (PLA-37 proteinase K 10D) evolved 9.3 g
of CO2 and showed a mineralization of c. 44.3% at 58°C. The improved degradation can be
attributed to the pretreatment with proteinase K at 37°C for 10 days. Proteinase K belongs
to serine endo peptidases that preferentially catalyze bond scission in the middle of the
substrate chain. The enzymes showing activity towards PLA belong to the protease type
group or peptidases. They can recognize the repeating L -lactic acid unit of PLA as the natural
homolog L -alanine unit of silk fibroin [2,4]. Though proteinase K is primarily known to cleave
peptide bonds, it can exhibit a certain level of non-specific hydrolytic activity towards PLA.
The PLA films were also pretreated with proteinase K enzyme at 58°C for 2 and 5
days, respectively (PLA-58 proteinase K 2D and PLA-58 proteinase K 5D) to evaluate the effect of
temperature and time on the efficiency of proteinase K’s enzymatic activity. PLA films
pretreated for 2 days showed CO2 evolution of 12.2 g and mineralization of c. 62.3%, whereas
PLA films pretreated for 5 days evolved 7.4 g of CO2 and exhibited a mineralization of c. 31%.
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All the films, when pretreated with proteinase K, followed similar trends and showed
considerable mineralization in comparison to the PLA control and PLA hydrolyzed films.
6.5
Conclusion
PLA films were pretreated with proteinase K to evaluate the effect of pretreatment
on the overall biodegradation of PLA films in simulated compost conditions (inoculated
vermiculite.) Subjecting PLA films to proteinase K pretreatment showed significant
improvements in CO2 evolution and mineralization rates, indicating accelerated degradation
compared to untreated PLA films. These findings suggest that enzymatic pretreatment offers
a promising approach to overcoming the challenges faced with PLA degradation, particularly
in composting facilities operating at lower temperatures. Furthermore, the study highlights
the potential of enzymatic hydrolysis as a sustainable and environmentally friendly method
for managing PLA waste. Future research efforts could optimize pretreatment conditions and
explore the scalability of enzymatic degradation processes for large-scale PLA waste
management applications.
6.6
Acknowledgments
P.M. acknowledges the School of Packaging at Michigan State University; R.A.
acknowledges the USDA National Institute of Food and Agriculture and Michigan State
University AgBioResearch, Hatch project number MICL02665, for partial study support.
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321
APPENDIX 6A: PHYSICOCHEMICAL CHARACTERISTICS
Some inoculated vermiculite was collected and sent to the Soil and Plant Testing
Laboratory at the University of Missouri (Columbia, MO, USA) to evaluate its
physicochemical parameters (dry solids, volatile solids, and C/N ratio) as described elsewhere
[23]. The physicochemical parameters are reported below in Table A6.1.
Table A6.1 Physicochemical parameters and total nutrient analysis of compost used in the
biodegradation test.
Parameter
Dry solids, %
Volatile solids, %
pH
C/N ratio
Carbon, %
Nitrogen, %
Phosphorus, %
Potassium, %
Calcium, %
Magnesium, %
Sodium, %
Sulfur, %
Iron, ppm
Zinc, ppm
Manganese, ppm
Copper, ppm
Boron, ppm
Compost
20.0
4.53
6.0-8.0
3.57
0.18
0.05
0.06
3.15
0.24
2.44
0.15
0.07
6811
17
60
59
24
Aluminum, ppm
4.05
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CHAPTER 7: CONCLUSIONS AND RECOMMENDATIONS FOR FUTURE WORK
7.1 Overall conclusion
Biobased, biodegradable polymers have emerged as a sustainable alternative to the
ever-growing escalating waste management and disposal problems due to the increased
fossil-based plastic production. Poly(lactic acid) - PLA, has garnered significant attention due
to its biobased renewable origin, low environmental footprint, compostable nature,
designable biodegradability, and because of its competitive price and expansion in different
sectors such as packaging, plasticulture, and medical industries. Despite being biodegradable
and compostable, the longer biodegradation timeframes associated with its degradation
compared to the readily degradable organic fractions such as starch and cellulose have
hindered PLA acceptance widely.
This dissertation addresses this issue to enhance PLA biodegradation under
mesophilic and thermophilic conditions, to expand its application to different environments
such as soil, and home/backyard composting, and to improve/ achieve a similar degradation
timeframe with that of organic matter encountered in industrial composting settings.
Chapter 2 of this dissertation provides an in-depth literature review of the
biodegradation of biodegradable polymers under mesophilic conditions, the different steps
involved in the process, the role of enzymes, the different test methods used to determine
degradation, key factors affecting the kinetic degradation rate. It also summarizes the
different approaches and combines various factors to provide a holistic understanding of the
complex biodegradation phenomena of biodegradable polymers, including PLA.
Chapter 3 of this dissertation investigated the reactive blending of PLA with
thermoplastic starch (TPS) as one of the methods to accelerate biodegradation under
mesophilic conditions, thereby simulating the home composting setting. The results revealed
that reactive blending ensured uniform dispersion of TPS into the PLA matrix. The presence
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of TPS as a food source for the microorganisms in the compost eliminated the extended lag
phase caused by the chemical hydrolysis process wherein the breakdown of PLA chains to
oligomers, dimers, and monomers, and a major reduction in the molecular weight is achieved.
Since TPS could be used immediately, structural imperfections were created in PLA, further
enhancing its biotic degradation rate. Without any blending, PLA underwent a long lag
phase, which lasted until the end of the test, which was 180 days. The test data from 58°C
biodegradation was also compared to illustrate and highlight the role of TPS in improving
the overall degradation of PLA at thermophilic conditions.
Chapters 4 and 5 explain the use of a biostimulation technique to enhance the
enzymatic biodegradation of PLA at 37°C. Different biostimulants, Fe3O4 nano-powder, skim
milk, gelatin, and ethyl lactate, were identified to target the chemical hydrolysis and the
biotic enzymatic degradation steps to facilitate and accelerate PLA degradation. Fe3O4 nano-
powder, skim milk, gelatin, and ethyl lactate were introduced into the compost media at 37°C,
and CO2 evolution, Mn, and Xc of PLA were monitored through 180 days test duration. The
introduction of individual biostimulants and a combination showed improved enzymatic
degradation of PLA compared to no using biostimulants. Adding biostimulants mainly
enhanced the enzymatic biotic phase, reflected in the improved kinetic degradation rate.
Chapter 6 focused on a pretreatment approach that involved pretreating PLA with
proteinase K enzyme to investigate the effect of enzymes in accelerating enzymatic
degradation. For the most part, in the reported research, enzymatic degradation has been
conducted for a minimal amount of PLA in a liquid media, wherein PLA was the only isolated
carbon source. In this chapter, PLA films were first enzymatically treated with proteinase K
at 37°C and 58°C for different intervals to achieve different Mn and Xc. After pretreatment,
the PLA films were introduced in an inoculated vermiculite solid matrix at 37° and 58°C to
replicate the home and industrial composting environment. The results depicted improved CO2
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evolution and a shortened lag phase for the treated samples compared to the untreated PLA
films.
The different approaches mentioned above offered practical strategies to accelerate
PLA biodegradation and advance the understanding of the complex biodegradation process
at lower temperatures, which are encountered in real end-of-life scenarios. The approaches
utilized above also provide a safe way of degrading PLA, unlike incineration and landfilling,
which bring many problems, including generating toxic gases and using higher temperatures,
thereby increasing pollution, energy consumption, and the leaching of microplastics into the
environment.
7.2 Recommendations for future work
The biodegradation of biodegradable polymers in different environments is a complex
phenomenon that requires an array of expertise to investigate, understand, and advance the
complex involved processes.
Starch has been extensively researched and studied in terms of polymer. Though
adding starch enhances the biodegradation of PLA, several setbacks are encountered while
processing starch into the PLA matrix on a larger scale. The current study only focused on
including starch as one of the additives and focused more on the biodegradation aspect of the
resulting blend. But overall, the blend should be able to provide the necessary properties
expected of a package or film (e.g., food packaging), until the end-of-life cycle. Different
additives and properties enhancement should be further considered and studied to provide a
holistic approach.
One of the main uncontrollable parameters encountered in the home composting
process is temperature due to the unreliable process control. The change from higher to lower
temperatures as we move from industrial to home composting hugely influences the
biodegradation phenomena. The dominant and rate-determining chemical hydrolysis phase
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for PLA proceeds very slowly. Different biostimulants could be explored and studied to
improve the hydrolytic phase without altering or modifying the PLA structure. In the case of
the biostimulation approach, further delving and fine-tuning the optimal concentrations of
biostimulants is essential to maximize their effectiveness while minimizing the associated
costs and environmental impacts of accelerating PLA biodegradation. Systematic studies
must be conducted by varying the concentrations of biostimulants to identify the optimal
concentration and maximize the biodegradation efficiency. To provide a realistic and
economic standpoint, laboratory-level studies should be scaled up to assess the feasibility and
compatibility of the novel methodologies with the existing home infrastructure in place,
specifically when compared to the industrial composting plants. Further testing of the
phylogenetic makeup and microbial interactions due to adding biostimulants during the
composting process should be conducted. This will help better understand the changes
happening in the compost since compostability is a desired end-of-life scenario for
contaminated PLA.
This dissertation used CO2 evolution tracking and molecular weight determination to
determine the acceleration of the biodegradation process, but further tests need to be
conducted to understand the changes taking place at the molecular level and to comprehend
better the process governing the interaction between the biostimulants, native microbial
population in the compost, and PLA as a polymer model system. Carbon 13 label of polymer
could also be a more expanded methodology to evaluate the biodegradation of PLA. Different
scientific methods, such as enzymatic activity assays, spectroscopy analysis, and scanning
electron microscopy imaging, can be complemented with computational modeling techniques
to examine further and unravel complex pathways involved in PLA biodegradation. Machine
learning can also be used to extract and analyze large existing microorganism datasets,
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identify patterns, and develop prediction models for optimizing PLA biodegradation before
running compostability tests.
The enzymatic pretreatment of PLA with proteinase K shows promising results, but
different enzymes, pretreatment conditions, and durations should be further explored. This
should be primarily conducted in a solid matrix such as compost as the current studies are
limited to liquid media, a small amount of PLA tested, and isolated with specific microbial
strains that do not replicate real-life disposal compost scenarios.
This dissertation has provided three different approaches that can be used to enhance
the biodegradation rate of PLA. From a broader perspective, future efforts should focus on
using these novel methodologies in collaboration with industries, governments, and
composting facilities to test the implications and improve PLA recovery.
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