a case study of tenebrio molitor a dissertation submitted

UNDERSTANDING NATURAL PLASTIC BIODEGRADATION:

A CASE STUDY OF TENEBRIO MOLITOR

A DISSERTATION

SUBMITTED TO THE DEPARTMENT OF CIVIL AND ENVIRONMENTAL

ENGINEERING

AND THE COMMITTEE ON GRADUATE STUDIES

OF STANFORD UNIVERSITY

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS

FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

Anja Malawi Brandon

July 2020

http://creativecommons.org/licenses/by-nc/3.0/us/

This dissertation is online at: http://purl.stanford.edu/cs239bh3255

Re-distributed by Stanford University under license with the author.

This work is licensed under a Creative Commons Attribution-Noncommercial 3.0 United States License.

I certify that I have read this dissertation and that, in my opinion, it is fully adequatein scope and quality as a dissertation for the degree of Doctor of Philosophy.

Craig Criddle, Primary Adviser

Richard Luthy

Weimin Wu

Approved for the Stanford University Committee on Graduate Studies.

Stacey F. Bent, Vice Provost for Graduate Education

This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file inUniversity Archives.

Abstract

We live in the “Age of Plastics,” an era in which plastic’s versatility, strength, and low

production costs have led to its ubiquity. Plastics are often designed for single use, leading

to the rapid accumulation of plastic waste. Much of this waste escapes formal collection

and accumulates in natural environments, especially the ocean. Increasing restrictions on

recycling and landfill space are rendering current waste management systems unable to

address current and future plastic waste. Finding a way to degrade plastic waste into valu-

able, sustainable resources and permanently eliminate it from the pollution cycle is critical

for the e↵ective remediation of the natural environment and development of a sustainable

future.

A promising solution to achieve this goal is microbial biodegradation of plastics. How-

ever, the rate of plastic degradation by most mixed microbial cultures and isolated microor-

ganisms is low. E↵orts to improve plastic biodegradation have identified an environment in

which a number of plastics can degrade at relatively rapid rates: the gut of mealworms (the

larvae of Tenebrio molitorLinnaeus). Understanding the mechanisms and conditions re-

quired for degradation within the mealworm could enable the scaling-up of this process as a

sustainable waste management strategy. The research described in this dissertation expands

our understanding of naturally occurring plastic biodegradation mechanisms, drawing on

insights from the study of the mealworm and its gut microorganisms to identify strategies

whereby that understanding can be harnessed in engineered systems. These contributions

advance the field of plastic biodegradation beyond a focus on isolated pure cultures to a

deeper understanding about environments and necessary conditions for plastic degrada-

tion. With this more holistic approach, it is possible to move past the existing paradigm of

single-use plastics to a future of sustainable, environmentally benign materials.

Chapter 2 examines the ability of mealworms to degrade multiple types of plastics,

specifically polyethylene (PE) and a mixture of two of the most common plastic wastes

(PE and PS). This study found that polyethylene, polystyrene, and mixtures of the plastics

degraded at comparable rates. Microbial analysis of the mealworm gut found several mi-

croorganisms strongly associated with the plastic-fed diets. These results provide the first

evidence of a non-specific degradation mechanism within the mealworm gut and highlight

the need for additional research to elucidate this mechanism.

Chapter 3 investigates the ex situ polystyrene degradation achieved by the gut micro-

biome of plastic-degrading mealworms. The gut microbiome was cultured with PS as the

only carbon source to enrich for e�cient plastic-degradation and to enable the isolation of

the bacteria involved. The mealworm gut environment was studied with and without the

gut microbiome to explore the role the insect host during plastic degradation. This research

revealed the important role of the mealworm host in secreting emulsifying factors that me-

diate plastic bioavailability. In addition, plastic-degrading microorganisms were identified

as well as microbial secreted factors that enhance degradation. Chapter 3 demonstrated

the ability to culture plastic-degrading bacteria, which may enable future investigations and

bioaugmentation for enhanced plastic degradation.

Chapter 4 explores the fate of chemical additives in plastics during degradation by

mealworms to assess whether there is evidence of bioaccumulation. Specifically, this study

examines the fate of hexabromocyclododecane (HBCD), a common flame retardant in PS

products. The results reveal that HBCD is rapidly excreted and does not bioaccumulate

within the mealworm biomass. PS-fed mealworm biomass was also used as a feed supplement

for shrimp without evidence of HBCD bioaccumulation. Chapter 4 presents the first

investigation into the fate of chemical additives during plastic degradation. This work

demonstrates the need for further investigations into the environmental impact of secreted

micro- and nano-plastic particles after plastic degradation and the continued need for green

chemistry in designing additives.

Chapter 5 presents a critical review on the state of plastic pollution and remediation

strategies that contextualizes the research conducted in this thesis. The review analyzes

the state of plastic pollution, bioplastic materials, and plastic biodegradation and presents

a framework for transitioning from recalcitrant plastics to biodegradable and non-toxic

replacement materials. This review provides a summary of the state of a rapidly expanding

field and proposes directions for future research to achieve a sustainable future.

The research summarized in this dissertation provides a comprehensive understanding

the rapid biodegradation of plastics in an unexpected natural system: the gut of meal-

worms. Results from these studies contribute to the growing field of plastic degradation

by illustrating how plastics are rapidly biodegraded within the mealworm gut and pro-

vides guidance on how this process might be replicated and scaled-up to enable a viable,

sustainable alternative for plastic waste.

Acknowledgments

I first need to thank my advisor Craig. Thank you for taking a chance on a passionate,

young undergraduate with limited research experience. Thank you for helping me turn an

idea into a PhD. Thank you for challenging me and for supporting me and, ultimately, for

helping me through this journey.

I also need to thank Dr. Wei-Min Wu. You were integral in helping bring this project

to Stanford and for creating its lasting impacts. Thank you for always looking out for me

and my best interests while balancing an increasingly complex, global project.

To the rest of my committee – thank you for taking the time to help me. Bob – thank

you for sharing your expertise and for your patience while I began my journey in polymer

science. Dick – thank you for your kindness and your support throughout this process.

Steve – thank you for making me think in di↵erent ways about the challenges facing the

world and for sharing your time with me.

To Royal – thank you for being the heart and soul of our program! Thank you for

always being willing to help in lab, to chat about nearly anything, and to take our o�ce

puppy out on walks. I cannot imagine making it through this journey without our daily

mezzanine chats, Friday beer laughs, and kitchen discussions. To the administrators who

keep our department and program running (especially Jack Chiueh and Jill Filice) – thank

you, thank you, thank you! From your kindness and patience in the face of frantic emails

to always having the right answer, I truly don’t know how I could have done this without

Throughout this journey I have been incredibly lucky to work with amazing collabora-

tors. I need to than Dr. James Flanagan for helping me develop methods and think through

any number of challenges early on in this project. Thank you to Dr. Shanshan Yang for

your help and your kindness. An incredibly big thank you to Dr. Yeo-Myoung Cho – your

expertise and support was invaluable! I absolutely could not have completed this project

with you. To Mfon – not only were you incredibly helpful with experiments, you brought

a joy and laughter to the lab that was always appreciated! Thank you for your patience,

kindness, and all your help.

To Alexa – you came into this project when I needed you most. You were a breath

of fresh air bringing new ideas, energy, and passion for the project and for science overall.

Thank you for always being open to bouncing around ideas, troubleshooting challenges,

and chatting about life amidst tedious lab work. I’m so glad to have worked with you as a

colleague and even more glad I get to keep you as a friend.

To Sahar – I could not have made it through this PhD without you. Thank you for

being a constant source of joy in the o�ce and for always being there to support me. I’m

so glad I moved up to the mezzanine level to be with you – I will miss swiveling around and

chatting about work (sometimes), life (frequently), and all things puppy-related (always).

I am forever grateful for your friendship these last five years – through the good times and

the bad, I knew you always had my back and that made all the di↵erence.

Andrew – thank you for always reminding me of basic environmental engineering pro-

tocols, being willing to teach me, and for always being there for a good chat.

To the rest of the Criddle group – thank you for making these last five years so great.

Thank you for always o↵ering help around the lab — form teaching me new protocols to

opening freezer doors for me when my back went out, you have all been so amazingly helpful.

I also appreciate all your great feedback over the years. Most importantly, thank you for

helping me keep my spirits up, from our BBQs to holiday parties to chats in the lab – you

are always kept me laughing!

I also need to thank everyone in the EES program for making my time here so great! An

extra big thank-you to the fun committee (and its original founders!) for helping cultivate a

welcoming and inclusive student environment. To Jill and Eily – thank you for role modeling

how to make it through grad school and for taking me under your wings for everything from

PCR assay development to finding a work-life balance, you ladies are amazing and I am

so glad I got to spend time with you! To Stephen – thank you for being an amazingly

thoughtful o�cemate! I will miss our political rants and morning check-ins. To Katy, Alex,

Kirin, and Nicole – thank you for your endless support, fun rants, and for all the laughs!

To my favorite group of hooligans – Wiley, James, and Christine, thank you all for being

such wonderful friends and people! Thank you for reading and editing my work and for

answering my frantic Zoom calls as I stressed about how to set up for a virtual defense.

More importantly, thank you for keeping me laughing, particularly during the very stressful

last few months of this process!

I have been incredibly lucky to have developed an amazingly supportive group of friends

during my time at Stanford. To John and Cassie – thanks for being our best couple and

for being there for me from the beginning of this journey! Thank you for being the best

cheerleaders and celebrating with me during moments of success, cheering me up in moments

of failure, and for hanging out with me during all the moments in between. Jesse - thank

you for your constant positivity and for being curious enough about my work to read the

papers I sent you! Thanks to the virtual reality gang for sanity-saving Zoom calls over these

crazy last few months filled with games, laughter, and cute kitties!

Kira - thank you for sticking with me over my entire Stanford career, we both grew

and evolved and I’m so glad I had you along with me for that process! Thank you for

reminding me to get out of the basement lab, supplying me with ca↵eine and Chinese take-

out, and for being the most supportive friend I could’ve ever asked for. Corey – thank you

for always being there for always being open to talk about science, STEM advocacy, fun

cooking ideas, and all things dogs. Thank you (and Andrei!) for being the most amazing

puppy-godparents to Kona and for all your help early on in our journey of training. I don’t

think I could’ve made it through this last year (especially shelter-in-place!) without you –

thanks for being there when I needed it and for keeping me smiling throughout (shout-out

to Bear for helping with this part too!).

This process has not been easy, beyond academic and research challenges, I, like many

of my peers, su↵ered from anxiety and imposter syndrome. I am thankful I was able to get

help and only after doing so did I realize how common it was. I want to use this space to

encourage anyone in the same position to seek out the help you need – there is no prize

for making it through this journey alone. Beyond mental health, I also struggled with a

major back injury that took me out of the lab (repeatedly) for several weeks and involved

a lengthy recovery. That recovery would not have been possible without the help of Zaldy

De La Cuesta from MORE Physical Therapy. Thank you for teaching me that, like grad

school, recovery is not a linear process and that setbacks are just part of the journey.

I also need to thank the people that helped inspire me to take this journey in the first

place. To Seattle Girls’ School - thank you for teaching me to think creatively and how

to stand and deliver, knowing I belong in any room. To the Seattle Aquarium, especially

the Youth Ocean Advocates program, Dave Glenn, and Katrina Bettis – thank you for

introducing me to the amazing world beneath the surface of our oceans and for teaching

me to lead with my passion.

To Mike, Mary, and Katie – thank you for welcoming me into your family, for supporting

me throughout this journey, and for taking me on much needed breaks! I couldn’t have

made it through this process without your love, kindness, and support. I am so excited for

more adventures together and for exploring even more wine regions!

To my family – I could not have done this without your unconditional support and love.

To Peace and DJ - thank you for putting up with all my lab-talk, for being my biggest

supporters, and for reminding me what’s really important in the world. To my parents

– thank you for always leading by example in encouraging me to follow my passion. To

Rob – thank you for being the absolutely best dad. Thank you for encouraging me to try

new things, even when it’s challenging, and for always being a calming presence when I

am anxious. To mom – you never got to finish your PhD (in no small part to me and the

other kids) so this degree is for you. Thank you for encouraging me to finish while I had

momentum, even when I thought I couldn’t keep going. You have always been my hero and

I love you.

I wouldn’t have made it through this journey without two more amazing helpers. To our

kitty Boba, thank you for keeping my lap warm during the long hours of working from home

and for always keeping me on my toes. To our puppy Kona, thank you for making me get

outside, making me smile always, for introducing me to the amazing world of dog-owners

on campus, and of course, for being the best girl.

Finally, to Kevin – thank you for being the best partner on this journey. Thank you for

editing my work, listening to hours of practice talks, and for talking through experiment

ideas with me – you have made me a better writer, presenter, and scientist. Thank you

for always being there to remind me that the only way to fail is to not try. Thank you for

keeping me sane, making me laugh, and always helping me see the bigger picture.

Contents

Abstract v

Acknowledgments ix

1 Introduction 1

1.1 Problem statement and motivation . . . . . . . . . . . . . . . . . . . . . . . 1

1.2 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

1.2.1 Overview of synthetic plastics . . . . . . . . . . . . . . . . . . . . . . 2

1.2.2 Recalcitrance of synthetic plastics . . . . . . . . . . . . . . . . . . . 3

1.2.3 Current plastic waste management systems . . . . . . . . . . . . . . 3

1.2.4 Emerging concerns related to plastic pollution . . . . . . . . . . . . 5

1.2.5 Alternative plastic materials . . . . . . . . . . . . . . . . . . . . . . . 5

1.2.6 Plastic biodegradation by microorganisms . . . . . . . . . . . . . . . 7

1.2.7 Plastic biodegradation by insects . . . . . . . . . . . . . . . . . . . . 7

1.3 Dissertation Organization . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

1.4 Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

1.5 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

2 Biodegradation of polyethylene and plastic mixtures in mealworms (larvae

of Tenebrio molitor) and e↵ects on the gut microbiome 15

2.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

2.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17

2.3 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19

2.3.1 Mealworm survival and plastic consumption . . . . . . . . . . . . . . 19

2.3.2 Plastic test materials . . . . . . . . . . . . . . . . . . . . . . . . . . . 20

2.3.3 Characterization of plastic degradation within egested frass . . . . . 20

2.3.4 Microbial community analysis . . . . . . . . . . . . . . . . . . . . . . 21

2.3.5 Statistical analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

2.4 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

2.4.1 PE consumption and e↵ects on survival . . . . . . . . . . . . . . . . 22

2.4.2 Evidence for depolymerization and biodegradation of PE . . . . . . 23

2.4.3 Evidence for mineralization via a mass balance . . . . . . . . . . . . 25

2.4.4 Biodegradation of mixed plastics . . . . . . . . . . . . . . . . . . . . 25

2.4.5 E↵ects of plastic consumption on gut microbiome . . . . . . . . . . . 26

2.5 Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

2.6 Supporting Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

2.7 Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

2.8 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

3 Enhanced polystyrene biodegradation in an ex situ gut microbiome en-

richment from Tenebrio molitor (mealworm larvae) 35

3.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

3.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37

3.3.1 Plastic materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

3.3.2 Mealworm maintenance . . . . . . . . . . . . . . . . . . . . . . . . . 39

3.3.3 Collection of gut microbiome and supernatant . . . . . . . . . . . . . 39

3.3.4 Enrichment and isolation of PS-degrading bacteria . . . . . . . . . . 40

3.3.5 Microbial activity analysis . . . . . . . . . . . . . . . . . . . . . . . . 41

3.3.6 Characterization methods . . . . . . . . . . . . . . . . . . . . . . . . 42

3.4.1 E↵ects of supernatant on ex situ microbial growth . . . . . . . . . . 43

3.4.2 Identification of endogenous emulsification activity . . . . . . . . . . 45

3.4.3 Enhanced ex situ microbial growth . . . . . . . . . . . . . . . . . . . 46

3.4.4 Identification of plastic degradation microbial community . . . . . . 48

3.5 Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49

3.7 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51

4 The fate of hexabromocyclododecane (HBCD), a common flame retar-

dant, in polystyrene-degrading mealworms: elevated HBCD levels in

egested polymer but no bioaccumulation 57

4.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

4.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59

4.3.1 Plastic test materials . . . . . . . . . . . . . . . . . . . . . . . . . . . 61

4.3.2 Mealworm growth conditions . . . . . . . . . . . . . . . . . . . . . . 61

4.3.3 HBCD quantification . . . . . . . . . . . . . . . . . . . . . . . . . . . 62

4.3.4 Aquaculture experimental conditions . . . . . . . . . . . . . . . . . . 63

4.4.1 E↵ects of HBCD on mealworm survival and plastic consumption . . 64

4.4.2 Exponential removal of HBCD from mealworm biomass . . . . . . . 65

4.4.3 Fate of HBCD in mealworm biomass over time . . . . . . . . . . . . 66

4.4.4 Lack of HBCD bioaccumulation in secondary trophic level . . . . . . 68

4.5 Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70

4.7 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72

5 The role of biotechnology for control and remediation of plastic pollution 77

5.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

5.2 Highlights . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 78

5.3 Current plastic production rates are not sustainable . . . . . . . . . . . . . 79

5.4 From tiny bioreactors to new mitigation strategies . . . . . . . . . . . . . . 80

5.5 From linear to fully circular economies . . . . . . . . . . . . . . . . . . . . . 80

5.6 Low-cost, tailored to the application, and Ecocyclable . . . . . . . . . . . . 81

5.7 Design for disassembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84

5.8 Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84

5.10 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

6 Conclusions and Future Work 91

6.1 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

6.2 Recommendations for future research . . . . . . . . . . . . . . . . . . . . . . 93

A Supporting Information for Chapter 2 97

A.1 Material and Methods Supplement . . . . . . . . . . . . . . . . . . . . . . . 98

A.1.1 Extraction method e�ciency assay . . . . . . . . . . . . . . . . . . . 98

A.1.2 Phasing amplicon sequencing library preparation . . . . . . . . . . . 98

A.1.3 E. coli K12 plastic characterization controls . . . . . . . . . . . . . . 99

A.2 Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

A.3 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

B Supporting Information for Chapter 3 115

B.1 Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116

B.2 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117

C Supporting Information for Chapter 4 121

C.1 Material and Methods Supplement . . . . . . . . . . . . . . . . . . . . . . . 122

C.1.1 Chemicals and reagents . . . . . . . . . . . . . . . . . . . . . . . . . 122

C.1.2 HBCD analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122

C.1.3 Litopenaus vannamei experimental conditions . . . . . . . . . . . . . 124

C.1.4 Mealworm depuration experimental conditions . . . . . . . . . . . . 125

C.2 Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127

C.3 Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128

D Supporting Information for Chapter 5 135

D.1 Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136

Bibliography 137

List of Tables

1.1 Five most common thermoplastics . . . . . . . . . . . . . . . . . . . . . . . 11

2.1 Characterization of PE and PS foams . . . . . . . . . . . . . . . . . . . . . 29

A.1 Overview of Illumina MiSeq data . . . . . . . . . . . . . . . . . . . . . . . . 101

A.2 1H-NMR peak information (PE samples) . . . . . . . . . . . . . . . . . . . . 102

A.3 1H-NMR peak information (PS samples) . . . . . . . . . . . . . . . . . . . . 103

A.4 FTIR peak information, PE samples . . . . . . . . . . . . . . . . . . . . . . 104

A.5 FTIR peak information, PS samples . . . . . . . . . . . . . . . . . . . . . . 105

A.6 FTIR peak information, plastic mixture samples . . . . . . . . . . . . . . . 106

A.7 Relative abundance of PE and PS associated OTUs . . . . . . . . . . . . . 107

B.1 Sequenced bacterial strains from microbial enrichment and isolates . . . . . 116

C.1 Characterization of the PS foams . . . . . . . . . . . . . . . . . . . . . . . . 127

D.1 Cost, material properties, and suitability for various manufacturing processes

for common plastics and plastic replacement materials . . . . . . . . . . . . 136

List of Figures

1.1 Characteristics within insect larvae that may accelerate biodegradation . . . 13

1.2 Characteristics within mealworms that may accelerate biodegradation . . . 14

2.1 Chapter 2 summary figure . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

2.2 Survival rate and plastic consumption by T. molitor by diet . . . . . . . . . 31

2.3 Characterization of polyethylene degradation within the mealworm gut . . . 32

2.4 Microbial community analysis of gut microbiome in di↵erent diets . . . . . 33

2.5 Di↵erential abundance analysis of gut microorganisms between experimental

diets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34

3.2 E↵ects of mealworm gut supernatant on ex situ gut microbiome growth on PS 52

3.3 Hydrophobicity of mealworm gut supernatant . . . . . . . . . . . . . . . . . 53

3.4 E↵ect of PS supernatant on microbial respiration . . . . . . . . . . . . . . . 54

3.5 Bacterial strain identifies from microbial enrichment and isolates . . . . . . 55

3.6 Bacterial strain identifies from microbial enrichment and isolates . . . . . . 56

4.2 Survival rate and HBCD consumption by T. molitor . . . . . . . . . . . . . 73

4.3 Reduction in T. molitor whole-body burden of HBCD with starvation . . . 74

4.4 Fate tracking of HBCD in T. molitor . . . . . . . . . . . . . . . . . . . . . . 75

4.5 Bioaccumulation e↵ects from plastic-fed mealworm biomass on L. vannamei 76

5.2 Projected increase in plastic production based on historic production data . 87

5.3 Current linear pathways for recalcitrant plastics and circular pathways of

future plastic materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88

5.4 A transition sequence for plastic materials . . . . . . . . . . . . . . . . . . . 89

A.1 Characterization of PS degradation within the mealworm gut . . . . . . . . 109

A.2 Characterization of plastic treated with E. coli K12 . . . . . . . . . . . . . 110

A.3 Mass balance on mealworms during the 32-day experiment . . . . . . . . . . 111

A.4 Characterization of mixed plastic (PE + PS) degradation . . . . . . . . . . 112

A.5 Principal Coordinate Analysis of microbial communities with outliers . . . . 112

A.6 E�ciency of extraction method used to recover residual polymer in the frass 113

B.1 Dose-response curves for supernatant and glucose equivalent . . . . . . . . . 118

B.2 Supernatant emulsification activity . . . . . . . . . . . . . . . . . . . . . . . 119

B.3 Enrichment biofilm formation . . . . . . . . . . . . . . . . . . . . . . . . . . 120

C.1 Schematic diagram of the mealworm experimental conditions . . . . . . . . 129

C.2 Schematic diagram of the L. vannamei experimental conditions . . . . . . . 129

C.3 Schematic diagram of sample preparation for HBCD analysis . . . . . . . . 130

C.4 Multiple regression analysis of predicted versus actual number of pupae . . 131

C.5 Fate tracking of HBCD in T. molitor in co-fed diets . . . . . . . . . . . . . 132

C.6 Recovery e�ciency of surrogate standard across sample types . . . . . . . . 132

C.7 Mass balance on HBCD in the mealworm experiments . . . . . . . . . . . . 133

C.8 Standard curves for the main stereoisomers of HBCD . . . . . . . . . . . . . 133

C.9 Log removal of HBCD from T. molitor over time . . . . . . . . . . . . . . . 134

Chapter 1

Introduction

1.1 Problem statement and motivation

Plastics are a growing concern for both the environment and waste management systems.

Plastic production has grown exponentially since the invention of synthetic polymers and

is now approximately 400 million metric tons per year (Fig. 5.2). As understanding of

polymer science has improved, polymers have been designed and modified to better suit

the needs of di↵erent industries.[139, 53] While some of these products have highly valuable

functions (such as sterile disposable tools in hospitals), much of the increase in plastic pro-

duction involves packaging and other products that lend themselves to wasteful, single-use

applications.[139, 3, 184] The properties that have led to the rampant increase in production

of plastics – versatility, durability, and low cost – also render these materials di�cult to

dispose of sustainably. These same features that make plastics desirable commercially (i.e.,

strength, durability, and inert nature) can contribute to their recalcitrance and ultimately

their accumulation within natural environments.

The incredible increase in plastic production has led to massive waste management

challenges. Globally, much of the waste that results from use of these products is misman-

aged, resulting in plastic waste accumulating in diverse natural environments, especially

the ocean. The most robust analysis of plastic inputs into the environment reported that

⇠ 32% of plastic waste is mismanaged and available to escape into the environment, with

an estimated 4.8 – 12.7 million metric tons of plastic entering the ocean annually.[79] Even

properly managed waste is subject to increasing challenges as landfills, the most common

2 CHAPTER 1. INTRODUCTION

end-of-life fate for plastics, are space-constrained and approaching capacity is some ur-

ban environments.[42] Thus, developing strategies for remediation and removal of discarded

plastic is increasingly critical. Biodegradation is one possible solution as it would break

down plastic waste and potentially enable the recovery of useful resources. However, cur-

rent rates of biodegradation are slow and often plastic-specific, limiting the use of microbial

biodegradation as a scalable solution for plastic waste. Identification of natural systems that

can accelerate plastic biodegradation is a critical first step to understanding of degradation

mechanisms and scale up of biodegradation as a sustainable end-of-life fate for plastic waste.

1.2 Background

1.2.1 Overview of synthetic plastics

Plastics are polymers – large macromolecules made up of smaller repeating units of smaller

molecules known as monomers. Conventional plastics are novel in that they are entirely syn-

thetic, enabling their physical and chemical properties to be tailored for di↵erent purposes.

Synthesis of these plastics (polymerization) typically yields a mixture of large polymers

with di↵erent molecular weights.[163] Controlling polymer chain length (molecular weight,

MW) and the distribution of sizes (polydispersity) can dramatically change the physical

and mechanical properties of the produced material.[163]

Plastics can be broadly classified as either thermoplastics or thermoset plastics. Ther-

moplastics soften upon heating and can therefore be formed into di↵erent shapes through

the application of heat and pressure. This property enables reuse (or recycling) of ther-

moplastics collected at end-of-life by melting and reprocessing.[3, 163] Thermoset plastics

are soft or even liquid polymers that upon heating become cross-linked, which solidifies the

polymer and prevents softening upon further exposure to heat.[3, 163] The most common

thermosets are foams like polyurethane or polyisocyanurate. Plastics can also be classi-

fied as either hydrolyzable, polymers that contain more readily degradable ester bonds, or

non-hydrolyzable plastics that are more resistant to enzymatic degradation.[78] The work

in this dissertation focuses on two of the most common thermoplastics: polyethylene (PE)

and polystyrene (PS), both of which are non-hydrolyzable.

1.2. BACKGROUND 3

1.2.2 Recalcitrance of synthetic plastics

The physical and molecular characteristics of plastics confer recalcitrance to abiotic and

biotic degradation.[149, 158] Abiotic processes include weathering, photooxidation by UV

light, and fragmentation due to mechanical stress.[158, 65] Biotic processes (i.e., biodegra-

dation) entail polymer degradation mediated by microorganisms – typically bacteria, fungi,

or algae.[149, 158] Microbial biodegradation can result in complete mineralization to CO2

and H2O and/or biomass production and partial degradation (partial depolymerized and

modified polymers).[158] Abiotic and biotic processes can operate in parallel in the envi-

ronment. The plastics listed in Table 1.1, along with most other synthetic plastics, are

all considered recalcitrant because they resist degradation in natural environments over

meaningful timescales.

Polymerization and manufacturing of many plastic products has been optimized to yield

stronger and more durable polymers. The addition of co-polymers, plasticizers, and other

additives have resulted in plastics that are more resistant to shattering, fragmentation, and

photo-oxidation, and thus less susceptible to abiotic degradation processes.[3]

Biotic degradation is likewise limited because plastic polymers are often large, bulky

high molecular weight molecules that cannot pass through cell membranes.[149] They must

first be depolymerized into lower molecular weight molecules before further biodegradation

and mineralization is possible. The inert chemical structure of plastics also limits biotic

degradation.[149] The most common bonds are C-C and C-H bonds, and the functional

moieties that enable biodegradation (e.g., alcohols, aldehydes, acids) are often lacking.[163]

The absence of such groups, contributes to the hydrophobic nature of most polymers, which

can also impedes microbial colonization.[163, 149, 65]

Synthetic polymers are relatively new in the environment, having only been discovered

a little over a century ago.[3, 1] By contrast, the mineralization of naturally-occurring

biopolymers (e.g., polyhydroxyalkanoates) has evolved over billions of years and thus occurs

rapidly in many environments.[143, 117] The work in this dissertation focuses on biotic

degradation (or biodegradation).

1.2.3 Current plastic waste management systems

A recent e↵ort to catalog the fate of all synthetic plastics ever made identified three major

end-of-life fates for plastics: discarded (landfilled or in the environment, 60%), incinerated

(12%), and recycled (9%).[66] The remaining fraction of plastics remain in use.

1. Landfilling. The vast majority of plastics (75.8% of properly collected waste in the

US[53]) end up in a landfill. If managed well, this can be a desirable end point as the

carbon is sequestered; however, space is finite, especially in urban areas where most

waste is generated.[42]

2. Incineration. The recovery of energy from waste plastic is typically achieved by com-

bustion, sometimes referred to as “quaternary or energy-recovery recycling.” This

process can be ine�cient, contribute to greenhouse gas emissions, and potentially

volatilize chemical additives; however, such emissions of additives or their derivatives

may be contained with proper engineering controls.[59]

3. Recycling. Plastics can be collected and recycled, but rates of plastic recycling are

the lowest of any material.[184] Recycling can take the following forms:[59]

(a) Primary/mechanical recycling: Production of new products with similar (pri-

mary) or di↵erent properties (secondary) to the original waste product. The

heating and re-melting of polymers can result in ‘downcycling,’ where the recy-

cled polymer has inferior qualities (MW, strength, durability) compared to virgin

material.

(b) Monomer or feedstock recycling (tertiary recycling): Production of chemical fuels

and feedstocks from waste plastic. This process works well for specific polymers

(e.g., polyethylene terephthalate and polypropylene) but remain limited in their

While recycling has been touted as a potential solution, in practice, recycling plastics

remains challenging. Proper collection, without contamination, remains a constant chal-

lenge. Once collected, plastics are di�cult to recycle given the variety of forms, additives,

and blends used in an increasing number of products.[59, 20] Further, the low cost of virgin

materials has led to economic disincentives that have prevented the revitalization of the re-

cycling system. China became the chief market for plastic recycling in the 1990s, importing

nearly 45% of all plastic waste generated since 1992.[20] Thus the 2017 decision by China

to ban the import of nonindustrial plastic waste further endangers global plastic recycling

systems.[20]

1.2. BACKGROUND 5

Current waste management systems are based on a linear economy in which fossil carbon

is transformed into recalcitrant plastics that accumulate in landfills or the environment after

use. In such a scheme, failure to capture plastic waste at the end-of-life inevitably leads to

rampant environmental problems. To address this issue a circular economy is needed, with

larger recycling loops that can sustainably break down plastic waste and recover value.

1.2.4 Emerging concerns related to plastic pollution

Recent research has highlighted the complex relationship between plastic debris and poten-

tially toxic chemicals, both from the plastic production process and from other sources.[52]

The hydrophobic nature of synthetic plastics allows plastic debris to sorb hydrophobic chem-

icals such as polychlorinated biphenyls (PCBs) from the environment.[9] Additionally, many

synthetic plastics are embedded with chemical additives such as plasticizers, stabilizers, and

flame retardants, many of which are hydrophobic and are of growing concern for human

health.[3, 7] For example, expanded and extruded polystyrene are embedded with the flame

retardants hexabromocyclododecane (HBCD),[11, 157] a known endocrine disruptor and

potential neurotoxin.[36, 80, 215, 129, 156]

New research has claimed that plastics can act as a vector for transporting these chemi-

cals into the food chain when ingested.[52, 9, 50, 114] This gives rise to many concerns about

the potential for bioaccumulation within the food web and within commercially important

species.[52, 9, 35]

Current systems for plastic production and recycling have yet to fully address the fate of

chemical additives, such as fire retardants and plasticizers. Strategies are needed to ensure

their removal from recycled materials and to develop green chemistry replacements.[7, 12]

Further, research is needed to understand the fate of contaminants within plastics when

they enter di↵erent environments, especially when ingested by diverse animals, including

humans, and enter the gut environment.[75, 172]

1.2.5 Alternative plastic materials

Sustainable replacement materials for petroleum-based plastics are needed to end the ram-

pant production and accumulation of plastic waste.[45, 170] Renewable plastic replacements

will need to match the in-use functionality of current plastics without leading to accumu-

lation, thus safeguarding the environment. To meet current plastic demands, a roughly

500-fold increase in renewable plastics production is required.[56] Production costs need to

be decreased by a factor of 2 – 4 to achieve cost parity with recalcitrant petroleum-based

plastics to enable the widespread use of renewable plastics (Table D.1).

Polylactic acid (PLA) is the most common renewable plastic currently in use, account-

ing for 10.3% of global polymer production in 2017.[56, 49] Most PLA is made from lactic

acid obtained through the fermentation of cultivated crops.[85] PLA readily degrades in

warm, aerobic environments (e.g., thermophilic composts) but in cool, anaerobic environ-

ments (e.g., soils and the marine environment) it is as recalcitrant as traditional petroleum-

based plastics.[86, 117] Thus, while PLA is an improvement over conventional plastics, it

is not an ideal replacement material given the lack of sustainability of using cultivated

crops and its failure to biodegrade in critical environments. These challenges can be ad-

dressed through advancements in polymer chemistry and biotechnology. Copolymerization

of PLA and the development of PLA composites have been demonstrated to improve ma-

terial properties and degradability.[49] Researchers are also investigating the use of micro-

bial carboxylesterases to rapidly depolymerize PLA, potentially enabling enzymatic control

strategies in the future.[70]

Another class of renewable polymers are naturally occurring biopolymers, polyhydrox-

yalkanoate (PHAs). PHAs are a vast family of moldable biopolymers that could serve as

“drop-in” replacements for traditional plastics in production and their in-use functionality.[217]

However, critical bioprocess challenges for PHA production such as feedstock and produc-

tion costs, the need to tailor polymer structure for specific applications, and polymer ex-

traction and purification remain, limiting the wide-spread use of these polymers. The use

of waste-derived feedstocks can reduce the cost of PHAs while creating a closed-loop cycle,

increasing the sustainability of these materials.[154] Research has demonstrated the ability

to tailor the polymer structure through the choice of co-substrates, application of selection

pressures, and genetic manipulation of biochemical pathways.[154, 123, 29] Biotechnology

has also been applied to overcome the challenges of conventional polymer extraction meth-

ods which are time, energy, and chemical-intensive.[154, 110] Recent advances such as the

development genetic modifications that enable direct secretion of PHA molecules and the

use of osmolysis to release PHA granules will further enable the scaling of PHA production

without the challenges of conventional extraction.[146, 54, 142]

Biotechnology has opened the door for improves in bioplastic design in terms of its

production, in-use properties, and end-of-life fate. Together these advances will be necessary

1.2. BACKGROUND 7

to transition away from the linear lifecycle of petroleum-based plastics to a sustainable,

circular lifecycle of renewable plastics.

1.2.6 Plastic biodegradation by microorganisms

Microbial enrichment and isolation studies have demonstrated that several bacterial isolates

are capable of degrading plastic wastes, as reviewed elsewhere.[65, 158, 93, 200, 124] Rates

of degradation vary but are typically slow, ranging from ⇠ 8% in two weeks to ⇠ 5% over six

months; however, the concentration of microorganisms, a critical variable in these systems,

is often not reported.[65, 158] Most of the identified microorganisms have been isolated

from environments contaminated with plastic waste such as landfills, waste water sludge,

or soils from disposal or recycling sites.[158, 65, 93, 214] To date, there has been little to

no research into endogenous environmental degradation of plastic, even in environments

heavily contaminated with waste.

Most studies of plastic biodegradation do not clearly identify the mechanism of de-

polymerization. Secreted enzymes such as oxygenases (monooxygenase and dioxygenase),

hydrolases, esterases, laccases, and peroxidases are hypothesized candidates for microbial-

mediated degradation of synthetic polymers.[93, 198, 78] However, without a firm un-

derstanding of the mechanism of depolymerization or environments that enable enhanced

degradation, it is di�cult to replicate and stimulate biodegradation at high rates and at

large scales. Given that individual bacterial strains have typically exhibited low plastic

biodegradation rates, future studies should evaluate whether microbial consortia can accel-

erate biodegradation and mineralization.

1.2.7 Plastic biodegradation by insects

Recent studies have demonstrated one environment in which plastics undergo rapid biodegra-

dation: the gut of insect larvae, specifically Tenebrio molitor (yellow mealworms),[212,

213, 210, 207] Tenebrio obscurus (dark mealworms),[134] Plodia interpunctella (Indianmeal

moth),[206] and Galleria mellonella (wax moths).[91, 147, 135, 105, 25] Of these, the most

well studied is mealworms, where researchers have established that within the 15 – 20 hour

gut retention time nearly 50% of ingested plastic is mineralized to CO2.[212, 210, 207]

There are several features common to these insect larvae that could contribute to their

unique ability to rapidly degrade plastic (Fig. 1.1). All these insects are omnivorous

and have evolved to utilize a variety of substrates, a trait that is thought to contribute

to their ability to opportunistically degrade plastic.[91, 51, 204] Further, the chewing and

fragmentation of plastic into smaller pieces via ingestion increases the surface area to volume

ratio, increasing the area exposed for enzymatic attack (Fig. 1.1). These insects are also

su�ciently small such that oxygen can passively di↵use throughout (or through most) of

the gut, allowing for oxidative attacks on the plastics (Fig. 1.1). A deeper understanding

of the degradation environment and mechanism, including the relative contributions of the

insect itself and the gut microbiome, could enable development of scalable enzyme-based

strategies for plastic waste management.

Overview of mealworms

Yellow mealworms (larvae of Tenebrio molitor) are the larval stage of a darkling beetle. T.

molitor has four life cycle stages: egg (4-18 days), larva (6-9 months), pupa (6-18 days),

and beetle (2-3 months).[212] Mealworms are omnivorous, and researchers hypothesize that

their gut bacteria play an important role in their ability to adapt to di↵erent foods.[194]

Originally from North America, the species is now found and reared by farmers throughout

the world as a protein source for agriculture and aquaculture.[10, 64, 165, 136] Recent

life cycle assessment studies have shown that mealworm protein has low greenhouse gas

emissions, energy requirements, and land requirements per gram of edible protein, which

has prompted researchers to consider mealworms as a potential protein source for humans

in the near future.[128]

The mealworm gut structure, pH, and microbiome have previously been studied by

entomologists.[194, 190, 169, 187] The gut structure is traditionally broken down into four

sections, which have been found to harbor di↵erent quantities of microorganisms in the

range of 105 to 106 colony forming units (CFU) per gut.[213, 194] Under normal conditions,

the anterior gut is dominated by facultative anaerobes of the genus Lactococcus, genus

Pantoea, and the family Bacillaceae while the posterior gut is more diverse and features

more anaerobic bacteria from the genera Spiroplasma, Clostridium, and Enterobacter (Fig.

1.2).[194] Another defining characteristic of the mealworm gut is a sharp pH gradient that

ranges from ⇠ 5.6 to ⇠ 7.9 moving from the anterior to the posterior, partially driven by

the loss of CO2 in the posterior spiracles (respiratory pores) of the animal as part of their

respiratory circuit (Fig. 1.2).[190, 169] Di↵erences in secreted enzyme activity within the

gut vary along this pH gradient.[190, 169]

1.3. DISSERTATION ORGANIZATION 9

While the mechanisms of plastic degradation within these insects are unknown, re-

searchers hypothesize that their diet of naturally occurring polymers (e.g., ligand, chitin,

etc.) play a role in the evolution of their unique degradation ability. At present, we lack a

comprehensive understanding of the mechanisms and specific conditions that lead to the ob-

served high rates of biodegradation within the mealworm gut, limiting our ability to utilize

this capacity to develop large-scale solutions for plastic waste. Understanding the environ-

ment in which plastic already degrades is important when considering how biotechnology

might be used to create more e�cient plastic degrading systems.

1.3 Dissertation Organization

This dissertation is comprised of six chapters. Chapter 1 (this chapter) provides an intro-

duction, including background and motivation for the work and the current understanding of

plastic biodegradation. Chapters 2 through 4 present three di↵erent experimental studies

conducted as a part of this dissertation. Each chapter contains an abstract, introduction,

methods, results, and discussion. Chapter 5 presents a critical review on the state of

biotechnology for the remediation and prevention of plastic waste. Tables and figures from

the manuscripts are embedded within the chapters and supplementary information is pre-

sented as appendices. Chapter 2 was published in May 2018, Chapter 4 was published

in January 2020, and Chapter 5 was published in June 2019. Chapter 3 is in preparation

for publication.

Chapter 2 presents an experimental study that assessed the ability of mealworms to

degrade polyethylene and a mixture of plastic wastes (PE and PS). The motivation of this

study was to expand our understanding of the biodegradation ability of the mealworm by

assessing its ability to degrade chemically dissimilar plastics. Results of this study suggest

a non-specific degradation mechanism within the mealworm gut.

Chapter 3 demonstrates ex situ polystyrene degradation by the gut microbiome of

mealworms. This study investigated the role of the mealworm host and the gut microbiome

in plastic biodegradation and utilized these insights to cultivate microorganisms capable of

enhanced polystyrene degradation. The system developed in this chapter will enable fu-

ture studies and bioengineering on the isolated microorganisms and will ultimately support

e↵orts to scale-up plastic degradation.

Chapter 4 investigated the fate of one common PS additive, HBCD, within plastic-

degrading mealworms using a mass-balance study coupled with body burden analysis using

analytical chemistry. HBCD was tracked as a representative plastic additive to assess poten-

tial bioaccumulation e↵ects within the mealworm. Bioaccumulation was further assessed

when plastic-degrading mealworm biomass was incorporated as an aquaculture feed for

shrimp (L. vannamei). We find that HBCD is rapidly excreted from the mealworm in its

frass and does not bioaccumulate withinin mealworm biomass.

Chapter 5 presents a brief critical review on the state of plastic pollution and biotech-

nology remediation strategies. This review includes the state of bioplastic degradation

and presents recommendations for future work. In addition, this review presents a frame-

work for transitioning from recalcitrant petroleum-based plastics to biodegradable, non-

bioaccumulative, and non-toxic plastic replacement materials.

Chapter 6 summarize the overall conclusions and contributions of this dissertation

research and provides suggestions for future research.

1.4. TABLES 11

1.4 Tables

Table 1.1: Five most common thermoplastics ranked by annual U.S. production (highest tolowest): showing chemical structure, applications, and methods of polymerization.[3, 184]

1.5 Figures

1.5. FIGURES 13

Gut wall

Ingestedplastic

Microbial activity

FrassDegradedresiduals

small size - concentrates

important reactants

oxygen permeabilitypH range

diverse & adaptablegut microbiome

pre-fragmentation -increased

surface area-to-volume ratio

recalcitrant plasticdegraded residuals

incorporation into biomass

observed degradation:

system characteristics influencing degradation rates:

15 - 20 hours

Figure 1.1: Characteristics within the insect larvae (drawn here as a mealworm) that mayenable the accelerated biodegradation of recalcitrant plastics.

Figure 1.2: T. molitor gut pH and microbial community structure (based on 16S rRNAsequencing) in the di↵erent regions of the gut constructed using data and images fromprevious studies on mealworms fed their natural diet (bran & vegetables). Image andrelative abundance data from Wang et al. 2015, annotated with additional data fromvarious sources.[194, 169, 190, 144]

Chapter 2

Biodegradation of polyethylene

and plastic mixtures in mealworms

(larvae of Tenebrio molitor) and

e↵ects on the gut microbiome

The results of this chapter were originally published in Environmental Science & Technology

in 2018, doi: 10.1021/acs.est.8b02301, reprinted here with permission, copywrite American

Chemical Society 2018. Drs. Shu-hong Gao, Renmao Tian, and Daliang Ning preformed

sequencing analysis and contributed to analysis of results under the supervision of Dr.

Jizhong Zhou. Dr. Shan-Shan Yang contributed to the design of the study. Dr. Wei-

Min Wu contributed to the design of the study and provided comments to improve the

manuscript.

16 CHAPTER 2. BIODEGRADATION OF POLYETHYLENE

2.1 Abstract

Recent studies have demonstrated the ability for polystyrene (PS) degradation within the

gut of mealworms (Tenebrio molitor). To determine whether plastics may be broadly sus-

ceptible to biodegradation within mealworms, we evaluated the fate of polyethylene (PE)

and mixtures (PE+PS). We find that PE biodegrades at comparable rates to PS. Mass

balances indicate conversion of up 49.0 ± 1.4% of the ingested PE into a putative gas frac-

tion (CO2). The molecular weights (Mn) of egested polymer residues decreased by 40.1 ±8.5% in PE-fed mealworms and by 12.8 ± 3.1% in PS-fed mealworms. NMR and FTIR

analyses revealed chemical modifications consistent with degradation and partial oxidation

of the polymer. Mixtures likewise degraded. Our results are consistent with a non-specific

degradation mechanism. Analysis of the gut microbiome by next-generation sequencing

revealed two OTUs (Citrobacter sp. and Kosakonia sp.) strongly associated with both PE

and PS as well as OTUs unique to each plastic. Our results suggest that adaptability of

the mealworm gut microbiome enables degradation of chemically dissimilar plastics.

Figure 2.1: Summary Figure.

2.2. INTRODUCTION 17

2.2 Introduction

Plastics are a growing concern for both the environment and waste management systems.

Global plastic production has tripled in the last 25 years to over 322 million tons in

2015.[139] This nearly exponential growth in production has contributed to waste man-

agement challenges including space limitations in landfills, which coupled with low re-

cycling rates has led to mismanagement of plastic waste and increased environmental

pollution.[139, 42, 184] Plastic is of especially great concern in marine environments where

its recalcitrance has led to accumulation and harmful e↵ects on wildlife and potentially

humans.[79, 172] To combat this growing problem, there has been a steady increase in

research on plastic biodegradation by bacteria and fungi.[158, 173, 93] Microbial enrich-

ment and isolation studies have demonstrated that several bacterial isolates are capable of

degrading plastics, but rates of degradation vary and are typically low.[158, 173, 93]

Recent work has demonstrated that mealworms (larvae of Tenebrio molitor), obtained

from various sources across the globe, readily ingest and biodegrade polystyrene (PS) to

CO2 and lower molecular weight compounds within their gut.[212, 213, 207] Antibiotic

studies implicated gut bacteria as agents of PS degradation, and Exugiobacterium sp. YT2,

a bacterium capable of PS degradation was isolated from the gut.[213] Mealworms were

shown to degrade nearly half of the ingested PS within the 12 – 15 hour retention time in

the gut, which is higher than the mass loss reported by the isolated Exugiobacterium sp.

YT2 and other plastic-degrading bacterial isolates.[212, 213, 69] A recent report found that

co-feeding PS with bran almost doubled the rate of PS degradation.[207]

Mealworms are omnivorous and researchers hypothesize that their gut bacteria play

an important role in their ability to adapt to di↵erent foods.[194] Bacterial concentrations

range from 105 to 106 colony forming units per gut.[213, 194] On a standard diet (e.g.,

bran), the anterior gut is dominated by facultative anaerobes of the genera Lactococcus and

Pantoea and by genera within the family Bacillaceae, while the posterior gut is more diverse,

featuring anaerobes from the genera Spiroplasma, Clostridium, and Enterobacter.[194]

To determine whether plastics may be broadly susceptible to biodegradation within

mealworms, we evaluated the fate of polyethylene (PE) within the gut. Assessing the

degradation of PE is of interest because PE has a markedly di↵erent chemical structure

than PS; PE lacks a benzene ring in the repeating monomer unit, which could impact

the resulting degradation. The chemical structure of polyethylene represents the simplest

homogenous carbon-to-carbon backbone.[3] Thus, the ability to degrade PE would suggest

that the mealworm gut microbiome is capable of degrading other plastics with a similar

chemical motif (e.g., polyvinyl chloride or polypropylene). In addition, the global production

of polyethylene (PE) is approximately four times that of PS, and PE plastics are among

the most common plastic pollutants.[3, 171]

There have been recent reports of PE degradation within the gut of other omnivorous

insect larvae (Indian meal moths, wax moths), which could suggest that the insect gut

broadly enables the degradation of recalcitrant plastics; however, evidence for degradation

is preliminary and warrants further study and confirmation.[206, 14] None these studies

reported the ability to degrade more than one type of plastic or plastic mixtures. Addition-

ally, few previously identified plastic-degrading systems, including isolated microorganisms,

are capable of degrading multiple types of plastic, suggesting that the plastic degradation

is typically plastic-specific.[158, 173, 93]

In this study, we investigate the biodegradation of polyethylene (PE) and mixed plastics

(PE and PS) in a previously untested strain of T. molitor. We find that PE degrades at

rates comparable to those of PS-fed larvae.[212, 207] Moreover, mixed plastics (PE and PS)

are consumed and degraded, suggesting that degradation is non-specific. Next-generation

sequencing indicates that this degradation is associated with changes in the gut microbiome.

2.3. MATERIALS AND METHODS 19

2.3 Materials and Methods

2.3.1 Mealworm survival and plastic consumption

Mealworms, larvae of T. molitor Linnaeus, (average weight 75-85 mg/worm) were purchased

online from Rainbow Mealworms (Compton, CA) and shipped overnight to the laboratories

at Stanford University. Prior to arrival, the mealworms were fed bran; after arrival, they

were subject to a 48-hour starvation period before initiating experimental diets. Natural

wheat bran was purchased from Exotic Nutrition (Newport News, VA).

Six experimental diets were compared: PE, PE + bran (1:1 [w/w]), PS, PS + bran

(1:1 [w/w]), PE + PS (1:1 [w/w]), and bran (control diet). To assess mealworm survival

rate and plastic mass loss, 120 randomly selected mealworms were placed in a food grade

polypropylene container (volume 475 mL) along with 1.80 g of plastic cut into 2 – 3 cm

cubes (PE, PS, or 0.90 g PE and 0.90 g PS). Bran-fed containers (PE + bran, PS + bran)

initially received 1.80 g bran plus plastic spread throughout the container. Additional bran

was added every 3 days to maintain a 1:1 ratio [w/w] of plastic to bran within each container.

Bran-fed controls initially received 1.80 g of bran and 1.80 g of additional bran every 3 days.

All tests were carried out in duplicate. Containers were stored in incubators maintained at

25 �C and 70% humidity.[212, 207]

Mealworm survival was evaluated approximately every 3 days for 32-days by counting

dead mealworms, which were then removed. Once a week, the mealworms were cleaned

with a stream of air to remove any residual plastic fragments and transferred to a clean

container to collect frass (excrement) for analysis. After 24 hours, the mealworms were

returned to their original container, and frass samples were weighed and stored at -20 �C.

To obtain su�cient frass for characterization, 1000 mealworms (from the same order)

were raised in larger “bulk-fed” food grade polypropylene containers (volume 780 mL) on

the six diets described above. These containers followed the same bran-supplementation

and frass-collection schedule as described above. Bulk-fed containers were also stored in

incubators maintained at 25 �C and 70% humidity. Frass samples from the end of the

32-day experiment are used in the analysis below.

2.3.2 Plastic test materials

To assess the degradation of commercially available plastic products, low-density polyethy-

lene foam, 1.3 cm thickness, was purchased online from the Foam Factory (Macomb, MI).

The polystyrene foam, 5.1 cm thickness, used was an expanded polystyrene insulation ma-

terial from Carlisle Construction Materials (Puyallup, WA). Plastic foam blocks were cut

into irregular 2 – 3 cm cubes and cleaned with a stream of air to remove any residues prior

to being placed in the appropriate container. The molecular weight and density of the con-

trol materials are shown in Table 2.1. The PE foam contained no chemical additives. The

PS foam contained less than 1% 1,2,5,6,9,10-hexabromocyclododecane (a common flame

retardant in PS materials).

2.3.3 Characterization of plastic degradation within egested frass

To characterize depolymerized polymer in the frass, the molecular weight (number-averaged

[Mn] and weight-averaged [Mw] molecular weight) of the polymer was quantified by high-

temperature gel permeation chromatography (HT-GPC) using previously established

methods.[207] Frass samples (50 mg) were gently crushed in a mortar and pestle prior

to a 2-hour extraction in 2 mL solvent aliquots with gentle heating (placed on a hot plate

on the lowest setting). For PE samples, the solvent was dichloromethane (� 99.9%, Thermo

Fisher Scientific Inc., Pittsburg, PA); for PS samples, the solvent was tetrahydrofuran (�99.9%, Thermo Fisher Scientific Inc., Pittsburg, PA) (SI Method A.1.1, Fig. A.6). After

2 hours, the solution was filtered using a 0.22 µm PVDF filter (Thermo Fisher Scientific

Inc., Pittsburg, PA) and transferred into a clean glass vial. The residual polymer in the fil-

tered solution was concentrated by rotary evaporation, and the residue (“residual polymer”)

was weighed to determine the extractable fraction (i.e., the fraction of the frass weight re-

covered). Residual polymer was dissolved in 1,2,4-trichlorobenzene (� 99%, Alfa Aesar,

Haverhill, MA) to obtain a final concentration of approximately 5 mg/mL. Triplicate anal-

yses for each sample were run at 180 �C with a 100 µL injection volume with an eluent

(1,2,4-trichlorobenzene) flow rate of 1.0 mL/min (EcoSEC High Temperature GPC System,

Tosoh Biosciences).

Proton nuclear magnetic resonance (1H-NMR) analysis was conducted to characterize

degradation in the egested frass. The control plastics and residual polymers from mealworms

fed each diet were analyzed. Trace residue extracted from the frass of bran-fed mealworms

was used as a control. Before conducting the liquid-state 1H-NMR analysis, frass samples

(50 mg) were extracted as described above. The residual polymer was re-suspended in

chloroform-D (� 99.8%, Cambridge Isotope Laboratories, Inc., Tewksbury, MA). Proton-

NMR spectra were obtained at 55 �C on a Varian Inova 500-MHz NMR spectrometer

(Agilent Technologies, Inc., Santa Clara, CA). The 1H-spectra [32 scans, delay time (d1)

= 0.0 s] were referenced to the residual deuterated-chloroform peak [7.26 ppm]. Spectra

were analyzed using MestReNova software (version 10.0.2), values are reported in parts per

million (ppm).

Additional characterization of the residual polymer was obtained using Fourier Trans-

form Infrared Spectroscopy (FTIR) on a Nicolet iS50 FTIR Spectrometer (Thermo Fisher

Scientific, Inc., Pittsburg, PA). Spectra were recorded from the residual polymers extracted

from the frass ground to a homogenous powder in absorbance mode and transformed into

transmittance for graphing. Spectra were recorded in the range of 4000-500 cm-1 with a

minimum of 16 scans with a spectral resolution of 0.482 cm-1. Peaks were identified using

OMNIC software (Thermo Fisher Scientific Inc., Pittsburg, PA). Proton-NMR and FTIR

analysis were run in duplicate for each diet on residual polymer collected from the bulk-fed

containers.

2.3.4 Microbial community analysis

At the end of the 32-day experiment, the gut content of each sample (four mealworms from

the same container pooled to eliminate individual variability) was harvested and washed

four times by vortexing the guts with 100 µL of DNA extraction bu↵er (0.1 M NaH2PO4, 0.1

M Na2HPO4, 0.1 M EDTA, 0.1 M Tris-HCl, 1.5 M NaCl, 1% CTAB).[219] Gut walls were

removed, and DNA was extracted using the MoBio PowerLyzer PowerSoil protocol with

a 25:24:1 phenol/chloroform/isoamyl alcohol modification to improve yield.[71] Phasing

amplicon sequencing was used to sequence the V4 region of 16S rRNA gene (SI Method

A.1.1).[203] Library of the 16S rRNA gene amplicon sequencing were constructed using

MiSeq reagent kit (Illumina, San Diego, CA) and DNA was sequenced using an Illumina

MiSeq platform.

Sequencing data were processed to combine pair-end reads and filter out poorly over-

lapped and unqualified sequences using the Amplicon Sequencing Analysis Pipeline (ASAP

version 1.3). The sequences (2 x 251 bp) were subjected to quality check with FastQC

(version 0.11.5). Pair-end sequences were merged based on the 3‘ overlap using PEAR (ver-

sion 0.9.10) with a quality score cuto↵ of 20 and minimum overlap length of 40 bp.[216]

Samples were demultiplexed using split libraries fastq.py from the QIIME package (ver-

sion 1.9.1) based on the barcodes (maximum barcode error of 0 and a trimming quality

score cuto↵ of 20).[24] Primer sequences were trimmed. Dereplication was performed using

USEARCH (version 9.2.64) with the command fastx uniques (with -sizeout for sequence

abundance output). Operational Taxonomic Units (OTUs) were clustered using UPARSE

(command -cluster otus of USEARCH) with OTU identity threshold of 97% and singletons

and chimeric sequences were removed.[47] Representative sequences of the OTUs were clas-

sified using RDP Classifier (training set 16, June 2016) with confidence cuto↵ of 0.8.[193]

Diversity was evaluated using the R package “vegan.”[126] Di↵erential abundance analysis

was conducted using the Bioconductor package DESeq2 in QIIME with the Benjamini-

Hochberg (BH) correction for multiple testing.[106, 8] Due to sample limitations there was

an uneven number of replicates for each diet, with more replicates available for the PE-fed

diets (Table A.1).

2.3.5 Statistical analysis

Statistical analyses were performed in Prism (GraphPad Software, version 7.0a). To assess

di↵erences in survival, plastic consumption, changes in molecular weight, and microbial

diversity ANOVAs were performed, followed by pairwise comparisons using Student’s t-

test with Tukey’s correction to assess di↵erences between diets. All p-values are adjusted

p-values and all error values are average ± standard deviation.

2.4 Results and Discussion

2.4.1 PE consumption and e↵ects on survival

At the end of the 32-day experiment, the survival ratio (SR) of the mealworms fed PE was

98.3% ± 0.0%, a value that was not significantly di↵erent (p = 0.92) from that of the bran

fed controls (96.3% ± 4.1%) (Fig. 2.2a). There was also no significant di↵erence (p = 0.65)

in SR of mealworms fed PE alone and mealworms fed PE + bran (95.0% ± 1.2%) (Fig.

1a). The SR of PS-fed mealworms (90.8% ± 2.4%) and PS + bran fed mealworms (91.3%

± 1.8%) were similar to values previously reported and were also not significantly di↵erent

2.4. RESULTS AND DISCUSSION 23

from those of mealworms fed PE (PS: p = 0.06, PS + bran: p = 0.08) PE + bran (PS: p =

0.44, PS + bran: p = 0.54), or the bran-fed controls (PS: p = 0.21, PS + bran: p = 0.27)

(Fig. 2.2a).[212, 207]

Consumption of PE and PS increased throughout the experiment (Fig. 2.2b). From

the initial 1.80 g PE, the total mass loss at the end of the experiment was 0.87 ± 0.0 g by

mealworms fed PE (Fig. 2.2c,d). For mealworms fed PS, the total PS mass loss was 0.57 ±0.12 g (Fig. 2.2c,d). For both PE- and PS-fed mealworms, the mass loss was significantly

greater when the mealworms received bran as a co-feed. For PE + bran, the mass loss was

1.10 ± 0.12 g and for PS + bran, the mass loss was 0.98 ± 0.11 g (Fig. 2.2c,d). Specific

rates of plastic consumption (mg plastic consumed per 100 worms per day) followed the

same pattern (Fig. 2.2d). The increase in specific consumption when co-fed with bran

supports previous findings.[207]

2.4.2 Evidence for depolymerization and biodegradation of PE

Residual polymer extracted from the frass of the bulk-fed containers was used to as-

sess biodegradation and depolymerization within the mealworm gut. The egested frass

contains an extractable fraction, consisting of non-degraded and partially degraded poly-

mer, and a non-extractable fraction (other biological waste not recovered in the extraction

process).[207] A decrease in the extractable fraction (“residual polymer”) suggests that more

of the ingested plastic is being completely degraded (mineralized) and/or is being incorpo-

rated into mealworm biomass.[207] The extractable portion of the frass from mealworms

fed PE or PS decreased over the course of the 32-day experiment (Fig. 2.3b).

Depolymerization of PE and PS was characterized using HT-GPC on the residual poly-

mers. HT-GPC analysis of the residual polymers from mealworms fed PE and PE + bran

showed a significant decrease in weight-averaged (Mw) and number-averaged (Mn) molec-

ular weight compared to the control PE (Fig. 2.3a). The residual polymer from PE-fed

mealworms showed an average reduction in Mw of 61.3 ± 5.0% and reduction in Mn of 40.1

± 8.5% (Fig. 2.3a; Table 2.1). The residual polymer from mealworms fed PE + bran showed

an average reduction in Mw of 51.8 ± 9.3% and reduction in Mn of 47.6 ± 8.5% (Fig. 2.3a;

Table 2.1), indicating significant depolymerization of PE occurred within the gut of the

mealworms fed PE and PE + bran. HT-GPC analysis of the residual polymer from meal-

worms fed PS and PS + bran also revealed significant decreases in Mw and Mn compared

to the control PS (Fig. A.1a; Table 2.1). The observed depolymerization in mealworms fed

PS and PS + bran is consistent with previous observations (Table 2.1).[212, 207]

Chemical modifications of the residual polymer were examined using 1H-NMR analysis.

Comparison of the control PE spectra to the spectra of the residual polymer from meal-

worms fed PE and PE + bran revealed a new peak around 5.3 ppm in a region associated

with alkene bonds (C=C-H) (Fig. 2.3c; Tables A.2, A.3). This peak was not observed in

spectra of control extractions from the frass of bran-fed mealworms, suggesting this peak

is not a result of contamination from the frass or the extraction method. Comparison of

the control PS spectra to the spectra of the residual polymers from PS and PS + bran

fed mealworms revealed new peaks associated with the incorporation of oxygen, previously

associated with plastic degradation (Fig. 2.3c; Tables A.2, A.3).[212, 207] To further assess

the possibility that the newly observed peaks were attributable to contamination by bac-

terial biomass or secreted proteins, samples of plastic (PE and PS) were separately treated

in a suspension of E. coli K12 for 48-hours then subject to the same extraction methods

(SI Method A.1.3).[196] The NMR spectra for PE and PS control samples incubated with

E. coli K12 did not reveal new peaks relative to the control spectra, supporting the con-

clusion that newly observed peaks in the residual polymers spectra (from PE- and PS-fed

mealworms) are evidence of chemical modifications resulting from plastic degradation and

are likely not due to contamination (Fig. A.2a,b).[196]

Additional evidence of chemical modifications in the residual polymer was obtained by

FTIR analysis. FTIR spectra from the residual polymers from mealworms fed PE and PE

+ bran revealed incorporation of oxygen as indicated by the appearance of peaks associated

with C-O stretching (1000–1200 cm-1) and alcohol groups (R-OH bend, 1300–1450 cm-1;

R-OH stretching, 3000–3500 cm-1) (Fig. 2.3d; Tables A.4, A.5, A.6). These peaks were not

observed in the control PE spectra. FTIR spectra for the residual polymers from mealworms

fed PS and PS + bran also revealed chemical modifications and the incorporation of oxygen

(Fig. A.1d; Tables A.4, A.5, A.6). Again, to assess whether the new peaks were byproducts

of microbial contamination, control plastic (PE and PS) incubated with E. coli K12 were

also subject to FTIR analysis (SI Method A.1.3). FTIR spectra of these controls did not

reveal any new peaks, supporting the conclusion that the newly observed peaks resulted

from plastic degradation within the mealworm gut and are likely not due to contamination

(Fig. A.2c,d; Tables A.4, A.5, A.6).[196]

2.4.3 Evidence for mineralization via a mass balance

A mass balance on the plastic-fed mealworms was conducted by measuring the weights of

system inputs (plastic and/or bran), outputs (the weight of the frass before extraction and

the weight of the extractable fraction), and the weight of accumulated biomass (changes in

the weight of surviving mealworms).[212] The mealworm containers were kept at a constant

humidity and all weights were measured as wet-weights to avoid errors due to losses or gains

in water vapor. Deviations from a perfect mass balance (e.g., if the outputs 6= the inputs +

changes within the container, “putative gas fraction”) would be due to losses into the gas

phase (e.g., mineralization), which were not directly measured.

The putative gas fraction (PGF) increases over the course of the experiment for PE-fed

mealworms while the extractable fraction from the frass decreased, both of which suggest

more degradation (and mineralization) occurred towards the end of the 32-day experiment

(Fig. 2.3b; Fig. A.3a). Mealworms fed plastic alone (PE or PS) gained less biomass

weight over the course of the experiment than mealworms co-fed with bran, an observation

consistent with previous studies of PS-fed mealworms (Fig. A.3).[207] By mass balance,

the PGF was 49.0 ± 1.4% at the end of the experiment for PE-fed mealworms (Fig. A.3a).

For mealworms fed PE + bran, the PGF was 24.3 ± 2.2% with a higher percent of the

ingested mass being incorporated into the mealworm biomass (Fig. S3b). The PGF in

PS-fed mealworms was 45.5 ± 2.9%, a value similar to that previously reported for PS

mineralization by mealworms (measured via mass balance and 13C-carbon tracing) (Fig.

A.3c).[212] The PGF of PE- and PS-fed mealworms were not significantly di↵erent.

2.4.4 Biodegradation of mixed plastics

When fed a diet of one plastic (PE or PS, with or without bran) there was no significant

di↵erence between plastic mass loss by the mealworms based on the plastic (Fig. 2.2b, c,

d). However, when mealworms were co-fed PE and PS (1:1 [w/w]), there was a significantly

higher mass loss of PE than PS (Fig. 2.2c, d). A previous report suggested that di↵erences

in plastic consumption rates among di↵erent types of PS foams may be due to density, with

a higher consumption rate associated with less dense plastics.[207] In this study, however,

the PE plastic had a higher density than the PS plastic, suggesting that other factors may

a↵ected the relative rates of mixed plastic consumption. Further work is needed to assess

what factors influence di↵erential plastic consumption rates.

The molecular weight distributions of the mixed polymers could not be di↵erentiated via

HT-GPC, therefore characterization of depolymerization was not included in this analysis.

However, FTIR analysis of the residual polymers revealed chemical modifications and the

incorporation of oxygen relative to the control PE and PS spectra, similar to those observed

in the residual polymers from mealworms fed PE or PS, which suggests degradation of the

mixed polymers occurred within the mealworm gut (Fig. A.4; Tables A.4, A.5, A.6). This

finding o↵ers further evidence that plastic degradation within the mealworm gut is non-

specific.

2.4.5 E↵ects of plastic consumption on gut microbiome

Next-generation sequencing was used to investigate the e↵ect of plastic diets on microbial

community structure (Table A.1). Relative abundance analysis revealed the same three

majority community members (OTUs) in all diets: Spiroplasma sp., Cronobacter sp., and

Enterococcus sp. (Fig. 2.4a). These three OTUs are common insect gut-associated bacteria

and are known members of the T. molitor gut microbiome.[194, 65]

We analyzed microbial diversities across all six experimental diets. The alpha diversity

of the microbial community, measured via the inverse Simpson index, was not significantly

di↵erent between the di↵erent diets (Fig. 2.4b). A principal coordinate analysis (PCoA)

based on Bray-Curtis dissimilarity index revealed clusters associated with di↵erent diets,

with clear clusters for PE-fed and bran-fed mealworms (Fig. 2.4c). A multi-response per-

mutation procedure test revealed a significant di↵erence in the microbial communities based

on diet (p = 0.001). This suggests that while the majority members of the microbial com-

munity do not di↵er dramatically in PE diets, the composition of the microbial community

is distinct from either bran-fed or PS-fed communities.

Di↵erential abundance analysis was used to assess whether particular OTUs were asso-

ciated with di↵erent diets (Fig. 2.5, Table A.7). This analysis revealed two OTUs that were

strongly associated (p < 0.05) with both the plastic diets (PE and PS): Citrobacter sp. and

Kosakonia sp. Both OTUs are members of the Enterobacteriaceae, a family known to con-

tain PE-degrading member Enterobacter absuriae YT1 isolated from the gut of the larvae of

Indian mealmoth.[206] Both OTUs can utilize oxygen (Citrobacter sp. are aerobic, Kosako-

nia sp. are facultative anaerobic), which could be further evidence for their involvement in

plastic degradation, as incorporation of oxygen is key in the accelerated biodegradation of

both PE and PS, as evident in the analysis of residual polymers and previous work (Fig.

2.3c, d; Fig. A.1c, d).[158, 173, 93, 65] Both Citrobacter sp. and Kosakonia sp. were more

abundant (based on relative abundance) in both of the plastic-only diets than the plastic +

bran fed diets and were also more abundant than the other OTUs identified via di↵erential

abundance analysis (Table A.7).

Two OTUs, both minority members of the microbial community, were significantly as-

sociated (p < 0.05) with PE-fed microbiomes: Sebaldella termitidis and Brevibacterium sp.

(Fig. 2.5b, c, d; Table A.7). Sebaldella termitidis is phylogenetically isolated within the

phylum Fusobacteria, is anaerobic, and is a known inhabitant of the posterior end of the

termite gut track.[72] Brevibacterium sp. are aerobic bacteria known to be associated with

hydrocarbon degradation, including n-alkanes.[27] Further work should assess the involve-

ment of Brevibacterium sp. in the degradation of polyethylene.

Seven OTUs, all minority members of the microbial community, were significantly asso-

ciated (p < 0.05) with the PS-fed gut microbiome: Listeria sp., Nitrospira defluvii, Pedomi-

crobium sp., Aquihabitans sp., unclassified Xanthomonadaceae, unclassified Saprospiraceae,

and unclassified Burkholeriales (Fig. 2.5c,d; Table A.7). Most of these PS-associated OTUs

are aerobic, which is important when considering their possible role in the degradation of

polystyrene. The increase in OTUs associated with the PS microbial community could be

indicative of a more diverse suite of daughter products created in PS degradation, likely due

to the more complex chemical composition of PS and the presence of benzene rings that

could degrade into a variety of daughter products. Changes in the PS-microbial community

could also be a↵ected by the presence of trace amounts (< 1%) of a chemical flame retardant

(present in most commercially available PS products). Further research is needed to assess

whether and how trace chemicals (especially flame retardants in PS materials) a↵ect the

microbial community.

Overall, di↵erential abundance analysis of the gut microbiome revealed several minority

OTUs strongly associated with the plastic diets. The gut microbiome, which previous

work has shown is necessary for PS degradation, shows changes in response to di↵erent

plastic diets, further suggesting the importance of the microbial community in the plastic

degradation process. While further work is needed to assess the role of individual OTUs

in the plastic degradation pathway, this analysis o↵ers an initial insight into what species

might be of interest in future studies.

2.5 Implications

This work is the first report to demonstrate that PE is depolymerized and undergoes chem-

ical modifications within the mealworm gut. Additionally, we demonstrated for the first

time that mixed plastics (PE and PS) undergo biodegradation within the gut. Application

of next-generation sequencing to the gut microbiome revealed two OTUs (Citrobacter sp.

and Kosakonia sp.) strongly associated with both PE and PS as well as OTUs unique to

each plastic. Our findings suggest that plastic degradation within the mealworm gut is not

plastic-specific. This could have implications for future waste management applications.

As the mealworms in this study were previously untested for plastic degradation, their

ability to degrade both PE and PS further suggests the ubiquity of plastic degradation

among mealworms. Further work is needed to assess whether other recalcitrant plastics that

resemble polyethylene (e.g., polyvinyl chloride and polypropylene) degrade in the mealworm

gut. Future work should focus on elucidating the mechanisms of degradation within the

mealworm to enable future applications.

2.6 Supporting Information

Appendix A contains more detailed information about the methods and results from this

study, including Tables A.1 through A.7 and Figures A.1 through A.6.

2.7. TABLES 29

2.7 Tables

Table 2.1: Characterization of PE and PS foams tested before and after degradation bymealworms (mean ± standard deviation, n = 4 for control samples, n = 3 for frass samples).

2.8 Figures

2.8. FIGURES 31

0 10 20 300

Bran OnlyPE OnlyPE + Bran (1:1)PS OnlyPS + Bran (1:1)PE + PS (1:1)

PE Only

PE + Bran (1:1)

PS Only

PS + Bran (1:1)

PE + PS (PE)

PE + PS (PS)0

0 10 20 300

(%) PE Only

PE + Bran (1:1)PS OnlyPS + Bran (1:1)PE + PS (PE)PE + PS (PS)

PE Only

PE + Bran (1:1)

PS Only

PS + Bran (1:1)

PE + PS (PE)

PE + PS (PS)0

(a) (b)

(c) (d)

Figure 2.2: Survival rate and plastic consumption by T. molitor by diet. (a) Survival rateof mealworms over 32-day experiment. (b) Mass loss in plastic (PE or PS) in the plastic-feddiets over 32-days. (c) The percent mass loss in the plastic by diet at the end of the 32-dayexperiment. (d) Average specific plastic consumption (mg plastic per 100 mealworms perday) over the 32-day experiment. All values represent mean ± SD, n = 2. Significance(Student’s t-tests, Tukey’s multiple test correction) p < 0.05 indicated by ⇤, p < 0.005indicated by ⇤ ⇤, no statistical significance indicated by ns. For mealworms fed PE + PS,the mass loss is displayed separately for each plastic.

1×104

2×104

3×104

4×104

5.0×104

1.0×105

1.5×105

2.0×105

2.5×105

olecular Weight (M

50010001500200025003000350040000

Wavenumber (cm-1)

ance PE Frass

PE + Bran FrassOHR

OHRstretch

stretch

PE ControlC Ostretch

likely C=C-H

Bran Frass

PE + Bran Frass

PE Frass

PE Control

(d)Week 1

Week 2Week 3

Week 40

PE Frass

PE + Bran Frass

PE Control

Figure 2.3: Characterization of polyethylene degradation within the mealworm gut. (a)Changes in molecular weight (Mn and Mw) within residual polymer from the frass versusthe control PE as measured by HT-GPC. Significance (Student’s t-tests, Tukey’s multipletest correction) p < 0.05 indicated by ⇤, p < 0.0005 indicated by ⇤ ⇤ ⇤, no statisticalsignificance indicated by ns. (b) Changes in the extractable fraction of the frass (% recoveredby extraction), a measure of residual non-degraded and partially degraded polymer in thefrass, over the 32-day experiment. (c) 1H-NMR spectra of residual polymer from the frass ofPE and PE + bran fed mealworms versus residuals extracted from bran-fed mealworms andthe control PE foam. The appearance of alkene derivatives is highlighted in grey. Detailedpeak information in Table S2. (d) FTIR spectra of residual polymer from the frass of PEand PE + bran fed mealworms versus the control PE, annotations show functional groupsassociated with key peaks based on wavenumber. Detailed peak information in Tables A.4,A.5, A.6.

2.8. FIGURES 33

●●

● ●

●●

● ●

PS_Bran1PE_PS2

PE1PE2

PE_Bran1PE_Bran3

PE_Bran4

PS2PE_Bran2

PE_PS1

PE7PE9PE6

PE_Bran5

PE_Bran4

PE8PE3

PE5Bran2 PE_Bran3

−0.2

−0.1

−0.2 0.0 0.2Axis.1 [28.5%]

Diet●●

●●

PE_Bran

PS_Bran

Bran PE

PE + BranPE + PS PS

PS + Bran0

Clostrium senso stricto sp. Lactobacillus sp.Lactococcus sp.un. BacteriaEnterococcus sp. Cronobacter sp. Spiroplasma sp.

Listeria sp.Bacillus sp. Sebaldella termitidis

Most Abundant OTUs:

Bran PE

PE + Bran

PE + PS PS

PS + Bran0

(a) (b)

Figure 2.4: Microbial community analysis of gut microbiome in di↵erent diets. (a) Changesin community composition by family by diet (average from all replicates, Table A.1), legendshows ten most abundant OTUs across all diets, un. indicates unclassified. (b) InverseSimpson Index of the gut microbiome by diet. No statistical significance (ANOVA, Tukey’smultiple test correction) indicated by ns. (c) Principal Coordinate Analysis (PCoA) ofmicrobial communities by diet based on Bray-Curtis distance, colored by diet and labeledwith the sample ID (two outliers removed: PS1, PS Bran2; PCoA with outliers Fig. A.5).

-10 -5 0 5 10

Clostrium sensu stricto sp.Clostrium sensu stricto sp.

Bacillus sp.Un. LachnospiraceaeBrachybacterium sp.

Un. EnterobacteriaceaeStaphylococcus sp.

Un. LachnospiraceaeBrevibacterium sp.

Sebaldella termitidisCorynebacterium sp.

Kosakonia sp.Citrobacter sp.

log2 Fold Change

PE vs. Bran

PE AssociatedBran Associated

-10 -5 0 5 10

Listeria sp. Nitrospira defluviiAquihabitans sp.

Un. XanthomonadaceaePedomicrobium sp. Un. BurkholderialesUn. SaprospiraceaeChondromyces sp.

Un. BurkholderialesSphingorhabdus sp.

Ferruginibacter sp. Brevibacterium sp.

Sebaldella termitidis

log2 Fold Change

PE vs. PS

PE AssociatedPS Associated-10 -5 0 5 10

Clostrium sensu stricto sp. Clostrium sensu stricto sp.

Bacillus sp. Un. Myxococcales

Un. SaprospiraceaeUn. Xanthomonadaceae

Pedomicrobium sp. Aquihabitans sp.

Un. BurkholderialesNitrospira defluvii

Citrobacter sp. Listeria sp.

Kosakonia sp.

log2 Fold ChangeBran Associated

PS vs. Bran

PS Associated

-10 -5 0 5 10

Clostrium sensu stricto sp.Bacillus sp.

Citrobacter sp.Un. Clostridiales

Clostrium sense stricto sp.Cronobacter sp.

Brachybacterium sp.Un. Enterobacteriaceae

Un. LachnospiraceaeKosakonia sp.

Brevibacterium sp.Sebaldella termitidis

Citrobacter sp.

log2 Fold ChangePE AssociatedPE + Bran Associated

PE vs. PE + Bran(a) (b)

(c) (d)

Figure 2.5: Di↵erential abundance analysis of gut microorganisms between experimentaldiets. OTUs shown significantly (BH adjusted p < 0.05) di↵ered between diets. Directionof fold change (log2) indicates which diet each OTU is more strongly associated (labeledbelow x-axis). (a) PE-fed microbiome versus bran-fed microbiome. (b) PE-fed microbiomeversus PE + bran-fed microbiome. (c) PE-fed microbiome versus PS-fed microbiome. (d)PS-fed microbiome versus bran-fed microbiome.

Chapter 3

Enhanced polystyrene

biodegradation in an ex situ gut

microbiome enrichment from

Tenebrio molitor (mealworm

larvae)

The results of this chapter are in preparation for publication. Alexa M. Gracia helped

design and perform laboratory experiments. Dr. Wei-Min Wu contributed to the design of

the study and provided comments to improve the manuscript.

36 CHAPTER 3. ENHANCED EX SITU DEGRADATION

3.1 Abstract

As the global threat of plastic pollution has grown in scale and urgency, so have e↵orts

to find sustainable and e�cient solutions. Research conducted over the past few years has

identified gut environments within insect larvae, including Tenebrio molitor (mealworms),

as microenvironments uniquely suited to rapid plastic biodegradation. However, there is

currently limited understanding of how the insect host and its gut microbiome collaborate

to create an environment conducive to plastic biodegradation. In this work, we provide

evidence that T. molitor secretes one or more emulsifying factor(s) (30 – 100 kDa) that

mediate plastic bioavailability. We also demonstrate that the insect gut microbiome se-

cretes factor(s) (<30 kDa) that enhance respiration on polystyrene. We appy these insights

to culture polystyrene-fed gut microbiome enrichments, with elevated rates of respiration

and degradation compared to the unenriched gut microbiome. Within the enrichment, we

identified eight unique microorganisms that are associated with polystyrene biodegradation

including Citrobacter freundii, Serratia marcescens, and Klebsiella aerogenes. Our results

demonstrate that the mealworm itself and its gut microbiome both contribute to the accel-

erated plastic biodegradation. This work provides new insights into insect-mediated mech-

anisms of plastic degradation and potential strategies for cultivation of plastic-degrading

microorganisms in future investigations and scale-up.

Mealworm host

Gut microbiome

T. molitorC C

Emulsifing agent

Respiration enhancing factor

Gut supernatant+ gut microbiome

Polystyrene

CO2 , H2O & degradedresiduals

3.2 Introduction

Plastic pollution is a well-known threat to human and environmental health. Strategies are

needed for e↵ective management of recalcitrant plastic wastes because current cradle-to-

grave lifecycles result in plastic accumulation in landfills and in the environment.[66] Thus,

there is an urgent need to permanently and completely eliminate plastic waste from this

linear lifecycle. Recent work to achieve this goal has focused on natural plastic biodegra-

dation, mainly identifying and isolating bacterial strains capable of biodegrading plastic

polymers.[158, 65, 93, 200, 124] Rates of plastic degradation vary among these microbes,

but are typically low, ranging from ⇠ 5% plastic degradation (by mass) within six months

(⇠ 0.03% per day) to ⇠ 8% within two weeks (⇠ 0.6% per day).[158, 65]

A fundamental limitations to microbial biodegradation of plastics is the limited bioavail-

ability of plastics to microbial attack.[158, 93, 197] Petroleum-derived plastics have high

molecular weights and are highly hydrophobic, both of which limit the ability of microor-

ganisms to interact with these materials, especially in aqueous environments.[158, 93, 197]

E↵orts to improve plastic biodegradation rates have revealed that the gut environment of

insect larvae can support comparatively rapid plastic degradation. To date, plastic degra-

dation has been demonstrated in several holometabolic insect species: Tenebrio molitor

(yellow mealworms),[213, 213, 210, 207, 18] Tenebrio obscurus (dark mealworms),[134] Zo-

phobas attratus (superworms),[133, 89, 140] Plodia interpunctella (Indianmeal moth),[206]

Achroia Grisella (lesser wax moth),[95] and Galleria mellonella (greater wax moths).[91,

147, 135, 105, 25] Of these, the best characterized is the yellow mealworm.[210] Prior re-

search has established that ingested plastic is retained within the gut of yellow mealworms

for 15 – 20 hours, during which time nearly 50% of the plastic is mineralized to CO2 (⇠60% per day).[212, 207, 207] Researchers have also demonstrated that the gut microbiome of

yellow mealworms can degrade several chemically-dissimilar plastics (including polystyrene

and polyethylene), suggesting an initial, non-specific attack.[18]

Plastic-degrading microorganisms demonstrate expedited degradation within the insect

gut environment, implying that the insect host may play a role in the plastic biodegrada-

tion process. When gut microbiome activity is suppressed, Galleria mellonella retains

the capacity to metabolize polyethylene.[91] In addition, marked changes are observed

in the proteins secreted by the insect’s salivary glands, underscoring its role in plastic

degradation.[91] When the gut microbiome is left intact, the microbial community compo-

sition shifts during degradation and one plastic-degrading bacterium was isolated: Enter-

obacter sp. D1.[147, 105] This suggests a complex system in which the insect host and its

gut microbiome collaborate to degrade plastic.

Mealworms are a useful model organism for plastic biodegradation studies because they

are readily cultivated and are the subject of a rapidly growing body of research.[210, 18,

177, 141] As with G. mellonella, prior research has demonstrated that the mealworm gut

microbiome undergoes notable changes during plastic degradation.[210, 18] Suppression of

gut microbiome by supplementing their diet with antibiotics indicates that the microbiome

is necessary for polystyrene (PS) biodegradation.[213] PS-degrading Exiguobacterium sp.

YT2 has been isolated from the mealworm gut microbiome[213] and researchers have inves-

tigated enzyme activity in both the mealworm and the gut microbiome, though the specific

mechanisms and the role of the mealworm itself have yet to be identified.[141]

In this work, we describe e↵orts to increase plastic biodegradation rates by enhancing

gut microbiome-derived enrichments outside the gut microenvironment. To do this, we first

investigated the roles of the mealworm and its gut microbiome in plastic degradation. We

then applied these insights to enhance plastic degradation in microbial enrichments outside

the mealworm.

3.3.1 Plastic materials

Commercially available expanded polystyrene (PS) foam (thickness: 5.2 cm, weight-averaged

molecular weight (Mw): 170700 ± 9800 Da, number-averaged molecular weight (Mn):

88500 ± 5900 Da) was used as a feed for the mealworms (Carlisle Construction Materi-

als, Puyallup, WA). Prior to use, the PS foam blocks were cut into 2 – 3 cm cubes and

cleaned with a stream of air to remove any residues. PS microplastics (size: ⇠ 150 µm, Mw:

76400 ± 4700 Da, Mn: 41500 ± 3300 Da) were used in microbial cultures (Jinshuowang

Plastic Materials Co., Ltd, Dongguan, Guangdong, China). PS microplastics were either

used directly in microbial cultures or heat-pressed at 260 �C using PHI Manual Compression

Press (PW-22 Series, Industry, CA) into uniform thin films (thickness: 17.6 ± 1.1 mm) for

use in microbial cultures.

3.3.2 Mealworm maintenance

Mealworms, larvae of Tenebrio molitor Linnaeus, (average weight: 75-85 mg/worm) were

purchased online from Rainbow Mealworms (Compton, CA) and shipped overnight to the

Environmental Engineering and Science laboratories at Stanford University (mealworms

from this source have previously been shown to degrade PS).[16, 17, 210, 207] Prior to

arrival, the mealworms were fed bran; after arrival, they were subject to a 48-hour starvation

period before initiating tests with the experimental diet of either natural wheat bran (Exotic

Nutrition, Newport News, VA) or PS foam. Mealworms (⇠ 1200 per experimental condition)

were reared in food grade polypropylene containers (volume: 780 mL) and kept in incubators

maintained at 25 �C and 70% humidity.[212, 207]

To understand the role of the mealworm and its gut microbiome as well as the en-

dogenous response of the mealworm to its natural diet versus an experimental plastic diet,

mealworms were reared under four conditions: (1) bran (control diet), (2) polystyrene (PS),

(3) bran + antibiotic,and (4) PS + antibiotic. Gentamicin sulfate (Acros Organics, Fisher

Scientific, Pittsburg, PA) was the antibiotic used to suppress the gut microbiome as de-

scribed in previous studies.[210, 213] Gentamicin was added at 30 mg/g bran to the bran

+ antibiotic group and an equivalent mass was added to the PS + antibiotic group. PS-fed

mealworms were initially given 2.6 g of PS foam. Bran-fed mealworms were initially given

12 g bran. Bran and gentamicin were supplemented every three days, and dead mealworms

were removed. Mealworms were maintained on the experimental diets for ⇠ 3 weeks before

use in any experiments. Prior to testing, mealworms were subjected to a 24-hour starvation

period to ensure removal of antibiotic residuals.

3.3.3 Collection of gut microbiome and supernatant

To collect the mealworm gut microbiome and produce a supernatant containing secreted

factors from the mealworm and its gut bacteria (henceforth, “supernatant”), the following

procedure was used. Fifty mealworms from each diet were scarified following established

methods.[213] Briefly, mealworms were immersed in 75% ethanol for one minute, then rinsed

with DI water. Mealworm guts were harvested via dissection into 0.6 mL sterile saline

(0.75% NaCl) in a sterile microcentrifuge tube. Samples were vigorously vortexed (Vortex-

Genie 2, MO BIO, Carlsbad, CA), and the liquid phase was collected and replaced with

sterile saline water. This was repeated ⇠ 3 – 4 times until the liquid phase was clear.

The collected liquid was then passed through a 40 µm nylon cell strainer (Falcon, Fisher

Scientific, Pittsburg, PA) and then centrifuged at 4000 rpm for 10 min to collect gut bacteria.

Following centrifugation, the supernatant was filtered through a 0.2 µm syringe filter

(25mm, nylon, Fisher Scientific, Pittsburg, PA) to remove remaining particulates. To better

understand the role(s) of secreted factors, supernatant was fractionated by molecular weight

using molecular weight spin columns (Amicon Ultra Centrifugal Filters, MilliporeSigma, St.

Louis, MO) into three fractions: <30 kDa, 30 – 100 kDa, and >100 kDa. All supernatant

samples were stored at 4 �C until used in experiments. Additional supernatant was collected

as needed to ensure samples were not stored for longer than three weeks.

3.3.4 Enrichment and isolation of PS-degrading bacteria

Amicrobial enrichment was used to study the potential for cultivation of key gut microbiome

members outside the mealworm gut. The gut microbiome was collected from 50 PS-fed

mealworms as described above. The collected gut bacteria were then inoculated into 30

mL of carbon-free Bushnell Haas (BH) media containing the following chemicals (g/L): 0.2

MgSO4, 0.02 CaCl2, 1.0 KH2PO4, 1.0 K2HPO4, 1.0 NH4NO3, 0.05 FeCl3.[23] PS film (0.5

g) was added to the culture along with the supernatant from PS-fed mealworms (0.18 mL,

0.6% v/v). The enrichment was grown in a round-bottomed glass culture tube (25mm outer

diameter, Fisher Scientific, Pittsburg, PA), sealed with a Kim-Kap (25mm outer diameter,

Fisher Scientific, Pittsburg, PA), and incubated at 30 � C on an orbital shaker table (150

rpm). The enrichment was cultured for three months weekly with 10 mL media changes

and supplemented with PS supernatant until a stable biofilm developed on the PS film (Fig.

Serum bottle assays were conducted to assess the volume of CO2 produced by PS-

degrading enrichments. Cultures were incubated in 25 mL serum bottles (Wheaton, Mealville,

NJ) capped with butyl-rubber stoppers (MilliporeSigma, St. Louis, MO) and crimp-sealed

(MilliporeSigma, St. Louis, MO) under an air headspace of one atmosphere. Liquid volume

was 15 mL and the headspace volume was 10 mL. Cultures were incubated at 30 � C on an

orbital shaker table (150 rpm). All serum bottle assays were conducted in triplicate. Serum

bottles were inoculated with either bacteria from the mealworm gut or bacteria from the

enrichment. To obtain su�cient inoculum, bacteria from either sample were first grown

in 15 mL tryptic soy media for 24 hours at 30 � C. The bacteria were then collected by

centrifugation, washed with BH media, and spun again prior to use.

To isolate bacterial strains involved in plastic degradation from the PS-degrading en-

richment, 30 µL liquid culture from the enrichment was plated on non-selective agar plates

(tryptic soy agar, Difco, Fisher Scientific, Pittsburg, PA). Colonies were picked with sterile

inoculum loops and streak-plated to isolate individual pure colonies, which were then were

grown in 3 mL tryptic soy media (MilliporeSigma, St. Louis, MO) for 24 hours at 30 � C.

A bacterial pellet was then collected by centrifugation, rinsed with BH media, and spun

again prior to use. The rinsed bacteria were then inoculated in 5 mL BH media with PS

film, supplemented with PS supernatant (30 µL, 0.6% v/v) and cultivated at 30 � C on an

orbital shaker (150 rpm).

To identify bacteria associated with plastic-degradation, colonies were isolated from

the enrichment and sequenced. Liquid media from the enrichment and isolates described

above was first plated on tryptic soy agar plates. For each isolate (16 total), an individ-

ual colony was randomly selected for sequencing. For the enrichment, individual repre-

sentative colonies were selected for sequencing (16 total). DNA was extracted from the

selected colonies using FastDNA SPIN Kit for Soil (MP Biomedicals, Santa Ana, CA,

USA) following manufacturer’s protocols. The 16s rRNA V4 region was then PCR ampli-

fied using 8F (5’AGAGTTTGATCCTGGCTCAG3’) and1492R (5’GGTTACCTTGTTAC-

GACTT3’). Amplification was confirmed by denaturing gel electrophoresis and clean PCR

products (QIAquick PCR Purification Kit, Qiagen, Hilden, Germany) were sent to MCLAB

(San Francisco, CA) for sequencing on ABI 3730xl automated sequencers. Pair-end reads

were aligned in Geneious Prime (2020.1.1) and strains were identified by BLAST sequence

similarity search through the NCBI 16s rRNA database.

3.3.5 Microbial activity analysis

Biolog MT2 Microplates were used (Biolog Cat. # 1013, Hayward, CA) were used to

screen the ability of cultivated bacteria and the mealworm gut microbiome to metabolize

PS as their sole carbon source. Each MT2 microplate is pre-loaded with bu↵ered nutrient

medium and a redox dye (tetrazolium violet), a well-established colorimetric indicator of

bacterial carbon oxidation (respiration) by bacteria.[63, 62] Bacterial cells from the gut

microbiome were collected via centrifugation as described above and resuspended in sterile

saline (0.75% NaCl). The cell suspension was inoculated into the MT2 microplate at 130

µL per well, followed by the addition of experimental supernatant and/or PS microplastics

(⇠ 3 mg) to the appropriate wells. All experimental conditions were tested in triplicate

with a negative control (loaded with inoculum and supernatant without PS) in duplicate.

The microplate was then incubated at 30 � C with gentle shaking in the Synergy HTX

Multi-Mode Plate Reader (BioTek Instruments, Winooski, VT) with hourly colorimetric

measurements (absorbance: 595 nm).[104, 111]

To assay the CO2 produced in the headspace of the serum bottles, 0.25 mL of gas from

each serum bottle was injected onto GOW-MAC gas chromatograph with an Altech CTR 1

column and a thermal conductivity detector. The following method parameters were used:

injector, 120 � C; column, 60 � C; detector, 120 � C; and current, 150 mV. The peak area of

CO2 was compared to standards and quantified using the software ChromPerfect (Justice

Laboratory Software, Denville, NJ, USA) to determine the concentration (measured as

percent of the headspace volume) CO2 in the headspace of each vial.

3.3.6 Characterization methods

Contact angle analyses was performed to quantitatively explore di↵erences in the super-

natant hydrophobicity for each experimental condition. A Contact Angle Goniometer

(Rame-Hart model-290, Succasunna, NJ) was used to measure supernatant hydrophobicity

(i.e., the ability of the tested supernatant to wet a hydrophobic surface, PS film), with

deionized (DI) water tested as a control. Increased surface wetting indicates increased hy-

drophobicity of the liquid. For each sample, PS film was placed on a level platform and 5

µL of liquid supernatant was dropped onto the film, in quadruplicate. Contact angle anal-

ysis was also utilized to characterize changes in PS films after incubation with microbial

cultures. Control and treated films were places on a level platform and 5 µL of DI water was

dropped onto the film, in quadruplicate, to assess changes in the hydrophobicity of the film

surface. Contact angles were quantified using the DROPimage Pro Software (Rame-Hart,

Succasunna, NJ).

To characterize chemical changes in the PS film after incubation with microbial cultures,

films were visualized using Fourier Transform Infrared Spectroscopy (FTIR) on a Nicolet

iS50 FTIR Spectrometer (Thermo Fisher Scientific, Inc., Pittsburg, PA), as previously

described.[18] Absorbance spectra were recorded and transformed into transmittance for

graphing. Spectra were recorded between 4000 – 500 cm-1 with at least 16 scans at a

spectral resolution of 0.482 cm-1. Peaks were identified using OMNIC software (Thermo

Fisher Scientific Inc., Pittsburg, PA).

Statistical analyses were performed in Prism (version 7.0a). To compare respiration activity

and supernatant contact angles between experimental conditions, ANOVA tests were per-

formed, followed by pairwise comparisons using Student’s t-test with Tukey’s correction.

All p-values are adjusted p-values and all error values are reported as average ± standard

deviation of experimental replicates.

3.4.1 E↵ects of supernatant on ex situ microbial growth

A Biolog MT2 plate assay was used to assess respiration by the gut microbiome immediately

after extraction from mealworms. This assay demonstrated that the gut microbiome was

able to respire on PS as the sole carbon source and respiration increased significantly in

cultures supplemented with the gut supernatant from the mealworms fed either PS or bran

(p 0.05 for both, Fig. 3.2a). No increase in respiration activity occurred after the

addition of gut supernatant from mealworms fed antibiotic-treated diets; rather inhibition

was observed (Fig. 3.2a). Inhibition could be due to carry-over of antibiotics in the gut

supernatant or secretion of other factors (e.g., immune responses proteins[51]) that are

inhibitory or toxic to the gut microbiome. To test whether inhibition was due to carry-

over of antibiotic residues, positive controls (glucose-fed wells) were supplemented with

supernatant from antibiotic-treated mealworms. No loss in activity was observed (p =

0.16). This is evidence that the gut microbiome produces and secretes factors into the

supernatant that enhance degradation.

To better understand the beneficial e↵ect of the supernatant on respiration, the su-

pernatant was fractionated by molecular weight into three fractions: <30 kDa, 30 – 100

kDa, and > 100 kDa. The two smallest of these fractions (<30, 30 – 100 kDa) from each

condition were then subject to the same respiration assay described above. This assay

revealed a two-fild increase in respiration activity in the lowest molecular weight fraction

(<30 kDa), from both PS and bran diets (Fig. 3.2a). Taken together, our results suggest

that the smallest supernatant fraction contains a respiration enhancing factor secreted by

the gut microbiome of mealworms fed PS or bran. This factor increases the ability of the

gut microbiome to respire on, and thus degrade, PS outside the mealworm gut.

After identifying the fraction of the supernatant that contributed to increased PS-

degradation, the amount of supernatant needed to achieve this enhanced e↵ect was assayed.

To do this, five dosages of supernatant were tested: 0.0, 0.05, 0.1, 0.5, 1 mL (0.0, 0.3, 0.6,

3.0, 5.6% v/v). Supernatant dosages were used to supplement the gut microbiome from

PS-fed mealworms cultured in carbon-free media with PS microplastics in serum bottles.

CO2 production was measured over two weeks. The ideal dose was determined as the con-

centration of supernatant that led to the largest net volume of CO2 produced over 14 days,

measured as the final minus initial concentration of CO2 in the headspace. The initial con-

centration of CO2 was measured 24 hours after initializing experiments to ensure gas-liquid

equilibrium. To assess whether the increase in respiration was due to addition of an exter-

nal carbon source (rather than the presence of degradation-enhancing secreted factors), the

CO2 production of cultures supplemented with an equivalent amount of glucose (based on

chemical oxygen demand) was tested for each supernatant dose (Fig. B.1). Plots of each

dose along with the negative controls (gut microbiome supplemented with supernatant but

no PS) and glucose equivalent are provided in the Supporting Information (Fig. B.1).

At the lowest doses (0.0, 0.05, 0.1 mL), the addition of the supernatant increases the

amount of CO2 produced compared to the negative control, again highlighting the benefit

of supernatant addition for PS biodegradation (Fig. B.1a-c). Supplementation with the two

highest doses (0.5, 1.0 mL), did not increase the respiration of PS overtime; instead, high

doses led to a rapid increase in increased CO2 production in the first 24 hours, after which

the CO2 concentration stabilized (Fig. B.1d,e). This suggests that supplementation at

high doses with supernatant or glucose likely selects for bacterial strains that grow quickly

on the available carbon but do not necessarily biodegrade PS. The final change in CO2

produced over the course of the experiment was plotted by dose to determine the ideal dose

of supernatant to support PS degradation (Fig. 3.2b). The lower doses of supernatant led

to greater increases than the highest two doses, with 0.1 mL (0.6% v/v) being the clear

ideal dose. These results reveal that supplementing ex situ gut microbiome cultures with

supernatant improves PS respiration beyond the leves achieved by addition of bioavailable

carbon, supporting the hypothesis that the supernatant contains factor(s) that enhance the

capacity of the gut microbiome to respire and biodegrade PS.

3.4.2 Identification of endogenous emulsification activity

In conducting the experiments described above, addition of gut supernatant resulted in

rapid emulsification of hydrophobic PS microplastics into the aqueous media (Fig. B.2a).

Further, this emulsification was stable over time, enabling the microplastics to settle in the

aqueous phase as they do when treated with Tween-80, a known surfactant (Fig. B.2b).

Emulsification was observed in all diets tested (PS, bran, ± antibiotic treatment), suggesting

that the production and secretion of an emulsifying agent is independent of both diet and

the gut microbiome, implying it must be endogenous to the mealworm itself. Emulsifying

surfactants are known to increase degradation rates of hydrophobic pollutants, including

plastics, where bioavailability of hydrophobic substrates is rating limiting.[6, 191, 112, 197]

It is possible that the supernatant acts similarly in the mealworm gut environment, coating

hydrophobic materials, such as plastic particles, with an emulsifying agent that increases

bioavailability and accelerates biodegradation.

To further quantify this emulsification activity, contact angle analysis was used to mea-

sure the ability of the supernatant to wet a hydrophobic surface (PS film) relative to DI

water as a control (Fig. 3.3). Increased surface wetting is analogous to increased hydropho-

bicity of the supernatant, which would implicate the supernatant as an emulsifying agent for

plastics. Whole supernatant from both PS- and bran-fed mealworms was more hydrophobic

than DI water (increased surface wetting on a hydrophobic surface, p < 0.001 for PS and

bran supernatant, Fig. 2a). When fractionated, the intermediate supernatant fraction (30 –

100 kDa) was significantly more hydrophobic than the other two fractions (<30 kDa, >100

kDa) for all four experimental diets: PS, PS + ab, bran, bran + ab (Fig. 3.3, PS: p < 0.0001

relative to both >100 kDa and <30 kDa; Bran: p = 0.002, p < 0.001 relative to >100 kDa,

<30 kDa respectively). In each case, the intermediate fraction was significantly better at

wetting the hydrophobic surface, as indicated by a distinctly larger smaller contact angle

(theta, �). This further confirms the emulsification activity is endogenous to the mealworm

itself and indicates that the emulsifying agent lies within the molecular weight range of 30

– 100 kDa, which will facilitate the identification of the factor.

This analysis reveals a significant role of the mealworm itself in the plastic biodegra-

dation process. The absence of emulsifying agents in ex situ microbial cultures could help

explain the low rate of PS biodegradation relative to in situ conditions. In addition to

enhancing plastic biodegradation by the gut microbiome, emulsifying agents could be a

valuable tool for remediation of other hydrocarbons. Further research into identify and

characterize this emulsifying agent is needed to maximize and exploit its potential utility.

3.4.3 Enhanced ex situ microbial growth

The insights from the previous sections were then applied to enhance the ex situ PS-

degradation using the extracted mealworm gut microbiome. Serum bottles were inoculated

with PS-fed mealworm gut microbiome in carbon-free media with PS microplastics added

as the sole carbon source. Cultures were supplemented with di↵erent fractions, tested sep-

arately and in combination as follows: <30 kDa, 30 – 100 kDa, <30 + 30 – 100 kDa, and

>100 kDa. Supernatant was supplemented at the previously determined ideal dose (0.6%

v/v) and CO2 production was measured over 10 days. When the supernatant fractions

were added separately, both the <30 kDa and 30 – 100 kDa fractions stimulated similar

increases in CO2 production (p � 0.99), both of which caused a significantly higher increase

than the >100 kDa fraction (p < 0.01 relative to both <30 kDa and 30 – 100 kDa, Fig.

3a). These results suggest that supplementation with either microbial secreted factors in

the <30 kDa fraction (Fig. 3.2) or mealworm-produced emulsifying agent in the 30 – 100

kDa fraction (Fig. 3.3) increases ex situ microbial respiration on plastic. When combined,

the <30 kDa + 30 – 100 kDa fractions promoted the highest increase in CO2 production

(p < 0.01 relative to both <30, 30 – 100 kDa, Fig. 3.4a). The only fraction from antibiotic

treated supernatant that increased activity was 30 – 100 kDa, further confirming that the

smallest molecular weight fraction provides no added benefit in the absence of the gut mi-

crobiome. These results demonstrate that the contributions of both the mealworm and its

gut microbiome are necessary to achieve enhanced plastic biodegradation.

To explore the long-term e↵ect of these respiration-enhancing factors on plastic biodegra-

dation, a microbial enrichment was inoculated with PS-fed mealworm gut microbiome in

carbon-free media supplemented with PS film (as the carbon source) and PS gut super-

natant. This mixed community enrichment was cultured for several months until a stable

biofilm formed (Fig. B.3). The PS respiration activity of the enrichment was then compared

to freshly extracted PS-fed mealworm gut microbiome to determine whether the enrichment

conditions selected for more e�cient plastic biodegrading microbes. This was done using

serum bottles seeded with inoculum (either the microbial community from the enrichment

or the freshly extracted gut microbiome) in carbon-free media, supplemented with 0.6%

v/v PS-fed mealworm gut supernatant and either PS film or PS microplastics as a carbon

source. Cultures were grown for 14 days. The microbial enrichment exhibited significantly

higher PS respiration activity (CO2 production) on both PS microplastics and PS film,

suggesting that the enrichment was selecting for more e�cient plastic degrading bacteria

than those found in the gut microbiome (Fig. 3.4b). This enhanced plastic biodegradation

activity was especially prominent when comparing the CO2 produced from respiration on

PS films; while the microbial enrichment produced slightly more CO2 on PS film than PS

microplastics, the gut microbiome produced significantly less CO2 on PS film (Fig. 3.4b).

This suggests that in addition to selecting for better PS-biodegrading bacteria, the con-

ditions of the microbial enrichment also selected for bacteria that thrive in biofilms, an

unsurprising result given that the microbial enrichment was cultivated with PS film and

formed a stable biofilm on its surface (Fig. B.3). These promising results highlight the

ability to extrapolate insights drawn from the mealworm gut environment (e.g., the addi-

tion of a gut supernatant supplement) to select for bacteria capable of enhanced plastic

biodegradation outside the mealworm.

To further understand the mealworm’s role(s) in PS biodegradation, supernatant from

the mixed community enrichment was subject to the same supernatant fractionation process

used for the mealworm gut supernatant. The enrichment supernatant fractions were then

assayed using the same serum bottle experiment described above (Fig. 3.4a). Unlike the

supernatant fractions from the PS-fed mealworm, only the smallest fraction (<30 kDa) from

the enrichment increased CO2 production. This provides additional confirmation that the

PS-degradation enhancing e↵ect of the 30 – 100 kDa supernatant fraction is due to factor(s)

secreted by the mealworm itself.

The e↵ects of PS degradation by the microbial enrichment were further assayed by mon-

itoring changes in functional groups indicative of oxidative attack and hydrophobicity of

PS films (via FTIR visualization and contact angle analysis), and mass loss. FTIR spectra

of the PS films incubated with the microbial enrichment revealed oxygen incorporation as

indicated by the appearance of peaks associated with R-OH stretch (3000–3500 cm-1) and

C-O stretching (1000–1300 cm-1), as well as C-H bending (700-750 cm-1) often associated

with monosubstituted benzene derivatives (Fig. 3.5a). Changes in the surface chemistry of

incubated PS films were further explored by measuring changes in contact angle of DI water

on control versus incubated PS films (Fig. 3.5b). These analyses revealed a significant dif-

ference between the surface hydrophobicity of the films (p < 0.001), with a marked increase

in surface wetting on the treated PS film. This is further evidence of changes in surface

chemistry of the PS film, including oxygen incorporation, which would facilitate increased

surface wetting by water. While the rate of plastic degradation observed (measured as the

percent of the initial PS mineralized to CO2) was still significantly slower than within the

mealworm gut (Fig. 3.5b), these results are an important first step towards higher rates

of plastic degradation outside the mealworm and suggest that further enhancements are

possible.

3.4.4 Identification of plastic degradation microbial community

Once the microbial enrichment formed a stable biofilm on the surface of the PS film, a

sample of the biofilm was collected, and streak-plated in order to isolate and identify strains

involved in plastic biodegradation. Sixteen individual representative colonies were randomly

selected for sequencing (”enrichment strains”). The same process was then repeated, and 24

individual representative colonies were randomly selected and grown overnight in carbon-

rich media. The cultures were then centrifuged, and the cultured bacteria transplanted

to carbon-free media with PS films, supplemented with PS gut supernatant at 0.6% v/v.

Cultures were monitored for biofilm development and changes in optical density as an

indication of growth and respiration on the PS film. After 14 days, media from each

isolate culture that showed signs of respiration on PS (16 total). To obtain isolated strains,

these cultures were plated, and representative colonies were picked for sequencing (“isolate

strains”).

Overall, eight unique bacterial species were isolated and identified (Fig. 3.6, Table

B.1). The identified species are predominantly aerobes or facultative anaerobes, supporting

previous findings that oxygen incorporation is critical for rapid plastic biodegradation.[18]

While the mixed enrichment and identified isolates included many of the same species,

they were not the same, suggesting that a di↵erent subset of species thrive in the mixed

enrichment than can survive alone on PS (Fig. 3.6). However, to fully understand this

shift, further information, such as that from next generation sequencing of the microbial

enrichment, is required.

Of the identified plastic-biodegrading strains, many are associated with the degradation

and bioremediation of hydrocarbons and polymeric substrates. In the mixed enrichment

(Fig. 3.6a), the most prevalent species was Citrobacter freundii, a bacterium known to

degrade long-chain polymers (e.g., tannic acid[94]), linear hydrocarbons (e.g., oil[77]), and

cyclic hydrocarbons (e.g., phenylenediamine,[131] 4-nitroaniline,[88] and pentachlorophenol[199]).

3.5. IMPLICATIONS 49

Further, Citrobacter sp. are significantly associated with gut microbiome in PS- and PE-fed

mealworms,[18] and several Citrobacter enzymes have been extracted and used for degrada-

tion of phenylenediamine including: 1,2-dioxygenase, dehydrogenase, alkaline phosphatase,

phenol hydroxylase, and acetate kinase.[131] These previously identified enzymes may also

be involved in PS biodegradation, suggesting possible directions for future research on the

enzymatic degradation of PS.

Of the identified isolates, the most prevalent strains are Serratia marcescens [120, 40, 2]

and Klebsiella aerogenes,[67, 153] both of which are associated with hydrocarbon degra-

dation and bioremediation. S. marscens, Klebsiella sp. (closely related to K. aerogenes),

and Pseudomonas aeruginosa (another identified isolate) were found in the gut micro-

biome of PS-degrading mealworms, but were incapable of PS degradation when cultured

in isolation.[177] This suggests that these strains rely upon mealworm-secreted factors to

enable PS biodegradation. P. aeruginosa is also known to degrade crude oil[41, 132] and

produce a rhamnolipid,[189] a biosurfactant that accelerates hydrocarbon degradation. In

addition, P. aeroginosa was recently isolated from the gut of PS-degrading superworms

(Zophobas atratus) and found capable of growth on PS alone.[89] Another identified iso-

late, Enterobacter asburiae, was able to degrade polyethylene after its isolation from the

gut microbiome of plastic-eating waxworms (Plodia interpunctella).[206] Stenotrophomonas

maltophilia, another identified species, is capable of cyclic hydrocarbon degradation,[31, 99]

and the rate of bioodegradation can be enhanced in the presence of surfactants, similar to

the results of this study.[15]

The results of this work suggest that bacterial strains already known for their ability

to degrade hydrocarbon xenobiotics are likely responsible for PS biodegradation. Further

investigation is required to optimize the conditions for biodegradation of PS and other

xenobiotics, and to elucidate the metabolic pathway(s) involved.

3.5 Implications

This work is the first to achieve ex situ cultivation of a mixed enrichment derived from

the mealworm gut microbiome for enhanced PS biodegradation. This was accomplished by

elucidating the respective roles of Tenebrio molitor and its gut microbiome. We demonstrate

a significant role for the insect in secreting an endogenous emulsifying agent, increasing the

bioavailability of PS, and enabling more rapid microbial attack. The gut microbiome was

also found to secrete factor(s) that enhance microbial respiration when PS is the sole carbon

source. Additionally, we identify eight unique bacterial species involved in PS-degradation,

many of which have previously been associated with the biodegradation of hydrocarbons and

other xenobiotics. Our results imply that naturally occurring components of the mealworm

gut – including those secreted by both the mealworm host and its gut microbiome – must

be present and work in concert with the microbiota themselves in order to achieve enhanced

plastic respiration and biodegradation. This work develops techniques for the ex situ growth,

investigations, and exploitations of plastic-degrading microorganism from the mealworm gut

microbiome, thereby providing guidance for future research and scale-up e↵orts.

Appendix B contains figures including supernatant dose response curves, images from emul-

sification assays, and images of stable biofilm formed in the microbial enrichment (Figs. B.1

- B.3). In addition, a table with details on the sequences isolates and most similar strains

from BLAST is available (Table B.1).

3.7. FIGURES 51

3.7 Figures

Figure 3.2: E↵ects of mealworm gut supernatant on ex situ gut microbiome growth on PS.(a) Respiration activity of the gut microbiome in Biolog MT2 plates was measured usingtetrazolium violet redox dye as a colorimetric indicator at absorbance: 595 nm. Bacteriawere cultured in carbon-free media supplemented with PS microplastics (carbon source)and supernatant fractions (<30 kDa, 30 – 100 kDa) from each experimental condition.Supernatant condition is indicated by color with antibiotic treated supernatant indicatedby ab. (b) Final concentration of CO2 respired (final – initial percent CO2 in the headspaceof serum bottles) by bacterial cultured supplemented with di↵erent doses of PS supernatant.Statistical significance is calculated relative to the gut microbiome not supplemented withsupernatant is indicated by: ⇤ for p 0.05, ⇤ ⇤ for p 0.01, ⇤ ⇤ ⇤ for p 0.001; nostatistical significance indicated by ns.

3.7. FIGURES 53

PS Supernatant Bran Supernatant

DI WaterPS Supernatant (PS Sup.)PS Sup. > 100 kDaPS Sup. 30 - 100 kDaPS Sup. < 30 kDaBran Supernatant (B Sup.)B Sup. > 100 kDaB Sup. 30 - 100 kDaB Sup. < 30 kDa

DI Water PS Supernatant PS Sup. 30 - 100 kDaPS Sup. >100kDa PS Sup. <30 kDa

PS + antibiotic Supernatant

Bran + antibioticSupernatant

Molecular Weight

(a) (b)

*** *** ****** *** *** *** ***

ns ns ns ns

Figure 3.3: Hydrophobicity of mealworm gut supernatant. Contact angle analysis on thesupernatant fractions from each of the mealworm diets, colored by mealworm diet for eachsupernatant and ordered by molecular weight fraction (<30 kDa, 30 – 100 kDa, >100 kDa,size gradient indicate below), legend on the right. DI water was used as a negative control.(a) Contact angle (� C) of the supernatant fractions from the PS and bran fed diets. (b)Contact angle (� C) of the supernatant fractions from PS and bran + antibiotic fed diets.(c) Representative photos of DI water, whole PS supernatant, and PS supernatant fractionson PS film. Statistical significance relative to the intermediate molecular weight fraction(30 – 100 kDa) fraction indicated by: ⇤ for p 0.05, ⇤ ⇤ for p 0.01, ⇤ ⇤ ⇤ for p 0.001;no statistical significance indicated by ns.

PS Sup. < 30PS Sup. 30 - 100

PS Sup. <30 + 30 - 100

PS Sup. > 100

PS + ab Sup. < 30PS + ab Sup. 30 - 100

PS + ab Sup. <30 + 30 - 100

PS + ab Sup. > 100

Enrich. Sup. < 30Enrich. Sup. 30 - 100

Enrich. Sup. <30 + 30 - 100

Enrich. Sup. > 100

Molecular Weight

5 10 15

O2 Gut Microbiome Control (no PS)

Gut Microbiome + PS MicroplasticGut Microbiome + PS Film

Enrichment Control (no PS)

Enrichment + PS MicroplasticEnrichment + PS Film

Figure 3.4: E↵ect of PS supernatant on microbial respiration. Microbial respiration onPS supplemented with supernatant. (a) Final percent change in CO2 produced (measuredin the headspace of serum bottles) by the gut microbiome supplemented with di↵erentsupernatant fractions. Supernatant fractions were tested separately and in combinationto assess the impact on respiration (final – initial percent CO2 in the headspace of serumbottles), colored by supernatant condition (PS in green, PS + ab in blue, enrichment inpurple) and ordered by molecular weight fraction (size gradient indicated below) legendon the right. (b) PS-degradation (represented by percent CO2 produced) of extracted gutmicrobiome (green) and cultured enrichment (purple) fed with PS microplastics (squares)or film (triangles), relative to control (no PS, circles). (c) Percent of initial PS mineralizedto CO2 over 14 days by experimental conditions described in b. Statistical significanceindicated by: ⇤ for p 0.05, ⇤ ⇤ for p 0.01; no statistical significance indicated by ns.

3.7. FIGURES 55

Gut Microbiome + PS MicroplasticGut Microbiome + PS Film

Enrichment + PS MicroplasticEnrichment + PS Film

500100015002000250030003500400050

Wavenumber (cm-1)

Enrichment PS FilmEnrichment PS FilmControl PS Film

OHRstretch

C Ostretch

out-of-planebending

Control PSFilm

Treated PSFilm

(b) (c)

Figure 3.5: Characterization of PS degradation by gut microbiome and cultured microbialenrichment through changes in polymer surface chemistry and mass loss. (a) FTIR spectra(in percent transmittance) of PS films after incubation with microbial enrichment (purple)and a microbial isolate (green) relative to control PS film, annotations show functionalgroups associated with key peaks based on wavenumber. (b) Contact angle analysis ofDI water on control PS film and PS film after incubation with the microbial enrichment.(c) Percent of initial PS mineralized to CO2 over 14 days in serum bottle experiments,colored by condition: extracted gut microbiome (green) and cultured enrichment (purple).Statistical significance calculated relative to control PS film indicated by: ⇤ ⇤ ⇤ for p 0.001.

n = 16 n = 16

Citrobacter freundiiKlebsiella aerogenesSerratia marcescensStenotrophomonas maltophilia

Pseudomonas aeruginosaEnterococcus faecalisEnterobacter asburiae

Bacillus thuringiensis

(a) (b)

Figure 3.6: Relative abundance of PS-degrading bacterial species identified by 16S rRNAsequencing of individually isolated colonies from (a) the mixed microbial enrichment (16representative colonies) and (b) pure cultured isolates capable of PS-degradation (16 total).Additional data on species and sequence information is available in Table B.1.

Chapter 4

The fate of

hexabromocyclododecane (HBCD),

a common flame retardant, in

polystyrene-degrading mealworms:

elevated HBCD levels in egested

polymer but no bioaccumulation

The results of this chapter were originally published in Environmental Science & Technology

in 2020, doi: 10.1021/acs.est.9b06501, reprinted here with permission, copywrite American

Chemical Society, 2020. Sahar H. El Abbadi and Uwakmfon A. Ibekwe helped design and

perform laboratory work with L. vannamei. Dr. Yeo-Myoung Cho assisted with develop-

ment of analytic methods. Dr. Wei-Min Wu contributed to the design of the study and

provided comments to improve the manuscript.

58 CHAPTER 4. FATE OF HBCD

4.1 Abstract

As awareness of the ubiquity and magnitude of plastic pollution has increased, so has interest

in the long-term fate of plastics. To date, however, the fate of potentially toxic plastic

additives has received comparatively little attention. In this study, we investigated the fate

of the flame retardant hexabromocyclododecane (HBCD) in polystyrene (PS)-degrading

mealworms and in mealworm-fed shrimp. Most of the commercial HBCD consumed by the

mealworms was egested in frass within 24 hours (1-log removal) with nearly a 3-log removal

after 48 hours. In mealworms fed PS containing high HBCD levels, only 0.27 ± 0.10%, of the

ingested HBCD remained in the mealworm body tissue. This value did not increase over the

course of the experiment, indicating little or no bioaccumulation. Additionally, no evidence

of higher trophic level bioaccumulation or toxicity was observed when L. vannamei (Pacific

whiteleg shrimp) were fed mealworm biomass grown with PS containing HBCD. Di↵erences

in shrimp survival were attributable to the fraction of mealworm biomass incorporated into

the diet, not HBCD. We conclude that the environmental e↵ects of PS ingestion need further

evaluation as the generation of smaller, more contaminated particles is possible, and may

contribute to toxicity at nanoscale.

4.2 Introduction

Plastic waste is a widespread environmental pollutant and a growing waste management

challenge. Polystyrene (PS), one of the five most common thermoplastics, is typically used

for packaging and insulation, and is among the least sustainable plastics.[3, 66] PS foams are

low density and bulky, making them di�cult and costly to transport and recycle.[66] Because

recycling centers typically do not accept PS wastes, they are generally landfilled or escape

into the environment where they persist and can accumulate due to their recalcitrance.[66]

Chemicals added to improve manufacturing properties (e.g., plasticizers or stabilizers)

or decrease flammability, for example in insulation in the case of PS, pose an additional

sustainability concern.[3, 7] Because these additives are not covalently bonded to the poly-

mer, they could potentially partition into fatty tissues and bioaccumulate in food chains or

the environment.[7, 11] For PS, the most common flame retardant is commercial hexabro-

mocyclododecane (HBCD).[11, 157] HBCD is hydrophobic, lipophilic, persistent in the en-

vironment, and has been known to bioaccumulate in marine organisms.[11, 36, 80, 215] It

is also an endocrine disruptor and potentially neurotoxic.[11, 157, 36, 129, 156] Due to its

toxic nature, HBCD has been the subject of regulatory action in the European Union (to

be phased-out and eventually banned)[174] and in the United States, HBCD is currently

the subject of a risk evaluation by the EPA.[176] Solutions are needed to address HBCD,

and other chemical additives, in PS waste.[16]

Biodegradation is of increasing interest for management of PS and other plastic wastes.

While researchers have isolated many plastic-degrading microorganisms (e.g., fungi and bac-

teria), rates of biodegradation are typically low.[158, 198, 161, 65] For plastic materials, one

rate-limiting factor is access to surface area.[93] Insects that chew natural polymers, such as

lignin and wax, convert macro-scale fragments into micron-scale particles, increasing spe-

cific surface area accessible to attack by secreted enzymes.[127] Within the insect gut (e.g.,

Tenebrio molitor,[18, 210, 212, 207] Plodia interpunctella,[206] and Galleria mellonella[91])

secreted enzymes and ingested particles are concentrated and co-located, enhancing rates

of biodegradation. Of the insects tested to date, mealworms (T. molitor) are the most

well-studied and can rapidly eat and degrade PS and polyethylene waste, with half-lives for

plastic conversion to CO2 on the order of 15 – 20 hr.[18, 210, 212, 207] For chemical fate and

e↵ect studies, mealworms are an attractive insect model: they are readily cultivated; have a

rapid life cycle; and are prey for other animals, enabling assessment of food chain impacts.

In addition to their plastic-degrading ability, mealworms are farmed as a valuable protein-

rich[84] feed supplement for birds and reptiles as well aquaculture (e.g., shrimp, prawns,

sea bass).[76, 10, 130, 64, 32, 58] Further, mealworms are being considered as a more sus-

tainable, lower greenhouse gas emitting source of edible protein for humans.[128, 159, 119]

Conversion of plastic wastes into a valuable feed-supplement is appealing, but the fate

of plastic additives, such as HBCD, must first be understood. If HBCD accumulates in the

tissues of mealworms, there is a risk of food chain contamination and bioaccumulation. On

the other hand, if HBCD passes through the mealworm, there is risk of concentration within

the remaining undegraded PS particles in the frass, creating a need for further remediation

of these particles, such as frass pyrolysis.[208, 209]

In this study, we investigated the fate of HBCD within PS-degrading mealworms using

a mass balance and body burden analysis to assess toxicity and bioaccumulation. We also

evaluated HBCD bioaccumulation at a secondary trophic level by feeding PS-fed mealworm

biomass to a model aquaculture organism, L. vannamei (Pacific whiteleg shrimp).

4.3.1 Plastic test materials

To determine the fate of HBCD in PS-degrading mealworms, two commercially available

expanded polystyrene foam packaging materials were acquired from local vendors. The

first PS foam was a commercial insulation material with a high concentration of HBCD,

⇠0.25 % [w/w] (2385.5 ± 353.0 µg HBCD/g PS) and was utilized as the PS material

high in HBCD (“PS-H”).[145] The second expanded PS foam was a commercial packing

material that contained only trace amounts of HBCD, ⇠ 8.0 x 10-5 % [w/w] (0.83 ± 0.20

µg HBCD/g PS) and was utilized as the PS material low in HBCD (“PS-L”). Table C.1

includes additional descriptive characteristics (e.g., molecular weights, density) of the two

PS foams used in this study, demonstrating that the concentration of HBCD is the main

di↵erence between them. The PS foam blocks were cut into irregular 2 – 3 cm cubes and

cleaned with a stream of air to remove any fine residues prior to being weighed and utilized

in experiments following established methods.[18, 210, 212, 207]

4.3.2 Mealworm growth conditions

Mealworms, larvae of T. molitor Linnaeus, (average weight 75-85 mg/worm) were purchased

online from Rainbow Mealworms (Compton, CA) and shipped overnight to the laboratories

at Stanford University. Prior to arrival, the mealworms were fed bran; after arrival, they

were subject to a 48-hour depuration (starvation) period before initiating experimental

diets.[18] Natural wheat bran used as a control diet[18, 210, 212, 207] was purchased from

Exotic Nutrition (Newport News, VA).

Five experimental diets were compared, in duplicate: PS-H, PS-H + bran (1:1 [w/w]),

PS-L, PS-L + bran (1:1 [w/w]), as well as a bran only control.[18, 210, 212, 207] Each repli-

cate (in 475 mL food grade polypropylene containers) started with 200 randomly selected

mealworms along with their respective feeds (Fig. C.1a). Plastic-fed containers began with

1.80 g of polystyrene cut into 2 – 3 cm cubes following established methods.[18] Containers

that received bran started with 1.80 g of bran, with bran added every 3 days to main-

tain a 1:1 ratio [w/w] of plastic to bran, including the bran-fed control.[18, 210, 212, 207]

Containers were stored in incubators maintained at 25 �C and 70% humidity.[18, 212]

Every 3 days, mealworm frass (excrement) in the containers was collected and weighed

and mealworm survival was evaluated for the duration of the 32-day experiment. Once

a week, the mealworms were cleaned with a stream of air to remove any residual plastic

fragments and transferred to a clean container to collect frass for HBCD analysis. After

24 hours, the mealworms were returned to their original container, and the frass samples

were weighed and stored at -20 �C. In addition, 10 mealworms from each container were

sacrificed and dissected (gut track and non-gut tissues) as representative weekly biomass

samples, which were then freeze-dried for > 72 hr prior to being stored for future analysis

(Fig. C.1a, SI Method C.1.2).

To prepare su�cient mealworm biomass for aquaculture feeding experiments, 500 meal-

worms (from the same order) were raised in larger “bulk-fed” food grade polypropylene

containers (volume 780 mL) on either PS-H, PS-L, or bran diets for 32 days following the

conditions described above (Fig. C.1b). At the end of 32 days, the mealworms were sac-

rificed and the biomass (all the mealworm tissues) were freeze-dried for > 72 hr prior to

being incorporated in aquaculture feed (SI Method C.1.3).

Depuration tests were conducted to investigate accumulation of HBCD in mealworm

tissues; full details are described in the Supporting Information (SI Method C.1.4). Briefly,

mealworms were fed either PS-H or PS-L for 72 hr and were then subject to depuration for 48

hr. Ten mealworms were collected at 0, 24, and 48 hr of depuration, in triplicate, for HBCD

analysis. The expected intake for each PS diet was calculated (SI Method C.1.4). Bran was

spiked with HBCD to achieve comparable expected intake as PS-H and PS-L, referred to

as Bran-H and Bran-L, respectively, to assess the e↵ects of PS on the bioaccumulation or

egestion of HBCD.

4.3.3 HBCD quantification

Full details of the HBCD analysis are described in the Supporting Information (SI Method

C.1.2, Fig. C.3). Briefly, analytical methods to quantify total HBCD were developed

following modified EPA methods using extraction via ultrasonication (sample extraction:

3350B[180]; sample cleanup: 3630C[178]) on lyophilized biomass or PS samples. Quan-

tification of total HBCD was performed using an Agilent gas chromatograph[188, 73, 102]

(model 6890) equipped with a micro electron capture detector (GC-µECD) following modi-

fied EPA methods 8082[182] and 8081.[181] Surrogate standards (a mixture of PCBs, BZ#

14, 65, 166; 20 µL of 400 µg/L) were used to assess recovery e�ciencies (Fig. C.6, accepted

range: 40 – 100%) and internal standards (a mixture of PCBs, BZ# 30, 204; 10 µL of

400 µg/L) were utilized for quantification.[215, 102] Standards of the three main stereoiso-

mers were tested independently and as a mixture. Retention times and response factors

of the three stereoisomers were statistically indistinguishable (Fig. C.8), justifying the use

of one HBCD signal to quantify total HBCD.[73, 102] Method detection limit (MDL) for

mealworm biomass samples was 0.20 ng HBCD/g dry weight and for shrimp biomass the

MDL was 1.00 ng HBCD/g dry weight. Our limit of detection (LOD) and limit of quanti-

tation (LOQ) were determined to be 40 pg and 1000 pg, respectively, based on established

definitions.[5, 116] To increase the robustness of our analysis, we do not utilize any values

below the MDL for statistical quantification, but we plot all values above our LOD.

4.3.4 Aquaculture experimental conditions

Postlarvae of L. vannamei were used as a model aquaculture organism. L. vannamei were

reared for two experimental tests. In the first, postlarvae were placed in individual contain-

ers to assess toxicity of experimental feeds through daily survival monitoring. The second

test evaluated total HBCB bioaccumulation. Postlarvae were grown in larger containers

with the same experimental diets to enable collection of su�cient biomass for HBCD analy-

sis (Fig. C.2). The experimental diets included bran-fed mealworm biomass (MWB), PS-L

fed MWB, PS-H fed MWB; each type of MWB was integrated into the shrimp diets at 3 dif-

ferent rates: 10%, 50%, and 100% w/w (for a total of 9 experimental diets). Control shrimp

were fed commercially available feed (FRiPPAK RW+500, INVE Aquaculture, Deception

Bay, Queensland, AUS). Detailed experimental conditions used to assess bioaccumulation

studies in aquaculture are available in the Supporting Information (SI Method C.1.3, Fig.

Statistical analyses were performed in Prism (GraphPad Software, version 8.1.1). To as-

sess di↵erences in plastic consumption and pupation rates, one-way ANOVAs were per-

formed, followed by pairwise comparisons using Student’s t-test with Bonferroni multiple

test correction to assess di↵erences between diets. Di↵erences in survival was assayed using

Kaplan-Meier survival analysis. All p-values are adjusted p-values and all error values are

average ± standard deviation.

4.4.1 E↵ects of HBCD on mealworm survival and plastic consumption

Mealworm survival rate was una↵ected by amount of total HBCD consumed in PS over the

course of the experiment (Fig. 4.2a,b). There was no significant di↵erence in final survival

curves among the experimental diets nor the control diet based on Kaplan-Meier survival

analysis (Fig. 4.2a). However, a pair-wise analysis of the final survival by diet found that

mealworms fed PS-L + bran had a significantly higher survival than mealworms fed PS-L

alone (p = 0.02). No other significant di↵erences in survival based on the inclusion of bran

in the diet were found (PS-H vs. PS-H + bran, PS-H vs. bran, PS-L vs. bran). Previous

studies have reported di↵erences in survival based on inclusion of bran.[210] Further research

is needed to determine why di↵erent types of PS may result in di↵erential survival outcomes

among mealworms when co-fed with bran. Importantly, a linear regression analysis shows

that total HBCD exposure from each diet was not a significant determinant of final survival

(Figure 1b, R2 = 0.13).

Mealworms consumed a similar amount of either type of PS, showing that higher con-

centration of HBCD in the PS did not a↵ect mealworm consumption patterns (Fig. 4.2c).

Mealworms fed plastic plus bran consumed significantly more plastic than those fed PS

alone, a trend established in previous studies[18, 210] (Fig. 4.2c). The increase in PS con-

sumption in bran-fed mealworms was not as large as previously reported,[18, 207] perhaps

due to feedstock di↵erences. Overall, mealworm plastic consumption rates in all experimen-

tal diets are comparable to those of previous studies.[18, 210, 207]

Mealworm pupation is a hormonally controlled process[150, 151] that is known to be

impacted by endocrine disrupting chemicals.[44, 162] Therefore, if HBCD (an endocrine

disrupting compound[157, 43]) were bioaccumulating in mealworm tissues, we would hy-

pothesize observable di↵erences in the rates of pupation across experimental diets on the

basis of HCBD exposure. However, our results indicate that the pupation rate was not

a↵ected by HBCD exposure (Fig. 4.2d). The major di↵erence in the rate of pupation

appeared to be driven by the inclusion of bran in the diet, rather than HBCD. Both PS

diets that included bran (PS-H + bran, PS-L + bran), had a comparable pupation rate as

the control group (Fig. 4.2d). This finding was further explored with a multiple regres-

sion model to assess the e↵ects of bran, plastic, and HBCD consumption on pupation rates

(Fig. C.4, model R2 = 0.95). Bran consumption was found to be a significant predictor of

pupation rate (p < 0.0001) while HBCD consumption was not (p > 0.05). This suggests

that di↵ering rates of pupation are driven by the added nutritional benefits from bran in

the diet rather than any e↵ect from HBCD.

In summary, these results demonstrate that HBCD in commercial PS was not toxic to

mealworms and did not a↵ect mealworm survival and pupation behavior. Taken together,

these results suggest that total HBCD does not bioaccumulate within mealworm tissue.

This finding was further explored through analysis of mealworm tissue.

4.4.2 Exponential removal of HBCD from mealworm biomass

Depuration tests post feeding of PS were conducted to investigate accumulation of HBCD

in mealworm tissues over time periods that were comparable to the retention time of PS in

the mealworm gut (15 – 20 hr).[18] This analysis revealed that HBCD was rapidly excreted

from the mealworm body (whole body tissues, including the gut track) when subject to

depuration (i.e., starvation) (Fig. 4.3). The HBCD concentration from both types of PS

(PS-H and PS-L) decreased significantly over 48 hr, in good agreement with a one-phase

exponential decay model (Fig. 4.3, R2 = 0.96 for PS-H, 0.93 for PS-L). From the fitted

decay models, the half-lives for HBCD in the mealworm gut was 5.3 and 7.6 hr for S-H and

PS-L, respectively. The decay rate constant (k) for PS-H is 0.13 hr-1 and for PS-L is 0.09

hr-1. This suggests that HBCD from both types of plastic are subject to similar processes

within the gut and are removed at comparable rates. After 24-hours of starvation, there was

a 1-log reduction in the HBCD body burden (ng HBCD/g dry weight of mealworm tissue)

from both types of plastic (Fig. 4.3). After 48-hours of starvation, tissue from mealworms

fed either type of PS experienced a nearly 3-log reduction in HBCD body burden (Fig. 4.3).

There was not a significant di↵erence in the log-reduction of HBCD between the PS-H and

PS-L diets at either time point (Fig. C.9).

To assess the impact of the presence of polystyrene on the egestion of HBCD, mealworms

were fed bran directly spiked with HBCD at a comparable expected intake of HBCD as

PS-H and PS-L referred to as Bran-H and Bran-L, respectively (SI Method C.1.4). The

concentration of HBCD in mealworm tissues were measured over the same depuration period

as mealworms fed PS (at 0, 24, and 48 hr) and were then fit to a one-phase decay model

(Fig. 4.3, R2 = 0.88 for Bran-H, 0.94 for Bran-L). The decay rate constants (k = 0.05

hr-1 for Bran-H and 0.04 hr-1 for Bran-L) and half-lives (13.7 hr for Bran-H and 16.0 hr

for Bran-L), indicate a significantly slower egestion of HBCD than mealworms fed PS. The

log-reduction of HBCD from both Bran-H and Bran-L diets was significantly lower than

the reduction of HBCD from either PS diet (Fig. C.9). These findings suggest that the

presence of PS within the gut may play a significant role in concentrating the HBCD in the

frass, leading to its egestion.

These findings suggest that the majority of the HBCD is transient and that when meal-

worms are subject to depuration periods longer than the gut retention time, HBCD passes

through the gut and is egested when PS is present. The rates of HBCD egestion are not

significantly di↵erent between PS-H and PS-L, suggesting that HBCD behaves similarly in

either PS diet. Previous studies of mealworm PS-degradation have demonstrated that a

fraction of the PS is egested in the frass as partially degraded polymer.[18, 212, 207] Given

the lipophilic nature of HBCD[157] and the rapid egestion of HBCD in mealworm tissues,

this could suggest that the HBCD is predominantly partitioning to and thus concentrating

within the egested PS polymer residues. The increased half-life of HBCD delivered without

PS (spiked into bran) further supports this hypothesis. Taken together, our findings suggest

that the presence of depolymerizing PS plays an important role in concentrating HBCD and

removing it from the mealworm; however, further work is needed to confirm the role of PS

in the egestion of HBCD.

4.4.3 Fate of HBCD in mealworm biomass over time

To further assess whether any of the ingested HBCD bioaccumulates in the mealworm

tissues, a mass balance was conducted to monitor the fate of HBCD in various mealworm

tissues over time. The amount of HBCD that entered the mealworm (based on consumption

of PS), the amount of HBCD that left the mealworm system (measured in the frass), and

the amount of HBCD remaining in mealworm (gut and non-gut tissues) was determined

on a weekly basis. This approach enabled the tracking of HBCD within the mealworms

over the course of the 32-day experiment to assess bioaccumulation (Fig. C.7). The results

indicate that the majority of the HBCD consumed by mealworms, fed either type of PS, is

egested in their frass (Fig. 4.4a,b). On average, of the amount of HBCD consumed from PS

on a weekly basis, 93.20 ± 1.45% was egested in PS-H fed-mealworms and 92.56 ± 4.17%

was egested from PS-L fed mealworms. This trend was also observed in PS diets co-fed

with bran, where the total HBCD consumption was higher than those fed PS alone (Fig.

C.5a,b). This further supports evidence that HBCD is rapidly excreted from mealworms

(Fig. 4.3). Moreover, while total consumption of HBCD varied over the course of the

experiment (as PS consumption by the mealworms varied), the HBCD recovered in frass

remained consistently high.

The weekly HBCD body burden analysis, based on 10 randomly selected representative

mealworms pooled together from each container, showed that little HBCD remained in the

mealworm tissues (Fig. 4.4c,d). Based on total PS weekly consumption and number of

mealworms per container, the expected weekly intake of HBCD (EWI, normalized to 10

mealworms) was calculated and compared to the amount of HBCD in mealworm tissues.

For mealworms fed PS-H, on average only 0.29 ± 0.11% of the EWI was found in the

mealworm gut (Fig. 4.4c). The representative mealworms were first subjected to a 24-hour

depuration, which from analysis in Fig. 4.3 suggests enables ⇠ 90% removal of transient

HBCD, explaining why trace amounts of HBCD may still have been detected within the

gut track. For the same mealworms fed PS-H, on average only 0.27 ± 0.10% of the EWI, or

25.6 ± 8.7 ng HBCD g-1 dry weight, was found in the non-gut tissues of the mealworms; this

amount remained consistently low throughout the experiment (Fig. 4.4c). For mealworms

fed PS-L, the HBCD concentration of the majority of body tissue samples were below

the method detection limit (MDL, 0.2 ng HBCD/g dry weight) and thus were not used

for quantification (plotted in Fig. 4.4d). The HBCD detected in the gut tissues is elevated

compared to the HBCD in the non-gut tissues, which again, was likely due to the limited (⇠90%) removal of transient HBCD during the 24-hour depuration period. For mealworms fed

either type of PS, the levels of HBCD in the non-gut tissue remained stable over the course

of the experiment, suggesting that HBCD was not bioaccumulating, as bioaccumulation

would likely have resulted in an increase in HBCD concentration in non-gut tissues over

Similar trends were observed in the PS plus bran co-fed diets (Fig. C.5). Previous

studies have established that mealworms co-fed bran gain more weight over the course of

the experiment than those fed PS alone.[18, 207] Despite the increased growth and metabolic

activity of mealworms fed these diets, mealworm tissues still only contained trace levels of

HBCD (0.24 ± 0.04% and 0.15 ± 0.12% of EWI for PH-H + bran remains in the gut and

non-gut tissues, respectively). As observed in the PS-L group, the HBCD concentration in

the tissues of mealworms fed PS-L plus bran were below the MDL, suggesting little or no

bioaccumulation. This also suggests that bioaccumulation of HBCD was not a↵ected by an

increase in metabolic activities.

Overall, the rapid excretion of HBCD from the gut and the consistently low body burden

analysis in all diets over the duration of the experiment suggest that HBCD is not bioaccu-

mulating in mealworm tissues. The mass balance instead demonstrates that total HBCD is

concentrating in the frass egested by the mealworms. Further research is needed to better

understand the mechanism driving rapid egestion of HBCD and to assess the environmental

impacts of HBCD within the smaller, more concentrated particles in the frass.

4.4.4 Lack of HBCD bioaccumulation in secondary trophic level

Mealworm biomass represents a valuable supplemental protein source in many agricultural

and aquaculture feeds.[76, 10, 130, 64, 32, 58] Therefore, we investigated the toxicity of

HBCD-exposed mealworm biomass (MWB) on the growth and survival of a model aqua-

culture organism, Pacific whiteleg shrimp (L. vannamei). Further, we investigated whether

HBCD from MWB was bioaccumulating through a body burden analysis of the shrimp

biomass. Mealworm biomass was incorporated into experimental shrimp diets both at a

realistic levels (10%)[76, 64, 136] and at higher level to assess the upper bound for HBCD

bioaccumulation and toxicity.

Our results from the toxicity experiments showed an increase in mortality with increasing

incorporation of MWB (Fig. 4.5a). The increase in mortality was observed for MWB from

mealworms fed PS-H as well as mealworms fed PS-L and bran-fed mealworms (Fig. 4.5a).

This suggests that changes in shrimp survival are driven by total amount of mealworm

biomass included in the diet, not by HBCD. The amount of MWB included in the shrimp-

feed, regardless of the diet of the mealworms, explained > 70% of the di↵erence in survival

(Fig. 4.5c, R2 = 0.73), with a greater inclusion of MWB leading to an increase in mortality.

This suggests that MWB of any type (bran, PS-H, or PS-L) a↵ects mortality at higher

inclusion rates, likely due to lack of nutrition, rather than presence of HBCD. Shrimp

mortality in this study was higher than has been previously observed when supplementing

L. vannamei diet with MWB.[130] Unlike previous work, we studied postlarvae, which are

more sensitive than adult shrimp,[218] likely explaining the higher mortality rate. Increases

in weight in the L. vannamei over the course of the experiment also appear to be driven by

percent of MWB incorporated in the diet rather than HBCD exposure (Fig. 4.5b).

To further confirm that total HBCD was not bioaccumulating in the shrimp, body

burden analysis was conducted on shrimp fed the same experimental diets, excluding the

high mortality 100% MWB diets, for two weeks (Fig. C.2b). The HBCD concentration

in all shrimp biomass including those fed PS-L and PS-H MWB were below the MDL and

not significantly di↵erent than zero. Moreover, the concentration of HBCD in the shrimp

biomass was una↵ected by the experimental diet (bran-fed, PS-H fed, or PS-L fed MWB).

Taken together, these results suggest no significant bioaccumulation of HBCD in the shrimp

fed mealworm biomass. While the starting concentration of HBCD in the shrimp feed was

likely relatively low (given the 24 hr depuration before collection of MWB), this case-study

demonstrates that that inclusion of PS-fed mealworm biomass under realistic aquaculture

growth conditions does not lead to the bioaccumulation of total HBCD.

4.5 Implications

This is the first study to assess the fate of chemical additives in a plastic biodegradation

system. This is important because chemical additives, such as HBCD, can have significant

environmental and health impacts. We demonstrated that total HBCD does not bioac-

cumulate in PS-degrading mealworms, but is instead egested in the frass where it likely

concentrated in the partially degraded PS. This study did not investigate the fate of in-

dividual stereoisomers of HBCD, such an analysis would not change the conclusions of

this study. Nevertheless, future research should assess whether the distribution of isomers

egested in the frass are di↵erent than those in the initial PS.[168]

This study serves as a proof-of-concept for deriving valuable biomass from plastic waste.

Through the egestion of HBCD, the proteinaceous mealworm biomass is preserved as a

potential resource, which in this study we demonstrated could be used as an aquaculture

feed supplement without leading to bioaccumulation at a higher trophic level. However,

only one chemical additive for one type of plastic was investigated. Other common chemical

additives (e.g., plasticizers, stabilizers, pigments, other types of flame retardants)[3, 7] have

variable chemical properties (e.g., hydrophobicity), which will result in a di↵erent fate within

plastic-degrading mealworms. This study lays the foundation for additional investigations

into the fate of plastic additives within plastic degrading systems. Future work should focus

on whether these findings are generalizable to other plastic additives and in other systems

that degrade plastic.

This research shows that in working to address the issue of plastic pollution, it is im-

portant to avoid creating a new concern as residual plastic particles become smaller and

smaller, increasingly concentrating hydrophobic contaminants in the particles. Future re-

search should focus on the environmental impacts of such particles. Further, this research

demonstrates the need for biodegradable plastic replacement materials and green chemistry

to ensure that future materials and additives are non-bioaccumulative and non-toxic.

4.6. SUPPORTING INFORMATION 71

Appendix C contains detailed methods on HBCD chemical analysis and quality assurance

measures, mealworm and aquaculture growth conditions, and sample preparation informa-

tion as well as Table C.1 and supporting Figs. C.1 – C.9.

4.7 Figures

4.7. FIGURES 73

0 10 20 300

BranPS-HPS-H + BranPS-LPS-L + Bran

✱✱

03×102

6×1029×102

1.2×106

1.5×106

1.8×10680

HBCD Exposure (ng)

Bran OnlyPS-H OnlyPS-H + Bran (1:1)PS-L OnlyPS-L + Bran (1:1)Linear Regression,R2 = 0.13

PS-H + Bran PS-L

PS-L + Bran0.0

✱✱

(a) (b)

(c) (d)

Figure 4.2: Survival rate and HBCD consumption by T. molitor. (a) Survival rate of meal-worms over 32-day experiment, significance analyzed using Kaplan-Meier survival analysis.(b) Scatter plot and linear regression (black line) with 95% confidence interval (grey lines),of final mealworm survival versus HBCD exposure, by diet. HBCD exposure was calculatedfor each experimental container as total plastic consumed over the course of the experiment[g] ⇥ the concentration of HBCD in the plastic [ng/g]. (c) Plastic mass loss [g] at the end ofthe experiment, used in the HBCD exposure calculation above, by diet. (d) Total numberof larvae that pupated over the course of the experiment by diet. All values represent meanSD, n = 2. Significance (one-way ANOVA, Bonferroni multiple test correction, for all testsnot already described) p 0.05 indicated by ⇤, p 0.005 indicated by ⇤ ⇤, no statisticalsignificance indicated by ns.

0 10 20 30 40-2000

/ g dw

Bran-H

PS-H decay model R2 = 0.96

Bran-Hdecay modelR2 = 0.88

expected intake

0 10 20 30 40-5

Bran-L

PS-L decay modelR2 = 0.93

Bran-Ldecay modelR2 = 0.94

expected intake

(a) (b)

Figure 4.3: Reduction in T. molitor whole-body burden of HBCD with starvation. Bodyburden [ng HBCD/g dry weight] over time (points) plotted with one-phase exponentialdecay model (solid lines) and 95% confidence interval (dashed lines) (a) for PS-H and forcontrol bran spiked with HBCD at a comparable concentration (b) for PS-L and for controlbran spiked with HBCD at a comparable concentration. From the models, the half-lives forHBCD in the mealworm gut are 5.3 and 7.6 hr for PS-H and PS-L. The half-lives for HBCDspiked directly into control bran are 14 hr and 16 hr for Bran-H and Bran-L, respectively.The rate constant (k) is 0.13 hr-1 for PS-H, 0.09 hr-1 for PS-L, 0.05 hr-1 for Bran-H and0.04 hr-1 for Bran-L. All values represent mean SD, n = 3.

4.7. FIGURES 75

Week 1 Week 2 Week 3 Week 40.0

1.5×105

3.0×105

4.5×105

6.0×105

7.5×105

Week 1 Week 2 Week 3 Week 40

80009000

Consumed in PSEgested in Frass

Week 1 Week 2 Week 3 Week 40.00.10.20.30.40.50.63.0

/ g dw

Gut trackBody tissuesMDL

(a) (b)

(c) (d)

Figure 4.4: Fate tracking of HBCD in T. molitor fed PS-H (left) or PS-L (right). (a, b)Amount of HBCD consumed in PS (grey bars) coupled with the amount of HBCD egestedin frass (purple bars) from each container weekly (a: PS-H, b: PS-L). Amount of HBCDconsumed was calculated as total plastic consumed per week [g] ⇥ the concentration ofHBCD in the plastic [ng/g]). Amount of HBCD egested was calculated as the concentrationof HBCD in the weekly representative clean frass sample [ng/g] ⇥ the total frass collectedfor the week [g]. (c, d) Body burden of HBCD [ng/g dry weight] in mealworm tissues (gutor non-gut tissues) weekly, plotted with the method detection limit (MDL) (c: PS-H, d:PS-L). Expected weekly intake (EWI) of HBCD was calculated as the average amount ofHBCD consumed for each diet [ng/week], normalized to 10 mealworms, and is plotted forcomparison. All values represent mean SD, n = 2.

10% 50% 100%Percent MWB in Diet

✱✱

Initial WeightControl Diet

PS-L MWBPS-H MWB

Bran MWB

10% 50%Percent MWB in Diet

0 20 40 60 80 1000

MWB in Diet (%)

Linear Regression,R2 = 0.73

0.200.80

10% 50%

Percent MWB in Diet

Control DietBran MWBPS-L MWBPS-H MWB

(a) (b)

(c) (d)

Figure 4.5: Bioaccumulation e↵ects from plastic-fed mealworm biomass on L. vannamei. (a,c) Survival of L. vannamei from toxicity experimental set-up. (a) Final survival proportionsfrom Kaplan-Meier survival analysis by diet normalized to control diet and (c) scatter plotand linear regression of final survival versus percent of mealworm biomass (MWB) includedin the diet. (b, d) Analysis of L. vannamei from bulk experimental set up. (b) Final bodyweight (dry weight, dw) based on diet. (d) Body burden of HBCD [ng/g dry weight] inshrimp biomass at the end of the bulk experiment. All samples are below the methoddetection limit (MDL, plotted in dashed line), thus raw data are shown but not used forquantification. All values represent mean SD, n = 3-5 depending on diet. Significanceversus control (one-way ANOVA, Bonferroni multiple test correction) p 0.05 indicated by⇤, p 0.005 indicated by ⇤ ⇤, no statistical significance indicated by ns.

Chapter 5

The role of biotechnology for

control and remediation of plastic

pollution

This chapter was originally published in Current Opinion of Biotechnology as Can biotech-

nology turn the tides on plastic?, doi: 10.1016/j.copbio.2019.03.020, reprinted here with

permission, copywrite Elsevier B.V. 2019.

78 CHAPTER 5. TURNING THE TIDE ON PLASTICS

5.1 Abstract

Accumulation of plastic pollution in aquatic ecosystems is the predictable result of high

demand for plastic functionalities, optimized production with economies of scale, and recal-

citrance. Strategies are needed for end-of-life conversion of recalcitrant plastics into useful

feedstocks and for transition to materials that are biodegradable, non-bioaccumulative,

and non-toxic. Promising alternatives are the polyhydroxyalkanoates (PHAs), a vast fam-

ily of polymers amenable to decentralized production from renewable feedstocks. Estab-

lishment of a global-scale PHA-based industry will require identification of PHAs with

tailored properties for use as ‘drop-in’ replacements for existing plastics; use of low-cost

renewable/waste-derived feedstocks; high productivity cultures that may be genetically-

modified microorganisms or non-axenic mixed cultures maintained by selection pressures

that favor high PHA-producing strains; and low-cost extraction/purification schemes.

5.2 Highlights

• Current fossil carbon-based plastics are accumulating at an unsustainable rate.

• Insight into enzymatic depolymerization could enable new mitigation incentives.

• The ‘Ecocyclable’ framework provides a path for sustainable plastic design.

• Replacement plastics will require waste as feedstock, design for disassembly.

• Replacement plastics will need tailored functionality, e�cient bioprocesses.

5.3. CURRENT PLASTIC PRODUCTION RATES ARE NOT SUSTAINABLE 79

5.3 Current plastic production rates are not sustainable

Plastic production has grown exponentially with a 14-year doubling time since the 1950’s.[66]

A conservative linear increase in plastic production going forward is 10.8 million metric tons

per year (Fig. 5.2). This increase far outpaces the expected increase in recycling of ⇠ 0.8

million metric tons per year for 2018.[66] The inevitable result is cradle-to-grave accumula-

tion in landfills and ecosystems (Fig. 5.3, 5.4).[66] While landfilling sequesters carbon, space

for landfills is limited, especially in dense urban environments[74], increasing the probability

that plastic wastes will enter sensitive ecosystems. This situation will be exacerbated by

China’s recent decision to halt plastic waste imports as the country has managed ⇠ 45%

of all plastic waste generated since 1992.[20] Accumulation of plastic in aquatic ecosystems

already threatens wildlife health.[100] Human health may also be harmed by exposure to

plastic additives, especially via microplastics in food, such as table salt.[202, 205]

Plastic-contaminated urban waters warrant special attention because these water bodies

are both hubs of recreational and economic activity and delivery points for plastic and

microplastics from rivers and storm water outfalls.[46, 113, 167] One of the few economic

assessments of marine debris estimated that communities on the west coast of the United

States spend $13 per resident annually to clean up trash.[164] If extrapolated to the current

global coastal population (40% of 7.6 billion people), such an e↵ort would incur an annual

price tag of $40 billion. These prohibitive costs and the well-documented harm to ecosystem

health [100] are compelling incentives to develop sustainable alternatives.

Current strategies for controlling plastic pollution in urban waters rely on wastewater

treatment. In these systems, large plastic debris is removed by screening then landfilled; mi-

croplastics are mostly removed by attachment to biosolids that are then transferred to solids

handling facilities [195]; a small fraction remains in the treated e✏uent.[113, 167]. When

these biosolids are applied to land, the plastics can potentially re-enter aquatic ecosystems

via runo↵.[195]. Release of plastic to such environments could be avoided if microplas-

tics were designed to biodegrade in aquatic systems, such as wastewater treatment plants.

Researchers have isolated plastic-degrading microorganisms from landfills and petroleum-

production sites [214, 68, 201, 200], but, to date, the rates achieved by isolated pure cultures

have been low and almost all isolates are restricted to growth on a single type of plastic.[200]

Microorganisms and enzymes are needed that can more rapidly and e�ciently degrade com-

plex plastic mixtures.[200]

5.4 From tiny bioreactors to new mitigation strategies

Recent research has identified one environment in which structurally diverse plastics do

undergo rapid degradation: the gut of arthropods, specifically insect larvae of Tenebrio

molitor (mealworms) [212, 207, 18], Plodia interpunctella (Indianmeal moth) [206], and

Galleria mellonella (wax moths) [91]. The most well studied are the mealworms, with half-

lives for the conversion of ingested plastic to CO2 on the order of 15 � 20 hours, much

faster than rates observed for microbial isolates.[213]. The mealworm gut microbiome can

also degrade multiple types of plastics and mixtures (e.g., polyethylene plus polystyrene),

suggesting an initial non-specific oxidative attack.[18] A deeper understanding of the under-

lying molecular mechanisms, including the relative contributions of the insect itself [91] and

the gut microbiome [206, 213], could enable development of scalable enzyme-based strate-

gies for plastic waste management.[200]. Given that the products of mealworm degradation

contain oxidized functional groups [207, 18], it is possible that such metabolites could serve

as substrates for future production of polyhydroxyalkanoates (PHAs), converting an other-

wise recalcitrant plastic into one that is biodegradable and sustainable.[124] This possibility

has been demonstrated for polyethylene terephthalate (PET): enzymatic-mediated PET hy-

drolysis yields terephthalic acid and ethylene glycol [214, 198], both of which can serve as

feedstocks for PHAs.[87, 60]

Systems for plastic production and recycling have yet to fully address the fate of chem-

ical additives, such as fire retardants and plasticizers.[137] Strategies are needed to ensure

their removal from recycled materials and to develop green chemistry replacements.[12]

Within the proposed ‘Ecocyclable’ framework (discussed later), chemical additives within

new materials would be evaluated through tests of toxicology and bioaccumulation. Such

assessments could be informed by non-profit organizations that provide screening tools and

services to assist in selection of safe replacement chemicals (e.g., Clean Production Action,

[33]).

5.5 From linear to fully circular economies

Figure 5.3 illustrates the potential transition from recalcitrant plastics in the current linear

economy to a circular economy, in which renewable plastics displace conventional recal-

citrant plastics. The inputs for the linear path of recalcitrant plastics are fossil carbon

5.6. LOW-COST, TAILORED TO THE APPLICATION, AND ECOCYCLABLE 81

feedstocks that required millions of years to accumulate, and recycle loops are small; the

outputs either accumulate in landfills or the environment [66]. By contrast, renewable plas-

tics fit within a circular economy with large recycle loops and feedstocks that are renewable

over a short time (< 10 years).[48] In order to meet current plastic demands, a roughly

500-fold increase in renewable plastics production is required.[56] In addition, production

costs need to be decreased by a factor of 2 � 4 to achieve cost parity with recalcitrant fossil

carbon-derived plastics (Table D.1).

Renewable plastic will need to match the in-use functionality of current plastics while

also safeguarding the environment and enabling disassembly at end-of-life (Table D.1). To

that end, an ‘Ecocyclable’ framework has been proposed in which new materials are com-

pared to reference polymers in terms of degradability, susceptibility to bioaccumulation,

and toxicity (https://ecocyclable.wm.edu, [117]). The reference polymers are cellulose, the

most abundant organic polymer on Earth, and polyhydroxybutyrate (PHB), an ancient

and ubiquitous biopolymer that is synthesized and degraded within numerous Bacteria and

Archaea.[143] Within the Ecocyclable framework, biodegradability is assessed by compari-

son of test materials to either cellulose or PHB in aerobic soils, anaerobic (methanogenic)

conditions, and aquatic environments.[117] Materials deemed ‘Generally Ecocyclable’ biode-

grade in all three environments; materials deemed ‘Conditionally Ecocyclable’ biodegrade

in one or more environment, but not all three.

5.6 Low-cost, tailored to the application, and Ecocyclable

Figure 5.4 presents a transition sequence from polyethylene terephthalate (PET) to poly-

lactic acid (PLA) to polyhydroxyalkanoates (PHAs). The order of this sequence reflects

increasingly renewable, biodegradable, and ‘Ecocyclable’ materials. Unlike other fossil

carbon-derived plastics (e.g., polystyrene), PET is recyclable without downcycling and typ-

ically does not contain toxic additives. It is recalcitrant under all environmental condi-

tions and therefore not ‘Ecocyclable.’ As noted previously, PET is susceptible to enzyme-

mediated degradation [214, 198]; thus, future innovation could conceivably lead to a form

that is ‘Ecocyclable.’

PLA is currently the most common renewable plastic.[56, 49] It is made from lactic acid

typically obtained from cultivated crops and would be classified as ‘Conditionally Ecocy-

clable’ because it degrades in some environments (e.g., thermophilic composts), but not

others (e.g., marine environment).[117, 86] Copolymerizing PLA or creating PLA com-

posites with other sustainable materials can result in better material properties and in-

creased degradability,[49] A recent study demonstrated PLA depolymerization by microbial

carboxylesterases [70], paving the way for future enzymatic control strategies like those

suggested above for PET or other fossil carbon-derived plastics, potentially leading to a

‘Generally Ecocyclable’ PLA.

PHAs are a vast family of moldable biopolymers that could serve as ‘drop-in’ replace-

ments for persistent plastics.[217] Most PHAs are likely to be ‘Generally Ecocyclable’ as

they are close relatives of the PHB reference polymer, first described in Bacillus megaterium

as a homopolyester of 3-hydroxybutyric acid.[101] Interest in PHB surged following a 1958

report showing that B. megaterium and B. cereus use PHB as a carbon and energy storage

polymer.[109] Studies thereafter focused on PHA synthesis in Cupriavidus necator (for-

merly known as Hydrogenomonas eutropha, Alcaligenes eutropha, and Ralstonia autropha)

[97] and on the development of a genetic toolkit for recombinant Escherichia coli.[160, 30]

More recently, whole genome sequencing and phylogenetic tools have greatly expanded un-

derstanding the biodiversity of Bacteria and Archaea that produce PHA granules and its

synthesis pathways. These data can be mined to identify PHA-producing organisms with

special capabilities of interest. Examples include extremophiles, such as Haloferax mediter-

ranei, a model strain for PHA metabolism in Archaea [39], and Halomonas, a halophilic

gammaproteobacterium, both of which can be grown at high salinities, a feature that could

decrease costs for media and PHA recovery.[148]

Critical bioprocess challenges for PHA production are feedstock and production costs,

the need to tailor polymer structure for specific applications, maximizing productivity (g

PHA/L-h), and optimizing downstream polymer extraction and purification. Costs can be

decreased through economies of scale and use of waste-derived feedstocks.[154] Economies

of scale could be achieved in large biorefineries where gas pipelines deliver renewable feed-

stocks, such as compressed biomethane, CO2, H2, and syngas.[154, 123] Alternatively,

economies of scale might be achieved through a decentralized approach in which production

facilities are co-located near wastewater treatment plants, landfills, or food waste man-

agement facilities.[217, 154] Waste-derived feedstocks (e.g., H2, CO2, CH4, glycerol, fatty

acids, lignocellulose, and potentially future degradation products of recalcitrant plastic

waste) avoid competition with cultivated crops for land and the associated water, energy,

and fertilizer requirements.[154]

5.6. LOW-COST, TAILORED TO THE APPLICATION, AND ECOCYCLABLE 83

Tailored PHAs are needed because PHB is brittle and has a narrow thermal processing

window, limiting its utility as a replacement material for fossil carbon-derived plastics.[123]

Fortunately, blends and copolymers, such as poly(3-hydroxybutyrate-co-3-hydroxyvalerate)

(PHBV) and more than 150 PHA variants, have material properties that can rival recalci-

trant plastics (Table D.1).[143, 103] The structure and in-use properties of PHA co-polymers

can be adjusted by the choice of co-substrates [154, 123], application of selection pressures

on mixed cultures [123], and genetic manipulation of biochemical pathways and enzyme

optimization.[29] Four classes of PHA synthase are known [30], and crystal structures are

now available for two Class I synthases.[28, 90] Knowledge of PHA synthase structure can

provide deep insights into its mechanisms [28, 90, 118] and facilitate enzyme re-engineering

for production of a diverse suite of PHA co-polymers with di↵ering monomer lengths and

sidechains and even PLA.[211]

Both synthetic biology and application of feast-famine selection pressures to non-axenic

mixed cultures can be used to achieve high PHA productivity.[123, 29, 82] High rates of PHB

production have been achieved with recombinant E. coli strain grown on agricultural wastes

(productivity > 2.13 g PHB/L-hr) [125] and by Cupriavidus necator DSM 545 grown on

waste glycerol (1.36 g PHB/L-hr).[121] Typically, PHA is produced in two steps: in the first,

growth rates are maximized during a period of balanced growth, and cells grow under replete

conditions, that is, with su�cient carbon source/electron donor, electron acceptor (typically

oxygen), and nutrients; in the second, PHA accumulation is induced under conditions of

unbalanced growth where carbon/electron donor is present in excess and growth of the

culture becomes limited by a nutrient (e.g., N, P, K) or an electron acceptor (e.g., oxygen,

nitrate).[143, 123] Ultimately, cells are harvested at high density, which can exceed 100

g/L.[110] Further increases in productivity are possible through both pure culture synthetic

biology and mixed culture selection strategies: mutants that generate PHA constitutively

could eliminate the need for two-stage processing [85] and imposition of feast-famine regimes

in non-axenic sequencing batch reactors can select for competitive dominance by high PHA-

producing strains, such as Plasticicumulans acidovorans, a gammaproteobacterium that can

accumulate > 85% PHB in a second-stage fed-batch bioreactor.[92, 83]

E�cient processing is also needed for recovery of PHA from the cells and its purifica-

tion. Current techniques involve heat treatment or freeze-thaw to disrupt the cell membrane

followed by one of several treatments: (1) solvents plus centrifugation, (2) chemical disrup-

tion plus centrifugation, or (3) use of enzymes, surfactants, and other agents to break the

cells plus dissolved-air flotation.[154, 110] These methods are time, energy, and chemical-

intensive.[154, 110] Biotechnology has the potential to improve this step through genetic

modifications that enable secretion of PHA molecules into the extracellular media [142];

feeding PHA-enriched microbial biomass to animals such as mealworms that digest the bac-

terial cell-wall, releasing relatively pure PHA granules in the excreted fecal pellets [122]; and

use of halophilic Archaea to produce PHA followed by transfer of PHA-rich cells to deionized

water where osmotic pressure causes cell lysis and the release of the polymer.[146, 55]

5.7 Design for disassembly

Insights from phylogenetics and metagenome screening can potentially be harnessed to

expand end-of-life options. The principle of ‘design for disassembly’ [107] has been cham-

pioned for years at the macro scale; advances in bioengineering now o↵er similar potential

at nanoscale. The natural architecture of PHA granules o↵ers inspiration with PHB de-

polymerases in the granule surface allowing access to amorphous regions susceptible to

depolymerization.[81] It remains to be seen whether such features (e.g., plastic degrading

enzymes) could be added to improve the ‘Ecocyclability’ of new materials (e.g., Carbiolice,

which hopes to commercialize plastic resins pre-seeded with biodegradation enzymes [13]).

Further knowledge of underlying biochemical degradation mechanisms might enable the de-

sign of plastics that are susceptible to disassembly upon exposure to specific environmental

conditions, such as high salinity, yielding harmless, or even useful products.

5.8 Outlook

Biotechnology opens the door for improved plastic design in terms of its production, in-use

properties, and end-of-life fate. E�cient strains and mixed cultures, low cost feedstocks,

optimized fermentations, and improved extraction schemes can usher in a new generation

of materials that can function as drop-in replacements for existing plastics. Deeper under-

standing of the mechanisms of recalcitrant plastic degradation can inform design of new

materials, perhaps enabling development of new materials that are readily converted into

useful feedstocks.

Widespread adoption of Ecocyclable standards or their equivalent can enable nuanced

regulation of plastics use, reuse, and disposal while encouraging innovation. With increased

5.9. SUPPORTING INFORMATION 85

public awareness of the capabilities of biotechnology and enabling legislation, biotechnology

is poised to play an important role in the remediation of plastic pollution and development

of new plastics and improved systems for plastics management.

Appendix D contains a supporting Table D.1 with details on the cost, material properties,

and suitability for various thermal manufacturing processes for common petroleum-derived

plastics and bioplastics.

5.10 Figures

5.10. FIGURES 87

19401960

19802000

20202040

Historic Plastic ProductionProjected exponential growthProjected linear growthbased on 1988 - 2015

Figure 5.2: Projected increase in plastic production based on historic production data.Historic data are shown in blue circles. Dashed lines show projected growth. The reddashed line shows projected plastic production based on an exponential growth model (R2

= 0.98). The purple dashed line shows projected production based on a linear regression(R2 = 0.99) from the last three decades. Historic plastic production data from Geyer et al.2017 [66].

CO2 BIOMASS

PHOTOSYNTHESIS

1 - 10 YEARS

>106 YEARS

craddle-to-grave

FOSSILCARBON

FEEDSTOCKS

407 MILLION TONNES

(global annual production,2015)

79% ACCUMULATE IN LANDFILLS OR

THE ENVIRONMENT

12% INCINERATED

9% RECYCLED

2 - 3%PROCESS

LOSSES

5% DOWNCYCLINGLOSSES

1 - 2%CLOSED-LOOP

RECYCLING

ENVIRONMENTALDEGRADATION

leakage

circular lifecycles

MICROBIAL DEGRADATION(bioreactors)

880 THOUSAND TONNES

(global annual production,2017)

PLAPHBV

PHYSICAL / CHEMICAL RECYCLING(recovery of resin materials)

RENEWABLECARBON

FEEDSTOCKS

RENEWABLEH2

COMPOST

Figure 5.3: Current linear cradle-to-grave pathways for recalcitrant plastics and circularpathways of future plastic materials. Data for recalcitrant plastics from Geyer et al. 2017[66] and Ellen MacArthur Foundation 2016 [48]. Data for biodegradable plastics fromEuropean Bioplastics 2017 [56].

5.10. FIGURES 89

CRADLE-TO-GRAVE CIRCULAR LIFECYCLE

Polylactic acid (PLA) Polyhydroxyalkanoates (PHAs)

Feedstock:Lactic acid

Precursor(lactide)

Plastic resins

Plastic products

CO2cultivated feedstocks(e.g., corn)

photo-synthesis

abioticpolymerization

Feedstocks:CH4, CO2, H2, Sugars, Fatty acids, Glycerol

Precursors(e.g., alkenoates,

hydroxyalkanoates)

Plastic products

Feedstock:Crude oil, coal,

natural gas

Precursors(e.g., ethylene glycol,

terephthalic acid)

Plastic resins

Plastic products

accumulation in landfills

recycling

Fossil Carbon-Based Plastics (e.g., PET)

cycling

agricultural byproducts

bio-basedabioticpolymerization

manufacturing

accumulation in environment

litter + mismanaged

wastedisposal

Plastic resins

PHA granules

accumulation in landfills

accumulation in environment

bioticpolymerization

PHA recovery

degradation in bioreactors and/or environment

g) PHA producingmicroorganisms

manufacturing

litter + mismanaged

wastedisposal

agricultural byproducts

hydrolysis & therm

al processing

sustainabledisposal

fermentation (future) plastic deg.

products

meltrecoverymanufacturingmelt

recovery

hydrolysis & therm

al processing

Figure 5.4: A transition sequence for plastic materials. Black arrows show conventionalpathways. Purple arrows show existing biotechnology-based pathways. Orange arrowsshow future biotechnology pathways. Timeline at the bottom shows the transition in thedominant type of plastic used toward an increasingly renewable and degradable materials.

Chapter 6

Conclusions and Future Work

6.1 Conclusions

The goal of this dissertation was to investigate plastic biodegradation in the mealworm

gut as a model system for plastic degradation. To date, the guts of mealworms and of

other insect larvae, such as waxworms, are the only environments in which diverse plastics

biodegrade on a timescale of hours. As such, they are of great interest for identifying mech-

anisms and rate-limiting steps of plastic degradation, potentially enabling development of

new strategies for disassembly of plastics at end-of-life. Faster plastic degradation would

decrease plastic pollution and mitigate many of its negative impacts, a global priority high-

lighted by the United Nation Sustainable Development Goals, NOAA, and by the US EPA,

as a global priority.[166, 185, 175] If scaled-up, more rapid plastic biodegradation would

revolutionize plastic management by enabling resource recovery from materials that would

otherwise accumulate in increasingly limited landfill spaces or the environment. Current

limitations include the ability to broadly degrade multiple types of plastic, to e�ciently cul-

ture plastic-degrading organisms, and to understand and manage chemical additives within

plastics.

The work fromChapter 2 demonstrates that the mealworm gut is capable of nonspecific

plastic degradation. Experiments were designed to assess the specificity of degradation

within the mealworm by testing multiple types of plastic and mixed plastics. The results

demonstrated that mealworms are capable of degrading polystyrene and polyethylene, as

well as mixtures of the two wastes. This finding is especially important given the need to

scale-up degradation as a viable alternative for all plastic wastes. This research highlights

92 CHAPTER 6. CONCLUSIONS AND FUTURE RESEARCH

the need for additional mechanistic studies to identify how and why plastics readily break

down in this environment.

To better understand plastic degradation mechanisms in the mealworm gut, Chapter

3 investigates the role of the mealworm host and the gut microbiome. This work reveals

the importance of the mealworm host in secreting emulsifying factors that mediate plastic

bioavailability, thus enabling microbial attacks. Additionally, microbial secreted factors and

key microorganisms are identified. The synergistic interactions identified were applied to

develop a system whereby the plastic-degrading ability of cultured microorganisms from the

gut microbiome can be further investigated for scale-up e↵orts.

Chapter 4 investigates the fate of chemical additives within plastic waste by tracking

hexacyclobromododecane, the most common polystyrene flame retardant during biodegra-

dation by mealworms. Surprisingly, these results demonstrate that HBCD does not bioaccu-

mulate. Instead, it is rapidly excreted and concentrated within polymer residuals within the

frass, preserving the valuable, proteinaceous mealworm biomass for use as an animal feed-

supplement. This work serves as a proof-of-concept for the transformation of plastic wastes

into mealworm biomass or other products of value. It also highlights the need for thought-

ful green chemistry to ensure that plastic additives are non-toxic and non-bioaccumulative.

Such considerations are required to facilitate the adoption of circular practices that account

for the fate and removal of additives in plastic waste management systems.

The critical review presented in Chapter 5 contextualizes the research conducted in

this thesis in the broader field of biotechnology. It investigates the current state of plastic

waste and the need for biotechnology advances to reduce current and future pollution.

This review highlights the significance of the nonspecific and rapid plastic degradation that

occurs naturally within the mealworm. Further, this work proposes a vision for transitioning

from traditional recalcitrant plastics to biodegradable replacement materials and identifies

specific criteria needed for replacement materials to ensure their sustainability. Finally, this

review demonstrates the significance of biotechnology in tackling plastic waste challenges

and directs the prioritization of research to reach a more sustainable future.

6.2. RECOMMENDATIONS FOR FUTURE RESEARCH 93

6.2 Recommendations for future research

The work presented in this dissertation significantly expands our understanding of naturally

occurring plastic biodegradation mechanisms and identifies strategies whereby that under-

standing can be harnessed in engineered systems. It also highlights areas requiring further

research, such as features within insect guts that enable rapid plastic degradation, and en-

zymes and biochemical pathways that enable large-scale rapid biodegradation to recover

value from otherwise recalcitrant plastics. Increasing our understanding of how biotech-

nology can develop and tailor drop-in replacements for petroleum-based plastics and green

chemistry in support of a sustainable, circular economy.

Gut microbiome as a tool

This research contributes to growing evidence that the insect gut environment is a valuable

resource for the discovery of novel degradation patterns and unique microorganisms. Until

recently, only one insect gut was widely studied and modeled for its degradation abilities:

termites and the symbionts within their hindgut that degrade plant lignose.[22, 21, 19] This

work, along with a growing number of other studies, demonstrate how the gut of insects

such as mealworms and waxworms are uniquely suited for enhanced plastic degradation,

likely because of their prior adaptation to the consumption of natural polymeric substances.

Further research is needed to assess the commonality of these gut environments that enable

degradation of complex, persistent polymers. These insect guts may also serve as a previ-

ously untapped resource for other biotechnology tools of interest, such as the endogenous

emulsifying agent identified in Chapter 3. Future studies should focus on identifying other

gut environments suitable for plastic biodegradation, especially those of other ecosystems,

such as the marine environment. These e↵orts would help characterize endogenous plastic

degradation in di↵erent ecosystems and identity microorganisms and environmental controls

that could enable rapid biodegradation.

Plastic biodegradation mechanisms

To date, there has been limited research into the mechanisms of plastic biodegradation in

either isolated microorganisms or in the gut of insects. While the research in this thesis

contributes to our understanding of degradation specificity and factors that enhance degra-

dation activity, it also highlights the need for enhanced understanding of biodegradation

pathways and the need for tools that can provide insight into these pathways in novel sys-

tems. Increased understanding of the degradation pathways and enzymes will enable future

e↵orts to scale-up biodegradation either through bioprocess engineering and/or synthetic

biology. This increased understanding may enable tailored biodegradation processes, such

as re-routing the degradation pathway to a di↵erent endpoint. Such tailoring could enable

the reclamation of valuable molecules or monomers, rather than letting the degradation pro-

ceed to complete mineralization. This possibility has been demonstrated for polyethylene

terephthalate (PET) where enzymatic PET hydrolysis yields terephthalic acid and ethylene

glycol,[198, 214, 87] both of which can serve as feedstocks for the production of bioplastics

such as polyhydroxyalkanoates.[87, 61] This process thus enables the use of by-products

from petroleum-based plastic degradation as substrates for production of biodegradable

bioplastics. This underscores the importance of identifying plastic degradation mechanisms

that can convert recalcitrant plastics into biodegradable and sustainable materials.

Scaling-up of plastic biodegradation

While a better understanding of the mechanisms of plastic degradation will support e↵orts

to scale-up these systems, more research is needed. There is a need to understand how to

increase the scale of plastic biodegradation can be accomplished without compromising on

the e�ciency or extent of degradation. Research is also needed to determine the optimal size

of future systems and to minimize losses that occur during collection and transportation

while maintaining economic feasibility. Further work is also needed to investigate how

mechanisms from biodegradation can be used at scale in the design and production of

new materials. As one example, further knowledge of enzyme structure and mechanism

might enable the production of plastics that are susceptible to enhanced degradation under

specific environmental conditions. This idea is already being explored commercially to

pre-seed materials with the necessary biodegradation enzymes to transform conditionally

biodegradable plastics, like polylactic acid, to fully biodegradable materials.[13] Utilizing

insights from biodegradation upstream in the design and production process could further

reduce waste management challenges and ultimately yield products that are less harmful to

the environment.

6.2. RECOMMENDATIONS FOR FUTURE RESEARCH 95

The need for sustainable replacement materials and green chemistry

To fully address the challenges of plastic pollution, there is a need to move from the linear

cradle-to-grave lifecycle of conventional plastics to renewable materials that fit into a circular

lifecycle. To make this transition economically feasible, research is needed to enhance the

material properties of proposed replacement materials to enable their ready use as drop-in

replacements for conventional plastics. Further biotechnology advances can help drive down

costs and overcome processing issues that currently limit widespread use of these plastics.

In addition, there is a need to overcome previous definitional issues surrounding replacement

materials. The Ecocyclable standard highlighted in Chapter 5 is a proposed option that

would enable a more nuanced definitions, and could be incorporated into legislation on

plastic pollution.[117, 195]

Current systems for plastic production and recycling fail to address and treat chemical

additives in plastic, such as fire retardants and plasticizers.[137] These additives are poten-

tially harmful to environmental and human health and require careful mitigation to ensure

they do not escape into the environment. They would pose similar threats if used in replace-

ment materials to achieve enhanced chemical properties. Strategies are therefore needed to

ensure their removal from recycled materials and to enable non-toxic and sustainable green

chemistry replacements.[12]

In combination with legislative actions (e.g., United States Microbead-Free Waters Act

of 2015), improved public awareness, and personal behavior changes, biotechnology is poised

to play an increasingly important role in the remediation of existing plastic pollution and

their replacement by materials that are environmentally benign.

Emerging challenges

As our understanding of plastic pollution in the environment grows, it becomes increasingly

evident that there is no ecosystem that remains undisrupted and unharmed. Macroplastic

pollution (e.g., plastic bags, water bottles, straws) are well documented to cause harm

through ingestion, entanglement, and exposure to chemical additives. Areas of emerging

concern that will necessitate further research include impacts from micro- and nano-plastics.

Research demonstrates that microplastic particles are ingested by a variety of zooplankton

species at the base of the marine food web, giving rise to concerns of bioaccumulation.[34, 35]

Health risks from the ingestion of these microplastics may result from their bioaccumulation

up the food chain or through other means of concentration, such as sea salt.[37, 202, 205]

Researchers have just begun examining the risks posed by nano-plastics.[152, 115] Clearly,

there is an imminent need to address plastic pollution in the environment without delay

and to develop remediation strategies across multiple scales.

Appendix A

Supporting Information for

Chapter 2

98 APPENDIX A. SUPPORTING INFORMATION FOR CHAPTER 2

A.1 Material and Methods Supplement

A.1.1 Extraction method e�ciency assay

The extraction method used to recover residual polymer from the frass in this study was

based on previously established methods.[207] To assess the e�ciency of the method, frass

samples from the bulk-fed containers (from the plastic only diets, PE and PS) along with

frass samples from the bran-fed controls were gently crushed in a mortar and pestle and

then divided into ⇠ 50 mg aliquots (4 for PE samples, 2 for PS samples, 4 for bran sam-

ples) and placed in clean glass vials with 2 mL of the extraction solvent and extracted

with gentle heating (placed on a hot plate on the lowest setting). The PS samples were

extracted in tetrahydrofuran (THF) (� 99.9%, Thermo Fisher Scientific Inc., Pittsburg,

PA) in duplicate. The PE and bran samples were extracted in THF in duplicate as well

as in dichloromethane (DCM) (� 99.9%, Thermo Fisher Scientific Inc., Pittsburg, PA) in

duplicate. Every 30 minutes for 3 hours the samples were allowed to settle, then the solvent

was recovered (leaving the frass particles in the vial) and filtered using a 0.22 µm PVDF

filter (Thermo Fisher Scientific Inc., Pittsburg, PA) into a clean pre-weighed glass vial.

Two mL of solvent was then re-added to the initial vials containing the frass to allow the

extraction to continue until the next time point. For the last time point, the vials were left

extracting for upwards of 12 hours before the solvent was recovered and filtered. The ex-

tracted residual polymer in the filtered solution was the concentrated by rotary evaporation

and the residual polymer was weighed to determine the extractable fraction (the percent of

the initial frass weight recovered in the extraction).

A.1.2 Phasing amplicon sequencing library preparation

The phasing amplicon sequencing (PAS) approach was used to sequence the V4 region

of 16S rRNA genes.[203] For the first-step PCR, 5 µL of 10⇥ PCR bu↵er II (Life Tech-

nologies, Carlsbad, CA), 2 µL of 10 µM forward and reverse target-only primers 515F

(5‘GTGCCAGCMGCCGCGGTAA3’) and 806R (5‘GGACTACHVGGGTWTCTAAT3’),

and ⇠10 ng template DNA were mixed in a 50 µL reaction. The DNA was denatured

at 94 �C for 1 min, and 10 cycles of 94 �C for 20 s, 53 �C for 25 s, and 68 �C for 45 s were

conducted before a final extension at 68 �C for 10 min. The PCR products were purified

with an Agencourt AMPure XP kit (Beckman Coulter, Beverly, MA) and were used in the

A.1. MATERIAL AND METHODS SUPPLEMENT 99

second-step PCR in which 2.5 µL 10⇥ PCR butter II, 2 µL forward and reverse phasing

primers (10 µM), and 15 µL of the first-step PCR products were mixed in a 25 µL reaction.

Twenty cycles of the above program were used and PCR products were detected with elec-

trophoresis and quantified with PicoGreen dsDNA assay Kit (Thermo Fisher Scientific Inc.,

Pittsburg, PA). Equal amount of product of the samples were pooled and purified with a

QIAquick gel extraction kit (Qiagen, Venlo, NL). Library of the 16S rRNA gene amplicon

sequencing were constructed using MiSeq reagent kit (Illumina, San Diego, CA) following

the manufacturer’s instruction, and DNA was sequenced using an Illumina MiSeq platform.

A.1.3 E. coli K12 plastic characterization controls

To test the possibility that the newly observed peaks in NMR and FTIR analysis were

attributable to contamination by bacterial biomass or secreted proteins, samples of plastic

(PE and PS) were separately treated in a suspension of E. coli K12, a common laboratory

bacterial culture not known to degrade plastics.[196] The E. coli K12 was grown in Luria-

Bertani (LB) broth (Thermo Fisher Scientific Inc., Pittsburg, PA) in 15 mL glass culture

tubes (Thermo Fisher Scientific Inc., Pittsburg, PA) at 30 �C with three approximately

0.1 g plastic pieces (either PE or PS) for 48-hours. After 24-hours, 5 mL of the broth was

removed from each tube and replaced with 5 mL of LB broth. After 48-hours, the plastic

samples were lightly rinsed with deionized water and allowed to dry completely prior to

being extracted and used for NMR or FTIR analysis.

A.2 Tables

A.2. TABLES 101

Table A.1: Overview of Illumina MiSeq data

Sequencing Data Feature:Minimum number of sequences (per sample): 9889Maximum number of sequences (per sample): 808059Mean number of sequences (per sample): 52532Resampling depth: 17460Resampled number of OTUs: 515Number of replicates per diet:

Bran 5PE 9

PE + Bran 7PS 2

PS + Bran 2PE + PS 2

Table A.2: 1H-NMR peak information (PE samples) based on analysis using MestReNovasoftware (version 10.0.2).

A.2. TABLES 103

Table A.3: 1H-NMR peak information continued (PS samples).

Table A.4: FTIR peak information for PE samples based on analysis using OMNIC software(Thermo Fisher Scientific Inc., Pittsburg, PA). Thermo Scientific Hummel Polymer andAdditives FTIR Spectral Library was used to obtain reference spectra of PE.

A.2. TABLES 105

Table A.5: FTIR peak information for PS samples, based on analysis using OMNIC software(Thermo Fisher Scientific Inc., Pittsburg, PA). Thermo Scientific Hummel Polymer andAdditives FTIR Spectral Library was used to obtain reference spectra of PS.

Table A.6: FTIR peak information for plastic mixture samples, based on analysis usingOMNIC software (Thermo Fisher Scientific Inc., Pittsburg, PA). Thermo Scientific HummelPolymer and Additives FTIR Spectral Library was used to obtain reference spectra of PEand PS.

A.2. TABLES 107

Table A.7: Relative abundance of PE and PS associated OTUs identified via di↵erentialabundance analysis. Average ± standard deviation reported by diet.

PE Associated Species Relative Abundance (% within sample) by DietOperational Taxonomic Units (OTU): PE Fed PE + Bran Fed PE + PS FedCitrobacter sp. 21.95 ± 9.45% 0.14 ± 0.08% 0.01 ± 0.02%Kosakonia sp. 17.94 ± 5.66% 1.17 ± 1.12% 5.40 ± 3.40%Sebaldella termitidis 1.78 ± 1.23% 0.04 ± 0.03% 0.11 ± 0.11%Brevibacterium sp. 0.12 ± 0.04% 0.01 ± 0.01% 0.03 ± 0.04%PS Associated Species Relative Abundance (% within sample) by DietOperational Taxonomic Units (OTU): PS Fed PS + Bran Fed PE + PS FedCitrobacter sp. 9.10 ± 9.08% 0.01 ± 0.01% 0.01 ± 0.02%Kosakonia sp. 13.14 ± 6.10% 0.01 ± 0.00% 5.40 ± 3.40%Listeria sp. 0.99 ± 1.40% 0.00 ± 0.00% 0.00 ± 0.00%Nistrospira defluvii 0.56 ± 0.40% 0.34 ± 0.24% 0.00 ± 0.00%Pedomicrobium sp. 0.19 ± 0.13% 0.11 ± 0.07% 0.01 ± 0.01%Aquihabitans sp. 0.20 ± 0.14% 0.18 ± 0.13% 0.00 ± 0.00%Un. Xanthomonodaceae 0.17 ± 0.12% 0.07 ± 0.05% 0.07 ± 0.05%Un. Saprospiraceae 0.15 ± 0.10% 0.03 ± 0.05% 0.01 ± 0.01%Un. Burkholderiales 0.23 ± 0.16% 0.09 ± 0.06% 0.02 ± 0.01%

A.3 Figures

A.3. FIGURES 109

6×104

8×104

1×105

1.0×105

1.5×105

2.0×105

olecular Weight (M

50010001500200025003000350040000

Wavenumber (cm-1)

ance PS Frass

OHRstretch

C Ostretch

aromaticstretch

out-of-planebending

PS + Bran Frass

PS Control

Bran Frass

PS + Bran Frass

PS Frass

PS Control

reduced aromatic region

Week 1Week 2

Week 3Week 4

PS Frass

PS + Bran Frass

PS Control

Figure A.1: Characterization of PS degradation within the mealworm gut. a) Changes inmolecular weight (Mn and Mw) within residual polymer from the frass versus the control PSas measured by HT-GPC. Significance (Student’s t-tests, Tukey’s multiple test correction)p < 0.05 indicated by ⇤, no statistical significance indicated by ns. (b) Changes in theextractable fraction, a measure of residual polymer in the frass, over the 32-day experiment.(c) 1H-NMR spectra of residual polymer extracted from the frass of PS and PS + branfed mealworms versus residuals extracted from bran fed mealworms, and the control PSfoam. The reduction in the aromatic region, appearance of phenol derivatives, and alcoholderivatives are highlighted in grey. Detailed peak information in Tables A.2, A.3. (d) FTIRspectra of residual polymer extracted from the frass of PS and PS + bran fed mealwormsversus the control PS, annotations show functional groups associated with key peaks basedon wavenumber. Detailed peak information in Tables A.4, A.5, A.6.

50010001500200025003000350040000

Wavenumber (cm-1)

PE E. Coli TreatedPE Control

50010001500200025003000350040000

Wavenumber (cm-1)

ance PS Control

PS E. Coli Treated

PE Control

PE E. Coli treated

PS Control

PS E. Coli treated

(a) (b)

(c) (d)

Figure A.2: Characterization of plastic treated with E. coli K12 as a control against micro-bial contamination (SI MethodA.1.2). (a & b) 1H-NMR spectra of control plastics comparedto E. coli K12 treated plastics (a: PE, b: PS). Detailed peak information in Tables A.2,A.3. (c & d) FTIR spectra of control plastics compared to E. coli K12 treated plastics (c:PE, d: PS). Detailed peak information in Tables A.4, A.5, A.6.

A.3. FIGURES 111

Week 1Week 2

Week 3Week 4

PE Fed

Week 1Week 2

Week 3Week 4

PE + Bran Fed

Week 1Week 2

Week 3Week 4

PS Fed

Week 1Week 2

Week 3Week 4

PS + Bran Fed

Week 1Week 2

Week 3Week 4

Frass - non-extractable fractionFrass - extractable fraction

Mealworm Biomass

PE + PS Fed

Week 1Week 2

Week 3Week 4

sFrass - non-extractable fractionFrass - extractable fraction

Mealworm Biomass

Bran Fed Control

(a) (b) (c)

(d) (e) (f)

Figure A.3: Mass balance on mealworms during the 32-day experiment. The percent ofingested mass (plastic & bran, depending on the diet) recovered in the frass, the extractableportion of the frass (e.g., residual polymer), and the mealworm biomass throughout the 4-week experiment (days 6, 15, 21, 27). The plastic only diets (a � c) show a steady decreasein the amount of plastic being recovered in the frass and an increase in the PGF (likely CO2)(a: PE, b: PS, c: PE + PS). The plastic diets co-fed with bran (d � e) also show a decreasein the amount of plastic being recovered in the extractable portion of the frass (d: PE +Bran, e: PS + Bran). The bran-fed control (f) shows little to no material was recovered inthe extractable fraction over the course of the experiment and that the amount of ingestedmaterial (bran) converted to gases increases overtime while the amount of ingested materialconverted into mealworm biomass decreased with time.

50010001500200025003000350040000

Wavenumber (cm-1)

stretch

C HOHRbend

stretch

C Ostretch

aromaticstretch

PE + PS Frass

out-of-planebending

PE ControlPS Control

Figure A.4: Characterization of mixed plastic (PE + PS) degradation. FTIR spectra ofresidual polymer from the frass of PE + PS fed mealworms versus the control polymers (PEand PS), annotations show functional groups associated with key peaks based on wavenum-ber. Detailed peak information in Tables A.4, A.5, A.6.

●●

PS_Bran1

PE_PS2PE1PE2

PE_Bran1

PE_Bran3

PS_Bran2

PE_Bran4

PE_Bran2

PE_PS1

PE9PE6

PE_Bran5

PE_Bran4

Bran1Bran3

PE_Bran3

●●

−0.2

−0.50 −0.25 0.00Axis.1 [27.7%]

Diet●a●

●a●

PE_Bran

PS_Bran

Figure A.5: Principal Coordinate Analysis of microbial communities by diet based onBray-Curtis distance, colored by diet and labeled with the sample ID with outliers (PS1,PS Bran2) included.

A.3. FIGURES 113

30 min60 min

90 min120 min

150 min180 min

>12 hour0

) PE in DCM - total extractable fraction 41.02 ± 0.64% PE in THF - total extractable fraction 5.89 ± 0.23%PS in THF - total extractable fraction 42.60 ± 0.78%

Time Extracted

Bran in THF - total extractable fraction 0.95 ± 0.22%Bran in DCM - total extractable fraction 1.11 ± 0.21%

Figure A.6: E�ciency of extraction method used to recover residual polymer in the frass.Extractable fraction (the percent of the initial frass weight recovered in the extraction, the“residual polymer”) recovered over time for frass from PE- and PS-fed mealworms as well asbran-fed controls. The residue recovered from the PE- and PS-fed samples was visible whiteand polymer-like while the residue recovered from the bran-fed controls was not, which isconsistent with previous applications of this method (not shown).[207]

Appendix B

Chapter 3

116 APPENDIX B. SUPPORTING INFORMATION FOR CHAPTER 3

B.1 Tables

Table B.1: Sequenced bacterial strains from microbial enrichment and isolates. Sequencesbacterial strains isolated from the cultivated enrichment (mixed community, “Enrichment”)and the bacterial isolates grown on PS film (“Isolates”) are shown with the most similarstrain (by sequence identity), the similarity grade (%), query coverage (%). All values arederived from BLAST comparison.

B.2. FIGURES 117

B.2 Figures

5 10 15

Control (Bacteria alone)Bacteria + PS

5 10 15-0.5

Control (Bacteria + 0.05 mL)0.05 mL Super. + PS0.05 mL glucose equivalent

5 10 15

5 10 15-1

5 10 15

-5-4-3-2-10123

(b) (c)

(d) (e)

Figure B.1: Dose-response curves for supernatant and glucose equivalent. Gut microbiomewas cultured in carbon-free media with PS microplastics and supplemented with varyingdoses of PS-fed supernatant (blue) or glucose equivalent based on chemical oxygen demand(red), controls (grey) were seeded with gut microbiome and supernatant but no PS. Respi-ration activity (CO2 production) was measured over 14 days. Five supernatant doses weretested: (a) 0 mL, (b) 0.05 mL, (c) 0.1 mL, (d) 0.5 mL), and (e) 1.0 mL.

B.2. FIGURES 119

(a) (b)

+20 min

Bran supernatant PS-fed supernatant+ - + -

Neg. Pos.Controls

Neg. Pos.Controlsantibiotic

treatment

Figure B.2: Supernatant emulsification activity. To assess the capacity for gut supernatantfrom the four experimental diets (PS and bran, with and without antibiotic treatment) toemulsify PS microplastics into aqueous solution. (a) Emulsifications formed immediatelyafter 2 minutes of vortexing, labeled by diet (top row) and antibiotic treatment (bottomrow). Negative control (DI water + PS microplastics) and positive control (DI water +PS microplastics + Tween 80, a common surfactant) are shown on the right. (b) Emulsi-fications remained 20 minutes after vortexing, demonstrating PS microplastics are stablyincorporated into aqueous phase.

Figure B.3: Enrichment biofilm formation. The microbial enrichment was cultured in car-bon free media with PS film and supplemented with PS supernatant. A stable biofilmformed on the PS film after 3 months of growth (shown).

Appendix C

Chapter 4

122 APPENDIX C. SUPPORTING INFORMATION FOR CHAPTER 4

C.1 Material and Methods Supplement

C.1.1 Chemicals and reagents

Hexane (>99.9%, HPLC grade) and methylene chloride (DCM, >99.9%, HPLC grade) were

purchases from Fischer Scientific Inc. (Pittsburg, PA). a-, b-, and g-HBCD (1,2,5,6,9,10-

hexabromocyclododecane, >98.0%, analytical standard grade) standards were purchased

from Sigma-Aldrich (now MilliporeSigma, St. Louis, MO). Sodium sulfate anhydrous

(99.0%) was purchased from EMD Millipore (now MilliporeSigman, St. Louis, MO) for

use sample preparation for HBCD analysis. GE Healthcare Whatman Binder-Free Glass

Microfiber Filters (Grade 934-AH, 70 mm) were purchased from Fischer Scientific Inc.

(Pittsburg, PA) for use sample preparation for HBCD analysis. Restek Normal Phase Re-

sprep SPE cartridges (packing material: Silica, 6mL, 1000mg) for the final clean-up step of

HBCD analysis was purchased from Fischer Scientific Inc. (Pittsburg, PA). Stock solutions

containing a mixture of polychlorinated biphenyls (PCBs) for use as a surrogate standards

and internal standards for HBCD chemical analysis (Surrogate standard: BZ#s 14, 65, 166;

Internal standard: BZ#s 30, 204) were purchased from Agilent ULTRA (Santa Clara, CA).

C.1.2 HBCD analysis

Analytical methods to quantify HBCD in mealworm and shrimp biomass as well as polystyrene

(PS) samples were developed following modified EPA methods[182, 181] for polychlorinated

biphenyls (PCBs) and organochloride pesticides because there is currently no accepted EPA

method for HBCDs and these methods have been used previously for the quantification of

other chlorinated and brominated compounds.[192, 96] Figure C.3 shows a schematic rep-

resentation of sample preparation, which is further explained below. To quantify HBCD in

mealworm biomass, mealworms were first sacrificed and then the gut track was dissected

out, separating the gut tissues from the non-gut tissues, these samples were pooled (10

mealworm gut tracks or non-gut tissues per sample). Samples were then freeze-dried for

>72 hr. Shrimp biomass was first sacrificed and pooled (5 – 6 shrimp/sample) prior to

freeze drying (>72 hr). These freeze-dried samples, as well as collected mealworm frass

samples, were then homogenized (via mortar and pestle). Sample extraction method was

based on EPA method 3350B.[180] The homogenized samples were then dried with sodium

C.1. MATERIAL AND METHODS SUPPLEMENT 123

sulfate, spiked with the surrogate standard (a mixture of PCBs, BZ# 14, 65, 166; 20 µLof 400 µg/L)[102, 215] prior to being extracted with methylene chloride (DCM, 20 mL)

using ultrasonic extraction (20 min in a Branson 2510 Ultrasonic bath). Polystyrene sam-

ples (0.05 g) were dissolved in 20mL DCM, spiked with the surrogate standard, and then

subject to the same ultrasonic extraction. Post sonication, samples were allowed to set-

tle, and the overlying DCM was decanted over a filter (containing approximately 1.0 g of

sodium sulfate) into a clean glass vial using a clean glass funnel. Twenty mL of DCM

was then re-added to the solid sample and re-extracted following the same methods. After

the solvent from the second extraction was filtered, samples were evaporated (using an N2

evaporator, N-EVAP 111) until the remaining volume was less than 2 mL, after which 4

mL of hexane was added to the vial and swirled to mix (repeated a second time after the

volume returns to 2 mL). The remaining volume (less than 2 mL) was then passed through

a silica SPE clean-up column (pre-conditioned with 4 mL hexane) following EPA method

3630C.[178] The column was then eluted with 4 mL of DCM and the flow-through collected

and concentrated to 1 mL using the N2-evaporator. The samples were then spiked with

internal standard (a mixture of PCBs, BZ# 30, 204; 10 µL of 400 µg/L) prior to instrumen-

tal analysis.[102, 215] PCBs were selected as surrogate and internal standards as they are

stable, chemically similar compounds to HBCD that do not interfere with quantifying the

peak of interest using our analytical methods and have been used previously as standards

for HBCD analysis.[215]

Quantification of total HBCD was performed using an Agilent gas chromatograph[102,

188, 73] (model 6890) with a fused silica capillary column (HP-5, 60 m ⇥ 0.25 mm ID)

and a micro electron capture detector (GC- µECD) based on EPA methods 8082[182] and

8081.[181] The main stereoisomers of HBCD present in PS are a-, b-, and g-HBCD; g-HBCD

is the predominant stereoisomer used in commercial products.[215, 183, 11] Standards of the

three main stereoisomers were tested independently and as a mixture. Retention times and

response factors of the three stereoisomers were statistically indistinguishable (Fig. C.8),

justifying the use of a single HBCD peak to quantify total HBCD.6,9,10 All quantifications

in this paper refer to total HBCD. A six-point calibration curve was prepared with g-

HBCD standard with the internal standard. HBCD concentration measured in unexposed

mealworms (under the control, bran-fed diet) and the bran used as feed were treated as

background (n = 3, all samples were below detection). Method detection limit (MDL) for

mealworm biomass samples was 0.20 ng HBCD/g dry weight and for shrimp biomass the

MDL was 1.00 ng HBCD/g dry weight. Our limit of detection (LOD) and limit of quanti-

tation (LOQ) were determined to be 40 pg and 1000 pg, respectively, based on established

definitions.[5, 116] To increase the robustness of our analysis, we do not utilize any values

below the MDL for statistical quantification, but we plot all values above our LOD. In ad-

dition, laboratory blanks were run with every batch (10 -– 20 samples/batch) of samples to

assess contamination from solvents, glassware, etc. To check for contamination in the GC,

a hexane blank is run at the beginning and end of a sample batch (10 – 20 samples/batch)

based on recommended frequency by the EPA[179] and best practices established by our

laboratory,[179, 108] no carry-over was observed. A preliminary assay was conducted to

assess average recovery e�ciency of surrogate standard across di↵erent sample types (Fig.

C.6a), we then used these results to establish an accepted range of recovery (40 – 100%) in

accordance with our laboratory established standard methods[108] (following EPA methods

that each laboratory should establish their own accepted range of recovery[182]). Recoveries

from this preliminary assay as well as from all the samples in this study are shown in Fig.

C.1.3 Litopenaus vannamei experimental conditions

L. vannamei postlarvae were purchased from Shrimp Improvement Systems LCC (Islam-

orada, FL). Following overnight shipment, the postlarvae were acclimated over 12 hrs fol-

lowing guidelines from the company to ensure changes in pH, salinity, and temperature were

less than 0.1 pH unit/15 min, 1ppt/15 min and 1�C/15 minutes, respectively. The postlar-

vae were kept in 30 L tanks (stocking rate ⇠ 80 postlarvae/L) with constant aeration, which

were maintained at 28 �C and 33 ppt salinity (with 50% water changes every 3 days) until

they were used in experiments. Artificial seawater (35 g/L) was made with Instant Ocean

(Blacksburg, VA). For the first 48 hr after arrival, the postlarvae were fed recently hatched

brine shrimp (premium grade Artemia franciscana cysts, Brine Shrimp Direct, Ogden, UT).

After 48 hr, their diet was switched to standard postlarvae feed for post larval stages 15 –

20 (FRiPPAK RW+500, INVE Aquaculture, Deception Bay, Queensland, AUS), which was

used in all the experiments as the control diet. The postlarvae were fed ⇠ 3% of biomass

per tank daily.

Two experimental set-ups were utilized to assess the impact of PS-fed mealworms on

survival and fate of HBCD within the aquaculture biomass (Fig. C.2). In the first experi-

mental setup (“toxicity & survival setup,” Fig. C.2a), individual postlarvae were transferred

C.1. MATERIAL AND METHODS SUPPLEMENT 125

to a cup (145mL sterile food-grade polypropylene) that enabled the survival tracking of in-

dividual shrimp and prevented cannibalism, following methods as described by Crab et

al.[38] Each cup contained 50 mL initially composed of 50% fresh aquarium water and 50%

water from their previous tank to ease acclimation. The individual cups were suspended

in a temperature-controlled water bath maintained at 28 �C using an aquarium heater (50

W submersible glass heater, Aqueon Products, Franklin, WI). Postlarvae were allowed to

acclimate for at least 48 hr in the cups prior to the start of the experiment, during which

time they were fed with the control diet. A 50% water change was performed daily to

maintain high quality water within the individual cups. Each group included 13 shrimp.

The experiment was run for 10 days and survival was monitored daily for Kaplan-Meier

survival analysis. After the acclimation period, shrimp were switched to the experimental

diets (fed at ⇠ 3% shrimp weight daily, ⇠ 0.003 g per day). The experimental diets in-

cluded: bran-fed mealworm biomass (MWB), PS-L fed MWB, PS-H fed MWB, each type

of MWB was integrated into the shrimp diets at 3 di↵erent rates: 10%, 50%, and 100%

(for a total of 9 experimental diets). Control shrimp were fed commercially available feed

(FRiPPAK RW+500, INVE Aquaculture, Deception Bay, Queensland, AUS).

Experimental feeds were prepared utilizing MWB from bulk experimental setup (Fig.

C.1). The mealworms were subject to a 24-hr depuration, sacrificed, and the biomass (whole

mealworm) freeze-dried for >72 hr. The dried biomass was then homogenized with a blade

grinder and mixed with the requisite amount of control feed for each diet. The experimental

feeds were prepared in bulk and the same batch of the feeds were utilized for all tests.

The second experimental setup (“bulk setup,” Fig. C.2b) enabled the collection of

su�cient shrimp biomass to conduct HBCD chemical analysis. Fifty postlarvae were placed

in a 20 L tank filled with 14 L of aquarium water with constant aeration. A 50% water

change was conducted every 3 days. The same experimental diets were tested as in the

toxicity setup, excluding the diets composed of 100% MWB due to the high mortality

caused by that diet (total of 6 experimental diets in addition to a control group, fed daily).

After 14 days, the surviving shrimp were sacrificed (transferred to ice-water bath for >10

mins) for HBCD analysis (SI Method C.1.2 and Fig. C.3 for further details).

C.1.4 Mealworm depuration experimental conditions

Depuration tests post feeding of PS were conducted to investigate accumulation of HBCD

in mealworm tissues over time periods that were comparable to the retention time of PS in

the mealworm gut (15 -– 20 hr).[18]

The expected intake of HBCD was determined for the mealworms fed PS-H and PS-L

(calculated as the average amount of HBCD consumed for each diet [ng/72 hr], normalized to

10 mealworms. To assess the impact of the presence polystyrene on HBCD bioaccumulation

and egestion, bran was spiked with HBCD at levels comparable to PS-H and PS-L and fed

directly to mealworms, referred to as Bran-H and Bran-L, respectively. HBCD was added at

a concentration of 30 µg/g or 0.05 µg/g for Bran-H and Bran-L comparisons, respectively,

to bran by spiking HBCD into 1 mL of DI water, which was then mixed and uniformly

applied to 4.5 g of bran and allowed to dry in a sealed container prior to being fed to

mealworms. Bran-H and Bran-L were analyzed for HBCD concentration (in triplicate,

following SI Method C.1.2) after drying and no loss of HBCD was detected through the

bran preparation process. After 72 hr of feeding, ten mealworms from all diets were collected

at 0, 24, and 48 hr of depuration (starvation conditions), in triplicate, for analysis of the

egestion of HBCD.

C.2. TABLES 127

C.2 Tables

Table C.1: Characterization of the two PS foams tested in this study (mean ± SD, n = 4)including concentration of HBCD, density, molecular weight (both number-averaged, Mn,and weight-averaged, Mw), and source. ESP — expanded polystyrene foam.

C.3 Figures

C.3. FIGURES 129

Figure C.1: Schematic diagram showing the two mealworm experimental setups utilized.(a) The mass balance experimental setup to track HBCD (see methods “mealworm growthconditions”) with 200 mealworms in individual containers in duplicate for each experimentaldiet. Maintenance (weighing of plastic and frass, addition of bran, survival) and sacrificeschedule described on the bottom. Data from this experimental setup utilized in Fig. 4.2 andFig. 4.4. (b) The bulk experimental setup to collect su�cient mealworm biomass to produceaquaculture feed for the higher trophic level bioaccumulation studies. Larger containers weremaintained under the same condition as the mass balance experience. Images not to scale.

Figure C.2: Schematic diagram illustrating the two experimental setups utilized to assesssecondary trophic level e↵ects of HBCD utilizing L. vannamei. (a) The toxicity and survivalexperimental setup. Postlarvae (PL 28) were placed in individual cups and then survivalwas monitored daily for Kaplan-Meier survival analysis in addition to daily water changesand feeding. (b) The bulk experimental setup to obtain su�cient L. vannamei biomassfor HBCD analysis. Fifty postlarvae were placed in a 20 L tank filled with 14 L of waterfed with the same experimental diets as those in the toxicity setup. A 50% water changewas conducted every 3 days. After 14 days, the surviving shrimp were sacrificed for HBCDanalysis. Images not to scale.

Figure C.3: Sample preparation for HBCD analysis via gas chromatography equipped witha micro electron capture detector (GC- µECD). Schematic diagram showing the sampleprocessing for HBCD analysis for the three types of samples studied. On the left, mealwormbiomass sample processing including the three types of biomass tested (gut track, non-guttrack, frass). In the middle, shrimp biomass sample processing. On the right, plasticcontrols sample processing. Addition of surrogate standard and internal standard in thesample analysis represented by Surg. Std. and Int. Std., respectively.

C.3. FIGURES 131

50 100 15050

Actual Number of Pupae

BranPS-HPS-H + branPS-LPS-L+branLine of identity

Figure C.4: Predicted versus actual number of pupae, based on a multiple regression analysisto assess the impact of amount of bran, PS, and HBCD consumed on pupation (model R2 =0.95), plotted by diet. The grey dashed line shows the line of identity (y = x). This analysisrevealed that bran consumption was a significant (p < 0.0001) predictor of pupation ratewhile HBCD consumption was not (p > 0.05)

2×105

4×105

6×105

8×105

100001200014000

/ g dw

Consumed in PS

Egested in Frass

Week 1 Week 2 Week 3 Week 40.00.10.20.30.40.50.63.03.54.0

/ g dw

) Gut trackBody tissuesMDL

(a) (b)

(c) (d)

Figure C.5: Fate tracking of HBCD in T. molitor in co-fed diets (PS-H + bran, PS-L +bran). (a, b) Amount of HBCD consumed in PS (grey bars) coupled with the amountof HBCD egested in frass (purple bars) from each container (a: PS-H + bran, b: PS-L +bran). Amount of HBCD consumed was calculated as total plastic consumed per week [g] ⇥the concentration of HBCD in the plastic [ng/g]. Amount of HBCD egested was calculatedas the concentration of HBCD in the weekly representative clean frass sample [ng/g] ⇥the total frass collected for the week [g]. (c, d) Body burden of HBCD [ng/g dry weight]in mealworm tissues (gut track or non-gut body tissues) weekly, plotted with the methoddetection limit (MDL) (c: PS-H + bran, d: PS-L + bran). Expected weekly intake (EWI)of HBCD was calculated as the average amount of HBCD consumed for each diet [ng/week],normalized to 10 mealworms, and is plotted for comparison. All values represent mean SD,n = 2.

PS-H PS-L Mealworm gut biomass

Mealworm biomass

Shrimp biomass

PS-H PS-L Mealworm gut-track

Mealworm non-gut biomass

Mealworm frass

Shrimp biomass

Accepted range 40 - 100%

(a) (b)

Figure C.6: Recovery e�ciency of surrogate standard in di↵erent sample types includingPS foams (PS-L, PS-H) as well as mealworm and shrimp biomass. (a) Results from thepreliminary analysis of each sample type to determine an accepted range for recovery e�-ciency. (b) Recovery e�ciencies across all samples in this study. Gray dashed lines showthe upper and lower bound of the accepted range, 40 -– 100%.

C.3. FIGURES 133

100PS-H

PS-H + Bran

PS-L + Bran

Figure C.7: Mass balance on HBCD in the mealworm experiments. Percent of ingestedHBCD recovered (combined) in the three types of mealworm tissues (frass, gut track, bodytissues) by diet for each week. Surrogate standard recovery e�ciencies (Fig. C.6b) wereused to estimate the HBCD recovered each week. Average HBCD recovered by diet, PS-H:94.14 ± 4.73%; PS-H + Bran: 94.65 ± 4.75%; PS-L: 92.60 ± 7.23%; PS-L + Bran: 94.48± 4.33%.

0 500 10000

HBCD Concentration (µg)

α-HBCDβ-HBCDγ-HBCDα-, β-, γ-HBCD (equal concentrationof each)

Figure C.8: Standard curves for the main stereoisomers of HBCD (a-, b-, and g-HBCD)individually and in combination (at equal concentrations of each) on the GC-ECD, plottedwith 95% confidence intervals. The slopes of response factors are statistically indistinguish-able (p = 0.16).

24-hour Starvation

48-hour Starvation

PS-HPS-Lns

Bran-HBran-L

Figure C.9: Log removal of HBCD at each sampling time for mealworms fed PS-H, PS-Lor bran-spiked with HBCD (Bran-H and Bran-L). All values represent mean SD, n = 3.Significance (two-way ANOVA, Bonferroni multiple test correction) p 0.001 indicated by⇤ ⇤ ⇤, no statistical significance indicated by ns.

Appendix D

Chapter 5

136 APPENDIX D. SUPPORTING INFORMATION FOR CHAPTER 5

D.1 Tables

Table D.1: Cost, material properties, and suitability for various thermal manufacturing pro-cesses for common synthetic petroleum-derived plastics and plastic replacement materials.Cost estimates for petroleum-derived plastic were obtained from the historic resin pricingdatabase for 2018, range of lowest to highest prices reported (a) [138]. Cost estimates for re-placement materials were obtained from the 2013 Bioplastics in California report (b) [155].References for thermal and material properties for each polymer listed after the polymername: (c) Castilho et al., 2009 [26]; (d) Andreeben et al., 2014 [4]; (e) Farah et al., 2016[57]. Suitability for manufacturing based on insights from (f) Applied Plastics EngineeringHandbook [98] as well as (g) 3D printing with biomaterials: Towards a sustainable andcircular economy [186].

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Relationship between virulence and enzymatic profiles in the cuticle of Tenebrio molitor by 2-deoxy-d-glucose-resistant mutants of Beauveria bassiana (Bals.) Vuill

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A Novel Tenebrio molitor Cadherin Is a Functional Receptor for Bacillus thuringiensis Cry3Aa Toxin

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A tapeworm molecule manipulates vitellogenin expression in the beetle Tenebrio molitor

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