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Environmental Chemistry of Commercial Fluorinated Surfactants:

Transport, Fate, and Source of Perfluoroalkyl Acid Contamination in the Environment

by

Holly Lee

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Department of Chemistry

University of Toronto

© Copyright by Holly Lee (2013)

ii

Environmental Chemistry of Commercial Fluorinated Surfactants:

Transport, Fate, and Source of Perfluoroalkyl Acid Contamination in the Environment

Doctor of Philosophy Degree, 2013

Holly Lee

Department of Chemistry, University of Toronto

ABSTRACT

Perfluoroalkyl carboxylates (PFCAs) and perfluoroalkane sulfonates (PFSAs) are

anthropogenic fluorinated surfactants that have been detected in almost every environmental

compartment studied, yet their production and applications are far outweighed by those of other

higher molecular weight fluorinated surfactants used in commerce. These fluorinated surfactants

are widely incorporated in commercial products, yet their post-application fate has not been

extensively studied. This thesis examines various biological and environmental processes

involved in the fate of these surfactants upon consumer disposal. Specific focus was directed

towards the environmental chemistry of polyfluoroalkyl phosphate esters (PAPs), perfluoroalkyl

phosphonates (PFPAs), and perfluoroalkyl phosphinates (PFPiAs), and their potential roles as

sources of perfluoroalkyl acids (PFAAs) in the environment. PAPs are established biological

precursors of PFCAs, while PFPAs and PFPiAs are newly discovered PFAAs in the

environment.

Incubation with wastewater treatment plant (WWTP) microbes demonstrated the ability

of PAPs to yield both fluorotelomer alcohols (FTOHs), which are established precursors of

PFCAs, and the corresponding PFCAs themselves. WWTP biosolids-applied soil-plant

microcosms revealed that PAPs can significantly accumulate in plants along with their

degradation metabolites. This has implications for potential wildlife and human exposure

iii

through the consumption of plants grown and/or livestock raised on farmlands that have been

amended with contaminated biosolids.

A number of compound-and environmental-specific factors were observed to

significantly influence the partitioning of PFPAs and PFPiAs between aqueous media and soil, as

well as, aquatic biota during sorption and bioaccumulation experiments respectively. In both

processes, PFPAs were primarily observed in the aqueous phase, while PFPiAs predominated in

soil and biological tissues, consistent with the few environmental observations of these

chemicals made to date.

Detection of the PAP diesters (diPAPs), PFPiAs, and fluorotelomer sulfonates (FTSAs),

all of which are used commercially, in human sera is evidence of human exposure to commercial

fluorinated products, but the pathways by which this exposure occurs remain widely debated.

Overall, this work presents novel findings on the environmental fate of commercial fluorinated

surfactants and each of the process studied shows a clear link between the use of commercial

products and the fluorochemical burden currently observed in the environment.

iv

ACKNOWLEDGEMENTS

With every obstacle I have encountered and overcome, I always think back to the quote,

“It takes an army to…”, because I am truly thankful for my personal army of family and friends

that have been rallying for me for the past five years and throughout my life.

I am truly grateful to Scott for his unbeatable creativity and enthusiasm, both of which

have inspired me to become the scientist that I am today. Not only has he been generous with his

encouragement during my Ph.D, but he has also given me many opportunities to travel abroad,

meet new people, and establish my own footing in our field. Thanks for always pushing me to

go above and beyond my limits. Through you, I have learned that no question is unanswerable;

it just depends on how hard you tackle it!

I also thank Frank and Jen for their continued support as my teachers and committee

advisors during my Ph.D, as well as the entire environmental chemistry faculty. Eric Reiner,

Derek Muir, and John Washington – thank you so much for your advice and generous support.

To the AIMS folks, thanks for being so patient and generous with helping me whenever our

instrument is down – special thanks to Michelle for always being there to share my LC pains!

Also a big thank you to Anna Liza for taking care of my big Ph.D milestones!

I thank my lucky stars everyday for the awesome group of people I got to work with for

the past five years. Amila, Craig, Cora, and Jess, you guys have not only been incredible role

models for me, but also great friends. Amila, I love how you can always make a good laugh out

of anything (“I do like it”). Craig, I can always count on you staying late at work and teaching

me those biodeg pathways! Cora, I’ll always remember our road trip to Ford and how you took

me shopping! Jess, where do I even begin? From day one, you have supported and believed in

me even when I didn’t believe in myself. You’ll always be my LC Yoda. Pablo, I wish you had

stayed so we can take another thumbs up picture together with our degrees, but I am so proud of

what you’ve accomplished today. Amy, my other half! I’m really happy we’ve become such

great friends and I’m constantly amazed at how you always manage to pull everything together –

it’s TIME. Derek, you are the walking Wikipedia that every group should have and I really

appreciate how generous you always are with helping me and others out. Anne, your laugh is so

infectious and I’m so glad you came back for your Ph.D! Keegan, your easy-going personality

v

has made working on the agro project a much less stressful experience and I look forward to

seeing our papers in press! Angela, I still remember thinking who is that smarty pants sitting

across from me in Frank’s modeling class and it has been a blast getting to know you better

through all our long chats! Leo the guru! Everyday I’m amazed at how hard you work and

despite how busy you always are, you never hesitate to help or even just to listen to my rants. I

will definitely miss working with you. Erin, Susanne, Shona, Rob, Lisa, Barbara, and Rui – it

has been a short, but sweet time working together and I wish you all the best! Many thanks also

to Alex, Hanin, Alicia, Ling, and Inthuja for working tirelessly whenever I needed extra help.

To everyone in environmental chemistry, you guys have given me five amazing years of

memories and I look forward to being friends for a long time to come! To Jeff, king of BP, how

is it possible that we went through undergrad without even knowing each other existed? You

have been an incredible walking-to-work buddy, coffee break/lunch partner, venting machine,

but most importantly, a close friend whom I’ll always hold dear to my heart. Stay on MSN!

Sarah, my partner-in doing everything last minute-crime, I’m going to miss our late-night chats!

To all my friends outside of chemistry, thanks for your love and support and always being there

whenever I’m ready to let my hair down and have fun! Toni, Barbara, and Lydia, I know I can

always count on you for advice, a shoulder to cry on,or just simply to make me laugh.

I am eternally grateful to my family and relatives for their unconditional love and support

throughout my life. To my beloved dog, Jai Jai, you’ll always have a special place in my heart.

To Aunt Josephine and Uncle Stanley, thank you for always being there and letting me know that

I’ll always have a home with you guys. To my brother, Billy, whom I’ve always looked up to as

a role model and who has always guided me through problems even when he is living halfway

across the world. To Karen, my sister-in-law, thanks for being the big sister that I never had!

And most importantly, to my parents who have provided a safe and wonderful environment for

my brother and me to grow up in, I don’t say this enough but I love the both of you very much.

Thanks for showing the good and bad of this world to me and allowing me to choose my own

walks of life. To Stephen’s family, thank you for taking care of me like I’m one of yours.

Lastly, to Stephen, thanks for keeping my head above the water and giving me a reality check

every once in a while to remind me what’s important in life. These pages are filled with your

love, patience, encouragement, and the occasional dose of your tasty mashed potatoes.

vi

TABLE OF CONTENTS

CHAPTER ONE – Overview of Perfluoroalkyl and Polyfluoroalkyl Substances 1

1.1 Overview 2

1.2 Industrial production and commercial applications of perfluoroalkyl and 4

polyfluoroalkyl substances

1.2.1 Electrochemical fluorination and telomerizaton 4

1.2.2 Application of fluorinated chemicals in commercial products - 6

Industrial trends and regulatory actions

1.3 Anthropogenic activities and use of commercial products as sources of 10

perfluoroalkyl and polyfluoroalkyl substances in the environment

1.3.1 Air-borne contamination with PFASs 11

1.3.1.1 Dust and indoor air 11

1.3.1.2 Outdoor air 14

1.3.2 Contamination in the aqueous environment 17

1.3.2.1 Surface water in freshwater, coastal, and marine bodies 17

1.3.2.2 Groundwater and drinking water 19

1.3.2.3 Wastewater treatment plant influents and effluents 20

1.3.3 Wastewater treatment plant sludge, sediments, and soil 21

1.3.4 PFAS contamination in humans 22

1.4 Fate of perfluoroalkyl and polyfluoroalkyl substances in the environment 25

1.4.1 Environmental and biological transformations 25

1.4.1.1 Atmospheric transformation of volatile polyfluoroalkyl 25

substances

1.4.1.2 Biological transformation of polyfluoroalkyl substances 31

1.4.2 Other environmental and biological processing of perfluoroalkyl and 37

polyfluoroalkyl substances

1.4.2.1 Environmental processes: Sorption and uptake into vegetation 37

1.4.2.2 Biological processes in aquatic organisms 37

1.4.2.2.1 PFAS contamination in aquatic wildlife 38

1.4.2.2.2 Bioaccumulation in aquatic organisms 42

1.4.2.2.3 Pharmacokineticsand distribution in aquatic 47

organisms

1.5 Goals and hypotheses 48

1.6 Literature cited 50

vii

CHAPTER TWO – Global Distribution of Polyfluoroalkyl and Perfluoroalkyl 80

Substances and their Transformation Products in Environmental Solids

Lee, H.; Mabury, S.A. Transformation Products of Emerging Contaminants in the Environment:

Analysis, Processes, Occurrence, Effects and Risks. 2012, to be submitted as a book chapter

2.1 Abstract 81

2.2 Introduction 81

2.3 Global contamination of PFASs in environmental solid matrices 84

2.3.1 Sediments 84

2.3.1.1 Temporal trends in sediment cores 88

2.3.2 Wastewater treatment plant sludge 90

2.3.3 Soil 92

2.3.3.1 Case study: Contamination of agricultural farmlands in Decatur, 94

Alabama

2.4 Fate of PFASs in environmental solids 97

2.4.1 Sorption 97

2.4.2 Leaching to surface waters and groundwater 100

2.4.3 Biodegradation in WWTP media and soils 101

2.4.4 Uptake into vegetation 101

2.5 Summary and future outlook 103


CHAPTER THREE – Biodegradation of Polyfluoroalkyl Phosphates (PAPs) as a 117

Source of Perfluorinated Acids to the Environment

Lee, H.; D’eon, J.; Mabury, S.A. Environ. Sci. Technol. 2010, 44, 3305-3310

3.1 Abstract 118

3.2 Introduction 118

3.3 Experimental section 120

3.3.1 Chemicals 120

3.3.2 Purging control experiment 120

3.3.3 Biodegradation experiments using aerobic WWTP microbes 122

3.3.4 Quality assurance of data 123

3.4 Results and discussion 124

3.4.1 Purging control experiment 124

3.4.2 Biodegradation of 6:2 monoPAP vs. 6:2diPAP (“Substitution study) 125

3.4.3 Biodegradation of the 4:2, 6:2, 8:2 and 10:2 monoPAP (“Chain length” 129

study)

3.5 Environmental implications 131

3.6 Acknowledgements 132


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CHAPTER FOUR – Biosolids Application as a Source of Polyfluoroalkyl 135

Phosphate Diesters and Their Metabolites in a Soil-Plant Microcosm:

Biodegradation and Plant Uptake

Lee, H.; Tevlin, A.G.; Mabury, S.A. Environ. Sci. Technol. 2012, to be submitted

4.1 Abstract 136



4.3.1 Materials 139

4.3.2 Soil-plant microcosm experiment 139

4.3.3 Sampling, extraction, and analysis 140


4.3.5 Data analysis 142


4.4.1 Amendment of WWTP biosolids and paper fiber biosolids as a source of 142

PFASs to soil

4.4.2 Metabolism of 6:2 diPAP in the soil-plant microcosm 146

4.4.3 Uptake and accumulation of PFCA metabolites in plants 149

4.5 Environmental implications 152



CHAPTER FIVE – Sorption of PerfluoroalkylPhosphonates and Perfluoroalkyl 158

Phosphonatesin Soil

Lee, H.; Mabury, S.A. Environ. Sci. Technol. 2012, to be submitted

5.1 Abstract 159



5.3.1 Chemicals 162

5.3.2 Soils used 162

5.3.3 Batch sorption experiments 163

5.3.4 Determination of distribution coefficients 164




5.4.1. Sorption kinetics and isotherms in different soils 167

5.4.2 Effect of soil properties on sorption 170

5.4.3 Effect of structural features on sorption and desorption 172

5.5 Implications for environmental distribution of PFPAs and PFPiAs 174



ix

CHAPTER SIX – Dietary Bioaccumulation of Perfluorophosphonates and

183Perfluorophosphinates in Juvenile Rainbow Trout: Evidence of Metabolism of

Perfluorophosphinates

Lee, H.; De Silva, A.O.; Mabury, S.A. Environ. Sci. Technol. 2012, 46, 3489-3497

6.1 Abstract 184



6.3.1 Chemicals 186

6.3.2 Food preparation 186

6.3.3 Fish care and sampling 187

6.3.4 Tissue distribution of PFPAs and PFPiAs 188

6.3.5 Extractions and instrumental analysis 188



6.3.8 Statistical analysis 190


6.4.1 Physical effects observed in fish 190

6.4.2 Uptake and depuration of PFPAs and PFPiAs 191

6.4.3 Assimilation of PFPAs and PFPiAs into different tissues 195

6.4.4 Effect of biotransformation on bioaccumulation parameters 196

6.5 Implications for environmental contamination 200



CHAPTER SEVEN – A Pilot Survey of Legacy and Current Commercial 207

Fluorinated Chemicals in Human Sera from United States Donors in 2009

Lee, H.; Mabury, S.A. Environ. Sci. Technol. 2011, 45, 8067-8074

7.1 Abstract 208


7.3 Materials and methods 210

7.3.1 Chemicals 210

7.3.2 Sera samples 212

7.3.3 Extractions and instrumental analysis 212


7.3.5 Statistical analysis 214


7.4.1 Concentrations in human sera 214

7.4.2 Detection of a new perfluorinatedacid in human sera 219

7.5 Current state of knowledge concerning exposure to commercial fluorinated 220

chemicals



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CHAPTER EIGHT – Summary, Conclusions, and Future Work 227

8.1 Summary and conclusions 228

8.2 Future research directions 232


xi

LIST OF TABLES

CHAPTER ONE

Table 1.1 Names, acronyms, and structures of perfluoroalkyl and polyfluoroalkyl

substances (PFASs) of interest 3

Table 1.2 Concentrations of volatile fluorinated species (pg/m3) in air samples collected

in WWTPs and over landfills 16

Table 1.3 Concentrations of PFSAs (C6 and C8) and PFCAs (C8–C12) in human blood,

sera, and plasma reported around the world 23

Table 1.4 Laboratory- and field-based metrics to evaluate bioconcentration,

bioaccumulation, and biomagnification of PFAAs 44

CHAPTER TWO

Table 2.1 Names, acronyms, and structures of PFASs 82

Table 2.2 Ambient concentrations of ΣPFCAs and ΣPFSAs (ng/g dry weight) observed

in freshwater, coastal, and marine sediments collected around the world 85

Table 2.3 Ambient concentrations of ΣPFCAs and ΣPFSAs (ng/g dry weight) reported

in selected WWTP monitoring campaigns conducted around the world 90

Table 2.4 Ambient concentrations of ΣPFCAs and ΣPFSAs (ng/g dry weight) reported

in selected soil monitoring campaigns conducted around the world 93

Table 2.5Organic carbon-normalized sorption distribution coefficients (logKOC) from

laboratory-based batch sorption experiments and field-based sediment and surface water

monitoring. Distribution coefficients in italics are not normalized to organic carbon

(logKd).

98

Table 2.6Plant-soil accumulation factors (PSAFs = Cplant/Csoil) calculated from the data

provided by Stahl et al. (128) and taken directly from Lechner and Knapp (38) and Yooet

al. (39).

102

CHAPTER THREE

Table 3.1 Structures, names, and acronyms of the target analytes in this study 121

CHAPTER FIVE

Table 5.1 Structures, full names, and acronyms of the target analytes monitored 161

CHAPTER SIX

Table 6.1 Structures, full names, and acronyms of the target analytes monitored 186

Table 6.2. Concentration of food (Cfood, in dry weight (dw)), depuration rate constant

(kd), depuration half-life (t1/2), assimilation efficiency (α), biomagnification factor (BMF)

of the dosed PFPAs and PFPiAs, and estimated time to achieve 90% steady state (tss).

The coefficient of correlation (r) for the linear regression analysis to determine kd is

shown in parentheses. The error is represented by ±1 standard error.

193

xii

CHAPTER SEVEN

Table 7.1 Structures, full names, and acronyms of the target analytes 211

xiii

LIST OF FIGURES

CHAPTER ONE

Figure 1.1 Industrial ECF production of perfluorooctanesulfonyl fluoride (POSF) and its

derivatives 5

Figure 1.2Telomerization production of fluorotelomer iodide (FTI) and its derivatives 6

Figure 1.3 Synthesis of perfluoroalkylphosphonates (PFPAs) and phosphinates

(PFPiAs) 9

Figure 1.4 Atmospheric transformation of volatile fluorotelomer-based precursors 26

Figure 1.5 Atmospheric transformation of volatile perfluoroalkanesulfonamido-based

precursors 30

Figure 1.6 Biological transformation of fluorotelomer-based precursors 32

Figure 1.7 Biological transformation of N-EtFOSE in rat subcellular fractions and

WWTP sludge 36

Figure 1.8 Global contamination of PFOS and PFOA in fish from selected data

summarized by Houdeet al. (174, 279) 39

Figure 1.9 Uptake and elimination processes of contaminants in fish 43

CHAPTER TWO

Figure 2.1 Environmental pathways of PFASs 83

Figure 2.2 Concentrations of diPAPs (ng/g) observed in WWTP sludge samples

collected from Ontario, Canada and in NIST SRM WWTP sludge samples 92

Figure 2.3 Concentrations of PFCAs, PFOS, and FTOHs observed in soils collected at

different depths in 2007 and 2009 from sludge-amended agricultural fields in Decatur,

Alabama. Data presented here were obtained from Washington et al. (101) and Yooet al.

(102)

95

Figure 2.4Concentrations of PFCAs observed in various plant species collected in 2009

from sludge-applied fields of Decatur, Alabama (left). Mean grass-soil accumulation

factors (GSAFs) calculated from five plant species (right). This data was obtained from

Yooet al. (39)

96

Figure 2.5 Concentrations of ΣPFCAs and ΣPFSAs observed in WWTP sludge collected

around the world. Note: Some of these concentrations were obtained by averaging total

concentrations reported in multiple monitoring campaigns within the same country to

yield an overall arithmetic mean for that country. *PFOS was the only PFAA monitored

in the Netherlands campaign; therefore, total ΣPFSA concentration = total PFOS

concentration

104

CHAPTER THREE

Figure 3.1 Proposed degradation pathway of 6:2 diPAP and 6:2 monoPAP. The solid

arrows represent pathways identified in this work. The dashed arrows represent microbial

and mammalian degradation pathways proposed in the literature

125

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Figure 3.2 Substitution study. (a) Degradation of 6:2 diPAP into 6:2 monoPAP and 6:2

FTOH, (b) Production of aqueous metabolites in 6:2 diPAP-dosed bottles, (c)

Degradation of 6:2 monoPAP into 6:2 FTOH, and (d) Production of aqueous metabolites

in 6:2 monoPAP-dosed bottles. Data are represented as arithmetic means (±standard

error) of triplicate incubations. Values less than the LOD are reported as zero and values

in between the LOD and LOQ were used unaltered and indicated with an asterisk (*) in

matching colours

127

Figure 3.3Chain length study. Degradation of (a) 4:2, (b) 6:2, (c) 8:2, and (d) 10:2

monoPAPs into FTOHs. Data are represented as arithmetic means (±standard error) of

triplicate incubations. Values less than LOD are reported as zero and values in between

the LOD and LOQ were used unaltered and indicated with an asterisk (*) in matching

colours

130

CHAPTER FOUR

Figure 4.1 Concentrations of diPAPs and PFCAs (ng/g) observed in control soil,

WWTP biosolids-amended soil, and WWTP biosolids- and paper fiber biosolids-

amended soil at 0, 3.5, and 5.5 months. Each data point represents the arithmetic mean

concentration of the triplicate (n = 3) sampling. The error bar represents the standard

error

144

Figure 4.2 Concentrations of 6:2 diPAP, 6:2 and 5:3 FTCAs and FTUCAs, C4–C7

PFCAs (ng/g) observed in soil and plants from 6:2 diPAP-supplemented microcosm at 0,

1.5, 3.5, and 5.5 months. Each data point represents the arithmetic mean concentration of

the triplicate (n = 3) sampling. The error bar represents the standard error

147

Figure 4.3 Correlation between the plant-soil accumulation factors (PSAFs, Cplant/Csoil)

and carbon chain length of the PFCAs analyzed in Treatments 2–4. Each data point

represents the arithmetic mean PSAF from averaging through individual PSAF measured

at each timepoint (1.5, 3.5, and 5.5 months). The error bar represents the standard error

151

CHAPTER FIVE

Figure 5.1 Sorption kinetics (left) of spiked PFPAs and PFPiAs displayed as their

percent mass fraction remaining in the aqueous phase upon equilibration with Soil A over

time. Sorption isotherms (right) of PFPAs and PFPiAs on Soil A. Each data point

represents the arithmetic mean of the triplicate (n = 3) samples. The error bar represents

the standard error

168

Figure 5.2 Dependence of logKOC on the number of perfluorinated carbons present in

PFSAs, PFCAs, PFPAs, and PFPiAs. LogKOC data for the PFSAs and PFCAs were

measured by Higgins et al. (16) and Ahrens et al. (19) in sediments

173

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Figure 5.3. Distribution of PFAAs in a simplified aquatic environment based on the

logKOC and logBMF measured for the PFPAs, PFPiAs, PFCAs, and PFSAs. LogKOC data

for the PFSAs and PFCAs were measured by Higgins et al. (16) and Ahrens et al. (19) in

sediments. LogBMF data for the PFSAs and PFCAs were measured by Martin et al. (24)

and logBMF data for the PFPAs and PFPiAs were measured by Lee et al. (25) in juvenile

rainbow trout

175

CHAPTER SIX

Figure 6.1. Growth-corrected whole-body homogenate concentrations (ng/g in wet

weight, (ww)) of C6, C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs in

rainbow trout during exposure and depuration phase. The top panels represent the data

collected from PFPA-dosed fish and the bottom panels represent the data collected from

PFPiA-dosed fish. Each data point represents the arithmetic mean concentration of the

triplicate (n = 3) sampling at each timepoint. The error bar represents the standard error

192

Figure 6.2. (A) Growth-corrected concentrations of PFPA metabolites (ng/g wet weight,

(ww)) observed in fish dosed with a mixture of C6/C6, C6/C8, and C8/C8 PFPiAs. (B)

Percent PFPA yield with respect to accumulated parent PFPiAs (mol basis) in fish dosed

with a mixture of C6/C6, C6/C8, and C8/C8 PFPiAs. Each data point represents the

arithmetic mean concentration of the triplicate (n = 3) sampling at each timepoint. The

error bar represents the standard error

197

Figure 6.3. Associations between the (A) depuration half-lives (t1/2) and (B) logBMFs

and the number of perfluorinated carbons present in PFSAs, PFCAs, PFPAs, PFPiAs, 8:2

FTAc, 8:2 FTCA, and 7:3 FTCA. Depuration half-lives and logBMFs for the PFSAs and

PFCAs were reported by Martin et al. (22). Note that the half-lives and logBMFs for 8:2

FTAc, 8:2 FTCA, and 7:3 FTCA were based on liver concentrations reported by Butt et

al. (61,62) and comparisons of these values to those of the other PFAAs should be treated

qualitatively.

199

CHAPTER SEVEN

Figure 7.1Arithmetic mean concentrations and standard error (μg/L) for all target

analytes detected in >20% of the single donor and pooled human sera samples (plotted on

a logarithmic scale). Note: Analytes denoted with an asterisk (*) were detected in <20%

of the samples, i.e. PFPeA (pooled); PFBS (single donor)

216

xvi

LIST OF APPENDICES

Appendix A – Supporting information for Chapter Three 238

Appendix B – Supporting information for Chapter Four 262

Appendix C – Supporting information for Chapter Five 277

Appendix D – Supporting information for Chapter Six 298

Appendix E – Supporting information for Chapter Seven 328

xvii

PREFACE

This thesis is organized as a series of manuscripts that have been published or are in preparation

for submission to be published in peer-reviewed scientific journals. As such, repetition of

introductory materials and methodology was inevitable. It should be noted that Chapters One

and Two together comprise the introduction to this thesis and for brevity, Chapter Two was

condensed from the original manuscript. All manuscripts were written by Holly Lee with critical

comments provided Scott Mabury. Contributions of all co-authors are provided in detail below.

Chapter One – Overview of Perfluoroalkyl and Polyfluoroalkyl Substances

Contributions – Prepared by Holly Lee with additional comments provided by Scott Mabury

Chapter Two – GlobalDistribution of Polyfluoroalkyl and Perfluoroalkyl Substances and their

Transformation Products in Environmental Solids

To be submitted to – As a book chapter to Transformation Products of Emerging Contaminants

in the Environment: Analysis, Processes, Occurrence, Effects and Risks

Author list – Holly Lee and Scott Mabury

Contributions – Prepared by Holly Lee with editorial comments provided by Scott Mabury

Chapter Three – Biodegradationof Polyfluoroalkyl Phosphates (PAPs) as a Source of

Perfluorinated Acids to the Environment

Published in – Environ. Sci. Technol. 2010, 44, 3305-3310

Author list – Holly Lee, Jessica D’eon, and Scott Mabury

Contributions – Prepared by Holly Lee with editorial comments provided by Jessica D’eon and

Scott Mabury. Holly Lee was responsible for designing and executing the biodegradation

experiments, LC-MS/MS method development, sample acquisition, and data interpretation.

Synthesis of the monoPAPs and diPAPs used for spiking in the biodegradation experiments and

the subsequent analysis of these chemicals by LC-MS/MS were performed by Holly Lee under

the guidance and training of Jessica D’eon.

xviii

Chapter Four - Biosolids Application as a Source of Polyfluoroalkyl Phosphate Diesters and

Their Metabolites in a Soil-Plant Microcosm: Biodegradation and Plant Uptake

To be submitted to – Environ. Sci. Technol.

Author list – Holly Lee, Alexandra G. Tevlin, and Scott Mabury

Contributions – Prepared by Holly Lee with editorial comments provided by Scott Mabury.

Holly Lee was responsible for designing the greenhouse microcosms in collaboration with Pablo

Tseng, performing the soil-plant biodegradation and uptake experiments, care and handling of

soil-plant systems during the experiment, method development, sample acquisition, and data

interpretation. Alexandra G. Tevlin assisted with sampling, extractions, and LC-MS/MS

analysis of plant samples with assistance from Holly Lee. Preparation of the manuscript by

Holly Lee involved adaptation of a report by Alexandra Tevlin.

Chapter Five – Sorption of PerfluoroalkylPhosphonates and PerfluoroalkylPhosphinates in Soil

To be submitted to – Environ. Sci. Technol.



Holly Lee was responsible for conceiving the experimental design, performing all sorption

experiments, method development, sample acquisition, and data interpretation.

Chapter Six – Dietary Bioaccumulation of Perfluorophosphonates and Perfluorophosphinates in

Juvenile Rainbow Trout: Evidence of Metabolism of Perfluorophosphinates


Author list – Holly Lee, Amila O. De Silva, and Scott Mabury

Contributions – Prepared by Holly Lee with editorial comments provided by Amila De Silva

and Scott Mabury. Holly Lee was responsible for conceiving the experimental design, care and

handling of animals during the experiments, performing all bioaccumulation experiments,

method development, sample acquisition, and data interpretation. Amila De Silva assisted in the

training of fish dissection and fish physiology.

xix

Chapter Seven – A Pilot Survey of Legacy and Current Commercial Fluorinated Chemicals in

Human Sera from United States Donors in 2009




Holly Lee was responsible for acquiring human sera samples, method development, sample

acquisition, and data interpretation.

Chapter Eight – Summary, Conclusions, and Future Work

Contributions – Prepared by Holly Lee with additional comments provided by Scott Mabury

xx

Other Publications DuringPh.D:

Rankin, K.R.; Lee, H.L.; Tseng, P.J.T.; Mabury, S.A. Investigating the Biodegradability of a

Fluorotelomer-Based Acrylate Polymer in a Soil-Plant Microcosm by Indirect and Direct

Analysis.2012, to be submitted to Environ. Sci. Technol.

Lee, H.L.; Rand, A.R.; D’eon, J. High Performance Liquid Chromatography-Tandem Mass

Spectrometry (HPLC-MS/MS) Analysis of Food Packaging Material as a Potential Source of

Human Exposure to Fluorochemicals: An Undergraduate Experiment.2012, to be submitted to J.

Chem. Ed.

xxi

GLOSSARY

This thesis describes the environmental chemistry of fluorinated chemicals used in commerce, those present as residual

impurities in commercial products, degradation intermediates, and terminal products. As such, the subsequent chapters involve the

use of numerous acronyms. This glossary provides a list of acronyms for the most commonly mentioned fluorinated chemicals, while

other fluorinated species will be specifically defined within each of the following chapters.

Perfluoroalkyl and polyfluoroalkylsubstance – PFAS

Perfluoroalkyl acid – PFAA Perfluorooctanesulfonyl fluoride – POSF

Perfluoroalkyl carboxylate – PFCA Perfluorooctane sulfonamide – FOSA

Perfluoroalkanesulfonate – PFSA N-Methyl perfluorooctane sulfonamide – MeFOSA

Perfluoroalkylphosphonate – PFPA N-Ethyl perfluorooctane sulfonamide – EtFOSA

Perfluoroalkylphosphinate – PFPiA N-Methyl perfluorobutanesulfonamidoethanol – MeFBSE

N-Ethyl perfluorobutanesulfonamidoethanol – EtFBSE

Fluorotelomer iodide – FTI N-Methyl perfluorooctanesulfonamidoethanol – MeFOSE

Fluorotelomer olefin – FTO N-Ethyl perfluorooctanesulfonamidoethanol – EtFOSE

Fluorotelomer alcohol – FTOH Perfluorooctanesulfonamidoacetate – FOSAA

Fluorotelomer acrylate – FTAC N-Methyl perfluorooctanesulfonamidoacetate – MeFOSAA

Fluorotelomer aldehyde – FTAL N-Ethyl perfluorooctanesulfonamidoacetate – EtFOSAA

Fluorotelomer unsaturated aldehyde – FTUAL N-Ethyl perfluorooctanesulfonamidoethyl

Fluorotelomer carboxylate – FTCA phosphate diester – SAmPAP

Fluorotelomer unsaturated carboxylate – FTUCA

Fluorotelomermercaptoalkyl phosphate diester – FTMAP

Fluorotelomersulfonate – FTSA

Polyfluoroalkyl phosphate ester – PAP

Polyfluoroalkyl phosphate monoester – monoPAP

Polyfluoroalkyl phosphate diester – diPAP

Polyfluoroalkyl phosphate triester – triPAP

1

CHAPTER ONE

Overview of Perfluoroalkyl and Polyfluoroalkyl Substances

Holly Lee and Scott A. Mabury

Contributions: Holly Lee prepared this chapter under the guidance of Scott Mabury

2

1.1 Overview

Perfluoroalkyl and polyfluoroalkyl substances (PFASs) are anthropogenic chemicals that

have a fluoroalkyl backbone (F(CF2)x) and a polar headgroup (R), both of which simultaneously

impart oleophobic and hydrophilic properties to these chemicals (1). The high surface activity of

these chemicals and their ability to repel water, oil, and stain has made them a crucial component

in non-stick, greaseproofing, and surface treatment applications. Commercial fluorochemical

production is largely dominated by high molecular weight (MW) polymers and surfactants (2, 3),

the latter of which will be the major focus of this thesis. However, the bulk of past scientific

research has predominantly focused on two classes of low MW perfluoroalkyl acids (PFAAs),

the perfluoroalkyl carboxylates (PFCAs) and perfluoroalkanesulfonates (PFSAs). PFCAs and

PFSAs have been ubiquitously detected in the environment despite their limited commercial use.

As they are fully fluorinated, they are recalcitrant to biological and environmental degradation

processes, but have themselves been observed as metabolites of commercial fluorinated

polymers and surfactants. The goal of this thesis is to investigate the distribution and fate of

commercial fluorinated surfactants as potential sources of the currently observed fluorochemical

contamination. Specifically, a number of biological (i.e. biotransformation, bioaccumulation)

and environmental (i.e. plant-soil uptake, sorption) processes are examined to characterize the

chemistry driving the distribution of these chemicals in the environment. Table 1.1 lists the

names, structures, and abbreviations of various PFASs that are of interest to this work.

3

Table 1.1 Names, acronyms, and structures of perfluoroalkyl and polyfluoroalkyl substances (PFASs) of interest.

Name Acronym Structure

Fluorotelomer-based Raw Materials

x:2 Fluorotelomer iodide x:2 FTI F(CF2)xCH2CH2I, x = 4, 6, 8, 10,…

x:2 Fluorotelomer olefin x:2 FTO F(CF2)xCH2=CH2, x = 4, 6, 8, 10,…

x:2 Fluorotelomer alcohol x:2 FTOH F(CF2)xCH2CH2OH, x = 4, 6, 8, 10,…

x:2 Fluorotelomer acrylate x:2 FTAC F(CF2)xCH2CH2OC(O)CH=CH2, x = 4, 6, 8, 10,…

Fluorotelomer-based Commercial Products

x:2 Polyfluoroalkyl phosphate monoester x:2 monoPAP F(CF2)xCH2CH2OP(O)O2-, x = 4, 6, 8, 10,…

x:2 Polyfluoroalkyl phosphate diester x:2 diPAP [F(CF2)xCH2CH2O]2P(O)O-, x = 4, 6, 8, 10,…

x:2 Polyfluoroalkyl phosphate triester x:2 triPAP [F(CF2)xCH2CH2O]3P(O), x = 4, 6, 8, 10,…

x:2 Fluorotelomermercaptoalkyl phosphate diester x:2 FTMAP [F(CF2)xCH2CH2SCH2]2(CCH2OP(O)(O

-)OCH2), x = 6, 8, 10,…

x:2 Fluorotelomersulfonate x:2 FTSA F(CF2)xCH2CH2SO3-, x = 4, 6, 8, 10,…

Fluorotelomer-based Biological and Environmental Transformation Intermediates

x:2 Fluorotelomer aldehyde x:2 FTAL F(CF2)xCH2CHO, x = 4, 6, 8, 10,…

x:2 Fluorotelomer unsaturated aldehyde x:2 FTUAL F(CF2)xCH=CHO, x = 4, 6, 8, 10,…

x:2 Fluorotelomer carboxylate x:2 FTCA F(CF2)xCH2CO2-, x = 4, 6, 8, 10,…

x:2 Fluorotelomer unsaturated carboxylate x:2 FTUCA F(CF2)x-1CF=CHCO2-, x = 4, 6, 8, 10,…

x–1:3 Fluorotelomer carboxylate x–1:3 FTCA F(CF2)x-1CH2CH2CO2-, x = 4, 6, 8, 10,…

x–1:3 Fluorotelomer unsaturated carboxylate x–1:3 FTUCA F(CF2)x-1CH=CHCO2-, x = 4, 6, 8, 10,…

PerfluoroalkaneSulfonamido-based Substances

Perfluorooctane sulfonamide FOSA F(CF2)8SO2NH2

N-Methyl perfluorooctane sulfonamide MeFOSA F(CF2)8SO2NH(CH3)

N-Ethyl perfluorooctane sulfonamide EtFOSA F(CF2)8SO2NH(CH2CH2)

N-Methyl perfluorobutanesulfonamidoethanol MeFBSE F(CF2)4SO2N(CH3)CH2CH2OH

N-Ethyl perfluorobutanesulfonamidoethanol EtFBSE F(CF2)4SO2N(CH2CH3)CH2CH2OH

N-Methyl perfluorooctanesulfonamidoethanol MeFOSE F(CF2)4SO2N(CH3)CH2CH2OH

N-Ethyl perfluorooctanesulfonamidoethanol EtFOSE F(CF2)4SO2N(CH2CH3)CH2CH2OH

Perfluorooctanesulfonamidoacetate FOSAA F(CF2)8SO2NH(CH2C(O)O-)

N-Methyl perfluorooctanesulfonamidoacetate MeFOSAA F(CF2)8SO2N(CH3)(CH2C(O)O-)

N-Ethyl perfluorooctanesulfonamidoacetate EtFOSAA F(CF2)8SO2N(CH2CH3)(CH2C(O)O-)

N-Ethyl perfluorooctanesulfonamidoethyl phosphate diester SAmPAP [F(CF2)8SO2N(CH2CH3)(CH2CH2O)]2P(O)O-

Perfluoroalkyl Acids (PFAAs)

Perfluoroalkyl carboxylate PFCA F(CF2)xCO2-, x = 1–13

Perfluoroalkanesulfonate PFSA F(CF2)xSO3-, x = 4, 6, 8, 10

Perfluoroalkylphosphonate CxPFPA F(CF2)xP(O)O2-, x = 6, 8, 10

Perfluoroalkylphosphinate Cx/CyPFPiA F(CF2)xP(O)(O-)((CF2)yF), x = 6, 8; y = 6, 8, 10, 12; x + y ≤ 18

4

1.2 Industrial Production and Commercial Applications of Perfluoroalkyl and

Polyfluoroalkyl Substances

1.2.1 Electrochemical Fluorination and Telomerization

Fluorochemical production has mainly proceeded by two manufacturing processes:

electrochemical fluorination (ECF) and telomerization (1).

3M Company was the major manufacturer of PFASs, who first employed ECF to produce

these chemicals in 1949, and remains active to date with three United States (U.S.)-based plants

in Minnesota, Illinois, and Alabama and one overseas plant in Antwerp, Belgium (2, 4, 5). In

ECF, a low-voltage electrical current (5–7 V) is applied to a hydrocarbon feedstock dissolved in

liquid anhydrous hydrogen fluoride to initiate fluorination whereby all hydrogen atoms in the

hydrocarbon are replaced by fluorine (1). From 1949 to 2002, ECF-based production largely

proceeded with the manufacture of perfluorooctanesulfonyl fluoride (POSF, F(CF2)8SO2F) as its

major starting material via fluorination of octane sulfonyl fluoride (H(CH2)8SO2F) (1, 2). POSF

functions as a basic building block where further derivatization of its sulfonyl fluoride moiety

would produce a suite of fluorinated materials with varying chemistries. Base-catalyzed

hydrolysis of POSF yields perfluorooctanesulfonate (PFOS, C8), while reactions with methyl

and ethyl amine yield N-methyl and N-ethyl perfluorooctane sulfonamide (MeFOSA and

EtFOSA) respectively, which may further react themselves with ethylene glycol carbonate to

form N-methyl and N-ethyl perfluorooctanesulfonamidoethanol (MeFOSE and EtFOSE)

respectively (Fig. 1.1) (1). Both FOSA and FOSE served as the primary starting materials of

3M’s fluorochemical production lines for surface treatments, paper and packaging protection,

and performance chemicals, as will be described in the next section (2). Perfluorooctanoate

(PFOA, C8) was similarly produced by ECF of octane acyl fluoride (H(CH2)7C(O)F), followed

by hydrolysis (1). As ECF is a relatively crude process (typically 34-40% yields of linear POSF)

(2), the final fluorinated products may be present as mixtures of odd, even, varying (C4–C9)

chain lengths, branched (30%) and linear (70%) isomers, and other byproducts (1, 2, 4, 6–8).

5

Figure 1.1 Industrial ECF production of perfluorooctanesulfonyl fluoride (POSF) and its

derivatives. H(CH2)8SO2F

F(CF2)8SO2F

ECF

F(CF2)8SO3-

PFOS

F(CF2)8SHO2N

H

CH3

F(CF2)8SHO2N

H

CH2CH3

MeFOSA EtFOSA

Hydrolysis CH3NH2 or CH3CH2NH2

OO

O

F(CF2)8SHO2N

CH2CH2OH

CH3

F(CF2)8SHO2N

CH2CH2OH

CH2CH3

MeFOSE EtFOSE

POSF

Telomerization was developed by E.I. du Pont de Nemours and Company in the 1940s (1,

9–11). The process begins by reacting pentafluoroethyl iodide (IF5) with tetrafluoroethylene

(CF2=CF2), in the presence of iodine (I2) and other catalysts to produce the telogen, n-

perfluoroethyl iodide (CF3CF2I) (Fig. 1.2) (1). Photochemically-catalyzed reactions convert the

telogen to a perfluoroalkyl radical (CF3CF2·), which may then iteratively react with the taxogen,

CF2=CF2, with the result of yielding a mixture of even-carbon-numbered telomer radicals of

varying chain lengths. Subsequent reactions of these radicals with I2 or perfluoromethyl iodide

(CF3I) produce a suite of perfluoroalkyl iodides (PFAIs, CF3CF2(CF2CF2)xI), where x depends on

the number of rounds of telomerization. The PFAIs are then converted via reactions with

ethylene (CH2CH2) to produce the x:2 fluorotelomer iodides (x:2 FTIs), which are the basic

building blocks for the production of fluorotelomer-based materials. The FTIs can be

functionalized to the corresponding alcohols (FTOHs), olefins (FTOs), thiols, thiocyanates, and

other functional groups, all of which may be used as intermediates for the production of

commercial fluorotelomer-based materials, as will be described next.

6

Figure 1.2 Telomerization production of fluorotelomer iodide (FTI) and its derivatives.

CF2=CF2

CF3CF2I

Telogen

IF5 I2

Taxogen

+ +

h

F(CF2)x

Perfluoroalkyl Radical

CF2=CF2Chain Propagation

I2 or CF3I

F(CF2)xI

Perfluoroalkyl Iodide

CH2=CH2

F(CF2)xCH2CH2I

Fluorotelomer Iodide (FTI)

F(CF2)xCH2CH2OH

Fluorotelomer Alcohol (FTOH)

F(CF2)xCH2CH2SH

Fluorotelomer Thiol

F(CF2)xCH2CH2SCN

Fluorotelomer Thiocyanate

F(CF2)xCH2=CH2

Fluorotelomer Olefin (FTO)Byproduct

F(CF2)8CO2-

Perfluorononanoate (PFNA)

Oxidation, x = 8

CO2/H2O, x = 8

F(CF2)7CO2-

Perfluorooctanoate (PFOA)

Oxidation, x = 8

Telomerization

1.2.2 Application of Fluorinated Chemicals in Commercial Products – Industrial Trends

and Regulatory Actions

During the period of 1949-2002, 3M Company was the dominant producer of POSF-

based materials and was responsible for ~80% (~4 million kg) of the total global production in

2000 (4). In 2000, the company decided to voluntarily phase out these chemicals due to

environmental concerns, with POSF-based production ceasing entirely in 2002 (12). 3M has

since transitioned their fluorochemical production to the perfluorobutyl-based chemistries (13).

Apart from the 3M plants in U.S. and Belgium, a number of other companies located in Italy,

Switzerland, United Kingdon, Brazil, Japan, China, India, and Russia have also been identified

7

as potential POSF producers, of which only Miteni S.p.A in Italy and Dainippon Ink &

Chemicals, Inc. in Japan have confirmed this independently (4).

Prior to the 1970s, production of POSF-based materials was low (5) until they replaced

the use of PFCAs in aqueous film-forming foams (AFFFs) for firefighting applications in the

1970s (14), from which point production increased by about five-fold between 1975 and 1989

(15), and remained relatively constant until their phase-out in 2000-2002. The majority of

POSF-based materials were produced by derivatizing FOSA and FOSE intermediates into high

molecular weight polymers and surfactants, which themselves may contain 1-2% of residual

PFOS, FOSAs, and FOSEs in the final products. Deliberate production of PFOS was estimated

to constitute only a very minor percentage (<0.5%) of the total production (4, 5, 16). PFOS was

primarily used in AFFF (2, 4, 17–20), but was also marketed under 3M’s Fluorad®

line of

performance chemicals as mining and oil surfactants, electronics and photography chemicals,

household cleaning and coating additives, chemical intermediates, and insecticide raw materials

(2, 4).

The remainder of 3M’s fluorochemical production is divided between the ScotchGard®

line of surface treatment chemicals (48% by weight) and the ScotchBan® line of paper and

packaging protectors (33% by weight) (2, 4, 8). The surface treatment products primarily

consisted of high molecular weight polymers, derivatized from MeFOSE acrylates

(F(CF2)8SO2N(CH3)CH2CH2OC(O)CH=CH2), and were used as protectors for carpets, fabric and

upholstery, apparel and leather, and other post market and consumer applications (2). The

ScotchBan® products were used as grease and water repellants in food contact paper and

packaging and also employed MeFOSE acrylate copolymers in this application, as well as,

mono- (10%), di- (85%), and tri- (5%) phosphate esters of EtFOSE (SAmPAPs) (2). The

SAmPAPs are of particular interest to this thesis and will be further discussed below and in

Chapter 7. Although these chemicals are currently regulated in the U.S. (17) and Europe (21,

22), POSF-based production still persists in Europe and Asia (4, 23). Since 2009, PFOS and

POSF have also been added to Annex B of the Stockholm Convention in which continued

production and use of these chemicals are regulated for specifically outlined purposes and

exemptions (24). In Asia, annual production has increased from <50 tons before 2004 to >200

tons from 2005 and onwards (25). POSF-based materials, including the SAmPAPdiester, are

currently commercially available from at least one Chinese manufacturer (26).

8

Fluorotelomer-based production began in the 1970s (27, 28) and has increased

significantly in the early 2000s (5-6 million kg/year in 2000-2002 (29)), presumably in response

to 3M’s phase-out of their POSF-based materials at that time. Approximately 80% of the

manufacturing process is directed towards the production of polymeric materials for surface

treatments of fabrics, carpets, and textiles, and the remaining 20% towards surfactants as

greaseproofing agents in food packaging and leveling and wetting agents in other household

products (29). Hydrolysis of FTIs yields FTOHs and to a lesser extent, the FTOs, as byproducts

(27). The majority of these FTOHs (~80%) are converted to the acrylate and methacrylate

monomers, which are then used as the building blocks for the synthesis of fluorotelomer-based

polymers. The remaining 20% are functionalized with different head groups to yield a suite of

fluorotelomer-based surfactants (3), such as the polyfluoroalkyl phosphate mono-, di-, and tri-

esters (mono-, di-, and triPAPs, collectively termed ―PAPs‖) (1, 30, 31). Alternatively, FTIs can

be converted to other surfactants, such as the fluorotelomersulfonates (FTSAs) and

fluorotelomerthiols (F(CF2)CH2CH2SH) (1), the latter of which are themselves intermediates and

may undergo further reactions to yield fluorotelomer-based surfactants for AFFF applications

and the fluorotelomer mercaptoalkyl phosphate diesters (FTMAPs) for oil repellency

applications (32). The application of these fluorinated surfactants, in particular the PAPs,

FTSAs, and FTMAPs, in commercial products will be a major focus of this work.

During fluorotelomer production, the polymeric materials and the surfactants are

primarily perfluorooctyl- and perfluorohexyl-based respectively (3, 29), but the final commercial

products are typically contaminated with a mixture of fluorotelomers of varying perfluoroalkyl

chain lengths (C4-C20) due to the inherent nature of the telomerization process. Nevertheless,

one of the major fluorotelomer-based manufacturers, DuPont, has transitioned from the

production of perfluorooctyl-based materials to the perfluorohexyl chain length, as evidenced

from the company’s new line of perfluorohexyl-based repellents and surfactants (33).

Direct application of ECF-based PFOA (1947-2002) was limited to its use as a

processing aid in the manufacture of fluoropolymers, such as polytetrafluoroethylene (PTFE) and

polyvinylidene fluoride (PVDF) (27, 34), while it is unclear where and in what capacity telomer-

based PFOA (2002–present), produced from the oxidation of perfluorooctyl iodide (F(CF2)8I)

(Fig. 1.2) (35), are currently used, as a number of fluorotelomer companies have asserted they do

not use PFOA in their manufacturing processes (34). Similarly, perfluorononanoate (PFNA,

9

C9), produced from either oxidation of 8:2 FTO (36) or carbonation of F(CF2)8I (Fig. 1.2) (37),

is only used in the manufacture of fluoropolymers. In contrast, perfluoroalkylphosphonates

(PFPAs) and phosphinates (PFPiAs) are the only high production volume PFAAs (4500–227000

kg/yr based on 1998 and 2002 data (38)) that currently have direct commercial applications as

leveling and wetting agents in household cleaning products (39), and historically as defoaming

agents in U.S. pesticide formulations (40), until their ban in this application in 2008 (41).

Synthesis of the PFPAs and PFPiAs begins with reacting PFAI with elemental phosphorus to

yield a mixture of perfluoroalkyl-phosphorus diiodides (F(CF2)xPI2) and di-(perfluoroalkyl)-

phosphorus iodides ([F(CF2)x]2PI) (42), which may then hydrolyze to produce

perfluoroalkylphosphonous (F(CF2)xP(OH)2) and phosphinous ([F(CF2)x]2P(OH)) acids

respectively (Fig. 1.3) (43). Further oxidation of these acid intermediates yields the

corresponding PFPAs and PFPiAs (Fig. 1.3) (43).

Figure 1.3 Synthesis of perfluoroalkylphosphonates (PFPAs) and phosphinates (PFPiAs).

F(CF2)xI

F(CF2)xPI2

Perfluoroalkyl Phosphorus Iodides

P

Perfluoroalkyl Iodide

+

Hydrolysis

Elemental Phosphorus

[F(CF2)x]2PI

Di-(Perfluoroalkyl) Phosphorus Iodides

Hydrolysis

F(CF2)xP(OH)2

Perfluoroalkyl Phosphonous Acids

Oxidation/Base

[F(CF2)x]2P(OH)

Perfluoroalkyl Phosphinous Acids

Oxidation/Base

F(CF2)xP(O)(O-)2

Perfluoroalkyl Phosphonate (PFPA)

[F(CF2)x]2P(O)(O-)

Perfluoroalkyl Phosphinate (PFPiA)

In 2004, the Canadian Ministers of Health and Environment imposed a temporary

prohibition on the import and manufacture of four new fluorotelomer-based polymers upon joint

10

assessment of their potential as PFCA sources (44). These prohibitions were due to expire in

2006-2007 and have since been extended as part of the Canadian action plan to prevent further

introduction of new fluorinated chemicals, especially those deemed as potential PFCA

precursors, into Canadian commerce (45). Further action to extend these prohibitions to other

fluorochemical substances, such as PFOA and long-chain PFCAs (≥9 perfluorinated carbons,

CFs) and their precursors, is currently under consideration (46, 47). Various PAP congeners and

the PFPAs and PFPiAs were among the 14 Domestic Substances listed as potential precursors to

long-chain PFCAs (47). Although PAPs are not regulated in Canada, the Minister of the

Environment has recently issued a significant new activity notice on these chemicals that

mandates importers and manufacturers to apply for approval for their use in all applications other

than those currently approved (48).

Environment Canada and Health Canada are currently working with various Canadian

fluorochemical companies (i.e. Arkema Canada Inc., Asahi Glass Company, Ltd., Ciba Canada

Ltd., Clariant Canada Inc., and E.I. du Pont Canada Company) towards reducing PFCA residuals

and their precursors that may be present as byproducts in the final sales products by 95% by the

end of 2010 and total elimination by the end of 2015 (49). In the U.S., a similar stewardship

program was established between the Environmental Protection Agency (EPA) and eight

fluorochemical manufacturers (i.e. Arkema, Asahi, BASF Corporation, Clariant, Daikin,

3M/Dyneon, DuPont, and Solvay Solexis), all of which have committed to reduce their

emissions and product content levels of PFCAs (≥7 CFs) and their precursors by 95% by 2010

and ultimately eliminate them by 2015 (50).

1.3 Anthropogenic Activities and Use of Commercial Products as Sources of

Perfluoroalkyl and Polyfluoroalkyl Substances in the Environment

Exposure to PFASs may occur through emissions of contaminated discharges from

fluorochemical manufacturers and the use and disposal of fluorinated consumer products. The

contribution of POSF-based and fluorotelomer-based materials to the observed burden of PFSAs

and PFCAs in the environment has been extensively discussed by Paul et al. (5) and

Prevedouroset al. (27). The following sections will discuss how anthropogenic activities,

including the use of commercial products, may contribute to the PFAS contamination observed

in different environmental compartments.

11

1.3.1 Air-borne contamination with PFASs

During fluorochemical production, the derivatization process of the starting raw materials

often carries forward unreacted materials or produces byproducts, both of which can be

incorporated into the consumer products at percent quantities (2, 27). Analysis of various

commercial fluorotelomer-based products revealed the presence of FTOHs as residual impurities

at 0.04–3.8%, while 0.4% of MeFOSE has been observed in a carpet protector from 3M (51).

PFAIs, FTIs, FTOs, FTOHs, fluorotelomer acrylates (FTACs), and other volatile fluorinated

impurities have also been observed in a commercial FTAC-based polymer (52), while 6:2, 8:2,

and 10:2 FTOHs have been measured at concentrations of 5–1200 µg/g in FTAC- and urethane-

based polymers (53). Prevedouros et al. estimated ~100 tons each of FTOHs and FTOs may be

present annually as residual materials in fluorotelomer-based products (27), and as such, the

release of these volatile materials from commercial products may represent a significant source

to the atmospheric fluorochemical burden. More importantly, human exposure may occur

through the offgassing of these materials from household commercial products, such as treated

carpets and home furnishings and paper products.

1.3.1.1 Dust and Indoor Air

Indoor measurements of volatile fluorinated species have recently been reviewed by

Harradet al. (54). PFAS contamination has been reported extensively in dust samples collected

from Canada (55–58), the U.S. (59), Sweden (60), Norway (61), and Japan (62, 63). The

majority of these samples were obtained from residential homes, offices, classrooms, daycare

centres, and cars. Concentrations of PFOA, PFOS, and perfluorohexanesulfonate (PFHxS, C6)

are typically within the mid ng/g to low µg/g range, although PFSA concentrations tend to be

higher than those reported for PFOA (55, 57, 59). As was suggested in all of these studies (55,

57, 59, 60, 62), the observed correlations among the concentrations of PFOA, PFOS, and PFHxS

point to a common exposure source, such as carpet and upholstery stain-repellents. Positive

correlations observed between the dust concentrations of PFAAs and the percentage of carpeting

found in Ottawa homes further support fluorochemically-treated carpets being a potential source

of the observed contamination (55).

Consistent with their lack of direct applications, detection of long chain PFCAs (≥8 CFs)

is occasional and if present, their concentrations are usually less than those observed for PFOA,

12

PFOS, and PFHxS (57, 59). In contrast, recent analysis of vacuum cleaner dust sampled from

Japanese homes revealed a distinct PFCA congener profile in which the odd carbon-chain

PFCAs (i.e. PFNA, perfluoroundecanoate (PFUnA, C11), and perfluorotridecanoate (PFTrA,

C13)) were present at higher concentrations than the adjacent, even-carbon PFCAs (i.e.

perfluorodecanoate (PFDA, C10), perfluorododecanoate (PFDoA, C12), and

perfluorotetradecanoate (PFTeA, C13)) (63). This is consistent with the manufacture of PFNA

in Japan in which fluorotelomer olefins are oxidized to produce odd carbon-chain PFCAs (27).

As discussed above, commercial ECF-based chemicals may contain 1-2% of residual

impurities like PFOS, FOSAs, and FOSEs (2). The statistical association between PFOS and

PFHxS is consistent with the presence of PFHxS as a potential residual byproduct in POSF-

based materials (64) and the use of perfluorohexanesulfonylfluoride (PHSF) to synthesize

perfluorohexyl-based materials for postmarket carpet treatment applications (2, 64). Similarly,

the observed correlation between PFOA and the two PFSAs may be due to the presence of PFOA

as a byproduct in POSF-based materials and/or ECF-based production of PFOA occurring in

tandem with that of POSF- and PHSF-based materials (27). PFOS and PFOA are among a

number of fluorinated additives that may be incorporated in aqueous stain-repellent emulsions

for treating carpets, upholstery, and home textiles (65). Furthermore, PFOA concentrations of 1–

39 µg/g have been measured in various fluorotelomer acrylate- and urethane-based polymers

used in surface treatment applications (53), and are similar to those reported in commercial

fluorotelomer-based formulations (1–80 µg/g), but typically 1–2 orders of magnitude higher than

those measured in the final treated carpets, apparel, and textiles (0.02–2 µg/g) (66, 67).

However, the demonstrated ability of residual volatile fluorinated materials to offgas from

products over time may be a more important contributor to indoor PFAS contamination due to

their percent quantities in commercial polymeric- and surfactant-based products (2, 51).

Shoeibet al. reported the first indoor air measurements of FTOHs in North America from

air samples collected in homes in Ottawa, Canada in 2002-2003 (68), followed by a second set of

measurements from Vancouver homes in 2007-2008 (57). In both studies, 8:2 FTOH was the

dominant FTOH congener observed (261–28900 pg/m3, Ottawa; 660–16080 pg/m

3, Vancouver),

followed by 10:2 FTOH (104–9210 pg/m3, Ottawa; 220–8160 pg/m

3, Vancouver) and 6:2 FTOH

(<LOD–22890 pg/m3, Vancouver) (57, 68). These concentrations were similar to those reported

13

by Barber et al. in Trømso, Norway (114 pg/m3, 4:2 FTOH; 2990 pg/m

3, 6:2 FTOH; 3424 pg/m

3,

8:2 FTOH; 3559 pg/m3, 10:2 FTOH) (69), Haug et al. in Norwegian houses (0.70–38 pg/m

3, 4:2

FTOH; 63–9414 pg/m3, 6:2 FTOH; 921–25323 pg/m

3, 8:2 FTOH; 377–28898 pg/m

3, 10:2

FTOH) and Fraser et al. in office environments in Boston, U.S. (<LOD–11000 pg/m3, 6:2

FTOH; 283–70600 pg/m3, 8:2 FTOH; 138–12600 pg/m

3, 10:2 FTOH) (70). In all of these

studies, FTOHs were observed as the dominant species, followed by the FOSEs and FOSAs.

From 2002 to 2009, a decline was observed in the indoor air concentrations of MeFOSE

and EtFOSE, consistent with the phase-out of POSF-based materials in 2002. Shoeib et al.

observed MeFOSE and EtFOSE at concentration ranges of 366–8190 pg/m3 and 227–7740

pg/m3, respectively in samples collected in 2002-2003 (56), while Barber et al. observed

MeFOSE at 6018 pg/m3 and EtFOSE at 5755 pg/m

3 in samples collected in 2005 (69). By

contrast, MeFOSE and EtFOSE concentrations were much lower in samples collected between

2007 and 2009, where Shoeib et al., Haug et al., and Fraser et al. all reported similar levels of

MeFOSE (289–380 pg/m3) and EtFOSE (18–97 pg/m

3) in indoor air sampled from Vancouver,

Norway, and Boston, respectively (57, 61, 70).

In most of these studies, positive correlations were observed in the concentrations within

the FTOHs and FOSEs themselves, but not between the two classes of compounds (57, 68, 70),

which suggests they may have separate sources. However in one study, EtFOSE indoor air and

dust concentrations did not correlate with those of MeFOSE and any of the other FOSAs

measured (57). These results suggest EtFOSE may be deriving from alternative applications,

such as the SAmPAPs used in food contact papers (2). In addition, indoor air concentrations

typically exceeded those measured outdoor by 1 to 2 orders of magnitude, which is consistent

with the extensive use of these chemicals in the indoor environment (2).

The distribution of FTOHs and FOSEs is preserved in the dust samples, but their

concentrations are typically lower than the PFOA, PFOS, and PFHxS observed in the same

samples (57, 59, 61). Air-dust partitioning coefficients, calculated from paired air and vacuum

dust samples collected from the same indoor environment (56), have been shown to significantly

underpredict dust concentrations of MeFOSE and EtFOSE. This discrepancy suggests the

observed dust contamination may not be limited to partitioning of volatile contaminants, but also

from the presence of other tightly bound FOSEs and/or FOSAs (56) and/or precursor materials

present in the commercial products used in homes. In fact, De Silva et al. recently reported the

14

detection of 8:2 diPAP at concentrations ranging from 2000 to 60000 ng/g in the same dust

samples collected by Shoeib et al. in 2007-2008 in Vancouver homes (57), as well as, the 6:2 and

10:2 diPAPs (58). DiPAPs are established biological precursors of PFCAs in mammalian

systems (71, 72), as will be discussed in the later sections. In the same samples, the PFPAs (3–

200 ng/g) and PFPiAs (0.1–52 ng.g) were also observed (58). These results suggest the wide

applicability of these fluorinated surfactants in commercial applications, as PAPs are not

exclusively used as greaseproofing agents in food contact papers (73, 74), but may also be found

in cosmetics, hair and personal care products, floor waxes, paints and finishes, and cleaning

fluids (75–79). Similarly, the PFPAs and PFPiAs may be found in waxes and coatings and

household cleaning products (39).

1.3.1.2 Outdoor Air

Atmospheric concentrations of volatile fluorinated species have been widely measured

(56, 57, 69, 80–87) and these data have been reviewed by Young and Mabury (88); therefore,

this section will only focus on air-borne contamination observed in near-source regions.

Air samples collected within a wastewater treatment plant (WWTP) and two landfills in

Ontario, Canada exhibited concentrations of volatile fluorinated species that were 4–11 times

and 2–36 times higher than those measured in background sites respectively (89). Total FTOH

concentrations (ΣFTOH) ranged from 1518 to 23706 pg/m3, while ΣFOSA+FOSE concentrations

were lower and ranged from 21 to 124 pg/m3 (Table 1.2). Air samples collected above the

primary clarifier and the aeration tanks typically exhibited higher concentrations of FTOHs,

FOSAs, and FOSEs than those collected above the secondary clarifier, which suggests these

chemicals or their fluorinated precursors may be degrading as the wastewater passes through the

microbially-enriched aeration tanks. Biodegradation of fluorinated chemicals will be discussed

in Section 1.4. Measurements performed in another Ontario WWTP showed similar

concentrations of FOSAs, FOSEs, and FTOHs (90), while much lower concentrations of these

analytes and other species, such as the FTACs, N-methyl perfluorobutanesulfonamidoethanol

(MeFBSE), and N-methyl perfluorobutane sulfonamide (MeFBSA), were observed in two

WWTPs near Lüchow, Germany (Table 1.2) (91). This may reflect geographical differences in

the composition of commercial products being used in North America and Europe.

15

In air samples collected at two landfills, ΣFTOH (2588–25994 pg/m3) and

ΣFOSA+FOSE (63–114 pg/m3) concentrations were 5–36 and 2–3 times higher than those

measured upwind of these sites (517–723 pg/m3, ΣFTOH ; 35–41 pg/m

3, ΣFOSA+FOSE) (Table

1.2) (89). These emissions likely derive from either the offgassing of residual fluorinated

materials present in the waste products deposited in the landfills or the degradation of fluorinated

precursors present in these products as they age. Together with the estimated annual emissions

of 2560 g/year for the WWTP and 99–1000 g/year for the two landfills, these results suggest

WWTPs and landfills may be important contributors to the atmospheric burden of PFASs.

Proximity to nearby manufacturers may also represent a major source of volatile

fluorinated chemicals to the atmosphere, as was observed in air samples collected near a

fluorotelomer production plant in China (92). Significant air contamination of PFAIs (1410–

30800000 pg/m3) and FTIs (1390–1320000 pg/m

3) was observed over various sampling sites in

the plant area (92). The dominance of perfluorooctyl iodide (PFOI) and perfluorohexyl iodide

(PFHxI) in these air samples, followed by perfluorodecyl iodide (PFDeI), and perfluorododecyl

iodide (PFDoI), is consistent with the perfluorooctyl- and perfluorohexyl-based chemistries that

are preferred by the fluorotelomer industry (3). As the PFAIs are synthetic precursors to the

FTIs (1), a similar distribution was observed in the FTI concentrations where 8:2 FTI was

dominant, followed by 6:2 and 10:2 FTIs. Following the phase-out of POSF-based materials in

2002, annual production of PFAIs increased dramatically to 4500 to 4.5 million kg per year (38),

while 5 to 6 million kg per year of FTIs were produced in 2000-2002 (29). Young et al.

demonstrated the conversion of FTIs to perfluoroalkyl aldehydes (PFALs), an atmospheric

precursor to PFCAs, may occur on a timescale of 5–10 days, which is sufficient time for these

volatile precursors to travel over long distances. (93). As such, fluorochemical emissions from

production plants are not restricted to local contamination, but may impact farther locations via

long-range transport.

16

Table 1.2 Concentrations of volatile fluorinated species (pg/m3) in air samples collected in WWTPs and over landfills.

Analyte

Air Concentrations (pg/m3)

Background

Sites for

WWTPs

WWTP Sampling Sites Background Sites

for Landfills

Landfill

Sampling Sites Location

Primary Clarifier Aeration Tank Secondary Clarifier

6:2 FTOH 90–605 5870–12286 2619–7739 895–1191 169–244 987–6462

Ontario,

Canada;

2009

(89)

8:2 FTOH 144–474 3413–10309 1597–3696 498–691 223–339 1290–17381

10:2 FTOH 70–115 521–1111 260–449 125–157 125–140 310–2151

MeFOSE 1–5 32–36 14–40 5–8 8–10 21–42

EtFOSE <LOD 15–20 6–18 <LOD 9–11 15–29

MeFOSA 6–8 14–15 16–48 8.8–9.9 9–11 15–27

EtFOSA 6–7 10–11 11–30 7–8 8–9 11–16

4:2 FTOH nd - nd–7 - - -

Lüchow,

Germany;

2009

(91)

6:2 FTOH 4–45 - 12–259 - - -

8:2 FTOH 7–176 - 36–419 - - -

10:2 FTOH 3–58 - 11–77 - - -

12:2 FTOH 2–24 - 6–34 - - -

6:2 FTAC nd–13 - nd–11 - - -

8:2 FTAC nd–6 - nd–49 - - -

10:2 FTAC nd–3 - 2–56 - - -

MeFBSE nd–5 - nd–7 - - -

MeFOSE nd–4 - 1–7 - - -

EtFOSE nd–3 - nd–9 - - -

MeFBSA nd–4 - nd–61 - - -

FOSA nd - nd - - -

MeFOSA nd–10 - 4–54 - - -

EtFOSA 1–9 - 3–69 - - -

6:2 FTOH - - 11000–12000 670–910 - -

Ontario,

Canada;

2010

(90)

8:2 FTOH - - 5700–5800 310–350 - -

10:2 FTOH - - 780–860 41–48 - -

MeFOSE - - 16–18 4.5–4.9 - -

EtFOSE - - 8.5–9.4 1.8–2.3 - -

FOSA - - 5–10 nd - -

MeFOSA - - 13–14 0.8–1 - -

EtFOSA - - 5.7–5.9 1.2–1.6 - -

17

1.3.2 Contamination in the Aqueous Environment

1.3.2.1 Surface Water in Freshwater, Coastal, and Marine Bodies

Contamination of PFASs has been observed in freshwater (94–108), coastal (102, 104, 109),

and marine (110–113) water bodies. Concentrations are typically in the tens to hundreds ng/L

range in freshwater and coastal systems, while oceanic concentrations are typically 1 order of

magnitude lower in the tens to hundreds pg/L range. Yamashita et al. showed that PFAA

concentrations may drastically differ between those measured in coastal waters and in the open

ocean, as exemplified by the high concentrations of PFOA (1800–192000 ng/L) and PFOS (338–

57700 ng/L) observed in Tokyo Bay and the much lower concentrations in Central Pacific Ocean

(15–62 pg/L, PFOA; 1.1–20 pg/L, PFOS) (110). Oceanic contamination appears to decrease from

the northern to southern hemisphere based on comparisons of PFAA concentrations measured in the

North, Baltic, and Norwegian Seas (10–4810 pg/L, PFOA; nd–6160 pg/L, PFOS) (113) with those

measured in equatorial Atlantic Ocean (100–439 pg/L, PFOA; 37–73 pg/L, PFOS) (110), south of

Australia (<5–11 pg/L, PFOA; <5–21 pg/L, PFOS) (111), and near Antarctica (<LOQ, PFOA and

all other PFCAs; 5–23 pg/L, PFOS) (111). Recently, Benskin et al. measured PFAAs in the North

and Southwestern Atlantic Ocean and the Canadian Arctic archipelago and also observed a general

decline in PFAA concentrations with latitude (114). ΣPFAA concentrations ranged from 280 to

980 pg/L in the Bay of Biscay and the Canary Islands and decreased to <210 pg/L in the southern

hemisphere, except for one hotspot location (350–540 pg/L) near Rio de la Plata, which was

attributed to the continued use of Sulfluramid (EtFOSA) in South America and proximity to urban

cities like Montevideo and Buenos Aires (114).

PFOA and PFOS are typically the dominant congeners observed, with exceptions in some

studies, as was observed by Simcik and Dorweiler who attributed the dominance of

perfluoroheptanoate (PFHpA, C7) in Lake Calhoun, Minnesota to nearby WWTP inputs (97), and

Nakayama et al. who ascribed the prevalence of perfluorobutanoate (PFBA, C4) in the Upper

Mississippi River Basin to local production of perfluorobutyl-based materials (106).

Proximity to urban development and industrialization is often associated with

fluorochemical contamination of nearby surface waters. The highest PFOS concentrations (198–

1090 ng/L) that have ever been measured in New York State waters were observed in Lake

Onondaga, a Superfund site located near several industries that receives a significant portion of a

18

local WWTP’s outflow (99). Similar contamination hotspots in the vicinity of heavy development

and industrialization have also been identified in Japan (96), Korea (100, 109), Hong Kong (109),

and China (109, 111). Point source contamination has also been identified in surface water sampled

in the Tennessee River located near a fluorochemical plant in Decatur, Alabama (94), in the Ruhr

and Moehne rivers located near WWTP biosolids-applied agricultural fields (101), in a creek that

received AFFF-contaminated effluents from an airport spill (115, 116), in ponds and streams near

farmlands in Decatur, Alabama that had received >10 years of WWTP biosolids (117), and in

monitoring wells and runoff water basins within a fluorochemical industrial park (118, 119).

Although total fluorine analysis revealed that seawater is predominantly composed of

inorganic fluoride (>90%), a significant proportion (60–90%) within the extractable organic

fluorine (EOF) fraction was not accounted for by the concentrations of known PFASs measured

(120). This suggests the presence of other unidentified fluorinated species in water. D’eon et al.

reported the first detection of PFPAs, a new class of PFAAs, in 80% of Canadian surface waters

sampled and in six of seven WWTP effluents sampled (105). The perfluorooctyl congener (C8

PFPA) was the predominant congener observed in surface water (88–3400 pg/L) and WWTP

effluents (760–2500 pg/L), followed by C6 PFPA (26–1200 pg/L, surface water; 330–6500 pg/L,

WWTP effluent) and C10 PFPA (41–870 pg/L, surface water; 380–460 pg/L, WWTP effluent)

(105). Unlike the PFCAs and PFSAs, PFPAs have no known precursors; therefore, the

contamination observed was presumably due to direct input. As mentioned above, the PFPAs are

applied in household cleaning products (39), and historically, in pesticide formulations (40) until

2008 in the U.S. (41). Given the surface water samples were collected in 2005 and 2007 during

which the use of PFPAs as inert additives in pesticides was still permitted, the contamination

observed in these samples may be due to extensive pesticides application in nearby agricultural

sites.

Perfluoro-4-ethylcyclohexanesulfonate (PFECHS), an ECF-based product by 3M, was

detected for the first time in the Great Lakes at concentrations ranging from 0.16 to 5.7 ng/L,

similar to those observed for PFOA (0.65–5.5 ng/L) (108). PFECHS was marketed for use as an

erosion inhibitor in aircraft hydraulic fluids (121), although the commercial formulation typically

contains other impurities, such as perfluoro-4-methylcyclohexane sulfonate (PFMeCHS), which

was also detected in the surface water at concentrations of 0.2–0.4 ng/L (108). Although

production of PFECHS has ceased since 3M’s phase-out of POSF-based materials, the use of this

19

chemical in aircraft hydraulic fluids is still permitted in Canada and the U.S. due to the lack of

alternatives and its anticipated minimal release to the environment (17). The detection of PFPAs

and PFECHS in surface waters represents the first environmental measurements of these new

classes of PFAAs in Canada and everywhere else to date.

1.3.2.2 Groundwater and Drinking Water

Groundwater contamination of PFASs has been attributed to the use of AFFFs during fire-

training exercises at nearby military bases (122, 123), wastewaters and street runoffs (124), and

proximity to nearby biosolids-applied farmlands (117). Both Moody et al. (122) and Schultz et al.

(123) observed similar concentrations of PFCAs and PFSAs (3–213 ng/L) in the groundwater wells

around the decommissioned Wurtsmith Air Force Base in northeast Michigan, but Schultz et al.

reported additional detection of the 4:2 FTSA (nd–7.3 ng/L), 6:2 FTSA (nd–14600 ng/L), and 8:2

FTSA (nd–17 ng/L) at this and two other bases (123). Murakami et al. observed 0.28–133 ng/L of

PFOS, 0.47–60 ng/L of PFOA, and 0.1–94 ng/L of PFNA as the major PFAAs in groundwater

collected in the Tokyo metropolitan area and estimated that 54–86% and 16–46% of this

contamination were due to wastewater and street runoffs respectively (124). In Decatur, Alabama,

groundwater sampled from 21 different farms that practiced biosolids application, some for as many

as 12 years, was significantly contaminated with PFAAs (117). In the most contaminated well,

PFCA concentrations ranged from 1260 ng/L of PFBA to 6410 ng/L of PFOA, all of which

exceeded U.S. EPA’s provisional health advisory level of 400 ng/L of PFOA in drinking water

(125). This contamination, also observed in soil (126, 127) and plants (128) collected from the

impacted fields, was traced back to the source of the biosolids, a local WWTP that had processed

effluents from nearby fluorochemical industries. This contamination is of concern as groundwater

supplies a substantial amount of water to both private wells and public drinking water facilities.

In fact, drinking water contamination has been observed in the U.S. (129, 130), Canada

(129), India (129), Japan (96, 129, 131), China (129, 132), and the Netherlands (133).

Concentrations are typically in the low ng/L range, except for those measured in drinking water

collected near known point sources. Skutlarek et al. reported ΣPFAA concentrations of 20–598

ng/L in drinking water sampled in the Rhine-Ruhr area, where the Ruhr and Moehne rivers

exhibited concentrations as high as 446 ng/L and 4385 ng/L of ΣPFAA due to contamination from

nearby biosolids-applied fields (101). Similar concentrations were observed in the drinking water

20

collected in Arnsberg, a district sandwiched between the Ruhr and Moehne rivers, where local

residents exhibited 4–8 times higher PFAA concentrations in their blood plasma, as compared to the

reference German population (134). The highest concentrations of PFOA (1500–7200 ng/L) ever

reported in U.S. public drinking water supplies were measured in the Little Hocking water system

in 2002–2005, which is situated close to a fluoropolymer manufacturing plant (135).

1.3.2.3 Wastewater Treatment Plant Influents and Effluents

In near-source regions, domestic, commercial, and industrial discharges have been identified

as a major source of PFASs to the wastewater environments.

Analysis of various consumer products revealed low levels of PFOA in PTFE-coated

cookware (4–75 ng/g), PFTE-based dental floss and tape (3–4 ng/g), PFTE sealant films (1800

ng/g), and popcorn bags (6290 ng/g) (136). PFOA was also observed at concentrations of 300–

1200 ng/g in a number of fluorochemically-treated food contact paper, such as containers and

wrappers for popcorn, muffins, croissants, hamburgers, sandwiches, French fries and pizza box

liners (137). Fluorinated greaseproofing agents, such as the PAPs, SAmPAPs, and FTMAPs, have

a demonstrated capacity to migrate out of microwaveable popcorn bags upon heating (136, 137).

The FTMAPs have been observed at 1400–3900 ng/g in post-heated microwaveable popcorn bags

purchased from the U.S. market (137), while recent surveys of European food contact materials

have also revealed the presence of PAPs and FTMAPs (138, 139). Treated carpeting, upholstery,

home and technical textiles, medical garments; stone, tile, and wood sealants; floor waxes and

paints; and home and office cleaners have also been shown to contain PFOA at concentrations as

high as 2000 ng/g (66). In addition to PFOA, the C5-C12 PFCAs have also been detected in a vast

array of U.S. consumer articles in the tens to thousands ng/g concentration range (67, 140), while

FTOHs have been measured in various household products (141) and in the gas phase released from

the use of PFTE-coated nonstick pans (142).

Such widespread use of fluorinated chemicals in industrial and commercial applications has

resulted in prevalent contamination of wastewaters in North America (105, 143–146), Europe (147–

149), and Asia (103, 104, 150–156). Concentrations vary between tens to thousands ng/L and

depend on proximity to nearby sources. For example, higher ΣPFAA concentrations were

measured in the influents (7–629 ng/L) and effluents (16–599 ng/L) of industrial WWTPs that

processed sewage from pharmaceutical, paper, and battery industries in Korea, as compared to

21

those measured (8–133 ng/L, influent; 7–101 ng/L, effluent) in municipal WWTPs that primarily

received domestic waste (152).

PFAA concentrations have been observed to increase from WWTP influent to effluent

(143–146, 148, 150–152, 154–156). In one of two WWTPs studied (145), significant correlation

was observed between the mass flows of PFOA and PFNA and between PFDA and PFUnA

following activated sludge treatment. In all wastewater samples, the concentrations of the even-

chain PFCAs were higher than those measured for the odd-chain lengths. This even>odd carbon

PFCA pair pattern is consistent with the biological production of PFCAs from fluorotelomer-based

materials in microbial and animal systems (157–165). The detection of fluorotelomer saturated

(FTCAs) and fluorotelomer unsaturated (FTUCAs) carboxylates in WWTP sludge and effluents

(145, 166, 167), both of which are intermediate metabolites observed during fluorotelomer

degradation to the PFCAs, further supports precursors are present in WWTPs. This has been

corroborated by the detection of diPAPs and FTSAs, both of which are established PFCA

precursors (71, 72, 168), in WWTP samples (143, 144, 169).

N-methyl and N-ethyl perfluorooctanesulfonamidoacetate (MeFOSAA and EtFOSAA),

perfluorooctanesulfonamidoacetate (FOSAA), and perfluorooctane sulfonamide (FOSA) are

intermediate metabolites observed during the transformation of perfluorooctanesulfonamido-based

precursors to PFOS, and have themselves been detected in WWTP media (143, 144, 148, 166, 170).

These intermediates typically exhibit increased mass flows from influent to effluent (143, 144),

consistent with their production from degradation occurring in the WWTP, while PFOS has been

shown to display inconsistent mass flows in different studies. In general, PFOS concentrations

have been observed to increase from influent to effluent (144–146, 148, 150, 151), but a number of

studies have also reported the opposite, in which removal of the chemical was attributed to sorption

to the sludge co-generated at these facilities (143, 152, 155, 156).

1.3.3 Wastewater Treatment Plant Sludge, Sediments, and Soil

Upon entering a WWTP, PFASs may either travel through effluent emissions to downstream

water bodies or sorb to the sludge generated at the facility. Disposal of the treated sludge or

biosolids may further transport the PFASs to landfills or agricultural farmlands during land

application. The global contamination of PFAS in WWTP sludge, sediments, and soil and a

discussion of their environmental pathways will be reviewed in Chapter 2.

22

1.3.4 PFAS Contamination in Humans

WWTP contamination of PFASs suggests humans may be exposed to fluorinated chemicals

as WWTPs primarily receive anthropogenic emissions and are generally considered as a useful

proxy for characterizing human exposure. Taves first reported the presence of organofluorine in

human blood in 1968 (171), but it was not until 2001 when the development of electrospray tandem

mass spectrometry (ESI-MS) allowed Hansen et al. to specifically identify PFOS, PFOA, and

PFHxS in human sera (172). Since then, numerous studies have reported the global detection of

PFASs in human blood, plasma, and sera, with PFOS generally observed as the dominant congener,

followed by PFOA and PFHxS. This profile corroborates the relatively long half-lives of these

chemicals in humans (5.4 years, PFOS; 3.8 years, PFOA; 8.5 years, PFHxS) (173). Human blood

contamination of PFASs has been reviewed by Houdeet al. in 2006 (174) and Vestergren and

Cousins in 2009 (175) respectively. D’eon et al. also recently examined various direct and indirect

exposure sources as potential contributors to human contamination (176).

PFASs in human blood are typically observed at µg/L concentrations. North American

populations (15, 64, 169, 172, 177–184) typically exhibit similar or higher concentrations than

those measured in Europe (70, 134, 178, 185–189), Asia (178, 190–192), and the southern

hemisphere (Table 1.3) (178, 193). Concentrations in developing nations typically range at the

lower end of those measured in industrialized countries (tens to hundreds µg/L), as was observed in

Colombia (8 µg/L, PFOS; 6 µg/L, PFOA; whole blood concentrations corrected to serum

concentrations by multiplying by 2) (178), Malaysia (13 µg/L, PFOS; 6 µg/L PFOA) (178), and

India (1–7.8 µg/L, PFOS; < 3–9.5 µg/L, PFOA) (178, 191), which indicates partial similarity in

human exposure to PFAS around the world, although this is highly dependent on the presence of

local emission sources.

A number of studies have demonstrated temporal PFAS trends in human blood that

correspond to the changes that have occurred in the fluorochemical manufacturing industry in the

past several decades (182, 183, 194–196, 188, 197, 198). Comparison of American Red Cross

blood data measured between 2000–2001 and 2006 revealed significant declines in the

concentrations of PFOS (-60%, 34 to 15 µg/L ), PFHxS (-30%, 2.2 to 1.5 µg/L), PFOA (-27%, 4.7

to 3.4 µg/L), PFHpA (-31%, 0.13 to 0.09 µg/L), and PFBA (-87%, 5.3 to 0.33 µg/L), while

concentrations of PFNA (+70%, 0.57 to 0.97 µg/L), PFDA (+112%, 0.16 to 0.34 µg/L), and PFUnA

(+80%, 0.10 to 0.18 µg/L) were observed to increase (183, 194). Similar declines ranging from 10

23

Table 1.3 Concentrations of PFSAs (C6 and C8) and PFCAs (C8–C12) in human blood, sera, and plasma reported around the world.

Matrix PFOA PFNA PFDA PFUnA PFDoA PFHxS PFOS Location, Year Reference

Sera 6.4 - - - - 6.6 28 USA, N/A (172)

Sera 4.6 - - - - 1.9 35 USA, 2000-2001 (64)

Sera and Plasma 2.4 - - - - 1.7 29 USA, 1974

(15) 5.7 - - - - 2.4 33 USA, 1989

Sera 4.2 - - - - 2.3 31 USA, 1999 (199)

Plasma 3.4 - - - - 1.5 15 USA, 2006 (183)

Sera 5.3 0.6 - - - 2.2 31 USA, 1999-2000 (200)

Sera 3.7 0.6 - - - 2.8 21 USA, 2001-2002 (180)

Sera 3.9 1.0 0.8* 0.6* - 1.9 21 USA, 2003-2004 (182)

Sera 4.4 0.8 0.2 - - - - USA, 2004-2005 (181)

Sera 4.2 0.6 0.2 0.1 - - 16.3 USA, 2004-2005

(169) 1.7 1.2 0.5 0.3 - - 9.9 USA, 2008

Sera 2.5 0.9 - - - 4.1 18.3 Urban Canada, 2004-2005 (184)

Plasma - - - - - 19 Remote Canada, 2004 (201)

Sera 2.2 0.8 0.2 0.1 <LOQ 0.8 9.4 Norway, 2007 (188)

Whole blood 1.6 - - - - 1.9 53 China, 2004 (192)

Whole blood nd-2.3 - - - - - 8.2 Japan, 2003 (190)

Sera 9.5 0.4 0.2 0.3 0.02 0.8 7.8 Urban Sri Lanka, 2003 (191)

Sera 0.5-9.1 0.04-0.09 0.02-0.05 0.04 0.002-0.008 0.1-0.8 1.0-6.3 Rural Sri Land, 2003

Sera 7.2 - - - - 7.6 22 Australia, N/A (193)

*95th Percentile

24

to 32% were also observed for PFOS, PFHxS, and PFOA in U.S. human sera samples analyzed

between 1999–2000 and 2003–2004 during the National Health and Nutrition Examination Survey

(NHANES), while concentrations of PFNA were observed to double (182). The substantial

declines observed for PFOS and PFHxS in these time periods corroborate the phase-out of POSF-

and PHSF-based materials in 2000–2002 (12). However, the relatively smaller decline observed for

PFOA and the fact that long chain PFCAs have become increasingly prevalent in human blood,

suggest the phase-out of ECF-based PFOA production may have only partially contributed to its

reduction, while an alternative exposure pathway, such as that from fluorotelomer-based products,

may be contributing to current exposure to PFOA and the long chain PFCAs. The fact that

telomerization often results in mixtures of different perfluoroalkyl chain lengths in the final

commercial products may also account for the distribution of different PFCAs observed.

Other studies covering a larger span of time (1960s–2010) have also reported similar

temporal trends for the PFSAs and PFCAs, as described above (188, 196, 197). Haug et al.

observed a 9-fold increase in the sera concentrations of a Norwegian population for PFOS and

PFOA between 1977 and the mid-1990s, followed by a decline between 2000 and 2007 (188).

Concentrations of PFNA, PFDA, and PFTrA were significantly correlated with those of PFOS and

PFOA, and were also observed to increase starting from 1977 to 2007 (188). This suggests humans

were historically exposed to PFOS and PFCAs at the same time, perhaps due to the concurrent

production of POSF- and fluorotelomer-based materials that began in the early 1950s and 1970s

respectively, until 2000-2002 at which point telomerization took over as the dominant

manufacturing process. Similarly, Wang et al. observed significant contamination of PFOS (42

µg/L, 1960s; 29 µg/L, 1980s) and to a lesser extent, PFHxS (1.6 µg/L, 1960s; 1 µg/L, 1980s) and

PFOA (0.3 µg/L, 1960s; 2.7 µg/L, 1980s) in Californian women blood sampled prior to the phase-

out, followed by lower concentrations (9 µg/L, PFOS; 0.9 µg/L, PFHxS; 2.1 µg/L, PFOA)

measured in 2009 (196). Analysis of sera collected from Swedish women shortly after their first

labour revealed doubling times of 6–18 years in the concentrations of PFBS, PFHxS, PFNA, and

PFDA between 1996 and 2010, while PFOSA, PFOS, PFDS, and PFOA were observed to eliminate

with half-lives of 3.1, 8.2, 6.6, and 22 years respectively (197).

In contrast, a temporal survey of a Chinese population in Shenyang, China revealed

increasing prevalence of PFOS and PFOA in human blood (0.0313 µg/L, PFOS; 0.073 µg/L,

PFOA) between 1987 and 2002 (198). After adjusting to sera concentrations, whole blood

25

concentrations of PFOS measured two years later in the same city were significantly higher (142

µg/L, males; 170 µg/L, females), although PFOA concentrations appeared to have declined (1.3

µg/L, males; 0.8 µg/L, females) (192). Contrary to the temporal trends observed in North American

(182, 183, 194–196) and European (188, 197) studies, the observed increase in human PFOS

contamination in China may be associated with the resurgence of POSF-based production in Asia in

the early 2000s in response to local and overseas demands that were no longer being met by North

American and European markets (25). Nevertheless, the consistency observed between the

temporal trends in human blood concentrations and changes in the production industry is evidence

of human exposure to PFASs through the use of commercial fluorinated products.

In addition to PFCAs and PFSAs, metabolic intermediates of perfluorooctanesulfonamido-

based materials, FOSA, FOSAA, MeFOSAA, and EtFOSAA have also been detected in human

blood (15, 64, 172, 180, 182, 183), whereas, FTCAs and FTUCAs have never been detected in this

matrix. Conversely, varying perfluoroalkyl chain lengths of the fluorotelomer-based diPAPs has

been observed at low µg/L concentrations, which represents the first observation of a commercial

fluorinated product in human sera (169).

1.4 Fate of Perfluoroalkyl and Polyfluoroalkyl Substances in the Environment

1.4.1 Environmental and Biological Transformations

1.4.1.1 Atmospheric Transformations of Volatile Polyfluoroalkyl Substances

The atmospheric transformation of volatile fluorinated chemicals has been reviewed by

Young and Mabury (88); therefore, a brief overview of the atmospheric chemistry of volatile

fluorotelomer- and perfluoroalkanesulfonamide-based substances will be presented here.

Smog chamber studies of FTIs (93), FTOHs (202–204), FTOs (205, 206), and most

recently, FTACs (207) have demonstrated their potential as atmospheric precursors of PFCAs.

These are all synthetic precursors of fluorotelomer-based materials, and have been measured as

residual impurities in a commercial fluorotelomer polymer (52). As shown in Fig. 1.4, the

atmospheric transformation of these various precursors begins differently until the formation of a

common intermediate, PFAI (F(CF2)xC(O)H), at which point the mechanism proceeds identically

towards PFCA formation. For FTI, FTOH, and FTAC, a separate pathway may also occur whereby

each precursor transforms to the fluorotelomer aldehyde (F(CF2)xCH2C(O)H, x:2 FTAL), which can

26

Figure 1.4 Atmospheric transformation of volatile fluorotelomer-based precursors.

F(CF2)xCH2CH2OH

x:2 FTOH

F(CF2)xCH2C(O)H

+OH/-H2O

+O2/-HO2

F(CF2)xCH2C(O)OO

x:2 FTAL

+OH/-H2O

+O2/-HO2

F(CF2)xCH2C(O)OH

x:2 FTCA +HO2/-O3 +RO2/-RO or

+NO/-NO2

F(CF2)xCH2C(O)O

-CO2

F(CF2)xCH2

+O2

F(CF2)xCH2OO

+h/-HCO

+RO2/-RO

+O2/-HO2

F(CF2)xC(O)H

PFAL

+OH/-H2O

+O2

F(CF2)xC(O)OO

+HO2/-O3

F(CF2)xC(O)OH

PFCA +RO2/-RO or

+NO/-NO2

F(CF2)xC(O)O

-CO2

F(CF2)x

+h/-HCO

+O2F(CF2)x or yOO

F(CF2)x or yO +RO2/-RO or

+NO/-NO2

F(CF2)y

-COF2

y = x-1, x-2, x-3,...

+O2

F(CF2)x or yOH

+RHO2/-RCHO

y = x-1, x-2, x-3,...

F(CF2)(x or y)-1C(O)F-HF

F(CF2)(x or y)-1C(O)OHH2O

PFCA

F(CF2)xCH2CH2I

h/-I

+O2

+RO2/-RO

+O2/-HO2

x:2 FTI

F(CF2)xCH2=CH2

+OH/-H2O

+RO2/-RO

-CH2OHx:2 FTO

F(CF2)xCH2CH2OC(O)CH=CH2

x:2 FTAC

F(CF2)xCH2CH2OC(O)C(O)H

x:2 FTGly

+O2

+NO/-NO2

1. +h/-HCO

2. +O2

3. +NO/-NO2/-CO2

4. +O2/-HO2

27

undergo further reaction itself to ultimately yield FTCA. The PFAL is expected to predominantly

photolyze to form a perfluoroalkyl radical (F(CF2)x·) (208), although it may also oxidize to form a

perfluoroalkyl acyl peroxy radical (F(CF2)xC(O)OO·), which itself may further react with HO2 to

yield a Cx+1 PFCA (F(CF2)xC(O)OH; e.g. 8:2 fluorotelomer precursor PFNA) or with an alkyl

peroxy radical (RO2) to yield the perfluoroalkyl radical (202).

Reaction of the perfluoroalkyl radical with oxygen would yield a perfluoroalkylperoxy

radical (F(CF2)xOO·), which in the presence of nitrogen oxides (NOx) under typical urban

atmospheric conditions, would further transform to a perfluoroalkoxy radical (F(CF2)xO·). This

perfluoroalkoxy radical can iteratively lose carbonyl fluoride (COF2) to produce perfluoroalkyl

radicals (F(CF2)x-1,x-2,x-3,…·) that are one carbon atom shorter, with this reaction cycling between

these three species until the perfluoroalkyl chain has fully unzipped to COF2. Further reactions of

these perfluoroalkyl radicals of varying chain lengths would yield a suite of Cx, Cx-1, Cx-2,… PFCAs

of different chain lengths (e.g. 8:2 FTOH trifluoroacetate (TFA, C2), perfluoroproprionate

(PFPrA, C3), PFBA, perfluoropentanoate (PFPeA, C5), perfluorohexanoate (PFHxA, C6), PFHpA,

PFOA, PFNA (202)). Alternatively, in rural environments where NOx concentrations are lower, the

perfluoroalkylperoxy radical may react with alkyl peroxy radicals with a hydrogen present on the

carbon alpha to the radical, to form a perfluoroalkyl alcohol (F(CF2)xOH), followed by

heterogeneous elimination to a perfluoroalkyl acyl fluoride (F(CF2)x-1C(O)F). Further hydrolysis of

the acyl fluoride yields the Cx PFCA (F(CF2)x-1C(O)OH).

These mechanisms are dependent on the concentration of NOx. In the presence of excess

NOx in polluted air, the overall yield of PFCA products may be suppressed due to competition

between perfluoroalkyl-based peroxy radicals and NOx for available alkyl peroxy radicals.

However, in rural locations where NOx concentrations are much lower, HO2 and other

peroxyradicals (RO2) may become more central to drive the atmospheric formation of PFCAs. For

example, Ellis et al. reported the production of PFOA (1.5% yield) and PFNA (1.5% yield), and a

suite of shorter chain PFCAs (TFA, PFPrA, PFBA, PFPeA, PFHxA, PFHpA, <0.5% yield

collectively) from smog chamber oxidation of 8:2 FTOH in the absence of NOx (202). This

suggests species other than NOx, such as HO2 and/or RO2, are capable of driving atmospheric

formation of PFCAs in low NOx environments. This has major implications for FTOHs and other

volatile fluorotelomer-based species that are capable of long-range transport, as potential precursors

to PFCAs observed in remote locations, such as the Arctic environment (209).

28

Neither photolysis nor wet and/or dry deposition is expected to be a significant loss process

for FTOHs as they do not absorb actinic radiation (210) and their lifetimes with respect to

atmospheric deposition ranges from 8 years (dry deposition) to 2.5 million years (wet deposition)

(211). As such, the dominant tropospheric fate of FTOHs is reaction with hydroxyl (·OH) radicals,

with a calculated lifetime of 20 days that is independent of the perfluoroalkyl chain length of the

parent FTOH (211). Within this timescale, FTOHs emitted from an urban point source can travel as

far as 7000 km based on the global average wind speed of ~14 km/h (211). Similarly, the lifetimes

of FTIs (1–7 days with respect to photolysis and reactions with ·OH (93)) and FTOs (8 days with

respect to reactions with ·OH and ozone (212)) may be sufficient to allow for their transport to rural

environments, while FTACs are rather short-lived (atmospheric lifetime ~1 day with respect to

reactions with ·OH (207)) and may not be able to travel as far.

Considerably less is known about the atmospheric chemistry of volatile

perfluoroalkanesulfonamide-based precursors, with only two studies published to date. Martin et

al. reported an atmospheric lifetime of 20–50 days for N-ethyl perfluorobutane sulfonamide

(EtFBSA, F(CF2)4SO2NH(CH2CH2)), with respect to reactions with ·OH radicals (213). The main

products of chlorine (Cl)-initiated oxidation of EtFBSA were a ketone (F(CF2)4SO2NHC(O)CH3),

an aldehyde (F(CF2)4SO2NHCH2C(O)H) (213), and an intermediate identified as F(CF-

2)4SO2N(C2H5O)- by high-resolution mass spectrometry (213), but later confirmed as F(CF-

2)4SO2N(OH)(CH2CH2) by theoretical studies (214). Together with the detection of COF2 and

sulfur dioxide (SO2) by Fourier transform infrared (FTIR) spectroscopy, the observed production of

TFA, PFPrA, and PFBA (~0.5% yield) suggests the carbon-sulfur (C–S) bond in the intermediate

species of EtFBSA, F(CF2)4SO2, may have cleaved to yield SO2 and a perfluoroalkyl radical,

F(CF2)4· (Fig. 1.5.) (213). Subsequent reactions of the perfluoroalkyl radical via the unzipping

cycle, as described above, would yield the observed short-chain PFCAs. Perfluorobutanesulfonate

(PFBS) was not detected as a product of F(CF2)4SO2, presumably due to the absence of ozone and

NOx in the smog chamber experiments (213), both of which have been reported to react with the

methanesulfonyl radical (CH3SO2) to yield the analogous methanesulfonic acid (CH3SO3H) (215).

In contrast, MeFBSE degrades faster in the atmosphere via reactions with ·OH radicals, with

a calculated lifetime of 2 days (216). The observed products from OH- and Cl-initiated oxidation of

MeFBSE included an aldehyde (F(CF2)4SO2N(CH3)CH2C(O)H), MeFBSA, PFBS, TFA, PFPrA,

and PFBA. The mechanism by which PFBS and the short-chain PFCAs were formed was proposed

29

to occur by an initial addition of an ·OH radical to the sulfone (S=O) double bond of MeFBSE to

yield an unstable sulfonyl radical, followed by scission of either the C–S or sulfur-nitrogen (S–N)

bond (Fig. 1.5.) (216). Cleavage of the S–N bond would yield PFBS, while cleavage of the C–S

bond would yield a perfluorobutyl radical that can undergo the same unzipping cycle, as described

above, to form PFBA, PFPrA, and TFA (216). The higher yield of PFCAs (10%), as compared to

that of PFBS (1%), was due to the formation of a more stable perfluorocarbon-centered radical

(F(CF2)x or y·) from C–S bond scission than the nitrogen-centered radical (·N(CH3)CH2CH2OH)

(216). These results represent the sole observation of PFSA formation from the atmospheric

breakdown of a volatile fluorinated precursor.

Given the short lifetime of MeFBSE (2 days), this compound is unlikely to travel far from

its point of emission, but its breakdown product, MeFBSA, may presumably have a similar

atmospheric lifetime to that calculated by Martin et al. for EtFBSA (i.e. 20–50 days) (213).

Considering MeFBSA has even fewer abstractable hydrogens than EtFBSA, MeFBSA is likely to

be less reactive with ·OH radicals. Similar to FTOHs, FTIs, and FTOs, MeFBSA and EtFBSA, and

their homologues (MeFOSA and EtFOSA), are expected to be sufficiently long-lived to travel and

contribute to the atmospheric burden observed in distant locations. This has been corroborated by

the atmospheric detection of FBSA and FOSA over the Atlantic Ocean (83), in the Canadian Arctic

(217), and Mace Head, Ireland (218).

In addition to fluorotelomer- and perfluoroalkanesulfonamido-based species, atmospheric

PFCA formation may derive from other precursors. Recent work by Jackson et al. reported minor

production of PFPrA from the hydrolysis of perfluoro-2-methyl-3-pentanone (PFMP), a fire-

fighting fluid marketed as Novec 1230 by 3M (219). The atmospheric lifetime of PFMP is

approximately 1-2 weeks, with photolysis solely dominating the breakdown of PFMP to COF2,

TFA, and PFPrA (219–221). Hydrolytic degradation of PFMP to PFPrA and HFC-227ea

(CF3CFHCF3), a long-lived greenhouse gas, was observed at environmentally relevant pH (5.6–

8.5), but this pathway was considered a negligible sink for PFMP due to the low proportion of

liquid water comprising the atmosphere (219).

30

Figure 1.5. Atmospheric transformation of volatile perfluoroalkanesulfonamido-based precursors.

F(CF2)xSO2N(R)CH2CH2OH

N-FAFSE

F(CF2)x-S-N(R)CH2CH2OH

+OH/-H2O

F(CF2)xSO3H

+O2

F(CF2)x or yOO

F(CF2)x or yO

-COF2

F(CF2)x or yOH

+RHO2/-RCHO

y = x-1, x-2, x-3,...

-HF

F(CF2)(x or y)-1C(O)OH

+H2O

PFCA

F(CF2)xSO2N(R)CH2C(O)H

O

O OH

+OH/-H2O

+O2/-HO2

F(CF2)xSO2NHR

+OH/-H2O

N-FAFSA-N(R)CH2CH2OH

PFSA

F(CF2)x or y

y = x-1, x-2, x-3,...

y = x-1, x-2, x-3,...

+RO2/-RO or

+NO/-NO2

y = x-1, x-2, x-3,...

-HOSO2N(R)CH2CH2OH

F(CF2)xSO2

-SO2

F(CF2)(x or y)-1C(O)F

31

1.4.1.2 Biological Transformations of Polyfluoroalkyl Substances

As PFCAs and PFSAs are perfluorinated, the strength in their carbon-fluorine (C–F) bonds

renders them recalcitrant to normal environmental and biological degradation processes (222–225).

During anaerobic incubations with WWTP sludge, PFOA (224, 225) and PFOS (223–225) were

observed to disappear over time, but this removal was not accompanied by a corresponding

detection of metabolites and/or fluoride ions released from their mineralization. As such, sorption

and bioaccumulation are likely the primary fate of these persistent chemicals. Much less is known

about the degradability of the other class of PFAAs, the PFPAs and PFPiAs. In a letter addressed to

the Office of Pesticides Program (226), the U.S. EPA cited concerns over the potential of PFPAs

and PFPiAs to biologically or abiotically degrade to other PFAAs, like the PFCAs. In fact, the

Canadian government has recently listed the PFPAs and PFPiAs of varying perfluoroalkyl chain

length as potential precursors to long-chain (≥8 CFs) PFCAs (47), but no investigations of their

biotransformation have been performed to date. This will be discussed in Chapter 6.

Numerous studies have demonstrated the biotransformation of fluorotelomer-based

substances to PFCAs in microbial and soil systems (52, 168, 227–237) and in vitro (158, 162, 165)

and in vivo (71, 72, 158–161, 163, 164, 238–240) animal models, with the majority of this work

centered on the FTOHs as the parent reactant. Butt et al. has previously reviewed these

biotransformation pathways in detail (240, 241); therefore, only a brief overview will be presented.

All of the x:2 FTOH-based biotransformation work has shown the production of Cx PFCA

and to a smaller extent, the Cx+1 PFCA and shorter-chain Cx-1,x-2 PFCAs (e.g. 8:2 FTOH PFOA

(C8), PFNA (C9), PFHxA (C6) and PFHpA (C7)) (Fig. 1.6.). A number of studies have proposed

β-oxidation or a similar mechanism as the dominant driver for the production of Cx PFCAs from x:2

FTOH biotransformation in microbial (227, 229, 230, 233) and animal (158, 160, 164, 238, 240)

systems, while minor contribution from α-oxidation has also been observed, primarily in animal-

based studies (158–164, 240), to the production of Cx+1 PFCAs. The initial steps of the mechanism,

by which x:2 FTOH first oxidizes to the transient x:2FTAL, followed by further transformation to

first x:2 FTCA, then x:2 FTUCA, were first proposed in the early 1980s (238), and have since been

widely corroborated by a number of studies (Fig. 1.6.). However, subsequent biotransformation of

x:2 FTUCA has been observed to diverge into different pathways (Fig. 1.6.).

32

Figure 1.6 Biological transformation of fluorotelomer-based precursors.

(F(CF2)xCH2CH2)nOR

x:2 Fluorotelomer-Based Precursor

R = -SO3H, -P(O)O3-nH, -C(O)CH=CH2, -C(O)C17H35, -(CH2CH2O)nH,

and/or -C(O)C(-Rbackbone-)(-Rbackbone-)

F(CF2)xCH2CH2OH

x:2 FTOH

F(CF2)xCH2C(O)H

x:2 FTAL

F(CF2)xCH2C(O)OH

x:2 FTCA

F(CF2)x(CH2CH2O)n-1CH2C(O)OH

x:2 FTEOnC

R = -(CH2CH2O)nH

F(CF2)x-1CF=CHC(O)OH

x:2 FTUCA

F(CF2)xC(O)OH

Cx+1 PFCA

F(CF2)x-1CF=CHC(O)H

x:2 FTUAL

F(CF2)x-1C(O)CH2C(O)H

x-1:3 -keto aldehyde

F(CF2)x-1C(O)CH2C(O)OH

x-1:3 -keto acid

F(CF2)x-1CH2CH2C(O)OH

x-1:3 FTCA

F(CF2)x-2CH2CH2C(O)OH

x-2:3 FTCA

F(CF2)x-1CH=CHC(O)OH

x-1:3 FTUCA

F(CF2)x-1C(O)CH3

x-1:2 ketone

F(CF2)x-1C(OH)CH3

x-1:2 sFTOH

F(CF2)x-1CH(OH)CH2C(O)OH

3-OH-x-1:3 FTCA

F(CF2)x-3C(O)OH

Cx-2 PFCA

F(CF2)x-2C(O)OH

Cx-1 PFCA

F(CF2)x-1C(O)OH

Cx PFCA

R = -SO3H

The discovery of novel metabolites and subsequent incubation studies with these and other

previously identified intermediate metabolites (158, 162, 231, 232, 240, 242) have also further

complicated the overall proposed mechanism for x:2 FTOH biotransformation, especially in the

33

context of PFCA production for which contradictory pathways have been proposed in the published

literature.

While the metabolite profiles are typically conserved among the biotransformation of

different chain lengths of FTOHs (158), the product yields of different PFCA products may differ.

Biotransformation of 8:2 FTOH typically yields PFOA as the dominant metabolite, with PFNA and

the shorter-chain PFHxA and PFHpA observed at much lower yields. This was exemplified in the

soil biotransformation of 8:2 FTOH in which PFOA accounted for 40% of the mass balance,

followed by 7:3 FTCA (18%), PFHxA (1–4%), and PFHpA (<1%) (231). The 7:3 FTCA has also

been identified as a metabolite of 8:2 FTCA, 8:2 FTUCA, and 7:3 FTUCA based on individual

dosing of these FTOH intermediates with isolated animal and human hepatocytes and microsomes

(162), in aerobic soils (198), and through dietary exposure to rainbow trout (240). In contrast to the

8:2 FTOH soil biotransformation where PFOA was observed as the major metabolite (231), Liu et

al. reported a yield of only 8% for the analogous metabolite, PFHxA, from 6:2 FTOH degradation

in the same soil, while PFPeA was observed as the dominant metabolite (30%), followed by 5:3

FTCA (15%) and PFBA (2%) (232). Similar PFCA congener distribution was observed in the soil

biotransformation of 5:2 sFTOH and 5:2 ketone, both intermediates of 6:2 FTOH, in which the

corresponding Cx-1 PFCA (i.e. PFPeA) was observed in higher yields (18–85%) than those (4–12%)

of the Cx PFCA (i.e. PFHxA) (232). However, this distribution was not conserved when 6:2 FTOH

and 5:2 sFTOH were incubated with mixed bacterial cultures in which PFHxA was either observed

as the more prominent metabolite or at much closer yields, as compared to the PFPeA (232). These

results suggest the occurrence of certain biotransformation pathways or the extent to which they

occur may depend on the perfluoroalkyl chain length of the parent fluorotelomer reactant, as well

as, the incubation matrix (i.e. soil vs. isolated bacterial cultures).

Nevertheless, the yields of PFCA products from x:2 FTOH biotransformation tends to be

low, ranging from <1% to 30% (158, 160–163, 227–233). These low yields are likely attributed to

decreased bioavailability via sorption of the parent reactant and/or intermediate metabolites to the

experimental system (i.e. septa and surfaces of the incubation vessels, soil, and sludge), the

extensive branching in the overall degradation that could lead to other terminal products, and the

formation of phase II conjugates in animal models. The last phenomenon was specifically probed

by Rand and Mabury who observed the formation of glutathione conjugates with FTUCAs and

fluorotelomer unsaturated aldehydes (FTUALs), both of which have been observed as intermediates

34

of FTOH metabolism (243). Following this work, Rand and Mabury investigated the reactivity of

FTUCA and FTUAL with various nucleophilic amino acids and two model proteins (244). Adduct

formation was observed between the amino acids of interest and both FTUCAs and FTUALs,

although only the FTUALs were observed to exhibit reactivity with apomyoglobin and serum

albumin (244). Together, these results suggest conjugation of FTOHs and their intermediate

metabolites with small biological nucleophiles and proteins may be quantitatively important in the

mass balance of fluorotelomer biotransformation in animal-based studies.

Recent research efforts have been directed towards investigating the biological fate of other

fluorotelomer-based precursors, particularly those present as active and/or inert ingredients in

commercial applications. A number of commercial fluorotelomer-based materials, such as the 6:2

FTSA (168), fluorotelomer acrylate- and urethane-based polymers (52, 236, 237), fluorotelomer

ethoxylates (FTEOs) (235), and 8:2 fluorotelomer stearate monoester (FTS) (245), have a

demonstrated capacity to biodegrade in either soil and/or WWTP media, with all, but the FTEOs,

established as PFCA precursors. Incubations of FTEOs with WWTP effluents resulted in their

oxidation to the fluorotelomer ethoxylate carboxylates (FTEOCs) as the terminal metabolites, while

the observed formation of PFHxA and PFOA were ascribed to the degradation of residual 6:2 and

8:2 FTOHs present at 0.3–0.5% in the commercial FTEO product used for dosing (235).

In contrast, animal-based biotransformation has only been investigated for the mono- and

diPAPs in rats (71, 72) and 8:2 FTAC in trout (164, 165). D’eon and Mabury proposed the

biotransformation of 8:2 mono- and diPAP occurred by enzyme-mediated cleavage of the

phosphate ester bond to yield 8:2 FTOH, which would then further oxidize to PFOA, as was

observed in the exposed rats (71). Oral gavage and intravenous injection experiments were later

carried out for the 4:2, 6:2, 8:2, and 10:2 mono- and diPAPs in rats in which metabolites profiles

were generally conserved across the different chain lengths, although the bioavailability of diPAPs

from absorption of the gut was observed to decrease with increasing chain length (72). Overall, the

production of FTCAs, FTUCAs, and PFCAs were consistent with the metabolite profiles reported

in previous FTOH biotransformation, although FTOHs were not monitored in these experiments

(71, 72). In contrast, Butt et al. have shown that 8:2 FTAC is rapidly hydrolyzed to 8:2 FTOH

either in the gut or within the internal tissues of exposed rainbow trout, followed by subsequent

production of 8:2 FTCA, 8:2 FTUCA, 7:3 FTCA, 7:3 FTUCA, PFHpA, PFOA, and PFNA (164).

35

Except for the FTEOs, the initial breakdown of these commercial fluorotelomer-based

substances has either been postulated or observed to produce FTOHs as the immediate metabolite

(52, 71, 72, 164, 165, 236, 237, 245), from which point subsequent biotransformation typically

follows the pathways as described in Fig. 1.6.

Considerably fewer studies have investigated the biotransformation of

perfluoroalkanesulfonamido-based materials, with the majority focused on EtFOSA (246–251) and

EtFOSE (252–254). PFOS is typically observed as the terminal metabolite in the incubation of

EtFOSA with trout liver microsomes (250); EtFOSE with rat liver slices (252), the whole rat (253),

and WWTP sludge (254); and FOSA through dietary exposure to rats (255).

Arrendale et al. (246), Manning et al. (247), Grossman et al. (248), and Vitayavirasuk and

Bowen (249) all observed rapid biotransformation of EtFOSA, an active component of the

insecticide, Sulfuramid, to FOSA in dosed rats, dogs, and sheep, but none of these studies

monitored for the production of PFOS or PFOA. PFOS and PFOA concentrations were also not

observed above background levels during incubations of a technical EtFOSA (~60% linear isomers)

standard with human microsomes and recombinant cytochrome P450s (251). These results contrast

the production of PFOS observed during incubations of EtFOSA with rainbow trout liver

microsomes (250).

In contrast, PFOS is consistently detected as a metabolite of EtFOSE biotransformation

(252–254). Incubations of EtFOSE and several of its established metabolites with rat liver

microsomes, cytosol, and slices were performed separately to help elucidate an overall

biotransformation mechanism for EtFOSE (252) (Fig. 1.7.). A two-step enzyme-mediated

dealkylation was responsible for the initial transformation of EtFOSE to first the deethylated FOSE,

then subsequently to FOSA (252) (Fig. 1.7.). Both the EtFOSE and FOSE were observed to

undergo O-glucuronidation, while FOSA was observed to form N-glucuronide conjugates (252). In

addition, oxidation of EtFOSE and FOSE resulted in the production of the corresponding EtFOSAA

in liver slices and FOSAA in the cytosol respectively, with no further transformation of either of

these metabolites observed. This contrasts the biotransformation of EtFOSAA previously observed

in spiked WWTP sludge to EtFOSA (254) and dosed worms to FOSA and PFOS (256).

Contradictory mechanisms have been proposed for the formation of PFOS in FOSA-spiked

rat liver slices (252) and WWTP sludge incubated with EtFOSE, EtFOSAA, FOSAA, EtFOSA,

FOSA, and perfluorooctane sulfinate (PFOSi) (254). Xu et al. suggested PFOS formation observed

36

in the rat liver slices proceeds through a N-glucuronide conjugate of FOSA in which the amine

moiety is sequentially converted to an iminium ion upon which SN2 attack by a hydroxide ion

would yield PFOS (252). On the other hand, Rhoads et al. identified PFOSi, an intermediate

metabolite of EtFOSE, as the direct hydrolytic precursor to PFOS in WWTP sludge (254). Similar

to fluorotelomer biotransformation, the occurrence of certain pathways to yield specific

perfluorooctanesulfonamido-based intermediates is dependent on the incubation medium, but the

overall biotransformation of EtFOSE generally proceeds through FOSA formation, followed by

subsequent transformation of this intermediate to PFOS (Fig. 1.7.). However, a similar degradation

mechanism of the perfluoroalkyl chain to produce shorter chain PFCAs that was observed for

fluorotelomer biotransformation was not operative here as all the metabolites retained the

perfluorooctyl chain length of the parent reactant in their structures. The lack of any PFOA

formation in these studies also suggests the sulfonate moiety of perfluorooctanesulfonamido-based

precursors is recalcitrant to degradation.

Figure 1.7 Biological transformation of EtFOSE in rat subcellular fractions and WWTP sludge.

F(CF2)8SO2N(CH2CH2)CH2CH2OH

EtFOSE

F(CF2)8SO2NH(CH2CH2) F(CF2)8SO2N(CH2CH2)(CH2C(O)OH)

EtFOSAAEtFOSA

F(CF2)8SO2NH(CH2CH2OH)

FOSE

F(CF2)8SO2NH(CH2C(O)OH)

FOSAA

F(CF2)8SO2NH2

FOSA

F(CF2)8SO3H

PFOS

PFOSA N-glucuronide

PFOSi

X

X

?

Observed Pathways in Rat Liver Microsomes, Cytosol, and Slices

Observed Pathways only in Rat Liver Slices

Unconfirmed Pathways in Rat Liver Microsomes, Cytosol, and Slices

Unobserved Pathways in Rat Liver Microsomes, Cytosol, and Slices

Observed Pathways in WWTP Sludge

Minor Pathways in WWTP Sludge

?

X

37

In vitro rat hepatocyte incubations of a commercial mixture, composed of primarily the

difluoroalkylated phosphate ester of EtFOSE (SAmPAPdiester) (~80%) and the monoalkylated

congener (SAmPAP monoester), yielded the O-glucuronide of EtFOSE, EtFOSAA, FOSAA,

FOSE, FOSA, and PFOS (257), similar to the metabolite profiles previously observed for EtFOSE

biotransformation in rat liver tissues (252). As the mass spectrometric conditions were not

optimized for EtFOSE, this compound was not detected in these experiments, but the initial

breakdown of SAmPAP can conceivably occur by enzyme-mediated hydrolysis of the phosphate

ester linkage to yield EtFOSE, analogous to that previously proposed for the PAPs (71), from which

point EtFOSE biotransformation would proceed as described above.

1.4.2 Other Environmental and Biological Processing of Perfluoroalkyl and Polyfluoroalkyl

Substances

1.4.2.1 Environmental Processes: Sorption and Uptake into Vegetation

PFASs have a demonstrated ability to sorb to anthropogenic inorganic materials, such as

activated carbon and zeolite (258, 259), and naturally occurring environmental solids, such as clay

minerals (260–262), sediments (263–267), soils (268, 269), and sludge (263, 270). This sorption

capacity has implications for the retention and potential release of these chemicals from these

environmental solids to their surrounding compartments, such as the aqueous and plant

environment. Groundwater contamination has already been discussed in Section 1.3. Monitoring

data for sediments, soil, and WWTP sludge will be reviewed in Chapter 2.

PFAS concentration data in plants are sparse, with only three studies published to date (128,

271, 272). However, plant uptake of PFCAs and PFSAs has been previously demonstrated in

spring wheat, oats, potatoes, maize, and ryegrass sown in PFOA- and PFOS-spiked soil (273), in

carrots, potatoes, and cucumbers sown in soil amended with WWTP biosolids in the laboratory

(274), and grass collected from farm fields that had been consistently treated with WWTP biosolids

(128). Laboratory- and field-based sorption and plant-uptake data will be reviewed in Chapter 2.

1.4.2.2 Biological Processes in Aquatic Organisms

PFASs have been measured in fish, birds, and mammals worldwide (275–278). Houde et al.

reviewed the global contamination of PFASs observed in wildlife and humans in 2006 (174), then

38

followed up in a second review with more updated data that specifically focused on aquatic biota in

2011 (279). Butt et al. has also recently reviewed concentration data measured in Arctic wildlife

(209). Due to the breadth of wildlife data in the literature, this section will primarily focus on

biological processes, such as bioaccumulation and pharmacokinetics, in aquatic organisms.

1.4.2.2.1 PFAS Contamination in Aquatic Wildlife

PFOS is the predominant PFAS observed in freshwater and marine species. PFOS and

PFOA contamination is relatively well-documented in the Arctic, North America, Europe, and Asia,

as compared to the southern hemisphere (Fig. 1.8). However, within the last several years, wildlife

monitoring has emerged in locations that were either never studied before or at least not

extensively. For example, PFOS and PFOA have been observed in Antarctica seal pups (9.4 ng/g

wet weight, ww, PFOS; <0.4 ng/g ww, PFOA) (280), as well as, in mussels, fish, fur seals and

dolphins from South Brazil (<0.5–91 ng/g ww, PFOS; <0.2–15 ng/g ww, PFOA) (281, 282).

Concentrations observed in South American aquatic biota are typically lower than those measured

in the northern hemisphere (281, 283). This suggests the occurrence of more intensive production

and use of commercial fluorinated products in the northern hemisphere, although evidence of

continued manufacture and widespread use of Sulfluramid in South America was recently

highlighted by the significant PFAA and FOSA contamination observed in coastal waters near Rio

de la Plata (114). Proximity to local emissions has also been linked to elevated concentrations in

aquatic biota, as was observed in fish sampled in Tokyo Bay that receives industrial and municipal

wastewaters and in Kin Bay near a military base that may have employed AFFF during fire-training

exercises (277).

PFAS contamination has been observed in polar bears, ringed seals, Arctic fox, various

birds and fish, and dolphins in remote environments, such as the Arctic (278, 284–286), high-

altitude lakes (287, 288), and in the ocean (289, 290). Overall, PFOS is consistently observed as

the predominant species in these studies, although long chain PFCAs (≥8 CFs) are also present.

Contrary to the predominance of PFOA in human blood data, the PFCA congener profile observed

in wildlife typically exhibits a characteristic odd>even chain length pattern in which the

concentration of the odd-chain PFCA exceeds that of the adjacent shorter even-chain PFCA (i.e.

[PFNA]>[PFOA]; [PFUnA]>[PFDA]) (278). Analysis of different Arctic trophic levels revealed

polar bears as the most contaminated species (8 ng/g, PFOA; 180 ng/g, PFNA; 56 ng/g, PFDA; 63

39

Figure 1.8 Global contamination of PFOS and PFOA in fish from selected data summarized by Houde et al. (174, 279).

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40

ng/g, PFUnA), likely due to their position as the apex predator (278). Lower trophic organisms,

such as ringed seals (<2 ng/g, PFOA; 4.9–5.9 ng/g, PFNA; 2.1–2.9 ng/g, PFDA; 2.0–3.3 ng/g,

PFUnA) and fish (<2 ng/g, PFOA; <0.5–6.2 ng/g, PFNA; 0.5–2.5 ng/g, PFDA; 1.1–5.7 ng/g,

PFUnA) also exhibit the odd>even pattern, but at much lower concentrations (278). Atmospheric

degradation of fluorotelomer-based precursors has been shown to produce equivalent yields of

adjacent PFCA pairs (i.e. 8:2 FTOH PFOA/PFNA; 10:2 FTOH PFDA/PFUnA) (Section

1.4.1.1) (202). Increased bioaccumulation of the longer odd-chain PFCA congener would result in

the observed odd>even pattern.

The disparity in the PFCA congener profiles observed between humans and wildlife is

reflective of different exposure sources. The predominance of PFOA in humans is consistent with

the use of commercial products in which the fluorochemical composition has been primarily

perfluorooctyl-based until recently (3, 29). The increased production of PFOA, as compared to

PFNA and other PFCAs, from the biotransformation of 8:2 fluorotelomer-based commercial

materials (Section 1.4.1.2) is another contributing factor. In contrast, animals that are far removed

from anthropogenic sources may become exposed via environmental transport to the relevant

compartments. Both Ahrens et al. (287) and Shi et al. (288) observed PFOS (0.2–9.0 ng/g ww) and

PFCAs (<0.1–30 ng/g ww) in fish sampled in alpine lakes and rivers in the French Alps and the

Qinghai-Tibetan Plateau respectively, which suggests atmospheric deposition of volatile fluorinated

species to these isolated water bodies as a potential source of the contamination observed. Skipjack

tuna sampled in the open Pacific ocean exhibited PFAA concentrations of <1.0–59 ng/g ww, where

PFOS and PFUnA were observed as the dominant congeners (290). In addition to PFCAs and

PFSAs, Houde et al. also reported the presence of 8:2 and 10:2 FTUCAs, both of which are

established fluorotelomer-based intermediates (Section 1.4.1.2), in bottlenose dolphins along the

Gulf of Mexico and in the Atlantic Ocean (289). Interestingly, FTUCA concentrations observed in

the open ocean dolphins (0.5–1.4 ng/g ww) were higher than those measured in coastal dolphins

(nd–<0.4 ng/g ww) (289). The occurrence of PFASs in oceanic biota may also derive from

atmospheric input, but likely in tandem with the continuous circulation of legacy contamination in

water masses from the shore to the open ocean. The odd>even pattern is present in the PFCA

congener profiles in all of these studies.

In near-source regions however, PFCA contamination in aquatic organisms may be affected

by local input sources. Instead of the distinct odd>even pattern observed in remote aquatic biota,

41

both Martin et al. (291) and Furdui et al. (292) observed similar concentrations among the PFCAs

monitored in various aquatic invertebrates and lake trout sampled from the Great Lakes, although

concentrations tend to decrease with increasing chain length.

Perfluoroalkanesulfonamido- and fluorotelomer-based degradation intermediates have been

detected in aquatic biota. FOSA is frequently detected, sometimes even at similar concentrations as

PFOS (185, 278, 291). FOSAA, MeFOSAA, and EtFOSAA were detected at low concentrations in

the intestines, stomach, and gills of Chinese sturgeons sampled from the Yangtze River, while 7:3

FTCA was also detected at concentrations of 0.13–1.4 ng/g ww in livers (293). In contrast, none of

the other FTCAs and FTUCAs monitored were detected, although Furdui et al. have observed very

low levels of 8:2 and 10:2 FTUCAs (<0.001–0.18 ng/g ww) in lake trout from the Great Lakes

(292). Similarly, Powley et al. also did not detect any 8:2 FTCA and 8:2 FTUCA in various Arctic

biota, although 7:3 FTCA was present at 0.5–2.5 ng/g ww in seal liver (294). As described in

Section 1.4.1.2, 7:3 FTCA is frequently observed as an intermediate metabolite of an 8:2

fluorotelomer-based precursor, and has been shown to biotransform to 7:3 FTUCA and/or PFHpA

at very low yields (<1%) (162, 240). Comparison of the elimination kinetics observed in rainbow

trout dosed separately with 8:2 FTCA, 8:2 FTUCA, and 7:3 FTCA showed that 7:3 FTCA was

eliminated much slower (t1/2: 5.1 days, blood; 10.3 days, liver), as compared to the other two

metabolites (t1/2: 0.4–1.2 days, blood; 1.3 days, liver* (*only 8:2 FTCA as 8:2 FTUCA was not

observed above the limits of detection in the liver)) (240). As such, 7:3 FTCA may be an

appropriate biomarker to characterize exposure to fluorotelomer-based materials, although more

data is necessary to evaluate the extent of its contamination in wildlife.

A recent survey of lake trout homogenates sampled in 2008–2010 revealed low

concentrations of 6:2 (98 pg/g ww) and 8:2 diPAPs (nd–310 pg/g ww), as well as, the C6/C6 (nd–9

pg/g ww) and C6/C8 (nd–12 pg/g ww) PFPiAs (295). This data represents the first set of wildlife

measurements for the diPAPs and PFPiAs. The low diPAP concentrations are not surprising

considering these chemicals are metabolically active (71, 72), but PFPiAs are perfluorinated and as

such, are expected to persist in the environment. Despite their detection in surface water (105), the

absence of the mono-alkylated PFPAs in the lake trout (295) suggests they may be less

bioaccumulative than the di-alkylated PFPiAs. This will be further explored in Chapter 6.

42

1.4.2.2.2 Bioaccumulation in Aquatic Organisms

In addition to persistence (P) and toxicity (T), the potential for a chemical to bioaccumulate

(B) in an organism is another important criterion for regulatory agencies to consider when

evaluating the environmental risks of emerging chemicals. In this thesis, the various endpoint

metrics used to characterize bioaccumulation are defined as described by Gobas et al.(296).

Bioaccumulation (BAF) is a field-based metric that is expressed as the ratio of the steady-

state concentration of a chemical in a water-respiring organism to that in the water (BAF =

Corganism/Cwater (L/kg)) and accounts for all possible routes of exposure within this environment (i.e.

respiratory uptake via the gills, dermal uptake, dietary uptake). Bioconcentration (BCF), on the

other hand, is a laboratory-based metric that is also expressed as a ratio of the concentration

measured in the organism to that in the water (BCF = Corganism/Cwater (L/kg)), but only considers

water-borne exposure to the test animal. Biomagnification is a quantitative measure of

predator/prey relationships and is expressed as the steady-state concentration of the chemical in the

organism to that in its food source (BMF = Corganism (predator)/Cfood (prey)). BMF can be measured in the

laboratory in which test animals are exposed to the chemical via only dietary uptake, whereas, field-

based BMFs account for all other exposure routes (i.e. air, water, diet). Whereas BMF only

considers a single predator/prey relationship, trophic magnification (TMF) is essentially an average

BMF that quantifies the change in chemical contamination in organisms feeding at different trophic

levels within a food web.

Contaminant uptake in aquatic species may be influenced by a number of competing

processes occurring at the same time, such as those described by Arnot and Gobas in Fig. 1.9 (297).

The interplay between these uptake and elimination processes is key to driving chemical

accumulation in the aquatic organism. As PFASs are surfactants by nature, they possess both

hydrophilic and hydrophobic properties and thus, their bioaccumulative behaviour cannot be

modeled using traditional logKow-based partitioning models. A number of studies have

demonstrated preferential partitioning of PFASs into proteinaceous compartments, such as blood,

liver, and kidneys (293, 298, 299), as opposed to fatty-rich tissues, which is consistent with the

ability of these chemicals to bind to serum proteins (300, 301).

43

Fig. 1.9 Uptake and elimination processes of contaminants in fish

Bioaccumulation metrics (i.e. BCF, BAF, and BMF) have been determined both in the

laboratory and from field data for different aquatic species (Table 1.4). Martin et al. reported the

first comprehensive sets of BCF and BMF data for a suite of PFSAs and PFCAs of varying

perfluoroalkyl chain lengths in rainbow trout exposed via the water (298) and diet (302). PFSAs

and PFCAs containing less than 6 and 7 CFs respectively were not detected in most tissues and

therefore, not considered to be significantly bioaccumulative. On the other hand, BCF and BMF

values for the longer chain congeners were observed to increase with increasing chain length, but

this relationship deviated from linearity for perfluorotetradecanoate (PFTeA, C14), the longest

PFAA studied, in the water-borne exposure experiments, perhaps due to decreased gill permeability

(298). In addition, PFSAs was consistently observed to be more bioaccumulative than PFCAs of

equal perfluoroalkyl chain length, a phenomenon also observed in other laboratory and field data

(Table 1.4). However, none of the calculated BMF values were statistically greater than 1 and as

such, Martin et al. concluded PFAAs do not biomagnify in juvenile trout from dietary exposure

(302). This contrasts the detection of PFOS and PFCAs, often at higher concentrations, in higher

trophic level animals (276–278, 285, 291, 294), as will be discussed below.

Dietary

Uptake

Respiratory

Uptake

Gill

Elimination

Metabolism

Growth

Dilution

Fecal

Elimination

44

Table 1.4 Laboratory- and field-based metrics to evaluate bioconcentration, bioaccumulation, and biomagnification of PFAAs.

Organism Location PFHxS PFOS PFOA PFNA PFDA PFUnA PFDoA PFTrA PFTeA Reference

Laboratory-Based Data

BCF = Corganism/Cwater (L/kg)

Fathead minnow Liver (Female) - 830 - - - - - - -

(303) Liver (Male) - 210 - - - - - - -

Mussels Whole-body (1 ppb) - 378 15 144 838 - - - -

(304) Whole-body (10 ppb) - 235 12 109 464 - - - -

Rainbow trout

Whole-body 9.6 1100 4 - 450 2700 18000 - 23000

(298) Blood 76 4300 27 - 2700 11000 40000 - 30000

Liver 100 5400 8 - 1100 4900 18000 - 30000

BMF = Corganism/Cfood

Rainbow trout Whole-body 0.14 0.32 0.038 - 0.23 0.28 0.43 - 1.0 (302)

Field-Based Data

BAF = Corganism/Cwater (L/kg)

Benthic invertebrate Whole-body - 1000 - - - - - - - (305)

Bluegill Liver - 41600 - - - - - - - (277)

All fish Liver - 8540 - - - - - - -

Common shiner* Liver - 6300-125000 - - - - - - - (115)

All fish* Whole-body 71 1995 7.6 112 2344 2951 - - -

(116) Liver 74 12589 25 427 5495 3388 - - -

BMF = Cpredator/Cprey

Lake trout/Alewife Whole-body - 3.7 0.6 5.3 4.4 6.4 1.9 3.1 >2.6

(291) Lake trout/Smelt Whole-body - 1.6 0.5 0.6 1.0 1.2 1.0 1.2 2.2

Lake trout/Sculpin Whole-body - 0.4 0.02 0.1 0.2 0.2 0.3 0.4 0.3

Smallmouth bass, Round

gobies/Algae, Cray fish Muscle, Whole-body - 2-4 - - - - - - -

(305)

Chinook salmon/Round gobies Liver, Whole-body - 10-20 - - - - - - -

Lower trophic fish/Zooplankton Whole-body 9.1-10 12-35 - - - - 2.5-156 - -

(306)

Seatrout/Lower trophic fish Whole-body - 1.5-2.8 - - - - 0.2-14 - -

Dolphin/All fish Whole-body 1.8-2 6.2-18 - - - - 0.1-2 - -

Seatrout/Pinfish Whole-body - 4.6 7.2 1.5 3.7 0.9 0.1 - -

Dolphin/All fish Whole-body 3.3-14 0.8-4 1.8-13 1.4-24 2.4-8.8 1.9-3.9 0.1-1.8 - -

Arctic cod/Zooplankton - - 8.7 - - 0.5 - 0.3 - - (294)

Seal/Arctic cod Blood - 7.0 - - 1.4 3.1 0.8 - -

*Fish were exposed to high levels of PFOS following an accidental spill of AFFF into the river from which the fish were sampled.

45

Interestingly, Martin et al. calculated higher BCF values from the data in the blood (76–

30000), and liver (100–30000) samples, as compared to the whole-body homogenates (excluding

the liver) (9.6–23000) (298). This is consistent with the tendency of PFASs to preferentially

accumulate in blood and liver, which often results in an overestimation of their bioaccumulative

potential in the organism if blood and liver concentrations were used to calculate BCF, BAF, and/or

BMF. This issue becomes especially important when comparing measured BCF and BAF with the

ranges deemed by regulatory agencies as sufficient for the chemical in question to be considered

bioaccumulative. For example, Martin et al. reported liver- and whole-body-based BCFs of 5400

and 1100 respectively for PFOS, where the former value would render PFOS as a bioaccumulative

substance according to Environment Canada (≥5000) and European Union (≥2000) (296), while the

latter would not. It is widely agreed that bioaccumulation metrics based on the analysis of whole-

body tissues are considered the most appropriate and bias-free (174, 279, 296), although this maybe

analytically difficult for larger animals, such as marine mammals. Instead, Houde et al.

recommended the use of whole-body burden estimates to measure bioaccumulation for larger

predators by extrapolating their tissue concentrations to the whole body based on the mass

distribution of the individual tissues analyzed (174). This was performed for quantifying BMF in a

bottlenose dolphin food web (306) and a Canadian Arctic marine food web (307).

Field-based BAFs and BMFs are consistently higher than the corresponding BCFs and

BMFs measured in the laboratory (Table 1.4). Whereas Martin et al. did not observe

biomagnification of any PFAAs studied in the laboratory (302), BMFs measured from field data are

often greater than 1, which implies the presence of biological and environmental variables that may

not be accounted for by laboratory experiments. For example, local inputs of fluorinated chemicals

may result in elevated concentrations of PFASs in wildlife and consequently, yield higher than

expected BAF or BMF values, as was observed for PFOS (6300–125000, BAF) in fish sampled in a

river following an accidental spill of AFFF (115, 116). Bioaccumulation, followed by

biotransformation of fluorinated precursor chemicals, such as those described in Section 1.4.1.2,

may also influence bioaccumulation. Butt et al. (164) and Brandsma et al. (239) both observed the

biotransformation of FTAC, FTOHs, and PFOSA in exposed rainbow trout to degradation products

that were more bioaccumulative than the precursors themselves.

Overall, PFOS is generally observed to biomagnify to top predators in both freshwater and

marine food webs, while the occurrence of biomagnification appears to be more variable for

46

PFCAs, with BMF typically increasing with increasing chain length. However, field-based BMF

can sometimes be suppressed when lower trophic organisms are more contaminated, as was

observed in the benthic invertebrate, Diporeia, and its natural predator, sculpin, in a freshwater food

web study in Lake Ontario (291). Higher PFOS (280–450 ng/g ww) and PFOA (44–90 ng/g ww)

concentrations were observed in Diporeia and sculpin, as compared to lake trout (170 ng/g ww,

PFOS; 1 ng/g ww, PFOA), the top predator of the food web. This contamination was speculated to

have derived from benthic uptake of contaminated sediments (291). However, when the BMF

values were normalized to the actual distribution of each prey item in the diet of the lake trout (90%

alewife, 7% smelt, 2% sculpin), the resulting diet-weighted trout/prey BMFs for PFOS and all

PFCAs all exceeded 1. This suggests the small proportion of sculpin that lake trout naturally

consumes would not significantly affect biomagnification and that trophic magnification of PFOS

(5.9, TMF) and PFCAs (2.5–4.7, TMF’s for PFDA, PFUnA, and PFTrA) was indeed occurring at

the top of the freshwater food web (291). This example demonstrates TMF may be a better

parameter to characterize biomagnification as it accounts for multiple predator/prey interactions

across the entire food web and is not subjected to as many variables as BMF.

Biomagnification may also differ significantly between aquatic and terrestrial food webs due

to differences in feeding ecology between poikilotherms and homeotherms, as well as, the various

processes, as described in Fig. 1.9, controlling chemical uptake in the organism. For example,

homeotherms have higher feeding rates than poikilotherms, with the result that birds and mammals

are typically more contaminated than aquatic invertebrates and fish. Respiration is an additional

mode of elimination of PFAAs for water-respiring organisms, whereas, this pathway would be

much less operative in air-respiring organisms due to the involatility of the anionic PFAAs.

However, terrestrial animals may be more exposed, especially in near-source regions, to nonvolatile

precursors via dermal and/or dietary uptake and volatile precursors via inhalation. In fact, wood

mice living near a fluorochemical plant exhibited some of the highest concentrations of PFOS

(470–180000 ng/g ww) to be ever reported for any organism (308). Caribou and wolves from

Northern Canada also exhibited low levels of PFOS and PFCAs (ΣPFAS: 0.25–2.4 ng/g ww,

muscle; 6.5–20 ng/g ww, liver) (272). In that same study, PFOS and all PFCAs (≥8 CFs) were

observed to biomagnify in the lichen-caribou-wolf food chain in which the TMFs (2.2–2.9) were

observed to be similar to those measured in a dolphin food web study (306), but less than those

47

measured in a Canadian Arctic marine food web (307). Nevertheless, these studies confirm that

PFAS may biomagnify in both aquatic and terrestrial ecosystems.

1.4.2.2.3 Pharmacokinetics and Distribution in Aquatic Organisms

The pharmacokinetic behaviour of PFSAs and PFCAs has been extensively reviewed by

Lau et al. (309), Andersen et al. (310), and most recently, Han et al. (311), although these reviews

primarily focused on mammalian studies. In general, PFAAs tend to predominate in protein-rich

tissues, such as blood, liver, and kidneys, as was observed in fish (293, 298, 299, 303) and harbour

seals (312), presumably due to their demonstrated affinity to serum proteins (300, 301). Once

absorbed, the recalcitrant PFAAs may persist in the organism via enterohepatic circulation (i.e.

biliary transport between the liver and the gastrointestinal tract).

Elimination in aquatic organisms may occur via the various processes described in Fig. 1.9.

Detection of PFASs in the gills of exposed rainbow trout (298) and Chinese sturgeons (293)

suggests this organ as a potential site of uptake and/or depuration via respiration. The occurrence of

PFASs in fish eggs (293, 299, 305), often at concentrations within the same order of magnitude as

the most contaminated liver tissue, also suggests oviparous transfer may decrease the body burden

in adult female fish. In addition, body growth during maturation may result in dilution of the

overall chemical burden in the organism.

Fecal egestion has been demonstrated in rainbow trout exposed to 8:2 FTAc via the diet in

which a number of intermediate metabolites (i.e. 8:2 FTCA, 8:2 FTUCA, 7:3 FTCA, and 8:2 FTOH

glucuronide) and the terminal PFOA and PFHpA were observed at concentrations ranging from

15to 3400 ng/g ww in the feces (164). Biliary excretion during enterohepatic circulation may act

as a source of the contamination observed in the feces and this was supported by the similar

concentrations (within 2–4-fold) observed in the bile (164), as well as, significant gallbladder

accumulation of PFCAs and PFSAs observed in fish from other studies (293, 298). As urine is

inherently difficult to sample from fish, urinary excretion of PFASs in fish is not well understood.

High kidney concentrations have previously been reported in rainbow trout exposed to 8:2 FTAC

(164) and PFAAs (298) via dietary and water-borne exposure respectively, although this may have

been an artifact of the perfusion of highly contaminated blood through this tissue. Nevertheless,

urinary excretion has been well established in other laboratory animals and has been implicated in

gender-associated differences in the renal elimination of PFCAs (311).

48

Gender differences have also been observed in the pharmacokinetics of PFOS and PFOA

separately in fathead minnows (303, 313). Faster depuration of PFOA was observed in female

minnows (6 h, t1/2), as compared to male minnows (69 h, t1/2) (313). Administration of the

androgen trenbolone to the female minnows resulted in slower elimination kinetics (25 h, t1/2),

although faster kinetics were not observed in males upon treatment with the estrogen

ethynylestradiol (313). This suggests PFOA elimination in fathead minnows may be partially

hormonally regulated, possibly due to differential gene expression of organic anion transport (OAT)

proteins between the two genders, as have been observed in rats (314–317). There is considerable

evidence that OAT proteins are involved in the active uptake and renal processing of PFCAs in rats

in which increased expression of resorptive OAT proteins in male rats would result in increased

retention of PFOA and PFNA, as compared to females, although these gender differences become

less pronounced or reversed for shorter and longer chain PFCAs (<7 CFs and >8 CFs) (314–318). In

contrast, oral exposure of PFOS to fathead minnows resulted in 2–3-fold higher concentrations

inthe blood, liver, and gonads of female minnows, as compared to the males, the reasons for these

gender differences were not apparent (303). It is unclear whether these gender differences would be

conserved in other aquatic species, as not all mammalian species studied have exhibited the same

patterns observed in rats.

Lastly, metabolism, as was observed in rainbow trout exposed to various fluorinated

precursors (164, 239), is another mechanism by which PFASs may be eliminated from an aquatic

organism. Overall, the elimination pathways discussed above are counteracted by dietary and

respiratory uptake, with the balance among these different processes ultimately controlling the level

of contamination in the organism.

1.5 Goals and Hypotheses

PFASs are dispersed in the environment via anthropogenic activities and the use of

commercial fluorinated products. Despite the limited direct applications of PFSAs and PFCAs, as

discussed in Section 1.2.2, these chemicals are often observed as the major species in the

environment. In contrast, commercial fluorinated surfactants comprise a significant component of

current fluorochemical production (~20% of the fluorotelomer industry (3, 29)) and are often

incorporated at percent quantities in the final sales products (2, 18, 39, 73, 76–79). Both

atmospheric and biological transformation of various commercial perfluoroalkanesulfonamido- and

49

fluorotelomer-based surfactants has been demonstrated to yield PFSAs and PFCAs of varying

perfluoroalkyl chain lengths (Section 1.4.1), but the processes involved in their distribution to the

relevant environmental compartments remain poorly understood. The motivation behind this thesis

is to examine the processes that are responsible for driving this distribution and ultimately,

understand how exposure to commercial materials may contribute to the current fluorochemical

contamination observed in humans, wildlife, and other abiotic media.

The distribution of PFAS via anthropogenic discharges to WWTPs is of particular interest to

this work. Once inside a WWTP, PFAS may disperse via two pathways: (1) relocation to landfills

and/or agricultural farmlands via disposal of biosolids generated at the WWTP, or (2) continued

transport to receiving waters located downstream from the facility. The relative importance of these

two pathways is reviewed in Chapter 2. Both of these pathways are examined as potential

contributors of commercial fluorinated materials, specifically phosphorus-based fluorinated

surfactants, such as the PAPs, PFPAs, and PFPiAs, as contaminants themselves, as well as sources

of PFAAs observed in these environments.

Chapter 3 investigates the potential for PAPs to contribute to the burden of PFCAs observed

in wastewater environments, as these chemicals, specifically the diPAPs, have been previously

detected at hundreds of ng/g concentrations in WWTP sludge (169). This hypothesis was tested by

performing biodegradation experiments of diPAPs and monoPAPs in the presence of WWTP

microbes. Their resulting degradation metabolites and the pathways to produce them were

elucidated. Chapter 4 builds upon these results and examines the potential of these fluorinated

surfactants to further transfer to the soil environment upon amendment of contaminated waste

materials, such as biosolids and paper fiber wastes, during agricultural land application. This

hypothesis was tested by a series of greenhouse biosolids-amended soil-plant microcosms in which

soil and plant biotransformation, as well as uptake of both the parent diPAPs and their degradation

products were investigated.

The next two chapters focus on the environmental chemistry of PFPAs and PFPiAs,

specifically in the aqueous environment and environmental solids, as PFPAs have been detected in

surface water and WWTP effluents (105), while PFPiAs have been found in WWTP sludge (319)

and lake trout (295). Chapter 5 examines the sorption and desorption behaviour of PFPAs and

PFPiAs in a diverse set of soils of varying geochemical properties. Structural features, such as the

perfluoroalkyl chain length and headgroup, were investigated to determine which congeners would

50

be more prone to remobilization into the aqueous environment and which would preferentially

remain bound to the solid phase. Regardless of which compartment the PFPAs and PFPiAs would

prefer to reside in, aquatic organisms can become exposed to these chemicals via respiratory and

dietary uptake. Biological processes, specifically biomagnification and depuration, of these

chemicals were investigated using juvenile rainbow trout as the test organism in Chapter 6.

Analysis of the total extractable organofluorine fraction in human blood revealed that

known fluorinated compounds, such as PFCAs and PFSAs, constituted only a small portion of the

actual fluorochemical contamination observed (320). This suggests the presence of other

unidentified fluorinated species. Efforts to provide a comprehensive evaluation of human blood

fluorochemical contamination were performed in Chapter 7 in which fifty North American human

sera samples were analyzed for forty different fluorinated analytes that included commercial

fluorinated surfactants, residual materials, degradation intermediates, and the terminal PFCA and

PFSA metabolites. In addition to the diPAPs which have been previously detected in human blood

(169), this study also surveyed for various fluorinated surfactants that were either not extensively or

never monitored before, including the FTSAs, SAmPAP, FTMAP, PFPAs, and PFPiAs.

The final chapter summarizes the overall contribution of the two aforementioned pathways

to the contamination observed in the various compartments studied in this work. Future research

directions to investigate how fluorinated surfactants are circulated in other environments are also

discussed.

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CHAPTER TWO

Global Distribution of Polyfluoroalkyl and Perfluoroalkyl Substances and their

Transformation Products in Environmental Solids


Submitted: As a book chapter for the invited review to “Transformation Products of

Emerging Contaminants in the Environment: Analysis, Processes, Occurrence, Effects and

Risks” by John Wiley & Sons, Ltd.

Contributions: Holly Lee prepared this manuscript with editorial comments provided by

Scott Mabury

This chapter has been condensed from the original manuscript, specifically in Section 2.4.3,

describing the biotransformation of PFASs, to reduce redundancy with Chapter 1.

Reproduced with permission from John Wiley & Sons Ltd.

Copyright John Wiley & Sons Ltd 2012

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2.1 Abstract

Perfluoroalkyl and polyfluoroalkyl substances (PFASs) have been ubiquitously detected

in environmental solids like sediments, wastewater treatment plant (WWTP) sludge, and soil,

with perfluorooctanoate (PFOA, C8) and perfluorooctanesulfonate (PFOS, C8) typically

observed as the dominant perfluoroalkyl acids (PFAAs). Urban and industrial discharges have

been identified as major contributors to current ambient levels of PFASs in near-source

environments, while a number of studies have also highlighted the contribution of known

fluorochemical point sources to regional contamination hotspots. In these near-source regions,

the high PFAS contamination observed in sediments and soils has been attributed to proximity of

airports and fire-training facilities using aqueous film-forming foams (AFFFs), discharges from

nearby fluorochemical production facilities, accidental spills, and application of contaminated

WWTP biosolids to agricultural farmlands. In the case of the biosolids-applied farmlands,

significant PFAS contamination was not only limited to soils, but was also observed in the plants

and groundwater collected in the vicinity. In addition, since the early 2000s, China has emerged

as a major fluorochemical producer, especially after the production phase-out of

perfluorooctylsulfonyl (POSF)-based materials in North America in 2000–2002. This shift in the

fluorochemical industry is reflected in global environmental surveys in which sediments, WWTP

sludge, and soil sampled in China and other Asian-Pacific countries often exhibit the highest

PFAS concentrations compared to those observed in Europe and North America.

2.2 Introduction

PFASsare anthropogenic chemicals that have a fluoroalkyl backbone and a polar

headgroup, both of which impart high surface activity and the ability to repel water, oil, and stain

to these chemicals (1). As such, PFASs are crucial components in non-stick, greaseproofing, and

surface treatment applications. Commercial fluorochemical production has largely proceeded by

two manufacturing processes, electrochemical fluorination (ECF) and telomerization(1), with the

bulk of the production centered on high molecular weight (MW) fluorinated polymers and

surfactants(2, 3) and a minor proportion directed towards the synthesis of specific

perfluoroalkanesulfonate (PFSA) and perfluoroalkyl carboxylate (PFCA) congeners. PFOS was

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the only PFSA deliberately produced to be used in AFFFs and various performance applications

(4, 5) until it was phased out of production in 2000–2002 (6). Among the PFCAs, PFOA and

perfluorononanoate (PFNA, C9) are primarily used as processing aids in the manufacture of

fluoropolymers(7), although other PFCAs of varying chain length have been detected as residual

impurities in commercial products (8).

Since the first discovery of PFOA and PFOS in human blood (9) and wildlife (10), these

PFAAs and other PFASs (Table 2.1) have emerged as common contaminants in surface water

(11), sediments (12), WWTP sludge (12), and soil (13).

Table 2.1.Names, acronyms, and structures of various PFASs of interest.

Name Acronym Structure

Fluorotelomer-based Substances

x:2 Fluorotelomer alcohol x:2 FTOH F(CF2)xCH2CH2OH, x = 4, 6, 8, 10,…

x:2 Fluorotelomer acrylate x:2 FTAC F(CF2)xCH2CH2OC(O)CH=CH2, x = 4, 6, 8, 10,…

x:2 Polyfluoroalkyl phosphate monoester x:2 monoPAP F(CF2)xCH2CH2OP(O)O2-, x = 4, 6, 8, 10,…

x:2 Polyfluoroalkyl phosphate diester x:2 diPAP [F(CF2)xCH2CH2O]2P(O)O-, x = 4, 6, 8, 10,…

x:2 Polyfluoroalkyl phosphate triester x:2 triPAP [F(CF2)xCH2CH2O]3P(O), x = 4, 6, 8, 10,…

x:2 Fluorotelomersulfonate x:2 FTSA F(CF2)xCH2CH2SO3-, x = 4, 6, 8, 10,…

Semifluorinatedx-alkane SFA or FxHy F(CF2)x(CH2)yH, x = 3–20, y = 3–20

Semifluorinated alkene SFAene or

FxHyene F(CF2)xCH=CH(CH2)y, x = 3–20, y = 3–20

x:2 Fluorotelomer carboxylate x:2 FTCA F(CF2)xCH2CO2-, x = 4, 6, 8, 10,…

x:2 Fluorotelomer unsaturated carboxylate x:2 FTUCA F(CF2)x-1CF=CHCO2-, x = 4, 6, 8, 10,…

PerfluoroalkaneSulfonamido-based Substances

Perfluorooctane sulfonamide FOSA F(CF2)8SO2NH2

N-Methyl perfluorooctane sulfonamide MeFOSA F(CF2)8SO2NH(CH3)

N-Ethyl perfluorooctane sulfonamide EtFOSA F(CF2)8SO2NH(CH2CH2)

Perfluorooctanesulfonamidoacetate FOSAA F(CF2)8SO2NH(CH2C(O)O-)

N-Methyl perfluorooctanesulfonamidoacetate MeFOSAA F(CF2)8SO2N(CH3)(CH2C(O)O-)

N-Ethyl perfluorooctanesulfonamidoacetate EtFOSAA F(CF2)8SO2N(CH2CH3)(CH2C(O)O-)

N-Ethyl perfluorooctanesulfonamidoethyl

phosphate diester SAmPAP [F(CF2)8SO2N(CH2CH3)(CH2CH2O)]2P(O)O

-

Perfluoroalkyl Acids (PFAAs)

Perfluoroalkyl carboxylate PFCA F(CF2)xCO2-, x = 1–13

Perfluoroalkanesulfonate PFSA F(CF2)xSO3-, x = 4, 6, 8, 10

Perfluoroalkylphosphonate CxPFPA F(CF2)xP(O)O2-, x = 6, 8, 10

Perfluoroalkylphosphinate Cx/CyPFPiA F(CF2)xP(O)(O

-)((CF2)yF),

x = 6, 8; y = 6, 8, 10, 12; x + y ≤ 18

Detection of PFASs has been reported worldwide andeven in remote environments like

the Arctic and Antarctica. PFASs may be directly released into the environment via emissions of

contaminated discharges from fluorochemical manufacturers and the disposal of commercial

83

products in which these chemicals are either present as active ingredients or as residual

impurities. Environmental degradation of commercial fluorinated polymers and surfactants

present in disposed products to the PFSAs and PFCAs also represents an indirect source of these

PFAAs to the environment. In addition, commercial manufacture of fluorinated chemicals is

typically a crude process during which unreacted starting materials or byproducts may be

incorporated into the consumer products (4, 7). In fact, analysis of various commercial

fluorinated products revealed the presence of fluorotelomer alcohols (FTOHs) and N-methyl

perfluorooctanesulfonamidoethanol (MeFOSE) as residual impurities at up to 4%quantities (14).

As FTOHs and MeFOSE are volatile, the release of these and potentially other volatile materials

via offgassing from commercial products may represent a significant source to the atmospheric

fluorochemical burden.

Atmospheric transport and degradation of volatile fluorinated precursors to PFCAs (15,

16) and PFSAs (17, 18) and the subsequent deposition of these degradation products may in part

contribute to the background levels of PFAAs observed in the environment (Fig.2.1.). Results

from modeling the formation of PFOA from the atmospheric oxidation of 8:2 FTOH predicted

ubiquitous PFOA pollution in the Northern hemisphere atmosphere, with higher concentrations

occurring in remote regions (e.g. Arctic) than in source regions (19). This distribution is

consistent with thehigh nitrogen oxide (NOx) environment of urban locations in which NOxmay

interfere with the atmospheric formation of PFCAs and thus, reduce their yields. As such, point

sources may be more important contributors to local PFAS contamination in near-source regions.

Figure 2.1.Environmental pathways of PFASs.

WWTPSurface

Water

Farmland

Manufacturer

Volatile Precursors

Sediment

Point Sources

Biosolids Application

Ocean

Sediment

Remote

Environments

Atmospheric TransportAtmospheric Transport

Atmospheric Oxidation

and DepositionAtmospheric Oxidation

and Deposition

Oceanic Transport

84

Upon exiting a WWTP, domestic and industrial effluents may further contaminate

receiving water bodies, as have been documented by the detection of PFASs in WWTP samples,

surface water, and sediments collected downstream from these facilities (Fig. 2.1.). However,

these data do not account for the potential release of PFASs from the treated sludge or biosolids

co-generated at these WWTPs. The disposal of these solid waste materials may have

implications for human and wildlife exposure, especially if they are applied as a soil fertilizer

onto agricultural fields (Fig.2.1.). PFASs have a demonstrated capacity to sorb to environmental

solid matrices, such as clay minerals (20–22), sediments (23–27), soils (28, 29), and sludge (30).

The fact that PFASs may sorb to these environmental solids has implications for the long-term

retention and release of these chemicals to the aqueous environment. A number of studies have

attributed land application of contaminated WWTP sludge (31, 32), AFFF use at fire-training

facilities (33, 34), and leaching of urban wastewater and runoffs (35, 36) as potential sources of

groundwater and surface water PFAS contamination. The primary concern of this

contamination, particularly in the groundwater, centers over its potential as a route of human

exposure to PFASs through drinking water. Human and wildlife exposure may also occur

through ingestion of contaminated field crops, as evidenced by recent experimental and field data

that demonstrated the transfer of PFCAs and PFSAs from contaminated soils to assorted plants

(37–39).

This review summarizes the monitoring data collected from sediment, WWTP sludge,

and soil samples collected around the world in the context of these environmental pathways.

Source elucidation of PFASs will also be evaluated by identifying diffuse sources (i.e.

atmospheric transport, urban/industrial discharges) as contributors to ambient levels and

distinguishing them from known fluorochemical point sources. A discussion of various

processes known to control the distribution of PFASs in the environment is also presented.

2.3 Global Contamination of PFASs in Environmental Solid Matrices

2.3.1 Sediments

The detection of PFCAs and PFSAs of varying chain lengths has been widely reported in

freshwater, coastal, and marine sediments collected around the world (Table 2.2).

85

Table 2.2.Ambient concentrations of ΣPFCAs and ΣPFSAs (ng/g dry weight, dw) observed in

freshwater, coastal, and marine sediments collected around the world.

Mean Concentration*

(ng/g dry weight, dw)

Country Type of Sediment ΣPFCAs ΣPFSAs Range of PFASs

observed Reference

Freshwater

Austria River and lake 3.98 0.14 nd-1.25 (40)

Arctic (Canada)** Lake 4.81 1.38 nd-2.78 (41)

Canada Lake 1.77 1.09 0.10-0.87 (42)

France River 3.60 4.80 nd-4.30 (43)

Hong Kong Channel and wetland 10.07 11.07 nd-9.06 (44)

Japan River 1.97 2.93 <LOQ-1.69 (45)

Kenya*** River 12.53 3.15 - (46)

Mainland China River 0.29 0.31 <LOQ-0.21 (47)

Mainland China River 0.35 0.56 0.02-0.48 (48)




Mainland China Water reservoir 0.29 nd nd-0.18 (52)

Mainland China River and lake 0.36 0.24 nd-0.15 (53)

Mainland China Lake 0.26 0.24 <LOD-0.24 (54)

Netherlands** River - 1.06 - (55)

Spain Canal 4.04 0.73 0.02-3.19 (56)

Taiwan

River upstream and

downstream of

WWTP

15.13 124.55 <LOQ-159.4 (57)

USA River downstream of

WWTP 0.56 2.02 0.06-1.24 (58)

Coastal

Australia Harbour river 0.96 2.20 nd-2.10 (59)

Japan Tidal flat 0.96 0.54 <LOD-0.96 (60)

Japan Bay 0.22 0.54 - (61)

Mainland China Bay and tributaries 0.40 0.20 <LOD-0.20 (62)

Mainland China*** River estuary - 236.20 - (63)

Spain WWTP and urban

emissaries 0.02 0.02 <LOD-0.02 (64)

USA Bay 0.65 1.56 <LOQ-1.05 (12)

Marine

Baltic and North

Seas Ocean 0.57 0.53 <LOQ-0.51 (65)

Baltic Sea Ocean 0.11 0.54 nd-0.38 (66)

*Concentrations reported in sediments collected from different sampling locations within the same study were

summarized here as an overall arithmetic mean. **Data from Char and Amituk Lakes only, data from Resolute

Lake discussed separately in the text. ***Only PFOA and/or PFOS was monitored.

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The data presented are considered to be background levels in this review and

representative of the diffuse sources in the environment. Total PFCA (ΣPFCA) and PFSA

(ΣPFSA) concentrations range from low (<1) to mid (~100) ng/g dry weight concentrations.

A suite of C2 (trifluoroacetate, TFA) to C14 (perfluorotetradecanoate, PFTeA) PFCAs

have been detected in sediments, with PFOA typically observed as the dominant congener,

followed by the longer chain PFCAs (≥8 perfluorinated carbons, CFs) in both concentration

levels and detection frequency. One exception to this distribution is the observation of TFA as

the dominant congener (22-90% of ΣPFASs) in sediments (45-127 ng/g dry weight), as well as in

sludge and soil collected from Shanghai, China (49). The fact that TFA and other short chain

PFCAs (<7 CFs) are usually not monitored in environmental samples is problematic given the

potential phytotoxicity of these compounds and that TFA has been previously observed at higher

concentrations than longer chain PFCAs in surface waters (67).

Similarly, the PFSA congener profile in sediments is dominated by PFOS, with

occasional detection of perfluorobutanesulfonate (PFBS, C4), perfluorohexanesulfonate (PFHxS,

C6), and perfluorodecanesulfonate (PFDS, C10). The observations of PFOA and PFOS as the

major PFAAs in sediments are consistent with their stronger affinity to sediments as compared to

shorter-chain PFAAs, as was observed by Higgins and Luthy(23), and the historically dominant

C8 chemistry in fluorochemical production.

To date, only one study has attempted to monitor for the C6, C8, and C10

perfluoroalkylphosphonates (PFPAs) in river sediments collected from the Netherlands, although

none of the congeners were detected in these samples (55). The concentrations of other

perfluoroalkanesulfonamidoacetates (FASAAs) (i.e. perfluorooctanesulfonamidoacetate

(FOSAA), N-methyl perfluorooctanesulfonamidoacetate (MeFOSAA), and N-ethyl

perfluorooctanesulfonamidoacetate (EtFOSAA)) are usually within the same order of magnitude

as those reported for the PFAAs in sediments.Recently, Benskinet al. reported for the first time,

N-ethyl perfluorooctanesulfonamidoethyl phosphate diester (SAmPAP) at concentrations of 40–

200 pg/g dw in marine sediments from a harbour in Vancouver, Canada (68). The SAmPAPs

were historically used as greaseproofing agents in food packaging materials (4) until they were

phased out in 2002 with other POSF-based materials by a major fluorochemical manufacturer in

North America (6). The fact that these chemicals remain detectable to date, despite the cessation

of their use a decade ago, suggests some persistence in the environment.

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Higher PFAS concentrations were typically observed in freshwater sediments collected

from rivers, lakes, canals, and other surface water bodies near urban and industrial regions, while

coastal and marine sediments were less contaminated. This spatial distribution may reflect

increased dilution of the PFAS contamination as the chemicals migrate towards the ocean,

althoughproximity to local urban and industrial emission sources of PFASs may also affect

regional contamination. For example, Pan and You measured significant PFOS concentrations

(73-537 ng/g dry weight) in sediments collected from Baozhen Port, a freight terminal and

passenger wharf, located in the Yangtze River estuary in China (63). These measurements

represent some of the highest PFOS sediment concentrations to be ever reported in coastal

environments and are reflective of the intensive anthropogenic activities occurring in this area.

Although dilution from marine waters plays a key role in redistributing PFASs as they travel

from freshwater towards the ocean, PFAS sorption to sediments may also increase with increased

water salinity (25), as was recently demonstrated by the increase in PFOS concentrations

observed in sediments collected from sites of low to high salinity in the same Yangtze River

estuary (63). This suggests that coastal estuaries may be an important sink for PFASs during

their transport from local aquatic environments (i.e. rivers, lakes, streams) to the ocean at which

point PFAS concentrations would become greatly diluted.

Sediment contamination may also arise from proximity to known fluorochemical point

sources. Stock et al. reported one of the earliest cases of significant sediment contamination in

the Canadian Arctic in which ~100 ng/g dry weight of total PFAS (ΣPFAS) were measured in

sediments collected from Resolute Lake, which continuously receives wastewater outflow from

the adjacent Meretta Lake and is located downstream of an airport at which AFFF may have

been used (41). These concentrations were 1–2 orders of magnitude higher than those measured

in sediments sampled from Char and Amituk Lakes (Table 2.2), both of which are isolated from

local emissions,such that the contamination observed within was proposed to be predominantly

due to atmospheric transport and degradation of volatile fluorinated precursors, followed by

subsequent deposition of their degradation products(41). A more recent case of contamination

was reported in Fuxin, China in which environmental samples collected near a fluorochemical

industrial park, were discovered to contain significant PFOA concentrations (up to 48 ng/g dry

weight in sediments; 668 ng/L in river water) (69). Accidental spills constitute a single pulse of

fluorochemical emission which may persist in the environment for a long time. One of the most

88

notable spills was recently highlighted by the elevated concentrations of PFOS (13 ng/g dry

weight) observed in sediments collected in 2009 from Spring Creek Pond which received AFFF

from an accidental spill from the Toronto International Airport almost a decade earlier (70).

As shown in Table 2.2, PFAS sediment contamination appears to be the highest in the

Asia-Pacific region (i.e. Hong Kong, mainland China, Taiwan) in which ΣPFCA and ΣPFSA

concentrations have been reported to exceed 100 ng/g dry weight. PFAS contamination in North

America, on the other hand, is much lower (<10 ng/g dry weight) and is generally similar to that

reported in other countries. This disparity may be related to the recent combined efforts of the

government and various fluorochemical manufacturers in North America to cease production of

certain fluorochemical product lines (3M phase-out of perfluorooctylsulfonyl (POSF)-based

materials in 2000) (6) and reduce and ultimately eliminate emissions from current manufacturing

processes and products (U.S. EPA 2010/2015 Global PFOA Stewardship Program) (71). Since

the early 2000s, there has been a resurgence in large-scale production of POSF-based materials

in China due to increasing local and overseas demands (72), which may be partially responsible

for the significant environmental contamination observed in Asia. However, the global

variability in the observed contamination of PFASs may also arise from differences in the

sampling techniques employed by these different studies, such as the chosen depth at which the

sediments were collected. As will be discussed next, PFAS concentrations can vary significantly

along a sediment core in which sectional analysis at different depths can produce a

contamination profile with respect to time.

2.3.1.1 Temporal Trends in Sediment Cores

The temporal trends of PFASs observed in sediment cores (41, 73–75) generally

correspond well with the major changes that occurred in fluorochemical production over the past

several decades. Analysis of sediment core slices collected from Arctic lakes showed higher

PFAS concentrations in the surface slices (0–1 cm, 1976–2003), as compared to those measured

at lower depths (1–2 cm, 1942–1996; 2–3.5 cm, 1908–1989), which is consistent with known

commercial fluorochemical production trends (41). In two core samples from Tokyo Bay,

Ahrens et al. observed an increase in ΣPFAS flux from 7 pg/cm2/year in 1956–1958 to 197

pg/cm2/year in 2001–2002, followed by a subsequent decline to 88 pg/cm

2/year beyond 2002

89

(73). PFOS concentrations doubled within 16 years between 1956 and 2008, but the increase

was then observed to slow down between 2001 and 2008 (73). Similarly, perfluorooctane

sulfonamide (FOSA) and EtFOSAA exhibited doubling times of 6 and 5 years respectively in

their sediment concentrations during the period of 1985–2001, followed by a rapid decline after

2001 (t1/2: 14 years, FOSA; 3 years, EtFOSAA) (73). These trends correspond to the use of ECF

to produce POSF-based materials that began in the late 1940s (5), followed by increased

production from the 1970s onward (76), until the complete cessation of POSF-based production

in 2002 (6), from which point telomer-based production increased significantly to assume the

fluorochemical market share left vacated by the phase-out (3). Concentrations of PFNA and

perfluoroundecanoate (PFUnA, C11) both of which are PFCA metabolites from fluorotelomer

degradation, were also observed to increase with doubling times of 4 and 5 years respectively

between 1990 and 2008 (73). Zushiet al. reported similar temporal trends in a sediment core also

collected in Tokyo Bay in which PFOS and its precursors, MeFOSAA and EtFOSAA exhibited

declines in their concentrations from the 1990s to 2004, while the opposing trend was observed

for PFOA and other long-chain PFCAs (74).

More recently, Benskinet al. observed good agreement between FTOH emission trends

and PFCA fluxes in sediment cores collected from two remote alpine lakes in the Canadian

Rocky Mountains that were purported to be predominantly influenced by atmospheric transport

of volatile fluorotelomer precursors (75). Specifically, substantial increases in FTOH emissions

that occurred in the period of 1999–2005 (30 to 156 tons/year) were accompanied by a

corresponding increase of ΣPFCA fluxes from 2 to 3.5 pg/cm2/year between 1985 and 2002 in

Lake Oesa and from 2 to 4.6 pg/cm2/year between 1989 and 2003 in Lake Opabin(75).

Subsequent decline in the fluxes for some PFCA congeners was also observed between mid-

2003 and 2008 in both lakes, which may be due to recent government and industry efforts to

reduce emissions of FTOHs and PFCAs in current telomer-based production, as well as, their

residuals in the final products (71). These results suggest that atmospheric oxidation of volatile

fluorinated precursors is a major source of fluorochemical contamination in remote

environments, as was previously demonstrated in two isolated lakes in the Canadian Arctic(41),

and corroborate previous work byYoung et al.(77). In that study, PFOA and PFNA were

measured in snow sampled from high-altitude ice caps in the Arctic and when their

concentrations were converted to yearly fluxes(77), the data corresponded well with

90

thosemodeled by Wallington et al.(19) based on estimated FTOH emissions and PFCA yields

from atmospheric degradation of FTOH. In addition, the strong correlation observed in the

levels between the atmospherically-derived PFCA product pairs (i.e. PFOA and PFNA; PFDA

and PFUnA) and the rapid response observed in the ice cap PFOS concentrations to the phase-

out of PFOS production both support atmospheric transport of volatile fluorinated precursors as

the sole source of contamination observed in the ice caps (77).

2.3.2 Wastewater Treatment Plant Sludge

As WWTPs receive influents from predominantly anthropogenic sources, the observed

contamination of PFASs in sludge, as shown in Table 2.3, is direct evidence of human exposure

to fluorinated chemicals. In comparison to sediment concentrations, total ΣPFCAs and ΣPFSAs

in sludge samples are consistently at least one order of magnitude higher, with concentrations as

high as 7500 ng/g dry weight reported in a primary sludge sample collected in Hong Kong (44).

Table 2.3.Ambient concentrations of ΣPFCAs and ΣPFSAs (ng/g dry weight) reported in

selected WWTP monitoring campaigns conducted around the world.

Mean Concentration*

(ng/g dry weight)

Country ΣPFCAs ΣPFSAs Range of PFASs

observed Reference

Canada 3.39 104.18 <LOD-203.9 (78)

Denmark 10.30 22.80 0.40-18.40 (79)

Korea 440.00 76.00 <LOD-190 (80)

Mainland China** 1517.38 1191.06 - (81)

Mainland China 502.07 46.20 0.41-279.00 (49)

Mainland China 600.21 49.86 <LOD-561.90 (82)

Netherlands** - 40.50 - (55)

Spain 59.78 89.89 <LOD-84.18 (83)

Spain 11.46 65.45 0.28-63.99 (84)

Switzerland 19.07 336.86 1.60-333.33 (85)

Thailand 795.65 571.90 3.10-474.75 (86)

USA 29.58 565.08 <LOQ-308.45 (12)

USA** - 144.00 - (87)

USA 182.75 31.00 <LOQ-107.00 (88)

*Concentrations reported in sludge samples collected from different locations within the same study were

summarized here as an overall arithmetic mean. **Only PFOA and/or PFOSwas monitored.

91

Similar to the global distribution observed in sediments, PFCA concentrations were the

highest in sludge samples from Asian-Pacific countries, such as Korea, China, and Thailand,

while PFOS concentrations in sludge appeared relatively consistent in most countries. As was

observed in sediments, long chain PFCAs (≥7 CFs) and PFOS are the dominant PFAAs observed

in sludge, except for those samples collected in Shanghai, China in which TFA and other short

chain PFCAs (<7 CFs) were measured at either higher or similar concentrations as compared to

the longer chain PFCAs (49). Recent attempts to monitor other classes of PFAAs did not detect

any PFPAs in sludge (55), although two perfluoroalkylphosphinate (PFPiA) congeners (C6/C6

and C6/C8 PFPiAs) were observed at ~2 ng/g (89).

A distinct pattern is typically present in the congener profile of the PFCAs detected in

sludge in which an even-carbon chain length PFCA (e.g. PFOA (C8), perfluorodecanoate

(PFDA, C10), or perfluorodecanoate (PFDoA, C12)) was often observed at higher concentrations

than the adjacent odd-carbon chain length PFCA (e.g. PFNA (C9), PFUnA (C11), or

perfluorotridecanoate (PFTrA, C13)). This even > odd-carbon chain PFCA pattern is consistent

with the biological production of PFCAs from fluorotelomer-based materials (90–93). The

discovery of fluorinated commercial products in WWTP sludge was first reported by D’eonet al.

who detected a suite of varying chain lengths (4:2 to 12:2) of polyfluoroalkyl phosphate diesters

(diPAPs) at concentrations ranging from <LOD to 200 ng/g (78) (Fig.2.2). These fluorotelomer-

based surfactants are used as greaseproofing agents in food contact paper, as well as leveling

agents in personal care and cosmetic products and have a demonstrated ability to degrade into

PFCAs in WWTP media (94). Detection of 6:2 and 8:2 fluorotelomer unsaturated carboxylates

(FTUCAs) in WWTP sludge (80, 85), both of which are metabolic intermediates of

fluorotelomer-based precursors, may also indicate previous exposure to diPAPs and other

fluorotelomer-based materials. The potential for these and other types of precursor materials to

enter WWTPs and undergo biodegradation is supported by observed increases in mass flows of

PFCA and PFOS concentrations from influent to effluent samples (88, 95).

92

Figure 2.2.Concentrations of diPAPs (ng/g) observed in WWTP sludge samples collected from

Ontario, Canada and in NIST SRM WWTP sludge sample.

The occurrence of PFASs in WWTPs may lead to contamination of farmlands upon land

application of waste materials generated at these facilities. A suite of PFASs has been measured

at total concentrations of 3-35 ng/g dry weight in compost and digestate samples collected from

commercial plants in Switzerland (96). Composting and digestion are common waste

management practices in Europe and the resulting organic waste materials are often applied to

agricultural soils. Application of WWTP materials to agricultural farmlands may lead to

significant contamination of soil and its surrounding environment, as will be discussed.

2.3.3 Soils

In comparison to sediments and sludge, considerably fewer studies have focused on

measuring PFASs in the soil environment. Table 2.4 summarizes the contamination of PFAAs

observed in selected soils collected worldwide.

4:2

diP

AP

4:2

/6:2

diP

AP

6:2

diP

AP

6:2

/8:2

diP

AP

8:2

diP

AP

8:2

/10:2

diP

AP

10:2

diP

AP

10:2

/12

:2 d

iPA

P

Co

ncen

trati

on

in

slu

dg

e (

ng

/g)

0

50

100

150

200

250

WWTP Sludge

NIST SRM Sludge

93

Table 2.4.Ambient concentrations of ΣPFCAs and ΣPFSAs (ng/g dry weight) reported in

selected soil monitoring campaigns conducted around the world.

Mean Concentration*

(ng/g dry weight)

Country ΣPFCAs ΣPFSAs Range of PFASs

observed Reference

Antarctica 2.02 1.29 <LOD-1.29 (97)

Japan 20.06 2.87 <LOQ-11.67 (98)

Mexico 0.76 10.10 <LOQ-10.10 (98)

Mainland China 175.15 9.71 0.01-135.96 (49)

Mainland China 0.36 0.62 nd-0.33 (51)

Mainland China 1.59 nd nd-0.61 (52)

Tierra del Fuego 0.98 0.46 <LOD-0.46 (97)

USA 36.47 1.59 <LOQ-21.26 (98)

*Concentrations reported in soil samples collected from different locations within the same study were summarized

here as an overall arithmetic mean.

The observed PFCA and PFSA concentrations are generally consistent among the

different countries, again with the exception of TFA, which was detected at concentrations at 1-2

orders of magnitude greater than the other PFAAs in soils collected from Shanghai, China (49,

82). Specific sources of this TFA contamination were not elucidated, although the authors

speculated precipitation and surface water contamination may be potential contributors. PFOS

was the dominant PFSA congener observed in most soils.

PFAS contamination in soils may be due to a combination of proximity to both diffuse

(i.e. urban and industrial outputs) and point (i.e. known fluorochemical outputs) sources. The

detection of various PFCAs and PFSAs in Antarctic soils (97) also highlights the role that

atmospheric transport plays in delivering volatile fluorinated precursors and their PFAA

degradation products to remote regions, as well as, contribute to the background burden observed

in soils and sediments collected worldwide. Davis et al. reported the earliest case of PFOA

contamination (up to 170 ng/g dry weight) in soils collected near a fluoropolymer manufacturing

facility in Parkersburg, West Virginia (13). More recently, environmental surveys of soil, water,

sediment, and biota collected around a training facility using AFFF reported high levels of

ΣPFCAs (24 ng/g dry weight), ΣPFSAs (86 ng/g dry weight), and 6:2 fluorotelomersulfonate

(FTSA,10 ng/g dry weight) (99). Use of commercial fluorinated products was also investigated

94

by Plassmannet al. who detected a suite of semi-fluorinated alkanes (SFAs) and alkenes

(SFAenes), chemicals that are used in ski waxes, at total concentrations of 7 ng/g dry weight in

soils collected from a ski area in Sweden (100).

2.3.3.1 Case Study: Contamination of Agricultural Farmlands in Decatur, Alabama

Application of treated sludge or biosolids to agricultural lands has been identified as a

major source of PFASs to soil, as evidenced by the elevated levels of PFAAs and FTOHs

observed in soil samples (101, 102) collected from farm fields in Decatur, Alabama. These

fields were previously amended with biosolids, some for as long as 12 years, from Decatur

Utilities, a WWTP facility known to have processed effluents from local fluorochemical

industries.

During two sampling periods in 2007 and 2009, ΣPFAA concentrations were measured at

6000 ng/g and 1300 ng/g dry weight respectively in the biosolids-applied soils (101), while total

fluorotelomer alcohols (ΣFTOHs) were measured at 140 ng/g dry weight in the 2009 soil

samples (102) (Fig. 2.3.). These concentrations represent some of the highest levels of PFAAs to

be ever reported in soils, typically 1-2 orders of magnitude higher than the ambient levels

presented in Table 2.4, as well as the background concentrations (typically <LOQ) observed in

soils collected from fields that were not amended with biosolids. PFAA concentrations observed

in the 2007 soil were typically higher than those in the 2009 soil (Fig.2.3.). The authors

speculated these differences may be due to variation in the PFAA concentrations present in the

different batches of biosolids applied to these fields and in the application rates, although the

decline may also be due to the WWTP’s decision to cease their practices of biosolids application

starting in November 2008 (http://www.epa.gov/region4/water/PFCindex.html).

PFAA contamination in the biosolids-amended soils decreased with depth (surface (0-10

cm) > 36-56 cm > 152-165 cm) (Fig. 2.3.). Comparison of the subsurface PFCA concentrations

to those observed at surface level shows higher distribution of perfluorohexanoate (PFHxA, C6)

and perfluoroheptanoate (PFHpA, C7) as compared to PFOA and the longer chain PFCAs, which

suggests the short chain PFCAs may percolate more easily through the soil environment. This

observation has implications for contamination of groundwater and public water supplies located

95

near soil environments that are more heavily contaminated with short chain PFCAs (<7 CFs), as

was observed in Shanghai, China (49).

Figure 2.3.Concentrations of PFCAs, PFOS, and FTOHs observed in soils collected at different

depths in 2007 and 2009 from sludge-amended agricultural fields in Decatur, Alabama. Data

presented here were obtained from Washington et al.(101) and Yooet al.(102).

The C6-C14 PFCAs were detected in all biosolids-amended soils, with PFOA and PFDA

as the dominant congeners observed. PFOS was also observed as a major contaminant (941 ng/g

dry weight, 2007 soil; 135 ng/g dry weight, 2009 soil), while PFHxS was only detected

occasionally in the subsurface soils. As was observed in sludge, concentrations of the even-

carbon chain PFCA congeners (PFOA (C8), PFDA (C10), and PFDoA (C12)) were higher than

the adjacent odd-carbon chain PFCAs (PFNA (C9), PFUnA (C11), and PFTrA (C13) in these

soil samples. This is consistent with the profile of PFCA degradation products from the

transformation of fluorotelomer-based precursor materials that may be present in the soil,

possibly through transfer from the biosolids upon application. This is further supported by the

detection of a suite of FTOHs (7:2 to 14:2), ranging in concentrations of 3-37 ng/g dry weight, in

the same soil samples (Fig. 2.3.). A number of biodegradation studies in both soils and WWTP

media have reported FTOH as an intermediate metabolite during the transformation of various

PF

HxA

PF

Hp

A

PF

OA

PF

NA

PF

DA

PF

Un

A

PF

Do

A

PF

TrA

PF

TeA

PF

OS

tota

l P

FA

S

Co

ncen

trati

on

in

so

il (

ng

/g d

ry w

eig

ht)

0

200

400

600

800

1000

1200

1400

1600

1800

20005000

6000

Sludge-amended soils (0-10 cm) 2007




6:2

FT

OH

7:2

sF

TO

H

8:2

FT

OH

9:2

sF

TO

H

10:2

FT

OH

11:2

sF

TO

H

12:2

FT

OH

13:2

sF

TO

H

14:2

FT

OH

tota

l F

TO

Hs

8:2

FT

Ac

0

20

40

60

80

100

120

140

160

180

200

96

fluorotelomer-based precursors, such as the diPAPs(94), acrylate-based polymers (103, 104), and

most recently, a fluorotelomer stearate monoester (105).

The discovery of elevated PFAS concentrations in these biosolids-applied fields in

Decatur spurred other monitoring studies in plants (39) and groundwater (106) collected in the

vicinity of these fields, all of which reported increased PFAS contamination. A suite of C6-C14

PFCAs and PFOS was observed at concentrations up to 200 ng/g dry weight in various plants

collected from the same fields where the WWTP sludge was applied, as shown in Fig.2.4.

Figure 2.4.Concentrations of PFCAs observed in various plant species collected in 2009 from

sludge-applied fields of Decatur, Alabama (left). Mean grass-soil accumulation factors (GSAFs)

calculated from five plant species (right). This data was obtained from Yooet al. (39).

Grass-soil accumulation factors (GSAFs) were calculated to evaluate transfers of PFAAs

from the sludge-applied soils to plants and were observed to decrease with increasing chain

length (Fig.2.4.). This is consistent with the higher mobility of short chain PFCAs to be taken up

into plants via transpiration of water migrating through the xylem of the plants.

Carbon Chain Length

6 8 10 12 14

Gra

ss

-So

il A

cc

um

ula

tio

n F

ac

tors

(G

SA

F,

Cp

lan

t/C

so

il)

0

1

2

3

4

PFCAs

PFSAs

PF

Hx

A

PF

Hp

A

PF

OA

PF

NA

PF

DA

PF

Un

A

PF

Do

A

PF

TrA

PF

Te

A

Co

nc

en

tra

tio

n i

n p

lan

ts (

ng

/g d

ry w

eig

ht)

0

50

100

150

200

250

Kentucky bluegrass

Tall fescue

Tall fescue

Tall fescue

Bermuda grass

97

2.4 Fate of PFASs in Environmental Solids

As was demonstrated by the case study, contamination is not necessarily contained to one

compartment, but may extend to the surrounding environment depending on the physicochemical

properties of the contaminant. Based on the experimentally-determined pKA’s of PFOA and

PFOS (<1) (107, 108), PFCAs and PFSAs are expected to primarily circulate as anions in the

environment and the extent to which they accumulate in specific compartments may vary

depending on environmental- and chemical-specific parameters. As PFCAs and PFSAs are

perfluorinated, the strength in their carbon-fluorine (C–F) bonds renders them recalcitrant to

environmental and biological degradation processes (109–112). As such, sorption and

bioaccumulation are likely the primary fate of these persistent chemicals. Upon accumulating in

sediments, soil, or WWTP matrices, these PFAAs and other PFASs may undergo a number of

different environmental processes, as will be described next.

2.4.1 Sorption

Both batch sorption experiments and field monitoring of sediments and surface water

have demonstrated the sorption capacity of PFASs. Table 2.5 summarizes the organic carbon-

normalized distribution coefficients (KOCs) that have been measured in the laboratory (20, 23–27,

29, 30, 113) and from field data (43, 50, 53, 56, 61, 63, 70, 73, 114–116). In the majority of

these studies, PFAA sorption exhibited a chain-length dependency such that their KOC values

would increase with the number of CFs present in the perfluoroalkyl chain of the PFAAs studied.

In addition, PFSAs were also observed to be more sorptive than PFCAs of equal perfluoroalkyl

chain length (e.g. logKOC: 2.39, PFNA (8 CF’s) < 2.57, PFOS (8 CF’s) (23)). These observations

are consistent with the distribution of PFAAs observed in environmental samples collected in

Tokyo Bay in which short-chain PFCAs (<7 CFs) were exclusively detected in pore water and

seawater, while the long-chain PFCAs (≥8 CFs), PFSAs (≥6 CFs), FOSA, and EtFOSAA were

predominantly observed in the suspended particulate and sediment samples (73, 114).

98

Table 2.5.Organic carbon-normalized sorption distribution coefficients (logKOC) from laboratory-based batch sorption experiments

and field-based sediment and surface water monitoring. Distribution coefficients in italics are not normalized to organic carbon

(logKd).

Sorption

Medium PFHxA PFHpA PFOA PFNA PFDA PFUnA PFBS PFHxS PFOS PFDS FOSA MeFOSAA EtFOSAA Reference

Laboratory-Based Sorption Data

Sediment - - 2.06 2.39 2.79 3.30 - - 2.57 3.53 - 3.11 3.23 (23)

Sediment - - - - - - - - 2.40-2.60 - - - - (20)

Sediment - - - - - - - - 2.94-3.25 - - - - (24)

Sediment - - - - - - - - 2.94-4.06 - - - - (25)

Sediment - - 2.20-2.60 - - - - - 3.00-4.20 - 3.70-5.00 - - (26)

Sediment - - - - - - - - 3.47 - - - - (27)

Soil - -0.01 0.40 0.99 1.69 - -0.54 - 1.39 - - - - (29)

Activated

Sludge - - 2.18-2.54 - - - - - 2.30-3.61 - - - - (30)

Dry

Sludge - - - - - - - - 1.89-2.44 - - - - (113)

Anaerobic

Sludge - - - - - - - - 2.16-2.32 - - - - (113)

Field-Based Sorption Data

Sediment - - - - - - - - - - - - 2.99 (73)

Sediment - - 1.90 2.40 3.60 4.80 - 3.60 3.80 - 4.30 - 4.80 (114)

Sediment - - 2.63 3.69 - - - - 3.16 - - - - (115)

Sediment - - - - - - - - 2.88-3.67 - - - - (63)

Sediment - - 3.40-5.50 - - - - - 4.50-5.90 - - - - (61)

Sediment - - 1.47 2.06 2.37 2.32 - 0.97 2.10 - 2.56 - - (70)

Sediment 2.10 2.10 - 2.90 3.80 4.70 - 2.20 3.70 - - - - (43)

Sediment 2.20 2.10 2.40 2.80 3.60 - 1.80 2.40 3.40 - - - - (116)

Sediment 2.70-4.70 2.40-4.00 2.60-4.20 3.10-4.30 3.80-4.70 4.00-4.80 - - 3.80-5.10 - - - - (50)

Sediment 2.62 2.70 2.98 3.56 3.74 - 2.79 - 3.58 4.51 - - - (56)

Sediment - - 2.28 - - - 2.16 - 2.88 - - - - (53)

99

80

Matrix-specific characteristics, such as the organic carbon fraction (fOC) of the soil or

sediment, pH of the solid matrices and their surrounding aqueous environment, and aqueous

salinity have also been observed to influence the sorption process of PFAAs. Higgins and Luthy

first demonstrated that PFAA sorption onto sediments was positively correlated with fOCand

aqueous concentrations of divalent cations, such as Ca2+

, and negatively correlated with aqueous

pH (23), and these effects have since been corroborated by other sorption studies in sediments

and soil (20, 25–27, 29). The fact that sorption has also been observed to positively correlate

with perfluoroalkyl chain length suggests the importance of hydrophobic interactions between

the perfluoroalkyl tail of PFAAs and the organic matter of soil and/or sediments. As most

sediments and soil typically carry a net negative surface charge, decreasing the aqueous pH

promotes protonation of oxides and other functional groups present on these solid surfaces and

thus, reduce the repulsion between the less negatively charged surface and the incident PFAA

anion. Similarly, increasing the concentrations of aqueous cations, particularly multivalent ones

like Al3+

, Fe3+

, Ca2+

, and Mg2+

, results in the formation of a cation interlayer that

functionssimultaneously as a barrier to the negatively charged solid surface and as a bridge to

electrostatically bind PFAA anions. These observations suggest both hydrophobic and

electrostatic interactions are important factors in controlling sorption of PFAAs, but it is unclear

as to which dominates the process.

Field-based sorption data are complicated by the heterogeneity of sediment- and aqueous-

specific conditions in the natural environment, as evidenced by the range of logKOC observed for

PFOA (1.47–5.50) and PFOS (2.10–5.90) that spans over 3 to 4 orders of magnitude (Table 2.5).

Although a number of studies have observed correlations between PFAA sorption and fOC in

field-collected sediments (73, 114–116), the occurrence of some of these correlations is limited

to specific PFCA and PFSA congeners, as was only observed for the linear isomers of PFOA and

PFOS by Kwadijket al.(115) and for the C10–C13 PFCAs and PFOS by Lasieret al. (116), while

others have also reported the lack of any correlations (50, 63, 70). Kwadijket al. also did not

observe any correlation between PFAA sorption and the pH and concentrations of Ca2+

of the

surface water sampled (115), which contrasts the observations by Pan and You (63) and Ahrens

et al. (73). These inconsistencies may be due to the diversity of geochemical parameters in the

natural environment, such as occasional nonequilibrium between the aqueous and solid phases;

variable organic carbon content, contaminant concentrations, and salinity; and the potential for

100

biodegradation by microbes and benthic biota to occur, all of which could significantly influence

the partitioning behaviour of PFASs in environmental solids. As such, field-based distribution

coefficients cannot be compared with those measured in the laboratory in which these

aforementioned parameters can be strictly controlled.

The potential for sorbed PFAAs to remobilize into the aqueous environment has been

examined in various desorption experiments (24, 25, 29). You et al. observed hysteresis for

PFOS at varying levels of salinity, with the desorption coefficients, logKdes, increasing with

increasing concentrations of aqueous CaCl2(25), as would be expected based on the cation-

bridging mechanism described above. In addition to high saline conditions, the presence of

cationic alkylammonium-based surfactants has also been demonstrated as an effective barrier for

immobilizing PFOS upon sorption to sediments (24). Desorption of PFAAs is also dependent on

the perfluoroalkyl chain length, as was observed by the increase in logKdesfrom 0.30 for PFHpA

(C7) to 1.71 for PFDA (C10) and from 0.08 for PFBS (C4) to 1.56 for PFOS (C8) by Enevoldsen

and Juhler(29). This suggests short-chain PFAAs (<7 CFs) may desorb more easily into the

aqueous environment, as compared to the longer-chain PFAAs (≥8 CFs).

2.4.2 Leaching to Surface Waters and Groundwater

Both laboratory and field studies have shown considerable evidence of PFAAs leaching

from soils, that have been spiked with PFAAs (117) or exposed to contaminated street runoffs

(118, 119) and WWTP sludge (106, 117, 120, 121), to groundwater and surface water.

Murakami et al. performed soil infiltration column tests in which PFAA-spiked artificial

street runoffs were fed either continuously or intermittently through a loamy soil for 80–160 days

(119). Removal of PFAAs by the soil was observed to increase from <20% for PFOA to >80%

for PFUnA and from PFCA (e.g. ~20% for PFNA) to PFSA (e.g. ~70% for PFOS) of equal

perfluorocarbon chain length(118, 119). Gellrichet al. performed similar flow-through column

experiments in which loamy sand was spiked once either with PFAAs or contaminated WWTP

sludge, followed by intermittent additions of water for two years (117). Analysis of the

percolating water revealed a chain length-dependency of PFAAs leaching from the soil, such that

short-chain PFCAs and PFSAs (≤7 CFs) eluted at speeds corresponding to their size (i.e. PFBA ~

PFBS ~ perfluoropentanoate (PFPeA, C5) ~ PFHxA>PFHpA ~ PFHxS> PFOA), while PFOS

and long-chain PFCAs (≥8 CFs) were not detected even after two years (117). Stronger retention

101

to the soil was also observed when contaminated sludge was used as the source of PFAAs in the

experiments, but the overall elution order remained the same (117). These results corroborate

the PFAA congener profiles typically observed in environmental samples. For example,

Lindstrom et al. calculated the ratios of the concentrations of PFCAs and PFSAs in groundwater

and surface water to those measured in soil from nearby contaminated biosolids-applied fields

and observed these ratios increased with decreasing perfluoroalkyl chain length (106), consistent

with the higher mobility of short-chain PFAAs.

2.4.3 Biodegradation in WWTP Media and Soils

In contrast to the persistent PFCAs and PFSAs, a number of studies have demonstrated

the biotransformation of fluorotelomer-based and perfluoroalkanesulfonamido-based substances

in WWTP-simulated and soil systems (90, 94, 103–105, 122–126), most of which results in the

production of PFCAs and PFSAs respectively. A more in-depth discussion of this topic is

provided in Chapter 1.4.1.2.

2.4.4 Uptake into Vegetation

To date, only three studies have measured PFASs in plants, with one that reported the

occurrence of FTOHs and PFAAs in grass collected from the contaminated farm fields in

Decatur, as was discussed above (39), and another that detected C5 to C11 PFCAs and C4, C6,

and C8 PFSAs in floating plants from Baiyangdian Lake in China (54). In the latter study, Shi et

al. observed similar ΣPFAS concentrations (11–19 ng/g dw) across the three different species of

aquatic plants sampled (54).Most recently, Müller et al. observed low levels of PFCAs (20–260

pg/g ww) and PFOS (2–62 pg/g ww) in lichen and plants sampled from Northern Canada (127).

PFOA and PFOS have a demonstrated capacity to transfer from contaminated soils to

plants (38, 39, 128). Stahl et al. observed uptake of both analytes in wheat, oats, corn, ryegrass,

and potatoes sown in PFOA- and PFOS-spiked soil, with preferential accumulation observed in

the vascular compartments, as compared to the internal storage organs (128). This was

evidenced by the plant-soil accumulation factors (PSAFs) calculated from the ratio of the plant

to soil concentrations of PFOA and PFOS concentrations reported by Stahl et al.(128), as shown

in Table 2.6.

102

Table 2.6.Plant-soil accumulation factors (PSAFs = Cplant/Csoil) calculated from the data provided

by Stahl et al.(128) and taken directly from Lechner and Knapp (38) and Yooet al.(39).

Plant Compartment

Plant-Soil Accumulation Factors

(PSAF = Cplant/Csoil) Reference

PFOA PFOS

Maize Corn Ears 0.006 ± 0.001 0.004 ± 0.001

(128)

Straw 0.244 ± 0.035 0.160 ± 0.020

Oat Grain 0.058 ± 0.016 0.008 ± 0.003

Straw 1.946 ± 0.850 0.447 ± 0.144

Potato Tuber 0.001 ± 0.000 0.000 ± 0.000

Peels 0.004 ± 0.001 0.012 ± 0.002

Spring Wheat Grain 0.062 ± 0.023 0.000 ± 0.000

Straw 2.762 ± 0.653 0.773 ± 0.247

Perennial

Wheatgrass

First Cutting 0.324 ± 0.060 0.128 ± 0.033

Last Cutting 4.456 ± 1.247 1.428 ± 0.726

Potato

Edible Parts 0.010 0.000

(38)

Peels 0.030 0.020

Leaves, Stalks,

and Roots 0.380 0.270

Carrot


Peels 0.040 0.030

Leaves, Stalks,

and Roots 0.530 0.320

Cucumber


Leaves, Stalks,

and Roots 0.760 0.120

Grass - 0.250 ± 0.103 0.070 ± 0.018 (39)

Similarly, Lechner and Knapp observed higher PSAFs in the transport compartments (i.e.

leaves, stalks, and roots) of potatoes, carrots, and cucumbers grown in biosolids-amended soil, as

compared to those measured in the edible parts of the vegetable (38) (Table 2.6). These results

suggest PFAA accumulation occurs more intensively in the vascular tissues that are responsible

for water-borne transport of nutrients within the plant via evatranspiration. This is consistent

with the inverse relationship between the PSAFs and carbon chain length observed for the C6–

C14 PFCAs (Fig. 2.4.) by Yooet al. (39) that demonstrates preferential plant uptake of the

shorter-chain and more water soluble PFCAs. The fact that PFOA tends to be taken up more

readily in the plant than PFOS, as was observed in these studies (Table 2.6), is also concurrent

with the stronger sorption capacity of PFOS to sediments (23) and soil (29), as compared to

PFOA.

103

These observations have important implications for assessing the risks of animal and

human exposure from consuming specific plant compartments that may be more or less

contaminated than others. Nevertheless, the demonstrated accumulation of PFAAs (38, 39, 128)

suggests plant uptake may be an important source of PFASs to the food chain and as such,

future monitoring should direct more efforts towards addressing the current paucity of data in

this environmental compartment.

2.5 Summary and Future Outlook

The global detection of PFASs in sediments, WWTP sludge, and soil described here is a

testament to the prevalent use of these chemicals in commercial applications and their

persistence in the environment. Atmospheric transport of volatile fluorinated precursor

materials, such as those shown to offgas from commercial products (14), and the subsequent

deposition of their PFAA degradation products via precipitation (129) may in part contribute to

the background contamination observed in sediments and soils sampled worldwide and

especially in remote regions (41, 97) where local anthropogenic inputs are considered minimal.

On the other hand, wastewater discharges of commercial fluorinated products are important point

sources to local contamination observed in urban and industrial locations. WWTP sludge may

serve as a useful proxy to determine environmental exposure to anthropogenic emissions in near-

source regions. Global PFAA contamination in WWTP sludge, as shown in Fig. 2.5, suggests

the Asia-Pacific region may be a hotspot for contamination, which is consistent with

environmental data generated from this region. As the North America shifts towards stricter

regulation and goals of decreasing emissions, contributions towards global PFAS contamination

from Asian countries may become more significant, especially in China where large-scale

production of POSF-based materials has resurged (72).

The extent to which the abovementioned environmental processes of PFASs occur is

dependent on the physicochemical properties of the contaminant, such as the perfluoroalkyl

chain length. Emission of short-chain (≤7 CFs) PFAAs was considered minimal in the past,

primarily as residual impurities in the predominantly C8-based fluorochemical industry. As

current manufacturing processes shift towards the perfluorohexyl (6 CFs)- and perfluorobutyl (4

CFs)-based chemistries, contamination of these short chain congeners may become increasingly

important, especially in the aqueous compartment, based on their demonstrated ability to desorb

104

much more rapidly from environmental solids, as compared to longer chain congeners. On

theother hand, remobilization of long-chain (≥8 CFs) PFAA into the aqueous environment may

eventually occur through gradual desorption of bound chemicals from legacy contamination and

the release of degradation products from the transformation of commercial precursor materials

present in environmental solids.

Figure 2.5.Concentrations of ΣPFCAs and ΣPFSAs observed in WWTP sludge collected around

the world. Note: Some of these concentrations were obtained by averaging total concentrations

reported in multiple monitoring campaigns within the same country to yield an overall mean for

that country. *PFOS was the only PFAA monitored in the Netherlands campaign; therefore,

total ΣPFSA concentration = total PFOS concentration.

A major limitation in interpreting monitoring data is the disparity in the fluorinated

analytes (i.e. different chain lengths, terminal PFAA degradation products versus intermediate

metabolites and/or precursor materials) that are currently being included for analysis. This is

especially problematic during comparisons of ΣPFAS concentration data, as the summed

contributions from the individual analytes may differ from study to study depending on which

congeners were chosen to be monitored. As such, while total concentration data may be useful

for comparing regional contamination, they may not be fully representative of the actual

fluorochemical contamination present in the sampled matrix. However, there has been increased

Can

ad

a

Den

mark

Germ

an

y

Ko

rea

M. C

hin

a

Neth

erl

an

ds

Sp

ain

Sw

itzerl

an

d

Th

ailan

d

US

A

P

FA

A c

on

ce

ntr

ati

on

(n

g/g

dry

weig

ht)

0

200

400

600

800

1000total PFCAs

total PFSAs

*

105

effort to broaden the range of target fluoroalkyl analytes in environmental monitoring to include

other emerging species, such as the diPAPs(78), SFAs and SFAenes(100), and PFPAs and

PFPiAs(55, 89). This is important as these chemicals are often present at percent quantities as

either the active or inert components in commercial products (130–132), yet their contribution as

direct and/or indirect sources to environmental PFAS contamination is currently not well

understood.


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117

CHAPTER THREE

Biodegradation of Polyfluoroalkyl Phosphates (PAPs) as a Source of Perfluorinated Acids

to the Environment

Holly Lee, Jessica D’eon, and Scott A. Mabury

Published as: Environ. Sci. Technol. 2010, 44, 3305-3310.

Contributions: Holly Lee was responsible for designing and executing the biodegradation

experiments, LC-MS/MS method development, sample acquisition, and data interpretation.

Synthesis of the monoPAPs and diPAPs used for spiking the biodegradation experiments and

the subsequent analysis of these chemicals by LC-MS/MS were performed by Holly Lee

under the guidance and training of Jessica D’eon. Holly Lee prepared this manuscript with

editorial comments provided by Jessica D’eon and Scott Mabury.

Reproduced with permission from Emvironmental Science and Technology

Copyright American Chemical Society 2010

118

3.1 Abstract

Wastewater treatment plants (WWTPs) have been identified as a major source of

perfluorocarboxylates (PFCAs) to aqueous environments. The observed increase in PFCA mass

flows from WWTP influent to effluent suggests the biodegradation of commercial fluorinated

materials within the WWTP. Commercial fluorinated surfactants are used as greaseproofing

agents in food-contact paper products, as well as leveling and wetting agents. As WWTPs are

likely the major fate of these surfactants, their biodegradation may be a source of PFCA

production. One class of commercial surfactants, the polyfluoroalkyl phosphates (PAPs), have

been observed in WWTP sludge. While PAPs have been shown to degrade into PFCAs in a rat

model, the present study investigates their microbial fate to determine whether the

biodegradation of PAPs within a WWTP-simulated system will contribute to the load of PFCAs

released. PAPs are applied commercially in mixed formulations of different chain lengths and

substitution at the phosphate center. The effect of chain length and phosphate substitution on the

biodegradation of PAPs was investigated by incubating mixtures of 4:2, 6:2, 8:2, and 10:2

monosubstituted PAPs (monoPAPs) in an aerobic microbial system, and by separately incubating

the 6:2 monoPAP and 6:2 disubstituted PAP (diPAP) for 92 days. Headspace sampling revealed

production of the fluorotelomer alcohols (FTOHs) from the hydrolysis of the PAP phosphate

ester linkages. Analysis of the aqueous phase revealed microbial transformation of the PAPs to

the final PFCA products was possible. The majority of the oxidation products observed were

consistent with previous investigations that have suggested fluorotelomer precursor compounds

degrade predominantly via a β-oxidation-like mechanism. However, in this study, the detection

of odd-chain PFCAs suggests that other pathways may be important. The present study

demonstrated microbially-mediated biodegradation of PAPs to PFCAs. This observation,

together with the diPAP concentrations observed in WWTP sludge, suggest PAPs-containing

commercial products may be a significant contributor to the increased PFCA mass flows

observed in WWTP effluents.

3.2 Introduction

In near-source regions, perfluorinated carboxylic acids (PFCAs) emitted from wastewater

treatment plants (WWTPs) have been identified as a major source of PFCA contamination to

119

aqueous environments (1, 2). PFCA concentrations have also been observed to increase from

WWTP influent to effluent (3–5). In one of two WWTPs studied in-depth, Sinclair and Kannan

(5) found strong correlations between the concentrations of perfluorooctanoate (PFOA) and

perfluorononanoate (PFNA) and between perfluorodecanoate (PFDA) and perfluoroundecanoate

(PFUnA), with higher concentrations of the even chain length PFCA observed as compared to

the odd chain lengths. This PFCA congener profile is consistent with the biological production

of PFCAs from fluorotelomer-based materials (6–11), and appears to result from biodegradation

within the WWTP, as no correlation was observed between PFDA and PFUnA before activated

sludge treatment. These studies together suggest that the biodegradation of fluorotelomer-based

materials within WWTPs may be a source of PFCAs to the environment.

Biotransformation of fluorotelomer alcohols (FTOHs) to PFCAs has been observed in

mixed microbial systems and WWTP sludge (6–8), soil (9), rat hepatocytes and microsomes (10,

11) and whole rat models (12). FTOHs have no known direct commercial application, but are

instead used as building blocks in the synthesis of fluorinated polymers and fluorinated

surfactants, which are themselves incorporated in the final sales products (13). Some evidence

of polymeric degradation into PFCAs was recently reported in two soil biodegradation studies of

a fluorotelomer acrylate polymer although the importance of this pathway is still widely debated

(14, 15). Fluorinated surfactants may also be potential precursors to PFCAs, as the surfactants

are expected to be less sterically constrained to microbial attack as compared to the polymers.

The polyfluoroalkyl phosphates (PAPs) are commercial fluorinated surfactants used primarily in

food-contact paper products and as leveling and wetting agents (16–19). PAPs have been

identified as a potential source of human PFCA exposure as these chemicals can leach out of

food packaging into food (20, 21). Biotransformation of PAPs to PFCAs was observed in a rat

model (22), and human exposure was confirmed by recent measurements of PAPs in human sera

at µg/L (ppb) concentrations (23). After consumer use, PAPs-containing products may be

released into WWTPs. Recent detection of the disubstituted PAP (diPAPs) in WWTP sludge at

levels (i.e. 50-200 ng/g) comparable to perfluorooctane sulfonic acid (PFOS) and far exceeding

the PFCAs demonstrates the potential for these chemicals to contribute to the PFCA

contamination observed in WWTPs (23). The present study investigated microbial

transformation of PAPs to PFCAs by incubating in-house synthesized monosubstituted and

120

disubstituted PAPs (monoPAPs and diPAPs) with aerobic microbes collected from a local

WWTP.

For oil repellency applications, PAPs are generally applied as a mixture of varying

fluoroalkyl chain lengths, as well as the mono-, di-, and tri-substituted phosphate congeners (21,

24). As a result, two studies were performed. The first, hereinafter called the ―substitution

experiment‖, was performed to compare the effect of substitution at the phosphate centre on

PAPs biodegradation and involved separate incubations of the 6:2 monoPAP and 6:2 diPAP.

The second, hereinafter called the ―chain length experiment‖, was performed to compare the

effect of chain length on PAPs biodegradation and involved the incubation of monoPAPs of four

different chain lengths (4:2, 6:2, 8:2, and 10:2). The PAP phosphate ester linkage is expected to

undergo microbially-mediated hydrolysis to produce the corresponding FTOH, which based on

previous investigations, is expected to oxidize to the PFCAs.

3.3 Experimental Section

3.3.1 Chemicals

The synthesis of the PAPs is described elsewhere (22). A list of all chemicals used in this

study is provided in the Supporting Information (SI) in Appendix A. All target analytes are

listed in Table 3.1.

3.3.2 Purging control experiment

Purging has been demonstrated to be an effective technique for removing unreacted

FTOHs from the synthesis of commercial fluorinated materials dissolved in the aqueous phase

(25). Analysis of the PAPs used in this study revealed the presence of FTOHs at significant

quantities in the starting monoPAPs and 6:2 diPAP. As a result, it was important to reduce the

levels of FTOHs in the starting PAP materials as much as possible before microbial inoculation

so that any FTOH or PFCA production in the biodegradation experiments could be attributed to

the degradation of PAPs. As PAPs are highly surface active, the effect of purging on the

aqueous concentrations of PAPs over time was investigated.

121

Table 3.1. Structures, names, and acronyms of the target analytes in this study.

Structure Name Acronym

P

O

HO

OH

OCH2CH2(CF2)xF

P

O

HO

OCH2CH2(CF2)6F

OCH2CH2(CF2)6F

HOCH2CH2(CF2)xF

HO CH2(CF2)xF

O

HO CH = CF(CF2)xF

O

HO CH2CH2(CF2)xF

O

HO (CF2)xF

O

Monosubstituted

polyfluoroalkyl phosphate

x:2 monoPAP

x = 4, 6, 8, 10

Disubstituted

polyfluoroalkyl phosphate 6:2 diPAP

Fluorotelomer alcohol x:2 FTOH

x = 4, 6, 8, 10

Saturated fluorotelomer

carboxylic acid

x:2 FTCA

x = 4, 6, 8, 10

Unsaturated fluorotelomer

carboxylic acid

(x+1):2 FTUCA

x = 3, 5, 7, 9


carboxylic acid

x:3 FTCA

x = 3, 5, 7, 9

Perfluorocarboxylic acid PFCA

x = 3 – 10

122

The experiment was performed in a purge-and-trap system described elsewhere (25), and

illustrated in Figure A1 (Appendix A). Briefly, 400 µg of 4:2, 6:2, 8:2, 10:2 monoPAPs, and 6:2

diPAP were spiked into two sets of polypropylene bottles containing 400 mL of deionized water,

with one set purged with air for 6-7 days, and the other set left to stand. The aqueous phase was

routinely sampled. At the end of the experiment, the gas diffuser tubes (only in the purging

bottles), septa, bottle caps, and bottles were sonicated in methanol at 60oC for 1 hour.

Experimental set-up, extraction, and chromatographic analysis are described in Appendix A.

3.3.3 Biodegradation experiments using aerobic WWTP microbes

Mixed liquor, a mixture of raw wastewater and sewage sludge, was collected from

Ashbridges Bay WWTP (Toronto, ON). Prior to being used as inocula or autoclaved as sterile

controls, the mixed liquor was aerated with in-house air to maintain viability. Both the chain

length and substitution experiments were performed in a purge-and-trap system with

polypropylene bottles containing a total volume of 400 mL of aqueous phase. The setup

included: (1) ―Mixed liquor only‖ control bottles (n = 2), with 10% v/v of washed mixed liquor

in mineral media, were included to monitor any production of FTOHs or PFCAs from potential

fluorinated materials present in the WWTP mixed liquor; (2) ―Sterile‖ control bottles (n = 2),

with 10% v/v of autoclaved mixed liquor, 400 µg of monoPAPs for the chain length experiment

or 400 µg of 6:2 monoPAP and diPAP for the substitution experiment, 300 mg of Hg2Cl2, and

mineral media, were included to quantify any non-microbial-mediated degradation (Hg2Cl2 was

subsequently added at various timepoints to maintain sterility); (3) ―PAPs only‖ control bottles

(n = 2), with 400 µg of monoPAPs for the chain length experiment or 400 µg of 6:2 monoPAP

and diPAP for the substitution experiment, and mineral media, were included to quantify abiotic

degradation; (4) ―Experimental‖ bottles (n = 3), with 10% v/v of washed mixed liquor, 400 µg of

monoPAPs for the chain length study or 400 µg of 6:2 monoPAP and diPAP, and media (Table

A2 in Appendix A). Prior to microbial inoculation, each bottle spiked with PAPs was purged for

5 days to strip the system of residual unreacted FTOHs that may have carried through the

synthesis. After purging, the FTOHs present in the starting PAPs were reduced to within their

detection limits. After microbial inoculation, each bottle was continuously purged with air for 92

days to strip volatile products (e.g. FTOHs) from the system. FTOHs were collected using

XAD-2 cartridges. The aqueous phase was sampled to monitor the production of nonvolatile

123

metabolites and disappearance of PAPs. Preparation of the mineral media, washing procedures

of the WWTP mixed liquor, extraction procedures, and chromatographic and instrumental

conditions are described in detail in Appendix A.

3.3.4 Quality assurance of data

PFCAs and the saturated and unsaturated fluorotelomer carboxylates (FTCAs and

FTUCAs) were quantified using internal calibration (Table A3 in Appendix A). Due to the lack

of native and internal standards for 3:3, 5:3, and 9:3 FTCAs, these analytes were quantified using

4:2, 6:2, and 10:2 FTCAs as surrogate standards. PAPs were quantified by external calibration

as no appropriate internal standards were available. To confirm that external calibration was

appropriate, the PAPs were spiked in mineral media treated with autoclaved mixed liquor and

analyzed by both standard addition and external calibration for comparison. Details are discussed

in Appendix A.

Recoveries for the FTOHs were in the range of 58 – 91% (Table A4 in Appendix A).

The XAD cartridges used included a second XAD plug that acted as a breakthrough to determine

any potential FTOH loss. The breakthroughs in all the vessels contained <10% of the total

FTOHs recovered by the XAD cartridges, except for 4:2 FTOH, where 23% of the total mass

produced was found in the breakthrough. Any levels found in the breakthroughs were summed

together with the sampling section to obtain a total amount of FTOH trapped in the cartridges.

Recoveries for the PFCAs, FTCAs, FTUCAs, and PAPs were in the range of 33 – 153% (Table

A3 in Appendix A). Values were reported as measured and were not corrected for recovery. The

spike and recovery procedures are described in Appendix A.

Contamination was accounted for using both instrumental blanks and procedural blanks

(n = 3, for the extraction of each timepoint). For analytes absent in the procedural blanks, the

limits of detection (LOD) were defined as the concentration with a signal-to-noise ratio (S/N) ≥

3, while the limits of quantitation (LOQ) were set at the concentration with a S/N of 10 (26). For

analytes present in the procedural blanks, the LODs and LOQs were calculated as 3 and 10

standard deviations of the mean blank signals respectively (26). The LODs and LOQs are listed

in Table A5 in Appendix A. Values less than the LOD were assigned a value of zero, while

values less than the LOQ were used unaltered to calculate arithmetic means (±standard error) of

the levels in the replicate bottles at each timepoint.

124

Viability of the microorganisms was verified by adding 6:2 FTUCA as a positive control

to bottles (n = 2) treated with active mixed liquor at day 21, day 51, and day 85. Each 6:2

FTUCA spike occurred when the amounts of the 6:2 FTUCA reactant had diminished to <1% of

the initial dose and the amounts of the 5:3 FTCA, perfluoropentanoate (PFPeA), and

perfluorohexanoate (PFHxA) products had leveled off. The 6:2 FTUCA degraded to produce 5:3

FTCA, PFPeA, and PFHxA, at 39%, 20%, and 37% yield respectively (Figure A4 in Appendix

A).

3.4 Results and Discussion

3.4.1 Purging control experiment

The effect of purging on the aqueous concentrations of 4:2, 6:2, 8:2, 10:2 monoPAP and

6:2 diPAP within this experimental setup was investigated, and a more detailed discussion of the

results is provided in Appendix A.

Purging of the system resulted in a decrease in the aqueous concentrations of the

monoPAPs over time, with the effect being more pronounced for the longer 8:2 and 10:2

monoPAP chains (Figure A5 in Appendix A). This decrease may be due to adsorption of the

PAPs to surfaces within the system, such as the bottle walls, caps, septa, and gas diffuser tubes.

Additionally, the surface-active PAPs may also be physically removed from the aqueous phase in

aerosols forming at the air-water interface and leaving the headspace of the bottles through the

XAD cartridges. At the end of the experiment, 62±4%, 37±16%, 26±10%, and 15±10%, of 4:2,

6:2, 8:2, and 10:2 monoPAPs respectively and 97±12% of 6:2 diPAP were accounted for from

both the aqueous phase and from sonication of the septa, the gas diffuser tubes, the bottle caps,

and the bottles themselves. Despite losing a portion of the starting materials from the aqueous

phase, purging successfully removed the bulk of the FTOH impurities, while sufficient amounts

of the PAPs remained in the dissolved phase to proceed with the biodegradation experiments.

After purging for 5 days, the aqueous concentration of PAPs was measured before

inoculating the bottles with microbes to begin the biodegradation experiments. The purge-and-

trap system inherently minimized the production of PFCAs as any FTOH produced from the

degradation of the PAPs would be largely removed from the aqueous phase. As a result, mass

balance calculations to account for the production of volatile and nonvolatile metabolites were

125

not performed. Instead, the product yield of FTOH, the immediate metabolite from microbial

hydrolysis of PAPs, was estimated based on the initial mass of PAPs added to the bottles prior to

purging.

3.4.2 Biodegradation of 6:2 monoPAP vs. 6:2 diPAP (“Substitution” Study)

The main degradation pathway of PAPs in WWTPs is likely to be microbial hydrolysis of

the phosphate ester bonds to produce FTOHs, which may further oxidize to produce the PFCAs

(6–12). Since FTOH production was not observed in any of the control bottles, degradation

observed in the experiments can be attributed to microbial transformation. A proposed

biodegradation pathway for 6:2 diPAP and 6:2 monoPAP is shown in Figure 3.1. While the

microbial pathway from FTOH to PFCA is well documented (6–9), microbial production of

FTOH from PAPs is investigated for the first time here.

Figure 3.1. Proposed degradation pathway of 6:2 diPAP and 6:2 monoPAP. The solid arrows

represent pathways identified in this work. The dashed arrows represent microbial and

mammalian degradation pathways proposed in the literature.

P

O

HOOCH2CH2(CF2)6F

OCH2CH2(CF2)6F P

O

HOOH

OCH2CH2(CF2)6F HOCH2CH2(CF2)6F

6:2 diPAP 6:2 monoPAP

6:2 FTOH

HO CH2(CF2)6F

O

6:2 FTCA

HO CH = CF(CF2)5F

O

6:2 FTUCA

HO CH2CH2(CF2)5F

O

5:3 FTCA

HO (CF2)4F

O

PFPeA

HO (CF2)5F

O

PFHxA

Pathway B:

Ref. (6,12)

HO (CF2)6F

O

PFHpA

Pathway A:

Ref. (10,11)

H3C (CF2)5F

O

5:2 Ketone

H3C (CF2)5F

OH

5:2 sFTOH

Legend

This work

Pathways proposed

in the literature

Legend

This work

Pathways proposed

in the literature

Ref. (6-12)

Ref.

(6-12)

Pathway D:

Ref. (28,29)

Ref.

(10,11)

Pathway C:

Ref. (27)

126

Degradation profiles of 6:2 monoPAP and 6:2 diPAP are shown in Figure 3.2. Following

microbial inoculation of both the 6:2 monoPAP- and 6:2 diPAP-dosed bottles, 6:2 FTOH was

observed in the XAD cartridges collected from the headspace. The production of the acid

metabolites was also observed in the aqueous phase, which suggests that PAPs microbial

degradation may involve a concerted hydrolytic mechanism to produce FTOH intracellularly,

followed by further oxidation.

The intermediate metabolite, 6:2 FTCA, was observed in the aqueous phase, and then

consumed, followed by formation of 6:2 FTUCA and PFHxA. This transformation was

consistent with a mechanism similar to β-oxidation of 8:2 FTCA to PFOA, as first proposed by

Hagen et al. in a mammalian system (12) and Dinglasan et al. in a microbial system (6) (Figure

3.1, Pathway B). Although no analytical standard for the 5:3 FTCA was available, a mass

transition (341.0>237.0) was inferred from the 7:3 FTCA. The 5:3 FTCA was detected

transiently in both the 6:2 monoPAP- and 6:2 diPAP-dosed bottles. Contrary to the previous

suggestion that 7:3 FTCA can undergo β-oxidation to form PFOA (8), recent biotransformation

experiments using 7:3 FTCA as the parent substrate in both microbial (9) and mammalian (11)

systems do not support this pathway. Here, the occurrence of the 5:3 FTCA coincided with the

production of PFPeA, alluding to a novel pathway recently reported by Butt et al. in which

rainbow trout dosed with 7:3 FTCA as the parent reactant was metabolized to form

perfluoroheptanoate (PFHpA) (27) (Figure 3.1, Pathway C). The production of PFPeA may also

be attributed to other precursors. In a soil biotransformation study of 6:2 FTOH, Liu et al.

proposed that 6:2 FTUCA may degrade into 5:2 fluorotelomer ketone (F(CF2)5C(O)CH3), which

can reduce to the 5:2 sFTOH (F(CF2)5CH(OH)CH3), which could then transform to the PFPeA

(28) (Figure 3.1, Pathway D). In another study, Fasano et al. also proposed that 8:2 FTUCA may

undergo hydroxylation, oxidation, and decarboxylation to form the 7:2 ketone (29), but no

literature precedent is currently available to explain the x:2 sFTOH to PFCA pathway.

Production of PFHpA was also observed in this work. In vitro hepatocyte incubations of 8:2

FTCA and 8:2 FTOH as the parent substrates resulted in the production of PFNA, PFHpA, and

even low levels of PFPeA (11), which supports the possibility of oxidation of the α-carbon in

FTCA to form odd-chain PFCAs (Figure 3.1, Pathway A). Furthermore, Martin et al. reported

127

Figure 3.2. Substitution study. (a) Degradation of 6:2 diPAP into 6:2 monoPAP and 6:2 FTOH, (b) Production of aqueous

metabolites in 6:2 diPAP-dosed bottles, (c) Degradation of 6:2 monoPAP into 6:2 FTOH, and (d) Production of aqueous metabolites

in 6:2 monoPAP-dosed bottles. Data are represented as arithmetic means (±standard error) of triplicate incubations. Values less than

the LOD are reported as zero and values in between the LOD and LOQ were used unaltered and indicated with an asterisk (*) in

matching colours.

0 20 40 60 80 100

0

10

2050

100

150

200

0 20 40 60 80 100

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PA

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:2 F

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6:2 monoPAP

6:2 FTOH

0 20 40 60 80 100

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sin

bo

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(n

mo

l)

0

5

10

15

20

PFHxA

6:2 FTCA

6:2 FTUCA

5:3 FTCA

PFPeA

PFHpA

* * * ** * *** ** **

***

**

* * * * *

a)

c)Time (days)

Time (days)

0 20 40 60 80 100

0

2

4

6

8

10

12

14

*** * **

**** *

* * *

* * *** * **

** * * * *

*

** *

*

** *

**

*

*

**

b)

d)

* * * * *

Time (days)

6:2 diPAP

6:2 monoPAP

6:2 FTOH

PFHxA

6:2 FTCA

6:2 FTUCA

5:3 FTCA

PFPeA

PFHpA

Legend

6:2 diPAP

6:2 monoPAP

6:2 FTOH

128

minor production of PFNA in whole rats and isolated rat hepatocytes dosed with 8:2 FTOH (10).

Neither Dinglasan et al. (6) nor Wang et al. (7, 8) observed the production of PFNA from the

biodegradation of 8:2 FTOH; however, the phosphate-free condition of the media used in this

study may have selected for a different microbial strain than were present in the other studies.

As 6:2 diPAP was consumed, 6:2 monoPAP was produced and then itself consumed, all

coinciding with the production of 6:2 FTOH (Figure 3.2a). This profile is consistent with

microbial hydrolysis of 6:2 diPAP to produce a unit of 6:2 FTOH and 6:2 monoPAP, which can

further hydrolyze to release an additional unit of 6:2 FTOH. Due to the steric hindrance at the

di-substituted phosphate center of 6:2 diPAP, 6:2 monoPAP was initially expected to be more

labile to microbial hydrolysis. However, 6:2 diPAP was observed to produce more 6:2 FTOH

with a yield of about 5% at the end of the experiment, as compared to a 1% yield from the 6:2

monoPAP. These yields should be treated as conservative estimates of the transformation of the

6:2 PAPs, because the bioavailability of the PAPs bound up within the system was unknown.

Furthermore, the experimental system used here was designed to probe mechanistically whether

the PAP phosphate ester linkage is susceptible to microbially-mediated hydrolysis, and not to

quantitatively mimic activated sludge treatment within a WWTP at the microscale. The

activated sludge in a WWTP would likely have significantly increased microbial activity and

likely contain more PAP substrates; hence the production of FTOH from PAP would also be

increased. The different reactivity between 6:2 monoPAP and 6:2 diPAP may be influenced by

differences in their binding affinities to the microbial biosolids and other surfaces, as well as

differences in the energy barrier of the respective hydrolysis reactions.

The amount of 6:2 monoPAP in the sterile controls rapidly decreased to less than its

detection limit upon the inoculation of autoclaved mixed liquor, while the amount of 6:2 diPAP

remained relatively consistent throughout the experiment (Figure A7a in Appendix A). The

dianionic phosphate center of 6:2 monoPAP may undergo unique interactions, such as those

previously observed for the sorption of glyphosate, a monosubstituted phosponate herbicide, to

sediments (30). Thus, the monoPAP may bind strongly to the biosolids in the mixed liquor

inoculum. On the other hand, 6:2 diPAP lacks a dianionic center and thus may be less associated

with surfaces. It is unclear whether the bound up fraction of PAPs may be biodegradable;

therefore, the relationship between bioavailability for degradation and sorption warrants further

investigation. In addition, extrapolation from uncatalyzed hydrolysis reaction rates suggests

129

phosphate diesters may be enzymatically degraded faster than the monoesters based on

substantial differences in their activation energies (31).

3.4.3 Biodegradation of the 4:2, 6:2, 8:2 and 10:2 monoPAP (“Chain length” study)

Production of FTOHs was observed in the headspace of the monoPAP-dosed bottles

during microbial incubation. This hydrolysis was microbially-mediated as the evolution of

FTOHs was not observed in the sterile controls. The production of FTCAs, FTUCAs, and

PFCAs in the aqueous phase of the experimental bottles suggests that some of the monoPAPs

were microbially transformed via a concerted mechanism that involved further oxidation of the

FTOH intermediate within the microbial cells.

The production of FTOHs from the monoPAPs is shown in Figure 3.3. Although the four

monoPAP congeners were observed to produce the corresponding FTOHs in relatively similar

yields of 1-2%, the rate of production was observed to decrease significantly as the chain length

of the monoPAP increased. Again, these yields should be treated as conservative estimates as

they were obtained under experimental conditions designed to investigate microbial degradation

and not mimic activated sludge treatment. In addition, the yields were calculated using the total

mass of monoPAPs added to the bottles rather than the monoPAPs measured in the aqueous

phase. Production of 4:2 FTOH leveled off within the first day of the experiment, while the

production of 6:2 and 8:2 FTOHs leveled off at about day 40 and 50 respectively. The

production of 10:2 FTOH did not level off within the length of the experiment. This difference

in the rate of production of FTOHs may be influenced by the distribution of the monoPAP

congeners within the aqueous phase and the steric factors imposed by the different chain lengths.

In the sterile controls, 4:2 monoPAP was consistently detectable throughout the experiment,

while the amounts of 6:2, 8:2, and 10:2 monoPAPs in the aqueous phase decreased substantially

(Figure A7b in Appendix A). This implies that the longer chain monoPAPs may have a stronger

binding affinity to microbial biosolids, although it is unclear whether this would hinder their

bioavailability for degradation. Alternatively, the longer chain monoPAPs may be more

sterically constrained to microbial attack. The slower rate of production of the longer chain

FTOHs implies that the longer chain monoPAPs may be less accessible or susceptible to

microbial degradation.

130

Figure 3.3. Chain length study. Degradation of (a) 4:2, (b) 6:2, (c) 8:2, and (d) 10:2 monoPAPs into FTOHs. Data are represented as

arithmetic means (±standard error) of triplicate incubations. Values less than LOD are reported as zero and values in between the LOD

and LOQ were used unaltered and indicated with an asterisk (*) in matching colours.

0 20 40 60 80 100

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* * * * *

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6:2 FTOH

* * * * *

Time (days)

0 20 40 60 80 100

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8:2 FTOH

0 20 40 60 80 100

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10:2 monoPAP

10:2 FTOH

a) Degradation of 4:2 monoPAP b) Degradation of 6:2 monoPAP

c) Degradation of 8:2 monoPAP d) Degradation of 10:2 monoPAP

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

Legend

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

Legend

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

4:2 monoPAP

4:2 FTOH6:2 monoPAP

6:2 FTOH

8:2 monoPAP

8:2 FTOH

10:2 monoPAP

10:2 FTOH

131

The formation of FTCAs, FTUCAs, and PFCAs in the aqueous phase of the monoPAPs-

dosed bottles is shown in Figure A8 in Appendix A. Both perfluorobutanoate (PFBA) and

PFHxA were observed as β-oxidation products of 4:2 and 6:2 monoPAPs degradation, while

degradation of the longer chain 8:2 and 10:2 monoPAPs ceased at the polyfluorinated

intermediates, 8:2 and 10:2 FTCAs/FTUCAs (Pathway B). On the other hand, while 3:3 FTCA

was not detected as a metabolite of 4:2 monoPAP degradation, production of the 5:3, 7:3, and

9:3 FTCAs were observed as degradation products of the longer chain monoPAPs. The

microbial transformation of the monoPAPs to the acid products appeared to be more operative

for the shorter chain monoPAPs as they were observed to fully degrade to the terminal PFCAs,

whereas the longer chain monoPAPs only partially degraded to the FTCA and FTUCA

intermediates. This difference in reactivity of the monoPAPs may be explained by the steric

constraint of the longer chain lengths to microbial attack and that the longer chain monoPAPs

may be preferentially associated with the various surfaces present in the experimental system, as

have been already discussed.

3.5 Environmental Implications

The biodegradation experiments performed here indicate that PAPs can undergo

microbially-mediated hydrolysis to produce FTOHs, which are known PFCA precursors. This

study is the first to establish a clear link between a commercial product and the production of

PFCAs within WWTPs. Individual diPAPs were observed in WWTP sludge at concentrations

up to 0.5 μg diPAP/g sludge (23), which translates into about 0.5 g diPAP/tonne of sludge

produced. If the conservative estimate of 5% diPAP to FTOH transformation observed here was

used to approximate transformation during activated sludge processing, this results in the

production of about 25 mg FTOH/tonne of sludge. If a 5% conversion of FTOH to PFCA

overtime r4was assumed, a value that is consistent with previous FTOH biodegradation studies

(6–8), this results in the production of approximately 1 mg PFCA/tonne of sludge treated. This

estimate of PFCA production from PAP biodegradation is conservative both because the

microbial activity of activated sludge treatment is expected to be significantly higher than that of

the experiments performed here, and because only the diPAP transformation was considered

here. Based on the composition of commercial products, monoPAPs and triPAPs may also be

present in the WWTP influent. Observed increases in PFCA concentrations between WWTP

132

influent and effluent range from no production to 1-10 g/day (5). The conservative estimate of 1

mg PFCA/tonne of sludge treated from diPAP transformation alone can, depending on the

amount of sewage processed by the facility, account for a significant portion of this PFCA

production. This study demonstrates the potential for biodegradation of commercial fluorinated

products within a WWTP to be a source of PFCAs to the environment.

3.6 Acknowledgements

We would like to thank Susanne Waaijers (University of Amsterdam, Amsterdam,

Netherlands), Hanin Issa and Alexandra Tevlin (University of Toronto, Toronto, ON) for

technical support, and Wellington Laboratories Inc. (Guelph, ON) for donation of mass-labelled

internal standards. This research was funded by a Natural Science and Engineering Research

Council of Canada (NSERC) PGS M to HL and the Ministry of the Environment Best in Science

grant.


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135

CHAPTER FOUR

Biosolids Application as a Source of Polyfluoroalkyl Phosphate Diesters and Their

Metabolites in a Soil-Plant Microcosm: Biodegradation and Plant Uptake

Holly Lee, Alexandra G. Tevlin, and Scott A. Mabury

In preparation: For submission to Environmental Science and Technology.

Contributions: Holly Lee was responsible for designing the greenhouse microcosms in

collaboration with Pablo Tseng, performing the soil-plant biodegradation and uptake

experiments, care and handling of soil-plant systems during the experiment, method

development, sample acquisition, and data interpretation. Alexandra G. Tevlin assisted with

sampling, extractions, and LC-MS/MS analysis of plant samples with assistance from Holly

Lee. Preparation of the manuscript by Holly Lee involved adaptation of a report by

Alexandra Tevlin. Holly Lee prepared this manuscript with editorial comments provided by

Scott Mabury

136

4.1 Abstract

Significant contamination of perfluoroalkyl acids (PFAAs), often observed in wastewater

treatment plant (WWTP) sludge, implicates the practice of applying treated sludge or biosolids

as a major entry route of these chemicals onto agricultural farmlands. Recent efforts to

characterize the sources of PFAAs in the environment have unveiled a number of fluorotelomer-

based materials that are capable of degrading to the perfluorocarboxylates (PFCAs), one of

which, the polyfluoroalkyl phosphate diesters (diPAPs), has been detected in various human

waste materials, such as WWTP sludge and paper fiber biosolids. Here, a greenhouse soil-plant

microcosm was used to investigate the behaviour of endogenous diPAPs and PFCAs present in

WWTP and paper fiber biosolids upon amendment of these materials with soil that has been

sown with Medicago truncatula plants. Biodegradation pathways and plant uptake of diPAPs

were further elucidated in a separate greenhouse microcosm supplemented with high

concentrations of the 6:2 diPAP congener. Biosolids-amended soil exhibited increased

concentrations of diPAPs (3.9–82.5 ng/g) and PFCAs (0.05–18.6 ng/g), as compared to control

soils (nd–1.4 ng/g) that did not receive biosolids amendment. A combination of sorption, plant

uptake, and biotransformaton contributed to the observed decline in diPAP soil concentrations

over time, the last of which was evidenced by the degradation of 6:2 diPAP to its corresponding

fluorotelomer intermediates and C4–C7 PFCAs. Substantial plant accumulation of endogenous

PFCAs present in the biosolids (0.1–138.4 ng/g) and those produced from 6:2 diPAP degradation

(0.1–58.3 µg/g) was observed within 1.5 month of biosolids application, with the congener

profile typically dominated by the short-chain PFCAs (C4–C6). This pattern was corroborated

by the inverse relationship observed between the plant-soil accumulation factor (PSAF,

Cplant/Csoil) and carbon chain length (p < 0.05, r = 0.90–0.97). Together, these results provide the

first evidence of soil biodegradation of diPAPs and their subsequent uptake, as well as their

metabolites into the plant environment.

4.2 Introduction

The high concentrations (ng/g) of perfluoroalkyl and polyfluoroalkyl substances (PFASs)

often reported in wastewater treatment plant (WWTP) sludge (1–5) and their demonstrated

capacity to sorb strongly to sludge (6) suggest this matrix may be a significant reservoir for these

137

chemicals in the environment. This is of concern because a significant fraction (40%) of treated

WWTP sludge or biosolids is directed towards agricultural land application in Ontario (ON),

Canada to increase soil fertility and supplement nutrients (7). Of the 130,000 tons of biosolids

generated at the largest WWTP facility in Toronto, ON in 2011, 37% was applied directly on

land, while another 37% was further processed via pelletization and chemical treatment for soil

amendment (8). Paper fiber biosolids are generated from the wastewater treatment process of

recycled paper products in pulp and paper mills and like WWTP biosolids, they are largely

applied (20%) as a soil conditioner on agricultural lands in Ontario (7). Analysis of WWTP

sludge and paper fibers collected across Ontario from 2002 to 2008 reported detection of varying

chain lengths of perfluorocarboxylates (PFCAs, 7–10 perfluorinated carbons (CFs)) and

perfluorooctanesulfonate (PFOS) at ng/g concentrations, and for the first time, significant

contamination of a commercial fluorinated product, the polyfluoroalkyl phosphate diesters

(diPAPs) (up to 860 ng/g and 2000 ng/g in WWTP sludge and paper fibers respectively) (5).

DiPAPs belong to a suite of commercial fluorotelomer-based materials, such as the

acrylate-based polymers (FTAcPs) (9, 10), sulfonates (FTSAs) (11), and stearate monoesters

(FTSs) (12), all of which have a demonstrated capacity to biodegrade to PFCAs in either soil

and/or WWTP-simulated environments (13). Detection of fluorotelomer saturated (FTCAs) and

unsaturated (FTUCAs) carboxylates in WWTP sludge and effluents (4, 14, 15), both of which

are metabolic intermediates of fluorotelomer-based precursor degradation to the PFCAs, also

suggest WWTPs are continuously exposed to contaminated influents containing diPAPs and

potentially other fluorotelomer-based materials. In addition to direct PFCA emission sources to

WWTPs, such as contaminated discharges from nearby fluorochemical industries (16) and

disposal of consumer products containing PFCAs (17–19), the degradation of commercial

fluorotelomer-based products in these facilities represents an additional source of PFCA

contamination in WWTP media. As such, the primary concerns of biosolids application onto

agricultural farmlands center over its potential as an exposure route of PFASs to soil and its

surrounding environment, as was observed in soil (20), tile drainage water (21), and assorted

plant crops (22) during laboratory and field experiments.

These very concerns were recently highlighted in agricultural farmlands in Decatur,

Alabama that have received >10 years of biosolids application from a local WWTP known to

have processed effluents from nearby fluorochemical manufacturers of PFASs. Some of the

138

highest soil concentrations of perfluoroalkyl acids (PFAAs) were reported in the biosolids-

amended soils (~2500 ng/g dry weight (dw) perfluorooctanoate (PFOA); ~1400 ng/g dw PFOS),

which were about 1–2 orders of magnitude higher than those measured in soils collected from

background fields that have never received biosolids application (23). Similarly, elevated PFAA

concentrations were observed in plants (10–200 ng/g dw PFOA; 1–20 ng/g dw PFOS) (24) and

surface and well water (up to 11000 ng/L in surface water; up to 6400 ng/L in well water) (25)

sampled in the vicinity of the impacted fields. The fact that varying chain lengths of

fluorotelomer alcohols (6:2 to 14:2 FTOHs) were also detected in the same plant samples (24)

and biosolids-amended soils (26) as above further supports potential biodegradation of

fluorotelomer-based materials that may be present in the soil through transfer from the WWTP

biosolids after application. A number of soil and WWTP-simulated biodegradation studies have

reported FTOH as a metabolite intermediate during the transformation of various fluorotelomer-

based precursors, such as the diPAPs (13), FTAcPs (9, 10), and FTSs (12). However, it has not

yet been demonstrated whether a similar transformation pathway may occur for these precursors

in a soil-plant environment.

Here, a greenhouse pot experiment was performed to investigate the fate of diPAPs in a

5.5-month soil-plant microcosm. Transformation metabolites of one diPAP congener (6:2) were

identified in biosolids-amended soil and the plant species, Medicago truncatula, sown in the

same pots. The influence of compound-dependent factors, such as the perfluoroalkyl chain

length and susceptibility to biodegradation, on the plant uptake of the parent diPAPs and their

corresponding metabolites was also examined. As diPAPs are marketed as greaseproofing

agents in food contact papers (27) and have been frequently found in European food packaging

material (28, 29), a separate greenhouse experiment was performed to investigate the potential

for endogenous diPAPs present in contaminated WWTP biosolids and paper fiber biosolids to

carry over to amended soils and subsequently, to plants grown on the same soils.

139


4.3.1 Materials

A list of all standards and reagents used in this study is provided in the Supporting

Information (SI) in Appendix B. All target analytes are listed in Table B1 in Appendix B. The

diPAPs (y = x) and 6:2 monoPAP used here were synthesized by methods described elsewhere

(30).

Dewatered biosolid material (30% solids) was collected from the North Toronto

Wastewater Treatment Plant (Toronto, Ontario (ON) in 2009. Paper fiber biosolids were

collected from an Ontario paper mill in 2008, and have been previously analyzed for diPAPs,

PFCAs, and PFSAs (5). Sandy loam soil was collected from an agricultural farmland

(Northumberland County, ON; 44o05’N, 78

o01’W) in 2009. Upon arrival at the laboratory, the

soil was sieved with a 2 mm stainless steel mesh and left to air-dry over several days. The soil

was analyzed by SGS AgriFood Laboratories (Guelph, ON) and selected characterization data

are as follows: pH 5.5; 1.8% organic matter; cation exchange capacity of 96 µmol/g; 49 mg/kg of

NaHCO3-extractable P; 63% sand, 32% silt, 5% clay. Alfafa plant seeds of the species, M.

truncatula, were obtained from the Western Regional Plant Introduction Station of the United

States Department of Agriculture Agricultural Research Service (USDA-ARS) (Pullman,

Washington).

4.3.2 Soil-Plant Microcosm Experiment

Using an OdjobTM

concrete mixer (Scepter Corporation, Toronto, ON), the WWTP

biosolids were mixed with soil at a rate of 16 g biosolids/kg of soil (≈ 8.7 metric dry tons/ha),

which was slightly higher than the maximal 5-year application rate of 8 tons/ha permitted in

Ontario (31). In a separate experiment, the same biosolids were mixed with paper fiber biosolids

at a ratio of 1:4 that corresponded to application rates of 16 g WWTP biosolids/kg of soil and 67

g paper fiber biosolids/kg of soil respectively. These biosolids-amended soils were then

transferred to pots (~600 g/pot) after which 5–10 manually scarified seeds of M. truncatula were

planted in each pot, followed by inoculation with a mixture of cultured rhizobia strains, known to

form symbiosis with M. truncatula in nature. Preparation of the rhizobia mixture is described in

140

Appendix B. As the pots contained holes at the bottom, a catch plate was placed under each pot

to capture any analytes that may have leached from the soil during watering.

The pots were grouped into four treatments, as shown in Figure B1 in Appendix B: (1)

soil without biosolids amendment (n = 1 per timepoint); (2) WWTP biosolids-amended soil sown

with plant seeds (n = 3 per timepoint); (3) soil amended with 1:4 mixture of WWTP biosolids

and paper fiber biosolids and sown with plant seeds (n = 3 per timepoint); and (4) WWTP

biosolids-amended soil sown with plant seeds and mixed with 100 mg of 6:2 diPAP from an

ethanol-based standard (n = 3 per timepoint). Treatment 1 served as the PFAS-free blank and

plant-free control, while treatments 2 and 3 were included to investigate the fate of endogenous

PFASs that may be present in the WWTP biosolids and paper fiber biosolids. In treatment 4, 6:2

diPAP was added as the parent reactant to monitor for its potential biodegradation and uptake

into the plants. Commercial greaseproofing formulations, containing fluorinated phosphate

surfactants like the diPAPs, are typically composed of a mixture of varying perfluoroalkyl chain

lengths (4–20 CF’s), as well as, the monofluoroalkylated (monoPAP) and trifluoroalkylated

(triPAP) phosphate esters (27, 32, 33). The use of 6:2 diPAP as the parent reactant here was

based on industry preference for the perfluorohexyl chain length in the manufacture of

fluorinated surfactants (34), and the fact that the diester typically exhibits the highest product

efficiency in oil repellency applications, as compared to the mono- and triesters (32). The pots

were watered daily and kept in a greenhouse (Earth Sciences Centre, University of Toronto, ON)

for 5.5 months under natural sunlight and supplementary illumination (200 µmol/m2/sec) and a

temperature regime of 25/21oC day/night.

4.3.3 Sampling, Extraction, and Analysis

Prior to plant growth, initial concentrations of diPAPs and their expected metabolites in

the soil were measured by sacrificing one pot (n = 1) from each of the four treatment groups. At

subsequent timepoints of 1.5, 3.5, and 5.5 months, triplicate pots (n = 3) were sacrificed for each

treatment, except for the blank soil, which was sampled as 1 pot at each timepoint. During each

sampling, the entire plant, including the roots, was harvested, shaken gently to remove any

adhering soil particles, and archived together in plastic bags. The soil was wholly removed from

each pot and mixed with 100–300 mg of sodium azide (NaN3) in a plastic bag to inhibit further

microbial activity. The plates placed under each pot were also archived in plastic bags. All

141

samples were stored at 4oC until analysis. Soil, plants, and catch plates were extracted by

methods described in Appendix B.

Sample analysis was performed with high pressure liquid chromatography-tandem mass

spectrometry (HPLC-MS/MS), using an Agilent 1100 HPLC coupled to an API4000 triple

quadrupole MS (Applied Biosystems/MDS Sciex) operating under negative electrospray

ionization mode. Chromatographic separation was performed using a GeminiNX C18 column

(4.6 x 50 mm, 3 µm; Phenomenex, Torrance, CA). Further instrumental parameters are provided

in Appendix B.

4.3.4 Quality Assurance of Data

Quantitation of the PFCAs, FTCAs and FTUCAs was performed using mass-labeled

internal standards, with the exception of those analytes, for which their corresponding mass-

labeled internal standards were not available at the time of the experiment, and thus were

quantified using internal standards of structurally similar analytes as surrogate standards (Table

B2 in Appendix B). As analytical standards for the 3:3, 5:3, 9:3 FTCAs and FTUCAs, and 7:3

FTUCAs were not commercially available at the time of analysis, these analytes were detected

and quantified using inferred mass transitions and native standards of the adjacent FTCAs and

FTUCAs as surrogate standards respectively (Table B2 in Appendix B). Due to the lack of

commercially available mass-labeled internal standards at the time of analysis, diPAPs were

quantified using matrix-matched calibration where control soil and plant served as the matrix.

Further details on preparation of the matrix-matched standards are described in Appendix B. As

no standards were synthesized for the mixed diPAPs (y = x + 2), each y = x + 2 diPAP was

quantified by a pseudo matrix-matched calibration curve that was created by averaging the

calibrations for the corresponding adjacent y = x diPAPs, as was performed previously (5).

Spike and recovery experiments were performed in triplicate (n = 3) in control soil and

plants and clean catch plates. All concentrations determined here were not corrected for

recovery. Details on the spike and recovery procedures and the recovery ranges (Table B2) are

provided in Appendix B.

The limits of detection (LODs) and limits of quantitation (LOQs) were defined as the

concentrations producing a signal-to-noise ratio of equal to or greater than 3 and 10 respectively.

142

The matrix-specific method LODs and LOQs for each analyte are listed in Table B3 in Appendix

B. All concentrations below the LOD were assigned a value of zero and values above the LOD,

but below the LOQ were used unaltered. All concentrations were reported as arithmetic means

with standard error.

Replicate procedural blanks (HPLC grade water, n = 5) were included in the extraction of

each timepoint. No contamination of analytes was observed above their corresponding LOQs in

the procedural blanks.

4.3.5 Data Analysis

The plant growth rates were calculated by fitting all plant mass data from Treatments 2–4

to an exponential model: ln(mass, g) = b·t + a; where b is the plant growth rate (/month), t is the

elapsed time (month), and a is a constant. All plant concentrations were corrected for growth

dilution by using the plant growth rates calculated for each Treatment, shown in Table B4 in

Appendix B.

The rate constant and half-life of 6:2 diPAP dissipation in the soil were calculated by

fitting the soil concentration data to the first-order decay model: ln(Csoil) = kd·t + a; where Csoil is

the soil 6:2 diPAP concentration, kd is the disappearance rate constant (/month), t is the time

(month), and a is a constant (StatsDirect, Version 2.7.8, 2010). The disappearance half-life (t1/2)

was calculated as ln(2)/kd.

All statistical analyses were performed using StatsDirect (Version 2.7.8, 2010). An α-

value of 0.05 was chosen as the criterion of statistical significance in all analyses.


4.4.1 Amendment of WWTP Biosolids and Paper Fiber Biosolids as a Source of PFASs to

Soil

Prior to biosolids amendment, background PFAS contamination was determined for the

control soil used in Treatment 1. The C6–C11 PFCAs were present at concentrations ranging

from 0.02 to 1.44 ng/g, with PFOA and the longer chain PFCAs as the more dominant congeners

in the soil. These concentrations are similar in range to those previously reported in various

sandy loam soils collected in Georgia, US (35). None of the polyfluoroalkyl PFCA precursors

143

(i.e. diPAPs, FTCAs, and FTUCAs) were detected in the control soil. Upon amendment of the

soil with WWTP biosolids (Treatment 2) and paper fiber biosolids (Treatment 3), significant

diPAP (up to 82 ng/g) and PFCA (up to 19 ng/g) concentrations were observed in the soil

(Figure 4.1, Figures B2 and B3 in Appendix B). As was previously observed in sludge sampled

across various Ontario WWTPs (5), a suite of diPAP congeners (6:2/8:2 to 10:2/12:2) were

detected in the WWTP biosolids-amended soil here at concentrations ranging from 3.9 ± 0.8 ng/g

for 6:2/8:2 diPAP to 51.1 ± 5.7 ng/g for 10:2 diPAP (Figure 4.1, Figure B2 in Appendix B).

Similar diPAP congeners (6:2 to 10:2/12:2) were also observed at concentrations ranging from

23.7 ± 4.5 ng/g for 6:2 diPAP to 82.5 ± 10.0 ng/g for 10:2 diPAP in soil amended at a 1:4 ratio of

WWTP biosolids and paper fiber solids respectively (Figure 4.1, Figure B3 in Appendix B).

Together with the significant diPAP contamination (up to 2600 ng/g) previously reported in

these same paper fiber solids (5), the high diPAP concentrations observed here in the paper fiber

biosolids-amended soil are consistent with the prevalent use of these chemicals in food contact

paper applications (27).

An increasing prevalence of the longer chain diPAPs was observed in both types of

treated soil, which is consistent with the sorption dependency on chain lengths that has been

previously reported for PFCAs and PFSAs in sediments (36) and soils (37). The observation of

different perfluoroalkyl chain lengths of diPAPs in the amended soil here and previously in

WWTP sludge and paper fiber biosolids (5) is also consistent with environmental exposure to

commercial fluorotelomer-based products.

A decline was observed in the concentrations of diPAPs in both types of treated soil over

time (Figure 4.1, Figures B2 and B3 in Appendix B), which may be due to a number of

pathways, such as sorption to the pots, soil, and/or biosolids, leaching to the catch plates during

watering, biodegradation, and translocation into plants. Accumulation of 104 ± 13 ng and 107 ±

20 ng of total diPAPs (ΣdiPAPs) were observed over time in the catch plates placed under the

WWTP biosolids-amended pots and WWTP- and paper fiber biosolids-amended pots

respectively. These masses corresponded to <0.5% losses (on a mole basis) of ΣdiPAPs present

in the soil at the end of the experiment, which suggest leaching may be a minor loss pathway

(Figure B4 in Appendix B).

144

Figure 4.1. Concentrations of diPAPs and PFCAs (ng/g) observed in control soil, WWTP

biosolids-amended soil, and WWTP biosolids- and paper fiber biosolids-amended soil at 0, 3.5,

and 5.5 months. Each data point represents the arithmetic mean concentration of the triplicate (n

= 3) sampling. The error bar represents the standard error.

Uptake of diPAPs at concentrations up to 30 ng/g was initially observed in plants

sampled from both types of biosolids-amended soil at 1.5 month, but subsequent analysis of

plants sampled at 3.5 and 5.5 months revealed either no detection or a decline in diPAP

concentrations (Figures B2 and B3 in Appendix B). Under WWTP-simulated conditions,

diPAPs have been shown to undergo microbially-mediated biodegradation to yield FTOHs of

Co

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100

1000

Soil without Biosolids Amendment

WWTP Biosolids-Amended Soil

WWTP and Paper Fiber Biosolids-Amended Soil

0.01

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6:2

diP

AP

6:2

/8:2

diP

AP

8:2

diP

AP

8:2

/10:2

diP

AP

10

:2 d

iPA

P

10

:2/1

2:2

diP

AP

0.01

0.1

1

10

100

1000

PF

BA

PF

Pe

A

PF

Hx

A

PF

Hp

A

PF

OA

PF

NA

PF

DA

PF

Un

A

PF

Do

A

PF

TrA

PF

Te

A

0.01

0.1

1

10

100

1000

0 Month

3.5 Month

5.5 Month

0 Month

3.5 Month

5.5 Month

145

corresponding chainlengths, followed by further oxidation of the FTOH intermediates and/or

continued biotransformation of the diPAPs themselves to the final PFCA products (13). The fact

that PFCA concentrations in both types of biosolids-amended soil were typically higher than

those measured in the control soil (e.g. 4.4–21.8 ng/g of PFOA in biosolids-amended soil vs. 1.4

ng/g of PFOA in control soil, Figure 4.1) suggests WWTP and paper fiber biosolids may be

significant sources. This is consistent with the high concentrations of endogenous PFCAs often

observed in North American WWTP sludge (1–5) and paper fiber biosolids (5), but the presence

of known PFCA precursors, such as FTSAs (2) and diPAPs (5), in WWTP media also implicates

commercial fluorotelomer-based materials as potential contributors to the observed

contamination. In addition, a distinct pattern was observed in the congener profile of the PFCAs

detected in both types of biosolids-amended soils in which an even-carbon chain length PFCA

(e.g. PFOA C8, perfluorodecanoate (PFDA, C10), or perfluorododecanoate (PFDoA, C12))

occurred at higher concentrations than the adjacent odd-carbon chain length PFCA (e.g.

perfluorononanoate (PFNA, C9), perfluoroundecanoate (PFUnA, C11), or perfluorotridecanoate

(PFTrA, C13) (Figure 4.1, Figures B2 and B3 in Appendix B). This even > odd carbon chain

pattern is consistent with the biological production of PFCAs from fluorotelomer-based materials

(38–41).

Despite the observed decline in diPAP concentrations in the soil and plant samples over

time, no consistent evolution of PFCAs was observed in either of these compartments. However,

the occasional detection of various FTCAs and FTUCAs in both the plants and soil is evidence

of biotransformation of some fluorotelomer-based precursor materials present in the system.

Identifying these specific precursors is complicated by the diverse functionalities incorporated in

the manufacture of commercial fluorotelomer-based chemicals (e.g. phosphates, sulfonates,

ethoxylates, polymers) (42) and the fact that these products may contain mixtures of different

chain lengths and other fluorotelomer-based residuals, like FTOHs (43). In the interest of

elucidating transformation kinetics, plant uptake, and metabolite profiles from potential soil

and/or plant degradation of a commercial fluorotelomer-based product, 6:2 diPAP was chosen as

the model parent reactant and added at high concentrations (mg/kg) to a soil-plant microcosm to

monitor its environmental fate in a simulated soil-plant microcosm.

146

4.4.2 Metabolism of 6:2 diPAP in the Soil-Plant Microcosm

The decline in 6:2 diPAP soil concentrations followed first order kinetics with a

calculated disappearance half-life of ~2 months (kdisappearance = 0.342 ± 0.002 /month; r = 1.00; p

< 0.0001) (Figure 4.2). As described above, the dissipation of diPAPs in soil may occur through

multiple pathways, such as leaching to the catch plates, sorption, uptake into the plants, and

biodegradation. As was observed above, leaching was a minor loss pathway for 6:2 diPAP as

only 712 ± 252 ng of 6:2 diPAP was observed to accumulate over time in the catch plates, which

corresponded to <0.1% (by moles) of the total 6:2 diPAP measured at 5.5 months in the catch

plate, soil, and plant compartments, while the majority of the 6:2 diPAP resided almost entirely

in the soil (99%), with minor uptake (1%) observed in the plants (Figure B4 in Appendix B).

Due to the difficulty of maintaining microbial sterility inside a greenhouse for 5.5 months, no

sterile controls were included here, which precluded assessing how much of the observed loss of

6:2 diPAP was due to sorption in the absence of soil or plant microbes capable of degrading the

chemical.

As was demonstrated in previous WWTP-simulated biodegradation experiments, the

main metabolic pathway for 6:2 diPAP is first microbially-mediated hydrolysis of the phosphate

ester bond to produce 6:2 FTOH, followed by further transformation of either the FTOH or the

diPAP itself to the final corresponding PFCAs (13). Analysis of FTOHs was not performed in

the microcosm here as the pots were open to the atmosphere of the greenhouse, which precluded

sampling of any volatile metabolites of diPAPs that may be offgassing from the plants and/or

soil. Given not all of the metabolites could be accounted for and the fact that any endogenous

diPAPs and/or other PFCA precursors present in the applied WWTP biosolids may additionally

contribute to the metabolites observed here, mass balance calculations were not performed in the

biotransformation of 6:2 diPAP here.

147

Figure 4.2. Concentrations of 6:2 diPAP, 6:2 and 5:3 FTCAs and FTUCAs, C4–C7 PFCAs

(ng/g) observed in soil and plants from 6:2 diPAP-supplemented microcosm at 0, 1.5, 3.5, and

5.5 months. Each data point represents the arithmetic mean concentration of the triplicate (n = 3)

sampling. The error bar represents the standard error.

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200

400

600

800

1000

1200

6:2 FTCA

6:2 FTUCA

5:3 FTCA

5:3 FTUCA

PFBA

PFPeA

PFHxA

PFHpA

Time (Months)

0 1 2 3 4 5 6

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6:2 FTCA

6:2 FTUCA

5:3 FTCA

5:3 FTUCA

Time (Month)

0 1 2 3 4 5 6

0

20000

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60000

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PFBA

PFPeA

PFHxA

PFHpA

6:2 diPAP

6:2 FTCA

6:2 FTUCA

5:3 FTCA

5:3 FTUCA

PFBA

PFPeA

PFHxA

PFHpA

6:2 diPAP

6:2 FTCA

6:2 FTUCA

5:3 FTCA

5:3 FTUCA

PFBA

PFPeA

PFHxA

PFHpA

Soil Plant

148

The observed production of 6:2 FTCA, 6:2 FTUCA, 5:3 FTCA, 5:3 FTUCA,

perfluorobutanoate (PFBA, C4), perfluoropentanoate (PFPeA, C5), and perfluorohexanoate

(PFHxA, C6) from 6:2 diPAP biodegradation in the soil-plant microcosm (Figure 4.2) is

consistent with the metabolite profiles, previously reported for 6:2 diPAP and 6:2 FTSA in

WWTP-simulated systems (11, 13), and 6:2 FTOH in aerobic soils (44). Consistent with the

beta-oxidation-like transformation of 8:2 FTCA to PFOA first observed in microbial (38) and

mammalian (45) degradation of 8:2 FTOH, 6:2 FTCA was the first intermediate detected in the

soil, then consumed, followed by formation of 6:2 FTUCA and PFHxA (Figure 4.2).

The consumption of first 6:2 FTCA and then 6:2 FTUCA coincided with formation of 5:3

FTCA as one of the major metabolites here and to a lesser extent, 5:3 FTUCA. Separate in vitro

and in vivo incubations of 8:2 FTCA and 8:2 FTUCA with mammalian hepatocytes and

microsomes (40) and in rainbow trout (41) have also produced 7:3 FTCA and 7:3 FTUCA,

which themselves have been shown to transform into one another (40). Here, the production of

5:3 FTCA was concurrent with that of PFPeA, which alludes to the demonstrated capacity of the

analog 7:3 FTCA to biotransform to PFHpA in rainbow trout (41). The mechanism by which

this pathway occurs was recently investigated in biodegradation experiments of 5:3 and 7:3

FTCAs as the parent reactants in WWTP activated sludge, in which 5:3 FTCA was observed to

undergo a series of dealkylation and defluorination steps to yield PFBA and PFPeA, while 7:3

FTCA appeared generally recalcitrant, with very low levels of PFHpA produced (46). In

contrast, the lack of detection of any metabolites during incubation of 5:3 and 7:3 FTCAs in

aerobic soils suggests their biodegradation was suppressed due to decreased bioavailabilty via

sorption to soil (44, 47). Instead, Liu et al. observed an alternative pathway in soil in which the

intermediate, 6:2 FTUCA, formed from 6:2 FTOH, was first transformed to 5:2 fluorotelomer

ketone, which itself degraded to 5:2 sFTOH, followed by consumption of this intermediate to

yield PFPeA and PFHxA as the dominant metabolites in that study (44). The initial step of this

pathway (x:2 FTUCA x–1:2 fluorotelomer ketone) has been corroborated by other studies

(40, 48), but no other work that further explores the mechanism of the x–1:2 sFTOH to x–1

PFCA pathway has been published.

The observation of PFBA as a metabolite here agrees with a number of studies that have

previously reported the removal of multiple –CF2- groups during biotransformation of

fluorotelomer-based substrates to yield PFCAs with two fewer CFs in their perfluorocarbon tails

149

(e.g. 6:2 FTOH PFBA (44); 8:2 FTOH PFHxA (40, 47)). The FTOH intermediates, 5:3

FTCA and 7:3 FTUCA, are both purported precursors to the corresponding PFBA and PFHxA

respectively (46, 47), but the mechanisms behind these pathways are currently not well

understood. Minor production of perfluoroheptanoate (PFHpA, C7) observed here in the soil is

also evidence of alpha-oxidation, which corroborates the pathway previously reported in

microbial and mammalian studies of 6:2 diPAP (13) and 8:2 fluorotelomer-based substrates (39,

40) transforming to the corresponding PFHpA and PFNA respectively, but contrasts with others

that did not observe this mechanism (11, 38, 44). As demonstrated above, there is considerable

variability in the multiple pathways associated with biotransformation of fluorotelomer-based

chemicals and a more comprehensive account of these pathways may be found elsewhere (40,

41, 44) and in Chapter 1.

4.4.3 Uptake and Accumulation of PFCA Metabolites in Plants

Plants sampled 1.5 month after WWTP biosolids (Treatment 2) and paper fiber biosolids

(Treatment 3) application to the soil exhibited PFCA concentrations ranging from 0.06 ± 0.03

ng/g for PFUnA to 138 ± 64 ng/g for PFBA, with no detection of PFTrA and PFTeA (Figures B2

and B3 in Appendix B). Possible sources of this contamination include uptake of endogenous

PFCAs already present in the applied biosolids and/or PFCA products formed from the soil or

plant metabolism of any fluorotelomer-based materials, such as the diPAPs observed here

(Figure 4.1, Figures B2 and B3 in Appendix B). The main uptake process of PFCAs into plants

is likely via transpiration of contaminated soil water through vegetative transport tissues, like the

xylem, from the roots to other plant compartments. This pathway is in part supported by two

laboratory studies investigating uptake of artificially spiked PFOS and PFOA and contaminated

biosolids from soil into various plant crops, both of which observed higher accumulation in the

vegetative compartments (i.e. leaves and stalks) than in the storage compartments inside the

edible portions of the plants (i.e. fruits, tubers, and grain) (22, 49). Alternative uptake may occur

by deposition of volatile PFCA precursors, such as FTOHs being produced and offgassing from

the soil, onto above-ground plant compartments inside which the FTOHs may further metabolize

to form PFCAs.

150

The observed decreases in the plant concentrations of 6:2 diPAP, 6:2 FTCA, 6:2 FTUCA,

and 5:3 FTCA coincided with the detection of PFBA, PFPeA, PFHxA, and PFHpA at

concentrations ranging from 0.025 ± 0.001 µg/g for PFHpA to 63.23 ± 9.04 µg/g for PFBA in

plants sampled from the 6:2 diPAP-spiked pots (Treatment 4) throughout the experiment (Figure

4.2). The occurrence of these analytes in the plants may be due to metabolism of plant-bound

6:2 diPAP and/or continuous uptake of PFCA metabolites from biodegradation of the 6:2 diPAP

initially spiked to the soil, as supported by the observed decline in PFCA soil concentrations at

5.5 months (Figure 4.2). It is unclear which of these pathways predominate in the microcosm

here, but it is likely both contributed to the PFCA contamination observed in the plants.

Plant-soil accumulation factors (PSAFs) were calculated for all three treatments (2–4),

based on the ratio of PFCA concentrations measured in plants to those measured in soil, and

plotted against carbon chain length in Figure 4.3. Plant uptake of the PFCAs observed here

demonstrated a chain-length dependency similar to that previously reported by Yoo et al. (24), in

which PSAFs decreased with carbon chain length (p < 0.05, r = 0.90–0.97) (Figure 4.3). This is

consistent with the predominant distribution of PFBA, PFPeA, and PFHxA observed in plants

sampled in all three treatments, as compared to the longer chain PFCAs (>C7) which prefer to

reside in the soil compartment, as shown in Figure B4 in Appendix B. The PSAFs calculated

here from plants sown in WWTP biosolids-amended soil (1.46 ± 0.49; PFOA) were typically

higher than those measured from carrots, potatoes, and cucumbers exposed to biosolids-amended

soil in the laboratory (0.01–0.05 from edible portions; 0.38–0.99 from leaves and stalks; PFOA)

(22) and grasses collected from the contaminated fields in Decatur, AL (0.09–0.65; PFOA) (24).

These differences may be due to variable uptake abilities across different plant species and may

also depend on the extent of the local fluorochemical contamination to which the plants are

exposed.

151

Figure 4.3. Correlation between the plant-soil accumulation factors (PSAFs, Cplant/Csoil) and

carbon chain length of the PFCAs analyzed in Treatments 2–4. Each data point represents the

arithmetic mean PSAF from averaging through individual PSAF measured at each timepoint

(1.5, 3.5, and 5.5 months). The error bar represents the standard error.

4.5 Environmental Implications

This study provides the first evidence of biotransformation of diPAPs and their

subsequent plant uptake, as well as, their metabolites in a soil-plant environment. This has

important implications for diPAPs and other commercial fluorotelomer-based chemicals that

have been discovered in the WWTP environment (2, 5), as land application of contaminated

3 4 5 6 7 8 9 10 11 12 13 14 15

Pla

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So

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F

ac

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AF

)

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50

100

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5000

10000

15000

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Cplant

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Cplant

/CWWTP- and Paper Fiber Biosolids-Amended Soil

3 4 5 6 7 8 9 10 11 12 13 14 15

log

(Pla

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Carbon Chain Length

3 4 5 6 7 8 9

Pla

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Cplant

/C6:2 diPAP-spiked in WWTP Biosolids-Amended Soil

3 4 5 6 7 8 9

log

(Pla

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Fa

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SA

F)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

152

biosolids may represent a significant source of these commercial materials and their metabolites

to soils.

Analysis of soil amended with both WWTP biosolids and paper fiber biosolids revealed

significant diPAP contamination (110–256 ng/g of ΣdiPAPs), while total PFCA concentrations

were as high as 32 ng/g and 561 ng/g in the soil and plants respectively. Translocation of PFCAs

into the plants was observed to favour the short-chain congeners, such as PFBA, PFPeA, and

PFHxA, although longer PFCAs (C7–C12) were also detected. Published data for PFASs in

edible plants are sparse, but the accumulation observed here and by others (22, 24, 49) suggest

plant uptake may be an important entryway for PFASs into the food chain through human

consumption of crops grown on contaminated fields. The discovery of elevated PFAS

concentrations in the biosolids-applied fields in Decatur, AL (23–26) has also triggered concern

over potential contamination of beef cattles that have been grazing on these fields for 12 years

(50). Analytical data on these animals have not been published, except for one raw milk sample,

obtained from a bulk tank supplied by these cattles, which reported a detectable concentration of

PFOS at 0.16 ng/g, but no other PFAAs monitored (51). Nevertheless, recent work by the

German Federal Institute of Risk Assessment (BfR) demonstrated significant accumulation of

perfluorosulfonates (PFSAs) and PFCAs of varying chain lengths in cows and pigs that have

been fed contaminated plant crops (0.3–1923 ng/g dw), which themselves were harvested from

PFAA-contaminated farmlands (52). These results and recent evidence of biomagnification of

PFAAs in the terrestrial lichen-caribou-wolf food chain (53) together suggest consumption of

contaminated herbivores may represent an additional route of human exposure to PFASs.

Assessing the magnitude of the soil-plant uptake pathway of PFASs and how that may

vary across different plant species and compartments is important when considering the risks of

animal and human exposure from consumption of only edible plant species and compartments.

Food-borne exposure to fluorinated chemicals has primarily focused on analyzing processed

food items (54, 55) and their packaging (28, 29, 56), but analysis of unrefined foods, such as raw

meat and produce, may better characterize the immediate impact of certain agricultural practices,

such as biosolids application, on the local environment.

153


We would like to thank Wellington Laboratories (Guelph, ON) for donating native and

mass-labeled internal standards, Pablo Tseng (University of Toronto, ON) and North Toronto

WWTP (Toronto, ON) for their assistance in collecting biosolids for this study, Katy Heath

(University of Toronto, ON) for assistance in provision of plant seeds and preparation of rhizobia

cultures, and Bruce Hall and Andrew Petrie (University of Toronto, ON) for assistance in

greenhouse set-up. The present study is funded by Natural Science and Engineering Research

Council of Canada (NSERC) and a NSERC Postgraduate Scholarship awarded to HL.




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158

CHAPTER FIVE

Sorption of Perfluoroalkyl Phosphonates and Perfluoroalkyl Phosphinates in Soil


In preparation: For submission to Environmental Science and Technology.

Contributions: Holly Lee was responsible for conceiving the experimental design,

performing all sorption experiments, method development, sample acquisition, and data

interpretation. Holly Lee prepared this manuscript with editorial comments provided by

Scott Mabury

159

5.1 Abstract

Perfluoroalkyl phosphonates (PFPAs) and perfluoroalkyl phosphinates (PFPiAs) are

newly discovered perfluoroalkyl acids (PFAAs) that have been recently detected in lake trout,

surface water and wastewater environments. Their presence in such varying matrices suggests

the environmental partitioning of PFPAs and PFPiAs is simultaneously governed by different

mechanisms, such as sorption and bioaccumulation, the latter of which has been recently

investigated in rainbow trout. The sorption of C6, C8, and C10 monoalkylated PFPAs and

C6/C6, C6/C8, and C8/C8 dialkylated PFPiAs was investigated in seven Canadian and American

soils of varying geochemical parameters. As have been previously observed for the

perfluoroalkyl carboxylates (PFCAs) and perfluoroalkane sulfonates (PFSAs) in sediments, the

sorption observed here was dependent on both perfluorocarbon chain length and the polar

headgroup. The organic carbon-normalized distribution coefficients, logKOC, ranged from 1.8 to

3.6 for the PFPAs and PFPiAs and were observed to increase with the number of perfluorinated

carbons present in the chemical. The logKOC of PFSAs (3.0–3.7), previously measured from

sediments, were similar in range to those calculated for the PFPiAs here (3.2–3.6), and greater

than those for the PFCAs (2.2–3.5) and PFPAs (1.8–3.1) of equal perfluorocarbon chain length.

No single soil-specific parameter, such as pH and organic carbon content, was observed to

significantly control the sorption of PFPAs and PFPiAs, the lack of which may be attributed to

competing interferences among different sorption-dependent parameters as they vary from soil to

soil. The PFPAs were observed to desorb to a greater extent, as compared to the PFPiAs, and

thus, are expected to primarily circulate as aqueous contaminants in the environment, while the

more bioaccumulative and sorptive PFPiAs would preferentially partition into biological

organisms and/or sorb to environmental solid phases.

5.2 Introduction

The pathway of urban discharges of perfluoroalkyl and polyfluoroalkyl substances

(PFASs) into wastewater treatment plants (WWTPs) and their receiving water bodies has been

well documented in wastewater, surface water, and sediment samples collected downstream from

these facilities (1–7). Analysis of WWTP sludge has consistently reported ng/g concentrations

of perfluoroalkyl carboxylates (PFCAs) and perfluoroalkane sulfonates (PFSAs), with

160

perfluorooctanoate (PFOA, C8 PFCA) and perfluorooctane sulfonate (PFOS, C8 PFSA) typically

observed as the dominant perfluoroalkyl acids (PFAAs), followed by longer chain PFCAs and

PFSAs (>8 perfluorinated carbons, CFs) (7–15). This is consistent with the demonstrated

capacity of PFAAs to sorb to environmental solid matrices, such as sediments (16–19), soils

(20), and sludge (21). The fact that PFAAs may sorb to these environmental solids has

implications for the potential retention and release of these chemicals to the aqueous

environment.

Laboratory batch experiments using freshwater sediments (16) and topsoils (20) indicated

the sorption of PFAAs exhibits a chain-length dependency in which their organic carbon-

normalized distribution coefficients (KOC) increase with the number of CFs present in the

perfluorocarbon tail of the PFAAs studied. In these experiments, PFSAs were also observed to

be more sorptive than PFCAs of equal perfluorocarbon chain length. These observations mirror

the distribution of PFAAs typically observed in environmental samples, such as those examined

by Ahrens et al. (22, 23), in which short chain PFCAs (≤7 CFs) were only detected in the

seawater and porewater collected from Tokyo Bay, Japan, while the long chain PFCAs (≥8 CFs)

and PFSAs (≥6 CFs) were only observed in the suspended particulate matter and sediment

samples. The influence of headgroup on the partitioning behaviour of PFAAs in the

environment was recently demonstrated by the different biomagnification factors (BMFs)

measured for the PFSAs, PFCAs, and perfluoroalkyl phosphonates (PFPAs) of equal

perfluorocarbon chain length in juvenile rainbow trout (24, 25).

PFPAs and perfluoroalkyl phosphinates (PFPiAs) constitute a new class of PFAAs, with

their perfluorocarbon tails attached through a carbon-phosphorus (C–P) bond to either a

phosphonate (Rx-P(O)O2-; Cx PFPA) or phosphinate (Rx-P(Ry)(O)O

-; Cx/Cy PFPiA) headgroup

(Table 5.1).

161

Table 5.1. Structures, full names, and acronyms of the target analytes monitored.

Structure Full Name Acronym

P

O

-OF

F F

O-x

Perfluorophosphonate

Cx PFPA

x = 6, 8, 10

3 congeners monitored

P

O

-O

F

F F

x F

FFy

Perfluorophosphinate

Cx/Cy PFPiA

x/y = 6/6; 6/8; 8/8


PFPAs and PFPiAs are currently marketed as leveling and wetting agents in household

cleaning products (26) and were historically incorporated as inert ingredients in United States

(US) pesticide formulations (27) until 2008 (28). Despite widespread observations of the C6,

C8, and C10 PFPAs in Canadian surface waters at pg/L concentrations (29), these chemicals

were not detected in any lake trout sampled from the Great Lakes , whereas the C6/C6 and

C6/C8 PFPiAs were observed at concentrations up to 32 pg/g in the same samples (30). This

difference is consistent with faster depuration kinetics previously observed for the C6, C8, and

C10 PFPAs in rainbow trout (4–5 days) (25) and rats (1–3 days) (31) than for the C6/C6, C6/C8,

and C8/C8 PFPiAs (6–53 days in rainbow trout; 2–4 days in rats) (25, 31). The C6/C6 and

C6/C8 PFPiAs have been observed at 2–3 ng/g in WWTP sludge (31), whereas analysis of

various Dutch sludge and sediment samples did not reveal any detection of the monitored C6,

C8, and C10 PFPAs (32). Together, these observations suggest the smaller molecular weight

(MW) PFPAs may be more water soluble than the higher MW PFPiAs, whereas, the PFPiAs

may preferentially partition to solid matrices.

The present study aimed to investigate the sorption behaviour of three PFPA (C6, C8, and

C10) and three PFPiA (C6/C6, C6/C8, and C8/C8) congeners in seven soils of varying

geochemical properties. The influence of structural features, such as the perfluorocarbon chain

length and headgroup, on potential hydrophobic and electrostatic interactions between the PFPAs

and PPFiAs and the soils studied was investigated by determining and comparing distribution

162

coefficients for the six congeners to those previously reported for the PFSAs and PFCAs (16,

20). The relationship between soil- and aqueous phase-specific parameters, such as pH and

organic carbon, and sorption was also examined. A number of laboratory and field studies have

shown considerable evidence of PFAAs leaching from soils, that have been artificially spiked

with PFAAs (33) or exposed to contaminated media, such as street runoffs (34) and WWTP

sludge (33, 35–37), to groundwater and surface water. As such, desorption experiments were

also performed with one soil type to determine which of the studied PFPAs and PFPiAs may be

prone to remobilization into the aqueous phase and which to desorption hysteresis.


5.3.1 Chemicals

Neat material (~1 mg for each congener) of C6 PFPA (>98%), C8 PFPA (>98%), C10

PFPA (>98%), C6/C6 PFPiA (>98%), C6/C8 PFPiA (>98%), and C8/C8 PFPiA (>98%) were

donated by Wellington Laboratories (Guelph, ON). However, the amount of chemical required

to perform all sorption experiments here precluded the use of these analytical standards. Instead,

the technical product, Masurf®-780 (Mason Chemical Company, Arlington, IL), was used for

spiking in the majority of these experiments. Masurf®

-780 is a technical product composed of a

mixture of PFPAs and PFPiAs. Using the analytical standards of C6, C8, and C10 PFPAs

(>99%) and C6/C6, C6/C8, and C8/C8 PFPiAs (>99%) from Wellington Laboratories, the

percent composition of this commercial material was determined by standard addition to be

6.9±0.4% C6 PFPA, 5.8±0.4% C8 PFPA, 3.2±0.5% C10 PFPA, 4.3±1.0% C6/C6 PFPiA,

4.7±0.9% C6/C8 PFPiA, and 0.6±0.1% C8/C8 PFPiA. This percent composition was used to

adjust the PFPA and PFPiA concentrations reported here. Details on how this percent

composition compared to that previously reported by D’eon and Mabury (31) and Lee and

Mabury (38) are provided in the Supporting Information (SI) in Appendix C, as well as a list of

all other chemicals used in this study.

5.3.2 Soils Used

Four soils (A–D) were collected from various locations in Southern Ontario, Canada,

while one soil (E) was collected in Athens, Georgia, US. Two reference soils, the Elliott silt

163

loam (F) and Florida Pahokee peat (G), were obtained, air-dried and pre-sieved, from the

International Humic Substances Society (Golden, Colorado), and used as received. All other

soils were sieved with a 2 mm stainless steel mesh prior to oven drying at 105oC and

homogenized finely using a mortar and pestle. Characterization data for each soil are provided

in Table C1 in Appendix C.

5.3.3 Batch Sorption Experiments

All sorption experiments were performed in compliance with the Organization for

Economic Cooperation and Development (OECD) guidelines for studying sorption and

desorption behaviour of a chemical in different soil types (39). Prior to all sorption experiments,

the soil was pre-equilibrated with 0.10 mM mercuric chloride (HgCl2) and 0.01 M calcium

chloride (CaCl2) by shaking overnight at 200 rpm.

A preliminary study using Soil A was performed to determine appropriate soil to solution

ratio at which >50% by mass of the chemical has sorbed to the soil and equilibration time for

sorption to occur. Three sets of 50 mL polypropylene tubes in triplicate (n = 3), containing 2, 5,

and 10 g of Soil A and 50 mL of 0.01 M calcium chloride (CaCl2) solution, were spiked with 10

μg/mL of Masurf and left to shake at 200 rpm. Polypropylene tubes were removed at 0.5, 2, 8,

24, 48, 72, and 144 hours and the soil and aqueous phase were separated and analyzed, as

described in the following section. Except for C6 PFPA, <50% of all other PFPA and PFPiA

congeners were observed to remain in the aqueous phase within the first sampling timepoint for

all three soil to solution ratios (Figure C1 in Appendix C). A soil to solution ratio of 1:10 (5 g:

50 mL) and a sorption equilibration time of 24 hours were chosen for the following experiments,

as these parameters were consistent with those used previously for investigating sorption of

PFAAs in sediments (16–19).

Sorption kinetics were determined in all seven soils by equilibrating 5 g of each soil type

with 50 mL of 0.01 M CaCl2, spiked with 10 μg/mL of Masurf, followed by sampling at 2, 4, 6,

8, and 24 hours. The percentage of analytes remaining in the aqueous phase and their

corresponding distribution coefficients in each soil type were determined based on separate

analysis of the aqueous and soil phase upon reaching equilibrium, as described in the following

section. To determine sorption isotherms, this experiment was repeated at four other

164

concentrations (0.5, 1, 5, and 50 μg/mL of Masurf), with the exception that the aqueous and soil

phases were sampled only once at the equilibration time of 24 hours.

Desorption kinetics were determined for one soil type by agitating 10 μg/mL of Masurf

with 5 g of Soil A and 50 mL 0.01 M CaCl2 until sorption equilibrium (i.e. 24 hours), after which

the aqueous phase was entirely removed and replaced with an equal volume of 0.01 M CaCl2

without the test chemicals. The new mixtures were further agitated with periodic removal for

analysis at 2, 4, 6, 24, 30, and 52 hours.

In all the described experiments, blanks consisting of soil and 50 mL 0.01 M CaCl2

without the test chemicals were also included to monitor for background contamination. A set of

triplicate control samples (n = 3), consisting of only Masurf in 0.01 M CaCl2, were treated in

parallel with the Masurf-spiked soil-aqueous phase mixtures to test for the potential of abiotic

degradation and adsorption to the surfaces of the container walls. Analysis of the aqueous

phases revealed <50% by mass of C8 and C10 PFPAs and all three PFPiA congeners spiked

were present in the dissolved phase at the time of equilibration (24 hours), but subsequent rinses

of the control containers with methanol were sufficient to fully recover the PFPAs and PFPiAs

adsorbed to the container walls (Figure C2 in Appendix C). Due to this observed surface

adsorption in the controls, calculations of distribution coefficients in the sorption experiments

were based on separate analysis of the soil and aqueous phases, as described below.

5.3.4 Determination of Distribution Coefficients

At each sampling timepoint in the above experiments, the soil and aqueous phases in the

Masurf-spiked samples were separated by centrifugation at 6000 rpm for 30 minutes. These

separated aqueous samples and those from the controls were diluted with methanol and analyzed

directly using liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS). For

soil-aqueous phase mixtures removed upon sorption equilibration (i.e. 24 hours) in each

experiment, 1 g of the separated soil was extracted using a modified ion-pair method developed

by Hansen et al. (40), while another aliquot was removed for oven drying at 105oC to determine

the soil moisture content. All soil concentrations are reported based on oven dry mass.

For the purposes of determining mass balances, a subset of the polypropylene tube walls

was each shaken with 25 mL of methanol for 30 minutes, after removing the aqueous phases and

165

as much of the soil as possible by scraping, followed by direct LC-MS/MS analysis of an aliquot

of the methanol extract. Detailed extraction and instrumental methods and parameters (TableC

2) are provided in Appendix C.


The C6, C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs were quantified

using matrix-matched calibration standards, with the blank soil-aqueous mixtures serving as the

matrix. This was necessitated by the lack of commercially available isotopically labeled

surrogates of the PFPAs and PFPiAs at the time of analysis. Further details on preparation of the

matrix-matched standards are available in Appendix C.

Spike and recovery experiments were performed in triplicate (n = 3) in the aqueous CaCl2

phase, soil, and empty polypropylene containers, all of which the spiked chemicals would come

into contact with during the experiments. Details on the spike and recovery procedures and the

recovery ranges for each phase (Table C3) are provided in Appendix C.

The limits of detection (LODs) and quantitation (LOQs) were defined as the

concentrations at which a signal-to-noise (S/N) ratio of equal to or greater than 3 and 10

respectively were obtained. The method LOD and LOQ for each PFPA and PFPiA congener in

soil and the aqueous phases are listed in Table C3 in Appendix C. For calculating arithmetic

means of each triplicate set of data, concentration values below the LOD were assigned a value

of zero and values greater than the LOD but below the LOQ were used unaltered. All reported

concentrations are presented here as arithmetic means with standard error.

Procedural blanks (n = 1 for each batch of extractions) were treated in parallel with the

separate extraction and analysis of the aqueous 0.01 M CaCl2 phase and soil. PFPA and PFPiA

contamination was not detected in these blanks.

Biotransformation of the PFPiAs to the corresponding PFPAs of equal perfluorocarbon

chain length has been previously observed in PFPiA-dosed rainbow trout (25). The mechanism

by which this biotransformation occurs is unknown, but microbial enzymes, such as C–P lyase,

have a demonstrated ability to cleave the C–P bond during in vitro incubations with organic

phosphonate and phosphinate compounds (41, 42). As such, the efficacy of HgCl2 as a biocide

here was tested by equilibrating triplicate (n = 3) tubes containing 5 g of Soil A and 100 ng/mL

of 8:2 fluorotelomer unsaturated acid (8:2 FTUCA), a known precursor to PFOA (43), in 0.01 M

166

CaCl2 for 24 hours. Concentrations of 8:2 FTUCA was observed to decrease in the aqueous

phase sampled right after spiking and at 24 hours without a corresponding increase in the

aqueous and soil levels of PFOA (Figure C3 in Appendix C), which suggests the observed

decrease in 8:2 FTUCA aqueous concentrations was primarily due to sorption to soil, not

biodegradation.

Individual batch experiments were also performed, using the neat material of C6, C8, and

C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs provided by Wellington Laboratories, by

equilibrating each congener with 5 g of Soil A in 0.01 M CaCl2 for 24 hours (Figure C4 in

Appendix C). No production of C6 and C8 PFPAs was observed in any of the PFPiA-spiked

soil-aqueous phase mixtures. Together with the data obtained from the 8:2 FTUCA experiments,

these results suggest either pre-equilibration of the soil-aqueous phases with HgCl2 effectively

inhibited microbial degradation or that biotransformation, if occurring, was not significant

enough to yield detectable products within the time required to reach sorption equilibrium (i.e.

24 hours).

5.3.6 Data Analysis

The percentage of analyte remaining in the aqueous phase upon equilibration with soil

was calculated based on the percent ratio of analyte mass detected in the aqueous phase to the

nominal mass spiked in the soil-aqueous phase mixture. This percent value was calculated at

each sampling timepoint and plotted versus time to determine the length of the sorption

equilibrium.

The distribution coefficient, Kd, was calculated based on the ratio between the analyte

concentration measured in the soil (Cs,e, ng/g dry weight (dw)) and that measured in the aqueous

phase (Caq,e, ng/mL), upon sorption equilibrium: Kd (mL/g) = Cs,e/Caq,e. The organic-carbon

(OC)-normalized distribution coefficient, KOC, was calculated by normalizing Kd to the organic

content of the corresponding soil (%OC): KOC (mL/g) = Kd·(100/%OC).

Data obtained from the sorption isotherm experiments performed at the concentration

range of 0.5–50 μg/mL of Masurf were fitted to the Freundlich sorption equation: logCs,e = logKF

+ 1/n·logCaq,e; where KF is the Freundlich sorption coefficient (mL/g, if n = 1; ng1-1/n

·(mL3 )

1/n·g

-

1, if n ≠ 1) and n is the regression coefficient that indicates the linearity of the isotherm.

167

Percent desorption (%D) was calculated based on the fraction of mass of analyte

desorbed in the aqueous phase (mdes) relative to that previously sorbed on the soil at equilibrium

(ms,e): %D = (mdes/ms,e)·100%.

All statistical tests were performed using StatsDirect (Version 2.7.8, 2010). An α-value

of 0.05 was chosen as the criterion for statistical significance in all analyses, unless specified

otherwise.


5.4.1 Sorption Kinetics and Isotherms in Different Soils

Except for C6 PFPA, all of the spiked PFPAs and PFPiAs were observed to sorb

significantly to the soils studied here (Figure 5.1, Figure C2 in Appendix C). For the majority of

the analytes, sorption equilibrium was rapid, typically within 6 to 24 hours, which was consistent

with the times observed in other experiments investigating sorption of PFAAs to sediments (17–

19). Analysis of the Masurf-spiked CaCl2 solution in the control samples revealed significant

losses (>75%) of the C8 and C10 PFPAs and all three PFPiAs to the surface of the container

walls upon equilibrium (Figure C2A in Appendix C). As expected, the presence of soil greatly

reduced this surface adsorption to <5%, as was observed in the subset of soil-aqueous phase

mixtures that was analyzed for mass balance distribution among the aqueous, soil, and container

phases (Figure C2B–H in Appendix C). Mass balances of C6 (400 amu) and C8 PFPAs (500

amu) from these three phases ranged from 50 to 118%, except in Soil G, in which the mass

balances were less than 50%. Mass balances for the C10 PFPA (600 amu) and PFPiAs (702–902

amu) were consistently lower than those calculated for the C6 (400 amu) and C8 PFPAs (500

amu) in the same soils (Figure C2 in Appendix C). This is consistent with the low mass

recoveries (<75%) previously determined for N-methyl and N-ethyl

perfluorooctanesulfonamidoacetates (N-MeFOSAA (571 amu) and N-EtFOSAA (585 amu)) in a

sediment sorption experiment (16).

168

Figure 5.1. Sorption kinetics (left) of spiked PFPAs and PFPiAs displayed as their percent mass fraction remaining in the aqueous

phase upon equilibration with Soil A over time. Sorption isotherms (right) of PFPAs and PFPiAs on Soil A. Each data point

represents the arithmetic mean of the triplicate (n = 3) samples. The error bar represents the standard error.

Time (hours)

0 5 10 15 20 25

% R

em

ain

ing

in

aq

ueo

us

ph

ase (

by m

ass)

0

20

40

60

80

100

120

140

C6 PFPA

C8 PFPA

C10 PFPA

C6/C6 PFPiA

C6/C8 PFPiA

C8/C8 PFPiA

log(Caq,e

, ng/mL)

-2 -1 0 1 2 3 4lo

g(C

s,e

, n

g/g

)0

1

2

3

4

5

C6 PFPA

C8 PFPA

C10 PFPA

C6C6 PFPiA

C6C8 PFPiA

C8C8 PFPiA

169

Poor mass recoveries (<50%) in sorption experiments may be due to inefficient extraction

from the soil clay interlayers into which the sorbed analyte may have irreversibly migrated

during equilibration, as was observed for oxytetracycline (44). Since the majority of the mass

balances observed were consistently less than 90%, subsequent determinations of Kd were based

on the analysis of both the soil and aqueous phases, as recommended by OECD guidelines (39).

This contrasts other PFAA sorption studies (16–20, 45), which have used the aqueous loss

method to calculate Kd by assuming the amount of analyte sorbed (ms,e) would fully account for

the difference between the initially spiked amount (mo) and that analytically measured in the

aqueous phase at equilibrium (maq,e). However, if irreversible sorption was the cause for the poor

mass balances observed here, determination of Kd based on direct soil analysis, instead of using

the aqueous loss approach, may underestimate the actual amount sorbed by excluding the

fraction lost to the soil inner layers that cannot be extracted efficiently using the methods

described here. This was evidenced by the consistently higher logKd and logKOC values (by 0.5–

1.2 log units), determined from the aqueous loss method, as compared to those measured by

direct soil analysis (Table C4 in Appendix C), although the difference was only significant for

C6 and C8 PFPAs and C6/C6 PFPiA (p < 0.05). Nevertheless, all subsequent discussions of

isotherms and Kd correlations with soil- and aqueous phase-specific parameters were based on

data obtained from direct soil analysis as per OECD recommendations.

The majority of the sorption isotherms measured for the PFPAs and PFPiAs in the

seven soils were nonlinear, with an overall n of 0.83 ± 0.04 (Table C5 and Figure C5 in

Appendix C). Nonlinearity in Freundlich isotherms is characteristic of saturation of the soil

surface sites available for sorption at high analyte concentrations, nonuniform interactions with

the soil organic matter and mineral surfaces, and competitive sorption in a multi-solute system.

Sediment sorption of PFCAs and PFSAs, present as a mixture (16) or in the presence of other

surfactants (17) at high concentrations (µg/L–mg/L), have been shown to exhibit nonlinear

isotherms, while sediments spiked separately with PFOA and PFOS at much lower

concentrations (ng/L) exhibited predominantly linear isotherms (19). Given the six PFPAs and

PFPiAs studied here constituted only 25% by mass of the Masurf used for spiking in the sorption

experiments, the remaining constituents, such as other PFPA (C12)and PFPiA (C6/C10, C8/C10,

C6/C12) (31, 46) congeners and other ingredients, may potentially interfere with the sorption

observed here. Sorption is typically suppressed in multi-solute systems due to competition for

170

surface sites among the different compounds present, as was observed in the sorption of atrazine

and metolachlor, applied as an analytical-grade mixture and a commercial pesticide formulation

to soils (47), and chlorinated aromatics in the presence of natural aromatic acids in soil (48).

Competitive sorption has also been demonstrated in the sorption of various PFCAs and PFSAs to

WWTP sludge (21) and kaolinite (49). Similarly, the distribution coefficients, logKd and

logKOC, calculated here for the PFPAs and PFPiAs, spiked from the Masurf technical product,

were, on average, 0.25 log units lower than those calculated from batch experiments spiked

individually with the analytical standards (Table C6 in Appendix C). As such, the Kd and KOC

values determined here for the C6,C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs in

all experiments using the Masurf as the spiking standard must be applied with caution as they

may be underestimated by the effects of competitive sorption, as was observed here.

5.4.2 Effect of Soil Properties on Sorption

To investigate the effect of soil-specific properties on the sorption of PFPAs and PFPiAs,

seven soils of varying geochemical parameters (i.e. pH 3.8–7.0; %OC 1.00–45.70; cation

exchange capacity (CEC) 96–335 µmol/g) were obtained from Southwestern Ontario and the U.S

(Table C1 in Appendix C). A number of sorption studies have demonstrated the influence of

organic carbon in driving the partitioning of PFAAs into sediments (16, 18, 45). Positive

correlations between Kd and the organic carbon fraction (fOC) of selected soils was only observed

to be significant at a lower significance level of α = 0.10 for C6 and C8 PFPAs and C6/C6

PFPiA (p < 0.10, r = 0.93–0.97, Table C7; Figure C6, Appendix C). Interestingly, Kd for the

higher MW PFPiAs appeared to decrease with increasing fOC, although this correlation was only

significant for C8/C8 PFPiA (p < 0.05, r = -0.97, Table C7; Figure C6, Appendix C). This is

consistent with the negative correlation of sorption with soil organic matter (SOM) observed for

glyphosate, an organophosphonate, in soils (50), although the humic fractions in SOM have also

been demonstrated to promote glyphosate sorption via hydrogen bonding of the phosphonate

moiety to the phenolic groups of humic and fulvic acids (51, 52). Weak correlation between

logKd and OC content has also been observed for PFOA and PFOS sorption to Japanese

sediments (19), although only three sediment types were considered in that study. Given the

small set of soils studied here, there is no conclusive evidence to support whether OC may

impact PFPA and PFPiA sorption in soil. Nevertheless, Kd was normalized to OC content to

171

obtain KOC (Table C4 in Appendix C) for comparison of PFPA and PFPiA sorption with other

PFCAs and PFSAs, as will be discussed.

Efforts to control the aqueous phase chemistry were avoided here, as was suggested by

Higgins and Luthy (16), to ensure analyte sorption to the soils was occurring under natural

exposure conditions. As such, the pH of the soil-aqueous systems measured prior to analyte

spiking and upon equilibration, varied from 3.8 to 7.0 across the seven soils. For the majority of

the analytes, the logKd data did not correlate well with the aqueous pH measured upon

equilibration with each of the seven soils (p > 0.05, r = -0.60–0.70, Table C6; Figure C7,

Appendix C), except for C10 PFPA and C6/C8 PFPiA, both of which exhibited a positive

correlation with pH (p < 0.05, r = 0.76–0.81, Table C6; Figure C7, Appendix C). This contrasts

the negative correlation of Kd with aqueous pH typically observed for PFCAs and PFSAs (16,

18, 53), which are expected to predominate in their anionic states at environmental pH based on

their low pKAs (<1) (54, 55). As most soils typically carry a net negative surface charge,

increasing the surrounding pH promotes deprotonation of oxides and other functional groups

present on the soil surfaces, such that the net soil surface charge becomes even more negative

(56). As such, sorption of organic anions is typically suppressed at higher pH due to electrostatic

repulsion with the increasingly negative charge on the soil surface, but under these same

conditions, the increase of negatively charged functional groups may also promote formation of a

cation interlayer in which solution cations, such as Al3+

, Fe3+

, Ca2+

, and Mg2+

, may offset the

negatively charged surfaces, as well as, act as a bridge to bind organic anions. The latter

phenomenon was proposed to account for the observed increase in PFOS sorption to sediments

as pH was adjusted from 7 to 8 (18), and is consistent with past observations of higher Kd values

of PFAAs with increasing salinity (16, 18, 45). Further corroboration of this sorption

dependency on aqueous cation concentrations was demonstrated by the significant correlation

observed between the logKd of C6 and C8 PFPAs and C6/C6 PFPiA and soil CEC (p < 0.05, r =

0.95–0.98, Table C6; Figure C8, Appendix C), although no correlation was observed for C10

PFPA and the higher MW PFPiAs.

Unlike previous experiments in which the pH dependence of PFAA sorption was

investigated by varying the solution pH in one sediment system (16, 18), the effect of pH on

PFPA and PFPiA sorption was studied here based on comparing Kd values measured from the

seven soil-aqueous systems, all of which not only varied in pH, but also in fOC and likely other

172

sorption-dependent parameters, such as aqueous salinity. Higgins and Luthy have previously

noted the difficulty of experimentally changing solution-specific parameters, like pH and salt

concentrations, in one sediment system without affecting one another (16); therefore, Kd

comparisons among multiple soil systems, as was performed here, may be even more

complicated due to the natural heterogeneity of soils. No single parameter was observed to

significantly influence the sorption of all the PFPAs and PFPiAs investigated, but this may be

due to interferences among the multiple parameters as they vary simultaneously from soil to soil.

5.4.3 Effect of Structural Features on Adsorption and Desorption

As was observed for PFCAs and PFSAs in previous batch sorption experiments in

sediments and soils (16, 19, 20), a significantly positive correlation was observed between the

mean log KOC, calculated from the seven soils used here, and number of CFs present in the

perfluorocarbon tail of the PFPAs and PFPiAs (p = 0.0059, r = 0.94) (Figure 5.2). This

relationship is also consistent with the size-dependent accumulation trend of PFPAs and PFPiAs

observed in juvenile rainbow trout upon dietary exposure (25) and rats upon intraperitoneal

injection of these chemicals (31), which suggest absorption may represent one potential mode of

partitioning into solid environmental and biological matrices for these chemicals.

Organic carbon-normalized desorption coefficients (logKdes) of the PFPAs and PFPiAs

were similarly observed to correlate significantly with the number of CFs present in their

perfluorocarbon tails (p = 0.0156, r = 0.90) (Figure C9 in Appendix C). Full remobilization of

the C6 PFPA sorbed to Soil A was observed in the aqueous phase within two hours of replacing

the previously Masurf-spiked aqueous phase with blank CaCl2 solution, while the propensity of

the other PFPA and PFPiA congeners to remobilize decreased with increasing molecular size

(Figure C9 in Appendix C). This desorption pattern further supports the predominance of the

smaller MW PFPAs in aqueous media, such as surface waters (29), as compared to the higher

MW PFPiAs, which are more likely to be detected in solid matrices, such as WWTP sludge (31).

173

Figure 5.2. Dependence of logKOC on the number of perfluorinated carbons present in PFSAs,

PFCAs, PFPAs, and PFPiAs. LogKOC data for the PFSAs and PFCAs were measured by Higgins

et al. (16) and Ahrens et al. (19) in sediments.

Based on the logKOC data obtained here for C6, C8, and C10 PFPAs and those reported

for other PFAAs (16, 19), PFSAs still exhibited the strongest sorption to soils and sediments, as

compared to the PFCAs and PFPAs of equal perfluorocarbon chain length (16, 19, 20), with their

logKOCs (3.0–3.7) similar to those observed for the higher MW PFPiAs (3.2–3.6) (Figure 5.2).

Differences between the sorption of PFCAs and PFPAs of equal perfluorocarbon chain length

were not as clearly distinguished, as the logKOCs of perfluorononanoate (PFNA, 8 CFs) and C8

PFPA were within error of one another (i.e. PFOS > PFNA ≈ C8 PFPA, logKOC), although an

increasing logKOC trend was observed for the C10 PFAAs in the order of PFDS > PFUnA > C10

PFPA. Unlike the singly charged PFCAs, PFSAs, and PFPiAs, the PFPAs primarily circulate as

dianions at environmental pH, and would presumably be more repelled by the negatively charged

soil and less inclined to sorb.

These results are consistent with previous studies that have shown the sorption of PFAAs

is controlled by both the hydrophobic effect of the perfluorocarbon chain length and the

functionality of the headgroup (16, 18, 49, 53, 57), but the relative importance of these effects is

Number of CF's

4 6 8 10 12 14 16 18

log

Ko

c (

mL

/g)

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

4.0

4.5

PFPA

PFPiA

PFCA

PFSA

174

currently not well understood. Higgins and Luthy have previously proposed absorption as the

dominant process of PFAA sorption to sediments, whereby the hydrophobic perfluorocarbon tail

has fully penetrated into the organic matter, while the headgroup may either be wholly or

partially embedded within the organic matter and oriented towards the aqueous phase (16).

However, recent evidence of preferential interactions between PFOA and the protein-derived

components of a peat soil (58) indicates organic matter is highly heterogeneous and contains

various domains, each with different propensity to sorb contaminants. This, together with the

demonstrated effects of cations, pH, and salinity on the headgroup’s potential to molecularly

interact with the surfaces of natural sorbents (16, 18, 45, 49, 53, 57), suggest PFAAs may be

undergoing dual-mode sorption, as described by Xing and Pignatello (59). In this mechanism,

soil organic matter is conceptualized as an amalgam of rubbery and glassy phases, both of which

are capable of absorbing contaminants based on their hydrophobicity, but the latter phase also

contains holes in which adsorption through site-specific covalent and/or ionic interactions may

occur (59). These cavities confer specificity to the sorption of different classes of PFAAs and

can account for the nonlinear isotherms and competitive sorption observed here for the PFPAs

and PFPiAs and elsewhere for the PFCAs and PFSAs (21, 49). The relative contribution of

absorption and adsorption processes to the overall sorption of PFAAs depends on the

composition (e.g. fOC, mineral content) of the sorbent of interest.

5.5 Implications for Environmental Distribution of PFPAs and PFPiAs

Together with the previously measured biomagnification factors (BMFs) for PFAAs in

rainbow trout (24, 25), the sorption data determined here for the PFPAs and PFPiAs and PFCAs

(2.11–3.47, logKOC) and PFSAs (2.68–3.66, logKOC) (16, 19) may be used to visualize the

distribution of PFAAs in a simplified aquatic environment, consisting of biota, represented by

fish here, and a soil or sediment phase. As shown in Figure 5.3, PFAAs containing ≤7 CFs are

predominantly found in the aqueous phase, while PFAAs with ≥8 CFs tend to partition into either

biota or environmental solid matrices. Environmental partitioning of PFAAs is also dependent

on head-group such that PFSAs tend to exhibit the highest KOC and BMF, as compared to the

PFCAs and PFPAs of equal perfluorocarbon chain length. This is consistent with the congener

distribution of PFCAs and PFSAs often observed in field measurements of surface waters,

sediments, and freshwater biota.(23, 60–63).

175

Figure 5.3. Distribution of PFAAs in a simplified aquatic environment based on the logKOC and logBMF measured for the PFPAs,

PFPiAs, PFCAs, and PFSAs. LogKOC data for the PFSAs and PFCAs were measured by Higgins et al. (16) and Ahrens et al. (19) in

sediments. LogBMF data for the PFSAs and PFCAs were measured by Martin et al. (24) and logBMF data for the PFPAs and PFPiAs

were measured by Lee et al. (25) in juvenile rainbow trout.

Number of CF's

6 8 10 12 14 16 18

log

KO

C (

mL

/g)

-4

-2

0

2

4

log

BM

F

-4

-2

0

2

4

PFPA

PFPiA

PFCA

PFSA

PFPA

PFPiA

PFCA

PFSA

PFPA

PFPiA

PFCA

PFSA

PFPA

PFPiA

PFCA

PFSA

PFPA

PFPiA

PFCA

PFSA

PFPA

PFPiA

PFCA

PFSA

Water Biota

Soil/Sediment

176

It is important to note that the distribution profile, shown in Figure 5.3, is flexible to

changes in the ambient environment and is not rigid in the partitioning of specific CF ranges of

PFAAs to a particular phase. Short-chain PFCAs and PFSAs (≤7 CFs) have been detected in

biota and sediments either through direct exposure or indirectly by biotransformation of some

precursor material present, while surface water contamination of longer-chain PFAAs (≥8 CFs)

has also been reported, but their relative concentrations in each matrix are often dependent on

their chain length and headgroup. Furthermore, greater sorption is expected to occur in sorbents

with larger surface areas, such as the sediment bed lying at the bottom of a water body, as

compared to suspended particulates present in the same environment. By observing the runoff

dynamics of PFAAs during snowmelt in an urban watershed, Meyer et al. identified a chain-

length threshold for PFAA solid-water partitioning whereby bulk streamwater concentrations of

PFAAs with <9 CFs were highly correlated with one another and corresponded to the water-

soluble fraction, while PFAAs with >9 CFs are primarily sorbed to the suspended particulate

phase (64). The fact that this threshold (9 CFs) is higher than that proposed in Figure 5.3 (7 CFs)

suggests the distribution of PFAAs is favoured towards the aqueous phase in the presence of

suspended particulates due to their inherently smaller volume capacity in an aqueous

environment, as compared to bottom lying sediments whose much larger capacity allows them to

sorb PFAAs with chain length as short as 7 CFs. As such, phase partitioning can significantly

change depending on the nature of the sorbent present in the environment.

The observed contamination of PFPAs in surface waters (29) and their lack of detection

in WWTP sludge and sediment samples (32) suggest these chemicals are primarily aqueous

contaminants. Desorption of C6 PFPA and to a lesser extent, the C8 and C10 PFPAs, as was

observed earlier, suggest these may be remobilized from the soil and permeate through the

porewater environment. Groundwater contamination of these chemicals has yet to be reported,

although the C8 PFPA was recently observed, at levels just below its detection limit of 95 pg/L,

in two tap water samples from the Netherlands (65). The partitioning coefficients in Figure 5.3

suggest the PFPiAs should predominate in biota, such as fish (30), and environmental solids,

such as soil, sediments, and WWTP sludge (31). However, the Kds and KOCs measured here for

the PFPAs and PFPiAs may be underestimated by the inability of the soil extraction methods

used to capture the entire amount of analytes sorbed, including that which may have become

embedded within the inner layers of the soil over time. This is important when considering how

177

much of the observed contamination would be bioavailable for degradation and whether these

chemicals may eventually remobilize into the environment over time.

Despite their high MW (>700 amu), suppressed PFPiA bioaccumulation, as compared to

PFCAs and PFSAs with much lower MW (<700 amu), has been reported in rainbow trout and

was in part attributed to metabolic transformation to their corresponding PFPAs (25). Although

PFPiA biotransformation was not observed here, perhaps due to microbial inhibition by HgCl2

and/or insufficient equilibration time with the soil, the presence of glyphosate-degrading bacteria

in soil, all of which are capable of cleaving the C–P bond (66), suggests this degradation

pathway is possible for the PFPiAs in soil and warrants further investigation.


We would like to thank Nicole Riddell and Wellington Laboratories (Guelph, ON) for

donating the PFPA and PFPiA neat material, native and mass-labeled standards; Eric Reiner

(Ministry of the Environment, Etobicoke, ON), Shane De Solla (Environment Canada,

Burlington, ON), Jeff Geddes and Geoff Stupple (University of Toronto, ON), John Kudlowsky

(EarthCo Soil, Concord, ON), and John Washington (U.S. EPA, Athens, GA) for collecting and

donating the soils used in this study; and Ling Li, Inthuja Selvaratnam, and Myrna Simpson

(University of Toronto, ON) for their assistance. This research is funded by the Natural Science

and Engineering Research Council of Canada (NSERC) and a NSERC Postgraduate Scholarship

awarded to H.L.


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CHAPTER SIX

Dietary Bioaccumulation of Perfluorophosphonates and Perfluorophosphinates in Juvenile

Rainbow Trout: Evidence of Metabolism of Perfluorophosphinates

Holly Lee, Amila O. De Silva, and Scott A. Mabury


Contributions: Holly Lee was responsible for conceiving the experimental design, care and

handling of animals during the experiments, performing all bioaccumulation experiments,

method development, sample acquisition, and data interpretation. Amila De Silva assisted in

the training of fish dissection and fish physiology. Holly Lee prepared this manuscript with

editorial comments provided by Amila De Silva and Scott Mabury

Reproduced with permission from Emvironmental Science and Technology


184

6.1 Abstract

The perfluorophosphonates (PFPAs) and perfluorophosphinates (PFPiAs) are high

production volume chemicals that have been observed in Canadian surface waters and

wastewater environments. To examine whether their occurrence would result in contamination

of organisms in aquatic ecosystems, juvenile rainbow trout (Oncorhynchus mykiss) were

separately exposed to a mixture of C6, C8, and C10 monoalkylated PFPAs and a mixture of

C6/C6, C6/C8, and C8/C8 dialkylated PFPiAs in the diet for 31 days, followed by 32 days of

depuration. Tissue distribution indicated preferential partitioning to blood and liver. Depuration

half-lives ranged from 3 to 43 days and increased with the number of perfluorinated carbons

present in the chemical. The assimilation efficiencies (α, 7–34%) and biomagnification factors

(BMFs, 0.007–0.189) calculated here for PFPAs and PFPiAs were lower than those previously

observed for the perfluorocarboxylates (PFCAs) and perfluorosulfonates (PFSAs) in the same

test organism. Bioaccumulation was observed to decreased in the order of PFSAs > PFCAs >

PFPAs of equal perfluorocarbon chain length and was dependent on the charge of the polar

headgroup. Bioaccumulation of the PFPiAs was observed to be low due to their rapid

elimination via metabolism to the corresponding PFPAs. Here, we report the first observation of

an in vivo cleavage of the carbon–phosphorus bond in fish, as well as, the first in vivo

biotransformation of a perfluoroalkyl acid (PFAA). As was previously observed for PFCAs and

PFSAs, none of the BMFs determined here for the PFPAs and PFPiAs were greater than one,

which suggests PFAAs do not biomagnify from dietary exposure in juvenile rainbow trout.

6.2 Introduction

Historical and current use of fluorinated chemicals has led to the widespread occurrence

of two classes of perfluoroalkyl acids (PFAAs), the perfluorocarboxylates (PFCAs) and

perfluorosulfonates (PFSAs), in aquatic wildlife (1) and their surrounding environments (2, 3).

Global contamination of PFCAs and PFSAs has been extensively reported in fish sampled from

U.S. rivers (4–6), the Great Lakes (1, 4, 7, 8), a German lake (9), the Mediterranean and Baltic

coasts (10, 11), the Japanese coasts (12, 13), the Chinese Yangtze river (14), Greenland and the

Faroe Islands (15), and the Arctic (16–19). Analysis of wildlife species at different trophic

levels consistently reports detection of PFOS and the longer chain PFCAs (≥7 perfluorinated

185

carbons, CFs) (4, 7, 16–20), with significant concentrations (mid to high ng/g wet weight (ww))

observed in both benthic feeders and predatory fishes at the top of the aquatic food web. This is

consistent with the bioaccumulation trend observed for PFAAs in which rainbow trout exposed

through water (21) and the diet (22) exhibited increased accumulation of longer chain PFSAs (≥

6 CFs) and PFCAs (≥7 CFs).

Total organofluorine analyses of freshwater (20) and marine (23) animals revealed that

known PFSAs and PFCAs may not fully account for the total fluorochemical contamination

observed in these samples, which implies the presence of other unidentified fluorinated

chemicals. Perfluorophosphonates (PFPAs) and perfluorophosphinates (PFPiAs) are newly

discovered PFAAs that structurally differ from the PFSAs and PFCAs in that their perfluorinated

carbon tails are attached through a carbon-phosphorus (C–P) bond to either a phosphonate (R-

P(O)O2-; PFPA) or phosphinate (R2-P(O)O

-; PFPiA) headgroup (Table 6.1). PFPAs and PFPiAs

are commercial fluorinated surfactants marketed for use as leveling and wetting agents in

household cleaning products(24) and defoaming agents in pesticide formulations (25), although

the latter application has been banned in the United States (U.S.) since 2008(26). Human

exposure to the PFPiAs was recently confirmed in U.S. human sera in which the C6/C6 and

C6/C8 congeners were observed at 4–38 ng/L concentrations (27). Given their high annual

production volumes (10,000–500,000 lbs) as reported in 1998 and 2002 (28), PFPAs and PFPiAs

are likely to be widely disseminated in the environment.

The PFPAs are prevalent contaminants in Canadian surface waters, with concentrations

ranging in the mid-to-high pg/L range (29). The presence of PFPAs in wastewater treatment

plant (WWTP) effluents (29) and PFPiAs in WWTP biosolids (30) suggests the potential of these

chemicals to partition from the aqueous phase into environmental solids like sediments, as was

previously observed for PFSAs and PFCAs (31). Benthic feeders, such as Lumbriculus

variegatus, have been observed to bioaccumulate PFAAs upon exposure to laboratory-spiked

and contaminated freshwater sediments (32, 33). Together with the significant PFSA and PFCA

contamination observed in other benthic organisms, such as the freshwater invertebrate,

Diporeia, and predatory fish, sculpin (7), these results suggest sediment may be an important

source of fluorinated chemicals to these and possibly higher trophic organisms within the aquatic

food web.

186

Table 6.1. Structures, full names, and acronyms of the target analytes monitored.

Structure Full Name Acronym

P

O

-OF

F F

O-x


Cx PFPA

x = 6, 8, 10


P

O

-O

F

F F

x F

FFy


Cx/Cy PFPiA

x/y = 6/6; 6/8; 8/8


OF

F F

O-x

Perfluorocarboxylate

Cx+1 PFCA

x = 4 – 10


The present research aims to evaluate the uptake and depuration of three PFPA (C6, C8,

and C10) and three PFPiA (C6/C6, C6/C8, and C8/C8) congeners in juvenile rainbow trout

(Oncorhynchus mykiss) upon dietary exposure for 31 days, followed by a 32-day depuration

phase.


6.3.1 Chemicals


Information (SI) in Appendix D. Neat material (~1 mg for each congener) of C6 n-PFPA

(>98%), C8 n-PFPA (>98%), C10 n-PFPA (>98%), C6/C6 n-PFPiA (>98%), C6/C8 n-PFPiA

(>98%), and C8/C8 n-PFPiA (>98%) were provided by Wellington Laboratories (Guelph, ON).

6.3.2 Food Preparation

Three batches of commercial fish feed (Silver Cup 1.5 mm extruded floating feed, Martin

Mills Inc., Elmira, ON) were prepared for the separate dosing of PFPAs and PFPiAs and the

187

control feed, as described in detail in Appendix D. Determination of the PFPA and PFPiA

concentrations in the dosed and control feed is described in Appendix D. The mean (± standard

error) concentrations in the PFPA- and PFPiA-dosed feed were 485 ± 28 ng/g C6 PFPA, 474 ±

37 ng/g C8 PFPA, and 533 ± 37 ng/g C10 PFPA; and 468 ± 12 ng/g C6/C6 PFPiA, 510 ± 24

ng/g C6/C8 PFPiA, and 420 ± 12 ng/g C8/C8 PFPiA respectively (Tables 2 and D1). The

PFPAs and PFPiAs were not detected in the control feed.

6.3.3 Fish Care and Sampling

Juvenile rainbow trout (5–7 g) were purchased from Humber Springs Trout Hatchery

(Orangeville, ON) and allowed to acclimate for two weeks prior to chemical exposure. The

animals were housed in three 475 L fiberglass tanks under flow-through conditions (4–8 L/min)

using carbon-filtered and dechlorinated water, maintained at 18oC, at the Aquatic Facility of the

Department of Cell and Systems Biology at the University of Toronto. A 12-hour daily

photoperiod was used. One tank was designated for the control fish, while the remaining two

tanks were designated for fish to be dosed separately with the PFPAs and PFPiAs. The initial

fish loadings in the three tanks were ~0.5 g/L. All animals in this research were treated and used

under approval by the University of Toronto Animal Care Committee and in accordance with the

guidelines of the Canadian Council on Animal care.

Prior to chemical exposure, fish were deprived of food for 24 hours to ensure an

aggressive first feeding. During the exposure phase, fish received daily feeding of the dosed or

control feed at 0.015 g feed (dry weight (dw))/g fish (ww), adjusted throughout the experiment

for growth, followed by a depuration phase during which the fish were fed untreated feed at the

same rate. Fish sampling always occurred before feeding. The fish were sampled on days -1

(predose), 1, 3, 6, 13, 20, and 31 of the exposure phase and days 1, 3, 8, 15, 25, and 32 of the

depuration phase. The fish were sampled in triplicate (n = 3) at each timepoint until days 25 and

32 of the depuration phase during which the control fish were sampled in duplicate (n = 2) and

the dosed fish were sampled in triplicate (n = 3) from their respective tanks. Each fish was

euthanized by a lethal overdose of tricaine methanesulfonate (MS-222, 4 g/L solution buffered to

pH 7 with sodium bicarbonate). After weighing, the fish were dissected to remove the livers and

subsequently minced into small carcass pieces. To minimize potential PFPA and PFPiA

188

contamination of the carcass from undigested food, the digestive tract, containing the esophagus,

stomach, pyloric caeca, and intestines, was discarded. The livers were weighed separately then

returned to their corresponding carcasses for further homogenization. All samples were archived

at -20oC until further analysis. The masses of the whole fish and their corresponding livers are

plotted as Figure D1.

6.3.4 Tissue Distribution of PFPAs and PFPiAs

On the last day of the exposure phase (i.e. day 31), three additional fish (n = 3) from each

of the control and dosed tanks were sampled to investigate tissue distribution. Fish were

euthanized by a 1 g/L solution of MS-222 (buffered to pH 7 with sodium bicarbonate). Whole

blood was collected through cardiac puncture with heparin-rinsed syringes and stored in

heparinized vials (BD Vacutainer, Franklin Lakes, NJ). Fish were dissected to remove the heart,

liver, kidneys, and gills. The remaining carcass was homogenized, as described above. All

samples were archived at -20oC until further analysis.

6.3.5 Extractions and Instrumental Analysis

Whole-fish homogenates, livers, kidneys, hearts, gills, and whole blood samples were

extracted using a modified version of the ion-pairing method developed by Hansen et al (34).

The livers, kidneys, and gills were homogenized in 1-2 mL of 1M tetrabutylammonium

hydrogen sulfate (TBAS) prior to extraction. Detailed extraction methods, chromatographic

gradients, instrumental conditions (Table D2 and D3), and sample chromatograms (Figure D2)

are provided in Appendix D.


The C5–C11 PFCAs were quantified using mass-labeled internal standards (Table D3).

The C6, C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs were quantified using

matrix-matched calibration standards where control fish homogenate served as the matrix. This

was necessary since isotopically-labeled surrogates of the PFPAs and PFPiAs were not available

189

at the time of analysis. Further details on preparation of the matrix-matched standards are

provided in Appendix D.

Spike and recoveries ranged from 73 to 126% for the PFPAs and PFPiAs in the different

fish tissues (Table D4a) and 68 to 123% for the C5-C11 PFCAs in whole-fish homogenates

(Table D4b). All reported tissue concentrations were not corrected for recovery. Further details

on the spike and recovery procedures are described in Appendix D.

The limits of detection (LOD) and limits of quantiation (LOQ) were defined as the

concentrations producing a signal-to-noise (S/N) ratio of equal to or greater than 3 and 10

respectively. The method LOD and LOQ values for each analyte in the different fish tissues are

listed in Table D4ab. For the purposes of calculating means, concentration values below the

LOD were assigned a value of zero and values greater than the LOD but below the LOQ were

used unaltered. All reported concentrations are presented as arithmetic means with standard

error.

One procedural blank (high pressure liquid chromatography (HPLC) grade water, n = 1)

was included in the extraction of each timepoint. No PFPA and PFPiA contamination was

observed in the procedural blanks.

The Canadian government recently listed the PFPAs and PFPiAs of varying

perfluorocarbon chain length as potential precursors to long chain PFCAs (≥8 CF’s) (35) and

therefore PFCAs were monitored in fish, although they were not a component in the dosing. No

production of PFCAs (Figure D3) was observed in PFPA- and PFPiA-dosed fish homogenate

extracts, as discussed in Appendix D.

6.3.7 Data Analysis

The whole-body and liver growth rates (Table D5) were calculated by fitting all fish and

liver mass data to an exponential model: (ln(mass, g) = b·t + a; where b is the growth rate (/day),

t is the time (day), and a is a constant. Whole-body concentrations in each treatment population

were corrected for growth dilution by using the individual whole-body growth rates shown in

Table D5. The liver somatic index (LSI) was calculated as LSI (%) = [liver mass (g)/whole fish

mass (g)] x 100%. Mean LSIs calculated for each batch of fish sampled from the three treatment

populations throughout the experiment are plotted as Figure D4.

190

Depuration rate constants (kd) from the whole-body homogenates were calculated by

fitting the growth-corrected concentration data in the depuration phase to the first-order decay

model: ln(Cfish) = kd·t + a; where Cfish is the growth-corrected whole-body concentration, kd is the

depuration rate constant (/day), t is the time (day), and a is a constant (StatsDirect, Version 2.7.8,

2010). Depuration half-lives (t1/2) were calculated as ln(2)/kd.

Assimilation efficiency (α) was determined by using iterative nonlinear regression to fit

the growth-corrected concentration data in the exposure phase to the integrated form of the

kinetic rate equation for constant dietary exposure (SigmaPlot, Version 9.01, 2004): Cfish =

(α·F·Cfood/kd)·[1 – exp(-kd·t)]; where Cfish is the growth-corrected whole-body concentration, F is

the feeding rate (0.015 g food dry wt/g of fish ww/day), Cfood is the concentration in the food,

and t is the time (day) (36). Assimilation efficiency is expressed as the ratio of the amount of

chemical absorbed to the amount fed. Biomagnification factors (BMFs) were calculated using

the kinetic equation method since steady-state was not achieved for all the analytes during the

exposure phase (BMF = α·F/kd).

The estimated time to achieve 90% steady-state (tss, day) for each analyte was calculated

by rearranging the above kinetic rate equation, as described in Appendix D.

6.3.8 Statistical Analysis

Analyte concentrations observed below their corresponding LODs in the depuration

phase were imputed as the LOD divided by square root of two, so that they can be fitted as

nonzero values to the first-order decay model described above for calculating kd and t1/2. All

tests were performed using StatsDirect (Version 2.7.8, 2010). An α-value of 0.05was chosen as

the criterion for statistical significance in all analyses. Further details describing the test results

and the p-values of the tests are provided in Appendix D.


6.4.1 Physical effects observed in fish

No mortality occurred in either of the dosed and control populations. Significantly higher

whole-body and liver growth rates (0.016 /day, whole-body and liver; p < 0.05, Tables D5, D6b)

were observed in the control population than in either of the dosed populations (0.0059–0.0096

191

/day). The fact that these growth rates were lower in the dosed populations than in the control

contrasts a number of studies in which rainbow trout, exposed to PFCAs and PFSAs at

concentrations similar to those used here, exhibited growth rates that were not significantly

different than those for the control (21, 22, 37). LSI factors were calculated since this is a

measure of liver enlargement and is used as an indicator of metabolic stress in an animal upon

chemical exposure. No significant difference was observed in the mean LSIs calculated between

the control and either of the dosed populations (p > 0.05, Table D7b) and no temporal trend was

observed in the LSIs from the control and dosed populations (Figure D4), both of which suggest

the absence of liver enlargement in the fish used in this experiment.

6.4.2 Uptake and depuration of PFPAs and PFPiAs

All six dosed PFPAs and PFPiAs were detected in the whole-fish homogenate samples

within 1 day of exposure (Figure 6.1). To statistically determine whether the PFPAs and PFPiAs

reached steady-state, Pearson’s correlation tests were performed on the last four to six

concentration data points of the exposure phase for each analyte. Regression of the C6, C8, C10

PFPAs and C6/C6 PFPiA data all produced slopes that were not significantly different from zero

(p > 0.05, Table D8). Together with their estimated times of <31 days to achieve 90% steady-

state (Table 6.2), these results are consistent with the plateau observed in the whole-fish

concentrations of the three PFPAs by day 13, although the C6/C6 PFPiA concentrations

appeared to rise towards the end of the exposure phase (Figures 6.1 and D5). On the other hand,

the regression slopes of the C6/C8, and C8/C8 PFPiA data were significantly different from zero

(p < 0.05, Table D8), which suggest these analytes did not reach steady-state. This is also

consistent with the uptake data observed for these two PFPiAs with estimated times to 90%

steady-state longer than the 31-day exposure phase (Figures 6.1 and D5, Table 6.2).

The assimilation efficiencies observed here for the PFPAs (9–16%, Table 6.2) and

PFPiAs (17–34%, Table 6.2) were lower than those reported for the PFCAs and PFSAs (59–

130%) in rainbow trout (22). The reduced uptake of the PFPiAs (MW >700 amu) is consistent

with the poor assimilation typically observed for chemicals with molecular weights greater than

600 amu (38), as was observed for the C16-chlorinated alkanes (39) and other organochlorines

with logKOW ≥ 7 (40, 41), and may in part be due to steric constraints in crossing biological

membranes during absorption into the gut.

192

Figure 6.1. Growth-corrected whole-body homogenate concentrations (ng/g in wet weight, (ww)) of C6, C8, and C10 PFPAs and

C6/C6, C6/C8, and C8/C8 PFPiAs in rainbow trout during exposure and depuration phase. The top panels represent the data collected

from PFPA-dosed fish and the bottom panels represent the data collected from PFPiA-dosed fish. Each data point represents the

arithmetic mean concentration of the triplicate (n = 3) sampling at each timepoint. The error bar represents the standard error.

Time (day)

0 10 20 30 40 50 60 70

Co

nc

en

tra

tio

n i

n f

ish

(n

g/g

ww

)

0.001

0.01

0.1

1

10

100

0 10 20 30 40 50 60 70

0.001

0.01

0.1

1

10

100

Exposure phase (Day 0 to 30)

Depuration phase (Day 31 to 63)

0 10 20 30 40 50 60 70

0.001

0.01

0.1

1

10

100

0 10 20 30 40 50 60 70

0.001

0.01

0.1

1

10

100

0 10 20 30 40 50 60 70

0.001

0.01

0.1

1

10

100

0 10 20 30 40 50 60 70

0.001

0.01

0.1

1

10

100

C6 PFPA C8 PFPA C10 PFPA

C6/C6 PFPiA C6/C8 PFPiA C8/C8 PFPiA

Exposure Depuration Exposure Depuration Exposure Depuration


193

Table 6.2. Concentration of food (Cfood, in dry weight (dw)), depuration rate constant (kd), depuration half-life (t1/2), assimilation

efficiency (α), biomagnification factor (BMF) of the dosed PFPAs and PFPiAs, and estimated time to achieve 90% steady state (tss).

The coefficient of correlation (r) for the linear regression analysis to determine kd is shown in parentheses. The error is represented by

±1 standard error.

Analyte Cfood

(ng/g dw) kd (/day) (r) t1/2 (day) α (%) BMF logBMF

tss

(day

)

Perfluorophosphonates (PFPAs)

C6 PFPA 485 ± 28 0.19 ± 0.03 (0.97) 3.7 ± 0.6 9 ± 4 0.007 ± 0.003 -2.13 ± 0.17 12

C8 PFPA 474 ± 37 0.16 ± 0.03 (0.96) 4.4 ± 0.7 7 ± 4 0.007 ± 0.003 -2.18 ± 0.21 14

C10 PFPA 533 ± 37 0.13 ± 0.02 (0.94) 5.3 ± 0.8 16 ± 6 0.018 ± 0.006 -1.74 ± 0.14 18

Perfluorophosphinates (PFPiAs)

C6/C6 PFPiA 468 ± 12 0.13 ± 0.01 (1.00) 5.5 ± 0.2 34 ± 6 0.041 ± 0.007 -1.39 ± 0.07 18

C6/C8 PFPiA 510 ± 24 0.03 ± 0.01 (0.88) 20.4 ± 4.9 24 ± 9 0.106 ± 0.033 -0.97 ± 0.13 77

C8/C8 PFPiA 420 ± 12 0.02 ± 0.01 (0.81) 52.7 ± 15.8 17 ± 15 0.189 ± 0.167 -0.72 ± 0.38 115

194

Although the pKA of PFPA is unknown, it is expected to be similar to the experimentally

determined pKA’s of PFOA and PFOS (<1) (42, 43), and corresponds to the first deprotonation

in the PFPA headgroup to form the monoanion. Similarly, the pKA of PFPiA may be inferred

from the reported pKA range of 1-2 for alkylphosphate diesters (44), although the inductive effect

of the perfluorinated chains in PFPiA should lower its pKA. Based on these pKA ranges (<1),

PFPAs and PFPiAs are expected to primarily circulate as anions in rainbow trout, even in acidic

compartments, such as the stomach (pH 2-4) (45, 46). Anionic chemicals are typically poorly

assimilated in animal tissues due to their reduced hydrophobicity and electrical repulsion from

the negative electrical potential inside animal cells, although there are carrier proteins that may

facilitate this transport (47). There is considerable evidence that organic anion transport (OAT)

proteins are involved in the active uptake of PFOA (C8 PFCA) into the liver (48) and the renal

transport of C6–C10 PFCAs between the kidneys and blood (49, 50) in rats. Gender differences

in PFOA elimination were also observed in sexually mature fathead minnows (51), which

suggest differential OAT protein expression may occur in sexually mature aquatic vertebrates.

However, the juvenile stage of the rainbow trout used in this experiment should preclude

activation of these hormonally-controlled transport mechanisms.

Depuration of PFPAs and PFPiAs followed first order kinetics with correlation

coefficients, r, greater than 0.80 (Table 6.2). Whole-body depuration rate constants ranged from

0.13 to 0.19/day for the PFPAs and 0.02 to 0.13/day for the PFPiAs, which corresponded to

depuration half-lives of 4 to 53 days (Table 6.2). These half-lives are within the range of those

previously observed in rainbow trout carcasses upon dietary exposure to the C8–C14 PFCAs (3–

35 days), PFHxS (9 days), and PFOS (13 days) (22) and in rainbow trout liver and blood upon

dietary exposure to a mixture of branched and linear isomers of PFOA and PFNA (3.7 days in

liver and 5.6 days in blood, n-PFOA; 6.0 days in liver and 15.9 days in blood, n-PFNA) (37).

As was observed for water-borne (21) and dietary (22) exposure to PFCAs and PFSAs,

the depuration half-lives observed here were positively correlated with the number of

perfluorinated carbons present in PFPAs and PFPiAs (p < 0.05, r = 0.94, Table D9, Figure 6.3).

This relationship is also consistent with the depuration trend observed in rats upon

intraperitoneal injection of a mixed dose of PFPAs and PFPiAs (30). It is important to note that

the difference between the headgroups of the doubly charged PFPAs and singly charged PFPiAs

may also contribute to the differences observed in their clearance rates. Furthermore, geometry

195

differences between the di-alkylated PFPiAs and the mono-alkylated PFPAs, PFCAs, and PFSAs

limit direct comparison of the clearance rates between the PFPiAs and the three other classes of

PFAAs.

6.4.3 Assimilation of PFPAs and PFPiAs into different tissues

The highest concentrations of PFPAs and PFPiAs occurred in the liver (35–572 ng/g ww)

and blood (33–116 ng/g) of rainbow trout collected on the last day of the exposure phase (Figure

D6, Table D10), which suggest these chemicals primarily accumulate in the enterohepatic

system. The high kidney concentrations also observed here (8–52 ng/g ww, PFPAs; 116–212

ng/g ww, PFPiAs; Figure D6, Table D10) are consistent with the demonstrated affinity between

renal proteins and PFAAs (49, 50). Although urine samples were not collected here, the

potential of urinary excretion as a major route of elimination for PFPAs has been previously

demonstrated by their observed high excretion efficiencies (up to 96% of the administered dose)

in the urine of dosed rats (30). In that same study, the higher molecular-weight (MW) PFPiAs

were not observed in any of the urine samples (30), the lack of which mirrored the low renal

excretion (<2% of dose) of the longer chain PFCAs (≥ C9) in rats (52). Digestive tissues and

feces were not analyzed here, but biliary excretion of the PFPAs and PFPiAs has been previously

reported in rats in which the excreted or unabsorbed chemicals were observed at <1–10% of the

administered dose in the feces (30).

The predominance of PFPAs and PFPiAs in the liver, blood, and kidneys (Figure D6) is

akin to the tissue distribution profile previously observed for PFCAs and PFSAs in fish (14, 21).

Liver-to-blood (LBRs), liver-to-carcass (LCRs), and blood-to-carcass (BCRs) concentration

ratios (Figure D7, Table D10) were calculated and generally exceeded one, which together

suggest PFPAs and PFPiAs are similar to other PFAAs in their tendency to predominate in

proteinaceous compartments like the liver and blood in fish. The magnitudes and trends of these

ratios are discussed in detail in Appendix D. Upon absorption into the bloodstream, some of the

PFPAs and PFPiAs may exit enterohepatic recirculation and enter systemic circulation in the

fish, as evidenced by their detection in the heart (nd–9 ng/g ww, PFPA; 42–57 ng/g ww, PFPiA)

and the gills (0.96–7 ng/g ww, PFPA; 34–57 ng/g ww, PFPiA) (Figure D6, Table D10). The

detection of PFPAs and PFPiAs in the gills suggests respiration may be an additional mode of

depuration of the PFPAs and PFPiAs from the fish.

196

6.4.4 Effect of biotransformation on bioaccumulation parameters

Production of the C6 and C8 PFPAs was first observed in the PFPiA-dosed fish

homogenates on day 6, with their concentrations increasing until the third day of the depuration

phase (Figure 6.2). Metabolism of the parent C6/C6 and C6/C8 PFPiAs may yield the C6 PFPA

either separately or synergistically, as with the C6/C8 and C8/C8 PFPiAs to C8 PFPA. Since the

PFPA metabolite here may derive from two potential parent compounds, metabolite yields were

estimated on a molar basis as the ratio of the amount of PFPA metabolite observed to the total

amount of the two parent PFPiAs observed, as described in Appendix D. These yields should be

treated conservatively as they are estimated by assuming equal contribution from the metabolism

of both parent PFPiAs to the corresponding PFPA. On average, 12% of the C6/C6 and C6/C8

PFPiAs observed in the fish was metabolized to C6 PFPA, while 4% of the accumulated C6/C8

and C8/C8 PFPiAs was metabolized to C8 PFPA (Figure 6.2). No PFPA contamination was

observed in the PFPiA neat material used to dose the fish feed, which further support the

detection of C6 and C8 PFPAs in the PFPiA-dosed fish as biotransformation products. The lack

of detection of C10 PFPA in the PFPiA-dosed fish is also consistent with the congener profile in

the dose in which none of the three PFPiA congeners contained a perfluorodecane (C10) linkage.

Abiotic hydrolysis of the carbon–phosphorus (C–P) bond in C1/C1, C2/C2, and C4/C4

PFPiA has been previously reported to yield the corresponding C1, C2, and C4 PFPAs under

aggressive conditions of long reaction times (~36 hours) and high temperatures (>100oC) (53),

but a biological degradation pathway has yet to be reported. Organic phosphonate and

phosphinate biodegradation has been primarily studied in in vitro systems using cultured

microbial enzymes (e.g. C–P lyase) that are known to be capable of cleaving the C–P bond (54,

55). The mechanism by which this microbial bond cleavage occurs is widely debated, but

various pathways, involving α-oxidation or free-radical dephosphorylation (55), have been

proposed. To our knowledge, this is the first observation of an in vivo production of a

phosphonate from a parent phosphinate in any organism, as well as, the first observation of an in

vivo biotransformation of a PFAA. Literature on whether PFAAs biodegrade is limited to the

observed disappearances of PFOA and PFOS in spiked WWTP sludge during anaerobic

incubations without further confirmation by the detection of PFOA and PFOS metabolites and/or

fluoride ions released from their mineralization (56).

197

Figure 6.2. (A) Growth-corrected concentrations of PFPA metabolites (ng/g wet weight, (ww)) observed in fish dosed with a mixture

of C6/C6, C6/C8, and C8/C8 PFPiAs. (B) Percent PFPA yield with respect to accumulated parent PFPiAs (mol basis) in fish dosed

with a mixture of C6/C6, C6/C8, and C8/C8 PFPiAs. Each data point represents the arithmetic mean concentration of the triplicate (n

= 3) sampling at each timepoint. The error bar represents the standard error.

Time (day)

Co

ncen

trati

on

in

fis

h (

ng

/g w

w)

0 10 20 30 40 50 60 70

0.001

0.01

0.1

1

10

100

C6 PFPA

C8 PFPA

C10 PFPA

Exposure Depuration

In PFPiA-dosed fish

0 10 20 30 40 50 60 70

Mo

lar

PF

PA

yie

ld f

rom

bo

th

po

ten

tial p

are

nt

PF

PiA

s (

%)

0

10

20

30

40

50

Exposure Depuration

%(moles of PFPA metabolite) (moles of 2 parent PFPiAs)

Time (day)

A B

198

As was also observed for their parent PFPiAs, the C6 and C8 PFPA products were

observed at the highest concentrations in the liver (27–36 ng/g ww), blood (10–13 ng/g ww), and

kidneys (5–14 ng/g) (Figure D8). This tissue distribution suggests the liver and kidneys may be

potential sites of biotransformation, although biotransformation in the gut cannot be precluded.

Higher PFPA yields with respect to their corresponding parent PFPiAs were observed in the liver

(17%, C6 PFPA; 12%, C8 PFPA) than in the kidneys (8%, C6 PFPA; 2%, C8 PFPA), which

suggest potentially higher metabolic activity in the liver. Given that certain bacterial strains have

been shown to cleave the C–P bond (54, 55), the microbial flora present in the digestive tract and

internal organs (e.g. liver and kidneys) of a fish (57) may also be responsible for the

biotransformation of PFPiAs observed here.

A number of studies have demonstrated that biotransformation of a chemical can

decrease its overall bioaccumulation potential in fish (58–60). The relatively low assimilation

efficiencies observed here for the PFPiAs (Table 6.2) are consistent with previous reports of

metabolizable compounds, like the short-chain polychlorinated alkanes (58, 59) and fipronil (60),

having small assimilation efficiencies due to confounding of this parameter by the rapid

metabolic depuration of these chemicals. The BMFs calculated here for the PFPiAs (0.041–

0.189) in rainbow trout were generally lower than those reported by Martin et al. for the PFCAs

(0.038–1.000) and PFSAs (0.14–0.32) (22) (Table 6.2, Figure 6.3).

Biotransformation of other fluorinated compounds, such as 8:2 fluorotelomer acrylate

(8:2 FTAc), 8:2 and 7:3 fluorotelomer carboxylates (8:2 and 7:3 FTCAs), was recently observed

in rainbow trout (61, 62). As expected for less persistent chemicals, their BMFs, calculated

based on liver concentrations, were lower than the carcass-based BMFs of PFCAs and PFSAs of

equal perfluorocarbon chain length (22) (Figure 6.3). This difference should only be treated

qualitatively, as a quantitative comparison between liver-based and carcass-based BMFs may not

be appropriate due to the potential magnitude of difference in the concentrations between these

two compartments.

199

Figure 6.3. Associations between the (A) depuration half-lives (t1/2) and (B) logBMFs and the number of perfluorinated carbons

present in PFSAs, PFCAs, PFPAs, PFPiAs, 8:2 FTAc, 8:2 FTCA, and 7:3 FTCA. Depuration half-lives and logBMFs for the PFSAs

and PFCAs were reported by Martin et al. (22). Note that the half-lives and logBMFs for 8:2 FTAc, 8:2 FTCA, and 7:3 FTCA were

based on liver concentrations reported by Butt et al. (61,62) and comparisons of these values to those of the other PFAAs should be

treated qualitatively.

Number of perfluorinated carbons

4 6 8 10 12 14 16 18

Dep

ura

tio

n h

alf

-lif

e (

day

)

0

20

40

60

80PFSAs

PFCAs

PFPAs

PFPiAs

8:2 FTAc

8:2 FTCA

7:3 FTCA

Number of perfluorinated carbons

4 6 8 10 12 14 16 18

log

BM

F

-5

-4

-3

-2

-1

0

A B

200

The smaller C6, C8, and C10 PFPAs (0.007–0.018, BMFs) were less bioaccumulative

than the PFPiAs and the corresponding PFCAs (i.e. PFNA and PFUnA) and PFSAs (i.e. PFHxS

and PFOS) of equal perfluorocarbon chain length (Figure 6.3). Direct BMF comparison between

C6 PFPA and PFHpA was not possible due to the lack of detection of PFHpA by Martin et al. in

their dosed fish (22), but the predicted BMF of 0.03 for PFHpA or any PFCA with six CFs,

estimated from extrapolating the logBMF vs. perfluoroalkyl chain length regression reported in

that study (22), still supports the PFCAs are more bioaccumulative than the corresponding

PFPAs of equal perfluorocarbon chain length. As was observed for the depuration half-lives, the

observed BMFs increased with the number of perfluorinated carbons present in the PFPAs and

PFPiAs (p < 0.05, r = 0.94, Table D9, Figure 6.3).

6.5 Implications for environmental contamination

The PFPAs were the least bioaccumulative compared to the corresponding PFCAs and

PFSAs of equal perfluorocarbon chain length studied so far in rainbow trout. Despite the

relatively similar MWs of the PFPAs (400–600 amu) with those of the other PFAAs (400–714

amu) studied (22), their reduced uptake may be due to the difference in the charge present in

their headgroup. Unlike the singly charged PFCAs, PFSAs, and PFPiAs, the PFPAs are doubly

charged at environmental pH (>5) and would presumably be more water-soluble and less

inclined to assimilate into animal tissues. As was observed for other metabolizable fluorinated

chemicals (61, 62), the biotransformation of the PFPiAs to PFPAs observed here resulted in their

reduced assimilation and bioaccumulation into rainbow trout despite their relatively large MWs

(>700 amu). In vivo biotransformation of a PFAA is reported for the first time here, although it

is unclear whether the fish or the bacterial flora present in the fish was responsible for this

observed metabolism.

In general, BMFs decreased in the order of sulfonate > carboxylate > phosphonate

headgroups of PFAAs of equal perfluorocarbon chain length. Direct BMF comparisons cannot

be made between the PFPiAs and other PFAAs, because no other PFAAs studied in rainbow

trout (22) had the same number of CFs as the PFPiAs and PFPiA bioaccumulation was

complicated by biotransformation, a problem that was not present with the other persistent

PFAAs. As was observed for the PFCAs and PFSAs (22), the BMFs of PFPAs and PFPiAs were

less than one, which suggest PFAAs, in general, do not biomagnify in juvenile rainbow trout

201

from dietary exposure. However, there is considerable evidence that trophic magnification of

PFOS and PFCAs (≥7 CFs) does occur in aquatic and marine food webs (4, 7, 15–20) and even in

a remote terrestrial food web in northern Canada (63). Despite the relatively low

bioaccumulation potential of PFAAs observed in fish, PFOS and PFCAs have been detected in

higher trophic level animals, such as birds, minks, seals, foxes, caribou, and polar bears (4, 7, 10,

12, 15–20, 63). This suggests the BMF data obtained here for the PFPAs, PFPiAs, and other

PFAAs (22) in rainbow trout cannot necessarily be extrapolated to predict the likelihood of their

biomagnification in higher trophic level biota. The presence of PFPAs and PFPiAs in aquatic

environments (29, 30) warrants their monitoring not only in aquatic biota, but also in terrestrial

wildlife to evaluate the potential of these chemicals to undergo trophic magnification upon

release into the environment.

6.6 Acknowledgement

We gratefully acknowledge Nicole Riddell and Wellington Laboratories (Guelph, ON,

Canada) for donating the PFPA and PPFiA neat material, native and mass-labeled standards,

Norman White, the staff at the Aquatic Facility, Alicia Sales De Andrade, Leo Yeung, Derek

Jackson (University of Toronto, ON, Canada), and Craig Butt (Duke University, NC, US) for

their assistance in this study. The present study was supported by the Environment Canada’s

Chemicals Management Plan, and funded by Natural Science and Engineering Research Council

of Canada (NSERC), and a NSERC Postgraduate Scholarship awarded to HL.


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207

CHAPTER SEVEN

A Pilot Survey of Legacy and Current Commercial Fluorinated Chemicals in Human Sera

from United States Donors in 2009



Contributions: Holly Lee was responsible for acquiring human sera samples, method

development, sample acquisition, and data interpretation. Holly Lee prepared this

manuscript with editorial comments provided by Scott Mabury

Reproduced with permission from Environmental Science and Technology


208

7.1 Abstract

Human biomonitoring has traditionally focused on analyzing the perfluorocarboxylates

(PFCAs) and perfluorosulfonates (PFSAs), although the presence of other unidentified

fluorinated chemicals has been demonstrated through total organofluorine analysis. Exposure to

legacy and current commercial fluorinated chemicals was investigated by analyzing fifty human

sera samples collected in 2009 from the United States for forty fluorinated analytes that included

the polyfluoroalkyl phosphate diesters (diPAPs), N-ethyl perfluorooctanesulfonamidoethanol-

based polyfluoroalkyl phosphate diester (SAmPAP), one fluorotelomer mercaptoalkyl phosphate

diester congener (FTMAP), fluorotelomer sulfonates (FTSs), perfluorophosphonates (PFPAs),

and perfluorophosphinates (PFPiAs). DiPAP concentrations (0.035–0.136 μg/L) for the more

dominant congeners (6:2, 6:2/8:2, 8:2) were lower than those reported in human sera samples

collected in 2004, 2005, and 2008. The SAmPAP and 6:2 FTMAP were not detected, but

exposure to SAmPAP was suggested based on the detection of one of its potential degradation

intermediates, N-ethyl perfluorooctanesulfonamidoacetate (N-EtFOSAA). PFPiAs were

detected for the first time in human sera, with C6/C6 and C6/C8 PFPiAs as the dominant

congeners, observed in >50% of the samples.

7.2 Introduction

Perfluorocarboxylates (PFCAs) and perfluorosulfonates (PFSAs) have been observed at

μg/L concentrations in human blood worldwide (1–7). The profile in human sera is typically

dominated by perfluorooctanesulfonate (PFOS, C8 PFSA), followed by perfluorooctanoate

(PFOA, C8 PFCA) and perfluorohexanesulfonate (PFHxS, C6 PFSA). One potential source of

this contamination is the metabolic transformation of commercial fluorinated materials into the

PFCAs and PFSAs. Fluorochemical production in North America has largely proceeded by

electrochemical fluorination (ECF) to produce perfluoroalkylsulfonamides (PFSAms) and

telomerization to produce fluorotelomer-based materials (8). After 3M announced the phase-out

of their perfluorooctylsulfonyl (POSF)-based materials in 2000, with production ceasing entirely

in 2002 (9), telomerization became the dominant manufacturing process of fluorochemicals in

North America (10). The PFSAm- and fluorotelomer-based starting raw materials produced

from these two processes are incorporated into polymers and surfactants for applications, such as

209

treating surfaces of fabrics, carpets, and textiles; greaseproofing food contact papers; and

leveling and wetting agents.

Fluorinated phosphate surfactants are used as greaseproofing agents in food contact

papers (11) and have a demonstrated potential to migrate into food (12, 13). The N-ethyl

perfluorooctanesulfonamidoethanol (N-EtFOSE)-based polyfluoroalkyl phosphate esters

(SAmPAPs) were used in food contact paper during the period of 1974-2000 (2, 14) until their

production ceased after the phase-out of POSF chemistries (9). Human exposure to SAmPAP is

consistent with the observed increase of N-ethyl perfluorooctanesulfonamidoacetate (N-

EtFOSAA) in human blood from 1974 to 1989 (4–6). Biotransformation of N-EtFOSE to PFOS

has been observed in rat liver microsomes, cytosol fractions, and liver slices (15), and so

SAmPAP may also represent a source of human exposure to PFOS exposure.

A family of fluorotelomer-based phosphate surfactants, the polyfluoroalkyl phosphate

diesters (diPAPs), was recently discovered at µg/L concentrations in human sera (16). DiPAPs

are established biological precursors of PFCAs in microbial and mammalian systems (17–19).

As biotransformation to PFCAs may be possible from any fluorotelomer backbone, research on

exposure to other types of fluorotelomer-based materials is warranted. The fluorotelomer

mercaptoalkyl phosphate esters (FTMAPs) have been commercialized for use in food packaging

in the United States (U.S.) since 1995 (20–22). Little is known about the potential for human

exposure and the environmental fate of these telomer-based phosphate surfactants. One possible

fate is enzyme-mediated cleavage of the carbon-sulfur (C-S) bond in the perfluoroalkylethylthio

moiety to produce the fluorotelomer sulfonates (FTSs). FTS concentrations have been observed

to increase from influent to effluent in 4 of 10 wastewater treatment plants (WWTP) studied

(23). This increase was potentially due to biodegradation of any precursors containing a

perfluoroalkylethylthio moiety. Significant groundwater contamination of FTSs (up to 14600

µg/L) near fire-training facilities has been attributed to the degradation of fluoroalkylthioamido

sulfonates (CF3(CF2)nCH2CH2SCH2CH2CONHC(CH3)2CH2SO3-) in aqueous film-forming

foams (AFFFs) used at these sites (24). Given that FTSs have been shown to biodegrade to the

PFCAs (25, 26), the FTMAPs may represent a potential new source of PFCAs to humans.

Perfluorophosphonates (PFPAs) and perfluorophosphinates (PFPiAs) are fluorinated

surfactants used as leveling and wetting agents in waxes and coatings, and as defoaming agents

in pesticide formulations (27, 28). However, in 2006, these chemicals were delisted as

210

ingredients allowed in pesticide formulations in the U.S. and effectively banned from this

application starting in 2008 (29). Widespread contamination of PFPAs was observed in 80% of

Canadian surface waters and WWTP effluents sampled (30). The C6/C6 and C6/C8 PFPiAs

were also detected at ~2 ng/g concentrations in a WWTP sludge sample (31). Oral gavage

experiments revealed that the elimination half-lives for these chemicals in rats (1-9 days) may

potentially translate to significant half-lives in humans (31). Given their prevalence in the

environment, there is the potential for human exposure to PFPAs and PFPiAs.

Commercial products are largely comprised of fluorinated polymers and/or surfactants

with percent quantities of residual PFSAm or fluorotelomer starting materials present (11, 27, 32,

33). In contrast, PFCAs and PFSAs have only been observed as trace (ppb) contaminants in

commercial products (12, 34). Previous analyses of the total extractable organofluorine fraction

in human blood revealed that known fluorinated chemicals, such as the PFCAs and PFSAs, may

not fully account for the total contamination observed (35). This suggests the presence of other

unidentified fluorinated chemicals. In this study, fifty North American blood samples were

analyzed for forty different fluorinated analytes that included commercial fluorinated surfactants,

residual materials, degradation intermediates, and final PFCAs and PFSAs degradation products.

This investigation is the first to examine human sera for the SAmPAP, one FTMAP congener,

PFPAs, and PFPiAs.

7.3 Materials and Methods

7.3.1 Chemicals


Information (SI) in Appendix E. Structures, full names, and acronyms of the target analytes are

shown in Table 7.1. DiPAPs (y = x) and 6:2 FTMAP were synthesized by methods described in

Appendix E. Due to a lack of authentic standards for the PFPiAs at the time of analysis, the

Masurf® 780 technical product was used for quantitation. Purity of the synthesized diPAPs (y =

x) and the chemical composition of the Masurf® were determined using analytical standards (6:2,

8:2, 10:2 diPAPs ; C6/C6, C6/C8, and C8/C8 PFPiAs) that became available after the analysis of

all samples, as described in Appendix E

211

Table 7.1. Structures, full names, and acronyms of the target analytes.

Fluorinated Precursors

Polyfluoroalkyl

phosphate diester

(diPAP)

x = 4, 6, 8, 10

y = x or x+2

If y=x, x:2 diPAP

If y=x+2, x:2/y:2 diPAP

6:2 fluorotelomer

mercaptoalkyl phosphate

diester

(6:2 FTMAP)

N-ethylperfluorooctanesulfonamidoethanol-

based polyfluoroalkyl phosphate diester

(SAmPAP)

Fluorinated Intermediates

If R=H, perfluorooctanesulfonamidoacetate (FOSAA)

If R=CH3, N-methyl perfluorooctanesulfonamidoacetate (N-MeFOSAA)

If R=CH2CH3, N-ethyl perfluorooctanesulfonamidoacetate (N-EtFOSAA)

Fluorotelomer sulfonate

(x:2 FTS)

x = 4, 6, 8

Perfluorinated Acids


(Cx PFPA)

x = 6, 8, 10


(Cx/Cy PFPiA)

x = 6, 8

y = 6, 8, 10, 12

x+y ≤ 18


(PFCA)

x = 4–14

Perfluorosulfonate

(PFSA)

x = 4, 6, 8, 10

212

7.3.2 Sera Samples

Fifty human sera samples were obtained from Golden West Biologicals, Inc. (Temecula,

CA). The samples were collected in the U.S. in 2009 from donors varying in age (18–70 years

old) and gender. Twenty samples were from individual male donors and twenty from individual

female donors. The remaining ten 2009 samples were pooled samples in which each sample

pool consisted of at least ten individual donors, with no overlap in donors between each pooled

sample. The rationale for analyzing both sample types is discussed in Appendix E. Calf serum

was purchased from Sigma Aldrich (Oakville, ON) for use as a recovery matrix. Sera samples

were stored at -20oC prior to extraction. A human serum standard reference material (SRM

1957: Organic Contaminants in Non-Fortified Human Serum) was obtained from the National

Institute of Standards and Technology (NIST) and analyzed for quality control.

7.3.3 Extractions and Instrumental Analysis

The sera samples (2–3 mL) were extracted using modified versions of the ion-pairing

method developed by Hansen et al. (1). Detailed extraction procedures, chromatographic

gradients, instrumental conditions, and multiple reaction monitoring (MRM) mass transitions are

provided in Appendix E.


The C4-C14 PFCAs, C4, C6, C8, and C10 PFSAs, perfluorooctanesulfonamidoacetate

(FOSAA), N-methyl perfluorooctanesulfonamidoacetate (N-MeFOSAA), and N-EtFOSAA were

quantified using mass-labelled internal standards (Table E2 in Appendix E). The diPAPs (y = x),

PFPAs, PFPiAs, FTSs, SAmPAP, and 6:2 FTMAP were quantified by standard addition as no

internal standards were available at the time of analysis. As no standards were synthesized for

the mixed diPAPs (y = x + 2), they were quantified as described previously (16), in which the

standard additions of the adjacent y = x diPAPs were used as matrix-matched standards.

Chromatograms of a standard addition analysis of a human sera sample for the PFPiAs are

provided in Appendix E.

213

Spike and recovery experiments were performed in triplicate by adding 1 ng of each of

the target analytes into the calf serum recovery matrix, and the samples were extracted and

analyzed as described in Appendix E. Analyte recoveries were corrected for background

concentrations present in the unspiked matrix and ranged from 71 to 125% (Table E3ab in

Appendix E). Of the 36 analytes measured, ~80% of the recoveries were within 10% of the

spiked concentrations. The reported concentrations in the human sera samples were not

corrected for recovery.

The limits of detection (LOD) and limits of quantitation (LOQ) were defined as the

concentrations producing a signal-to-noise (S/N) ratio of equal to or greater than 3 and 10

respectively. The method LOD and LOQ values for each analyte are listed in Table E3ab in

Appendix E. Values below the LOD were reported as nondetect (nd). For the purposes of

calculating means, values below the LOD were assigned a value of zero and values below the

LOQ were used unaltered. All reported concentrations are presented as arithmetic means with

standard error.

Each human sera sample was extracted in duplicate with one procedural blank (HPLC

grade water) extracted in company to each sample (n = 50). The procedural blanks (n = 50) were

analyzed to check for contamination during the extractions. The average relative standard errors

for the duplicate analysis of sera samples observed at concentrations above the analyte-specific

method LOQ were in the range of 11-36% for analytes quantified by standard addition and 3-

24% for analytes quantified using internal standards. Few analytes were detected in the blanks,

and when detected, their concentrations were consistently below the analyte-specific LOQs, with

the exception of PFHxA (0.011±0.007 µg/L), PFHpA (0.008±0.004 µg/L), and PFOA

(0.011±0.004 µg/L). The sera concentrations for these analytes are at least one order of

magnitude higher than their corresponding LOQs. Analysis of methanol rinses of items used for

blood collection by the commercial supplier showed no contamination by any of the target

analytes, except for PFOA and 6:2 diPAP, which were observed at concentrations below their

corresponding instrumental LOQs. Details of the rinse procedure are described in Appendix E.

The methods used in this study were evaluated by analyzing the NIST SRM 1957 serum

sample and comparing the concentrations of the C7-C11 PFCAs, PFHxS, and PFOS to those

reported on the certificate of analysis and in an interlaboratory study (36). The SRM sample was

also analyzed for the full suite of analytes monitored in this study and the data set is provided in

214

Table E4ab in Appendix E. The percent errors of the concentrations measured in the serum

SRM were <15%, with the exception of PFOS (35%). Although the concentration of PFOS

observed in this study was lower (13.7±0.8 µg/L) as compared to the NIST value (21.1±1.2

µg/L), the relative standard error of the replicate analysis (n = 4) was low (6%). Here, the

chromatographic peak area corresponding to only the linear isomer of PFOS was integrated to

match the use of a linear PFOS standard for quantitation, whereas, the NIST approach also

included the branched isomers in their integration. The NIST values are concentrations of total

PFOS isomers and should be higher than the concentrations of linear PFOS only.

7.3.5 Statistical Analysis

For all statistical tests, any concentrations below the LOD were imputed as the LOD

divided by the square root of two. All tests were performed using StatsDirect (Version 2.7.8,

Cheshire, UK). A p-value of 0.05 was chosen as the criterion for statistical significance in all

analyses. A summary of the descriptive statistics calculated for all detected analytes is provided

in Table E6a-e in Appendix E. Further details describing the test results are provided in

Appendix E and in Table S5-9 in Appendix E.


7.4.1 Concentrations in human sera

The 6:2 diPAP was detected at mean concentrations of 0.13±0.04 µg/L in all of the

pooled sera samples and 0.072±0.015 µg/L in about 80% of the single donor samples (Figure

7.1, Table E6a in Appendix E). The 6:2/8:2 diPAP (0.049±0.019 µg/L, pooled; 0.035±0.009

µg/L, single donor) and 8:2 diPAP (0.13±0.04 µg/L, pooled; 0.11±0.05 µg/L, single donor) were

also detected, but less frequently (30–60%) (Figure 7.1, Table E6a in Appendix E). The 4:2 and

4:2/6:2 diPAPs were detected in <20% of the samples, while 10:2 diPAP was not detected at all.

The distribution of diPAPs observed here is similar to the profile reported for human sera

samples collected in 2008 (16), where the 6:2, 6:2/8:2, and 8:2 diPAPs were found to be the most

prevalent congeners. However, the concentrations of 6:2 and 6:2/8:2 diPAPs measured here

were significantly lower than previous measurements in pooled sera samples collected in the

period of 2004–2005 and in 2008 (Mann–Whitney U test, p<0.05), while no significant change

215

was observed between the concentrations of 8:2 diPAP measured here and those in the 2008

samples (Mann–Whitney U test, p=0.78) (16).

Contamination-driven issues precluded the detection of 8:2 diPAP in any of the 2004-

2005 human sera samples (16), but a downward temporal trend is likely given the decrease

observed for the 6:2 and 6:2/8:2 diPAPs, as well as the disappearance of the other diPAP

congeners (4:2, 4:2/6:2, 8:2/10:2, and 10:2 diPAPs), that were previously detected in the

2004/2005 samples (16), from the 2008 samples (16) and the 2009 samples measured here.

Although PAPs are used as greaseproofing agents in food packaging materials (11), human

exposure to PAPs may also result from their incorporation into personal care and cosmetic

products (32, 37). While the levels of diPAPs in humans may be declining, it is important to

note that even low-level exposure from day-to-day contact with all of these different PAPs-based

products may still result in PFCA contamination in humans (19).

The SAmPAPs and FTMAPs were also applied as greaseproofing agents in paper and

paperboard used for food packaging (11–14), but unlike the diPAPs, their potential as PFCA

precursors is relatively unexplored. As commercial SAmPAP formulations predominantly

consisted of the diester (85%), followed by the mono- (10%) and the tri- (5%) esters (14), the

diester was monitored, but was not detected. Considering the span of 10 years since the phase-

out of these chemicals in North America, exposure to SAmPAPs is expected to decline, although

exposure may still occur via use of products purchased before the phase-out. Continued

exposure may also occur in certain European and Asian countries where POSF-based production

remains active (38, 39). The SAmPAP is currently commercially available in China (40).

216

Figure 7.1. Arithmetic mean concentrations and standard error (µg/L) for all target analytes detected in >20% of the single donor and

pooled human sera samples (plotted on a logarithmic scale). Note: Analytes denoted with an asterisk (*) were detected in <20% of the

samples, i.e. PFPeA (pooled); PFBS (single donor).

Co

nce

ntr

ati

on

of

An

aly

tes (

g/L

)

0.001

0.01

0.1

1

10

* *

diPAPs

FOSAA

N-MeFOSAA

N-EtFOSAA

6:2, 8:2 FTS

C6/C6,

C6/C8

PFPiAsPFCAs

PFSAs

6:2 diPAP

6:2/8:2 diPAP

8:2 diPAP

FOSAA

LEGEND

N-MeFOSAA

N-EtFOSAA

8:2 diPAP 6:2 FTS

FOSAA 8:2 FTS

C6/C6 PFPiA

C6/C8 PFPiA

PFBA

PFPeA

PFHxA

PFHpA

PFOA

PFNA

PFDA

PFUnA

PFBS

PFHxS

PFOS

PFDS

Pooled (n = 10)Single donor (n = 40)

217

Previous investigations by 3M revealed that in vitro incubations of the SAmPAP

monoester with rat and human hepatocytes resulted in the production of N-EtFOSE and other

metabolites, like N-EtFOSAA, FOSAA, and PFOS (41). In this study, the FOSAA and N-

EtFOSAA metabolites were detected at mean concentrations in the range of 0.050–0.069 µg/L,

which are about 1 to 2 orders of magnitude lower than previous measurements in human sera (2,

4–6) (Figure 7.1, Table E6b in Appendix E). This decline is consistent with that observed for N-

EtFOSAA and N-MeFOSAA in American Red Cross blood samples collected between 2000 and

2006 (6). During an investigation on dietary exposure to the SAmPAP, Tittlemier et al. observed

a similar decline in the levels of N-ethyl perfluorooctanesulfonamide (N-EtFOSA) in food

samples collected between 1992 and 2004 where concentrations peaked in 1998, followed by a

yearly decline until 2002, the year of the final phase-out of POSF-based materials, after which no

more detects were observed (42). It is important to note that the source of these chemicals in

human sera is not limited to food packaging, but may also include inhalation of volatile PFSAm-

based precursors offgassing from other non-food contact applications (33). N-

methylperfluorooctanesulfonamidoethanol (N-MeFOSE) was typically polymerized with

urethane, acrylate, and/or adipate monomers and used as stain repellants in textiles, personal

apparel, and home furnishings, with some application as protectors for food packaging (14).

Considering the extensive use of these chemicals in the indoor environment, it is not

surprising that the indoor air concentrations of N-MeFOSE and N-EtFOSE typically exceeded

the levels observed in outdoor air by up to 2 orders of magnitude (43, 44). Compared to FOSAA

and N-EtFOSAA, higher concentrations of N-MeFOSAA (0.44±0.11 µg/L, pooled; 0.36±0.07

µg/L, single donor) were observed in the human sera samples (Mann Whitney U test, p<0.0001)

(Figure 7.1, Table E6b in Appendix E). This distribution is mirrored in the higher concentrations

of N-MeFOSE (1500–2600 pg/m3) observed in the indoor air of Canadian homes as compared to

N-EtFOSE (740–770 pg/m3) (43, 44). A report from 3M indicated that historical production of

N-MeFOSE-based polymers exceeded that of the N-EtFOSE surfactants by weight (14),

although higher use patterns and presumably, higher stability of the N-MeFOSE-based polymers

may also account for the dominance of N-MeFOSAA observed in human sera as compared to

FOSAA and N-EtFOSAA.

The 6:2 FTMAP was not detected in any of the human sera samples. Both the diPAPs

and FTMAPs are approved food contact additives and regulated by the U.S. Food and Drug

218

Administration (FDA) (11). However, since the late 1990s, Ciba Specialty Chemicals Co. has

discontinued their LodyneTM

line of perfluoroalkylthio-based surfactants (22). As of 2004,

Chemguard (Mansfield, TX) has acquired the business rights to sell the remaining stockpile of

the discontinued LodyneTM

fluorosurfactant series from Ciba (22). Although they are still

commercially available according to a recent Ciba 2007 product guide (21), it is unclear where

and in what capacity these chemicals are being used in North America. The FTMAPs were

observed at high concentrations (1.4–3.9 µg/g) in microwaveable popcorn purchased in the U.S.,

although the sample ages were not discussed (13). No published literature on the fate of

FTMAPs is known, but one possible degradation pathway is cleavage of the carbon-sulfur bond

to release the fluorotelomer appended to the cyclic phosphate moiety, which upon further

oxidation, may yield either FTOH or FTS depending on which side of the sulfur atom the

cleavage occurs.

The 8:2 FTS was the dominant congener observed in human sera (<LOD (0.005 µg/L)–

0.231 µg/L; >95% of the samples), followed by 6:2 FTS (<LOD (0.005 µg/L)–0.047 µg/L;

>54%) and 4:2 FTS (<LOD (0.005 µg/L)–0.018 µg/L; <20%) (Figure 7.1, Table E6b in

Appendix E). Only one other study detected 6:2 FTS and 8:2 FTS in pooled human sera and

plasma collected in 2002 (45). The concentrations observed in that study (<LOD–0.109 µg/L)

do not differ significantly from those observed in the pooled samples here (Mann-Whitney U

test, p>0.05, Table E8 in Appendix E). The 6:2 FTS is marketed as a wetting and/or foaming

agent in commercial products (46), but it has also been identified as a major constituent (~1600

µg/L) in some AFFF formulations (24), and a proposed breakdown product of the active

materials in AFFF, the fluorotelomerthiol-based polyacrylamides (47, 48). Predominance of 6:2

FTS contamination in groundwater collected near sites of high AFFF use (24) was consistent

with the high purity of 6:2 fluorotelomer surfactant (>99.5%) in most AFFF formulations, with

the remaining ~0.5% comprised of the 8:2 and higher homologues (48). While exposure to

AFFF surfactants may partially account for the FTS contamination observed here, it seems an

unlikely source to which the general population would be chronically exposed. The observation

of different perfluoroalkyl chain lengths of FTS in human sera here is consistent with exposure

to fluorotelomer-based products. The sources of this contamination may include exposure to

commercial products containing the FTS themselves, or to other fluorotelomerthiol-based

products, such as FTMAPs.

219

7.4.2 Detection of a new perfluorinated acid in human sera

Using the recently released analytical standards of three PFPiA congeners from

Wellington Laboratories, the percent composition of the Masurf® 780 was determined to be

36.9±0.1% C6/C6 PFPiA, 33±6% C6/C8 PFPiA, and 27±3% C8/C8 PFPiA (Appendix E). This

percent composition was used to adjust the concentrations of C6/C6, C6/C8, and C8/C8 PFPiAs

reported here (Figure 7.1, Table E6c in Appendix E). The detection of the remaining PFPiA

congeners (C6/C10, C8/C10, C6/C12) will be briefly discussed.

A suite of six PFPiA congeners were detected for the first time in human sera. The mean

concentrations of the two most prevalent congeners (C6/C6 and C6/C8 PFPiAs) ranged from

0.004 to 0.038 µg/L (>50% of the samples) (Figure 7.1). The C6/C10 and C6/C12 PFPiAs were

also detected in >40% and <20% of the samples respectively. Detection of the C8/C8 and

C8/C10 PFPiAs was infrequent (5–10%), and in the case of detects for C8/C8 PFPiA, the

concentrations were significantly lower than those of the other PFPiAs (Mann Whitney U test,

p<0.05). The predominance of the perfluorohexyl-based (C6) PFPiAs is consistent with

previous detection of only the C6/C6 and C6/C8 PFPiAs in WWTP sludge (31). It is unclear

whether this distribution is an artifact of the composition used in commercial products or some

unique aspect of human pharmacokinetics that would result in preferential uptake of the C6

congeners. A significant correlation among the concentrations of the C6-based PFPiA congeners

was observed in both the single donor and pooled sera samples (Spearman’s rank correlation, r

=0.48–0.83, p<0.05, Table E9 in Appendix E). This suggests that human exposure to the PFPiAs

may derive from a common source.

The C6, C8, and C10 PFPAs were not detected in this study. This contrasts the

widespread contamination observed for these chemicals in surface waters and wastewaters (30).

Intraperitoneal dosing of rats with Masurf® 780 demonstrated higher urinary (≤96% of the dose)

excretion of the PFPAs, as compared to PFPiAs (0%) (31). Together with the shorter half-lives

observed for the PFPAs (~1-3 days) as compared to the PFPiAs (~2-9 days) in rats (31), these

results suggest faster excretion kinetics of PFPAs, which may account for their lack of detection

in human sera here. Furthermore, analysis of paired rat whole blood and plasma samples

collected in the same study demonstrated PFPAs may bind to cellular components in whole

blood (31). As such, PFPA contamination in humans may be underestimated if plasma or sera

samples were analyzed, as was done in this study, instead of whole blood.

220

The concentrations of C6/C6 and C6/C8 PFPiAs appear to be greater in males as

compared to females, although the difference was only significant at a higher significance level

of p=0.10 (Mann-Whitney U test, p<0.10 (one-sided), Table E7 in Appendix E). The only other

analyte that showed significantly higher concentrations in males as compared to females was

PFOA (Mann-Whitney U test, p=0.0061 (one-sided), Table E7 in Appendix E). This is

consistent with other studies in which serum concentrations of PFOS and PFOA were observed

to be higher in males than in females (3–6), although no statistical difference has been reported

(7). Variations in geographical location, lifestyle, exposure pathways, and pharmacokinetics,

may contribute to the gender-associated differences observed in the occurrence of perfluorinated

acids in humans.

Consistent with other human sera measurements (1–7), PFOS was present at the highest

concentrations (4.44±0.46 µg/L, pooled; 12.26±3.79 µg/L, single donor), followed by PFOA

(1.76±0.31 µg/L, pooled; 2.00±0.18 µg/L, single donor) and PFHxS (1.19±0.18 µg/L, pooled;

1.25±0.20 µg/L, single donor). PFBA, PFPeA, PFHxA, and PFBS were also observed at

concentrations ranging from <LOD (0.001–0.005 µg/L) to 0.073 µg/L. These short chain

perfluorinated acids are typically not monitored in human sera analysis, but in the case of

detection, the concentrations are usually below or close to the LOQ (5–7, 49). Despite their

rarity in humans, monitoring for the short chain PFCAs and PFSAs is necessitated by the shift in

fluorochemical manufacturing processes to the perfluorobutyl- and perfluorohexyl-based

chemistries. No correlations were observed between the PFPiAs and any of the diPAPs, PFCAs,

and PFSAs; therefore, humans may be exposed to the PFPiAs via different exposure sources.

7.5 Current state of knowledge concerning exposure to commercial fluorinated

chemicals.

Following the discovery of diPAPs in human sera (16), the detection of PFPiAs observed

here represents the second observation of a commercial fluorinated product and the first

observation of this class of perfluorinated acids in human sera. Unlike the PFCAs and PFSAs,

there are no known PFPiA precursors in production. Exposure to these chemicals may occur

through day-to-day use of common household products, such as carpet and upholstery cleaners,

and cleaning fluids for the bathroom (27). Ingestion of foods cultivated in the presence of

pesticides containing PFPiAs is not expected to be a major source as the use of these chemicals

in pesticide formulations has been discontinued (29). The observation here of PFPiAs in human

221

sera and previous observations of PFPAs in wastewaters (30) and PFPiAs in WWTP sludge (31)

are evidence of human exposure to these chemicals.

Despite the percentage quantities of chemicals (i.e. diPAPs, SAmPAPs, FTMAPs,

PFPAs, PFPiAs) typically present in commercial products (11, 27, 32), PFOS, PFHxS, and the

longer chain PFCAs were the major fluorinated contaminants observed here. This disparity in

sera concentrations may be due to differences in pharmacokinetic behavior and/or differences in

metabolic fates among these different classes of chemicals. Uptake of diPAPs and their

subsequent metabolism to produce the biologically persistent PFCAs have been demonstrated in

rats (18, 19). If FTMAP can biologically degrade to the FTS, as hypothesized above, this

chemical may potentially be a new fluorotelomer-based source of PFCAs, as biotransformation

of 6:2 FTS was recently demonstrated to produce the C4-C6 PFCAs in a microbial system (26).

Exposure to PFPAs and PFPiAs are limited to direct sources and their prevalence in commercial

applications is unknown. Exposure to commercial fluorinated chemicals is also not well

understood in other locations, although legislated efforts to characterize their sources are

underway in Europe (50). Certain European and Asian populations may still be exposed to

POSF-based materials, such as SAmPAP (38–40), while the diPAPs and FTMAPs are approved

for use in food contact materials in Germany (51).

Despite the low concentrations (sub ppb) of diPAPs, FTSs, and PFPiAs observed here,

their presence in human sera provides direct evidence of human exposure to commercial

fluorinated products. The absence of SAmPAP and 6:2 FTMAP observed here also does not

preclude previous exposure. The paucity of pharmacokinetic and toxicological data on these

chemicals prevents assessment of their persistence in humans and potential health risks at the

levels currently observed in blood. We believe a comprehensive evaluation of human sera

fluorochemical contamination is necessary to properly characterize human exposure. Here, we

report a pilot set of human sera data on a small North American population’s exposure to a suite

of fluorinated chemicals that range from those present as active or residual materials in products

to their potential degradation intermediates and products.


The authors would like to thank Susanne Waaijers (University of Amsterdam,

Netherlands), Alexandra Tevlin and Barbara Weiner (University of Toronto, Toronto, ON) for

222

synthesizing the diPAPs and 6:2 FTMAP, Timothy Begley (U.S. FDA, College Park, MD) for

providing the SAmPAP standard, Amila De Silva (Environment Canada, Burlington, ON) for

providing the analytical diPAP standards, Wellington Laboratories (Guelph, ON) for donating

native and mass-labelled internal standards, and Tennessee Blood Services Corp. (Memphis, TN)

for donating blood collection items. This research was funded by the Natural Science and

Engineering Research Council of Canada (NSERC), the Ministry of the Environment Best in

Science grant and a NSERC Postgraduate Scholarship (PGS) awarded to H.L.







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Fluorochemicals in Human Blood Samples from China. Environ. Sci. Technol. 2006, 40,

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227

CHAPTER EIGHT

Summary, Conclusions, and Future Work


Contributions: Holly Lee prepared this chapter with additional comments provided by Scott

Mabury

228

8.1 Summary and conclusions

This thesis explored the fate of commercial fluorinated surfactants as potential sources to

the fluorochemical contamination currently observed in humans, wildlife, and the environment.

Anthropogenic discharges of domestic and industrial origin have been identified as major

sources of perfluoroalkyl and polyfluoroalkyl substances (PFASs) to wastewater treatment plant

(WWTP) environments. Elucidation of the post-WWTP fate of commercial fluorinated materials

and their degradation products is of particular interest to this work. Specifically, the interest in

elucidating the diverging pathways of PFAS transport to agricultural farmlands via biosolids

application and surface water environments have established two branches of study in this work.

Biological (i.e. biotransformation and bioaccumulation) and environmental (i.e. soil-plant uptake

and sorption) processes were investigated at the molecular level in an effort to understand how

these processes contribute to the fluorochemical burden observed in the relevant environmental

compartments. A specific focus of this work was the role of phosphorus-based commercial

fluorinated surfactants, the polyfluoroalkyl phosphate esters (PAPs), the

perfluoroalkylphosphonates (PFPAs), and the perfluoroalkylphosphinates(PFPiAs) as sources of

perfluoroalkyl acids (PFAAs) in the environment.

This connection was first investigated in the environmental degradation of PAPs in the

presence of WWTP microbes, as presented in Chapter 3. PAPs are primarily used as

greaseproofing agents in food packaging (1, 2), although they may also be found in a wide

variety of other products (3–7). For oil repellency applications, commercial greaseproofing

formulations are primarily composed of the di-fluoroalkylated PAP congener (diPAP) due to its

high efficiency (2), while the mono-fluoroalkylated congener (monoPAP) may be present as

byproducts. The microbial fate of monoPAPs and diPAPs was investigated through

biodegradation experiments in a WWTP-simulated system. Headspace analysis of the

experimental system revealed production of fluorotelomer alcohols (FTOHs) of varying

perfluoroalkyl chain lengths that corresponded to the hydrolysis of their parent PAP phosphate

ester linkages.This suggests PAPs can undergo microbially-mediated hydrolysis to produce

FTOHs, which are established PFCA precursors in microbial systems (8–14), although the

production yields of FTOHs from PAPs are quite low (1–5%).Analysis of the aqueous phase also

revealed production of the perfluoroalkyl carboxylates (PFCAs). The majority of the

intermediate metabolites and final PFCA products observed were consistent withthe metabolite

229

profiles typically produced from a β-oxidation-like mechanism, as was previously observed in

the degradation of otherfluorotelomer-based precursors (8, 10, 11, 14). However, the detection

of odd-chain PFCAs (i.e. 6:2 diPAP/6:2 monoPAPperfluoroheptanoate (PFHpA, C7)) also

suggests other pathways may play a minor role in PFCA production. This study is the first to

establish a connection between a commercial product and the production of PFCAs within

WWTPs. This observation, together with the diPAP concentrations previously observed in

WWTP sludge (15), suggest PAPs-containing commercial products may contribute to the

increased mass flows of PFCAs often observed between WWTP influents and effluents (16).

In addition to WWTP sludge, diPAPs have also been found at hundreds of ng/g

concentrations in other anthropogenic waste materials, such as paper fiber solids (15). In

Chapter 4, a series of greenhouse biosolids-applied soil-plant microcosm was used to simulate

the fate of PFASs that may be present in WWTP and paper fiber biosolids in farmlands that have

been amended with these waste materials. Higher concentrations of diPAPs and PFCAs were

observed in biosolids-amended, as compared to control soils that were not amended with any

biosolids. Soil biodegradation of diPAPs was examined using 6:2 diPAP as the test parent

reactant and its observed degradation to the corresponding fluorotelomer intermediates and

PFCAs was consistent with the metabolite profiles previously observed in Chapter 3 and those

described for 6:2 FTOH degradation in soil (13, 14). Plant uptake of diPAPs and PFCAs from

the biosolids-applied soils was also observed, with preferential accumulation of the short-chain

PFCAs (C4–C6) observed. This pattern is consistent with the predominance of short-chain

PFCAs (<C8) previously observed in grass samples collected from farmfields that were

demonstrated to be highly contaminated with PFASs from biosolids application(17). This work

is the first to demonstrate the biodegradation pathway ofdiPAPs to PFCAs in soil and alsothe

subsequent uptake of these chemicals and their metabolites in plants. The plant accumulation of

PFCAs observed here and previously by others (17–19) has major implications for potential

migration of these chemicals and other PFASs intoterrestrial food chains.

WWTP effluents containing anthropogenic discharges of commercial fluorinated

surfactants may further contaminate downstream aqueous environments. PFPAs and PFPiAs are

newly discovered PFAAs that have recently been detected in WWTP effluents and sludge (20,

21), surface water (20), and fish (22). Chapters 5 and 6 examined how sorption and

bioaccumulation influenced the partitioning of these chemicals between aqueous media and an

230

environmental solid and between aqueous media and biological tissues respectively. In Chapter

5, the C6, C8, and C10 monoalkylated PFPAs and C6/C6, C6/C8, and C8/C8

dialkylatedPFPiAswere observed to sorb to seven different soils at varying degrees. The organic

carbon-normalized distribution coefficients, logKOCs, were observed to increase with the number

of perfluorinated carbons present in the PFPAs and PFPiAs, consistent with the trend that has

been previously demonstrated in the sorption of PFCAs and perfluoroalkanesulfonates (PFSAs)

to sediments (23) and soils (24). Comparison oflogKOCvalues measured in this study with those

reported for other PFAAs(23) revealed similar sorption capacity between the PFSAs and PFPiAs

to environmental solids, while the PFCAs and PFPAs were generally less sorptive than the

PFSAs of equal perfluoroalkyl chain length. The PFPAs desorbed more rapidly and to a greater

extent from soil, as compared to the PFPiAs, which suggests the PFPAs are more prone to

remobilization into the aqueous environment, while the more sorptivePFPiAs are likely to

remain bound in environmental solid phases. These results represent the first set of sorption data

for PFPAs and PFPiAs and they aregenerally consistent with the observed distribution of these

chemicals in environmental media. The PFPAs have been detected in surface waters (20), but

not in WWTP sludge and sediments (25). Conversely, the PFPiAs have been measured at ng/g

concentrations in WWTP sludge(21), but have yet to be monitored for in any aqueous media.

PFPiAs have also been observed at low pg/g concentrations in lake trout, whereas the

PFPAs were not detected. This suggests bioaccumulation of these chemicals may also be

governed by structural features, as was observed in their sorption to soils. In Chapter 6, juvenile

rainbow trout (Oncorhynchusmykiss) were separately exposed to a mixture of C6, C8, and C10

PFPAs and a mixture of C6/C6, C6/C8, and C8/C8 PFPiAs in the diet. Depuration half-lives

ranged from 4 to 5 days for the PFPAs and 6 to 53 days for the PFPiAs and were observed to

increase with the number of perfluorinated carbons present in the chemical. Biomagnification

factors (BMFs) were calculated from whole-body homogenate concentrations as opposed to

blood or liver concentrations, as tissue analyses revealed preferential accumulation of all target

analytes in the blood and liver, as was previously observed in other studies (26–28). The

calculated BMFs were lower than those previously determined in rainbow trout exposed to

PFCAs and PFSAs also via the diet (29). In general, the PFSAs were the most bioaccumulative

in rainbow trout, followed by PFCAs, and then PFPAs of equal perfluoroalkyl chain length,

which suggests bioaccumulation may be dependent on the charge of the polar headgroup. Unlike

231

the other singly charged PFAAs studied, PFPAs predominate as dianions at environmental pH

and are likely more water soluble and less inclined to assimilate into biological tissues, as

compared to the other PFAAs. The metabolism of PFPiAs to the corresponding PFPAs was also

the first observation of an in vivo cleavage of the carbon–phosphorus bond in fish and the first in

vivo metabolism of a PFAA observed anywhere. Similar to the BMFs measured for the PFCAs

and PFSAs in rainbow trout(29), the BMFs of PFPAs and PFPiAsalso did not exceed one, which

suggests PFAAs do not biomagnify from dietary exposure in juvenile rainbow trout.

Environmental circulation of commercial fluorinated surfactants through the various

processes discussed above may potentially result in human exposure. Human exposure may

occur through the consumption of vegetable crops grown and livestock raised on farmlands that

have been contaminated with PFASs via biosolids application. Alternatively, combined aquatic

and terrestrial biomagnification may result in the ultimate transfer of PFASs to humans at the

apex of these food webs, although in the case where contaminated seafood comprises part of the

diet of humans living near these food sources, direct consumption may represent an additional

mode of exposure (30).Direct exposure may also occur through the use of commercial

fluorinated products. Regardless of the relative contribution of these exposure pathways, human

contamination was evidenced by the detection of diPAPs and PFPiAs at ng/L concentrations in

fifty North American human sera samples, as shown in Chapter 7. Concentrations of the 6:2,

6:2/8:2, and 8:2 diPAPs were lower than those reported in human sera samples collected in 2004,

2005, and 2008 (15), which suggests diPAP levels in humans may be declining, possibly due to a

transition to other chemistries in commercial products. The C6/C6 and C6/C8 PFPiAs were also

observed for the first time in >50% of the human sera sampled, at concentrations of 4 to 38 ng/L.

As was observed in lake trout (22), none of the monitored PFPAs were detected in human sera,

which is consistent with their faster depuration kinetics previously observed in rainbow trout in

Chapter 6 and in rats (21). However, the analysis of PFPAs in serum may underestimate the

actual levels in whole blood as PFPAs may bind to cellular components in blood, which may

possibly result in their diminished presence in plasma or sera (21). Other commercial surfactants

that have never beenmonitored for in human blood or at least not extensively, such as the N-

ethylperfluorooctanesulfonamidoethyl phosphate diester (SAmPAP), 6:2

fluorotelomermercaptoalkyl phosphate diester (FTMAP), and fluorotelomersulfonates (FTSAs),

were also surveyed, but only the FTSAs were detected at concentrations as high as 231 ng/L.

232

Despite the low concentrations of diPAPs, FTSAs, and PFPiAs observed, their occurrence in

human sera is direct evidence of human exposure to commercial fluorinated products.

8.2 Future research directions

This thesis presents novel findings on the environmental chemistry of commercial

phosphorus-based fluorinated surfactants, including a number of processes that were observed to

play a role in circulating these chemicals among different environmental compartments. As the

diPAPs, PFPAs, and PFPiAs are considered emerging chemicals in the field of fluorochemical

research, future work should be directed towards further understanding of the environmental fate

of these novel PFASs, as well as, addressing various questions that were raised in this work.

The mechanism by which diPAPsfirst biotransformto their corresponding FTOHs,

followed by subsequent oxidation to the terminal PFCAs involves the production of various

intermediate metabolites, the fluorotelomer saturated (FTCAs) and unsaturated (FTUCAs)

carboxylates. Their occurrence during biological transformation of fluorotelomer precursors is

of particular concern, as both FTCAs and FTUCAs have been found to be orders of magnitude

more toxic than the corresponding PFCAs (31) in aquatic biota. Not monitored in this work

were the fluorotelomer saturated (FTALs) and unsaturated (FTUALs) aldehydes, both of which

are also intermediates of FTOH metabolism. The demonstrated affinity of these intermediate

metabolites to small biological nucleophiles and proteins (32–35) may contribute a significant

portion, in addition to the generally low yields from metabolite production, to the mass balance

in fluorotelomer biotransformation. However, the relative importance of this pathway is

dependent on the extent of intermediate production during the biotransformation of commercial

fluorotelomer materials, which has not been extensively studied due to the analytical challenges

involved in detecting and quantifying these transient intermediates. Future conjugate and/or

protein binding studies should focus more on using commercial fluorinated materials, such as the

diPAPs, instead of the intermediates as the parent reactantsas that would better represent human

exposure to fluorinated chemicals through the use of consumer products.

The low metabolite yields observed for the biotransformation of diPAPs in Chapter 3 and

other fluorotelomer-based precursors may also be attributed to sorption of the parent reactant

and/or intermediate metabolite to the incubation medium (i.e. soil, sludge). While the sorption of

PFCAs and PFSAs to environmental solids has been widely demonstrated (23, 24, 36, 37), only

233

one study has investigated the sorption behaviour of FTOH in soil to date (38), with no work

published on the sorption of other fluorotelomer-based precursors, including the diPAPs.

Evaluation of the sorption capacity of diPAPs in environmental solids is important as these

chemicals are present at significant concentrations in WWTP sludge (15) and may be transported

to soil during agricultural land application.

Biotransformation of 6:2 diPAP was observed in the greenhouse soil-plant microcosm,

although it is unclear whether the transformation was predominantly occurring in the soil or in

the plants as the parent reactant and its degradation metabolites were observed in both

compartments. Both pathways are likely contributors, but plant metabolism must be investigated

separately to assess its potential as a long term sink for PFASs that have accumulated in this

compartment. Hydroponics experiments are ideal for examining contaminant uptake and

metabolism within a plant system in the absence of soil influence.

Biomonitoring of aquatic and terrestrial food webs is also necessary to examine the

potential for PFPAs and PFPiAs to undergo trophic magnification. Despite their low BMFs (<1)

measured in rainbow trout, the PFPiAs have been detected in lake trout (22), the apex predator of

most aquatic food webs, and have a demonstrated capacity to sorb to environmental solids.

Benthic uptake of contaminated sediments may represent a potential route for these chemicals to

accumulate in benthic biota and subsequently undergo biomagnification up the food web, as was

observed for other PFAAs in an aquatic food web in Lake Ontario (39). Biomonitoring of

terrestrial animals, especially those located in near-source regions, is especially important

considering the extensive use of these chemicals in commercial products (40, 41). Terrestrial

food webs should be studied for PFPAs and PFPiAs, more so in temperate regions rather than

remote environments, as these chemicals have no known volatile precursors like the PFCAs and

PFSAs and therefore, would not be expected to occur significantly at locations far from emission

sources.

Given the PFPiAs can metabolize in fish, biotransformation studies in other systems are

necessary. Previous investigations of the pharmacokinetics of PFPAs and PFPiAs in rats

employed a technical mixture of both classes of these chemicals for dosing, which precluded any

elucidation of biotransformation pathways (21). Future experiments should employ separate

standards of the PFPAs and PFPiAs to probe their biological and environmental fate without the

influence from one another. Furthermore, the use of juvenile rainbow trout in Chapter

234

6precluded the study of gender differences in the biological handling of these chemicals. Gender

differences have been observed for the uptake and elimination of PFOS and PFOA in fathead

minnows (42, 43) and as such, future analysis of both male and female aquatic organisms may

help to elucidate the mechanisms involved in the differential pharmacokinetic behaviour

observed between these two genders.

Finally, human exposure to commercial fluorinated surfactants was discussed in Chapter

7, but the various pathways involved in this contamination remain widely debated. Both direct

and indirect sources have been discussed (44), but the relative importance of these sources to the

overall fluorochemical contamination observed remains uncertain. In addition, the majority of

human analysis has traditionally focused on the PFCAs and PFSAs with some focus on

intermediate metabolites, but little to no attention devoted to characterizing the actual fluorinated

chemicals applied in commercial products. Measurements of commercial fluorinated surfactants,

such as those surveyed in Chapter 7 and others, in human blood, as well as, the relevant products

in which they are incorporated (e.g. food contact materials, household products, personal care

products) would greatly assist in ascribing the relative importance of different exposure

pathways to the overall fluorochemical burden observed in humans.

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238

APPENDIX A

SUPPORTING INFORMATION FOR CHAPTER THREE

Biodegradation of Polyfluoroalkyl Phosphates (PAPs) as a Source of Perfluorinated Acids

to the Environment

239

LIST OF TABLES AND FIGURES

Figure A1. Diagram of the purge-and-trap system used in the purging control and

biodegradation experiments 246

Table A1. Description of bottles in the purging control experiment 247

Table A2. Description of experimental and control bottles in the biodegradation

experiments 248

Table A3. Internal standards, multiple reaction monitoring (MRM) mass transitions, and

recovery results for all target aqueous analytes 250

Table A4. Single ion monitoring (SIM) molecular ions, dwell time, and recovery results

for FTOHs in the headspace 252

Figure A2. Typical chromatograms of monoPAPs in standard and sample extract 253

Figure A3. Recoveries of PAPs in mineral media treated with mixed liquor 254

Table A5. Limits of detection (LOD) and limits of quantitation (LOD) for the target

analytes 255

Figure A4. Positve control to assess microbial viability during the 92-day

biodegradation 257

Figure A5. Results from the purging control experiment 258

Figure A6. Recovery of PAPs from water, septa, gas diffuser tubes, and bottles at the

end of the purging control experiment 259

Figure A7. PAPs in sterile controls during the 92-day biodegradation 260

Figure A8. Degradation of monoPAPs in chain length study 261

240

EXPERIMENTAL

Chemicals.

4:2 fluorotelomer alcohol (4:2 FTOH, 95%), 6:2 fluorotelomer alcohol (6:2 FTOH, 95%),

and perfluoropentanoic acid (PFPeA, 97%) were purchased from Oakwood Products, Inc. (West

Columbia, SC). 8:2 fluorotelomer alcohol (8:2 FTOH, 95%), 10:2 fluorotelomer alcohol (10:2

FTOH, 95%), and 7:3 fluorotelomer acid (97%, 7:3 FTCA) were purchased from SynQuest

Labs, Inc. (Alachua, FL). 4:2 fluorotelomer acid (4:2 FTCA) and 4:2 unsaturated fluorotelomer

acid (4:2 FTUCA) were synthesized by our group according to Achilefu et al. (1) with purity of

>95%. 6:2, 8:2, 10:2 fluorotelomer acids (6:2 FTCA, 8:2 FTCA, 10:2 FTCA, >98%), 6:2, 8:2,

10:2 unsaturated fluorotelomer acids (6:2 FTUCA, 8:2 FTUCA, 10:2 FTUCA, >98%), and

perfluorobutanoic acid (PFBA, 98%) were donated from Wellington Laboratories (Guelph, ON).

Perfluorohexanoic acid (PFHxA, >97%), perfluoroheptanoic acid (PFHpA, 99%), phosphorus

(V) oxychloride (POCl3, 99%), tetrabutylammonium hydrogen sulfate (TBAS, 99%), and

sodium carbonate (Na2CO3, >99.5%) were purchased from Sigma Aldrich (Oakville, ON).

Perfluorooctanoic acid (PFOA, 96%), perfluorononanoic acid (PFNA, 97%), perfluorodecanoic

acid (PFDA, 98%), and perfluoroundecanoic acid (PFUnA, 95%) were purchased from Aldrich

Chemical Co. (Milwaukee, WI). Triethylamine (TEA) was purchased from ACP Chemicals, Inc.

(Montreal, QB). Anhydrous ethyl alcohol was purchased from Commercial Alcohols, Inc.

(Brampton, ON). DriSolv® tetrahydrofuran (THF) was purchased from EMD Chemicals, Inc.

(Gibbson, NJ), while methanol (Omnisolv, >99%), water (Omnisolv, >99%), methyl-tert-butyl

ether (MTBE, Omnisolv, >99%), formic acid (Omnisolv, 98%), and ammonia (30%) were

purchased from EMD Chemicals, Inc. (Mississauga, ON).

Mass-labeled internal standards were donated from Wellington Laboratories (Guelph,

ON) and they included 13

C2-PFHxA (>99%), 13

C4-PFOA (>99%), 13

C5-PFNA (>98%), 13

C2-

PFDA (>98%), 13

C2-FHUEA (6:2 FTUCA, >99%), 13

C2-FOUEA (8:2 FTUCA, >98%), and 13

C2-

FDUEA (10:2 FTUCA, >98%).

Purging control experiment.

From a mixed standard of 4:2, 6:2, 8:2, and 10:2 monoPAPs at 100 ugmL-1 and a

standard of 6:2 diPAP at 660 ugmL-1, 4 mL (400 ug) and 0.6 mL (400 ug) respectively were

241

spiked in duplicate into 500 mL polypropylene bottles (Nalgene®, VWR International, Toronto,

ON), sealed with in-house drilled caps and septa to fit 100mg Orbo Amberlight XAD-2

cartridges (Supelco, Bellefonte, PA) and gas diffuser tubes (Pyrex, VWR International Ltd.,

Mississauga, ON), containing 400 mL of 18Ω Milli-Q deionized water. Upon spiking, one set of

bottles were purged with carbon-filtered in-house air, while another set was left to stand with no

purging. The treatment of these bottles is described in Table A1. The aqueous phase was

sampled to monitor the aqueous concentrations of monoPAPs and 6:2 diPAP over a period of 6-7

days. At the end of the experiment, the gas diffuser tubes (only present in the purging bottles),

septa, bottle caps, and bottles were sonicated in methanol at 60oC for 1 hour.

Biodegradation experiments.

The mineral medium was prepared using a phosphate-free Tris buffer (6.05 gL-1

of Tris

HCl, 1gL-1

of NH4Cl, 0.68 gL-1 sodium acetate), 1% (v/v) solution of 19.9 μgL-1

of FeCl2•4H2O,

0.9 mgL-1

p-aminobenzoate, 0.9 mgL-1

nicotinic acid, and 1% (v/v) solution of 20 mgL-1

of

(NH4)6Mo7O24•4H2O, 50 mgL-1

of H3BO3, 30 mgL-1

of ZnCl2, 3 mgL-1

of CoCl2•6H2O, 10 mgL-

1 of (CH3COO)2Cu•H2O, 20 mgL

-1 of FeCl2•6H2O, at pH ~7. Phosphate-free conditions were

used to promote the growth of microbes capable of utilizing organophosphates (i.e. PAPs) as

their sole source of phosphorus. The media and mixed liquor (for sterile controls) were

autoclaved for 30 min. at 121oC in a Steris SG-120 Scientific Gravity Sterilizer. Mixed liquor

used as inocula was first shaken to resuspend the biosolids, and then 40 mL were centrifuged at

3000 rpm and the supernatant removed. The isolated biosolids were then washed twice with

media with centrifugation in between washings, and then resuspended in 10% of the total volume

of media used in the biodegradation system.

Extraction Procedure.

For the analysis of FTOHs (4:2, 6:2, 8:2, and 10:2), the XAD resin and glass wool in

each XAD-2 cartridge were extracted with two 2 mL aliquots of ethyl acetate and the combined

fractions were transferred to autosampler vials for analysis by gas chromatography-mass

spectrometry (GC-MS).

242

All water samples were extracted by one of two different methods, depending on the

target analytes, and analyzed by high performance liquid chromatography-mass spectrometry

(HPLC-MS/MS). The choice of using which method was based on spike and recovery

experiments (data not shown), where better efficiencies were obtained for different analytes

depending on the extraction method. For the analysis of PFCAs (C4 – C8), 4:2 and 6:2 FTCAs

and FTUCAs, 3:3 and 5:3 FTCAs, and 4:2, 6:2, and 8:2 monoPAPs, the 0.5 mL water samples

collected at each timepoint were mixed with an equal volume of HPLC-grade methanol and

spiked with the appropriate internal standards (Table A3). The sample/MeOH mixtures were

drawn up using disposable plastic syringes (Norm-Ject®, Tuttlingen, Germany), filtered through

0.25 μm nylon syringe filters (Chromatographic Specialties, Brockville, ON) and transferred to

1.2 mL low-temperature cryo vials (VWR International Ltd., Mississauga, ON) for storage at -

20oC until analysis.

For the analysis of PFCAs (C9 – C11), 8:2 and 10:2 FTCAs and FTUCAs, 7:3 and 9:3

FTCAs, 10:2 monoPAP, and 6:2 diPAP, the 0.5 mL water samples were extracted using the ion-

pairing method developed by Hansen et al. (2). Briefly, 2 mL of 0.25M Na2CO3 solution, 1 mL

of TBAS solution adjusted to pH 10, and 2 mL of MTBE were added to the 0.5 mL water sample

in a 15 mL polypropylene tube (BD Biosciences, Franklin Lakes, NJ). After shaking vigorously

by hand for 5 min. and centrifuged at 3300 rpm for 5 min., the MTBE layer was transferred to a

clean polypropylene tube and a second 2 mL aliquot of MTBE was added to the original water

sample to repeat the extraction for another time. The combined MTBE extracts were evaporated

to dryness under nitrogen, reconstituted in 0.5 mL methanol, vortexed for 30 sec., filtered

through 0.25 μm syringe filters into 1.2 mL cryo vials, and stored at -20oC until analysis.

Instrumental Analysis.

Gas chromatography details.

Analysis of FTOHs was performed using a Hewlett-Packard 6890 GC coupled to a 5973

inert MS (Agilent Technologies, Wilmington, DE) under electron impact ionization (EI) mode.

Quantification proceeded under EI in single ion monitoring mode and their molecular ions are

listed in Table A4. FTOH separation proceeded with the use of a ZebronTM ZB-WAX column

(30 m x 0.25 mm x 0.25 um) (Phenomenex®, Torrence, CA) and the following oven program:

243

the initial oven temperature was 60oC and remained for 2 min. before ramping at 10

oC/min to

95oC. Without holding, the temperature continued to ramp from 95

oC to 240

oC at 30

oC/min and

held for 1 min. The total run time of the oven program is 11.33 min. The carrier gas was helium

at a flow rate of 1.8 mL/min. Injections of 1 uL in pulsed splitless mode were performed at an

initial pressure of 25 psi and at 220oC, followed by a return to 16.16 psi in 1.2 min. Linear

regression of the calibration curves was typically achieved with R2 > 0.99.

Liquid chromatography details.

Separation was performed using a GeminiNX C18 column (4.6 x 50 mm, 3 μm)

(Phenomenex®, Torrance, CA) and the analytes were identified using an API 4000 triple

quadrupole mass spectrometer (Applied Biosystems/MDS Sciex) operating under negative

electrospray ionization mode, coupled to an Agilent 1100 autosampler. Two HPLC-MS/MS

methods were developed and used, depending on the target analyte. For the analysis of PFCAs

(C4 – C11), FTCAs and FTUCAs, and 6:2 diPAP, the samples were injected as 50 μL injections

and analyzed by the following gradient method at 360 μL/min using HPLC grade methanol and

water, each prepared into 10 mM ammonium acetate mobile phases: the initial solvent

composition at t = 0 min. was 40:60 methanol: water, which changed to 95:5 over a period of 6

min. at t = 6 min. and held for 3 min. to t = 9 min. before returning to the initial composition of

40:60 methanol:water at t = 10 min. The column was allowed to reequilibrate for 3.5 min for a

total run time of 13.5 min.

Analysis of monoPAPs was complicated by its highly interactive dianionic phosphate

moiety and a separate chromatographic method at low pH was developed. In order to drive the

conversion of monoPAPs to its monoanionic form, all water samples were acidified with 98%

formic acid (5% v/v). The samples were then injected as 50 μL injections, and analyzed by the

following gradient method at 500 μL/min using HPLC grade methanol and water as mobile

phases, each containing 0.5% (v/v) formic acid: the initial solvent composition at t = 0 min. was

40:60 methanol:water, which changed to 95:5 over a period of 2.5 min. and held for 7.5 min.

before returning to the initial composition of 40:60 methanol:water at t = 10.5 min. The column

was allowed to reequilibrate for 2.5 min. for a total run of time of 13.0 min. Sample

chromatograms of the monoPAPs in a 1 ppb standard and sample extract are shown as Figure S2.

244

Quality assurance of data.

Quantification of 6:2 diPAP and monoPAPs using external calibration. To justify the use

of external calibration, 2 ug of 4:2, 6:2, 8:2, 10:2 monoPAPs and 6:2 diPAP were spiked into

mineral media treated with autoclaved mixed liquor. After 1 day, aliquots of the aqueous phase

were extracted and analyzed by both standard addition and external calibration using PAP

standards prepared in methanol and PAP matrix-matched standards prepared in the same media

used for the spike and recovery. Recoveries obtained by the two quantification methods are

presented as Figure A3. The use of standard addition or external calibration to quantify PAPs

was found to have no statistical difference (p<0.05) using the student’s t-test (SigmaPlot 9.01,

Systat Software, Inc. 2004); therefore, the latter method was chosen.

Spike and recovery procedure.

A spike and recovery (n =4) was performed to validate the extraction efficiency of XAD

cartridges to trap purged FTOHs. A mass of 20 ug of 4:2, 6:2, 8:2, and 10:2 FTOHs was spiked

into purge-and-trap bottles containing 400 mL of mineral media treated with autoclaved mixed

liquor and the bottles were purged for 1 day. A recovery experiment (n = 4) was performed for

the PFCAs, FTCAs, FTUCAs, and PAPs by spiking 50 ng of the available standards into 50 mL

of mineral media treated with autoclaved mixed liquor, followed by extraction after 1 day.

Recovery results for all target analytes are provided in Table A3 and A4.

RESULTS AND DISCUSSION

Purging control experiment.

The aqueous concentrations of PAPs in the bottles left to stand, i.e. no purging, stayed

relatively consistent, except for 10:2 monoPAP, where only 33±7% of the mass that was initially

added was measured in the aqueous phase on the last day (Figure A5). This observed loss

occurred during the first 2 days of the experiment, suggesting initial rapid adsorption to the bottle

walls. In a sediment sorption experiment, Higgins and Luthy also observed that the 2-(N-

methyl-) and 2-(N-ethylperfluorooctanesulfonamido) acetate (N-MeFOSAA, N-EtFOSAA)

sorbed to polystyrene vial walls in controls with no sediment added and were difficult to recover

245

from the aqueous phase (3). At the end of the experiment, 57±13% of 10:2 monoPAP was

accounted for from the aqueous phases and after sonication of the septa, caps, and bottles

themselves. The 4:2, 6:2, 8:2 monoPAPs and 6:2 diPAP were mostly retained in the aqueous

phase (Figure A6).

In the bottles undergoing purging, 62±4%, 37±16%, 26±10%, and 15±10% of 4:2, 6:2,

8:2, 10:2 monoPAPs and 97±12% of 6:2 diPAP were accounted for from the aqueous phase,

bottles, septa, and gas diffuser tubes at day 6 (Figure A6). Purging appeared to decrease the

aqueous concentrations of the PAPs over time, especially those of the monoPAPs (Figure A5).

One explanation is that the surface-active PAPs may have been enriched at the air-water

interface and partitioned into bubbles formed during purging. As these bubbles eject into the

headspace, they may break on contact with any surfaces (eg. bottle walls, bottom of caps, septa,

and gas diffuser tubes) and release PAPs for sorption to these surfaces. The gas diffuser tubes,

which were absent in the bottles left to stand, i.e. no purging, also contain porous glass tips to

disperse air into the water and may represent a significant amount of surface area to irreversibly

bind the PAPs.

LITERATURE CITED

(1) Achilefu, S.; Mansuy, L.; Selve, C.; Thiebaut, S. Synthesis of 2H,2H-Perfluoroalkyl and

2H-Perfluoroalkenyl Carboxylic Acids and Amides. J. Fluor. Chem. 1995, 70, 19–26.



Sci. Technol. 2001, 35, 766–770.


Sci. Technol. 2006, 40, 7251–7256.

246

Figure A1. Diagram of the purge-and-trap system used in the purging control and

biodegradation experiments.

CARBON AIR FILTER

Active Active Active Active Blank Blank

In-house air

XAD

cartridge

247

Table A1. Description of bottles in the purging control experiment.

Control Type (n

= 2)

Volume Fraction of Added Component per Bottle Total

Volume per

Bottle

1

(1 ugmL-1

PAPs;

no purging)

4:2 monoPAP

4 mL of 100 ugmL-1

stock

= 400 ug

400 mL

6:2 monoPAP

8:2 monoPAP

10:2 monoPAP

6:2 diPAP 0.606 mL of 660 ugmL

-1 stock

= 400 ug

18Ω deionized water 395 mL

2

(1 ugmL-1

PAPs;

purging)

4:2 monoPAP

4 mL of 100 ugmL-1

stock

= 400 ug

400 mL

6:2 monoPAP

8:2 monoPAP

10:2 monoPAP


-1 stock

= 400 ug

18Ω deionized water 395 mL

248

Table A2. Description of experimental and control bottles of the biodegradation experiments.

Control Type Volume Fraction of Added Component per

Bottle

Total

Volume per

Bottle

Mixed liquor only

(n = 2)

- 10% v/v washed

mixed liquor

4:2 monoPAP 0 mL

400 mL

6:2 monoPAP 0 mL

8:2 monoPAP 0 mL

10:2 monoPAP 0 mL

6:2 diPAP 0 mL

Mineral media 360 mL

Washed mixed

liquor

40 mL

Hg2Cl2 0 mL

Sterile control

(n = 2)

- 1 ugmL-1

of PAPs

- 10% v/v of

autoclaved mixed

liquor

- 300 mg of Hg2Cl2

4:2 monoPAP 0.505 mL of 792 ugmL

-1 stock

= 400 ug

400 mL


-1 stock

= 400 ug


-1 stock

= 400 ug


-1 stock

= 400 ug


-1 stock =

400 ug

Mineral media 351 – 353 mL

Sterile mixed

liquor

40 mL

Hg2Cl2 0 mL5.6 mL of 54 mgmL-1

stock

= 300 mg

PAPs only

(n = 2)

- 1 ugmL-1

of PAPs


-1 stock

= 400 ug

400 mL


-1 stock

= 400 ug


-1 stock

= 400 ug


-1 stock

= 400 ug


-1 stock =

400 ug

Mineral media 397 – 399 mL

Washed mixed

liquor

0 mL

Hg2Cl2 0 mL

249

Control Type Volume Fraction of Added Component per

Bottle

Total

Volume per

Bottle

Experimental

(Chain length study)

(n = 3)

- 1 ugmL-1

of PAPs

- 10% v/v of washed

mixed liquor


-1 stock

= 400 ug

400 mL


-1 stock

= 400 ug


-1 stock

= 400 ug


-1 stock

= 400 ug


Washed mixed

liquor

40 mL

Hg2Cl2 0 mL

Experimental

(Substitution study)

(n = 3)

- 1 ugmL-1

of PAPs

- 10% v/v of washed

mixed liquor


-1 stock

= 400 ug

400 mL 6:2 diPAP

0.606 mL of 660 ugmL-1

stock =

400 ug


Washed mixed

liquor

40 mL

Hg2Cl2 0 mL

250

Table A3. Internal standards, multiple reaction monitoring (MRM) mass transitions, and

recovery results for all target analytes in the aqueous phase.

Analyte Internal Standard MRM transition Recovery (%)

(n = 4)

PFBA (C4) 13

C2-PFHxA 212.8 > 168.9 111 ± 3

PFPeA (C5) 13

C2-PFHxA 262.8 > 219.0 130 ± 3

PFHxA (C6) 13

C2-PFHxA 313.0 > 268.8 116 ± 3

PFHpA (C7) 13

C4-PFOA 362.9 > 318.8 153 ± 5

PFOA (C8) 13

C4-PFOA 413.0 > 368.9 91 ± 3

PFNA (C9) 13

C5-PFNA 462.9 > 419.0 104 ± 11

PFDA (C10) 13

C2-PFDA 513.0 > 468.9 72 ± 8

PFUnA (C11) 13

C2-PFDA 562.8 > 519.0 95 ± 14

4:2 FTCA 13

C2-6:2 FTUCA 276.9 > 192.9 112 ± 2

6:2 FTCA 13

C2-6:2 FTUCA 376.9 > 292.9 105 ± 3

8:2 FTCA 13

C2-8:2 FTUCA 477.0 > 393.0 98 ± 7

10:2 FTCA 13

C2-10:2 FTUCA 577.0 > 492.9 68 ± 3

4:2 FTUCA 13

C2-6:2 FTUCA 256.9 > 192.8 116 ± 3

6:2 FTUCA 13

C2-6:2 FTUCA 356.9 > 292.9 85 ± 3

8:2 FTUCA 13

C2-8:2 FTUCA 457.0 > 393.0 107 ± 9

10:2 FTUCA 13

C2-10:2 FTUCA 557.0 > 492.9 78 ± 6

3:3 FTCA*

13C2-6:2 FTUCA 241.0 > 137.0 -

5:3 FTCA* 13

C2-6:2 FTUCA 341.0 > 237.0 -

7:3 FTCA 13

C2-8:2 FTUCA 441.0 > 337.0 82 ± 5

251

9:3 FTCA* 13

C2-10:2 FTUCA 541.0 > 437.0 -

4:2 monoPAP** - 343.0 > 96.9 96 ± 17

6:2 monoPAP** - 443.0 > 96.9 80 ± 21

8:2 monoPAP** - 543.0 > 96.9 40 ± 3

10:2 monoPAP** - 643.0 > 96.9 33 ± 8

6:2 diPAP** - 789.0 > 96.9 102 ± 5

* Spike and recovery experiments were not performed due to lack of available analytical

standards at the time of experiment.

** Quantitation was performed by external calibration as no appropriate internal standards were

available at the time of experiment.

252

Table A4. Single ion monitoring (SIM) molecular ions, dwell time, and recovery results for

FTOHs in the headspace.

Analyte SIM Dwell time (ms) Recovery (%)

(n = 4)

4:2 FTOH 263.0 100 58 ± 23

6:2 FTOH 363.0 100 91 ± 30

8:2 FTOH 463.0 100 83 ± 27

10:2 FTOH 563.0 100 63 ± 4

253

Figure A2. Typical chromatograms of monoPAPs in a 1 ppb (ngmL-1

) standard and Day 0 water

extract from control bottle spiked with only monoPAPs in mineral media.

X Data

0 2 4 6 8 10 12 14

X Data

0 2 4 6 8 10 12 14

1 ppb mixed standard Day 0 sample in 'PAPS only' control after inoculation with sludge

4:2 monoPAPS

6:2 monoPAPS

8:2 monoPAPS

10:2 monoPAPS

Time (days)

0 2 4 6 8 10 12 14


4:2 monoPAPS

6:2 monoPAPS

8:2 monoPAPS

10:2 monoPAPS

Time (days)

0 2 4 6 8 10 12 14


4:2 monoPAPS

6:2 monoPAPS

8:2 monoPAPS

10:2 monoPAPS

Area count = 7690

Area count = 9120

Area count = 18400

Area count = 13500

Area count = 519000

Area count = 293000

Area count = 52200

Area count = 56600

254

Figure A3. Recoveries of PAPs in mineral media treated with autoclaved mixed liquor,

analyzed by standard addition and external calibration using matrix-matched standards. Error

bars represent the standard error of spike and recovery experiment performed in triplicate.

Analyte

4:2 MP 6:2 MP 8:2 MP 10:2 MP 6:2 DP

% R

eco

very

0

20

40

60

80

100

120

140

160

Standard addition

External calibration

Mineral media treated with mixed liquor

255

Table A5. Limits of detection (LOD) and limits of quantitation (LOQ) for the target analytes. In

all figures, values less than the LOD were reported as not detected (nd) and assigned a value of

zero; while values less than the LOQ were reported without alteration. Unless indicated with an

asterisk, all LODs and LOQs were calculated based on contamination in the procedural blanks.

Analyte LOD (ngmL-1

) LOQ (ngmL-1

)

PFBA (C4)* 0.02 0.10

PFPeA (C5)* 0.02 0.10

PFHxA (C6) 0.08 0.26

PFHpA (C7) 1.72 5.74

PFOA (C8) 0.57 1.91

PFNA (C9) 0.87 2.90

PFDA (C10) 0.11 0.37

PFUnA (C11) 0.22 0.66

4:2 FTCA* 0.02 0.10

6:2 FTCA 0.12 0.38

8:2 FTCA 0.12 0.42

10:2 FTCA 0.03 0.09

4:2 FTUCA 0.05 0.17

6:2 FTUCA 0.06 0.21

8:2 FTUCA 0.17 0.57

10:2 FTUCA 0.11 0.38

3:3 FTCA** 0.02 0.10

5:3 FTCA** 0.12 0.38

7:3 FTCA* 0.02 0.10

256

9:3 FTCA** 0.03 0.09

4:2 monoPAP 2.34 7.79

6:2 monoPAP 1.15 3.83

8:2 monoPAP 0.41 1.38

10:2 monoPAP 1.03 3.43

6:2 diPAP 2.00 6.68

4:2 FTOH* 10 25

6:2 FTOH* 10 25

8:2 FTOH* 10 25

10:2 FTOH* 10 25

* LOD and LOQ were empirically derived as the concentrations giving a signal-to-noise ratio ≥

3 and ≥ 10 respectively.

** As there were no native and mass-labeled standards available for 3:3, 5:3, and 9:3 FTCAs,

these analytes were quantified based on the calibration for 4:2, 6:2, and 10:2 FTCAs and

therefore, shared the same LODs and LOQs as these surrogate standards.

257

Figure A4. Positive control to assess microbial viability during the 92-day biodegradation.

Disappearance of 6:2 FTUCA spiked and subsequent production of 5:3 FTCA, PFPeA, and

PFHxA in control bottles containing active mixed liquor at day 21, day 51, and day 85. Data are

represented as arithmetic means (±standard error) of duplicate incubations, except for PFHxA

where data was collected from one incubation.

Time (days)

0 20 40 60 80 100

Am

ou

nt

pe

r b

ott

le (

nm

ol)

0

2

4

6

8

10

12

14

6:2 FTUCA (N=2)

5:3 FTCA (N=2)

PFHxA (N=1)

PFPeA (N=2)

258

Figure A5. Changes in levels of 4:2, 6:2, 8:2, 10:2 monoPAPs and 6:2 diPAP in

purging control experiment: ( ) Control 1, no purging, and ( ) Control 2, purging. Data are

represented as arithmetic means (±standard error) of duplicate incubations.

Time (hours)0 20 40 60 80 100 120 140

Am

ou

nt

in b

ott

le (

nm

ol)

0

200

400

600

800

1000

1200

1400

1600

1800

Control 1: PAPs left to stand (no purging)

Control 2: PAPs under purging

Time (hours)0 20 40 60 80 100 120 140

0

200

400

600

800

1000

1200

Time (hours)

0 20 40 60 80 100 120 140

0

200

400

600

800

1000

1200

Time (hours)

0 20 40 60 80 100 120 140

0

200

400

600

800

4:2 monoPAPS

6:2 monoPAPS

8:2 monoPAPS

10:2 monoPAPS

Time (hours)

0 20 40 60 80 100 120 140 160 180

Am

ou

nt

in b

ott

le (

nm

ol)

0

200

400

600

800

6:2 diPAPS

Am

ou

nt

in b

ott

le (

nm

ol)

259

Figure A6. Total recoveries of monoPAPs and 6:2 diPAP summed together from individual

extractions of water, septa, gas diffuser tubes, and bottles at the end of the purging control

experiment. ( ) Control 1, no purging, and ( ) Control 2, purging. Data are represented as

arithmetic means (±standard error) of duplicate incubations.

Control 1 Control 2

% R

ec

ove

ry

0

20

40

60

80

100

Control 1: No purging

Control 2: Purging

Control 1 Control 2 Control 1 Control 2 Control 1 Control 2

Control 1 Control 2

% R

ec

ove

ry

0

20

40

60

80

100

120

140

4:2 monoPAPS6:2 monoPAPS

8:2 monoPAPS

10:2 monoPAPS

6:2 diPAPS

260

Time (days)

0 20 40 60 80 100

Am

ou

nts

of

6:2

diP

AP

an

d

mo

no

PA

Ps in

bo

ttle

(n

mo

l)

0

50

100

150

200

250

3006:2 diPAPS

6:2 monoPAPS

0 20 40 60 80 100

0

50

100

150

1000

2000

4:2 monoPAPS

6:2 monoPAPS

8:2 monoPAPS

10:2 monoPAPS

Figure A7. Levels of (a) 6:2 diPAP and 6:2 monoPAP and (b) 4:2, 6:2, 8:2, and 10:2

monoPAPs in sterile controls over the 92-day biodegradation experiments. Data are represented

as arithmetic means (±standard error) of duplicate incubations.

261

Figure A8. Production of FTCAs, FTUCAs, and PFCAs in the aqueous phase of monoPAP-

dosed bottles in the chain length experiment. (a) Transformation of 4:2 monoPAP, (b)

Transformation of 6:2 monoPAP, (c) Transformation of 8:2 monoPAP, (d) Transformation of

10:2 monoPAP. Data are represented as arithmetic means (±standard error) of triplicate

incubations. Values less than LOD are reported as zero and values in between the LOD and LOQ

were used unaltered and indicated with an asterisk (*) in matching colours.

0 20 40 60 80 100

Am

ou

nt

in b

ott

le (

nm

ol)

0.0

1.0

2.0

3.0

4.0

PFBA

4:2 FTCA

4:2 FTUCA

**** * *** * * * * * * * *

0 20 40 60 80 100

0.0

0.5

1.0

1.5

2.0

PFHxA

6:2 FTCA

6:2 FTUCA

5:3 FTCA

*

*

* * * *

** *

* **

* *

*

0 20 40 60 80 100

0.0

0.5

1.0

1.5

2.0

PFOA

8:2 FTCA

8:2 FTUCA

7:3 FTCA

** * * *

* * * *** * * * **

Time (days)

0 20 40 60 80 100

0.0

0.2

0.4

0.6

0.8

1.0

PFDA

10:2 FTCA

10:2 FTUCA

9:3 FTCA

* ** *

**

** *

* * * * * * * *

(a) Degradation of 4:2 monoPAP

(b) Degradation of 6:2 monoPAP

(c) Degradation of 8:2 monoPAP

(d) Degradation of 10:2 monoPAP

262

APPENDIX B

SUPPORTING INFORMATION FOR CHAPTER FOUR

Biosolids Application as a Source of Polyfluoroalkyl Phosphate Diesters and Their

Metabolites in a Soil-Plant Microcosm: Biodegradation and Plant Uptake

263


Table B1.Target analytes of interest monitored in this study 269

Figure B1.Experimental set-up of soil-plant microcosm 270

Table B2. Multiple reaction monitoring (MRM) transitions for all target analytes and

their internal standards. Matrix recoveries in different compartments for all target

analytes.

271

Table B3.Limits of detection (LODs) and limits of quantitation (LOQs) in soil and plants

for the target analytes. 272

Table B4.Plant growth parameters (mean ± standard error) 273

Figure B2.Concentrations of diPAPs and PFCAs in the soil and plants sampled from

Treatment 2 (WWTP biosolids-amended soils) 274

Figure B3.Concentrations of diPAPs and PFCAs in the soil and plants sampled from

Treatment 3 (WWTP- and paper fiber biosolids-amended soils) 275

Figure B4.Molar distribution (%) of diPAPs and PFCAs in the catch plates, soil, and

plant of Treatments 2–4 microcosms 276

264

EXPERIMENTAL

Chemicals.

Perfluorobutanoate (PFBA (C4), >99%), perfluoropentanoate (PFPeA (C5), >99%),

perfluorohexanoate (PFHxA (C6), >99%), perfluoroheptanoate (PFHpA (C7), >99%),

perfluorooctanoate (PFOA (C8), >99%), perfluorononanoate (PFNA (C9), >99%),

perfluorodecanoate (PFDA (C10), >99%), perfluoroundecanoate (PFUnA (C11), >99%),

perfluorododecanoate (PFDoA (C12), >99%), perfluorotridecanoate (PFTrA (C13), >99%),

perfluorotetradecanoate (PFTeA (C14), >99%), 6:2, 8:2, and 10:2 fluorotelomer carboxylates

(FTCAs, >98%), and 6:2, 8:2, and 10:2 fluorotelomer unsaturated carboxylates (FTUCAs,

>98%) were obtained from Wellington Laboratories (Guelph, ON). The 4:2 FTCA and 4:2

FTUCA were synthesized as per the methods reported by Achilefuet al,1 with final purities of

>95%. The 7:3 FTCA (>97%) was purchased from SynQuest Labs, Inc. (Alachua, FL). Mass-

labeled internal standards were donated from Wellington Laboratories and they included: 13

C2-

PFHxA (>99%), 13

C4-PFOA (>99%), 13

C5-PFNA (>99%), 13

C2-PFDA (>99%), 13

C2-PFUnA

(>99%), 13

C2-PFDoA (>99%), 13

C2-6:2 FTUCA (>99%), 13

C2-8:2 FTUCA (>98%), and 13

C2-

10:2 FTUCA (>98%).

The 4:2, 6:2, 8:2, and 10:2 polyfluoroalkyl phosphate diesters (diPAPs) were synthesized

by methods described elsewhere.2

Methanol (Omnisolv, >99%), water (Omnisolv, >99%), methyl-tert-butyl ether (MTBE,

Omnisolv, >99%), and ammonium hydroxide (30%) were purchased from EMD Chemicals, Inc.

(Mississauga, ON). Sodium azide (NaN3) was purchased from Anachemia Sciences (Montreal,

ON).

Preparation of Rhizobia Inoculum.

Six different Sinorhizobium strains were selected to be cultured on agar plates. The agar

media contained 18 g/L agar, 60 g/L urea, and 29 g/L sodium chloride (NaCl) in distilled water.

After autoclaving at 120oC for 20 min., the agar media were poured into 6 different petri plates

and allowed to solidify. A sterile stainless steel loop was used to transfer each rhizobium strain

to each petri plate and the plates were subsequently incubated at 30oC for 5 days. Single

colonies of each of the 6 rhizobium strains were then transferred and incubated separately in

265

autoclaved liquid YG growth media, which contained 5 g/L tryptone (Difo, Bioshop), 3 g/L yeast

extract (Difco, Bioshop), and 1.5 g/L calcium chloride (Sigma), adjusted to pH 7, for 4 days.

The cell density of each rhizobium strain in the final rhizobia inoculum mixture was adjusted to

~108 cells/mL (based on optical density 600 nm) by diluting the 6 liquid cultures with distilled

water.

Extraction of Soil and Plant Samples for diPAPs, FTCAs, FTUCAs, and PFCAs.

At each timepoint, 2–5 g of soil were sampled in triplicate (n = 3) from each pot and each

subsample was sonicated in 5 mL of basic methanol (containing 1% (v/v) ammonium hydroxide)

at 60oC for 15–20 min. After centrifuging at 6000 rpm and transferring the supernatant to a new

polypropylene tube, the sonication step was repeated and the 2 aliquots of basic methanol were

combined and evaporated to dryness under nitrogen. Following reconstitution with 2 mL of

methanol, the sample was filtered with 0.2 µm syringe filters and stored in low-temperature

cryovials at -20oC until analysis.

Plant matter (2 g) was first freeze-dried with liquid nitrogen, and then homogenized

finely using a mortar pestle. The resulting powder was sonicated in 10 mL of basic methanol at

60oC for 15–20 min, followed by centrifuging and removal of the supernatant. This step was

repeated and the combined aliquots of basic methanol were then concentrated to 5 mL for further

clean-up using ENVI Carb cartridges (Supelclean, 1 mL/100 mg). The cartridges were

preconditioned with 3 aliquots of basic methanol, then loaded with the samples, and finally

rinsed with 2 mL of basic methanol to elute the target analytes. After evaporating the methanol

extract to dryness, the sample was reconstituted in 2 mL methanol, filtered, and stored as

described above.

The catch plates were rinsed with 10 mL of basic methanol, which was then transferred to

a polypropylene tube and evaporated to dryness. The sample was then reconstituted in 2 mL of

methanol, filtered, and stored as described above.


Chromatographic separation was performed using a GeminiNX C18 column (50 x 4.6

mm, 3 µm; Phenomenex, Torrance, CA). Analyte quantitation was performed using an API4000

266

triple-quadrupole mass spectrometer (Applied Biosystems/MDS Sciex) in the negative

electrospray ionization mode, coupled to an Agilent 1100 high pressure liquid chromatography

(HPLC) system. Three HPLC gradient methods were used for the analysis of the target analytes.

For the analysis of the diPAPs, the samples were injected as 50 µL injections and

analyzed by the following gradient method at 500 µL/min using HPLC grade methanol and

water, each prepared into 50 mM ammonium acetate mobile phases: the initial solvent

composition at t = 0 min. was 60:40 water:methanol, which to changed to 5:95 over a period of

2.5 min. at t = 2.5 min. and held for 3.5 min. to t = 6.0 min., before returning to the initial

composition of 60:40 water:methanol at t = 6.5 min. The column was allowed to reequilibrate

for 1.5 min. for a total run time of 8 min.

For the analysis of PFBA, PFPeA, PFHxA, PFHpA, PFOA, 3:3, 4:2, 5:3, 6:2, 7:3, and

8:2 FTCAs and FTUCAs, the samples were injected as 25 µL injections and analyzed by the

following gradient method at 500 µL/min, using the same mobile phases as above: the initial

solvent composition at t = 0 min. was 80:20 water:methanol, which changed to 10:90 over a

period of 3 min. at t = 3.0 min. and held for 2 min. to t = 5.0 min., before returning to the initial

composition of 80:20 water:methanol at t = 5.5 min. The column was allowed to reequilibrate

for 1.5 min. for a total run time of 7 min.

For the analysis of PFNA, PFDA, PFUnA, PFDoA, PFTrA, PFTeA, 9:3 and 10:2 FTCAs

and FTUCAs, the samples were injected as 25 µL injections and analyzed by the following

gradient method at 500 µL/min, using the same mobile phases as above: the initial solvent

composition at t = 0 min. was 25:75 water:methanol, which changed to 5:95 over a period of 2

min. at t = 2.0 min. and held for 2 min. to t = 4.0 min., before returning to the initial composition

of 25:75 water:methanol at t = 4.5 min. The column was allowed to reequilibrate for 1.5 min. for

a total run time of 6 min.

A list of the analyte-specific multiple reaction monitoring (MRM) transitions for all

target analytes and their corresponding mass-labeled internal standards is provided in Table B2.

267

Quality Assurance of Data.

Preparation of Matrix-Matched Calibration Standards.

Matrix-matched calibration standards were prepared by extracting control soil and plant

samples in the same manner as the experimental samples, followed by spiking of the target

diPAPs at 7 different concentrations. The control soil was sampled from Treatment 1 pots in

which the soil did not receive any biosolids amendment. The control plants were sampled from

the same agricultural farmland (Northumberland County, ON; 44o05’N, 78

o01’W) from which

the soil used here was collected. These control matrices were also used in the spike and recovery

experiments, as described below. Endogenous contamination of the diPAPs was not observed in

either of the control matrices, except for the 6:2 diPAP, which was detected at small quantities in

the control soil. Background concentration of the 6:2 diPAP was determined in the control soil

and used to correct the concentrations of the matrix-matched calibration standards.

Spike and Recovery Procedures in Soil, Plants, and Catch Plates.

Spike and recovery experiments in soil were performed in triplicate (n = 3) by adding 200

ng of 4:2, 6:2, 8:2, and 10:2 diPAPs and 100 ng of 4:2, 6:2, 8:2, and 10:2 FTCAs and FTUCAs,

7:3 FTCAs, and C4–C14 PFCAs to 1 g of control soil. Plant recovery experiments were

performed in triplicate (n = 3) by adding 200 ng of the diPAPs, 100 ng of the FTCAs and

FTUCAs, and 10 ng of the C4–C14 PFCAs to 2 g of control plant material. These spiked

matrices were extracted and analyzed as described above. Spike and recovery experiments were

also performed in triplicate (n = 3) in the catch plates by adding 200 ng of the diPAPs, 100 ng of

the FTCAs and FTUCAs, and 10 ng of the C4–C14 PFCAs to clean catch plates containing 5 mL

of MTBE. After gently swirling the plates, the MTBE was allowed to evaporate overnight in a

fumehood and the plates were rinsed with methanol as described in the manuscript the following

day to extract the spiked analytes. The matrix-specific recoveries for each analyte are listed in

Table B2.

268

LITERATURE CITED

(1) Achilefu, S.; Mansuy, L.; Selve, C.; Thiebaut, S. Synthesis of 2H,2H-Perfluoroalkyl and

2H-Perfluoroalkenyl Carboxylic Acids and Amides. J. Fluor. Chem.1995, 70, 19–26.



Routes of Human Contamination. Environ. Sci. Technol.2007, 41, 4799–4805.

269

Table B1. Target analytes of interest monitored in this study.

Structure Name Acronym

F

F

F F

F F

F F

F F

O

O

P

O

O-

x

y

Polyfluoroalkyl phosphate diester

DiPAP

x = 4, 6, 8, 10

y = x or x + 2

If y = x, x:2 diPAP

If y = x + 2, x:2/y:2 diPAP

F

F F O

O-x


carboxylate

x:2 FTCA

x = 4, 6, 8, 10

F

F F

x

F

O

O-


carboxylate

x:2 FTUCA

x = 4, 6, 8, 10

F

F F

xO

O-


carboxylate

x:3 FTCA

x = 3, 5, 7, 9

F

F F

xO

O-


carboxylate

x:3 FTUCA

x = 3, 5, 7, 9

OF

F F

O-x

Perfluorocarboxylate PFCA

x = 3–13

270

Figure B1. Experimental set-up of soil-plant microcosm.

Timepoint

(Month)

Treatment

1 2 3 4 5

Soil only

(n = 1)

Biosolids-amended

soil/Plant

(n = 3)

Biosolids- and paper

fiber solids-amended

soil/Plant

(n = 3)

Biosolids-amended

soil/Plant/100 mg

6:2 monoPAP

(n = 3)

Biosolids-amended

soil/Plant/100 mg

6:2 diPAP

(n = 3)

0

1.5

3.5

5.5

271

Table B2. Multiple reaction monitoring (MRM) transitions for all target analytes and their

internal standards. Matrix recoveries in different compartments for all target analytes.

Target Analyte Mass

transition Internal Standard

Mass

transition

Recovery (%) (n= 3)

Soil Plant Catch

Plate

Polyfluoroalkyl phosphate diester (diPAP)

4:2 diPAP 589.1>96.9

- - 79 ± 14 78 ± 7 117 ± 23 589.1>343.0

4:2/6:2 diPAP 689.0>96.9 - - - - -

6:2 diPAP 789.0>96.9

- - 91 ± 7 37 ± 9 83 ± 16 789.0>443.0

6:2/8:2 diPAP 889.0>96.9 - - - - -

8:2 diPAP 989.0>96.9

- - 115 ± 37 55 ± 4 39 ± 11 989.0>543.0

8:2/10:2 diPAP 1089.0>96.9 - - - - -

10:2 diPAP 1189.0>96.9

- - 82 ± 38 110 ± 14 * 1189.0>643.0

10:2/12:2 diPAP 1289.0>96.9 - - - - -

Fluorotelomer saturated and unsaturated carboxylate (FTCA and FTUCA)

4:2 FTCA 276.9>192.9 13

C2-6:2 FTUCA 359.0>294.0 57 ± 11 24 ± 5 36 ± 1

6:2 FTCA 376.9>292.9 13

C2-6:2 FTUCA 359.0>294.0 100 ± 5 40 ± 4 26 ± 12

8:2 FTCA 477.0>393.0 13

C2-8:2 FTUCA 459.0>394.0 132 ± 8 87 ± 26 32 ± 7

10:2 FTCA 577.0>492.9 13

C2-10:2 FTUCA 559.0>494.0 114 ± 8 57 ± 4 91 ± 1

3:3 FTCA 241.0>137.0 13

C2-6:2 FTUCA 359.0>294.0 - - -

5:3 FTCA 341.0>237.0 13

C2-6:2 FTUCA 359.0>294.0 - - -

7:3 FTCA 441.0>337.0 13

C2-8:2 FTUCA 459.0>394.0 51 ± 5 - -

9:3 FTCA 541.0>437.0 13

C2-10:2 FTUCA 559.0>494.0 - - -

4:2 FTUCA 256.9>192.8 13

C2-6:2 FTUCA 359.0>294.0 84 ± 10 34 ± 3 64 ± 3

6:2 FTUCA 356.9>292.9 13

C2-6:2 FTUCA 359.0>294.0 87 ± 1 68 ± 7 48 ± 8

8:2 FTUCA 457.0>393.0 13

C2-8:2 FTUCA 459.0>394.0 100 ± 3 135 ± 18 69 ± 2

10:2 FTUCA 557.0>492.9 13

C2-10:2 FTUCA 559.0>494.0 114 ± 8 93 ± 2 89 ± 26

3:3 FTUCA 239.0>169.0 13

C2-6:2 FTUCA 359.0>294.0 - - -

5:3 FTUCA 339.0>269.0 13

C2-6:2 FTUCA 359.0>294.0 - - -

7:3 FTUCA 439.0>369.0 13

C2-8:2 FTUCA 459.0>394.0 - - -

9:3 FTUCA 539.0>469.0 13

C2-10:2 FTUCA 559.0>494.0 - - -

Perfluorocarboxylate (PFCA)

PFBA 212.8>168.9 13

C2-PFHxA 314.8>269.8 84 ± 4 107 ± 14 70 ± 21

PFPeA 262.8>218.97 13

C2-PFHxA 314.8>269.8 96 ± 6 53 ± 11 66 ± 27

PFHxA 312.8>268.9 13

C2-PFHxA 314.8>269.8 104 ± 6 109 ± 7 105 ± 60

PFHpA 362.8>319.0 13

C4-PFOA 417.0>372.0 108 ± 4 101 ± 2 73 ± 21

PFOA 413.0>368.9 13

C4-PFOA 417.0>372.0 73 ± 12 73 ± 5 76 ± 13

PFNA 462.9>419.0 13

C5-PFNA 468.0>423.0 127 ± 8 118 ± 17 81 ± 9

PFDA 513.0>470.0 13

C2-PFDA 515.0>470.0 94 ± 22 104 ± 5 83 ± 2

PFUnA 562.8>519.0 13

C2-PFUnA 564.8>520.0 89 ± 4 90 ± 2 84 ± 13

PFDoA 612.8>569.0 13

C2-PFDoA 614.8>570.0 74 ± 6 112 ± 10 96 ± 5

PFTrA 662.8>619.0 13

C2-PFDoA 614.8>570.0 27 ± 8 58 ± 8 116 ± 5

PFTeA 712.8>669.0 13

C2-PFDoA 614.8>570.0 50 ± 5 52 ± 4 159 ± 10

272

Table B3. Limits of detection (LODs) and limits of quantitation (LOQs) in soil and plants for

the target analytes.

Target Analyte

Soil Plant

Method Method

LOD LOQ LOD LOQ

(ng/g) (ng/g)

Polyfluoroalkyl phosphate diester (diPAP)

4:2 diPAP 0.02 0.07 0.25 0.83

4:2/6:2 diPAP - - - -

6:2 diPAP 0.07 0.23 0.47 1.57

6:2/8:2 diPAP - - - -

8:2 diPAP 0.13 0.42 0.42 1.40

8:2/10:2 diPAP - - - -

10:2 diPAP 7.59 25.30 2.20 7.33

10:2/12:2 diPAP - - - -

Fluorotelomer saturated and unsaturated carboxylate (FTCA and FTUCA)

4:2 FTCA 0.03 0.09 0.10 0.33

6:2 FTCA 0.05 0.16 0.09 0.29

8:2 FTCA 0.04 0.12 0.12 0.39

10:2 FTCA 0.02 0.06 0.74 2.48

3:3 FTCA - - - -

5:3 FTCA - - - -

7:3 FTCA 0.01 0.02 0.01 0.03

9:3 FTCA - - - -

4:2 FTUCA 0.01 0.02 0.02 0.07

6:2 FTUCA 0.002 0.01 0.01 0.03

8:2 FTUCA 0.001 0.003 0.01 0.03

10:2 FTUCA 0.01 0.03 0.03 0.11

3:3 FTUCA - - - -

5:3 FTUCA - - - -

7:3 FTUCA - - - -

9:3 FTUCA - - - -

Perfluorocarboxylate (PFCA)

PFBA 0.02 0.07 0.07 0.23

PFPeA 0.01 0.03 0.03 0.10

PFHxA 0.002 0.01 0.01 0.03

PFHpA 0.003 0.01 0.02 0.05

PFOA 0.003 0.01 0.01 0.03

PFNA 0.01 0.04 0.07 0.22

PFDA 0.01 0.02 0.05 0.18

PFUnA 0.004 0.01 0.07 0.25

PFDoA 0.003 0.01 0.04 0.13

PFTrA 0.004 0.01 0.09 0.30

PFTeA 0.005 0.02 0.14 0.47

273

Table B4. Plant growth parameters (mean ± standard error).

aThe growth rates were calculated by fitting all plant mass data to an exponential model (ln(mass, g) = a + bt, where

a is a constant, b is the growth rate (g/month) and t is the time (month). The coefficients of correlation, r, for the

model are shown in parentheses.

Treatment Growth Rate (g/month)

(r)a

Plant Mass (g)

1.5 Month 3.5 Month 5.5 Month

2 0.56 ± 0.02 (1.00) 1.28 ± 0.09 4.24 ± 0.78 12.00 ± 2.67

3 0.27 ± 0.17 (0.85) 2.26 ± 0.76 6.97 ± 1.99 6.74 ± 0.70

4 0.76 ± 0.08 (0.99) 0.63 ± 0.16 3.82 ± 0.17 13.08 ± 1.20

274

Figure B2.Concentrations of diPAPs and PFCAs in the soil and plants sampled from Treatment 2 (WWTP biosolids-amended

soils).Each data point represents the arithmetic mean concentration of the triplicate (n = 3) sampling at each timepoint. The error bar

represents the standard error.

PF

BA

PF

PeA

PF

Hx

A

PF

Hp

A

PF

OA

PF

NA

PF

DA

PF

Un

A

PF

Do

A

PF

TrA

PF

TeA

0

200

400

600

800

1000

1200

1400

0

5

10

15

20

25

30C

on

ce

ntr

ati

on

of

PF

AS

sin

So

il (

ng

/g)

0

10

20

30

40

50

600 Month

1.5 Month

3.5 Months

5.5 Months


6:2

diP

AP

6:2

/8:2

diP

AP

8:2

diP

AP

8:2

/10:2

diP

AP

10:2

diP

AP

10:2

/12:2

diP

AP

Co

nc

en

tra

tio

n o

f P

FA

Ss

in P

lan

ts (

ng

/g)

0

10

20

30

40

50

601.5 Month Plant

3.5 Month Plant

5.5 Month Plant

275

Figure B3. Concentrations of diPAPs and PFCAs in the soil and plants sampled from Treatment 3 (WWTP- and paper fiber biosolids-

amended soils).Each data point represents the arithmetic mean concentration of the triplicate (n = 3) sampling at each timepoint. The

error bar represents the standard error.

PF

BA

PF

Pe

A

PF

Hx

A

PF

Hp

A

PF

OA

PF

NA

PF

DA

PF

Un

A

PF

Do

A

PF

TrA

PF

Te

A

0

50

100

150

200

2500

2

4

6

8

10

Co

nc

en

tra

tio

n o

f P

FA

Ss

in S

oil

(n

g/g

)

0

20

40

60

80

1000 Month Soil

1.5 Month Soil

3.5 Month Soil

5.5 Month Soil

6:2

diP

AP

6:2

/8:2

diP

AP

8:2

diP

AP

8:2

/10

:2 d

iPA

P

10

:2 d

iPA

P

10

:2/1

2:2

diP

AP

Co

nc

en

tra

tio

n o

f P

FA

Ss

in P

lan

ts (

ng

/g)

0

5

10

15

20

25

30

351.5 Month Plant

3.5 Month Plant

5.5 Month Plant

WWTP- and Paper Fiber Biosolids-Amended Soil

276

Figure B3. Molar distribution (%) of diPAPs and PFCAs in the catch plates, soil, and plant of Treatments 2–4 microcosms.Each data

point represents the arithmetic mean percent of the triplicate (n = 3) sampling at each timepoint. The error bar represents the standard

error.

% (Mole Basis) Distribution in Soil-Plant Microcosm

0 20 40 60 80 100

PFOA

PFHpA

PFHxA

PFPeA

PFBA

6:2 diPAP

% Catch Plates

% Soil

% Plant

% (Mole Basis) Distribution in Soil-Plant Microcosm

0 20 40 60 80 100

PFTeAPFTrA

PFDoAPFUnA

PFDAPFNAPFOA

PFHpAPFHxAPFPeAPFBA

10:2/12:2 diPAP10:2 diPAP


6:2/8:2 diPAP

% Catch Plates

% Soil

% Plant


0 20 40 60 80 100

PFTeAPFTrA

PFDoAPFUnA

PFDAPFNAPFOA

PFHpAPFHxAPFPeAPFBA




% Catch Plates

% Soil

% Plant

WWTP- and Paper Fiber Biosolids-Amended Soil

6:2 diPAP-Spiked and WWTP Biosolids-Amended Soil

277

APPENDIX C

SUPPORTING INFORMATION FOR CHAPTER FIVE

Sorption of Perfluoroalkyl Phosphonates and Perfluoroalkyl Phosphinates in Soil

278


Table C1. Soil characteristics 283

Figure C1. Percent of PFPAs and PFPiAs (by mass) remaining in the aqueous phase

upon equilibration with 10 g, 5 g, and 2 g of Soil A. Mass balances of each PFPA and

PFPiA congener in the aqueous, soil, and container surface phases

284

Figure C2. Sorption kinetics of PFPAs and PFPiAs in seven different soils. Mass

balances of each PFPA and PFPiA congener in the aqueous, soil, and container surface

phases

285

Table C2. Multiple reaction monitoring (MRM) transitions and mass spectrometry

parameters for all target PFPAs and PFPiAs 286

Table C3. Limits of detection (LODs), limits of quantitation (LOQs), and matrix

recoveries (% Rec) in different phases for the PFPAs and PFPiAs 287

Figure C3. Concentrations of 8:2 FTUCA and PFOA in aqueous and soil phases

sampled after equilibration of Soil A with 0.01 M CaCl2 and 0.10 mM HgCl2 288

Figure C4. Aqueous and soil concentrations of PFPAs and PFPiAs in individually-

spiked soil-aqueous mixtures after 0.5 and 24 hours of equilibration 289

Table C4. Freundlich sorption coefficients (logKF), regression coefficients of sorption

isotherms (n), and distribution coefficients and their organic-carbon normalized analog

(logKd and logKOC) of the PFPAs and PFPiAs in each soil

290

Figure C5. Sorption isotherms of PFPAs and PFPiAs in seven different soils 291

Table C5. Distribution coefficients (logKd and logKOC) of each PFPA and PFPiA,

spiked either from the Masurf®-780 or as analytical-grade standards to Soil A

292

Table C6. Pearson’s correlation test results of the correlation between the logKd of each

PFPA and PPFiA and %OC, pH, and CEC 293

Figure C6. Dependence of distribution coefficients (logKd) on soil organic carbon 294

Figure C7. Dependence of distribution coefficients (logKd) on measured pH of the

aqueous phase equilibrated with each of the seven soils 295

Figure C8. Dependence of distribution coefficients (logKd) on soil cation exchange

capacity (CEC) 296

Figure C9. Desorption kinetics of the PFPAs and PFPiAs in Soil A. Dependence of

desorption coefficient (logKdes) on number of perfluorinated carbons in the PFPAs and

PFPiAs

297

279

EXPERIMENTAL

Chemicals.

Perfluorooctanoic acid (n-PFOA, >99%), perfluorononanoic acid (n-PFNA, >99%), 8:2

unsaturated fluorotelomer acid (8:2 FTUCA, >98%), C6 perfluorohexylphosphonate (C6 n-

PFPA, >99%), C8 perfluorooctylphosphonate (C8 n-PFPA, >99%), C10

perfluorodecylphosphonate (C10 n-PFPA, >99%), C6/C6 bis(perfluorohexyl)phosphinate

(C6/C6 n-PFPiA, >98%), C6/C8 perfluorohexylperfluorooctylphosphinate (C6/C8 n-PFPiA,

>98%), and C8/C8 bis(perfluorooctyl)phosphinate (C8/C8 n-PFPiA, >98%) were donated from

Wellington Laboratories Inc. (Guelph, ON). Mass-labeled internal standards were also donated

from Wellington Laboratories and they included: 13

C4-n-PFOA (>99%), 13

C5-n-PFNA (>99%),

and 13

C2-8:2 FTUCA (>98%).

Tetrabutylammonium hydrogen sulfate (TBAS, 99%) was purchased from Sigma Aldrich

(Oakville, ON). Methanol (Omnisolv, >99%), water (Omnisolv, >99%), and methyl-tert-butyl

ether (MTBE, Omnisolv, >99%) were purchased from EMD Chemicals, Inc. (Gibbstown, NJ).

Calcium chloride (CaCl2) was purchased from Fisher Chemicals, Fisher Scientific (Fairlawn,

NJ).

Using the analytical standards of C6, C8, and C10 PFPAs (>99%) and C6/C6, C6/C8,

and C8/C8 PFPiAs (>99%) from Wellington Laboratories (Guelph, ON), the percent

composition of the Masurf®

-780 was determined by standard addition to be 7% C6 PFPA, 6%

C8 PFPA, 3% C10 PFPA, 4% C6/C6 PFPiA, 5% C6/C8 PFPiA, and 1% C8/C8 PFPiA. This

percent composition, particularly for the PFPiAs, differs from that previously reported by D’eon

and Mabury1 (10%, C6 PFPA; 8%, C8 PFPA; 5%, C10 PFPA; 10%*, C6/C6 PFPiA; 6%*,

C6/C8 PFPiA; 5%*, C8/C8 PFPiA) and Lee and Mabury2 (8%, C6 PFPA; 7%, C8 PFPA; 5%,

C10 PFPA; 37%, C6/C6 PFPiA; 33%, C6/C8 PFPiA; 27%, C8/C8 PFPiA). It is important to note

that D’eon and Mabury determined their percent composition of the PFPiAs in the Masurf based

on peak area comparisons and not by quantification due to the lack of available analytical

standards for the PFPiAs at the time of analysis.1 These differences may be related to the use of

different lots of Masurf for preparing the standards, as well as, inter-batch variability between the

standards prepared here and those in the other two studies. Nevertheless, the percent

280

composition determined here was used to adjust the PFPA and PFPiA concentrations in this

study.

Extraction of Soil.

Briefly, 1 mL of 0.5 M TBAS solution (pH ~3) was added to 1 g of soil, followed by two

rounds of extraction with 4 mL of MTBE. The MTBE fractions were combined, evaporated to

dryness under nitrogen, and reconstituted in 1 mL of methanol.


Liquid Chromatography Details.

Chromatographic separation was performed using a Kinetex C18 column (50 x 4.6 mm,

2.6 μm; Phenomenex®, Torrance, CA). Analytes were quantified using an API4000 triple-

quadrupole mass spectrometer (MS/MS) (Applied Biosystems/MDS Sciex) in the negative

electrospray ionization mode, coupled to a Waters Acquity ultra-high pressure liquid

chromatography (UPLC) system. For the analysis of the PFPAs and PFPiAs, the samples were

injected as 30 μL injections and analyzed by the following gradient at 600 µL/min: the initial

solvent composition at t = 0 min. was 70:30 water:methanol, which changed to 5:95 over a

period of 5 min. at t = 5.00 min. and held for 2 min. to t = 7.00 min., before returning to the

initial composition of 70:30 water:methanol at t = 7.50 min. The column was allowed to

reequilibrate for 2.50 min. for a total run time of 10 min.

Mass Spectrometry Details.

A list of the analyte-specific multiple reaction monitoring (MRM) transitions and mass

spectrometry parameters for the studied PFPAs and PFPiAs is provided in Table C2.

281


Preparation of Matrix-matched Calibration Standards.

Matrix-matched calibration standards were prepared by extracting blank soils or aqueous

phases that have been equilibrated with the blank soils in the same manner as the Masurf-spiked

samples, followed by spiking of the Masurf at six different concentrations. No endogenous

contamination of any of the target PFPAs and PFPiAs was observed in these blank extracts at the

dilution factors used here.

Spike and Recovery Procedures in Aqueous CaCl2 Phase, Soil, and Polypropylene Containers.

For the aqueous phase, spike and recovery experiments were performed by adding 250

µg of Masurf into 50 mL of 0.01 M CaCl2 solution, and the samples were diluted with methanol

and injected directly onto the UPLC-MS/MS. For the soil, 100–250 µg of Masurf was added to

1 g of each soil type (A–G) that has previously been equilibrated with 0.01 M CaCl2 for four

hours prior to spiking, then extracted as described above. Analyte recovery from the container

walls was assessed by spiking 250 µg of Masurf into polypropylene tubes containing 50 mL of

MTBE and shaking for 30 min., after which the MTBE was evaporated to dryness and the empty

containers were rinsed with 25 mL of methanol. An aliquot of this methanol was then diluted

and analyzed directly on the UPLC-MS/MS. All spike and recovery experiments were

performed in triplicate (n = 3) and the resulting recoveries for each phase are provided in Table

C3.

282

LITERATURE CITED




Chemicals in Human Sera from United States Donors in 2009. Environ. Sci. Technol.

2011, 45, 8067–8074.

(3) Wascher, H. L.; Veale, P. T.; Odell, R. T. Will County Soils - Soil Report 80; University

of Illinois Agricultural Experiment Station: Urbana, Illinois, 1962.

(4) De Solla, S. R.; Martin, P. A. Toxicity of Nitrogenous Fertilizers to Eggs of Snapping

Turtles (Chelydra serpentina) in Field and Laboratory Exposures. Environ. Toxicol.

Chem. 2007, 26, 1890.

(5) Washington, J. W.; Ellington, J. J.; Jenkins, T. M.; Evans, J. J. Analysis of Perfluorinated

Carboxylic Acids in Soils: Detection and Quantitation Issues at Low Concentrations. J.

Chrom. A. 2007, 1154, 111–120.

(6) Soil Survey of Glades County, Florida; U.S. Department of Agriculture and Natural

Resources Conservation Service: Washington, DC, 1989.

283

Table C1. Soil characteristics.

Soil pH

%

Organic

Carbon

(OC)*

%

Sand**

%

Silt**

%

Clay**

CEC

(μmol/g)* Texture Sample Location

A

5.2 1.00 63 32 5 96 Sandy loam

Agricultural farm field in

Northumberland County, ON

Canada

B 6.6 1.10 25 38 37 335 Clay loam Highway 50 in Brampton, ON,

Canada

C 7.0 1.10 28 48 22 233 Loam Turtle garden in Burlington,

ON, Canada

D 3.8 3.60 53 41 6 238 Sandy loam Haliburton forest in Algonquin

Highlands, ON, Canada

E 4.4 10.20 58 20 22 113 Sandy clay

loam

Wooded area in Athens,

Georgia, US

F 6.1 2.63 10 61 27 242 Silt loam Army ammunition plant in

Joliet, Illinois, US

G 4.5 45.7 - - - - Peat Agricultural peat soil of Florida

Everglades, Florida, US *

% organic carbon (OC) and cation exchange capacity (CEC) data were measured by SGS AgriFood

Laboratories (Guelph, ON) for soils A–E. %OC and CEC data for soil F were obtained from the National

Cooperative Soil Survey, National Cooperative Soil Characterization Database

(http://ncsslabdatamart.sc.egov.usda.gov) and a soil report published by the University of Illinois Agricultural

Experiment Station.1 Data for soil G were obtained from the International Humic Substances Society website

(http://www.humicsubstances.org). **

Soil texture (%sand, %silt, %clay) was measured by SGS AgriFood Laboratories for soils A, B, and D.

Soil texture data for soils C, E, and F were obtained from De Solla and Martin,2 Washington et al.,

3 and the National

Cooperative Soil Survey, National Cooperative Soil Characterization Database

(http://ncsslabdatamart.sc.egov.usda.gov) respectively. As soil G is largely composed of organic matter (75–90%),4

it is considered a mineral-free soil and typically not analyzed for sand fractions.

284

Figure C1. Percent of PFPAs and PFPiAs (by mass) remaining in the aqueous phase upon equilibration with different masses of Soil

A (10 g, (A); 5 g, (B); 2 g, (C)) over time. Mass balances of each PFPA and PFPiA congener, as determined from their percent

distribution in the aqueous, soil, and container surface phases, were calculated after 0.5 (D) and 24 (E) hours of equilibration at the

1:10 soil:solution ratio. Each data point represents the arithmetic mean percent of the triplicate (n = 3) samples. The error bar

represents the standard error.

Time (hours)

0 20 40 60 80 100 120 140 160

% R

em

ain

ing

in

aq

ue

ou

s

ph

as

e (

by

ma

ss

)

0

20

40

60

80

100

120

140

C6 PFPA

C8 PFPA

C10 PFPA

C6/C6 PFPiA

C6/C8 PFPiA

C8/C8 PFPiA A. 1:5 Soil:Solution

0 20 40 60 80 100 120 140 160

B. 1:10 Soil:Solution

0 20 40 60 80 100 120 140 160

C. 1:25 Soil:Solution

C6

PF

PA

C8

PF

PA

C1

0 P

FP

A

C6

/C6 P

FP

iA

C6

/C8 P

FP

iA

C8

/C8 P

FP

iA

% D

istr

ibu

tio

n i

n

dif

fere

nt

ph

as

es

0

20

40

60

80

100

120

140 % Remaining in aqueous phase

% Adsorbed to soil

% Recovered from container walls

% Mass balance

C6

PF

PA

C8

PF

PA

C1

0 P

FP

A

C6

/C6 P

FP

iA

C6

/C8 P

FP

iA

C8

/C8 P

FP

iA

D. Mass balance of 1:10 soil:solution samples at t = 0.5 hrs

E. Mass balance of 1:10 soil:solution samples at t = 24 hrs

285

Figure C2. Sorption kinetics of PFPAs and PFPiAs in seven different soils. Percent of PFPAs and

PFPiAs (by mass) remaining in the aqueous phase upon equilibration with each soil type over time (left).

Mass balances of each PFPA and PFPiA congener, as determined from their percent distribution in the

aqueous, soil, and container surface phases (right). Each data point represents the arithmetic mean percent

of the triplicate (n = 3) samples. The error bar represents the standard error.

0 5 10 15 20 25

% R

em

ain

ing

in

aq

ue

ou

sp

ha

se

(b

y m

as

s)

0

20

40

60

80

100

120

140

C6 PFPA

C8 PFPA

C10 PFPA

C6/C6 PFPiA

C6/C8 PFPiA

C8/C8 PFPiA

0 5 10 15 20 25

% R

em

ain

ing

in

aq

ue

ou

s

ph

as

e (

by

ma

ss

)

0

20

40

60

80

100

120

140

% Remaining in aqueous phase

% Recovered from container walls

% Adsorbed to soil

% Mass balance

0 5 10 15 20 25

0

20

40

60

80

100

120

140

0 5 10 15 20 25

% R

em

ain

ing

in

aq

ue

ou

s

ph

as

e (

by

ma

ss

)

0

20

40

60

80

100

120

140

0 5 10 15 20 25

% R

em

ain

ing

in

aq

ue

ou

s

ph

as

e (

by

ma

ss

)

0

20

40

60

80

100

120

140

C6

PF

PA

C8

PF

PA

C1

0 P

FP

A

C6

/C6

PF

PiA

C6

/C8

PF

PiA

C8

/C8

PF

PiA

0 5 10 15 20 25

0

20

40

60

80

100

120

140

0 5 10 15 20 25

0

20

40

60

80

100

120

140

Time (hours)

0 5 10 15 20 25

0

20

40

60

80

100

120

140

C6

PF

PA

C8

PF

PA

C1

0 P

FP

A

C6

/C6

PF

PiA

C6

/C8

PF

PiA

C8

/C8

PF

PiA

(A) Control

(B)Soil A

(C) Soil B

(D) Soil C

(E) Soil D

(F) Soil E

(G) Soil F

(H) Soil G

286

Table C2. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target PFPAs and PFPiAs.

Analyte Acronym Mass

Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy,

CE

(V)

Collision

Cell Exit

Potential,

CXP (V)

Perfluorophosphonates (PFPAs) and perfluorophosphinates (PFPiAs)

C6 perfluorophosphonate C6 PFPA 399.0>79.0 40 -60 -75 -10



C6/C6 perfluorophosphinate C6/C6 PFPiA 701.0>401.0 40 -100 -75 -10


801.0>501.0 40 -100 -85 -10


287

Table C3. Limits of detection (LODs), limits of quantitation (LOQs), and matrix recoveries (% Rec) in different phases for the

PFPAs and PFPiAs. The LODs and LOQs in soil are reported on a dry weight (dw) basis. All spike and recovery experiments were

performed in triplicate (n = 3).

Analyte

Instrumental

(on column) CaCl2 aqueous phase

Polypropylene container

wall

LOD LOQ LOD LOQ % Rec (±SE) % Rec (±SE)

(pg) (ng/mL)

C6 PFPA 1.37 3.43 0.03 0.06 112 ± 5 87 ± 3

C8 PFPA 0.12 0.58 0.12 0.23 81 ± 4 94 ± 2

C10 PFPA 3.22 4.83 0.06 0.19 52 ± 2 91 ± 1

C6/C6 PFPiA 0.09 0.21 0.01 0.03 107 ± 4 90 ± 2

C6/C8 PFPiA 0.47 2.34 0.02 0.09 108 ± 6 95 ± 4

C8/C8 PFPiA 1.14 2.28 0.05 0.09 102 ± 12 94 ± 5

Analyte

Soil A Soil B Soil C Soil D Soil E Soil F Soil G

LOD LOQ % Rec

(±SE)

LOD LOQ % Rec

(±SE)

LOD LOQ % Rec

(±SE)

LOD LOQ % Rec

(±SE)

LOD LOQ % Rec

(±SE)

LOD LOQ % Rec

(±SE)

LOD LOQ % Rec

(±SE) (ng/g dw) (ng/g dw) (ng/g dw) (ng/g dw) (ng/g dw) (ng/g dw) (ng/g dw)

C6

PFPA 0.04 0.09 81 ± 2 0.04 0.10 61 ± 3 0.05 0.12 34 ± 3 0.09 0.19 36 ± 0 0.04 0.09 30 ± 2 0.04 0.10 75 ± 1 0.08 0.19 34 ± 4

C8

PFPA 0.003 0.02 81± 0 0.003 0.02 73 ± 5 0.004 0.02 48 ± 6 0.003 0.02 31 ± 3 0.003 0.02 68 ± 5 0.003 0.02 85 ± 3 0.01 0.03 49 ± 3

C10

PFPA 0.08 0.13 83 ± 0 0.09 0.14 73 ± 3 0.11 0.16 62 ± 5 0.09 0.13 47 ± 4 0.08 0.12 73 ± 4 0.10 0.15 87 ± 2 0.18 0.27 85 ± 1

C6/C6

PFPiA 0.002 0.01 85 ± 5 0.002 0.01 86 ± 2 0.003 0.01 87 ± 5 0.002 0.01 69 ± 13 0.002 0.01 86 ± 9 0.003 0.01 93 ± 6 0.01 0.01 104 ± 7

C6/C8

PFPiA 0.01 0.06 88 ± 2 0.01 0.07 93 ± 8 0.02 0.08 76 ± 10 0.01 0.06 64 ± 15 0.01 0.06 97 ± 14 0.01 0.07 109 ± 9 0.03 0.13 102 ± 7

C8/C8

PFPiA 0.03 0.06 105 ± 0 0.03 0.07 106 ± 17 0.04 0.08 68 ± 14 0.03 0.06 51 ± 15 0.03 0.06 84 ± 18 0.01 0.02 108 ± 8 0.06 0.13 99 ± 16

288

Figure C3. Concentrations of 8:2 FTUCA and PFOA in aqueous and soil phases sampled after 0.5

and 24 hours of equilibration of 5 g of Soil A in 0.01 M CaCl2 containing 0.10 mM HgCl2. Each

data point represents the arithmetic mean concentration of the triplicate (n = 3) samples. The error

bar represents the standard error.

8:2 FTUCA PFOA

Co

ncen

trati

on

in

aq

ueo

us p

hase (

ng

/mL

) o

r so

il p

hase (

ng

/g d

w)

0

20

40

60

80

100

120

0.5h aqueous phase

0.5h soil phase

24h aqueous phase

24h soil phase

289

Figure C4. Aqueous and soil concentrations of PFPAs and PFPiAs observed in individually-spiked soil-aqueous mixtures after 0.5 and 24 hours of

equilibration of Soil A with 0.01 M CaCl2 (left). Mass balances of each PFPA and PFPiA congener (right). Each data point represents the arithmetic mean

concentration or percent of the triplicate (n = 3) samples. The error bar represents the standard error.

C6 P

FP

A

C8 P

FP

A

C10 P

FP

A

C6/C

6 P

FP

iA

C6/C

8 P

FP

iA

C8/C

8 P

FP

iA

Co

nc

en

tra

tio

n i

n a

qu

eo

us

ph

as

e (

ng

/mL

) o

r s

oil p

ha

se

(n

g/g

dw

)

0

100

200

300

400

500

600

0.5h aqueous phase

0.5h soil phase

24h aqueous phase

24h soil phase

C6 P

FP

A

C8 P

FP

A

C10 P

FP

A

C6/C

6 P

FP

iA

C6/C

8 P

FP

iA

C8/C

8 P

FP

iA

% D

istr

ibu

tio

n o

f A

na

lyte

s

0

20

40

60

80

100

120

% Remaining in aqueous phase

% Adsorbed to soil

% Mass balance

290

Table C4. Comparison of distribution coefficients and their organic-carbon normalized analog (logKd and logKOC) of the PFPAs and PFPiAs, measured by

direct soil analysis and the aqueous loss method.

Soil Sorption

Parameters

C6 PFPA C8 PFPA C10 PFPA C6/C6 PFPiA C6/C8 PFPiA C8/C8 PFPiA

Direct Soil

Analysis

Aqueous

Loss

Method

Direct

Soil

Analysis

Aqueous

Loss

Method

Direct

Soil

Analysis

Aqueous

Loss

Method

Direct

Soil

Analysis

Aqueous

Loss

Method

Direct

Soil

Analysis

Aqueous

Loss

Method

Direct

Soil

Analysis

Aqueous

Loss

Method

A logKd ± SE -0.38±0.18 0.14±0.35 0.45±0.15 1.26±0.03 1.11±0.15 1.91±0.02 1.30±0.13 2.08±0.04 1.95±0.20 2.80±0.10 2.10±0.34 3.14±0.25

logKOC ± SE 1.62±0.18 2.14±0.35 2.45±0.15 3.26±0.03 3.11±0.15 3.91±0.02 3.30±0.13 4.08±0.04 3.95±0.20 4.80±0.10 4.10±0.34 5.14±0.25

B logKd ± SE -0.33±0.07 - 0.68±0.11 1.30±0.04 1.88±0.11 2.40±0.00 1.49±0.15 1.98±0.02 2.59±0.14 3.22±0.05 2.21±0.16 2.71±0.02

logKOC ± SE 1.63±0.07 - 2.64±0.11 3.26±0.04 3.84±0.11 4.36±0.00 3.45±0.15 3.94±0.02 4.55±0.14 5.18±0.05 4.17±0.16 4.67±0.02

C logKd ± SE 0.49±0.30 2.53±0.35 1.34±0.29 2.17±0.28 2.05±0.15 2.61±0.13 2.06±0.35 2.62±0.34 2.53±0.10 2.99±0.07 2.82±0.44 3.10±0.43

logKOC ± SE 2.45±0.30 4.49±0.35 3.30±0.29 4.12±0.28 4.01±0.15 4.57±0.13 4.02±0.35 4.58±0.34 4.49±0.10 4.95±0.07 4.78±0.44 5.06±0.43

D logKd ± SE 1.42±0.14 2.00±0.14 1.61±0.15 1.94±0.15 0.95±0.05 1.02±0.05 1.82±0.08 2.41±0.08 1.49±0.07 2.01±0.06 1.31±0.07 1.81±0.04

logKOC ± SE 2.86±0.14 3.45±0.14 3.06±0.15 3.39±0.15 2.40±0.05 2.46±0.05 3.26±0.08 3.85±0.08 2.94±0.07 3.46±0.06 2.76±0.07 3.26±0.04

E logKd ± SE 0.37±0.07 - 1.06±0.09 1.64±0.01 1.58±0.10 1.99±0.02 1.82±0.08 2.41±0.02 2.37±0.11 2.84±0.06 2.19±0.11 2.57±0.05

logKOC ± SE 1.30±0.07 - 2.05±0.09 2.45±0.01 2.58±0.10 2.98±0.02 2.82±0.08 3.40±0.02 3.36±0.11 3.83±0.06 3.18±0.11 3.56±0.05

F logKd ± SE 0.07±0.04 0.85±0.11 1.03±0.10 1.47±0.07 2.13±0.05 2.58±0.01 1.60±0.04 2.33±0.04 2.15±0.03 2.89±0.02 1.81±0.07 2.58±0.02

logKOC ± SE 1.65±0.04 2.43±0.11 2.61±0.10 3.05±0.07 3.71±0.05 4.16±0.01 3.18±0.04 3.91±0.04 3.73±0.03 4.47±0.02 3.39±0.07 4.16±0.02

G logKd ± SE 1.03±0.03 2.57±0.03 1.70±0.05 2.70±0.02 1.46±0.09 2.21±0.06 1.98±0.09 2.81±0.04 2.02±0.10 2.71±0.06 2.00±0.13 2.69±0.04

logKOC ± SE 1.37±0.03 2.91±0.03 2.04±0.05 3.04±0.02 1.80±0.09 2.55±0.06 2.32±0.09 3.15±0.04 2.36±0.10 3.05±0.06 2.34±0.13 3.03±0.04

Mean logKd ± SE 0.37±0.06 1.62±0.08 1.13±0.06 1.76±0.05 1.60±0.04 2.10±0.02 1.72±0.06 2.38±0.05 2.16±0.04 2.78±0.02 2.06±0.09 2.66±0.07

logKOC ± SE 1.84±0.06 3.08±0.08 2.59±0.06 3.22±0.05 3.06±0.04 3.57±0.02 3.19±0.06 3.84±0.05 3.62±0.04 4.25±0.02 3.53±0.09 4.13±0.07

- Kd and KOC were not obtained for C6 PFPA in soils B and E due to the presence of interfering artifacts during analysis

291

Table C5. Freundlich sorption coefficients (logKF), regression coefficients of sorption isotherms (n), and distribution coefficients and their organic-carbon

normalized analog (logKd and logKOC) measured for the PFPAs and PFPiAs in each soil.

Soil

Type

Sorption

Parameters

Analyte

C6 PFPA C8 PFPA C10 PFPA C6/C6 PFPiA C6/C8 PFPiA C8/C8 PFPiA

A

logKF ± SD -0.76 ± 0.12 -0.12 ± 0.38 0.66 ± 0.16 1.27 ± 0.20 1.82 ± 0.09 1.08 ± 0.13

n ± SD (R2) 0.86 ± 0.04 (1.00) 0.75 ± 0.17 (0.95) 0.79 ± 0.10 (0.98) 0.95 ± 0.14 (0.95) 0.92 ± 0.08 (0.98) 0.91 ± 0.14 (0.97)

logKd ± SE -0.38 ± 0.18 0.45 ± 0.15 1.11 ± 0.15 1.30 ± 0.13 1.95 ± 0.20 2.10 ± 0.34

logKOC ± SE 1.62 ± 0.18 2.45 ± 0.15 3.11 ± 0.15 3.30 ± 0.13 3.95 ± 0.20 4.10 ± 0.34

B

logKF ± SD -0.99 ± 0.77 -1.52 ± 0.52 - 0.04 ± 0.32 - -

n ± SD (R2) 0.76 ± 0.28 (0.88) 0.52 ± 0.21 (0.96) - 0.55 ± 0.20 (0.96) - -

logKd ± SE -0.33 ± 0.07 0.68 ± 0.11 1.88 ± 0.11 1.49 ± 0.15 2.59 ± 0.14 2.21 ± 0.16

logKOC ± SE 1.63 ± 0.07 2.64 ± 0.11 3.84 ± 0.11 3.45 ± 0.15 4.55 ± 0.14 4.17 ± 0.16

C

logKF ± SD 0.90 ± 0.15 -0.90 ± 0.17 - 1.25 ± 0.04 2.47 ± 0.05 -

n ± SD (R2) 1.17 ± 0.06 (0.99) 0.60 ± 0.06 (0.98) - 0.93 ± 0.02 (1.00) 1.05 ± 0.04 (0.99) -

logKd ± SE 0.49 ± 0.30 1.34 ± 0.29 2.05 ± 0.15 2.06 ± 0.35 2.53 ± 0.10 2.82 ± 0.44

logKOC ± SE 2.45 ± 0.30 3.30 ± 0.29 4.01 ± 0.15 4.02 ± 0.35 4.49 ± 0.10 4.78 ± 0.44

D

logKF ± SD 1.46 ± 0.09 1.55 ± 0.27 - 1.03 ± 0.24 1.48 ± 0.33 -

n ± SD (R2) 1.01 ± 0.04 (0.99) 0.87 ± 0.17 (0.94) - 0.72 ± 0.16 (0.96) 0.81 ± 0.22 (0.91) -

logKd ± SE 1.42 ± 0.14 1.61 ± 0.15 0.95 ± 0.05 1.82 ± 0.08 1.49 ± 0.07 1.31 ± 0.07

logKOC ± SE 2.86 ± 0.14 3.06 ± 0.15 2.40 ± 0.05 3.26 ± 0.08 2.94 ± 0.07 2.76 ± 0.07

E

logKF ± SD 0.71 ± 0.15 1.27 ± 0.18 1.59 ± 0.17 1.37 ± 0.30 2.01 ± 0.18 -

n ± SD (R2) 1.11 ± 0.06 (0.99) 1.03 ± 0.08 (0.98) 1.01 ± 0.10 (0.97) 0.70 ± 0.23 (0.93) 0.79 ± 0.16 (0.95) -

logKd ± SE 0.31 ± 0.07 1.06 ± 0.09 1.58 ± 0.10 1.82 ± 0.08 2.37 ± 0.11 2.19 ± 0.11

logKOC ± SE 1.30 ± 0.07 2.05 ± 0.09 2.58 ± 0.10 2.82 ± 0.08 3.36 ± 0.11 3.18 ± 0.11

F

logKF ± SD -0.09 ± 0.19 0.19 ± 0.39 0.96 ± 0.26 1.81 ± 0.13 1.71 ± 0.11 -

n ± SD (R2) 0.96 ± 0.07 (0.99) 0.73 ± 0.18 (0.95) 0.46 ± 0.26 (0.96) 1.03 ± 0.10 (0.97) 0.60 ± 0.14 (0.98) -

logKd ± SE 0.07 ± 0.04 1.03 ± 0.10 2.13 ± 0.05 1.60 ± 0.04 2.15 ± 0.03 1.81 ± 0.07

logKOC ± SE 1.65 ± 0.04 2.61 ± 0.10 3.71 ± 0.05 3.18 ± 0.04 3.73 ± 0.03 3.39 ± 0.07

G

logKF ± SD 0.33 ± 0.51 1.27 ± 0.06 1.45 ± 0.08 2.00 ± 0.15 1.99 ± 0.17 2.14 ± 0.09

n ± SD (R2) 0.80 ± 0.22 (0.91) 0.80 ± 0.04 (1.00) 0.95 ± 0.05 (0.99) 1.01 ± 0.13 (0.95) 0.94 ± 0.14 (0.95) 0.93 ± 0.12 (0.97)

logKd ± SE 1.03 ± 0.03 1.70 ± 0.05 1.46 ± 0.09 1.98 ± 0.09 2.02 ± 0.10 2.00 ± 0.13

logKOC ± SE 1.37 ± 0.03 2.04 ± 0.05 1.80 ± 0.09 2.32 ± 0.09 2.36 ± 0.10 2.34 ± 0.13

292

Figure C5. Sorption isotherms of PFPAs and PFPiAs in seven different soils. Equilibrium

concentrations measured in the soil and aqueous phases are denoted as Cs,e (ng/g dry weight (dw)) and

Caq,e (ng/mL) respectively. Each data point represents the arithmetic mean concentration of the triplicate

(n = 3) samples. The error bar represents the standard error.

log(Caq,e

, ng/mL)

-1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

1

2

3

4

5

log(Caq,e

, ng/mL)

-1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

1

2

3

4

5log(C

aq,e, ng/mL)

-2 -1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

0

1

2

3

4

5

C6 PFPA

C8 PFPA

C10 PFPA

C6C6 PFPiA

C6C8 PFPiA

C8C8 PFPiA

log(Caq,e

, ng/mL)

-1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

0

1

2

3

4

5

log(Caq,e

, ng/mL)

-1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

1

2

3

4

5

log(Caq,e

, ng/mL)

-1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

1

2

3

4

5 log(Caq,e

, ng/mL)

-1 0 1 2 3 4

log

(Cs

,e, n

g/g

dw

)

1

2

3

4

5

293

Table C6. Distribution coefficients (logKd and logKOC) measured for each PFPA and PFPiA, spiked

either from the commercial product, Masurf®-780, or as analytical-grade standards to Soil A.

Table S7. P-values and r-values from the parametric method, Pearson’s correlation test, to evaluate the

correlation between the logKd determined for each target PFPA and PPFiA and various soil-specific

parameters (i.e. %OC, pH, and CEC) (α = 0.05).

Analyte

Type of Comparison

logKd vs. %OC logKd vs. pH logKd vs. CEC

p-value r p-value r p-value r

C6 PFPA 0.0723 0.93 0.1576 -0.60 0.0036 0.98

C8 PFPA 0.0253 0.97 0.3735 -0.40 0.0023 0.98

C10 PFPA 0.7878 -0.21 0.0257 0.81 0.4897 0.36

C6/C6 PFPiA 0.0623 0.94 0.8095 -0.11 0.0043 0.95

C6/C8 PFPiA 0.2885 -0.71 0.0464 0.76 0.8718 0.09

C8/C8 PFPiA 0.031 -0.97 0.0788 0.70 0.6467 0.24

Analyte

Commercial product

Masurf®-780

Analytical PFPA and PFPiA

standards

logKd ± SE logKOC ± SE logKd ± SE logKOC± SE

C6 PFPA -0.38 ± 0.18 1.62 ± 0.18 -0.15 ± 0.11 1.85 ± 0.11

C8 PFPA 0.45 ± 0.15 2.45 ± 0.15 0.89 ± 0.12 2.89 ± 0.12

C10 PFPA 1.11 ± 0.15 3.11 ± 0.15 1.01 ± 0.13 3.01 ± 0.13

C6/C6 PFPiA 1.30 ± 0.13 3.30 ± 0.13 1.70 ± 0.11 3.70 ± 0.11

C6/C8 PFPiA 1.95 ± 0.20 3.95 ± 0.20 2.09 ± 0.10 4.09 ± 0.10

C8/C8 PFPiA 2.10 ± 0.34 4.10 ± 0.34 1.96 ± 0.03 3.96 ± 0.03

294

Figure C6. Dependence of distribution coefficients (logKd) on soil organic carbon. Each data point represents the arithmetic mean logKd of

the triplicate (n = 3) samples. The error bar represents the standard error.

0.00 0.01 0.02 0.03 0.04 0.05

log

Kd

-1

0

1

2

3

0.00 0.01 0.02 0.03 0.04 0.05 0.00 0.01 0.02 0.03 0.04 0.05

0.00 0.01 0.02 0.03 0.04 0.05

-1

0

1

2

3

Fraction of organic carbon

0.00 0.01 0.02 0.03 0.04 0.05 0.00 0.01 0.02 0.03 0.04 0.05



295

Figure C7. Dependence of distribution coefficients (logKd) on measured pH of the aqueous phase equilibrated with each of the seven soils.

Each data point represents the arithmetic mean logKd of the triplicate (n = 3) samples. The error bar represents the standard error.

3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5

log

Kd -1

0

1

2

3

3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5

3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5

-1

0

1

2

3

pH

3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5



296

Figure C8. Dependence of distribution coefficients (logKd) on soil cation exchange capacity (CEC). Each data point represents the arithmetic

mean logKd of the triplicate (n = 3) samples. The error bar represents the standard error.

50 100 150 200 250 300 350

log

Kd -1

0

1

2

3

50 100 150 200 250 300 350 50 100 150 200 250 300 350

50 100 150 200 250 300 350

-1

0

1

2

3

CEC (g/mol)

50 100 150 200 250 300 350 50 100 150 200 250 300 350



297

Figure C9. Desorption kinetics of the PFPAs and PFPiAs in Soil A (left). Dependence of desorption coefficient (logKdes) on number of

perfluorinated carbons in the PFPAs and PFPiAs (right). Each data point represents the arithmetic mean percent or logKdes of the triplicate (n

= 3) samples. The error bar represents the standard error.

Time (hours)

0 10 20 30 40 50 60

%D

es

orb

ed

fro

m s

oil (

by

mas

s)

0

20

40

60

80

100

120

140

C6 PFPA

C8 PFPA

C10 PFPA

C6/C6 PFPiA

C6/C8 PFPiA

C8/C8 PFPiA

Number of CF's

4 6 8 10 12 14 16 18

log

Kd

es

0.5

1.0

1.5

2.0

2.5

PFPA

PFPiA

298

APPENDIX D

SUPPORTING INFORMATION FOR CHAPTER SIX

Dietary Bioaccumulation of Perfluorophosphonates and Perfluorophosphinates in

Juvenile Rainbow Trout: Evidence of Metabolism of Perfluorophosphinates

299


Table D1. Dosed concentrations for all target PFPAs and PFPiAs in the fish feed 310

Figure D1. Fish whole-body and liver masses (g) during uptake and depuration

phases from control, PFPA-dosed, and PFPiA-dosed populations 311

Table D2. Multiple reaction monitoring (MRM) transitions and mass spectrometry

parameters for all target analytes 312

Table D3. Multiple reaction monitoring (MRM) transitions and mass spectrometry

parameters for all internal standards used for quantifying PFCAs 313

Figure D2. Sample chromatograms of PFPAs and PFPiAs in various tissues removed

from fish sampled on day 31 of uptake phase 314

Table D4a. Limits of detection (LODs), limits of quantification (LOQs), and matrix

recoveries in different tissues for the PFPAs and PFPiAs 315

Table D4b. Limits of detection (LODs), limits of quantification (LOQs), and matrix

recoveries for the PFCAs 316

Figure D3. Concentrations of C5-C11 PFCAs in control, PFPA-, and PFPiA-dosed

whole-fish homogenate extracts at different timepoints 317

Figure D4. Liver somatic indices (LSI, %) during uptake and depuration phases from

control, PFPA-dosed, and PFPiA-dosed populations 318

Table D5. Whole-body and liver growth parameters of juvenile rainbow trout

exposed to C6, C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs separately 319

Table D6a. P-values from Shapiro-Wilk W test to analyze fish whole-body and liver

masses from each treatment population for evidence of non-normality 320

Table D6b. P-values from the parametric method of grouped linear regression with

covariance analysis to compare the fish whole-body and liver growth rates among the

PFPA-dosed, PFPiA-dosed, and control populations

320

Table D7a. P-values from Shapiro-Wilk W test to analyze overall mean liver somatic

indices (LSIs) calculated from each treatment population for evidence of non-

normality

321

Table D7b. P-values from the parametric method of unpaired two-sample Student t-

test to compare the overall mean LSI calculated throughout the length of the

experiment between the control and each of the PFPA-dosed and PFPiA-dosed

population

321

Figure D5. Whole-body homogenate concentrations (ng/g wet wt) of C6, C8, and

C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs in rainbow trout during exposure

phase

322

Table D8. P-values from parametric method, Pearson’s correlation test, to assess

whether steady state was achieved within the last 4 to 6 timepoints of the exposure

phase for each analyte

323

Table D9. P-values and r-values from the parametric method, Pearson’s correlation

test to evaluate the correlation between the depuration half-life and logBMF observed

for each target PFPA and PFPiA and the number of perfluorinated carbons in their

corresponding structures

323

Figure D6. Concentrations (ng/g wet wt) of C6, C8, and C10 PFPAs ((A) PFPA-

dosed fish) and C6/C6, C6/C8, and C8/C8 PFPiAs ((B) PFPiA-dosed fish) in various

fish tissues collected on the last day (day 31) of the exposure phase

324

300

Figure D7. Ratios of liver-to-blood and liver-to-carcass concentrations for the C6,

C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs based on tissue

concentrations measured in rainbow trout collected on last day of exposure phase

325

Table D10. Concentrations of C6, C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8

PFPiAs in different fish tissues (ng/g ww) analyzed on the last day of the exposure

phase (day 31) and their corresponding liver-to-blood (LBR), liver-to-carcass (LCR),

and blood-to-carcass (BCR) ratios calculated based on these concentrations

326

Figure D8. Concentrations of C6, C8, and C10 PFPAs (ng/g wet wt) observed in

different tissue extracts removed from PFPiA-dosed fish sampled on the last day of the

exposure phase (day 31)

327

301

EXPERIMENTAL

Chemicals.

Perfluoropentanoic acid (n-PFPeA, >99%), perfluorohexanoic acid (n-PFHxA, >99%),

perfluoroheptanoic acid (n-PFHpA, >99%), perfluorooctanoic acid (n-PFOA, >99%),

perfluorononanoic acid (n-PFNA, >99%), perfluorodecanoic acid (n-PFDA, >99%),

perfluoroundecanoic acid (n-PFUnA, >99%), C6 perfluorohexylphosphonate (C6 n-PFPA,

>99%), C8 perfluorooctylphosphonate (C8 n-PFPA, >99%), C10 perfluorodecylphosphonate

(C10 n-PFPA, >99%), C6/C6 bis(perfluorohexyl)phosphinate (C6/C6 n-PFPiA, >98%),

C6/C8 perfluorohexylperfluorooctylphosphinate (C6/C8 n-PFPiA, >98%), and C8/C8

bis(perfluorooctyl)phosphinate (C8/C8 n-PFPiA, >98%) were donated from Wellington

Laboratories Inc. (Guelph, ON). Mass-labeled internal standards were also donated from

Wellington Laboratories and they included: 13

C2-n-PFHxA (>99%), 13

C4-n-PFOA (>99%),

13C5-n-PFNA (>99%),

13C2-n-PFDA (>99%), and

13C2-n-PFUnA (>99%).

Neat material (~1 mg for each congener) of C6 PFPA (>98%), C8 PFPA (>98%), C10

PFPA (>98%), C6/C6 PFPiA (>98%), C6/C8 PFPiA (>98%), and C8/C8 PFPiA (>98%) were

donated by Wellington Laboratories (Guelph, ON) to be used for dosing the fish feed.

Tetrabutylammonium hydrogen sulfate (TBAS, 99%) and ethyl 3-aminobenzoate

methanesulfonate (MS-222, 98%) were purchased from Sigma Aldrich (Oakville, ON). Sodium

bicarbonate (>99%) was purchased from ACP Chemicals Inc. (Montreal, QC). Methanol

(Omnisolv, >99%), water (Omnisolv, >99%), methyl-tert-butyl ether (MTBE, Omnisolv,

>99%), and acetone (Omnisolv, >99%) were purchased from EMD Chemicals, Inc.

(Gibbstown, NJ). Sodium heparin was purchased from LEO Pharma Inc. (Thornhill, ON) for

rinsing the syringes used for sampling whole blood from fish.

Food Preparation.

Three batches of ~100 g of commercial fish feed (Silver Cup 1.5 mm extruded floating

feed, Martin Mills Inc., Elmira, ON) were prepared for the separate dosing of PFPAs and

PFPiAs and the control feed. Each batch was placed in a 500 mL round bottom flask,

followed by the addition of small volumes (<150 µL) of either a mixed standard of C6, C8,

and C10 PFPAs or C6/C6, C6/C8, and C8/C8 PFPiAs dissolved in methanol and 150 mL of

302

acetone. After shaking the mixture for 1 hour followed by 15 minutes of sonication, the bulk

of the acetone solvent was removed by rotary evaporation in a 50oC water bath. The fish feed

were then left to dry overnight in the oven at 60oC. The control feed was treated with acetone,

as described above, except without the spiking of the PFPAs or PFPiAs. The dosed and

control feed were stored in the dark and at room temperature.

Extraction of Dosed and Control Feed.

Approximately 0.5 g of each of the dosed and control feed were extracted in triplicate

(n = 3) with two sequential additions of 4 mL of methanol. After evaporating the combined 8

mL of methanol to dryness under nitrogen, the sample was reconstituted in 1 mL methanol

and filtered through 0.25 µm nylon filters (Chromatographic Specialties, Brockville, ON) into

low-temperature cryo vials (VWR International Ltd., Mississauga, ON). The concentrations

in the PFPA- and PFPiA-dosed feed were 485 ± 28 ng/g C6 PFPA, 474 ± 37 ng/g C8 PFPA,

and 533 ± 37 ng/g C10 PFPA; and 468 ± 12 ng/g C6/C6 PFPiA, 503 ± 21 ng/g C6/C8 PFPiA,

and 420 ± 12 ng/g C8/C8 PFPiA respectively (Table D1). The PFPAs and PFPiAs were not

detected in the control feed. These dosing concentrations are consistent with those used in

similar experiments performed by Martin et al. (1) and De Silva et al. (2).

Extraction of Tissue Samples for PFPAs, PFPiAs, and PFCAs.

Briefly, 1 mL of 0.5M TBAS solution (pH ~3) was added to each subsample of whole-

fish homogenate (0.5–1g), liver (~0.1 g), kidneys (~0.05–0.1 g), heart (~0.01 g), gills (~0.5–

0.7 g), or whole blood (~0.05–0.2 g), followed by extraction with two 4 mL aliquots of

MTBE. The MTBE aliquots were combined, evaporated to dryness under nitrogen, and

reconstituted in 0.5–1 mL of methanol. The livers, kidneys, gills, and fish carcasses were

homogenized in 1–2 mL of TBAS first before extraction. One (n = 1) procedural blank

(HPLC grade water) was extracted with each sampling timepoint during the uptake and

depuration phase.

303



Chromatographic separation was performed using a Kinetex C18 column (50 x 4.6

mm, 2.6 μm; Phenomenex®, Torrance, CA). Analyte quantitation was performed using an

API4000 triple-quadrupole mass spectrometer (Applied Biosystems/MDS Sciex) in the

negative electrospray ionization mode, coupled to a Waters Acquity UPLC system. Two high

performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) methods

were used for the analysis of the target analytes.

For the analysis of the PFPAs and PFPiAs, the samples were injected as 30 µL

injections and analyzed by the following gradient method at 600 µL/min: the initial solvent

composition at t = 0 min. was 70:30 water: methanol, which changed to 5:95 over a period of

5 min. at t = 5.00 min. and held for 2 min. to t = 7.00 min., before returning to the initial

composition of 70:30 water:methanol at t = 7.50 min. The column was allowed to


For the analysis of PFPeA, PFHxA, PFHpA, PFOA, PFNA, PFDA, and PFUnA, the

samples were injected as 25 µL injections and analyzed by the following gradient method at

600 µL/min: the initial solvent composition at t = 0 min. was 65:35 water:methanol, which

changed to 5:95 over a period of 3 min. at t = 3.00 min. and held for 2 min. to t = 5.00 min.,

before returning to the initial composition of 65:35 water:methanol at t = 5.50 min. The

column was allowed to reequilibrate for 2.50 min. for a total run time of 8 min.


A list of the analyte-specific multiple reaction monitoring (MRM) transitions and mass

spectrometry parameters for all target analytes and their corresponding internal standards is

provided in Tables D2 and D3. The PFPAs fragment exclusively to PO3- (79 m/z) (3), while

the PFPiAs fragment to [F(CF2)xPO2F]- (4).

304


Preparation of Matrix-matched Calibration Standards.

Matrix-matched calibration standards were prepared by extracting control fish tissue

extracts (i.e. whole-fish homogenate, liver, kidneys, gills, heart, and whole blood) in the same

manner as the experimental samples, followed by spiking of the target PFPAs and PFPiAs at

six different concentrations. No endogenous contamination of the C10 PFPA and PFPiAs was

observed in these control tissue extracts, whereas background concentrations of the C6 and C8

PFPAs were determined in these control matrices and used to correct the concentrations of the

matrix-matched standards.

Spike and Recovery Procedures in Different Fish Tissues.

Spike and recovery experiments were performed in triplicate (n = 3) by adding 30 ng

each of the PFPAs and PFPiAs or 10 ng of the C5–C11 PFCAs into ~0.5 g whole-body

homogenate subsamples prepared from extra control fish, and the samples were extracted and

analyzed as described above. For the tissue distribution study, spike and recovery

experiments were also performed in triplicate (n = 3) by adding 1–5 ng of the PFPAs and

PFPiAs into liver, kidneys, gills, heart, and whole blood subsamples prepared from extra

control fish, and the samples were extracted and analyzed as described above.

Monitoring for Production of PFCAs in the Dosed Fish.

The Canadian government recently listed different chain lengths of PFPAs and PFPiAs

as potential precursors to long chain PFCAs (≥8 carbons) (5). Metabolism of the C6, C8, and

C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs would most likely occur at the carbon–

phosphorus bond, which would result in the release of a perfluorohexane (C6),

perfluorooctane (C8), or perfluorodecane (C10) tail. As such, the C5–C11 PFCAs were

monitored in the dosed rainbow trout at occasional timepoints in case these perfluorocarbons

undergo further biotransformation to the PFCAs. No distinct increase in PFCA concentrations

was observed in either the dosed or control fish, although significant PFDA contamination

was observed in the control fish, the reason for which is unknown (Figure D3). As none of

the dosed PFPA and PFPiAs congeners were detected in the control fish, the observed PFDA

305

contamination is likely due to a water-borne source, although water samples were not

analyzed to confirm this. These results suggest PFPAs and PFPiAs will not biotransform to

the PFCAs in rainbow trout.

Data Analysis.

Estimation of time to achieve 90% steady-state in the exposure phase.

As described by Martin et al. (1), the time to reach 90% steady-state (tss, day) can be

estimated for each analyte by rearranging the kinetic rate equation as follows:

(1) Cfish/Cfood = (α·F/kd)·[1 – exp(-kd·tss)]

(2) Cfish/Cfood = BMF·[1 – exp(-kd·tss)]

(3) 0.90·BMF = BMF·[1 – exp(-kd·tss)]

(4) 0.10 = exp(-kd·tss)

(5) tss = ln(0.10)/(-kd)

where Cfish is the growth-corrected whole-body concentration, F is the feeding rate, Cfood is

the food concentration, BMF is the biomagnification factor, and kd is the depuration rate

constant for each analyte.

Statistical Analysis.

Analyte concentrations observed below their corresponding LODs in the depuration

phase were imputed as the LOD divided by square root of two, so that they can be fitted as

nonzero values to the first-order decay model described above for calculating kd and t1/2. All

data were tested for evidence of non-normality using the Shapiro-Wilk W test (p-values in

Tables D6a, D7a) to determine whether they should be analyzed using parametric (normal

distribution) or nonparametric (non-normal distribution) methods.

306

Comparison of fish whole-body and liver growth rates between control and dosed

populations.

Fish whole-body mass data from the control population (p = 0.0127) showed evidence

of non-normality, while the mass data from the PFPA-dosed (p = 0.0599), and PFPiA-dosed

(p = 0.5816) populations showed no evidence of non-normality (Table D6a). Reanalysis of

the logarithmically transformed data from all three populations with the Shapiro-Wilk W test

showed no evidence of non-normality (p = 0.0884 – 0.9489, Table D6a); therefore, the whole-

body growth rates among the three populations were statistically compared using the

parametric method of grouped linear regression with covariance analysis. The test for the

overall difference between the slopes of the three populations was significant (p = 0.0198),

which is mainly due to the statistically significant differences observed in the whole-body

growth rates between the control and PFPA-dosed populations (p = 0.0134) and the control

and PFPiA-dosed populations (p = 0.0161) (Table D6b). For the fish liver mass data, the

Shapiro-Wilk W test results showed evidence of non-normality for both the control (p =

0.0290) and PFPA-dosed populations (p = 0.0356), but no evidence of non-normality for the

PFPiA-dosed population (p = 0.8024) (Table D6a). Logarithmic transformation of the liver

mass data from all three populations resulted in Shapiro-Wilk W test results of no evidence of

non-normality (p = 0.1132–0.6223, Table D6a). Grouped linear regression with covariance

analysis of the liver mass data showed statistically significant differences in the liver growth

rates observed between the control and PFPA-dosed populations (p = 0.0048) and the control

and PFPiA-dosed populations (p = 0.0164) (Table D6b).

Comparison of liver somatic indices (LSIs) between control and dosed populations.

The liver somatic indices calculated from the whole-body and liver data from the

control, PFPA-dosed, and PFPiA-dosed populations all showed no evidence of non-normality

(p = 0.1884–0.8867, Table D7a); therefore, the parametric unpaired two-sample Student t-test

was used to compare the overall mean LSI calculated throughout the experiment between the

control and each of the dosed populations. No significant difference was observed in the

mean LSIs calculated between the control and the dosed populations (Control vs. PFPA-

dosed, p = 0.0841 (two-sided); Control vs. PFPiA-dosed, p = 0.5691 (two-sided), Table D7b).

307

Correlation between half-life and perfluorinated carbon chain length and between logBAF

and perfluorinated carbon chain length for the target PFPAs and PFPiAs detected in whole-

body homogenates.

Results from Pearson’s correlation tests showed the whole-body depuration half-lives

and logBAFs calculated for the PFPAs and PFPiAs detected in the rainbow trout were

correlated with perfluorinated carbon chain length (Table D9).

Partitioning of PFPAs and PFPiAs in Different Tissue Compartments.

Liver-to-blood (LBRs), liver-to-carcass (LCRs), and blood-to-carcass (BCRs) ratios

(Figure D6, Table D10) were calculated to evaluate partitioning of the PFPAs and PFPiAs in

these compartments. LBRs for the PFPAs (0.61–5.02) and PFPiAs (3.24–4.63) (Figure D6,

Table D10) detected in the day 31 tissues were similar to the LBRs of PFPAs and PFPiAs

observed in rats (0.02–39).(4) In rainbow trout, the C10 PFPA and all three PFPiA congeners

exhibited preferential partitioning into the liver from blood based on their LBRs (>1), while

no distinct preference was observed for the liver-to-blood partitioning of the C6 and C8

PFPAs (LBRs ≤1), as was also observed for these analytes in rats (4). The LCRs calculated

here for the PFPAs (37.70–138.27) and the PFPiAs (7.18–7.99) (Figure D6, Table D10) were

within the range of the liver-to-muscle ratios (LMRs) observed for PFOS (61.5) and the C11–

C13 PFCAs (11.1–63.4) in Chinese sturgeon (6). Similarly, the BCRs calculated here for the

PFPAs and PFPiAs all exceeded 1 (Table D10). Together these ratios (>1) suggest PFPAs

and PFPiAs are similar to other PFAAs in their tendency to predominate in proteinaceous

compartments like the liver and blood, potentially through interactions with proteins, as was

observed between serum albumin and PFOS (7) and PFOA (8).

Both the fish LBRs and LCRs of the PFPAs were observed to increase with chain

length, but this trend was not conserved for the PFPiAs (Figure D7). The plateau observed in

the lower LBRs and LCRs of PFPiAs mimics the deviation from the linear relationship

between uptake rates of water-borne PFAs in rainbow trout and chain length observed by

Martin et al. (9), in which C14 PFCA (MW 714 amu), the most hydrophobic PFA tested, was

taken up to a lesser extent than expected based on extrapolation from the liver, blood, and

carcass concentrations of the shorter PFCAs. Reduced uptake of hydrophobic organic

contaminants with MW in excess of 600 amu (10) is well documented (11–13), and is

presumably due to size-based exclusion during membrane permeation. Here, the MW

308

threshold, above which uptake into the liver is no longer directly proportional to chain length,

occurs between 600 amu (C10 PFPA) and 702 amu (C6/C6 PFPiA).

Calculation of PFPA metabolite yield.

The production yield of C6 and C8 PFPA metabolites in the PFPiA-dosed fish was

quantitatively expressed as the percentage of the molar sum of the accumulated two parent

PFPiAs containing the corresponding perfluorocarbon tail of equal chain length.

% C6 PFPA yield = (moles of C6 PFPA observed at each timepoint) x 100

(moles of C6/C6 PFPiA + moles of C6/C8 PFPiA)

% C8 PFPA yield = (moles of C8 PFPA observed at each timepoint) x 100

(moles of C6/C8 PFPiA + moles of C8/C8 PFPiA)

The PFPA metabolite yields were calculated at each timepoint throughout the

experiment and plotted against time in Figure 2. It is important to treat these yields

conservatively as they rely on the assumption that the metabolism of both PFPiA congeners

(e.g. C6/C6 and C6/C8 PFPiAs) is contributing equally to the production of the corresponding

PFPA metabolite (e.g. C6 PFPA). These yields also do not account for how metabolic

depuration and depuration by excretion may affect each other.

Literature.


Perfluorinated Acids in Juvenile Rainbow Trout (Oncorhynchus mykiss). Environ.

Toxicol. Chem. 2003, 22, 189–195.

(2) De Silva, A. O.; Benskin, J. P.; Martin, L. J.; Arsenault, G.; McCrindle, R.; Riddell, N.;

Martin, J. W.; Mabury, S. A. Disposition of Perfluorinated Acid Isomers in Sprague-

Dawley Rats; Part 2: Subchronic Dose. Environ. Toxicol. Chem. 2009, 28, 555.


Perfluorinated Phosphonic Acids in Canadian Surface Waters and Wastewater

Treatment Plant Effluent: Discovery of a New Class of Perfluorinated Acids. Environ.

Toxicol. Chem. 2009, 28, 2101.

309

(4) D’eon, J. C.; Mabury, S. A. Uptake and Elimination of Perfluorinated Phosphonic

Acids in the Rat. Environ. Toxicol. Chem. 2010, 29, 1319–1329.

(5) Draft Ecological Screening Assessment Report - Long-Chain (C9-C20)

Perfluorocarboxylic Acids, their Salts, and their Precursors; Environment Canada,

2010.

(6) Peng, H.; Wei, Q.; Wan, Y.; Giesy, J.P.; Li, L.; Hu, J. Tissue Distribution and Maternal

Transfer of Poly- and Perfluorinated Compounds in Chinese Sturgeon (Acipenser

sinensis): Implications for Reproductive Risk. Environ. Sci. Technol. 2010, 44, 1868–

1874.

(7) Jones, P. D.; Hu, W.; De Coen, W.; Newsted, J. L.; Giesy, J. P. Binding of

Perfluorinated Fatty Acids to Serum Proteins. Environ. Toxicol. Chem. 2003, 22, 2639.

(8) Han, X.; Snow, T. A.; Kemper, R. A.; Jepson, G. W. Binding of Perfluorooctanoic Acid

to Rat and Human Plasma Proteins. Chem. Res. Toxicol. 2003, 16, 775–781.

(9) Martin, J. W.; Mabury, S. A.; Solomon, K. R.; Muir, D. C. G. Bioconcentration and

Tissue Distribution of Perfluorinated Acids in Rainbow Trout (Oncorhynchus mykiss).


(10) Niimi, A. J.; Oliver, B. G. Influence of Molecular Weight and Molecular Volume on

Dietary Absorption Efficiency of Chemicals by Fishes. Can. J. Fish. Aquat. Sci. 1988,

45, 222–227.

(11) Fisk, A. T.; Bergman, Åk.; Cymbalisty, C. D.; Muir, D. C. G. Dietary Accumulation of

C12- and C16-Chlorinated Alkanes by Juvenile Rainbow Trout (Oncorhynchus

mykiss). Environ. Toxicol. Chem. 1996, 15, 1775–1782.

(12) Gobas, F.; Muir, D.; Mackay, D. Dynamics of Dietary Bioaccumulation and Faecal

Elimination of Hydrophobic Organic Chemicals in Fish. Chemosphere. 1988, 17, 943–

962.

(13) Fisk, A. T.; Norstrom, R. J.; Cymbalisty, C. D.; Muir, D. C. G. Dietary Accumulation

and Depuration of Hydrophobic Organochlorines: Bioaccumulation Parameters and

Their Relationship with the Octanol/Water Partition Coefficient. Environ. Toxicol.

Chem. 1998, 17, 951–961.

310

Table D1. Dosed concentrations for all target PFPAs and PFPiAs in the fish feed.

Target Analyte Acronym Dosed Concentration

(ng/g)

C6 perfluorophosphonate C6 PFPA 485 ± 28



C6/C6 perfluorophosphinate C6/C6 PFPiA 468 ± 12



311

Figure D1. Fish whole-body and liver masses (g) during uptake and depuration phases from

control (), PFPA-dosed (), and PFPiA-dosed () populations. Each data point represents

the arithmetic mean (n = 3) of the triplicate sampling at each timepoint, except for the last two

timepoints during which the control fish were sampled in duplicate (n = 2) and the dosed fish

were sampled in triplicate (n = 3). Each error bar represents the standard error.

Time (day)

0 10 20 30 40 50 60 70

Ma

ss

of

fis

h (

g)

0.0

0.2

0.4

5.0

10.0

15.0

20.0

25.0

30.0

Whole body masses from PFPA-dosed population

Whole body masses from PFPiA-dosed population

Whole body masses from Control population

Liver masses from PFPA-dosed population

Liver masses from PFPiA-dosed population

Liver masses from Control population

Uptake Depuration

312

Table D2. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.


Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy,

CE

(V)

Collision

Cell Exit

Potential,

CXP (V)

Perfluorophosphonates (PFPAs) and perfluorophosphinates (PFPiAs)






801.0>501.0 40 -100 -85 -10


Perfluorocarboxylates (PFCAs)

Perfluoropentanoate PFPeA (C5) 262.8>218.97 20 -20 -13 -15

Perfluorohexanoate PFHxA (C6) 312.8>268.9 20 -20 -13 -15

Perfluoroheptanoate PFHpA (C7) 362.8>319.0 20 -27 -13 -15

Perfluorooctanoate PFOA (C8) 413.0>368.9 20 -35 -15 -15

Perfluorononanoate PFNA (C9) 462.9>419.0 20 -35 -15 -15

Perfluorodecanoate PFDA (C10) 513.0>468.9 20 -45 -15 -15

Perfluoroundecanoate PFUnA (C11) 562.8>519.0 20 -45 -15 -15

313

Table D3. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all internal standards used for

quantifying PFCAs.

Target Analyte Internal

Standard

Mass

Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy, CE

(V)

Collision Cell

Exit Potential,

CXP (V)

Perfluorocarboxylates (PFCAs)

PFPeA (C5) 13

C2-PFHxA 314.8>269.8 20 -20 -13 -15

PFHxA (C6) 13

C2-PFHxA 314.8>269.8 20 -20 -13 -15

PFHpA (C7) 13

C4-PFOA 417.0>372.0 20 -35 -15 -15

PFOA (C8) 13

C4-PFOA 417.0>372.0 20 -35 -15 -15

PFNA (C9) 13

C5-PFNA 468.0>423.0 20 -35 -15 -15

PFDA (C10) 13

C2-PFDA 515.0>470.0 20 -45 -15 -15

PFUnA (C11) 13

C2-PFUnA 564.8>520.0 20 -45 -15 -15

314

Figure D2. Sample chromatograms of PFPAs and PFPiAs in various tissues removed from fish sampled on day 31 of uptake phase.

Time (min)

0 2 4 6 8 10

0

60000

120000

In PFPiA-dosed fishC8/C8 PFPiA901 > 501Area 270000

Re

sp

on

se

0

8000

16000

0

8000

16000

0

8000

16000

0

60000

120000

0

60000

120000

In PFPA-dosed fishC6 PFPA399 > 79Area 21600



Day 31 of uptake phase inwhole-body homogenate samples



Day 31 of uptake phase in liver samples

0

60000

120000

0

60000

120000

0

60000

120000

0

250000

500000

0

250000

500000

0 2 4 6 8 10

0

250000

500000







0

6000

12000

Day 31 of uptake phase in gill samples

0

6000

12000

0

6000

12000

0

150000

300000

0

150000

300000

0 2 4 6 8 10

0

150000

300000







Day 31 of uptake phasein blood samples

0

500

1000

0

500

1000

0

500

1000

0

40000

80000

0

40000

80000

0 2 4 6 8 10

0

40000

80000







315

Table D4a. Limits of detection (LODs), limits of quantification (LOQs), and matrix recoveries in different tissues for the PFPAs and

PFPiAs. The LODs and LOQs are reported on a wet weight (ww) basis.

Analyte

Instrumental

(on column)

Whole-fish homogenate Liver Gills Blood

Method Recovery

(%) Method

Recovery

(%) Method

Recovery

(%) Method

Recovery

(%)

LOD LOQ LOD LOQ (n = 3) LOD LOQ (n = 3) LOD LOQ (n = 3) LOD LOQ (n = 3)

(pg) (ng/g ww) (ng/g ww) (ng/g ww) (ng/g ww)

C6 PFPA 0.75 1.50 0.02 0.04 86 ± 1 0.80 2.01 86 ± 5 0.20 0.50 84 ± 2 0.14 0.28 87 ± 6

C8 PFPA 0.75 1.50 0.02 0.10 98 ± 3 0.04 0.20 113 ± 16 0.01 0.05 73 ± 15 0.06 0.14 87 ± 19

C10 PFPA 1.50 7.50 0.21 0.42 112 ± 12 0.40 2.01 126 ± 33 0.01 0.05 78 ± 11 0.06 0.28 74 ± 7

C6/C6 PFPiA 0.15 0.30 0.02 0.04 82 ± 3 0.04 0.08 102 ± 16 0.01 0.02 81 ± 5 0.03 0.06 80 ± 4

C6/C8 PFPiA 0.30 0.75 0.04 0.10 95 ± 7 0.04 0.08 116 ± 23 0.02 0.05 76 ± 8 0.03 0.06 82 ± 6

C8/C8 PFPiA 0.30 0.75 0.04 0.10 105 ± 17 0.04 0.08 110 ± 22 0.01 0.02 75 ± 2 0.03 0.06 89 ± 8

Analyte

Kidneys Heart

Method Recovery

(%) Method

Recovery

(%)

LOD LOQ (n = 3) LOD LOQ (n = 3)

(ng/g ww) (ng/g ww)

C6 PFPA 0.48 0.96 94 ± 3 5.88 11.76 86 ± 3

C8 PFPA 0.19 0.96 85 ± 11 2.94 11.76 88 ± 9

C10 PFPA 0.19 0.96 74 ± 14 5.88 29.41 88 ± 11

C6/C6 PFPiA 0.10 0.19 82 ± 4 0.59 1.18 84 ± 6

C6/C8 PFPiA 0.10 0.19 89 ± 1 1.18 2.94 89 ± 6

C8/C8 PFPiA 0.10 0.19 90 ± 13 0.59 2.94 101 ± 8

316

Table D4b. Limits of detection (LODs), limits of quantification (LOQs), and matrix recoveries for the

PFCAs.

Analyte

Whole-fish homogenate

Instrumental

(on column) Method

Recovery (%)

(n = 3) LOD LOQ LOD LOQ

(pg) (ng/g)

PFPeA (C5) 0.25 1.25 0.04 0.20 68 ± 10

PFHxA (C6) 0.13 0.25 0.02 0.04 112 ± 8

PFHpA (C7) 0.13 0.25 0.02 0.04 82 ± 9

PFOA (C8) 0.13 0.25 0.02 0.04 93 ± 14

PFNA (C9) 0.13 0.25 0.02 0.04 98 ± 6

PFDA (C10) 0.13 0.25 0.02 0.04 92 ± 18

PFUnA (C11) 0.13 0.63 0.02 0.10 123 ± 13

317

Figure D3. Concentrations of C5-C11 PFCAs in control (A), PFPA- (B), and PFPiA-dosed (C) whole-fish homogenate extracts at

different timepoints. Each data point represents the arithmetic mean (n = 3) of the triplicate sampling at each timepoint, except for the last

two timepoints during which the control fish were sampled in duplicate (n = 2) and the dosed fish were sampled in triplicate (n = 3). Each

error bar represents the standard error.

Time (day)

0 10 20 30 40 50 60

Co

nce

ntr

ati

on

in

fis

h (

ng

/g w

w)

0

10

20

30

40

50

PFPeA

PFHxA

PFHpA

PFOA

PFNA

PFDA

PFUnA

0 10 20 30 40 50 60

0

1

2

3

4

5

0 10 20 30 40 50 60

0

1

2

3

4

5


(A) Control fish (B) PFPA-dosed fish (C) PFPiA-dosed fish

318

Figure D4. Liver somatic indices (LSI, %) during uptake and depuration phases from control (),

PFPA-dosed (), and PFPiA-dosed () populations. Each data point represents the arithmetic mean (n

= 3) of the triplicate sampling at each timepoint, except for the last two timepoints during which the

control fish were sampled in duplicate (n = 2) and the dosed fish were sampled in triplicate (n = 3).

Each error bar represents the standard error.

Time (day)

0 10 20 30 40 50 60 70

Liv

er

so

mati

c in

de

x (

LS

I, %

)

0.0

0.5

1.0

1.5

2.0

2.5

PFPA-dosed population

PFPiA-dosed population

Control population

Uptake Depuration

319

Table D5. Whole-body and liver growth parameters (mean ± standard error) of juvenile rainbow trout exposed to C6, C8, and C10

PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs separately.

a The growth rates were calculated by fitting all whole-body and liver mass data to an exponential model (ln(mass, g) = a + bt, where a is a constant, b is the growth

rate (/day) and t is the time (day). The coefficients of correlation, r, for the model are shown in parentheses. b The liver somatic index (LSI) was calculated as the ratio of the fish liver mass to the whole-body mass. The LSIs shown here are the overall means of the LSI

calculated at each timepoint for each population.

Population

Growth Rate (/day) (r)a

Fish Mass (g) Liver

Somatic

Index

(LSI, %)b

%

Mortality Whole-body Liver Predose

(1 day) Day 63

Control 0.0163 ± 0.0018

(0.94)

0.0161 ± 0.0026

(0.88) 6.58 ± 0.33 20.87 ± 9.51 1.24 ± 0.04 0

PFPA-dosed 0.0094 ± 0.0023

(0.77)

0.0059 ± 0.0022

(0.63) 6.67 ± 0.73 13.66 ± 1.59 1.39 ± 0.07 0

PFPiA-dosed 0.0096 ± 0.0013

(0.91)

0.0075 ± 0.0024

(0.68) 6.02 ± 0.49 13.08 ± 1.39 1.21 ± 0.04 0

320

Table D6a. P-values from Shapiro-Wilk W test to analyze fish whole-body and liver masses from each

treatment population for evidence of non-normality (α = 0.05).

Treatment

Population Type of Sample

p-values

Data without log transformation Log-transformed data

PFPA-dosed Whole-body 0.0599

0.0884

Liver 0.0356 0.1132

PFPiA-dosed Whole-body 0.5816 0.9489

Liver 0.8024 0.1745

Control Whole-body 0.0127 0.1578

Liver 0.0290 0.6223

Table D6b. P-values from the parametric method of grouped linear regression with covariance analysis

to compare the fish whole-body and liver growth rates between the two dosed populations and the

control population (α = 0.05). This method provides an analysis of variance that shows whether or not

there is a significant difference between the growth rates calculated from each treatment population as a

whole, and then further compares all of the growth rates individually. In each cell, the top row

represents the test performed on the fish whole-body masses among the three treatment populations and

the bottom row represents the test performed on the fish liver masses among the three treatment

populations.

Whole-body PFPA-dosed PFPiA-dosed

Liver

Control 0.0134 0.0161

0.0048 0.0164

Overall Difference 0.0198

0.0105

321

Table D7a. P-values from Shapiro-Wilk W test to analyze overall mean liver somatic indices (LSIs)

calculated from each treatment population for evidence of non-normality (α = 0.05).

Treatment Population Mean LSI (%)

throughout experiment p-values

PFPA-dosed 1.39 ± 0.07 0.7822

PFPiA-dosed 1.21 ± 0.04 0.8867

Control 1.24 ± 0.04 0.1884

Table D7b. P-values from the parametric method of unpaired two-sample Student t-test to compare the

overall mean LSI calculated throughout the length of the experiment between the control and each of the

PFPA-dosed and PFPiA-dosed populations (α = 0.05).

Treatment Population

Type of Comparison

Control Population vs. Dosed Population

p-value (two-sided)

PFPA-dosed 0.0841

PFPiA-dosed 0.5691

322

Figure D5. Growth-corrected whole-body homogenate concentrations (ng/g wet wt) of C6, C8, and C10 PFPAs and C6/C6, C6/C8, and

C8/C8 PFPiAs in rainbow trout during exposure phase. Left panel corresponds to PFPA-dosed fish and right panel corresponds to PFPiA-

dosed fish. Each data point represents the arithmetic mean concentration of the triplicate (n = 3) sampling at each timepoint. The error

bar represents the standard error.

Time in exposure phase (day)

0 5 10 15 20 25 30 35Co

nc

en

tra

tio

n in

fis

h (

ng

/g w

et

wt)

0

2

4

6

8

1040

50

C6 PFPA

C8 PFPA

C10 PFPA

0 5 10 15 20 25 30 35

0

10

20

30

40

50

C6/C6 PFPiA

C6/C8 PFPiA

C8/C8 PFPiA

323

Table D8. P-values from parametric method, Pearson’s correlation test, to assess whether steady state

was achieved within the last 4 to 6 timepoints of the exposure phase for each analyte (α = 0.05).

Target Analyte

Number of concentration

data/time points prior to end

of exposure phase

95% CI for

population value of

slope

p-values

C6 PFPA 6 -0.090 to 0.119 0.8081

C8 PFPA 6 -0.064 to 0.058 0.8998

C10 PFPA 4 -0.370 to 0.299 0.6930

C6/C6 PFPiA 4 -0.125 to 1.245 0.0721

C6/C8 PFPiA 4 0.314 to 2.043 0.0278

C8/C8 PFPiA 4 0.114 to 1.720 0.0390

* All data for analytes were deemed normally distributed using the Shapiro-Wilk W test (α = 0.05, p >

0.05).

Table D9. P-values and r-values from the parametric method, Pearson’s correlation test, to evaluate the

correlation between the depuration half-life and logBAF observed for each target PFPA and PFPiA and

the number of perfluorinated carbons in their corresponding structures (α = 0.05).

Type of Comparison

95% CI for

population value

of slope

p-values r

t1/2 vs. # of perfluorinated carbons 0.33 to 6.83 0.0377 0.84

logBAF vs. # of perfluorinated carbons 0.11 to 0.20 0.0007 0.98

324

Figure D6. Concentrations (ng/g wet wt) of C6, C8, and C10 PFPAs ((A) PFPA-dosed fish) and C6/C6, C6/C8, and C8/C8 PFPiAs ((B)

PFPiA-dosed fish) in various fish tissues collected on the last day (day 31) of the exposure phase. Each data point represents the

arithmetic mean concentration of the triplicate (n = 3) sampling. The error bar represents the standard error. Analyte concentrations

observed below the LOD are indicated with an asterisk (*) (i.e. C6 PFPA in heart).


Co

ncen

trati

on

in

fis

h t

issu

es

(n

g/g

, w

et

wt)

0.1

1

10

100

1000

Carcass Liver Blood Kidneys Heart Gills

C6/

C6

PFP

iA

C6/

C8

PFP

iA

C8/

C8

PFP

iA0.1

1

10

100

1000A B

*

325

Figure D7. Ratios of liver-to-blood and liver-to-carcass concentrations for the C6, C8, and C10 PFPAs

and C6/C6, C6/C8, and C8/C8 PFPiAs based on tissue concentrations measured in rainbow trout

collected on last day of exposure phase. Each data point represents the arithmetic mean ratios reflective

of the triplicate (n = 3) sampling at that timepoint. The error bar represents the standard error.

C6 P

FP

A

C8 P

FP

A

C10 P

FP

A

C6/C

6 P

FP

iA

C6/C

8 P

FP

iA

C8/C

8 P

FP

iALiv

er:

Blo

od

an

d L

iver:

Carc

as

s R

ati

os

0.01

0.1

1

10

100

1000

Liver:Blood

Liver:Carcass

326

Table D10. Concentrations of C6, C8, and C10 PFPAs and C6/C6, C6/C8, and C8/C8 PFPiAs in different fish tissues (ng/g ww) analyzed

on the last day of the exposure phase (day 31) and their corresponding liver-to-blood (LBR), liver-to-carcass (LCR), and blood-to-carcass

(BCR) ratios calculated based on these concentrations.

Analyte

Concentration in fish tissues (ng/g ww) Liver-to-

blood

ratio

(LBR)

Liver-to-

carcass

ratio

(LCR)

Blood-to-

carcass

ratio

(BCR)

Liver Blood Kidneys Gills Heart Carcass

C6 PFPA* 35.2 ± 14.4 32.5 ± 26.6 8.3 ± 2.1 1.1 ± 0.5 nd 0.93 ± 0.52 1.08 37.70 34.78

C8 PFPA* 70.3 ± 31.9 115.7 ± 110.7 9.8 ± 3.6 0.96 ± 0.55 2.3 ± 0.6 0.98 ± 0.77 0.61 71.77 118.16

C10 PFPA* 571.6 ± 282.9 113.9 ± 102.6 51.7 ± 13.2 7.0 ± 1.9 9.1 ± 0.4 4.1 ± 2.9 5.02 138.27 27.56

C6/C6 PFPiA¥ 168.0 ± 25.4 36.3 ± 3.6 116.7 ± 40.7 34.2 ± 4.0 41.9 ± 5.7 21.0 ± 5.0 4.63 7.99 1.73

C6/C8 PFPiA¥ 248.5 ± 33.1 60.5 ± 11.3 212.4 ± 87.2 56.8 ± 6.9 57.3 ± 9.8 34.6 ± 9.2 4.11 7.18 1.75

C8/C8 PFPiA¥ 151.2 ± 14.5 46.7 ± 13.4 126.2 ± 47.9 39.3 ± 3.2 35.2 ± 7.2 20.6 ± 5.0 3.24 7.35 2.27

* Concentrations of these PFPAs were from PFPA-dosed fish

¥ Concentrations of these PFPiAs were from PFPiA-dosed fish

327

Figure D8. Concentrations of C6, C8, and C10 PFPAs (ng/g wet wt) observed in different tissue

extracts removed from PFPiA-dosed fish sampled on the last day of the exposure phase (day 31).

Each data point represents the arithmetic mean concentration of the triplicate (n = 3) sampling.

The error bar represents the standard error. Detection of C10 PFPA in the liver and blood, as

represented by the symbol (ɵ), may be due to endogenous contamination in the fish, as C10

PFPA was not detected in the whole-fish homogenates at any timepoint during the experiment

and none of the dosed PFPiA congeners contained a perfluorodecane (C10) linkage in their

structures to produce C10 PFPA upon C–P bond cleavage. Analyte concentrations observed

below the LOD are indicated with an asterisk (*) (i.e. C6 PFPA in heart; C10 PFPA in carcass,

kidneys, heart, and gills).


0.1

1

10

100

Co

nc

en

tra

tio

n in

fis

h t

iss

ue

s (

ng

/g, w

et

wt)

Carcass Liver Blood Kidneys Heart Gills

PFPAs in tissues removed from PFPiA-dosed fish

*

* ***

328

APPENDIX E

SUPPORTING INFORMATION FOR CHAPTER SEVEN

A Pilot Survey of Legacy and Current Commercial Fluorinated Chemicals in

Human Sera from United States Donors in 2009

329


Table E1a-c. Multiple reaction monitoring (MRM) transitions and mass

spectrometry parameters for all target analytes 339

Table E2. Multiple reaction monitoring (MRM) transitions and mass

spectrometry parameters for all internal standards 342

Figure E1. Chromatograms of a standard addition analysis of a human sera

sample for the suite of PFPiAs 343

Table E3ab. Limits of detection (LODs), limits of quantification (LOQs), and

matrix recoveries for the analytes of interest 344

Table E4a. Concentrations of all monitored PFCAs and PFSAs observed in

NIST SRM 1957 human sera from NIST Certificate of Analysis, an

interlaboratory study, and this study

346

Table E4b. Concentrations of all other target analytes observed in NIST SRM

1957 human sera from this study 347

Table E5ab. P-values from Shapiro-Wilk W test to analyze data for evidence of

non-normality 348

Table E6a-e. Summary of descriptive statistics for all detected analytes 350

Table E7. P-values from Mann-Whitney U test to compare concentrations

between single donor and pooled sera samples and for gender differences 355

Table E8. P-values from Mann-Whitney U test to compare concentrations of 6:2

and 8:2 FTS observed in pooled human sera collected in 2002 and 2009 356

Table E9. Spearman’s rank correlation coefficient r-values and p-values from

Spearman’s rank correlation test to analyze two groups of data for correlation 357

330

EXPERIMENTAL

Chemicals.

Perfluorobutanoic acid (PFBA, >99%), perfluoropentanoic acid (PFPeA, >99%),

perfluorohexanoic acid (PFHxA, >99%), perfluoroheptanoic acid (PFHpA, >99%),

perfluorooctanoic acid (PFOA, >99%), perfluorononanoic acid (PFNA, >99%),

perfluorodecanoic acid (PFDA, >99%), perfluoroundecanoic acid (PFUnA, >99%),

perfluorododecanoic acid (PFDoA, >99%), perfluorotridecanoic acid (PFTrA, >99%),

perfluorotetradecanoic acid (PFTeA, >99%), perfluorobutanesulfonate (PFBS, >99%),

perfluorohexanesulfonate (PFHxS, >99%), perfluorooctanesulfonate (PFOS, >99%),

perfluorodecanesulfonate (PFDS, >99%), perfluorooctanesulfonamidoacetate (FOSAA,

>99%), N-methylperfluorooctanesulfonamidoacetate (N-MeFOSAA, >99%), N-

ethylperfluorooctanesulfonamidoacetate (N-EtFOSAA, >99%), 4:2, 6:2, and 8:2

fluorotelomer sulfonates (4:2, 6:2, 8:2 FTS, <99%), C6 perfluorohexylphosphonate (C6

PFPA, >99%), C8 perfluorooctylphosphonate (C8 PFPA, >99%), C10

perfluorodecylphosphonate (C10 PFPA, >99%), C6/C6 bis(perfluorohexyl)phosphinate

(C6/C6 PFPiA, >98%), C6/C8 perfluorohexylperfluorooctylphosphinate (C6/C8 PFPiA,

>98%), and C8/C8 bis(perfluorooctyl)phosphinate (C8/C8 PFPiA, >98%) were obtained

from Wellington Laboratories Inc. (Guelph, ON). Mass-labeled internal standards were

donated from Wellington Laboratories and they included: 13

C4-PFBA (>99%), 13

C2-

PFHxA (>99%), 13

C4-PFOA (>99%), 13

C5-PFNA (>99%), 13

C2-PFDA (>99%), 13

C2-

PFUnA (>99%), 13

C2-PFDoA (>99%), 18

O2-PFHxS (>99%), and 13

C4-PFOS (>99%), d3-

N-MeFOSAA (>99%) and d3-N-EtFOSAA (>99%).

Due to a lack of authentic standards at the time of analysis, the Masurf®

FS-780

technical product was purchased from Mason Chemical Co. (Arlington Heights, IL) to be

used as a standard for the following chemicals: C6/C6, C6/C8, C8/C8, C6/C10, C8/C10,

and C6/C12 perfluorophosphinates (PFPiA, no purity information available). The

recently released authentic standards of C6/C6 PFPiA, C6/C8 PFPiA, and C8/C8 PFPiA

(Wellington Laboratories, Guelph, ON) were used to determine the percent composition

of these three PFPiAs in the Masurf® 780 technical product, as 36.9±0.1% C6/C6 PFPiA,

33±6% C6/C8 PFPiA, and 27±3% C8/C8 PFPiA. The concentrations of these three

PFPiAs reported in human sera here, as determined by using the Masurf® as the standard,

331

were corrected for based on this percent composition. The C6/C10, C8/C10, and C6/C12

PFPiAs were also detected in the Masurf®, but the lack of authentic standards precluded

the determination of their percent composition in the product. As such, the

concentrations of C6/C10, C8/C10, and C6/C12 PFPiAs, as determined by using the

Masurf® as the standard, were reported as is in the Supporting Information here and

should be treated as relative concentrations. All concentrations of the PFPiAs, whether

corrected or not, were used in the statistical tests, as described below.

Potassium chlorate (K2CO3, 99%) was purchased from Caledon Laboratory Ltd.

(Georgetown, ON). Dibromoneopentyl glycol (HOCH2C(CH2Br)2CH2OH, 98%), 2-

pentanone (CH3COCH2CH2CH3, >99%), phosphorus (V) oxychloride (POCl3, 99%), and

tetrabutylammonium hydrogen sulfate (TBAS, (CH3CH2CH2CH2)4N(HSO4), 99%) were

purchased from Sigma Aldrich (Oakville, ON; St. Louis, MO). Dichloromethane (CH2Cl2,

>99%) was purchased from Aldrich Chemical Co., Inc. (Milwaukee, WI). Toluene

(C6H5CH3, >99%), acetone (CH3COCH3, >99%), and m-xylene (C6H4(CH3)2) were

purchased from Fisher Scientific (Fairlawn, NJ). Methanol (Omnisolv, >99%), water

(Omnisolv, >99%), methyl-tert-butyl ether (MTBE, Omnisolv, >99%), and ammonia

(NH3, 30%) were purchased from EMD Chemicals, Inc. (Mississauga, ON).

The 4:2, 6:2, 8:2, and 10:2 polyfluoroalkyl phosphate diesters (diPAPs, y = x

only) were synthesized to be used as standards, as described elsewhere (1). Authentic

standards for the diPAPs became available after the analysis of all samples (Chiron AS,

Trondheim, Norway). The 6:2 (94%), 8:2 (98%), and 10:2 diPAPs (95%) were used to

determine the purities of the synthesized 6:2, 8:2, and 10:2 diPAPs as 94±5%, 98±7%,

and 39±5% respectively. The lack of an authentic standard for 4:2 diPAP at the time of

analysis precluded purity determination of the synthesized 4:2 diPAP. The

concentrations of the diPAPs reported in human sera here were not corrected for based on

these purities.

Synthesis of 6:2 fluorotelomer mercaptoalkyl phosphate diester (6:2 FTMAP).

The synthesis was performed as a bench-scale version of two patented processes

(2, 3). The reaction scheme of the two-step synthesis is shown below.

332

Step 1:

FF

F

FF

F

FF

F F

FF

F

SH +OH

OH

Br

Br

1. K2CO3,

2-pentanone, F

FF

F F

F F

F F

F F

F F

F

F

F F

F F

F F

F F

F F

S

S

OH

OHF

A mixture of 1H,1H,2H,2H-perfluorooctanethiol (CAS# 34451-26-8; 5.00 mmol,

2.00 eq.), dibromoneopentyl glycol (CAS# 3296-90-0; 2.50 mmol, 1.00 eq.), K2CO3

(CAS# 3811-04-9; 8.03 mmol, 3.21 eq.), and 2.50 mL of 2-pentanone (CAS# 107-87-9;

solvent) was reacted under a nitrogen atmosphere at 105oC for 16 hours. After cooling

the mixture to 70oC, 4.00 mL of H2O was added and the entire mixture was transferred to

a separatory funnel to separate the aqueous and organic phases. Evaporation of the

organic phase and two rounds of recrystallization with toluene produced the white solid

product of bis-(1H,1H,2H,2H-perfluorooctanethiolmethyl)-1,3-propanediol (1.70 mmol,

1.46 g, 68% pure). Product identification was confirmed by 1H,

19F, and

13C NMR

analysis: 1H NMR (CD3OD, 400 MHz): δ = 2.45-2.61 (m, 8H, CH2), 2.80-2.87 (m, 8H,

CH2); 13

C NMR (CD3OD, 101 MHz): δ = 25.0 (C), 30.7 (CH2), 35.4 (CH2), 46.4 (CH2),

63.8 (CH2); 19

F NMR (CD3OD, 377 MHz): δ = 81.5 (t, 3F, CF3), -114.4 (t, 2F, CF2), -

122.0 (mc, 2F, CF2), -123.0 (mc, 2F, CF2), -123.5 (mc, 2F, CF2), -126.5 (mc, 2F, CF2).

Step 2:

F

FF

F F

F F

F F

F F

F F

F

F

F F

F F

F F

F F

F F

S

S

OH

OHF

F

FF

F F

F F

F F

F F

F F

F

F

F F

F F

F F

F F

F F

S

S

O

OF

P

O

OH

2. POCl3, CH2Cl2,

Acetone/H2O,

The bis-(1H,1H,2H,2H-perfluorooctanethiolmethyl)-1,3-propanediol (0.20 mmol,

1.0 eq.) was dissolved in 5.0 mL of anhydrous CH2Cl2 under a nitrogen atmosphere.

Excess POCl3 dissolved in 0.50 mL of dry CH2Cl2 was added dropwise to the above

mixture. After refluxing for 21 hours, the reaction mixture was evaporated under

333

vacuum, and the residue was redissolved in 5.0 mL of 90:10 mixture of acetone:H2O and

refluxed for another 24 hours. Any residual acetone was removed by a rotary evaporator.

After recrystallization with m-xylene, a white solid product of 6:2 FTMAP was obtained

(95% pure). Product identification was confirmed by 1H,

19F, and

31P NMR analysis:

1H

NMR (CD3OD, 400 MHz): δ = 2.45-2.63 (m, 4H, CH2), 2.86-2.93 (m, 8H, CH2), 4.34 (d,

2J = 12.3 Hz, 4H, CH2);

19F NMR (CD3OD, 377 MHz): δ = -81.5 (t, 3F, CF3), -114.4 (t,

2F, CF2), -122.0 (mc, 2F, CF2), -123.0 (mc, 2F, CF2), -123.5 (mc, 2F, CF2), -126.5 (mc,

2F, CF2); 31

P NMR (CD3OD, 162 MHz): δ = -5.57.

Extraction procedures of sera samples.

Briefly, 1 mL of 0.5M TBAS solution, either adjusted to pH 10 with 30% aqueous

NH3 or without pH adjustment (pH ~3), was added to 2-3 mL of sera, followed by

extraction with two 4 mL aliquots of MTBE. The MTBE aliquots were combined,

evaporated to dryness under nitrogen, and reconstituted in 0.14–0.15 mL of methanol.

For the analysis of the PFPiAs, the sera samples were extracted using the TBAS solution

adjusted to pH 10. For the analysis of all other analytes, the sera samples were extracted

using the TBAS solution without pH adjustment. Each of the fifty human sera sample

was extracted in duplicate with one procedural blank (HPLC grade water) extracted in

company to each sample (n = 50).



Chromatographic separation was performed using a Kinetex C18 column (50 x

4.6 mm, 3 μm; Phenomenex®, Torrance, CA). Analyte quantitation was performed using

an API4000 triple-quadrupole mass spectrometer (Applied Biosystems/MDS Sciex) in

the negative electrospray ionization mode, coupled to an Agilent 1100 LC system. Four

high performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS)

methods were used for the analysis of the target analytes.

For the analysis of the diPAPs, SAmPAP, 6:2 FTMAP, 4:2 FTS, 6:2 FTS, and 8:2

FTS, the samples were injected as 35 µL injections and analyzed by the following

334

gradient method at 500 µL/min using HPLC grade methanol and water, each prepared

into 10 mM ammonium acetate mobile phases: the initial solvent composition at t = 0

min. was 60:40 water:methanol, which changed to 5:95 over a period of 2.5 min. at t =

2.50 min. and held for 3.5 min. to t = 6.00 min., before returning to the initial

composition of 60:40 water:methanol at t = 6.50 min. The column was allowed to


For the analysis of the PFPAs and PFPiAs, the samples were injected as 35 µL

injections and analyzed by the following gradient method at 500 µL/min: the initial

solvent composition at t = 0 min. was 70:30 water: methanol, which changed to 5:95 over

a period of 5 min. at t = 5.00 min. and held for 2 min. to t = 7.00 min., before returning to

the initial composition of 70:30 water:methanol at t = 7.50 min. The column was allowed

to reequilibrate for 2.50 min. for a total run time of 10 min.

For the analysis of PFBA, PFPeA, PFHxA, PFHpA, and PFBS, the samples were

injected as 25 µL injections and analyzed by the following gradient method at 500

µL/min: the initial solvent composition at t = 0 min. was 80:20 water:methanol, which

changed to 5:95 over a period of 3 min. at t = 3.00 min. and held for 2 min. to t = 5.00

min., before returning to the initial composition of 80:20 water:methanol at t = 5.50 min.

The column was allowed to reequilibrate for 2.50 min. for a total run time of 8 min.

For the analysis of PFOA, PFNA, PFDA, PFUnA, PFDoA, PFTrA, PFTeA,

PFHxS, PFOS, PFDS, FOSAA, N-MeFOSAA, and N-EtFOSAA, the samples were

injected as 25 µL injections and analyzed by the following gradient method at 500

µL/min: the initial solvent composition at t = 0 min. was 35:65 water:methanol, which

changed to 5:95 over a period of 3 min. at t = 3.00 min. and held for 2 min. to t = 5.00

min., before returning to the initial composition of 35:65 water:methanol at t = 5.50 min.

The column was allowed to reequilibrate for 2.50 min. for a total run time of 8 min.


A list of the analyte-specific multiple reaction monitoring (MRM) transitions and

mass spectrometry parameters for all target analytes and their corresponding internal

standards is provided in Table E1a-c and E2. For the analysis of diPAPs (y = x only),

SAmPAP, and FTSs, two MRM transitions were monitored for quantitation and identity

335

confirmation for each analyte. Three MRM transitions were monitored for 6:2 FTMAP.

The most sensitive transition of 6:2 FTMAP (921.0>79.0; [PO3-]) was frequently

encumbered with interference peaks, especially at low concentrations; therefore, two

additional transitions (921.0>318.7; [CF3(CF2)4CF2-] and 921.0>575.0; loss of one 6:2

fluorotelomer tail) were simultaneously monitored.. The peak ratios between the

different MRM transitions were consistent within <15% relative standard deviation

(RSD) for all the analytes, except for 10:2 diPAP (25% RSD). The PFPAs fragment

exclusively to PO3- (79 m/z) (4), while the PFPiAs fragment to [F(CF2)xPO2F]

- (5). Each

of these transitions was monitored for quantitation of the PFPAs and PFPiAs and

chemical identification was internally confirmed by standard addition.

Comparison of using single donor versus pooled samples in human sera analysis.

Pooled sera samples have been used to obtain representative population-based

estimates of concentrations of polyfluorinated and perfluorinated chemicals in humans

(7-10). The advantages of pooled samples are reduced analytical costs and lower

biosafety costs, since the samples are typically pre-screened for hepatitis and HIV by the

commercial supplier. However, human sera analysis using pooled samples does not

provide information on the contamination present in individual donors. In this study, a

higher number of detects was typically observed in the pooled samples than in the single

donor samples, especially for the analytes present in the sub-ppb (µg/L) concentration

ranges, such as the diPAPs, FOSAA, N-EtFOSAA, FTSs, PFPiAs, the short chain PFCAs

(C4–C6), and PFBS. For the majority of the analytes, no significant differences were

observed in the concentrations between the pooled and single donor samples (Mann

Whitney U test, p>0.05, Table S7), except for 6:2 diPAP, N-EtFOSAA, 6:2 FTS, C6/C6

PFPiA, C6/C8 PFPiA, and PFUnA. The choice between using pooled and single donor

samples may be dependent on analyte, as well as, the type of data desired (i.e.

population-based estimate of the contamination vs. individual contamination).

336


Methanol rinses of blood collection items.

All blood collection items, including storage tubes, bottles, collection bags,

needles, and tubings were provided by Tennessee Blood Services Corp. (Memphis, TN).

The storage tubes (10 mL) and bottles (250 mL) were rinsed with 3 mL and 50 mL

aliquots of HPLC grade methanol respectively. The blood collection bags and the

tubings and the needle attached to these bags were cut into small pieces with methanol-

rinsed scissors and transferred to 50 mL polypropylene tubes (BD Biosciences, Franklin

Lakes, NJ), followed by addition of 40 mL of HPLC grade methanol. All rinses were

performed in triplicate (n = 3). From the methanol rinses of each item, a 1 mL aliquot

was filtered through 0.25 μm nylon syringe filters (Chromatographic Specialties,

Brockville, ON) into 1.2 mL low-temperature cryo-vials (VWR International Ltd.,

Mississauga, ON) and analyzed directly by HPLC-MS/MS without further concentration.

Statistical Analysis.

For all statistical tests, any concentrations below the LOD were imputed as the

LOD divided by the square root of two. All data were tested for evidence of non-

normality using the Shapiro-Wilk W test (p-values in Table E5ab). Data from the single

donor samples were largely non-normally distributed (~90% of the analytes), while data

from the pooled samples showed more frequent cases of normal distribution (~60% of the

analytes). Non-normally distributed data were logarithmically transformed and re-tested

with the Shapiro-Wilk W test, but normality only improved for ~10% of the transformed

data. The assumption of normality in the data was minimized by using nonparametric

methods, such as the Mann-Whitney U test to compare analyte concentrations (i.e.

temporal, gender, analyte vs. analyte) and the Spearman rank correlation test to test for

possible correlations among the target analytes. A p-value of 0.05 was chosen as the

criterion for statistical significance in all analyses. All statistical tests were performed

using StatsDirect (Version 2.7.8, Cheshire, UK). A summary of the descriptive statistics

calculated for all detected analytes is provided in Table E6a-e. A significant

concentration difference was observed between the single donor and pooled samples for

337

6:2 diPAP, N-EtFOSAA, 6:2 FTS, C6/C6 PFPiA, C6/C8 PFPiA, and PFUnA (Mann-

Whitney U test, p<0.05, Table E7), and so their concentrations were considered

separately. No significant difference was observed for the remaining analytes (Mann-

Whitney U test, p>0.05, Table E7), and so the concentrations in both sample types were

combined for Spearman’s rank correlation analyses.

Literature Cited

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(PFCAs) from the biotransformation of polyfluoroalkyl phosphate surfictants

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2. Falk, R. A. C., K.P.; Karydas, A.; Jacobson, M. (Co., C.-G.).Heteroatom

containing perfluoroalkyl terminated neopentyl glycols and compositions

therefrom. U.S Patent 5,045,624; Ardsley, NY, 1991.

3. Falk, R. A., Clark, K.P. (AG, C.-G.).5,5-Bis(perfluoroalkylheteromethyl)-2-

hydroxy-2-oxo-1,3,2-dioxaphosphiranes, derived acyclic phosphorus acids and

salts or esters thereof. European Patent 0,453,406,A1; New City, NY; Bethel, CT,

1991.

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A., Perfluorinated Phosphonic Acids in Canadian Surface Waters and Wastewater

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5. D'eon J, C.; Mabury, S. A., Uptake and elimination of perfluorinated phosphonic

acids in the rat. Environ Toxicol. Chem. 2010, 29, (6), 1319-1329.

6. Keller, J.M.; Calafat, A.M.; Kato, K.; Ellefson, M.E.; Reagen, W.K.; Strynar, M.;

O'Connell, S.; Butt, C.M.; Mabury, S.A.; Small, J.; Muir, D.C.G.; Leigh, S.D.;

Schantz, M.M. Determination of perfluorinated alkyl acid concentrations in

human serum and milk standard reference materials. Anal. Bioanal. Chem. 2010,

397, (2), 439-451.

7. Hansen, K.J.; Clemen, L.A.; Ellefson, M.E.; Johnson, J.O. Compound-specific,

quantitative characterization of organic fluorochemicals in biological matrices.

Environ. Sci. Technol. 2001, 35, 766-770.

8. Calafat, A.M.; Kuklenyik, Z.; Caudill, S.P.; Reidy, J.A.; Needham, L.L.

Perfluorochemicals in pooled serum samples from United States residents in 2001

and 2002. Environ. Sci. Technol. 2006, 40, 2128-2134.

338

9. Haug, L.S.; Thomsen, C.; Becher, G. Time trends and the influence of age and

gender on serum concentrations of perfluorinated compounds in archived human

samples. Environ. Sci. Technol. 2009, 43, 2131-2136.

10. D’eon, J.C.; Crozier, P.W.; Furdui, V.I.; Reiner, E.J.; Libelo, E.L.; Mabury, S.A.

Observation of a commercial fluorinated material, the polyfluoroalkyl phosphoric

acid diesters, in human sera, wastewater treatment plant sludge, and paper fibers.

Environ. Sci. Technol. 2009, 43, 4589-4594.

11. Connolly, P.; Decker, E.; Zhu, X.; Keller, R. Analysis of pooled human sera and

plasma and monkey sera for fluorocarbons using Exygen method ExM-023-071.

Prepared for 3M Environmental Laboratory. AR226-1152.

339

Table E1a. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.


Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy,

CE

(V)

Collision

Cell Exit

Potential,

CXP (V)

Polyfluoroalkyl phosphate diester

4:2 polyfluoroalkyl phosphate diester 4:2 diPAP 589.1>96.9 30 -50 -50 -15

589.1>343.0 20 -50 -25 -15

4:2/6:2 polyfluoroalkyl phosphate diester 4:2/6:2 diPAP 689.0>96.9 30 -60 -60 -15


789.0>443.0 20 -65 -27 -15



989.0>543.0 20 -70 -33 -15



1189.0>643.0 40 -80 -40 -15


Fluorotelomer mercaptoalkyl phosphate diester

6:2 fluorotelomer mercaptoalkyl phosphate

diester 6:2 FTMAP

921.0>79.0 40 -95 -99 -15

921.0>318.7 40 -95 -70 -15

921.0>575.0 40 -95 -50 -15

N-ethyl perfluorooctanesulfonamidoethanol-based phosphate diester

N-ethyl perfluorooctanesulfonamidoethanol-

based phosphate diester SAmPAP

1203.0>526.0 30 -190 -68 -15

1203.0>650.0 30 -190 -57 -15

340

Table E1b. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.


Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy,

CE

(V)

Collision

Cell Exit

Potential,

CXP (V)

Fluorotelomer sulfonate

4:2 fluorotelomer sulfonate 4:2 FTS 327.0>81.0 20 -95 -53 -15

327.0>306.8 20 -95 -30 -18


427.0>406.8 20 -100 -32 -10


527.0>506.8 20 -100 -40 -15

Perfluorooctanesulfonamidoacetate, N-methyl & N-ethyl perfluorooctanesulfonamidoacetate

Perfluorooctanesulfonamidoacetate FOSAA 559.9>419.0 20 -40 -45 -15

N-methyl perfluorooctanesulfonamidoacetate N-MeFOSAA 570.0>419.0 20 -40 -36 -15

N-ethyl perfluorooctanesulfonamidoacetate N-EtFOSAA 584.0>419.0 20 -50 -36 -15

Perfluorophosphonate and perfluorophosphinate










341

Table E1c. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.

Compound Acronym Mass

Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy,

CE

(V)

Collision

Cell Exit

Potential,

CXP (V)


Perfluorobutanoate PFBA (C4) 212.8>168.9 40 -25 -13 -15

Perfluoropentanoate PFPeA (C5) 262.8>218.97 40 -20 -13 -15

Perfluorohexanoate PFHxA (C6) 312.8>268.9 20 -20 -13 -15

Perfluoroheptanoate PFHpA (C7) 362.8>319.0 20 -27 -13 -15

Perfluorooctanoate PFOA (C8) 413.0>368.9 20 -35 -15 -15

Perfluorononanoate PFNA (C9) 462.9>419.0 20 -35 -15 -15

Perfluorodecanoate PFDA (C10) 513.0>470.0 20 -45 -15 -15

Perfluoroundecanoate PFUnA (C11) 562.8>519.0 20 -45 -15 -15

Perfluorododecanoate PFDoA (C12) 612.8>569.0 20 -45 -15 -15

Perfluorotridecanoate PFTrA (C13) 662.8>619.0 20 -45 -15 -15

Perfluorotetradecanoate PFTeA (C14) 712.8>669.0 20 -45 -15 -15

Perfluorosulfonate

Perfluorobutanesulfonate PFBS (C4) 299.0>99.0 20 -55 -65 -15

Perfluorohexanesulfonate PFHxS (C6) 399.0>99.0 20 -55 -65 -15

Perfluorooctanesulfonate PFOS (C8) 499.0>99.0 20 -120 -80 -15

Perfluorodecanesulfonate PFDS (C10) 599.0>99.0 20 -120 -80 -15

342

Table E2. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all internal standards.

Target Analyte Internal

Standard

Mass

Transition

Dwell

(ms)

Declustering

Potential, DP

(V)

Collision

Energy, CE

(V)

Collision Cell

Exit Potential,

CXP (V)

Perfluorooctanesulfonamidoacetate, N-methyl & N-ethyl perfluorooctanesulfonamidoacetate

FOSAA d3-N-MeFOSAA 573.0>419.0 20 -40 -36 -15

N-MeFOSAA d3-N-MeFOSAA 573.0>419.0 20 -40 -36 -15

N-EtFOSAA d5-N-EtFOSAA 589.0>419.0 20 -50 -36 -15

Perfluorinated acids

PFBA (C4) 13

C4-PFBA 217.0>172.0 40 -25 -13 -15

PFPeA (C5) 13

C2-PFHxA 314.8>269.8 20 -20 -13 -15

PFHxA (C6) 13

C2-PFHxA 314.8>269.8 20 -20 -13 -15

PFHpA (C7) 13

C4-PFOA 417.0>372.0 20 -35 -15 -15

PFOA (C8) 13

C4-PFOA 417.0>372.0 20 -35 -15 -15

PFNA (C9) 13

C5-PFNA 468.0>423.0 20 -35 -15 -15

PFDA (C10) 13

C2-PFDA 515.0>470.0 20 -45 -15 -15

PFUnA (C11) 13

C2-PFUnA 564.8>520.0 20 -45 -15 -15

PFDoA (C12) 13

C2-PFDoA 614.8>570.0 20 -45 -15 -15

PFTrA (C13) 13

C2-PFDoA 614.8>570.0 20 -45 -15 -15

PFTeA (C14) 13

C2-PFDoA 614.8>570.0 20 -45 -15 -15

PFBS (C4) 18

O2-PFHxS 403.0>103.0 20 -55 -65 -15

PFHxS (C6) 18

O2-PFHxS 403.0>103.0 20 -55 -65 -15

PFOS (C8) 13

C4-PFOS 503.0>99.0 20 -120 -80 -15

PFDS (C10) 13

C4-PFOS 503.0>99.0 20 -120 -80 -15

343

Time (min)

Sample

Sample + 1x

Sample + 2x

Sample + 5x

Sample + 10x

701>401 801>501 901>501 901>601 1001>601 1001>701

Figure E1. Chromatograms of a standard addition analysis of a human sera sample for the suite of PFPiAs.

344

Table E3a. Limits of detection (LODs), limits of quantification (LOQs), and matrix

recoveries for the analytes of interest.

Analyte

Instrumental

(on column)

Method

(20X) Recovery (%)


(pg) (μg/L)

Fluorinated Precursors

4:2 diPAP 1.75 3.50 0.008 0.015 107 ± 20

6:2 diPAP 1.75 3.50 0.008 0.015 109 ± 22

8:2 diPAP 17.50 26.25 0.075 0.113 87 ± 21

10:2 diPAP 8.75 17.50 0.038 0.075 100 ± 27

6:2 FTMAP 1.75 3.50 0.015 0.038 97 ± 17

SAmPAP 1.75 3.50 0.008 0.02 101 ± 8

Fluorinated Intermediates

FOSAA 0.88 1.75 0.011 0.023 90 ± 5

N-MeFOSAA 0.18 0.35 0.002 0.005 94 ± 6

N-EtFOSAA 0.35 0.88 0.005 0.011 94 ± 2

4:2 FTS 0.35 0.88 0.005 0.011 90 ± 20

6:2 FTS 0.35 0.88 0.005 0.011 100 ± 21

8:2 FTS 0.35 0.88 0.005 0.011 94 ± 15

345

Table E3b. Limits of detection (LODs), limits of quantification (LOQs), and matrix

recoveries for the analytes of interest.

Analyte

Instrumental

(on column)

Method

(20X) Recovery (%)


(pg) (μg/L)

Perfluorinated Acids

C6 PFPA 3.50 8.75 0.009 0.023 86 ± 12

C8 PFPA 1.75 3.50 0.005 0.009 90 ± 11

C10 PFPA 26.25 35.00 0.070 0.093 89 ± 10

C6/C6 PFPiA 0.32 0.65 0.001 0.002 101 ± 32

C6/C8 PFPiA 0.29 0.58 0.001 0.002 105 ± 32

C8/C8 PFPiA 0.47 0.95 0.001 0.003 100 ± 38

C6/C10 PFPiA* 0.88 1.75 0.002 0.005 95 ± 26

C8/C10 PFPiA* 1.75 3.50 0.005 0.009 93 ± 27

C6/C12 PFPiA* 1.75 3.50 0.005 0.009 98 ± 34

PFBA (C4) 0.35 0.88 0.005 0.011 114 ± 17

PFPeA (C5) 0.18 0.35 0.002 0.005 96 ± 9

PFHxA (C6) 0.04 0.18 0.001 0.002 125 ± 11

PFHpA (C7) 0.04 0.18 0.001 0.002 71 ± 3

PFOA (C8) 0.18 0.35 0.002 0.005 91 ± 8

PFNA (C9) 0.18 0.35 0.002 0.005 93 ± 14

PFDA (C10) 0.18 0.35 0.002 0.005 114 ± 15

PFUnA (C11) 0.26 0.35 0.003 0.005 96 ± 13

PFDoA (C12) 0.35 0.88 0.005 0.011 111 ± 23

PFTrA (C13) 0.35 0.88 0.005 0.011 92 ± 18

PFTeA (C14) 0.35 0.88 0.005 0.011 85 ± 24

PFBS (C4) 0.35 0.88 0.005 0.011 80 ± 8

PFHxS (C6) 0.35 0.88 0.005 0.011 106 ± 20

PFOS (C8) 0.18 0.35 0.002 0.005 97 ± 14

PFDS (C10) 0.35 0.88 0.005 0.011 94 ± 15

* Concentrations were not corrected based on corresponding percent

distribution in Masurf® 780 standard

346

Table E4a. Concentrations of all monitored PFCAs and PFSAs observed in NIST SRM

1957 human sera from NIST Certificate of Analysis, an interlaboratory study, and this

study.

Analyte

Reported Concentrations (µg/L) Measured

Concentrations3

(µg/L) NIST Certificate

of Analysis1

Interlaboratory

Study2

PFBA * <LOD or <LOQ nd

PFPeA * <LOD or <LOQ 0.23±0.09

PFHxA * <LOD or <LOQ 0.08±0.02

PFHpA 0.305±0.036 0.28–0.33 0.27±0.10

PFOA 5.00±0.40 4.08–5.86 5.06±0.86

PFNA 0.880±0.068 0.76–0.97 0.88±0.10

PFDA 0.39±0.10 0.29–0.53 0.33±0.06

PFUnA 0.174±0.031 0.11–0.22 0.15±0.02

PFDoA * 0.16–0.20 0.02±0.01

PFTrA * <LOD or <LOQ 0.03±0.00

PFTeA * <LOD or <LOQ nd

PFBS * <LOD or <LOQ nd

PFHxS 4.00±0.75 3.01–6.49 3.49±0.94

PFOS 21.1±1.2 19.5–38.0 13.66±1.13

PFDS * 0.15–0.49 0.22±0.05 1 Data obtained from certificate of analysis available on the NIST website:

www.nist.gov/srm. 2 Data obtained from ref. (6).

3 Data obtained from replicate analysis (n = 4) of SRM1957 in the present study.

* Concentrations of PFBA, PFPeA, PFHxA, PFDoA, PFTrA, PFTeA, PFBS, and

PFDS are not reported on the NIST certificate of analysis.

nd = nondetects (i.e. analytes were either not detected or concentrations were

below their corresponding LODs)

347

Table E4b. Concentrations of all other target analytes monitored in NIST SRM 1957

human sera from this study (n = 4).

Analyte Measured Concentrations (µg/L)

4:2 diPAP 0.05±0.01

4:2/6:2 diPAP 0.15±0.04

6:2 diPAP 0.31±0.09

6:2/8:2 diPAP 0.13±0.05

8:2 diPAP 0.14±0.05

8:2/10:2 diPAP nd

10:2 diPAP nd

6:2 FTMAP nd

N-EtFOSE phosphate nd

FOSAA 0.16±0.02

N-MeFOSAA 0.74±0.06

N-EtFOSAA 0.15±0.01

4:2 FTS 0.03±0.01

6:2 FTS 0.02±0.01

8:2 FTS 0.09±0.03

C6 PFPA nd

C8 PFPA nd

C10 PFPA nd

C6/C6 PFPiA 0.003±0.001

C6/C8 PFPiA 0.006±0.001

C8/C8 PFPiA nd

C6/C10 PFPiA* 0.011±0.001

C8/C10 PFPiA* nd

C6/C12 PFPiA* nd

nd = nondetects (i.e. analytes were either not detected or concentrations were

below their corresponding LODs)

* Concentrations were not corrected based on corresponding percent

distribution in Masurf® 780 standard

348

Table E5a. P-values from Shapiro-Wilk W test to analyze data for evidence of non-

normality. A p-value of 0.05 is the chosen criterion of statistical significance such that if

the test statistic is below 0.05 (p<0.05), the null hypothesis may be rejected, and the data

are unlikely to be normally distributed. If the test statistic is above 0.05 (p>0.05), the

Shapiro-Wilk W test can only conclude there is no evidence of non-normality.

Analyte Type of

Sample

Type of Data

Data without log

transformation Log-transformed data

4:2 diPAP Single donor <0.0001 <0.0001

Pooled * *

4:2/6:2 diPAP Single donor <0.0001 <0.0001

Pooled <0.0001 <0.0001

6:2 diPAP Single donor <0.0001 0.0547

a

Pooled 0.0122 0.2566b

6:2/8:2 diPAP Single donor <0.0001 0.0001

Pooled 0.0027 0.0494

8:2 diPAP Single donor <0.0001 <0.0001

Pooled 0.0486 0.0476

FOSAA Single donor <0.0001 0.001

Pooled 0.0242 0.6837b

N-MeFOSAA Single donor <0.0001 <0.0001

Pooled 0.0009 0.5203b

N-EtFOSAA Single donor 0.0001 <0.0001

Pooled 0.1183b

0.7616b

4:2 FTS Single donor <0.0001 <0.0001

Pooled <0.0001 <0.0001

6:2 FTS Single donor <0.0001 <0.0001

Pooled 0.4333b

0.061a

8:2 FTS Single donor <0.0001 0.2636

b

Pooled 0.0324 0.2043b

C6/C6 PFPiA Single donor <0.0001 <0.0001

Pooled <0.0001 0.0109

C6/C8 PFPiA Single donor <0.0001 0.0065

Pooled <0.0001 0.0023


Pooled <0.0001 <0.0001


Pooled <0.0001 0.1284b


Pooled <0.0001 <0.0001


Pooled <0.0001 <0.0001

* Test cannot be performed due to 100% non-detection in the samples. a Test was not quite significant; cannot assume there is no evidence of non-normality.

b No evidence of non-normality.

349

Table E5b. P-values from Shapiro-Wilk W test to analyze data for evidence of non-

normality. A p-value of 0.05 is the chosen criterion of statistical significance such that if

the test statistic is below 0.05 (p<0.05), the null hypothesis may be rejected, and the data

are unlikely to be normally distributed. If the test statistic is above 0.05 (p>0.05), the

Shapiro-Wilk W test can only conclude there is no evidence of non-normality.

Analyte Type of

Sample

Type of Data

Data without log

transformation Log-transformed data

PFBA (C4) Single donor <0.0001 <0.0001

Pooled 0.7788b

0.8161b

PFPeA (C5) Single donor <0.0001 0.0002

Pooled * *

PFHxA (C6) Single donor <0.0001 <0.0001

Pooled 0.0048 0.0301

PFHpA (C7) Single donor <0.0001 <0.0001

Pooled 0.5644b

0.5239b

PFOA (C8) Single donor 0.1385

b 0.0338

Pooled 0.2385b

>0.9999b

PFNA (C9) Single donor 0.0784

a 0.0997

a

Pooled 0.0827a 0.6286

PFDA (C10) Single donor 0.0001 <0.0001

Pooled 0.2721b

0.6403b

PFUnA (C11) Single donor <0.0001 0.0022

Pooled 0.1059b

0.9254b

PFBS (C4) Single donor <0.0001 <0.0001

Pooled <0.0001 <0.0001

PFHxS (C6) Single donor <0.0001 0.1724

b

Pooled 0.7754b

0.3600b

PFOS (C8) Single donor <0.0001 0.0139

Pooled 0.9626b

0.9799b

PFDS (C10) Single donor 0.0001 <0.0001

Pooled 0.0031 0.0349

* Test cannot be performed due to 100% non-detection in the samples. a Test was not quite significant; cannot assume there is no evidence of non-normality.

b No evidence of non-normality.

350

Table E6a. Summary of descriptive statistics for all detected analytes. For the purposes

of calculating means, values below the LOD were assigned a value of zero and values

below the LOQ were used unaltered. For analytes that were detected in <20% of the

samples, mean concentrations were not calculated and only the range is reported.

Concentrations are reported in ng/L (ppt).

(ng/L) Analyte

4:2 diPAP 4:2/6:2 diPAP 6:2 diPAP 6:2/8:2 diPAP 8:2 diPAP

All single donor samples (n = 40)

Mean * * 72.07 34.65 110.31

SE * * 15.13 8.68 48.05

Range <LOD–21.51 <LOD–100.54 <LOD–388.55 <LOD–303.05 <LOD–1801.74

% <LOD 88 85 18 48 68

% <LOQ 98 90 50 93 78

Male single donor samples (n = 20)

Mean * * 87.14 42.85 91.96

SE * * 25.53 16.60 39.45


% <LOD 85 80 25 55 70

% <LOQ 95 90 30 85 75

Female single donor samples (n = 20)

Mean * * 57.00 26.46 128.65

SE * * 16.24 5.19 88.82


% <LOD 90 90 10 40 65

% <LOQ 100 90 20 100 80

All pooled samples (n = 10)

Mean * * 131.81 49.06 133.59

SE * * 37.85 19.20 38.80

Range - <LOD–163.94 30.97–346.46 <LOD–157.03 <LOD–323.36

% <LOD 100 90 0 40 40

% <LOQ 100 90 0 70 50

* Mean concentrations and standard error were not reported due to the low frequency of

detection in the samples (<20%).

- Range was not reported due to 100% non-detection in the samples

351

Table E6b. Summary of descriptive statistics for all detected analytes. For the purposes





(ng/L) Analyte

FOSAA N-MeFOSAA N-EtFOSAA 4:2 FTS 6:2 FTS 8:2FTS


Mean 64.30 356.99 50.28 * 7.64 37.75

SE 15.22 71.24 6.92 * 1.26 6.16

Range <LOD–432.35 <LOD–

1997.72

<LOD–

173.54 <LOD–17.94 <LOD–29.54 <LOD–162.49

% <LOD 35 10 18 90 46 5

% <LOQ 43 10 18 95 69 13


Mean 44.24 241.29 41.25 * 5.91 45.18

SE 16.25 72.15 7.93 * 1.70 9.56

Range <LOD–305.55 <LOD–

1509.43

<LOD–

134.61 <LOD–15.83 <LOD–18.39 <LOD–154.12

% <LOD 50 15 25 95 58 11

% <LOQ 50 15 25 100 68 11


Mean 84.36 472.69 59.31 * 9.28 30.68

SE 25.38 119.24 11.20 * 1.82 7.78

Range <LOD – 432.35 <LOD–

1997.72

<LOD–

173.54 <LOD–17.94 <LOD–29.54 7.32 – 162.49

% <LOD 20 5 10 85 35 0

% <LOQ 35 5 10 90 70 15

All pooled donor samples (n = 10)

Mean 64.20 443.66 69.19 * 23.74 73.68

SE 13.56 108.22 6.78 * 5.37 21.03

Range 25.93–166.58 146.76–

1355.58 43.27–119.86 - <LOD–47.25 9.12 – 230.70

% <LOD 0 0 0 100 20 0

% <LOQ 0 0 0 100 30 20




352

Table E6c. Summary of descriptive statistics for all detected analytes. For the purposes





(ng/L)

Analyte

C6/C6

PFPiA

C6/C8

PFPiA

C8/C8

PFPiA

C6/C10

PFPiAa

C8/C10

PFPiAa

C6/C12

PFPiAa


Mean 3.65 7.67 * 19.88 * 12.19

SE 1.32 1.91 * 4.77 * 6.01

Range <LOD–

50.24

<LOD–

60.96

<LOD–

22.19

<LOD–

133.95

<LOD–

48.73

<LOD–

225.12

% <LOD 50 28 95 58 95 80

% <LOQ 58 30 98 58 98 80


Mean 5.71 9.74 * 26.59 * 20.87

SE 2.51 3.19 * 7.73 * 11.56

Range <LOD–

50.24

<LOD–

60.96

<LOD–

22.19

<LOD–

133.95

<LOD–

48.73

<LOD–

225.12

% <LOD 40 15 95 45 95 70

% <LOQ 45 15 95 45 95 70


Mean 1.60 5.60 * 13.18 * *

SE 0.65 2.07 * 5.38 * *

Range <LOD–

12.02

<LOD–

36.67 -

<LOD–

86.56 <LOD–5.68

<LOD–

47.86

% <LOD 60 40 100 70 95 90

% <LOQ 70 45 100 70 100 90


Mean 23.20 37.86 * 140.35 * *

SE 19.81 27.35 * 115.98 * *

Range <LOD–

201.41 4.36–283.38

<LOD–

50.73

<LOD–

1182.50

<LOD–

891.02

<LOD–

957.44

% <LOD 10 0 90 30 90 90

% <LOQ 20 0 90 30 90 90



- Range was not reported due to 100% non-detection in the samples a Concentrations were not corrected based on corresponding percent distribution in

Masurf® 780 standard

353

Table E6d. Summary of descriptive statistics for all detected analytes. For the purposes of calculating means, values below the LOD

were assigned a value of zero and values below the LOQ were used unaltered. For analytes that were detected in <20% of the samples,

mean concentrations were not calculated and only the range is reported. Concentrations are reported in ng/L (ppt).

(ng/L) Analyte

PFBA PFPeA PFHxA PFHpA PFOA PFNA PFDA PFUnA


Mean 35.06 72.59 49.62 97.16 2001.42 694.89 416.75 218.67

SE 7.81 17.91 18.94 15.27 182.67 59.95 59.63 48.49

Range <LOD–

227.56

<LOD–

502.04

<LOD–

718.22

<LOD–

416.60

190.15–

5163.96

108.23–

1581.31

<LOD–

1561.41

<LOD–

1439.77

% <LOD 43 35 40 13 0 0 5 20

% <LOQ 45 35 40 13 0 0 5 20


Mean 45.76 68.65 36.58 100.59 2466.50 782.63 464.47 188.66

SE 14.09 24.39 15.26 20.11 285.47 93.99 92.70 52.03

Range <LOD–

227.56

<LOD–

402.86

<LOD–

288.42

<LOD–

299.50

329.87–

5163.96

108.23–

1581.31

<LOD–

1561.41

<LOD–

757.02

% <LOD 45 40 50 15 0 0 5 30

% <LOQ 45 40 50 15 0 0 5 30


Mean 24.36 76.53 62.67 93.73 1536.34 607.14 369.04 248.68

SE 6.29 26.85 34.95 23.49 180.89 71.48 75.94 82.77

Range <LOD–

89.91

<LOD–

502.04

<LOD–

718.22

<LOD–

416.60

190.15–

3650.99

129.22–

1456.65

25.31–

1172.39

<LOD–

1439.77

% <LOD 40 30 30 10 0 0 0 10

% <LOQ 45 30 30 10 0 0 0 10


Mean 37.46 * 38.61 83.18 1760.65 703.72 294.84 261.56

SE 3.88 * 2.13 13.58 307.54 80.05 15.44 42.52

Range 37.65 –

57.30 -

32.52 –

55.98

24.64–

161.75

613.75–

3978.95

444.92–

1303.42

229.10–

405.90

121.53–

577.79

% <LOD 0 100 0 0 0 0 0 0

% <LOQ 0 100 0 0 0 0 0 0

* Mean concentrations and standard error were not reported due to the low frequency of detection in the samples (<20%).


354

Table E6e. Summary of descriptive statistics for all detected analytes. For the purposes of calculating means,

values below the LOD were assigned a value of zero and values below the LOQ were used unaltered. For

analytes that were detected in <20% of the samples, mean concentrations were not calculated and only the range

is reported. Concentrations are reported in ng/L (ppt).

(ng/L) Analyte

PFBS PFHxS PFOS PFDS


Mean * 1249.05 12263.19 39.89

SE * 202.69 3794.29 6.36

Range <LOD–59.60 27.99 – 6795.84 143.96 – 119559.05 <LOD–155.26

% <LOD 85 0 0 35

% <LOQ 85 0 0 38


Mean * 1419.63 13295.01 36.53

SE * 250.53 5991.11 8.14

Range <LOD–59.60 185.63–4362.86 143.96 – 119559.05 <LOD–118.74

% <LOD 85 0 0 35

% <LOQ 85 0 0 35


Mean * 1078.46 11231.37 43.25

SE * 320.68 4805.89 9.93

Range <LOD–53.68 27.99–6795.84 778.07–75979.13 <LOD–155.26

% <LOD 85 0 0 35

% <LOQ 85 0 0 40


Mean 16.78 1193.81 4442.95 51.34

SE 8.55 177.07 462.25 3.76

Range <LOD–58.64 353.24–2039.20 2318.33 – 7209.94 40.76–82.39

% <LOD 70 0 0 0

% <LOQ 70 0 0 0

* Mean concentrations and standard error were not reported due to the low frequency of detection in the

samples (<20%).


355

Table E7. P-values from Mann-Whitney U test to compare concentrations between single donor and pooled sera

samples and for gender differences. A p-value of 0.05 is the chosen criterion of statistical significance such that

if the test statistic is below 0.05 (p<0.05), the null hypothesis may be rejected, and there is a significant difference

between the two groups of data. If the test statistic is above 0.05 (p>0.05), there is no significant difference

between the two groups of data. The Mann-Whitney U test was used to compare the concentrations observed in

the single donor and pooled sera samples, and the concentrations observed in male and female single donor

samples. In the gender comparison analysis, a one-sided p-value was calculated to test whether the

concentrations observed in female donors were lower as compared to male donors.

Analyte

Type of Comparison

Single donor vs. Pooled Female (F) vs. Male (M)

p-value

(two-sided)

p-value

(two-sided)

p-value

(one-sided; F<M)

4:2 diPAP 0.6211 0.6050 0.3025

4:2/6:2 diPAP 0.8581 0.5335 0.2668

6:2 diPAP 0.0246 0.9734 0.4867

6:2/8:2 diPAP 0.6848 0.6105 0.3053

8:2 diPAP 0.0751 0.6423 0.3212

FOSAA 0.1914 0.1382 0.0691

N-MeFOSAA 0.0699 0.2661 0.1331

N-EtFOSAA 0.0264 0.2999 0.1499

4:2 FTS 0.2589 0.5768 0.2884

6:2 FTS 0.0051 0.1995 0.0998

8:2 FTS 0.1285 0.2354 0.1177

C6/C6 PFPiA 0.0377 0.1496 0.0748

C6/C8 PFPiA 0.0065 0.1233 0.0617

C8/C8 PFPiA 0.4612 0.4872 0.2436

C6/C10 PFPiA 0.1707 0.1302 0.0651

C8/C10 PFPiA 0.4612 * *

C6/C12 PFPiA 0.8322 0.1257 0.0628

PFBA (C4) 0.2010 0.4484 0.2242

PFPeA (C5) * 0.7476 0.3738

PFHxA (C6) 0.1187 0.3486 0.1743

PFHpA (C7) 0.7469 0.7581 0.3790

PFOA (C8) 0.6242 0.0122 0.0061

PFNA (C9) 0.8392 0.1738 0.0869

PFDA (C10) 0.8734 0.4568 0.2284

PFUnA (C11) 0.0363 0.5871 0.2935

PFBS (C4) 0.2005 0.8984 0.4492

PFHxS (C6) 0.4224 0.1081 0.0540

PFOS (C8) 0.8955 0.4612 0.2306

PFDS (C10) 0.1780 0.7748 0.3874

* Test was not performed due to 100% non-detection in the samples.

356

Table E8. P-values from Mann-Whitney U test to compare concentrations of 6:2 and 8:2 FTS observed in

pooled human sera collected in 2002 and 2009. A p-value of 0.05 is the chosen criterion of statistical significance

such that if the test statistic is below 0.05 (p<0.05), the null hypothesis may be rejected, and there is a significant

difference between the two groups of data. If the test statistic is above 0.05 (p>0.05), there is no significant

difference between the two groups of data. The Mann-Whitney U test was used to compare the concentrations

observed in the single donor and pooled sera samples, and the concentrations observed in male and female single

donor samples. In the gender comparison analysis, a one-sided p-value was calculated to test whether the

concentrations observed in female donors were lower as compared to male donors.

Analyte

Type of Comparison

2002 pooled seraa vs. 2009 pooled sera

b

p-value (two-sided)

6:2 FTS 0.3915

8:2 FTS 0.8968

a Data obtained from ref. (11).

b Data obtained from this study.

357

Table E9. Spearman’s rank correlation coefficient r-values and p-values from Spearman’s rank correlation test to analyze two groups of

data for correlation. A p-value of 0.05 is the chosen criterion of statistical significance such that if the test statistic is below 0.05 (p<0.05),

the null hypothesis may be rejected, and there is a significant correlation between the two groups of data. If the test statistic is above 0.05

(p>0.05), there is no significant correlation between the two groups of data. The value of r always falls between -1 and +1. The closer r

falls to +1 or -1, the greater the correlation. The closer r is to 0, the lesser the correlation. In each cell, the top row represents the test

performed on the concentrations between single donor samples and the bottom row represents the test performed on the concentrations

between the pooled samples.

Single donor C6/C6 PFPiA C6/C8 PFPiA

C8/C8

PFPiA C6/C10 PFPiA

C8/C10

PFPiA C6/C12 PFPiA

Pooled

C6/C6 PFPiA n/a r=0.76; p<0.0001

* r=0.66; p<0.0001

* r=0.48; p=0.0019

r=0.83; p=0.0047 r=0.50; p=0.1548 *

C6/C8 PFPiA r=0.76; p<0.0001

n/a * r=0.78; p<0.0001

* r=0.56; p=0.0002

r=0.83; p=0.0047 r=0.83; p=0.0047 *

C8/C8 PFPiA * * n/a * * *

C6/C10 PFPiA r=0.66; p<0.0001 r=0.78; p<0.0001

* n/a * r=0.60; p<0.0001a

r=0.50; p=0.1548 r=0.83; p=0.0047

C8/C10 PFPiA * * * * n/a *

C6/C12 PFPiA r=0.48; p=0.0019 r=0.56; p=0.0002

* r=0.60; p<0.0001a

* n/a * *

n/a Correlation tests were not performed for the concentrations of the same analyte.

* Correlation tests were not performed due to the large number of non-detects observed for these analytes. a The correlation test to compare C6/C10 and C6/C12 PFPiA was performed on the concentrations combined from the single donor

and pooled samples as the Mann-Whitney U test showed no significant difference in their concentrations from both sample types

(p>0.05, Table S7).

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