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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)
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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
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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.
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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
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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.
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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
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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
2.6 Literature cited 105
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
3.7 Literature cited 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
4.2 Introduction 136
4.3 Experimental section 139
4.3.1 Materials 139
4.3.2 Soil-plant microcosm experiment 139
4.3.3 Sampling, extraction, and analysis 140
4.3.4 Quality assurance of data 141
4.3.5 Data analysis 142
4.4 Results and discussion 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
4.6 Acknowledgements 153
4.7 Literature cited 153
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
5.2 Introduction 159
5.3 Experimental section 162
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
5.3.5 Quality assurance of data 165
5.3.6 Data analysis 166
5.4 Results and discussion 167
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
5.6 Acknowledgements 177
5.7 Literature cited 177
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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
6.2 Introduction 184
6.3 Experimental section 186
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
6.3.6 Quality assurance of data 188
6.3.7 Data analysis 189
6.3.8 Statistical analysis 190
6.4 Results and discussion 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
6.6 Acknowledgements 201
6.7 Literature cited 201
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
7.2 Introduction 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
7.3.4 Quality assurance of data 212
7.3.5 Statistical analysis 214
7.4 Results and discussion 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
7.6 Acknowledgements 221
7.7 Literature cited 222
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CHAPTER EIGHT – Summary, Conclusions, and Future Work 227
8.1 Summary and conclusions 228
8.2 Future research directions 232
8.3 Literature cited 234
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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).
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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.
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CHAPTER SEVEN
Table 7.1 Structures, full names, and acronyms of the target analytes 211
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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
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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
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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.
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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.
Author list – Holly Lee and Scott Mabury
Contributions – Prepared by Holly Lee with editorial comments provided by Scott Mabury.
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
Published in – Environ. Sci. Technol. 2012, 46, 3489-3497
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.
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xix
Chapter Seven – A Pilot Survey of Legacy and Current Commercial Fluorinated Chemicals in
Human Sera from United States Donors in 2009
Published in – Environ. Sci. Technol. 2011, 45, 8067-8074
Author list – Holly Lee and Scott Mabury
Contributions – Prepared by Holly Lee with editorial comments provided by Scott Mabury.
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
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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.
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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
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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
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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.
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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
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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).
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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.
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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
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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).
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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,
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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
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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.
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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,
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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
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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
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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.
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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.
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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 - -
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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
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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
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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
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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
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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.
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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
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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
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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
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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
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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
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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).
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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
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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).
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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
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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.).
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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
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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
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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).
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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
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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
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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
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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
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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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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,
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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.
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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).
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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
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Elimination
Metabolism
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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.
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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
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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
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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).
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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
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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
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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.
1.6 Literature Cited
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51
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(30) Brace, N. O.; Mackenzie, A. K. Polyfluoroalkyl Phosphates 1963.
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(45) Perfluorinated Carboxylic Acids (PFCAs) and Precursors: An Action Plan For Assessment
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(78) DuPont Zonyl FSP Fluorosurfactant, technical information; DuPont.
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CHAPTER TWO
Global Distribution of Polyfluoroalkyl and Perfluoroalkyl Substances and their
Transformation Products in Environmental Solids
Holly Lee and Scott A. Mabury
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
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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
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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).
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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 River 102.91 2.90 0.13-63.43 (49)
Mainland China River 2.95 2.94 1.35-2.94 (50)
Mainland China River 0.96 0.41 0.01-0.35 (51)
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
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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
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(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
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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.
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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).
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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
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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
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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
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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
Sludge-amended soils (0-10 cm) 2009
Sludge-amended soils (36-56 cm) 2009
Sludge-amended soils (152-165 cm) 2009
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
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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
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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).
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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)
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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
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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
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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.
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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
Edible Parts 0.050 0.050
Peels 0.040 0.030
Leaves, Stalks,
and Roots 0.530 0.320
Cucumber
Edible Parts 0.030 0.000
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.
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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
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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
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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.
2.6 Literature Cited
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(131) DuPont Zonyl FSE Fluorosurfactant, technical information; DuPont.
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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
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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
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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
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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.
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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
Saturated fluorotelomer
carboxylic acid
x:3 FTCA
x = 3, 5, 7, 9
Perfluorocarboxylic acid PFCA
x = 3 – 10
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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
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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.
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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
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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)
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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
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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.
0 20 40 60 80 100
0
10
2050
100
150
200
0 20 40 60 80 100
Am
ou
nt
of
6:2
PA
Ps in
aq
ueo
us p
has
e
an
d 6
:2 F
TO
H c
um
ula
tively
str
ipp
ed
fro
m s
yste
m (
nm
ol)
0
50
100
150
200
6:2 diPAP
6:2 monoPAP
6:2 FTOH
0 20 40 60 80 100
Am
ou
nt
of
aq
ueo
us m
eta
bo
lite
sin
bo
ttle
(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
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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
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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.
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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
Am
ou
nt
of
mo
no
PA
Ps in
aq
ueo
us p
hase
an
d F
TO
Hs c
um
ula
tively
str
ipp
ed
fro
m s
yste
m (
nm
ol)
0
20
40
600
800
1000
1200
4:2 monoPAP4:2 FTOH
* * * * *
0 20 40 60 80 100
0
20
40
60
80
100
120
6:2 monoPAP
6:2 FTOH
* * * * *
Time (days)
0 20 40 60 80 100
0
10
20
30
40
50
8:2 monoPAP
8:2 FTOH
0 20 40 60 80 100
0
10
20
30
40
50
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
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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
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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.
3.7 Literature Cited
(1) Boulanger, B.; Peck, A. M.; Schnoor, J. L.; Hornbuckle, K. C. Mass Budget of
Perfluorooctane Surfactants in Lake Ontario. Environ. Sci. Technol. 2005, 39, 74–79.
(2) Huset, C. A.; Chiaia, A. C.; Barofsky, D. F.; Jonkers, N.; Kohler, H.-P. E.; Ort, C.; Giger,
W.; Field, J. A. Occurrence and Mass Flows of Fluorochemicals in the Glatt Valley
Watershed, Switzerland. Environ. Sci. Technol. 2008, 42, 6369–6377.
(3) Schultz, M. M.; Barofsky, D. F.; Field, J. A. Quantitative Determination of Fluorinated
Alkyl Substances by Large-Volume-Injection Liquid Chromatography Tandem Mass
SpectrometryCharacterization of Municipal Wastewaters. Environ. Sci. Technol. 2006, 40,
289–295.
(4) Schultz, M. M.; Higgins, C. P.; Huset, C. A.; Luthy, R. G.; Barofsky, D. F.; Field, J. A.
Fluorochemical Mass Flows in a Municipal Wastewater Treatment Facility. Environ. Sci.
Technol. 2006, 40, 7350–7357.
(5) Sinclair, E.; Kannan, K. Mass Loading and Fate of Perfluoroalkyl Surfactants in
Wastewater Treatment Plants. Environ. Sci. Technol. 2006, 40, 1408–1414.
(6) Dinglasan, M. J. A.; Ye, Y.; Edwards, E. A.; Mabury, S. A. Fluorotelomer Alcohol
Biodegradation Yields Poly- and Perfluorinated Acids. Environ. Sci. Technol. 2004, 38,
2857–2864.
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(7) Wang, N.; Szostek, B.; Folsom, P. W.; Sulecki, L. M.; Capka, V.; Buck, R. C.; Berti, W.
R.; Gannon, J. T. Aerobic Biotransformation of 14C-Labeled 8-2 Telomer B Alcohol by
Activated Sludge from a Domestic Sewage Treatment Plant. Environ. Sci. Technol. 2005,
39, 531–538.
(8) Wang, N.; Szostek, B.; Buck, R. C.; Folsom, P. W.; Sulecki, L. M.; Capka, V.; Berti, W.
R.; Gannon, J. T. Fluorotelomer Alcohol Biodegradation - Direct Evidence that
Perfluorinated Carbon Chains Breakdown. Environ. Sci. Technol. 2005, 39, 7516–7528.
(9) Wang, N.; Szostek, B.; Buck, R. C.; Folsom, P. W.; Sulecki, L. M.; Gannon, J. T. 8-2
Fluorotelomer Alcohol Aerobic Soil Biodegradation: Pathways, Metabolites, and
Metabolite Yields. Chemosphere. 2009, 75, 1089–1096.
(10) Martin, J. W.; Mabury, S. A.; O’Brien, P. J. Metabolic Products and Pathways of
Fluorotelomer Alcohols in Isolated Rat Hepatocytes. Chem. Biol. Interact. 2005, 155,
165–180.
(11) Nabb, D. L.; Szostek, B.; Himmelstein, M. W.; Mawn, M. P.; Gargas, M. L.; Sweeney, L.
M.; Stadler, J. C.; Buck, R. C.; Fasano, W. J. In Vitro Metabolism of 8-2 Fluorotelomer
Alcohol: Interspecies Comparisons and Metabolic Pathway Refinement. Toxicol. Sci.
2007, 100, 333–344.
(12) Hagen, D. F.; Belisle, J.; Johnson, J. D.; Venkateswarlu, P. Characterization of Fluorinated
Metabolites by a Gas Chromatographic-Helium Microwave Plasma Detector—The
Biotransformation of 1H,1H,2H,2H-Perfluorodecanol to Perfluorooctanoate. Anal.
Biochem. 1981, 118, 336–343.
(13) Telomer Research Program Update; U.S. EPA Public Docket AR226-1141; U.S. EPA
OPPT: Washington, DC, 2002.
(14) Russell, M. H.; Berti, W. R.; Szostek, B.; Buck, R. C. Investigation of the Biodegradation
Potential of a Fluoroacrylate Polymer Product in Aerobic Soils. Environ. Sci. Technol.
2008, 42, 800–807.
(15) Washington, J. W.; Ellington, J. J.; Jenkins, T. M.; Evans, J. J.; Yoo, H.; Hafner, S. C.
Degradability of an Acrylate-Linked, Fluorotelomer Polymer in Soil. Environ. Sci.
Technol. 2009, 43, 6617–6623.
(16) DuPont Zonyl FSE Fluorosurfactant, technical information; DuPont.
(17) DuPont Zonyl UR Fluorosurfactant, technical information; DuPont.
(18) DuPont Zonyl RP Paper Fluorosurfactant, technical information; DuPont.
(19) Indirect Food Additives: Paper and Paperboard Components.; Code of Federal
Regulations, 21 CFR 176.170; U.S. Food and Drug Administration; U.S. Government
Printing Office: Washington, DC, 2003.
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(20) Begley, T. H.; White, K.; Honigfort, P.; Twaroski, M. L.; Neches, R.; Walker, R. A.
Perfluorochemicals: Potential Sources of and Migration from Food Packaging. Food
Addit. Contam. 2005, 22, 1023–1031.
(21) Begley, T. H.; Hsu, W.; Noonan, G.; Diachenko, G. Migration of Fluorochemical Paper
Additives from Food-Contact Paper into Foods and Food Simulants. Food Addit. Contam.
2008, 25, 384–390.
(22) D’eon, J. C.; Mabury, S. A. Production of Perfluorinated Carboxylic Acids (PFCAs) from
the Biotransformation of Polyfluoroalkyl Phosphate Surfactants (PAPS): Exploring
Routes of Human Contamination. Environ. Sci. Technol. 2007, 41, 4799–4805.
(23) 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.
(24) Brace, N. O.; Mackenzie, A. K. Polyfluoroalkyl Phosphates 1963.
(25) Dinglasan, M. J. A.; Mabury, S. A. Significant Residual Fluorinated Alcohols Present in
Various Fluorinated Materials. Environ. Sci. Technol. 2006, 40, 1447–1453.
(26) Keith, L. H.; Crummett, W.; Deegan, J.; Libby, R. A.; Taylor, J. K.; Wentler, G. Principles
of Environmental Analysis. Anal. Chem. 1983, 55, 2210–2218.
(27) Butt, C. M.; Muir, D. C. G.; Mabury, S. A. Elucidating the Pathways of Poly- and
Perfluorinated Acid Formation in Rainbow Trout. Environ. Sci. Technol. 2010, 44, 4973–
4980.
(28) Liu, J.; Wang, N.; Szostek, B.; Buck, R. C.; Panciroli, P. K.; Folsom, P. W.; Sulecki, L.
M.; Bellin, C. A. 6-2 Fluorotelomer Alcohol Aerobic Biodegradation in Soil and Mixed
Bacterial Culture. Chemosphere. 2010, 78, 437–444.
(29) Fasano, W. J.; Sweeney, L. M.; Mawn, M. P.; Nabb, D. L.; Szostek, B.; Buck, R. C.;
Gargas, M. L. Kinetics of 8-2 Fluorotelomer Alcohol and Its Metabolites, and Liver
Glutathione Status Following Daily Oral Dosing for 45 Days in Male and Female Rats.
Chem. Biol. Interact. 2009, 180, 281–295.
(30) Borggaard, O. K.; Gimsing, A. L. Fate of Glyphosate in Soil and the Possibility of
Leaching to Ground and Surface Waters: A Review. Pest Manag. Sci. 2008, 64, 441–456.
(31) Wolfenden, R.; Ridgway, C.; Young, G. Spontaneous Hydrolysis of Ionized Phosphate
Monoesters and Diesters and the Proficiencies of Phosphatases and Phosphodiesterases as
Catalysts. J. Am. Chem. Soc. 1998, 120, 833–834.
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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
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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
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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
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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.
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4.3 Experimental Section
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
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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
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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.
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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 Results and Discussion
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
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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).
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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
nc
en
tra
tio
n o
f P
FA
Ss
in
So
il (
ng
/g)
0.01
0.1
1
10
100
1000
Soil without Biosolids Amendment
WWTP Biosolids-Amended Soil
WWTP and Paper Fiber Biosolids-Amended Soil
0.01
0.1
1
10
100
1000
0.01
0.1
1
10
100
1000
0.01
0.1
1
10
100
1000
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
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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.
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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.
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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.
Co
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n o
f 6
:2 d
iPA
P i
n S
oil
(n
g/g
)
0
10000
20000
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60000
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6:2 diPAP
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f F
TC
As
, F
TU
CA
s,
an
d P
FC
As
in
So
il (
ng
/g)
0
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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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:2 d
iPA
P i
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lan
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/g)
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10000
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TC
As
, F
TU
CA
s,
an
d P
FC
As
in
Pla
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(n
g/g
)
0
500
1000
1500
6:2 FTCA
6:2 FTUCA
5:3 FTCA
5:3 FTUCA
Time (Month)
0 1 2 3 4 5 6
0
20000
40000
60000
80000
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
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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
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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.
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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.
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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
nt-
So
il A
ccu
mu
lati
on
F
ac
tor
(PS
AF
)
0
50
100
150
5000
10000
15000
20000
25000
Cplant
/CWWTP Biosolids-Amended Soil
Cplant
/CWWTP- and Paper Fiber Biosolids-Amended Soil
3 4 5 6 7 8 9 10 11 12 13 14 15
log
(Pla
nt-
So
il A
cc
um
ula
tio
n
Fa
cto
r, P
SA
F)
-2
-1
0
1
2
3
4
5
Carbon Chain Length
3 4 5 6 7 8 9
Pla
nt-
So
il A
cc
um
ula
tio
n
Fa
cto
r (P
SA
F)
0
10
20
30
40
50
300
400
Cplant
/C6:2 diPAP-spiked in WWTP Biosolids-Amended Soil
3 4 5 6 7 8 9
log
(Pla
nt-
So
il A
cc
um
ula
tio
n
Fa
cto
r, P
SA
F)
0.0
0.5
1.0
1.5
2.0
2.5
3.0
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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.
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4.6 Acknowledgements
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.
4.7 Literature Cited
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Metals in Tile Drainage and Groundwater Following Applications of Municipal Biosolids
to Agricultural Fields. Sci. Tot. Environ. 2010, 408, 873–883.
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Perfluorochemicals and Fluorotelomer Alcohols in Plants from Biosolid-Amended Fields
using LC/MS/MS and GC/MS. Environ. Sci. Technol. 2011, 45, 7985–7990.
(25) Lindstrom, A. B.; Strynar, M. J.; Delinsky, A. D.; Nakayama, S. F.; McMillan, L.; Libelo,
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Compound Contamination of Surface and Well Water in Decatur, Alabama, USA.
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(26) Yoo, H.; Washington, J. W.; Ellington, J. J.; Jenkins, T. M.; Neill, M. P. Concentrations,
Distribution, and Persistence of Fluorotelomer Alcohols in Sludge-Applied Soils near
Decatur, Alabama, USA. Environ. Sci. Technol. 2010, 44, 8397–8402.
(27) Indirect Food Additives: Paper and Paperboard Components.; Code of Federal
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Printing Office: Washington, DC, 2003.
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(30) D’eon, J. C.; Mabury, S. A. Production of Perfluorinated Carboxylic Acids (PFCAs) from
the Biotransformation of Polyfluoroalkyl Phosphate Surfactants (PAPS): Exploring
Routes of Human Contamination. Environ. Sci. Technol. 2007, 41, 4799–4805.
(31) Guidelines for the Utilization of Biosolids and Other Wastes on Agricultural Land;
Ministry of Environment and Ministry of Agriculture, Food, and Rural Affairs, 1996.
(32) Yoshida, T.; Iida, S. Process for Preparing Di(Fluoroalkyl Containing Group-Substituted
Alkyl) Phosphate Salt 1991.
(33) DuPont Zonyl FSE Fluorosurfactant, technical information; DuPont.
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(34) Telomer Research Program Update; U.S. EPA Public Docket AR226-1141; U.S. EPA
OPPT: Washington, DC, 2002.
(35) Washington, J. W.; Henderson, W. M.; Ellington, J. J.; Jenkins, T. M.; Evans, J. J.
Analysis of Perfluorinated Carboxylic Acids in Soils II: Optimization of Chromatography
and Extraction. J. Chrom. A. 2008, 1181, 21–32.
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(37) Enevoldsen, R.; Juhler, R. K. Perfluorinated Compounds (PFCs) in Groundwater and
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(38) Dinglasan, M. J. A.; Ye, Y.; Edwards, E. A.; Mabury, S. A. Fluorotelomer Alcohol
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(42) EPA - DuPont Telomers Degradation Technical Meeting; Washington, DC, 2004.
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(46) Wang, N.; Buck, R. C.; Szostek, B.; Sulecki, L. M.; Wolstenholme, B. W. 5:3
Polyfluorinated Acid Aerobic Biotransformation in Activated Sludge Via Novel ―One-
Carbon Removal Pathways.‖Chemosphere. 2012, 87, 527–534.
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Fluorotelomer Alcohol Aerobic Soil Biodegradation: Pathways, Metabolites, and
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CHAPTER FIVE
Sorption of Perfluoroalkyl Phosphonates and Perfluoroalkyl Phosphinates in Soil
Holly Lee and Scott A. Mabury
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
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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
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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).
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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
3 congeners monitored
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
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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 Experimental Section
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
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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
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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
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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.
5.3.5 Quality Assurance of Data
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
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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.
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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 Results and Discussion
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).
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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
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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
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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
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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
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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).
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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
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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).
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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.
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
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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
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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.
5.6 Acknowledgements
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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Occurrence and Pathways of Degradation. Appl. Microbiol. Biotechnol. 1995, 43, 545–
550.
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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
Published as: Environ. Sci. Technol. 2012, 46, 3489-3497.
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
Copyright American Chemical Society 2012
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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
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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.
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Table 6.1. 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
3 congeners monitored
OF
F F
O-x
Perfluorocarboxylate
Cx+1 PFCA
x = 4 – 10
7 congeners monitored
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 Experimental Section
6.3.1 Chemicals
A list of all standards and reagents used in this study is provided in the Supporting
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
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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
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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.
6.3.6 Quality Assurance of Data
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
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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.
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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 Results and Discussion
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
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/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.
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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
Exposure Depuration Exposure Depuration Exposure Depuration
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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
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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
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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.
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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).
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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
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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.
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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
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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
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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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Depuration of Individual C10-, C11- and C14-Polychlorinated Alkanes by Juvenile
Rainbow Trout (Oncorhynchus mykiss). Aquat. Toxicol. 1998, 43, 209–221.
(59) Fisk, A. T.; Tomy, G. T.; Cymbalisty, C. D.; Muir, D. C. G. Dietary Accumulation and
Quantitative Structure-Activity Relationships for Depuration and Biotransformation of
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Biotransformation, and Metabolite Formation of Fipronil and Chiral Legacy Pesticides in
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Acrylate in Rainbow Trout. 1. In vivo Dietary Exposure. Environ. Toxicol. Chem. 2010, 29,
2726–2735.
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Gamberg, M.; Muir, D. C. G. Biomagnification of Perfluorinated Compounds in a Remote
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8673.
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CHAPTER SEVEN
A Pilot Survey of Legacy and Current Commercial Fluorinated Chemicals in Human Sera
from United States Donors in 2009
Holly Lee and Scott A. Mabury
Published as: Environ. Sci. Technol. 2011, 45, 8067-8074.
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
Copyright American Chemical Society 2011
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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
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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
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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
A list of all standards and reagents used in this study is provided in the Supporting
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
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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
Perfluorophosphonate
(Cx PFPA)
x = 6, 8, 10
Perfluorophosphinate
(Cx/Cy PFPiA)
x = 6, 8
y = 6, 8, 10, 12
x+y ≤ 18
Perfluorocarboxylate
(PFCA)
x = 4–14
Perfluorosulfonate
(PFSA)
x = 4, 6, 8, 10
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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.
7.3.4 Quality Assurance of Data
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.
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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
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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 Results and Discussion
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
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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).
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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)
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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
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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.
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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.
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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
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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.
7.6 Acknowledgements
The authors would like to thank Susanne Waaijers (University of Amsterdam,
Netherlands), Alexandra Tevlin and Barbara Weiner (University of Toronto, Toronto, ON) for
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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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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, 439–451.
(37) Perfluoroalcohol Phosphate Treatment, technical literature; Pft-001; Kobo Products:
Plainfield, NJ, U.S.
(38) Hazard Assessment of Perfluorooctane Sulfonate (PFOS) and Its Salts;
ENV/JM/RD(2002)17/FINAL; OECD Environment Directorate: Paris, France, 2002.
(39) Results of the 2006 Survey on Production and Use of PFOS, PFAS, PFOA, PFCA, their
Related Substances and Products/Mixtures Containing These Substances;
ENV/JM/MONO(2006)36, JT03219292; OECD: Paris, France, 2006.
(40) Perfluorooctyl Organic Phosphate; Wuhan Defu Economic Development Co., Ltd.:
Wuhan, China, 2007.
(41) Mulvana, D. E.; Henion, J. Qualitative Investigation of the In Vitro Metabolism of T-6292,
T-6293, T-6294, and T-6295 by Rat and Human Hepatocytes Using Ion Spray LC/MS and
LC/MS/MS; AR226-0328; Advanced Bioanalytical Services, Inc.: Ithaca, NY, 1996.
(42) Tittlemier, S. A.; Pepper, K.; Edwards, L. Concentrations of Perfluorooctanesulfonamides
in Canadian Total Diet Study Composite Food Samples Collected between 1992 and 2004.
J. Agric. Food Chem. 2006, 54, 8385–8389.
(43) Shoeib, M.; Harner, T.; Ikonomou, M.; Kannan, K. Indoor and Outdoor Air
Concentrations and Phase Partitioning of Perfluoroalkyl Sulfonamides and
Polybrominated Diphenyl Ethers. Environ. Sci. Technol. 2004, 38, 1313–1320.
(44) Shoeib, M.; Harner, T.; Wilford, B. H.; Jones, K. C.; Zhu, J. Perfluorinated Sulfonamides
in Indoor and Outdoor Air and Indoor Dust: Occurrence, Partitioning, and Human
Exposure. Environ. Sci. Technol. 2005, 39, 6599–6606.
(45) Connolly, P. D. E.; Zhu, X.; Keller, R. Analysis of Pooled Human Sera and Plasma and
Monkey Sera for Fluorocarbons Using Exygen Method ExM-023-071; AR226-1152;
Prepared for 3M Environmental Laboratory: St. Paul, MN, 2002.
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(46) DuPont Performance Chemicals. Zonyl FS-62 Fluorosurfactant, technical information;
DuPont.
(47) Chemguard FS-220B MSDS; Chemguard Inc.
(48) Dynax DX5022 Fluorochemical Foam Stabilizer, technical information; DynaxCo.
(49) Yeung, L. W. Y.; Taniyasu, S.; Kannan, K.; Xu, D. Z. Y.; Guruge, K. S.; Lam, P. K. S.;
Yamashita, N. An Analytical Method for the Determination of Perfluorinated Compounds
in Whole Blood Using Acetonitrile and Solid Phase Extraction Methods. J. Chrom. A.
2009, 1216, 4950–4956.
(50) Commission Recommendation of 17 March 2010 on the Monitoring of Perfluoroalkylated
Substances in Food (Text with EEA Relevance); 2010/161/EU; EFSA, European Union:
Brussels, Belgium, 2010.
(51) Recommendation XXXVI. Paper and Board for Food Contact; Federal Institute for Risk
Assessment, 2009.
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CHAPTER EIGHT
Summary, Conclusions, and Future Work
Holly Lee and Scott A. Mabury
Contributions: Holly Lee prepared this chapter with additional comments provided by Scott
Mabury
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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
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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
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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
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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.
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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
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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
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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.
8.3 Literature cited
(1) Indirect Food Additives: Paper and Paperboard Components.; Code of Federal
Regulations, 21 CFR 176.170; U.S. Food and Drug Administration; U.S. Government
Printing Office: Washington, DC, 2003.
(2) Brace, N. O.; Mackenzie, A. K. Polyfluoroalkyl Phosphates 1963.
(3) Perfluoroalcohol Phosphate Treatment, technical literature; Pft-001; Kobo Products:
Plainfield, NJ, U.S.
(4) DuPont Zonyl FSE Fluorosurfactant, technical information; DuPont.
(5) DuPont Zonyl FSJ Fluorosurfactant, technical information; DuPont.
(6) DuPont Zonyl FSP Fluorosurfactant, technical information; DuPont.
(7) DuPont Zonyl UR Fluorosurfactant, technical information; DuPont.
(8) Dinglasan, M. J. A.; Ye, Y.; Edwards, E. A.; Mabury, S. A. Fluorotelomer Alcohol
Biodegradation Yields Poly- and Perfluorinated Acids. Environ. Sci. Technol.2004, 38,
2857–2864.
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(9) Wang, N.; Szostek, B.; Buck, R. C.; Folsom, P. W.; Sulecki, L. M.; Capka, V.; Berti, W.
R.; Gannon, J. T. Fluorotelomer Alcohol Biodegradation - Direct Evidence that
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(10) Wang, N.; Szostek, B.; Folsom, P. W.; Sulecki, L. M.; Capka, V.; Buck, R. C.; Berti, W.
R.; Gannon, J. T. Aerobic Biotransformation of 14C-Labeled 8-2 Telomer B Alcohol by
Activated Sludge from a Domestic Sewage Treatment Plant. Environ. Sci. Technol.2005,
39, 531–538.
(11) Liu, J.; Lee, L. S.; Nies, L. F.; Nakatsu, C. H.; Turco, R. F. Biotransformation of 8:2
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(12) Wang, N.; Szostek, B.; Buck, R. C.; Folsom, P. W.; Sulecki, L. M.; Gannon, J. T. 8-2
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(13) Liu, J.; Wang, N.; Szostek, B.; Buck, R. C.; Panciroli, P. K.; Folsom, P. W.; Sulecki, L. M.;
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Bacterial Culture. Chemosphere.2010, 78, 437–444.
(14) Liu, J.; Wang, N.; Buck, R. C.; Wolstenholme, B. W.; Folsom, P. W.; Sulecki, L. M.;
Bellin, C. A. Aerobic Biodegradation of [14C] 6:2 Fluorotelomer Alcohol in a Flow-
Through Soil Incubation System. Chemosphere.2010, 80, 716–723.
(15) 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.
(16) Sinclair, E.; Kannan, K. Mass Loading and Fate of Perfluoroalkyl Surfactants in
Wastewater Treatment Plants. Environ. Sci. Technol.2006, 40, 1408–1414.
(17) Yoo, H.; Washington, J. W.; Jenkins, T. M.; Ellington, J. J. Quantitative Determination of
Perfluorochemicals and Fluorotelomer Alcohols in Plants from Biosolid-Amended Fields
using LC/MS/MS and GC/MS. Environ. Sci. Technol.2011, 45, 7985–7990.
(18) Stahl, T.; Heyn, J.; Thiele, H.; Hüther, J.; Failing, K.; Georgii, S.; Brunn, H. Carryover of
Perfluorooctanoic Acid (PFOA) and Perfluorooctane Sulfonate (PFOS) from Soil to Plants.
Arch. Environ. Contam. Toxicol.2008, 57, 289–298.
(19) Lechner, M.; Knapp, H. Carryover of Perfluorooctanoic Acid (PFOA) and Perfluorooctane
Sulfonate (PFOS) from Soil to Plant and Distribution to the Different Plant Compartments
Studied in Cultures of Carrots (Daucus carota ssp. Sativus), Potatoes (Solanum tuberosum),
and Cucumbers (Cucumis Sativus). J. Agric. Food Chem.2011, 59, 11011–11018.
(20) D’eon, J. C.; Crozier, P. W.; Furdui, V. I.; Reiner, E. J.; Libelo, E. L.; Mabury, S. A.
Perfluorinated Phosphonic Acids in Canadian Surface Waters and Wastewater Treatment
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Plant Effluent: Discovery of a New Class of Perfluorinated Acids. Environ. Toxicol.
Chem.2009, 28, 2101.
(21) D’eon, J. C.; Mabury, S. A. Uptake and Elimination of Perfluorinated Phosphonic Acids in
the Rat. Environ. Toxicol. Chem.2010, 29, 1319–1329.
(22) Guo, R.; Reiner, E. J.; Bhavsar, S. P.; Helm, P. A.; Mabury, S. A.; Braekevelt, E.;
Tittlemier, S. A. Determination of Polyfluoroalkyl Phosphoric Acid Diesters,
Perfluoroalkyl Phosphonic Acids, Perfluoroalkyl Phosphinic Acids, Perfluoroalkyl
Carboxylic Acids and Perfluoroalkane Sulfonic Acids in Lake Trout from the Great Lakes
Region. Anal. Bioanal. Chem.In press.
(23) Higgins, C. P.; Luthy, R. G. Sorption of Perfluorinated Surfactants on Sediments. Environ.
Sci. Technol.2006, 40, 7251–7256.
(24) Enevoldsen, R.; Juhler, R. K. Perfluorinated Compounds (PFCs) in Groundwater and
Aqueous Soil Extracts: Using Inline SPE-LC-MS/MS for Screening and Sorption
Characterisation of Perfluorooctane Sulphonate and Related Compounds. Anal. Bioanal.
Chem.2010, 398, 1161–1172.
(25) Esparza, X.; Moyano, E.; de Boer, J.; Galceran, M. T.; van Leeuwen, S. P. J. Analysis of
Perfluorinated Phosphonic Acids and Perfluorooctane Sulfonic Acid in Water, Sludge and
Sediment by LC–MS/MS. Talanta.2011, 86, 329–336.
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Distribution of Perfluorinated Acids in Rainbow Trout (Oncorhynchus mykiss). Environ.
Toxicol. Chem.2003, 22, 196–204.
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Transfer of Poly- and Perfluorinated Compounds in Chinese Sturgeon (Acipenser sinensis):
Implications for Reproductive Risk. Environ. Sci. Technol.2010, 44, 1868–1874.
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Alcohol in the Rat. Toxicol. Sci.2006, 91, 341–355.
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Fluorotelomer Alcohols in Isolated Rat Hepatocytes. Chem. Biol. Interact.2005, 155, 165–
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Fluorotelomer Unsaturated Acids and Aldehydes with Glutathione. Cell Biol. Toxicol.2012,
28, 115–124.
(35) Rand, A. A.; Mabury, S. A. In Vitro Interactions of Biological Nucleophiles with
Fluorotelomer Unsaturated Acids and Aldehydes: Fate and Consequences. Environ. Sci.
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APPENDIX A
SUPPORTING INFORMATION FOR CHAPTER THREE
Biodegradation of Polyfluoroalkyl Phosphates (PAPs) as a Source of Perfluorinated Acids
to the Environment
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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
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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
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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).
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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:
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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.
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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
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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.
(2) Hansen, K. J.; Clemen, L. A.; Ellefson, M. E.; Johnson, H. O. Compound-Specific,
Quantitative Characterization of Organic Fluorochemicals in Biological Matrices. Environ.
Sci. Technol. 2001, 35, 766–770.
(3) Higgins, C. P.; Luthy, R. G. Sorption of Perfluorinated Surfactants on Sediments. Environ.
Sci. Technol. 2006, 40, 7251–7256.
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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
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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
6:2 diPAP 0.606 mL of 660 ugmL
-1 stock
= 400 ug
18Ω deionized water 395 mL
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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
6:2 monoPAP 0.862 mL of 464 ugmL
-1 stock
= 400 ug
8:2 monoPAP 0.336 mL of 1190 ugmL
-1 stock
= 400 ug
10:2 monoPAP 0.820 mL of 488 ugmL
-1 stock
= 400 ug
6:2 diPAP 0.606 mL of 660 ugmL
-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
4:2 monoPAP 0.505 mL of 792 ugmL
-1 stock
= 400 ug
400 mL
6:2 monoPAP 0.862 mL of 464 ugmL
-1 stock
= 400 ug
8:2 monoPAP 0.336 mL of 1190 ugmL
-1 stock
= 400 ug
10:2 monoPAP 0.820 mL of 488 ugmL
-1 stock
= 400 ug
6:2 diPAP 0.606 mL of 660 ugmL
-1 stock =
400 ug
Mineral media 397 – 399 mL
Washed mixed
liquor
0 mL
Hg2Cl2 0 mL
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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
4:2 monoPAP 0.505 mL of 792 ugmL
-1 stock
= 400 ug
400 mL
6:2 monoPAP 0.862 mL of 464 ugmL
-1 stock
= 400 ug
8:2 monoPAP 0.336 mL of 1190 ugmL
-1 stock
= 400 ug
10:2 monoPAP 0.820 mL of 488 ugmL
-1 stock
= 400 ug
Mineral media 357 mL
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
6:2 monoPAP 0.862 mL of 464 ugmL
-1 stock
= 400 ug
400 mL 6:2 diPAP
0.606 mL of 660 ugmL-1
stock =
400 ug
Mineral media 359 mL
Washed mixed
liquor
40 mL
Hg2Cl2 0 mL
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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
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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.
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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
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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
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
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
Area count = 7690
Area count = 9120
Area count = 18400
Area count = 13500
Area count = 519000
Area count = 293000
Area count = 52200
Area count = 56600
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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
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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
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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.
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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)
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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)
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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
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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.
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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
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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
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263
LIST OF TABLES AND FIGURES
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
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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
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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.
Instrumental Analysis.
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
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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.
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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.
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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.
(2) D’eon, J. C.; Mabury, S. A. Production of Perfluorinated Carboxylic Acids (PFCAs) from
the Biotransformation of Polyfluoroalkyl Phosphate Surfactants (PAPS): Exploring
Routes of Human Contamination. Environ. Sci. Technol.2007, 41, 4799–4805.
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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
Saturated fluorotelomer
carboxylate
x:2 FTCA
x = 4, 6, 8, 10
F
F F
x
F
O
O-
Unsaturated fluorotelomer
carboxylate
x:2 FTUCA
x = 4, 6, 8, 10
F
F F
xO
O-
Saturated fluorotelomer
carboxylate
x:3 FTCA
x = 3, 5, 7, 9
F
F F
xO
O-
Unsaturated fluorotelomer
carboxylate
x:3 FTUCA
x = 3, 5, 7, 9
OF
F F
O-x
Perfluorocarboxylate PFCA
x = 3–13
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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
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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
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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
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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
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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
WWTP Biosolids-Amended Soil
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
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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
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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
8:2/10:2 diPAP8:2 diPAP
6:2/8:2 diPAP
% Catch Plates
% Soil
% Plant
WWTP Biosolids-Amended Soil
0 20 40 60 80 100
PFTeAPFTrA
PFDoAPFUnA
PFDAPFNAPFOA
PFHpAPFHxAPFPeAPFBA
10:2/12:2 diPAP10:2 diPAP
8:2/10:2 diPAP8:2 diPAP
6:2/8:2 diPAP6:2 diPAP
% Catch Plates
% Soil
% Plant
WWTP- and Paper Fiber Biosolids-Amended Soil
6:2 diPAP-Spiked and WWTP Biosolids-Amended Soil
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277
APPENDIX C
SUPPORTING INFORMATION FOR CHAPTER FIVE
Sorption of Perfluoroalkyl Phosphonates and Perfluoroalkyl Phosphinates in Soil
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278
LIST OF TABLES AND FIGURES
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
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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
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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.
Instrumental Analysis.
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.
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281
Quality Assurance of Data.
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.
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282
LITERATURE CITED
(1) D’eon, J. C.; Mabury, S. A. Uptake and Elimination of Perfluorinated Phosphonic Acids
in the Rat. Environ. Toxicol. Chem. 2010, 29, 1319–1329.
(2) Lee, H.; Mabury, S. A. A Pilot Survey of Legacy and Current Commercial Fluorinated
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.
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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.
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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
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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
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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
C8 perfluorophosphonate C8 PFPA 499.0>79.0 40 -70 -75 -10
C10 perfluorophosphonate C10 PFPA 599.0>79.0 40 -80 -90 -10
C6/C6 perfluorophosphinate C6/C6 PFPiA 701.0>401.0 40 -100 -75 -10
C6/C8 perfluorophosphinate C6/C8 PFPiA 801.0>401.0 40 -100 -95 -10
801.0>501.0 40 -100 -85 -10
C8/C8 perfluorophosphinate C8/C8 PFPiA 901.0>501.0 40 -100 -95 -10
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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
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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
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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
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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
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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
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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
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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
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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
C6 PFPA C8 PFPA C10 PFPA
C6/C6 PFPiA C6/C8 PFPiA C8/C8 PFPiA
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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
C6 PFPA C8 PFPA C10 PFPA
C6/C6 PFPiA C6/C8 PFPiA C8/C8 PFPiA
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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
C6 PFPA C8 PFPA C10 PFPA
C6/C6 PFPiA C6/C8 PFPiA C8/C8 PFPiA
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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
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APPENDIX D
SUPPORTING INFORMATION FOR CHAPTER SIX
Dietary Bioaccumulation of Perfluorophosphonates and Perfluorophosphinates in
Juvenile Rainbow Trout: Evidence of Metabolism of Perfluorophosphinates
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LIST OF TABLES AND FIGURES
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
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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
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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
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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.
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Instrumental Analysis.
Liquid Chromatography Details.
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
reequilibrate for 2.50 min. for a total run time of 10 min.
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.
Mass Spectrometry Details.
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).
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Quality Assurance of Data.
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
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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.
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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).
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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
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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.
(1) Martin, J. W.; Mabury, S. A.; Solomon, K. R.; Muir, D. C. G. Dietary Accumulation of
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.
(3) D’eon, J. C.; Crozier, P. W.; Furdui, V. I.; Reiner, E. J.; Libelo, E. L.; Mabury, S. A.
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.
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(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).
Environ. Toxicol. Chem. 2003, 22, 196–204.
(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.
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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
C8 perfluorophosphonate C8 PFPA 474 ± 37
C10 perfluorophosphonate C10 PFPA 533 ± 37
C6/C6 perfluorophosphinate C6/C6 PFPiA 468 ± 12
C6/C8 perfluorophosphinate C6/C8 PFPiA 510 ± 24
C8/C8 perfluorophosphinate C8/C8 PFPiA 420 ± 12
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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
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Table D2. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.
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
C8 perfluorophosphonate C8 PFPA 499.0>79.0 40 -70 -75 -10
C10 perfluorophosphonate C10 PFPA 599.0>79.0 40 -80 -90 -10
C6/C6 perfluorophosphinate C6/C6 PFPiA 701.0>401.0 40 -100 -75 -10
C6/C8 perfluorophosphinate C6/C8 PFPiA 801.0>401.0 40 -100 -95 -10
801.0>501.0 40 -100 -85 -10
C8/C8 perfluorophosphinate C8/C8 PFPiA 901.0>501.0 40 -100 -95 -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
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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
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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
In PFPA-dosed fishC8 PFPA499 > 79Area 18000
In PFPA-dosed fishC10 PFPA599 > 79Area 52900
Day 31 of uptake phase inwhole-body homogenate samples
In PFPiA-dosed fishC6/C6 PFPiA701 > 401Area 263000
In PFPiA-dosed fishC6/C8 PFPiA801 > 501Area 169000
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
In PFPA-dosed fishC6 PFPA399 > 79Area 21800
In PFPA-dosed fishC8 PFPA499 > 79Area 136000
In PFPA-dosed fishC10 PFPA599 > 79Area 615000
In PFPiA-dosed fishC6/C6 PFPiA701 > 401Area 1530000
In PFPiA-dosed fishC6/C8 PFPiA801 > 501Area 705000
In PFPiA-dosed fishC8/C8 PFPiA901 > 501Area 565000
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
In PFPA-dosed fishC6 PFPA399 > 79Area 5180
In PFPA-dosed fishC8 PFPA499 > 79Area 12300
In PFPA-dosed fishC10 PFPA599 > 79Area 44900
In PFPiA-dosed fishC6/C6 PFPiA701 > 401Area 859000
In PFPiA-dosed fishC6/C8 PFPiA801 > 501Area 452000
In PFPiA-dosed fishC8/C8 PFPiA901 > 501Area 410000
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
In PFPA-dosed fishC6 PFPA399 > 79Area 5050
In PFPA-dosed fishC8 PFPA499 > 79Area 2530
In PFPA-dosed fishC10 PFPA599 > 79Area 2840
In PFPiA-dosed fishC6/C6 PFPiA701 > 401Area 216000
In PFPiA-dosed fishC6/C8 PFPiA801 > 501Area 97300
In PFPiA-dosed fishC8/C8 PFPiA901 > 501Area 64800
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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
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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
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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
Exposure Depuration Exposure Depuration Exposure Depuration
(A) Control fish (B) PFPA-dosed fish (C) PFPiA-dosed fish
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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
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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
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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
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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
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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
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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
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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).
C6 PFPA C8 PFPA C10 PFPA
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
*
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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
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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
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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).
C6 PFPA C8 PFPA C10 PFPA
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
*
* ***
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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
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329
LIST OF TABLES AND FIGURES
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
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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,
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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.
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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
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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).
Instrumental Analysis.
Liquid Chromatography Details.
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
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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
reequilibrate for 3.5 min. for a total run time of 10 min.
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.
Mass Spectrometry Details.
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
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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).
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336
Quality Assurance of Data.
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
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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
1. D'eon, J. C.; Mabury, S. A., Production of perfluorinated carboxylic acids
(PFCAs) from the biotransformation of polyfluoroalkyl phosphate surfictants
(PAPS): Exploring routes of human contamination. Environ. Sci. Technol. 2007,
41, (13), 4799-4805.
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.
4. D'eon, J. C.; Crozier, P. W.; Furdui, V. I.; Reiner, E. J.; Libelo, E. L.; Mabury, S.
A., Perfluorinated Phosphonic Acids in Canadian Surface Waters and Wastewater
Treatment Plant Effluent: Discovery of a New Class of Perfluorinated Acids.
Environ. Toxicol. and Chem. 2009, 28, (10), 2101-2107.
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.
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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.
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Table E1a. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.
Analyte Acronym Mass
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
6:2 polyfluoroalkyl phosphate diester 6:2 diPAP 789.0>96.9 30 -65 -65 -15
789.0>443.0 20 -65 -27 -15
6:2/8:2 polyfluoroalkyl phosphate diester 6:2/8:2 diPAP 889.0>96.9 30 -70 -70 -15
8:2 polyfluoroalkyl phosphate diester 8:2 diPAP 989.0>96.9 30 -80 -75 -15
989.0>543.0 20 -70 -33 -15
8:2/10:2 polyfluoroalkyl phosphate diester 8:2/10:2 diPAP 1089.0>96.9 30 -80 -80 -15
10:2 polyfluoroalkyl phosphate diester 10:2 diPAP 1189.0>96.9 30 -80 -85 -15
1189.0>643.0 40 -80 -40 -15
10:2/12:2 polyfluoroalkyl phosphate diester 10:2/12:2 diPAP 1289.0>96.9 30 -80 -85 -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
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Table E1b. Multiple reaction monitoring (MRM) transitions and mass spectrometry parameters for all target analytes.
Analyte Acronym Mass
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
6:2 fluorotelomer sulfonate 6:2 FTS 427.0>81.0 20 -100 -65 -15
427.0>406.8 20 -100 -32 -10
8:2 fluorotelomer sulfonate 8:2 FTS 527.0>81.0 20 -100 -72 -14
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
C6 perfluorophosphonate C6 PFPA 399.0>79.0 40 -60 -75 -10
C8 perfluorophosphonate C8 PFPA 499.0>79.0 40 -70 -80 -10
C10 perfluorophosphonate C10 PFPA 599.0>79.0 40 -80 -90 -10
C6/C6 perfluorophosphinate C6/C6 PFPiA 701.0>401.0 40 -95 -75 -10
C6/C8 perfluorophosphinate C6/C8 PFPiA 801.0>501.0 40 -99 -85 -10
C8/C8 perfluorophosphinate C8/C8 PFPiA 901.0>501.0 40 -97 -90 -10
C6/C10 perfluorophosphinate C6/C10 PFPiA 901.0>601.0 40 -92 -90 -10
C8/C10 perfluorophosphinate C8/C10 PFPiA 1001.0>601.0 40 -97 -97 -10
C6/C12 perfluorophosphinate C6/C12 PFPiA 1001.0>701.0 40 -92 -98 -10
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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)
Perfluorocarboxylate
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
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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
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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.
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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 (%)
(n = 3) LOD LOQ LOD LOQ
(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
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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 (%)
(n = 3) LOD LOQ LOD LOQ
(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
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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)
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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
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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
C8/C8 PFPiA Single donor <0.0001 <0.0001
Pooled <0.0001 <0.0001
C6/C10 PFPiA Single donor <0.0001 <0.0001
Pooled <0.0001 0.1284b
C8/C10 PFPiA Single donor <0.0001 <0.0001
Pooled <0.0001 <0.0001
C6/C12 PFPiA Single donor <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.
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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.
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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
Range <LOD–21.51 <LOD–100.54 <LOD–388.55 <LOD–303.05 <LOD–777.24
% <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
Range <LOD–9.23 <LOD–55.54 <LOD–328.29 <LOD–70.54 <LOD–1801.74
% <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
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351
Table E6b. 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
FOSAA N-MeFOSAA N-EtFOSAA 4:2 FTS 6:2 FTS 8:2FTS
All single donor samples (n = 40)
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
Male single donor samples (n = 20)
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
Female single donor samples (n = 20)
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
* 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
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352
Table E6c. 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
C6/C6
PFPiA
C6/C8
PFPiA
C8/C8
PFPiA
C6/C10
PFPiAa
C8/C10
PFPiAa
C6/C12
PFPiAa
All single donor samples (n = 40)
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
Male single donor samples (n = 20)
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
Female single donor samples (n = 20)
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
All pooled donor samples (n = 10)
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
* 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 a Concentrations were not corrected based on corresponding percent distribution in
Masurf® 780 standard
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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
All single donor samples (n = 40)
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
Male single donor samples (n = 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
Female single donor samples (n = 20)
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
All pooled donor samples (n = 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%).
- Range was not reported due to 100% non-detection in the samples
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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
All single donor samples (n = 40)
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
Male single donor samples (n = 20)
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
Female single donor samples (n = 20)
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
All pooled donor samples (n = 10)
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%).
- Range was not reported due to 100% non-detection in the samples
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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.
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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.
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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).