CAPILLARY ELECTROPHORESIS METHOD FOR DETERMINATION OF
BISPHENOL A IN DRINKING WATER AND IN BRITISH COLUMBIA MUSSELS
by
Elizabeth Marie Andrucson
A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR
THE DEGREE OF MASTER OF SCIENCE IN ENVIRONMENTAL SCIENCE
In the Department of Physical Sciences (Chemistry)
Thesis Examining Committee:
Kingsley Donkor (PhD), Professor and Thesis Supervisor, Physical Sciences (Chemistry)
Heidi Huttunen-Hennelly (PhD), Associate Professor and Committee Member, Physical
Sciences (Chemistry)
Dipesh Prema (PhD), Associate Professor (Teaching) and Committee Member, Physical
Sciences (Chemistry)
James Kariuki (PhD), External Examiner, Professor, University of Alberta, Augustana
Campus

August 2021
Thompson Rivers University
(Kamloops)
© Elizabeth Marie Andrucson, 2021

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ABSTRACT

Bisphenol A has been used around the world for over 50 years and traces of it can be found
everywhere: in the air, soil, living things, and water. It is very useful for manufacturing
polycarbonate plastics, epoxy resins, and countless other products but there are concerns about
negative effects, especially on human health since it has been classified as an endocrine
disruptor. As such, knowing exactly where it is found is important information. Municipal
water supply from various locations in Kamloops, British Columbia were tested by a capillary
electrophoresis method developed in this study to detect bisphenol A as low as 5 parts per
million. Mussel samples from coastal British Columbia were also tested. None of the water
samples tested showed the presence of bisphenol A at the concentrations that were detectable
by this method. Mussel tissue analysis was inconclusive and requires further investigation to
confirm BPA concentrations around 5 parts per million. Other future work will include refining
the capillary electrophoresis method to detect lower concentrations and to attempt to detect
bisphenol analogues which are being used to replace bisphenol A in many products.

Keywords: capillary electrophoresis, bisphenol A, water, mussel

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TABLE OF CONTENTS:
Abstract .............................................................................................................................. ii
Table of Contents .............................................................................................................. iii
Acknowledgements .............................................................................................................v
Dedication ......................................................................................................................... vi
List of Abbreviations ........................................................................................................ vii
List of Figures ................................................................................................................. viii
List of Tables .................................................................................................................... ix
Chapter 1. Introduction .......................................................................................................1
Bisphenol A .............................................................................................................2
Microplastic Particles and Mussels .........................................................................4
Chapter 2. Capillary Electrophoresis ...................................................................................6
Factors Influencing CE Analysis ...........................................................................10
Chapter 3. Materials and Methods .....................................................................................14
Experimental ..........................................................................................................15
Reagents ..............................................................................................................15
Instrumentation ...................................................................................................15
Solution and Buffer Preparation ........................................................................16
Capillary Electrophoresis Method ......................................................................17
Sample Collection and Preparation ........................................................................18

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Water Samples ....................................................................................................18
Mussel Samples ..................................................................................................19
Chapter 4. Results and Discussion ....................................................................................20
BPA Standards .......................................................................................................21
Samples ..................................................................................................................22
Water Analysis ....................................................................................................22
Mussel Sample Analysis .....................................................................................23
Chapter 5. Conclusions and Future Work .........................................................................27
Conclusions ............................................................................................................28
Future Work ...........................................................................................................29
References ..........................................................................................................................30
Appendix ............................................................................................................................33
Mussel Sample Data ..............................................................................................33
Sample Electropherograms ....................................................................................34

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ACKNOWLEDGEMENTS
Unending gratitude to my supervisor, Dr. Kingsley K. Donkor. With his wisdom and
compassion, he has provided invaluable guidance and support in my research and in my life.
He has been there for me when the lab results were discouraging and when life got in the way
of going forward. I have learned by example how to be a better chemist, scholar, teacher, and
person.
I wish to acknowledge the amazing support I have received from so many at Thompson Rivers
University. The chemistry department, my committee members and the MSc committee and
coordinators went above and beyond to help me make this a reality. My instructors have taught
me so much both in the classroom and out, including much appreciated concern and advice
offered during chance encounters in the community.
My family and friends are everything to me in all I do. Support for my further education has
been especially strong from my parents, my children Dennis, Thomas, and Rebecca (even
though you are mostly adults) and Jacqueline. You keep me grounded in my sense of self and
my place in the world. Never underestimate my appreciation for when you asked how I was
doing, or you made me a meal or a coffee. Thank you.
I gratefully acknowledge funding that supported me throughout my studies which include the
Natural Sciences and Engineering Research Council (NSERC) of Canada, TRU Graduate
Teaching Assistant program, and the Donkor Research Group. I also gratefully acknowledge
Canada Foundation of Innovation (CFI) for funding for the purchase of the capillary
electrophoresis instrument. Thanks to Dr. John Church and TRU ALNA group for the support
in the early part of my program. The work of Mae Frank and Dr. Louis Gosselin was
fundamentally important to my research; thank you both.

vi

To my Dad and to David with love
For where I started and what the future holds

vii

LIST OF ABBREVIATIONS

AU

Absorbance Unit

BGE

Background Electrolyte

BPA

Bisphenol A

CE

Capillary Electrophoresis

CMC

Critical Micelle Concentration

EFSA

European Food Safety Authority

EOF

Electroosmotic flow

FDA

Food and Drug Administration

M

Molarity (moles/litre)

MEKC

Micellar Electrokinetic Capillary Chromatography

MP

Microplastic Particles

NSERC

Natural Sciences and Engineering Research Council of Canada

PC

Polycarbonate

R2

linear correlation coefficient

SDS

Sodium Dodecyl Sulphate

TDI

Tolerable Daily Intake

UV

ultraviolet

viii

LIST OF FIGURES

Figure 1.1. Chemical structure of bisphenol A ....................................................................2
Figure 2.1. Double layer formed within capillary................................................................7
Figure 2.2. A diagram depicting EOF ..................................................................................8
Figure 2.3. Schematic diagram of capillary electrophoresis ................................................9
Figure 2.4. Ohm’s law plot ................................................................................................12
Figure 3.1. Picture of capillary electrophoresis system used .............................................16
Figure 4.1. Calibration curve for BPA concentrations from 5 ppm to 50 ppm ................21
Figure 4.2. The electropherogram of the 10 ppm BPA standard .......................................22
Figure 4.3. Electropherogram for CE analysis of water sample ........................................23
Figure 4.4. Electropherogram of mussel sample #109 ......................................................24
Figure 4.5. CE electropherogram of mussel sample #116 .................................................25
Figure 4.6. CE electropherogram of mussel sample #116, basic pH .................................26
Figure B.1.Water sample electropherograms .....................................................................34
Figure B.2. Expanded view of Figure B.1. electropherograms .........................................34
Figure B.3. Mussel sample electropherograms ..................................................................35
Figure B.4. Mussel sample electropherograms ..................................................................35

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LIST OF TABLES

Table 3.1. Preparation of BPA standards for calibration. ..................................................17
Table 3.2. Optimized CE instrumentation parameters for BPA analysis ..........................18
Table 3.3. Sample sites around Vancouver Island, BC......................................................19
Table 4.1. Potential BPA concentration in mussel samples...............................................26
Table A.1 Mussel sample locations and microplastic particle counts ...............................33

1

CHAPTER 1

INTRODUCTION

2

CHAPTER 1. INTRODUCTION

Bisphenol A
Bisphenol A (BPA) is ubiquitous in the modern world and has been shown to have negative
effects on human health. This small organic monomer (Figure 1.1) was first described and
synthesized in 1891 by a Russian chemist (Jalal et al., 2018; Xiao et al, 2020). It was
investigated and later abandoned as a possible synthetic estrogen replacement in the 1930s due
to its hormone effects; it is classified as an endocrine disruptor (Rubin, 2011). Since the 1940s,
it has been used in manufacturing plastics and today it is produced in huge amounts in making
of polycarbonate (PC) plastics and epoxy resins; these account for roughly 95% of BPA usage
(Rubin, 2011). Other products that use BPA include thermal paper, water-pipes, electronics,
toys, and food packaging. Yearly worldwide production of BPA is over 3.8 million tons
(Michałowicz, 2014). Use in Canada is reported from 100 000 to 1 million kg annually
(Environment and Climate Change Canada, 2018).
BPA is not found naturally; the only source is from manufacturing, and it is now detectable
almost everywhere. It is seen in the atmosphere, dust, water, and in human tissues, organs, and
fluids (Michałowicz, 2014). BPA concentrations have been measured in the atmosphere across
the entire globe over cities, rural areas, oceans, and polar regions (Fu & Kawamura, 2010).
CH3
OH

HO
CH3

Figure 1.1. The chemical structure of bisphenol A, (2,2-bis(4-hydroxyphenyl)propane, CAS
registry number 80-05-7).

Primary concern about the safety of BPA stems from its known disruption of human hormones.
There is a large body of research into the effects on human health and the environment. Human
health effects include disruption of DNA strands and signaling pathways, dysregulation of
calcium ion homeostasis of cells (Jalal et al., 2018), social impairments of 4 year old children

3

with measured pre-and post-natal BPA exposure (Lim et al., 2017), and promotion of ovarian
cancer cell development (Shi et al., 2017). The risks of obesity, diabetes, liver cell damage,
immunotoxicity, and coronary heart diseases may also increase with BPA exposure
(Michałowicz, 2014). Animal studies show changes in neonatal brain development,
development of reproductive organs in males and females, disruptions in sperm production and
prostate health (Talsness et al., 2009). Susceptibility to drug addiction has also been linked to
BPA exposure (Jones & Miller, 2008).
BPA is found in most people tested (Rubin, 2011). Oral ingestion is the primary source of
exposure through food and drink, primarily due to the linings of containers for foods and drinks
made using BPA and epoxy resins (Rubin, 2011). BPA levels are higher in women than men
and higher in younger age brackets than older. Levels have been found as high as 1-104 ng/g
of tissue in placenta and fetus (Jalal et al., 2018). Maximum daily exposure has been estimated
at about 1 µg/kg BW/day but negative effects from BPA may be experienced at daily exposure
as low as 0.025 µg/kg BW/day (Kang et al., 2006).
Due to its ubiquitous presence in the environment, there have also been studies into its effects
there. Aquatic species studied by Alexander et al in 1988 showed toxic effects from BPA at
levels between 1 and 10 ppm of water and a study by Mihaich et al (2009) found no-effect
concentrations (NOECs) as low as 0.025 ppm. Xiao et al (2020) have recently written a review
in the Journal of Hazardous Materials of scores of articles studying how BPA effects growth
and development of plants and some research into using plants for bioremediation of BPA
pollution. BPA does break down but there is always more entering the environment, so it
appears as a persistent contaminant.
Evidence that there is widespread concern about the effects of BPA on human health and on
the environment can be seen in the existence of government regulations and guidelines. In
Canada, there are guidelines and one regulation regarding BPA. The Government of Canada
through Environment and Climate Change Canada has set a Federal Environmental Quality
Guideline (FEQG) as a measure of acceptable levels of BPA in environmental waters at
3.5 μg/L (Environment and Climate Change Canada, 2018). FEQGs are not regulatory or
binding; they are merely set to provide a guide for what is deemed to be safe. No maximum
acceptable concentration (MAC) has been set in the Guidelines for Canadian Drinking Water

4

Quality as BPA has not been found in drinking water in levels that are considered “of possible
human health significance” (Health Canada, 2020). The first regulation concerning BPA was
set in Canada in the Canada Consumer Product Safety Act of 2010 which includes a prohibition
against the sale of PC baby bottles made with bisphenol A (Government of Canada, 2010).
Infants up to six months old fed formula from polycarbonate bottles were seen to have the
highest BPA dietary exposure per kilogram of body weight (Srivastava et al., 2015) so there
was sufficient concern to mandate reduced sources of BPA. The United States Environmental
Protection Agency (US EPA) established a chronic oral Reference Dose (RfD) and the
European Food Safety Authority (EFSA) set a Tolerable Daily Intake (TDI) of BPA. They are
both at 50 μg/kg∙bw/day (Shelnutt et al., 2013). These measurements are both estimates of safe
daily exposure over a lifetime. In 2015 the EFSA reduced that to 4 μg/kg∙bw/day because of
concerns that the data did not conclusively show that the higher limit was a reasonable level
of exposure; in September 2018, the EFSA put out a “Call for data relevant to the hazard
assessment of Bisphenol A (BPA)” (European Food Safety Authority, 2018) to better assess
the effects of BPA and set a TDI based on current research. There have been no changes to the
guidelines reflecting new information from this research.

Microplastic Particles and Mussels
Microplastics are small, less than 5 mm, bits of plastic (Anderson et al., 2016) that are found
in water everywhere including the ocean and other water systems (Frank, 2018). There are an
estimated 15 to 51 trillion particles, up to 236 thousand metric tons, of microplastics in the
ocean (Van Sebille et al., 2015). Some microplastic particles are intentionally manufactured
but much of the pollution arises from larger plastics breaking down into tiny pieces through
use, such as plastic fibers breaking off fabrics being laundered (Frank et al., 2016) or
degradations in the environment (Anderson et al., 2016; Desforges et al., 2014). Microplastics
found in the ocean are composed of many types of plastic including ones made with BPA
(Tang, Rong, et al., 2020) and microplastics may even be absorbing BPA from the water (Tang,
Zhou, et al., 2020).

5

One of the concerns is that these particles are about the size of food particles for phytoplankton
and other marine creatures, especially filter-feeders (Desforges et al., 2014). Filter-feeding
shellfish are marine animal that includes mussels, clams, and oysters. They ingest food, mainly
phytoplankton, by siphoning in water which contains phytoplankton but can also have varying
levels of pollutants, including microplastics and chemicals that adsorb to microplastics such as
BPA, (Davidson & Dudas, 2016; Desforges et al., 2014; Frank et al., 2016). Thus,
microplastics are ingested as particles and as contaminants in the phytoplankton (Van
Cauwenberghe & Janssen, 2014). Many of these filter-feeders are shellfish species which are
economically significant. Farmed and wild shellfish production in Canada was over 3 billion
dollars in 2018 (Canada, 2019). Understanding and protecting shellfish is critical to the
economy and the environment.
The risks to marine life that ingest microplastics are both physical and chemical. Physical
hazards include obstructions, deformations, malnutrition, and others (Tang, Rong, et al., 2020).
Chemically, there is some research that suggests increased toxic effects from other pollutants
in the presence of microplastics (Tang, Rong, et al., 2020) and there is speculation that the
presence and possible release of BPA from these plastics could have deleterious effects beyond
the physical trauma (Frank et al., 2016; Tang, Zhou, et al., 2020).
Davidson and Dundas (2016) found plastic microparticles in all samples of wild and cultured
Manila Clams (Vererupis philippinarum) collected from Baynes Sound on the east coast of
Vancouver Island, British Columbia. Research has also confirmed the presence of microplastic
particles in the majority, 28 of 36, of wild Mytilus trossulus samples collected various locations
between the mainland and the west coast of Vancouver Island, British Columbia (Frank et al.,
2016). In Frank’s work, there were zero to six particles counted in each specimen with an
average of 2 microplastic particles in each of the mussels examined.

6

CHAPTER 2

CAPILLARY ELECTROPHORESIS

7

CHAPTER 2. CAPILLARY ELECTROPHORESIS
Capillary electrophoresis (CE) is a relatively new analytical separation technique that is based
on a solute’s ability to move through a solution, usually an aqueous buffer, under the influence
of an electric field (Harris, 2007). In CE, the separations take place within a tubular capillary
with internal diameter ranging from 25 to100 µm. Before the capillary can be used for analysis,
the inner walls of the capillary need to be electrically charged. This is accomplished by flushing
the capillary with sodium hydroxide (NaOH) solution followed by a buffer solution. On
passing the NaOH solution, the silanol (Si-OH) groups lining the inner walls of the capillary
are ionized to negatively charged silanoate (Si-O-) groups. The cations within the buffer get
attracted towards the negatively charged silanoate groups. This results in the formation of a
double layer composed of a fixed layer and a mobile layer (Figure 2.1).

Figure 2.1. Double layer formed within capillary (Baker, 1995)

The fixed layer is composed of cations held tightly to the silanoate groups. The mobile layer
is composed of cations further away from the silanoate groups, and thus can slowly migrate
towards the cathode if a voltage is applied. Solvation of anions to cations forming the mobile
layer causes the bulk buffer to flow towards the cathode. The flow of the bulk buffer towards
the cathode is called the electroosmotic flow (EOF) (Baker, 1995).
Once the capillary is electrically charged, the sample containing the analyte can be injected
into the buffer filling the capillary. As the electric current is applied, the buffer moves towards

8

the cathode and the sample containing the analyte will move within the buffer and be separated
based on differing electrophoretic mobilities, which are related to size-to-charge ratios. Once
the analyte reaches the UV detector, it is detected and processed by a computer in the form of
an electropherogram (Figure 3). The electropherogram shows peaks, representing different
analytes, at different migration times. The area under the peak is proportional to the amount of
analyte and the migration time is unique to each compound.
Capillary electrophoresis (CE) is an analytical method that differentiates and separates charged
species on the basis of mobility under the influence of an electric field gradient (Chu et al.,
1995) or electrophoretic mobility. The analytes are separated and analysed based on size and
charge. Smaller ions travel more easily through the buffer resulting in faster migration within
the buffer toward the respective electrode. Increased charge on the ion increases attraction and
thus the flow toward the respective electrode. Therefore, small positive ions are detected before
small negatively charged ions. Neutral species migrate at the rate of the EOF. This process can
be seen below in Figure 2.2.

Figure 2.2. A diagram depicting EOF including the separation of ions based on size and charge
between the anode and cathode (Laboratoire Suisse d’Analyse du Dopage, 2008) .

A basic CE system is shown in Figure 2.3. It consists of several main components namely a
controllable high voltage power supply, inlet and outlet buffer vials, a capillary with optical
viewing window, a detector, and a data display device like a computer. Most CE units also are
equipped with a cooling ability to control or dissipate heat inside the capillary. The power
supply can be set at a constant voltage up to 30 kV or a constant current up to 300 µA as well

9

as a constant power up to 6 W. The polarity can be also reversed. Because the migration time
varies if the voltage changes, it is very common to operate at constant voltage. The most
important variable in CE is the composition of the buffer, which influences the EOF. Any small
change in pH or concentration of buffer can affect the migration time of solutes. Capillaries in
CE system usually have very small inner diameters, around 25-100 μm. It is very common to
use a fused silica capillary instead of the glass ones because they are still transparent at shorter
wavelengths than 280 nm. An external coating of polyimide covers the capillary, making them
stronger, more flexible, durable, and not easily broken.

Capillary

Capillary

Inlet

Outlet

Figure 2.3. Schematic diagram of capillary electrophoresis (Donkor, 2021).

UV detection or PDA detection can be used for direct or indirect detection of an analyte. Direct
detection is used for compounds that can absorb UV light strongly, resulting in high positive
peaks in the electropherogram without any software manipulation. For solutes that do not
possess strong UV absorbance, indirect UV detection can be used. In this approach, the buffers
would contain UV absorbing compounds, i.e., chromophoric ions, that serve as visualization
agents to make the background electrolyte absorbent with indirect UV detection. In the
electropherogram, the absorbance of buffer with those UV absorbing compounds will appear

10

as the baseline with a high background signal. When the non-UV absorbing solutes pass
through the detector, their peaks would be a negative dip compared with the baseline. These
negative peaks will be flipped to become positive peaks in the electropherogram by a software
that reverses detector’s output polarity (Baker, 1995).
Micellar Electrokinetic Capillary Chromatography (MEKC) is a mode of capillary
electrophoresis that allows for interactions with micelles and the separation of electrically
neutral compounds becomes possible (Harris, 2007). A surfactant is added to the buffer above
the critical micelle concentration (CMC), the threshold where micelles can form. These
micelles form a pseudostationary phase. Very hydrophobic molecules spend all their time in
the micelles and migrate at the same rate as the micelles. Molecules that are partially soluble
in the micelles will partition between the buffer and the micelles at a rate proportional to their
hydrophobicity. Hydrophilic, water-soluble molecules stay in the aqueous buffer and do not
interact with the micelles. The interactions with the pseudostationary phase are rapid compared
to interactions with a stationary phase and do not increase analysis time because the EOF
dominates the migration rate. Sodium dodecyl sulphate forms anionic micelles above the CMC
of 8.2 mM; cationic, non-ionic, and zwitterionic surfactants form corresponding micelles.
Anionic micelles migrate within the buffer toward the anode but the EOF is stronger and,
ultimately, the micelles flow toward the cathode.
Factors Influencing CE Analysis
(i) Capillary Inner Diameter
The capillary diameter influences the convective diffusion and it is important to minimize it in
analysis. The smaller the capillary radius is, the less convective diffusion is generated due to
the smaller temperature difference. This leads to narrower zones, and better separation of the
zones. The temperature difference, ∆T, between the center and the wall of the capillary, is
given by the equation below (Baker, 1995):
0.239Q 2
)r
4k
Q: power density (Watt/m3)
k: thermal conductivity
r: capillary radius
∆T = (

11

A reduced capillary diameter offers higher electrical resistance and less current for the applied
voltage, resulting in less Joule heat. In addition, a small capillary dissipates heat faster because
of a larger inner surface area-to-volume ratio. In a smaller capillary, the solutes tend to move
to the detector as single zones. In a larger capillary, the solutes around the warm center will
move through the tube with different speeds compared to the ones at the cooler outer wall,
leading to two distinct zones for the same solute. Therefore, for CE analysis, it is desirable to
use a capillary with the smallest diameter possible, as it can provide good separation as well
as reduce Joule heat and increase heat dissipation (Baker, 1995).
(ii) Chemical Modification of the Capillary Wall (Coating)
The capillary wall can be altered by coating or chemical bonding to reduce or eliminate the
EOF. These modifications reduce the zeta potential by shielding the surface charges on the
inner wall, thereby resulting in a reduction or elimination of EOF. The modifications also
increase the viscous drag on the buffer at the wall, thus reducing the rate of EOF. A coating
can also minimize the adsorption of solute to the capillary wall and leading to better results in
some cases. By chemically treating the capillary, a reasonable detection time is obtained., but
adequate EOF is still needed and the level of EOF required depends on the analytes (Baker,
1995).
(iii) Voltage
High voltages of up to 30 kV can be used because the capillary has a small diameter and can
dissipate heat quickly. The EOF is proportional to the electrical field which depends on the
applied voltage. The higher the voltage applied, the shorter the migration time of samples
because of the increase in EOF. High voltage also provides faster separation and better
efficiencies. Theoretically, it is better to use the highest voltage because of faster analysis time
and narrow peaks. Nevertheless, Joule heat will be increased by the higher voltage, a decrease
of the buffer’s viscosity or an increase of flow rate. The Joule heat may cause broader peaks,
unstable migration times, solution decomposition or denaturation, or buffer boiling which is a
cause of electrical discontinuity. Therefore, the applied voltage should be reasonable so that
the heat can dissipate well in capillary. The maximum optimized voltage can be investigated
and determined based on a plot of Ohm’s law, which is a plot of the observed currents versus

12

the applied voltages as shown in Figure 2.4. The point at which nonlinearity starts represents
the maximum voltage that can be applied to prevent Joule heating (Baker, 1995).

Figure 2.4. Ohm’s law plot (Baker, 1995).

(iv) Source and Destination Vials Buffers
Source and destination vials are filled with the same buffers with even levels to prevent
changes in migration time or laminar flow due to the siphoning of the buffer. Even with that,
repeated analysis might also change the concentration and pH of the buffer because of the
electrolysis of water, where hydrogen ions are formed at the cathode and the hydroxyl ions at
the anode. Moreover, after repeated analysis, the buffers in outlet vial may have a different
composition because solute ions in the capillary might elute and accumulate into the outlet
buffer vial, which can change the electrical field and lead to non-reproducible migration times.
Therefore, buffer replenishment is very important by rinsing and refilling the vials and
capillary to achieve good reproducibility. Especially in indirect absorbance detection, any
change in the chromophore’s concentration or composition might lead to a drifting baseline
and more noise (Baker, 1995).
There are some criteria for selecting the buffer or chromophoric ion in CE analysis. Firstly, the
mobility of a chromophoric ion or any ions in the buffers should be similar to the solute’s
mobility to prevent asymmetrical peaks. Secondly, in indirect absorbance detection, the

13

chromophore should have a high molar absorptivity at the specific wavelength so that when
non-absorbing solutes pass through the detector, there will be a large decrease, indicated by a
negative dip peak. In other words, the chromophoric ions should absorb light strongly at the
wavelength that the solutes cannot absorb. Lastly, the buffer or chromophoric ion must be
stable and nonreactive with the capillary or any components in the samples (Baker, 1995).
(v) pH of the Buffer
Buffer pH can change the zeta potential, thus affecting strongly the EOF. At higher pH, the
inner wall of the capillary will be highly negatively-charged because of the high dissociation
of Si-OH to Si-O-. The surface’s high charge is proportional to the zeta potential, resulting in
increased EOF or electroosmotic velocity. In contrast, at low pH (< 2), the EOF is eliminated
because most of the silanol groups cannot be deprotonated. In addition, buffer pH will also
affect the electrophoretic mobility of the solutes. Depending on the analysis, the required buffer
pH can be selected by providing the best separation or optimum EOF flow (Baker, 1995).
(vi) Buffer Concentration
For a stable capillary temperature, a high buffer concentration will decrease the zeta potential
and result in lower EOF. Analysis time will be shorter with lower concentration of buffers, but
unreasonably low concentration may lead to broader and asymmetric peaks as well as cause
adsorption of solutes. Another noticeable distortion of electrical field can occur when the
buffer concentration is not higher than the solute concentration. This results in broad, skewed
peaks. As a general rule, the run buffer concentrations are in the range of 10-100 mM (Baker,
1995).
(vi) Temperature
Temperature is one of the important factors in CE analysis. Unstable temperature can lead to
various migration times, zone spreading, and sample decomposition. The EOF rate also
increases when the temperature elevates. It is always advisable to control the temperature.
There are many different cooling systems to help the heat dissipate quickly and keep the
temperature stable, either by air or liquid cooling. The most common method is liquid cooling
in which a coolant is circulated through a jacket placed around the capillary (Baker, 1995).

14

CHAPTER 3

MATERIALS AND METHODS

15

CHAPTER 3. MATERIALS AND METHODS

Experimental
Reagents
Bisphenol A, 97+%, was purchased from Alfa Aesar, Ward Hill, MA, USA. Sodium
tetraborate decahydrate, Na2B4O7∙10H2O, and sodium dodecyl sulfate (SDS) were purchased
from Sigma-Aldrich Chemical Co, St Louis, MO, USA. Sodium dodecyl sulfate (SDS) was
obtained from BDH Biochemicals, Toronto, Canada. Sodium hydroxide was purchased from
EMD Chemicals, Gibbstown, New Jersey, USA.
Instrumentation
The Beckman Coulter P/ACE MDQ capillary electrophoresis system (Beckman Coulter, Brea,
CA, USA) equipped with ultraviolet detector and interfaced with the 32-Karat software for
data acquisition is shown in Figure 3.1. An uncoated fused-silica capillary with 50 µm internal
diameter and total length of 60 cm was used. The separation is done in fused silica capillary
from Polymicro Technologies, AZ, USA. The 25 mm Nylon® 0.45 µm syringe filter was
purchased from Canadian Life Science, Ontario, Canada. The pH meter used was Mettler
Toledo FE20- FiveEasyTM from Grelfensee Switzerland, purchased from USA.

16

Figure 3.1. Picture of capillary electrophoresis system used in the Donkor research laboratory.

Solution and Buffer Preparation
All water used for standards and buffer preparation and capillary rinsing was 18 MΩ prepared
by the TRU Chemistry department. Glassware was all cleaned with manual detergent in hot
water, rinsed with tap water three times, deionized water three times, followed by three rinses
with 18 MΩ water. They were dried either in an oven at 100°C or at ambient temperature.
The buffer used for analysis was 60 mM borate, 20 mM SDS prepared in 18 MΩ water and
pH adjusted to 9.5 with 1.0 M NaOH. Buffer solutions were prepared in a volumetric flask
using 18 MΩ water and sonicated when necessary to fully dissolve the solids. The pH of the
buffer solutions was adjusted with 0.1 M NaOH. All stock solutions were filtered when
prepared and before injection into the CE instrument using 0.45 μm Nylon syringe filters.
Sodium hydroxide solutions (0.1 M and 1.0 M) were prepared in a volumetric flask using
18 MΩ water.

17

Standards were prepared in 10 % methanol to allow the BPA to fully dissolve and keep the
organic solvent to a minimum to avoid solvent effects in the analysis. A 50 mM BPA stock
solution was prepared in a 50 mL volumetric flask by adding 5.00 mL methanol by volumetric
pipette. When the BPA was fully dissolved, the flask was topped up with 18 MΩ water,
thoroughly mixed and then filtered with a 0.45 μm Nylon syringe filter. Dilutions were made
directly in 2 mL CE vials by using the 50 mM standard and 18 MΩ water using micropipettes
of the appropriate volume range. No precipitation of BPA was observed upon dilution with the
water. Table 3.1 shows the volumes of BPA and 18 MΩ water used to prepare the standards.
Table 3.1. Preparation of BPA standards for calibration.
Standard Concentration

Volume of 50 ppm BPA

Volume of 18 MΩ water

50 ppm

2 mL

0 mL

40 ppm

1.6 mL

0.4 mL

30 ppm

1.2 mL

0.8 mL

20 ppm

0.8 mL

1.3 mL

10 ppm

0.4 mL

1.6 mL

5 ppm

0.2 mL

1.8 mL

Capillary Electrophoresis Method
All analysis was performed using a Beckman P/ACETM MDQ capillary electrophoresis
instrument (Beckman Coulter, Inc., Fullerton, CA) using an ultraviolet (UV) detector. The
capillary used was uncoated fused-silica with 50 μm inner diameter, 365 μm outer diameter,
60 cm total length with 50 cm to the detector (Polymicro Technologies, Phoenix, AZ).
Galden®

HT-110

perfluoropolyether

coolant

(Ideal

Vacuum

Products

LLC,

Albuquerque, NM) was used in the capillary cartridge to keep the buffer and samples at 25 °C
during migration through the capillary. Nylon filters, 0.45 μm, from Canadian Life Science,
Peterborough ON, were used to filter all solutions prior to injection. The temperature was held
constant at 25 °C. Direct UV detection wavelength was set at 214 nm.

18

New capillaries were conditioned at 20 psi for 10 min with water, 60 min with 1M NaOH,
30 min with 0.1 M NaOH followed by 20 min with water again. Each day the capillary was
initially rinsed with 0.1 M NaOH for 15 min and then with the buffer that would be used in the
analysis for another 15 min. This daily rinse was also at 20 psi. Capillaries were always stored
with both ends immersed in 18 MΩ water when not in use.
Bisphenol A analysis was ultimately conducted with the method shown in Table 3.2.
Table 3.2. Optimized CE instrumentation parameters for BPA analysis

Rinse Pressure

20 psi

5 min

0.1 M NaOH

Rinse Pressure

20 psi

3 min

18 MΩ water

Rinse Pressure

20 psi

8 min

buffer

Injection

0.5 psi

5s

Separate voltage 20 kV
Pressure

20 min

0.17 min ramp

normal polarity

Sample Collection and Preparation
Water Samples
Water for analysis was collected from the Thompson River at Riverside Park and from the
Kamloops city water supply. Tap water was from the north end of the city, Rayleigh, the south
end, Aberdeen, and central to Kamloops at Thompson Rivers University. At TRU, samples
were collected from water taps in the Ken Lepin Science building and from the oldest campus
building, Old Main, in a staff kitchen and two washrooms.
Water was collected from taps after the water was allowed to run for a full one minute. All
water collection followed the same procedure; amber glass vials were rinsed three times with
the sample, filled to the shoulder of the sample bottle and sealed with a screw top. The only
exceptions were the domestic tap water provided from Rayleigh in a clear glass, nylon
stoppered sample vial. Samples were left unopened at room temperature until they were filtered
immediately before analysis.

19

Mussel Samples
Mussel samples, Mytilus trossulus, were collected from 12 sites around the southern half of
Vancouver Island (Table 3), all of which were accessible by vehicle. All mussels were between
3 and 4 cm and their tissues were digested with nitric acid to remove organic matter while
leaving non-tissue materials, such as plastics. Microplastic particles were documented and the
samples were stored at 4°C until they were pH adjusted with 1M NaOH for analysis by CE.
(Frank, 2018).
Table 3.3. Sample sites around Vancouver Island, BC. (Frank, 2018).
Site

Community

Latitude/ Longitude

Coast

Environment

Human
population

1

Tofino

49°09.161’N, 125°54.196’W

West

Dock

1,932

2

Bamfield

48°50.081’N, 125°08.144’W

West

Dock

155

3

Port Renfrew

48°33.338’N, 124°24.818’W

West

Dock

144

4

Jordan River

48°25.254’N, 124°03.299’W

West

Intertidal

100

5

Sooke

48°22.978’N, 123°42.344’W

South

Intertidal

367,770

6

Colwood

48°20.030’N, 123°27 .230’W

South

Intertidal

367,770

7

Oak Bay

48°27.065’N, 123°17.856’W

South

Dock

367,770

8

Duncan

48°47.795’N, 123°36.145’W

East

Dock

44,451

9

Departure
Bay

48°11.396’N, 123°56.983’W

East

Dock

104,936

10

Parksville

49°20.899’N, 124°21.516’W

East

Dock

28,922

11

Comox

49°40.319’N, 124°55.699’W

East

Dock

54,157

12

Campbell
River

50°01.952’N, 125°14.650’W

East

Dock

37,861

20

CHAPTER 4

RESULTS AND DISCUSSION

21

CHAPTER 4. RESULTS AND DISCUSSION
BPA Standards
The capillary electrophoresis analysis of water and mussel samples for the presence of
bisphenol A (BPA) was undertaken. A CE method was developed and optimized until it was
able to detect BPA concentrations in standard solutions between 5 ppm and 50 ppm.
Following the optimization, a calibration curve was developed to quantify any BPA that was
detected in the samples. The calibration curve obtained is shown in Figure 4.1. The calibration
curve showed reasonable linearity with a R2 of 0.9867.

Figure 4.1. The calibration curve for BPA concentrations from 5 ppm to 50 ppm developed
with the method used in this study.

The BPA identifying peak in standard solutions from 5 ppm to 50 ppm using ultraviolet direct
detection appeared in the electropherograms between 13.5 and 14 min. Below, in Figure 4.2,
is an electropherogram of the 10 ppm BPA standard.

22

Figure 4.2. The electropherogram of the 10 ppm BPA standard as a sample of the data used to
generate the calibration curve shown in Figure 7.

Samples
Water Analysis
Water samples from municipal water supply from several sources around Kamloops were
analysed. No BPA was detected in these samples. A representative electropherogram shows
no peak in the BPA region. An example of the electropherogram of water sample taken from
TRU Old Main Staff lunchroom kitchen cold water tap is shown in Figure 4.3. As it can be
seen no significant peak was obtained in the region for the detection of BPA.
Electropherograms for other water samples displayed similar results. This observation
indicates that there was no measurable amount of BPA in any of the Kamloops municipal water
supply that was analysed or the amounts were below the detection limit of the developed
method.

23

Figure 4.3. Sample electropherogram for CE analysis of Kamloops municipal tap water taken
from the staff lunchroom in the Old Main building on the Thompson Rivers University campus
showing no BPA detected. The peak for BPA is expected at about 13 min. The insert shows an
expanded view from 12 min to 14 min.

Mussel Sample Analysis
The samples of mussel tissue tested were identified by area of collection and the number of
microplastic particles (MP) counted per mussel. There were up to three MPs in each of the
samples tested and up to 6 MPs across all of the samples available. On the chromatograms of
the mussel samples, the absorbance units (AU) scale is very much smaller than the other graphs
and no peaks appeared at the expected migration time for BPA. Small peaks in the
electropherograms for the mussel samples had inconsistent and shorter migration times. This
may be due to a change in the EOF of the analysis for the mussel samples. The samples were
provided in nitric acid with pH less than one. Different pHs of the mussel samples were also
investigated. One mussel sample was subjected to extreme acidic (pH < 1) and basic
conditions, and then analysed. Electropherograms are shown in Figures 4.4 to Figures 4.6. The

24

BPA was detected at approximately 7.5 min, 11 min for the acidic samples and 10 min for the
basic mussel sample. Adjusting the pH of the sample did not greatly affect the results. The
sample that was adjusted to a basic pH had a peak detected at 10 min, representing 1 min
shorter migration time than when the same sample was acidic. This is within the variation seen
for the detection of the standards and does not necessarily represent any significance.

Figure 4.4. Electropherogram of mussel sample #109 collected from Oak Bay, BC. The sample
was digested in nitric acid for microplastic particle analysis (2 MP count) and analysed by CE
without pH adjustment. The large peak at 7.3 min was labeled by the CE analysis software
with an area of 7472.

25

Figure 4.5. CE electropherogram of mussel sample #116 (3 MP count) collected from Duncan,
BC, acidified for microplastic particle analysis and provided for BPA analysis. The pH of the
sample was less than one and left unadjusted for CE analysis. The area of the peak at
approximately 11 min has an area of 2617.

26

Figure 4.6. CE electropherogram of mussel sample #116 (3 MP count) collected from Duncan,
BC, acidified for microplastic particle analysis and provided for BPA analysis. The pH of the
sample was adjusted with NaOH and tested to be basic. The area of the peak at approximately
10 min was found to be 6957.

In the mussel analysis, the peaks of interest which may indicate the presence of BPA can be
interpreted to indicate low levels of BPA. Table 4.1 shows calculated concentration from the
peak areas.

Table 4.1 Potential BPA concentration in mussel samples.
Figure
4.4
4.5
4.6

Area of CE peak
7472
2617
6957

BPA concentration (ppm)
8.5
3.0
7.9

27

CHAPTER 5

CONCLUSIONS AND FUTURE WORK

28

CHAPTER 5. CONCLUSIONS AND FUTURE WORK
Conclusions
Bisphenol A (BPA) was not detected in any of the water samples tested by the capillary
electrophoresis (CE) method developed and used in this study. The lowest concentration
standard used was 5 ppm. With safe levels of BPA exposure set between 4 μg/kg∙bw/day by
the EFSA and 50 μg/kg∙bw/day by the US EPA (Shelnutt et al., 2013), a BPA detection at
5 ppm is not sensitive enough to determine if the BPA levels are of concern unless the levels
were extremely high. Walpole et al (2012) estimated the average adult body mass at 62 kg
across the world and 80.7 kg in North America; therefore, a total exposure for an adult could
be considered safe between (62x4=248 μg/day and 4035 μg/day ) 0.248 and 4.035 mg/day. If
BPA were detected at 5 ppm in drinking water, as little as 0.05 to 0.8 L per day could result in
an unsafe level of exposure. Considering all the other sources of BPA exposure, a level that
could be detected and quantified with the current method would be very alarming.
The analysis of mussel tissue had more interesting results that were quantifiable by
interpolation using the equation of the calibration line. Assuming the prominent peaks
represent BPA, the concentrations ranged from 3.0 ppm to 8.5 ppm in the samples. There is no
apparent correlation between BPA concentration and the number of microplastic particles; the
sample with the larger concentration had two microplastic particles counted while the smaller
area peak, was in a sample with 3 MPs.
The peaks for the mussel samples had shorter and varied migration times which means they
cannot be definitively identified as being a response to the presence of BPA. In fact, they were
initially assumed to not represent detection of BPA and it is only upon further reflection that
they are possibly indications of BPA in the samples. Without confirmation that the peaks
actually represent detection of BPA, no definitive conclusions can be made about the presence
of BPA in the mussels. The presence of microplastic particles shows there is plastic
contamination of the marine environment around Vancouver Island but the present research
has not verified BPA as a component of the problem. Given the ubiquitous nature of BPA in
the environment, it would be surprising if these filter feeders did not have some BPA in their
tissues.

29

Future Work
Using the current method to further analyse the mussel samples could provide interesting
information. Spiking the samples with known concentrations of BPA could reveal if the peaks
are due to the presence of BPA. Using the method of standard addition with the mussel samples
would be valuable future work given the complex matrix of these samples.
A first approach to detecting lower concentrations of BPA would be using the existing
parameters to test lower concentration BPA standards. This might allow development of a
lower concentration calibration curve and lower the level of detection. This could allow the
smaller area peaks in the mussel samples to be quantified.
One of the limitations for UV detection in CE is the narrow capillary diameter provides a short
pathlength for the UV detection and thus limits its sensitivity. There are various other CE
techniques that could also be investigated such as large volume sample stacking to enhance the
sensitivity.
Using CE to detect analogues of BPA is a natural next step in this line of research. As the
health and environmental hazards have been documented, BPA has been replaced with similar
compounds such as bisphenol S. It will be important to know where these compounds are and
what concentrations are there.

30

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33

APPENDIX
Appendix A: Mussel samples
Table A.1. Mussel samples by location and microplastic particle (MP) count
Sample number

Collection site

MP count

3
7
8
12
15
18
23
24
30
31
34
36
44
45
46
52
57
58
64
65
66
75
76
77
84
86
89
91
94
99
107
108
109
115
116
117

Bamfield
Bamfield
Bamfield
Departure Bay
Departure Bay
Departure Bay
Parksville
Parksville
Parksville
Jordan River
Jordan River
Jordan River
Campbell River
Campbell River
Campbell River
Port Renfrew
Port Renfrew
Port Renfrew
Tofino
Tofino
Tofino
Comox
Comox
Comox
Sooke
Sooke
Sooke
Colwood
Colwood
Colwood
Oak Bay
Oak Bay
Oak Bay
Duncan
Duncan
Duncan

0
5
3
3
6
1
0
3
2
1
0
1
1
1
0
2
4
3
0
0
1
3
2
2
3
3
2
0
1
5
1
0
2
2
3
4

*Analysed by CE

*
*
*
*

*

*
*

34

Appendix B: Sample Electropherograms
Figure B.1. Water samples from Rayleigh (black), Old Main, hot kitchen tap (grey), Old
Main cold kitchen tap (blue), Old Main 2nd floor ladies washroom (pink), and Old Main
gender neutral washroom automatic tap (green).

Figure B.2 Water sample electropherograms from Figure B.1 (see above) expanded to show
11.5 min to 15.5 min more closely.

35

Figure B.3. Mussel sample electropherograms for samples: #58 pH unadjusted <1 (grey); #58
pH adjusted >14 (blue); #64 pH unadjusted (pink); #64 pH adjusted >14 (dark green) and
#77 pH unadjusted (bright green). The pH of samples unadjusted were analysed as provided,
pH <1.

Figure B.4. Mussel sample electropherograms for samples: #109 pH unadjusted (black);
#116 pH unadjusted (grey); #116 pH adjusted >6 (blue); #52 pH 6 (pink); #46 pH 3 (dark
green); #46 pH 6 (bright green) and #46 pH unadjusted (purple). The pH of samples
unadjusted were analysed as provided, pH <1.