Trends and exposure of naturally produced brominated substances in Baltic biota

Trends and exposure of naturally produced
brominated substances in Baltic biota
- with focus on OH-PBDEs, MeO-PBDEs and PBDDs
Karin Löfstrand
Department of Materials and Environmental Chemistry
Stockholm University
Stockholm 2011
Doctoral Thesis 2011
Department of Materials and Environmental Chemistry
Stockholm University
SE-106 91 Stockholm
Sweden
Abstract
The semi-enclosed and brackish Baltic Sea has become heavily polluted by
nutrients, anthropogenic organic and inorganic chemicals via human activities.
Persistent organic pollutants (POPs) have been thoroughly investigated due to
their linkage to toxic effects observed in Baltic biota. There has been far less
focus on semi-persistent pollutants e.g. naturally produced oraganohalogen
compounds (NOCs) and their disturbances in the environment. This thesis is
aimed on assessment of levels and trends of naturally produced brominated
compounds in Baltic biota; more specifically on hydroxylated polybrominated
diphenyl ethers (OH-PBDEs), methoxylated PBDEs (MeO-PBDEs) and
polybrominated dibenzo-p-dioxins (PBDDs). These, NOCs, may originate from
production in algae and cyanobacteria. OH-PBDEs and MeO-PBDEs may also
be formed as metabolites of polybrominated diphenyl ethers (PBDEs), i.e. wellknown commercial flame retardants.
High levels of OH-PBDEs, MeO-PBDEs and PBDDs are shown within Baltic
biota (cyanobacteria, algae, mussels, fish), often in much higher concentrations
than PBDEs which are possible anthropogenic precursors of OH- and MeOPBDEs. The levels of OH-PBDEs, MeO-PBDEs and PBDDs are higher in the
Baltic Sea than on the west coast of Sweden. Temporal and seasonal variations
show fluctuations in concentrations of OH-PBDEs, MeO-PBDEs and PBDDs,
possibly related with macroalgal life-cycles. OH-PBDEs, MeO-PBDEs and
PBDDs are present in several filamentous macroalgae species, but considering
the levels quantified, the time of peak exposure and the species life-cycle the
macroalgae, Pilayella, Ceramium and Cladophora are suggested as major
natural producers of OH-PBDEs and PBDDs.
The high levels of OH-PBDEs, MeO-PBDEs and PBDDs in the Baltic Sea may
affect numerous organisms in the ecosystem. The toxic effects of OH-PBDEs
and PBDDs are of particular concern. This thesis stress the importance of
assessing and monitoring these substances, since the exposure to OH-PBDEs
and PBDDs, during summer, may cause acute effects in Baltic fish and wildlife.
© Karin Löfstrand
ISBN 978-91-7447-226-4
Universitetsservice US-AB, 2011
ii
Till min älskade familj
i väntans tider
iii
List of papers
This thesis is based on the following papers, which are referred to in the text by
their roman numerals, I-IV. Paper I, II and III are reproduced with the kind
permissions of the publishers. Some unpublished results are also included in the
thesis.
I
Löfstrand K., Malmvärn A., Haglund P., Bignert A., Bergman Å.,
Asplund L. (2010). Brominated phenols, anisoles and dioxins present in
blue mussels from the Swedish coast line. Environmental Science and
Pollution Research, 17, 1460-1468.
II
Haglund, P., Löfstrand K., Malmvärn A., Bignert A., Asplund L. (2010).
Temporal variations of polybrominated dibenzo-p-dioxin and
methoxylated diphenyl ether concentrations in fish revealing large
differences in exposure and metabolic stability. Environmental Science
and Technology, 44, 2466-2473.
III
Löfstrand K., Liu X., Lindqvist D., Jensen S., Asplund L. (2010).
Seasonal variations of hydroxylated and methoxylated brominated
diphenyl ethers in blue mussels from the Baltic Sea. Chemosphere
in press, doi10.1016/j.chemosphere.2011.01.001.
IV
Löfstrand K., Haglund P., Bergman Å., Kautsky L., Asplund L. (2011).
Hydroxylated and methoxylated polybrominated diphenyl ethers and
polybrominated-p-dioxins in macroalgae and blue mussels from the
Swedish coast line –patterns and correlations. Manuscript.
iv
Table of contents
Abstract .............................................................................................................. ii
List of papers..................................................................................................... iv
Table of contents................................................................................................ v
Abbreviations................................................................................................... vii
1. Introduction ................................................................................................... 9
1.1 Aim............................................................................................................ 9
2. Background.................................................................................................. 11
2.1 Baltic Sea ................................................................................................ 11
2.2 Biogenic production of natural organohalogen compounds.................... 11
2.3 Hydroxylated polybrominated diphenyl ethers ....................................... 15
2.4 Methoxylated polybrominated diphenyl ethers....................................... 17
2.5 Polybrominated dibenzo-p-dioxins and dibenzofurans........................... 18
2.6 Polybrominated phenols and anisoles ..................................................... 19
2.7 Biological description of studied species ................................................ 20
2.7.1 Algae ........................................................................................................... 20
2.7.2 Cyanobacteria.............................................................................................. 21
2.7.3 Blue mussels................................................................................................ 21
2.7.4 Baltic clam .................................................................................................. 22
2.7.5 Perch............................................................................................................ 22
2.7.6 Flounder ...................................................................................................... 22
2.7.7 Grey seal ..................................................................................................... 23
3. Analytical methods ...................................................................................... 24
3.1 Samples and sampling............................................................................. 24
3.1.1 Algae ........................................................................................................... 24
3.1.2 Cyanobacteria.............................................................................................. 25
3.1.3 Mussels........................................................................................................ 25
3.1.4 Fish.............................................................................................................. 25
3.1.5 Seal.............................................................................................................. 26
3.1.6 Sediment...................................................................................................... 26
3.2 Extraction methods.................................................................................. 26
3.3 Determination of extractable material and carbon content ..................... 27
3.4 Lipid removal .......................................................................................... 28
3.5 Separation of substance groups ............................................................... 31
3.5.1 Separation of neutral and phenolic compounds........................................... 31
3.5.1 Separation of non-planar and planar compounds ........................................ 32
3.6 Derivatisation .......................................................................................... 32
3.7 Instrumental analysis............................................................................... 33
v
3.8 Quality Assurance/Quality Control......................................................... 33
4. Additional results......................................................................................... 35
4.1 Flounders................................................................................................. 35
4.2 PBDD concentration in Askö samples taken at different trophic levels . 37
4.3 Samples from New Zealand .................................................................... 37
4.4 Herring and seal blood concentrations .................................................... 39
5. Discussion ..................................................................................................... 41
5.1 Data normalization .................................................................................. 41
5.2 Trends...................................................................................................... 41
5.2.1 Temporal variations..................................................................................... 41
5.2.2 Seasonal variations...................................................................................... 42
5.2.3 Geographical distribution ............................................................................ 45
5.3 Food web distribution ............................................................................. 47
5.4. Exposure and uptake .............................................................................. 49
5.5 Origin ...................................................................................................... 50
5.6 Ecological perspective ............................................................................ 51
6. Future perspectives ..................................................................................... 52
7. Acknowledgements ...................................................................................... 53
8. References .................................................................................................... 55
vi
Abbreviations
ADP
ASE
ATP
BCF
BFRs
BMF
CYP
DNA
ECD
ECNI
EI
EOM
GC
GPC
HRMS
l.w.
LOD
Log Kow
LOQ
LRMS
MeO-PBDEs
MS
n.a.
NOCs
OC
OHCs
OH-PBDEs
OH-PCBs
PBAs
PBDDs
PBDEs
PBDFs
PBPs
PCBs
PCDDs
PCDFs
pKa
PLE
POPs
psu
Adenosine diphosphate
Accelerated solvent extraction
Adenosine triphosphate
Bioconcentration factor
Brominated flame retardants
Biomagnification factor
Cytochrome P-450
Deoxyribonucleic acid
Electron capture detector
Electron capture negative ionisation
Electron ionization
Extractable organic matter
Gas chromatography
Gel permeation chromatography
High resolution mass spectrometry
Lipid weight
Limit of detection
Octanol-water partition coefficient
Limit of quantification
Low resolution mass spectrometry
Methoxylated polybrominated diphenyl ethers
Mass spectrometry
Not analysed
Natural organohalogen compounds
Organic carbon
Organohalogen compounds
Hydroxylated polybrominated diphenyl ethers
Hydroxylated polychlorinated biphenyls
Polybrominated anisoles
Polybrominated dibenzo-p-dioxins
Polybrominated diphenyl ethers
Polybrominated dibenzofurans
Polybrominated phenols
Polychlorinated biphenyls
Polychlorinated dibenzo-p-dioxins
Polychlorinated dibenzofurans
Acid dissociation constant
Pressurized liquid extraction
Persistent organic pollutants
Practical salinity units
vii
S/N
SIM
TMF
TTR
w.w.
Signal to noise
Selected ion monitoring
Trophic magnification factor
Transthyretin
Wet weight
viii
1. Introduction
In the second phase of the industrial revolution, i.e. in the first half of the 20th
century, organohalogen compounds (OHCs) started to be produced on a
commercial basis, to aid in everyday life. Several of these chemicals were
designed to be stable to enable long lasting use in their applications.
Unfortunately, the chemical stability soon proved to have its drawbacks, also
being stable in the environment, i.e. persistent. The persistency of these
compounds resulted in increasing concentrations in the environment and they
were soon shown to cause adverse effects in wildlife, in particular at high
trophic levels.
During the last Century, the Baltic area was subjected to major discharges of
anthropogenic chemicals and became heavily contaminated. Monitoring
programs were initiated in the Baltic region, including several marine and
terrestrial wildlife species. The research within the monitoring programs was
focused to what we today may call traditional anthropogenic contaminants,
persistent organic pollutants (POPs). In addition to the POPs there are several
thousand of substances, with chemical and structural similarities, but formed
through natural processes [1]. Such chemicals, natural products, are either
formed via biogenic synthesis or metabolic transformations. It is reasonable to
believe that natural products and anthropogenic chemicals may act through
similar mechanisms leading to potentially adverse effects in wildlife. Far less is
known about semi-persistent pollutants, including anthropogenic contaminants
and those being natural products.
1.1 Aim
This thesis focuses on polybrominated compounds in Baltic biota, more
specifically, in biota collected in areas along the Swedish coastline. The aim
was to evaluate origin, assess concentrations, geographical distribution and
indicate ecological relevance of mainly three groups of chemicals; hydroxylated
polybrominated diphenyl ethers (OH-PBDEs), sometimes referred to as
polybrominated phenoxyphenols; methoxylated PBDEs (MeO-PBDEs), also
known as polybrominated phenoxyanisols; and polybrominated dibenzo-pdioxins (PBDDs). The thesis objectives include studies of time trends and
geographic and inter-species distribution of these chemicals. The specific
objectives of the individual papers are described hereunder.
Paper I: The exposure of brominated compounds, in particular OH-PBDEs,
MeO-PBDEs, PBDDs and simple brominated phenols and anisoles were
investigated in a filtrating species, the blue mussel. The study was aimed to
9
assess differences and similarities in the chemical composition in blue mussels
collected at the west coast of Sweden and in the Baltic proper.
Paper II: The original aim was to investigate whether the levels of PBDDs and
MeO-PBDEs in Perch from Kvädöfjärden in the Baltic Sea have increased
during the last two decades and if there were any correlation between the two
substance groups within the studied species. During the data evaluation the
focus was shifted to discuss the individual congener retention and metabolic
stability in perch versus their molecular structures.
Paper III: The goal of this study was to determine any seasonal variation,
within the summer season (May-September), of OH-PBDEs and MeO-PBDEs,
and to discuss possible correlations with the lifecycle of some primary
producers, i.e. algae and cyanobacteria.
Paper IV: This article objective was to study the seasonal and geographic
distribution of OH-PBDEs, MeO-PBDEs and PBDDs in several algae species
and blue mussels as well as inter-species differences between the algae. The
paper also aims to investigate possible relations between the mussel
concentrations and the algae species to try to identify major producers of OHPBDEs, MeO-PBDEs and PBDDs.
10
2. Background
2.1 Baltic Sea
The Baltic Sea including its large bays, the Gulf of Bothnia, Gulf of Finland and
Gulf of Riga, provides a coastal zone for nine countries; Sweden, Finland,
Russia, Estonia, Latvia, Lithuania, Poland, Germany and Denmark. The
drainage area is even greater, also including Belarus, Czech Republic, Norway,
Slovakia and Ukraine. The surrounding lands have approximately two-hundred
rivers giving a yearly fresh water runoff of about 500 km3, a large contribution
of water to this semi-enclosed sea, holding a total water volume of 21 760 km3.
The only inlet of saline waters to the Baltic Sea comes from the North Sea via
the small sounds in Denmark and between Denmark and Sweden. This results in
a low salinity in the Baltic Sea in general, but also gives the sea a gradient from
the southwest to the north and the east of eight to one practical salinity units
(psu). In addition, a large part of the Baltic Sea is vertically stratified into two
parts, with the saltier, heavier and oxygenated water from the North Sea at the
bottom. This barrier prevents the mixing of oxygen and nutrients in the Sea,
leading, in the long run, to dead zones. The salt gradient together with the
parallel temperature gradient affects the flora and fauna, limiting the
biodiversity in the Baltic Sea [2].
Eutrophication is a large problem in most parts of the Baltic Sea [3]. The
nutrient load, mainly originating from municipal and rural human sources and
agriculture, has significantly increased during the 20th century [3]. A slight
decrease in nutrient load in the open Baltic proper has been observed in the
beginning of the 21th century [4]. Regional differences in eutrophication occur,
especially in the coastal waters [5]. The more obvious effects seen in the Baltic
Sea, due to the nutrient enrichment are, the large-scale cyanobacteria blooms
that occur during summer months [6,7], large amounts of macroalgae ending up
on the shores [8], reduced habitat of some species in favour of other more
adaptable species [2] and oxygen depletion [2].
2.2 Biogenic production of natural organohalogen compounds
Formation of natural organohalogen compounds (NOCs) is a common
phenomenon in both the terrestrial and marine environment, with over 3800
identified NOCs produced by either abiotic processes or by biota [9]. The
marine environment is by far the most important source of biogenic NOCs, with
producers such as algae, sponges, corals, tunicates and bacteria as summarized
by Gribble [9]. In the terrestrial environment NOCs may be produced by plants,
fungi, lichen, bacteria, insects, and even in some higher animals including
humans [9].
11
NOCs may include any of the four halogen elements but over 95 % of all NOCs
contain either bromine and/or chlorine [9]. Structures of four common NOCs
are presented, as examples thereof, in Figure 2.1. Bromomethane (1, Figure 2.1)
is a good example of the simple haloalkanes found in both marine and in
terrestrial plants [9]. Indoles (2), bipyrolles (3) and MeO-PBDEs (4) are
commonly detected in the marine biota. These compounds are according to
present knowledge produced by marine sponges, bacteria and fungi, [9].
1
2
3
4
Figure 2.1. Examples of some brominated NOCs. 1) bromomethane; 2) 3,6dibromoindole, 3) 1,1’-dimethyl-3,3’,4,4’,5,5’-hexabromo-2,2’-bipyrrole; 4) 6-methoxy2,2’,4,4’-tetrabromodiphenyl ether
2.2.1 Biosynthesis
In general, the hydrocarbon skeleton of the NOC is formed first, followed by
halogenation. More complex structures can also be formed through fusion of
smaller NOCs. The hydrocarbon skeleton of aromatic organic compounds can
be biosynthesized by three main pathways; the acetate pathway forming
acetogenins, the mevalonate pathway forming terpenoids and sterols or the
shikimate pathway forming aromatic compounds e.g. phenols [10]. Most
brominated simple phenols are produced by the shikimate pathway (starting
with D-glucose), either via 4-hydroxybenzoate or phenol. Since all NOCs
included in this thesis probably are derived via phenol, only the shikimate
pathway is further discussed. This pathway is schematically shown in Figure 2.2
[10,11]. The reaction is enzyme driven and energy consuming, however the
exact mechanism of which the enzyme participates is not known.
12
On enzyme
surface
ATP
+
Enz:
-HO
-H2O
Figure 2.2. Schematic presentation of the formation of 4-hydroxybenzoate and phenol
through the shikimate pathway, starting with phosphoenolpyruvate and erythrose-4phosphate, both originating from D-glucose. The figure is modified from a figure
presented by Nielson [10].
The bromination mechanism of phenolic compounds seems to be a
bromoperoxidase catalysed cationic reaction in the presence of bromine and
hydrogen peroxide [10]. This synthesis begins with an enzymatically catalysed
reaction of hydrogenperoxide and bromine to form the reagents, hypobromous
acid as shown in Figure 2.3. The reagents then undergo an electrophilic reaction
with high electron density centres (Figure 2.3), e.g. an electrophilic aromatic
substitution reaction. The halogen atom is usually introduced in the ortho- or
para-position to the phenolic group in naturally occurring brominated phenolic
compounds. It is also plausible for substrates with low electron density to go
through an anionic bromination by direct insertion of bromide, e.g. brominated
hydroquinones (with hydroxyl groups in the para-positions) [10].
½ H2O2 + Br
bromoperoxidase
HOBr
HOBr +
+ H2O
Figure 2.3. Example of cationic bromination of naturally produced bromophenols.
13
After halogenation, dimerization is achieved through peroxidase catalysed
radical reactions of single ringed, aromatic compounds, forming products such
as biphenyls, diphenyl ethers, bis-indoles, dibenzofurans and dibenzo-p-dioxins
(examples in Figure 2.4) [10]. The peroxidase initiate the coupling reaction of
phenolic dimerization products by a one-electron oxidation in the presence of
hydrogen peroxide, giving a phenoxy radical in the ortho- or para- position
(Figure 2.4). Generally, the oxidation is catalysed by peroxidase, but coupling
may also be catalysed by the cytochrome P-450 system in vascular plants as
shown for the formation of diphenyl ether alkaoids by Berberis stolonifers [12].
O
O
H
O
Figure 2.4. Example of dimerization reactions of phenolic radicals, forming an orthohydroxylated diphenyl ether (left) and an ortho-dihydroxylated biphenyl (right). The figure
is modified from a figure presented by Nielson [10].
14
Among all possible NOCs in the environment, this thesis concentrates on a few
groups thereof.
2.3 Hydroxylated polybrominated diphenyl ethers
OH-PBDEs can be formed as metabolites of PBDEs as reviewed by Haak and
Letcher [13] and as presented in more recent articles [14-16]. However, OHPBDEs are also known to be natural products [1]. The naturally formed OHPBDEs identified so far, have the hydroxyl group in an ortho-position while
PBDE metabolites seem to preferentially have the hydroxyl group in either a
meta- or para-position [13,16]. The ortho-substituted 2’-OH-BDE28, 6-OHBDE47 and 2’-OH-BDE66, identified as minor PBDE congener metabolites,
are exceptions [16].
Naturally formed OH-PBDEs are widespread and have been identified in
several species as exemplified in Table 2.1. They are believed to be synthesised
by primary producers such as algae, marine sponges and cyanobacteria [1,17].
Additional suggested sources are the formation of OH-PBDEs from the
corresponding naturally produced MeO-PBDE congener via biogenic
demethylation [18], or by abiotic oxidation in the atmosphere via reaction of
PBDEs with hydroxyl radicals [19].
Table 2.1. Examples of the worldwide distribution of detected OH-PBDEs in marine
species.
Species
Detected in
Cyanobacteria
Baltic Sea [20]
Algae
Baltic Sea [21], Philippines [22]
Marine Sponge
Indo-pacific ocean [23], Indonesia [24], Palau [25,26],
Mozambique [27]
Mussels
Baltic Sea [21]
Fish
Baltic Sea [28,29], Detroit River [30]
Seal
Baltic Sea [31], Svalbard [31], East Greenland [32]
Polar bear
Norway [33], East Greenland [32]
15
The OH-PBDEs have log Kow values varying between 5 and 9 (Table 2.2)
depending on congener structure but also on method of calculation or software
applied. Although the log Kow values are high and indicates a high
hydrophobicity, the OH-PBDEs do not bioaccumulate in lipid tissue. OHPBDEs behaves like many other halogenated phenolic compounds, such as
halophenols and hydroxylated PCBs (OH-PCBs), by association to blood
proteins, e.g. transthyretin [34]. Further, the water solubility of OH-PBDEs may
increase due to the inverse relationship between water solubility and pKa (Table
2.2) of the phenolic compounds. At natural pH in the marine waters, at least half
of the OH-PBDE concentrations may be present in its ionic form. The range in
pKa values are from approx. 5 to 7 which may result in different uptake of OHPBDE congeners and leading to congener specific exposure. In addition to
being a factor in exposure via direct uptake from water, the pKa value is also
relevant for uptake via the diet.
The OH-PBDEs are linked to several toxicological effects. For example, 6-OHBDE47 is confirmed to be acutely toxic in developing and adult zebrafish at
concentrations in the nanomolar range [35]. The effects are contributed to
disruption of oxidative phosphorylation [35], i.e. inhibiting the phosphorylation
of adenosine diphosphate (ADP) to adenosine triphosphate (ATP), eventually
leading to energy depletion. OH-PBDEs also have a potential to disrupt the
endocrine system [36]. 6-OH-BDE47 and 4'-OH-BDE49 are shown to have
competitive binding to transthyretin (TTR) [37]. In vitro studies in human cells
suggest that meta- and para-OH-substituted PBDEs have 160-1600 higher
relative binding potencies to TTR than BDE-47 [38]. OH-PBDEs are shown to
have both estrogenic effects, through interactions with the estrogen receptor
[39], and anti-estrogenic effects by inhibition of estradiol sulfotransferase [38].
Several OH-PBDEs are also found to inhibit CYP17 and CYP19 (aromatase)
activity in human adrenocortical carcinoma (H295R) cells in micromolar
concentrations [40] and aromatase activity in the human placenta [41]. Further,
6-OH-BDE47 and 6-OH-BDE85 are shown to be cytotoxic in micromolar
concentrations in H295R cells, but do not generate DNA-damage [42].
Dingemans et al. found 6-OH-BDE47 to be neurotoxic, by disrupting the
calcium ion homeostasis in pheochromocytoma cells [43].
16
Table 2.2. Calculated Log Kow and pKa values [17,44] for some of the most common
naturally former OH-PBDEs and log Kow values [45] of naturally occurring MeOPBDEs
Compound
Log Kow
pKa
ACD
Experimental
ACD
6-OH-BDE47
6.8 ± 0.2
5.82 ± 0.03
6.8 ± 0.2
2’-OH-BDE68
7.2 ± 0.6
5.36 ± 0.04
6.6 ± 0.2
6-OH-BDE85
8.3 ± 0.7
5.83 ± 0.02
6.2 ± 0.2
6-OH-BDE90
8.2 ± 0.7
5.83 ± 0.03
5.7 ± 0.2
6-OH-BDE99
8.4 ± 0.7
2-OH-BDE123
8.3 ± 0.7
5.82 ± 0.03
5.7 ± 0.2
6-OH-BDE137
9.3 ± 0.8
6.45 ± 0.03
4.7 ± 0.2
5.2 ± 0.2
6-MeO-BDE47
6.44 ± 0.01
2’-MeO-BDE68
6.16 ± 0.02
6-MeO-BDE85
6.26 ± 0.01
6-MeO-BDE90
6.65 ± 0.03
2-MeO-BDE123
6.62 ± 0.01
6-MeO-BDE137
6.98 ± 0.03
ACD/LABs TM software
2.4 Methoxylated polybrominated diphenyl ethers
MeO-PBDEs have been identified as natural products by determination of
radiocarbon (14C) content of two MeO-PBDEs (6-MeO-BDE47 and 2’-MeOBDE68) isolated from a True's beaked whale (Mesoplodon mirus) [46]. MeOPBDEs have also been identified in a whale oil sample from 1921, sampled
before any industrial production of OHCs started [47]. For many years no MeOPBDEs metabolites have been indicated due to PBDE exposure, and
accordingly considered to be solely of natural origin. Lately however, Feng et
al. reported MeO-PBDEs in rainbow trout after exposure to decabromodiphenyl
ether [48]. This is to my knowledge the only study supporting the metabolic
formation of MeO-PBDEs from PBDEs. Further, microbial methylation of OHPBDEs may occur in e.g. sediments by microorganisms [49-54]. MeO-PBDEs
are known to bioaccumulate in tissue as indicated by their log Kow (Table 2.2)
and have been identified in many species worldwide, some of which are
summarized in Table 2.3. The log Kow values, varying from 6-7 (Table 2.2.), are
higher for the MeO-PBDEs than the OH-PBDEs.
17
Table 2.3. Examples of the worldwide distribution of detected MeO-PBDEs in marine
species.
Species
Detected in
Cyanobacteria
Baltic Sea [20]
Algae
Baltic Sea [21], Australia [55-57], China [58], Philippines [22]
Mussels
Baltic Sea [21], China [59], Canadian Arctic [60]
Fish
Baltic Sea [28,29,61,62], Canadian Arctic [60], Mediterranean
Sea [63], Detroit River [30]
Seal
Baltic Sea [61], Canadian Arctic [60], East Greenland [32],
Southern North Sea [64]
Polar bear
Norway [33], East Greenland [32]
The toxicity of MeO-PBDEs is low , but some studies have reported effects in
cell cultures [40,41,65-67] with very high but not environmentally relevant
levels. For example, 6-MeO-BDE47 inhibits the CYP17 [65] and the aromatase
(CYP19) [40,67] activity but does not affect the sex hormone production [67] or
show any cytotoxcity [40,65]. However, the possibility of demethylation
[18,68-70], forming the corresponding OH-PBDE, may have toxicological
implications.
2.5 Polybrominated dibenzo-p-dioxins and dibenzofurans
PBDDs and polybrominated dibenzofurans (PBDFs) are coplanar compounds,
formed as by-products in brominated flame retardant (BFR) production [71] and
combustion of BFR containing products [71-74]. PBDFs can also be formed
though photolytic transformation of decaBDE [75,76]. PBDDs/Fs undergo
photolysis more rapidly than polychlorinated dibenzo-p-dioxins (PCDDs) in sun
and indoor light [77]. In addition, PBDDs may be formed via photolysis of OHPBDEs [78].
The PBDDs formed in combustion are dominated by tetra- and pentaBDDs, but
the congener composition may differ with material and combustion temperature
[72]. High levels of lower brominated dibenzo-p-dioxins (Br1-Br4), in Baltic Sea
biota have led to a discussion of natural formation of these compounds
[17,20,79-81] (Paper I, II and IV). Non-halogenated dibenzofurans are
common among identified and reported natural products [10,82], but
halogenated dibenzofurans and dibenzo-p-dioxins are not [10]. One example of
a natural PBDF is the 2-bromodibenzofuran found in the sponge
18
Chelonaplysilla sp. [10]. Further, derivatives of PBDDs have been isolated from
marine sponges [83,84] and derivatives of PBDFs from red algae [85].
Naturally produced PBDDs and PBDFs may be formed through diaryl coupling
of phenolic radicals, as presented in chapter 2.2.1 and Figure 2.4.
PBDDs/Fs have e.g. been found in cyanobacteria [20], algae [20], mussel
[79,81], marine sponge [86] and fish [79] from the Baltic Sea, in shellfish and
fish from the west coast of Sweden [79], in marine shellfish from United
Kingdom [87], as well as, in human adipose tissue from Japan [88] and in
human breast milk [89]. PBDDs/Fs have also been detected in sediments
[90,91].
Information on biological effects of PBDDs/Fs is limited and often deduced
from the knowledge obtained from studies of PCDDs and polychlorinated
dibenzofurans (PCDFs). In vivo toxicity studies of PBDDs show biological
effects associated with PCDDs/Fs, i.e. lethality, wasting, thymic atrophy,
tetratogenicity, reproductive effects, chloracne, immunotoxicity, enzyme
induction, decrease in T4 and vitamin A and increased hepatic porphyrins [71].
In addition, in vitro enzyme induction and anti-estrogenic activity are linked to
PBDDs/Fs [71]. PBDDs/Fs are also potent inducers of microsomal
monooxygenase activity, aryl hydrocarbon hydrolase and ethoxyresorufin-odeethylase (EROD) both in vitro and in vivo [92]. PBDDs/Fs can bind to the
aryl hydrocarbon receptor (AhR) [93,94], but the binding affinity is generally
half compared to the chlorinated analogues as reviewed by Birnbaum et al. [95].
The toxic equivalent system used to compare PCDD toxicity is not yet
developed to include the PBDDs.
2.6 Polybrominated phenols and anisoles
PBPs are produced in several anthropogenic processes. 2,4,6-TriBP is produced
in large scale and is by far the most common PBP in the world. In 2001 the
worldwide annual production was 9500 tonnes. 2,4,6-triBP is used as a wood
preservative and, both 2,4-diBP and 2,4,6-triBP are employed as reactive flame
retardant intermediates [96-99]. PentaBP has been used as a mulluscicide [100]
as well as an intermediate in the production of pentabromophenoxy compounds.
2-monoBP, 2,4-diBP, 2,6-diBP, 2,4,6-triBP has been identified in vehicle
emission of leaded petrol [101].
PBPs are also produced naturally in large quantities in marine biota [1]. For
example, the acorn worm Balanoglossus biminiensis produces up to 15 mg 2,6diBP per animal as a defensive secretion [102]. Several species of marine algae
are known to contain [56,57] and biosynthesis [55,103] brominated phenols.
The abundance of PBPs is both spatially and temporally correlated with the
19
abundance of infauna that produces these metabolites [104]. Although there is a
worldwide anthropogenic production and use of 2,4,6-triP, the amounts released
into the environment from natural sources is proposed to be more abundant.
PBPs can also be formed via biodegradation of other pollutants, such as
brominated benzenes and some brominated diphenyl ethers [1,18,105]. Further,
PBPs can be formed from demethylation of polybrominated anisoles (PBAs)
under anaerobic conditions [100].
The estimated bioaccumulation potential of PBPs increases with the degree of
bromination, as indicated by the log Kow values presented in Table 2.4.
Predicted bioconcentration factors (BCFs) (Table 2.4) suggest some potential
for bioaccumulation [100]. However, at natural marine pH both the 2,4,6-triBP
and the pentaBP are mostly dissociated (Table 2.4), and accordingly the route of
uptake differs between congeners.
Table 2.4. Reported log Kow and pKa [106] and BCF [100] values for some PBPs.
Log Kow
4-monoBP
2,4-diBP
2,4,6-triBP
Penta-BP
1
2
1
pKa
2.62
3.48
4.24
5.30
1
9.17
7.79
6.08
4.40
BCF
2
20
24
120
3100
at 25ºC [106]
Calculated using Bcfwin [100]
PBAs are formed in methylation processes in the marine environment, e.g.
2,4,6-triBA can be formed as a fungal metabolite of 2,4,6-triBP [10] or by Omethylation in bacteria [53].
2.7 Biological description of studied species
A short introduction to the biology of the species analysed and discussed within
this thesis is given here. More detailed information of the samples and sampling
can be found in chapter 3.1 and the Papers I-IV.
2.7.1 Algae
The family of algae is vast, including marine autotrophic and eukaryotic
organisms, ranging from unicellular to multicellular organisms. This thesis is
20
concentrated on macroscopic, multicellular, benthic marine algae, summarized
from now on as filamentous macroalgae. The term includes members of red,
brown and green algae. The difference in colour of the algae is attributed to
their pigments being optimized to absorb light at their habitat sea depth. The
green algae grow closest to the surface followed by the brown algae and the red
algae.
The red algae are one of the oldest and the largest of the groups of eukaryotic
algae, with somewhere between 6000 and 10000 species. Most of which are
filamentous macroalgae with sexual reproduction. The red colour is given by
the accessory pigments, phycobiliproteins. The brown algae is a large group of
mostly marine multicellular algae with about 1500-2000 species. They play an
important role in marine environments both as food, and for the habitats they
form. Most brown algae contain the pigment fucoxantin (brown pigment) and
chlorophyll (green pigment), giving them their characteristic greenish-brown
colour. Brown algae reproduce by means of both flagellate spores and gametes.
The green algae are usually single cell organisms, while others form colonies,
long filaments or macroscopic seaweeds. There are about 8000 species of both
fresh water (7000) and marine green algae (1000). The green colour is given
from chlorophyll a and b and the reproduction is commonly sexual.
2.7.2 Cyanobacteria
Cyanobacteria constitute a large and diverse group of bacteria capable of
oxygen photosynthesis and are found in most waters worldwide. Cyanobacteria
are unicellular or filamentous and can form colonies or aggregates.
In the Baltic Sea there are three main species of cyanobacteria; Nodularia
spumigena, Aphanizomenon flos-aquae and Anabaena spp. The cyanobacteria
blooms are initiated by calm and sunny weather, elevated surface water
temperature and thermal stratification. Nitrogen fixating cyanobacteria such as
Aphanizomenon and Nodularia are also depending on phosphate avalability.
The Aphanizomenon flos-aquae or hepatoxin containing Nodularia spumigena
usually dominated the large cyanobacteria blooms formed during the summer in
the Baltic.
2.7.3 Blue mussels
The blue mussel (Mytilus edulis) is a suspension-feeding marine bivalve
mollusc found worldwide in temperate and cold oceans. The blue mussels attach
themselves to hard surfaces such as cliffs, rocks or tongs with their byssus
threads. They are very robust and can stand large variations in temperature and
salt content. The blue mussel will reach reproductive age at one year.
Reproduction occurs from early spring into the autumn by releasing their
21
gametes into the surrounding waters. The larvae are pelagic and swim for 2-3
weeks before attaching themselves to any surface.
In the Baltic Sea the blue mussels have adapted to the brackish water, but they
are much smaller than in areas like the North Sea where the salinity is higher.
The differences are genetic [107], morphologic [108] and physiologic [109].
Some even argue that the Baltic blue mussel belong to a different sub-species
and has been named Mytilus edulis trossulus [110]. In this thesis, however, the
sub-species of the blue mussels have not been considered as a factor since the
species are only used to monitor exposure of contaminants.
2.7.4 Baltic clam
The Baltic clam (Macoma baltica) is a bivalve living in sandy and clayey Sea
bottoms. It is so named since its habitat is the entire Baltic Sea. The Baltic clam
lives buried in the sediments eating small plant and animal parts from the Sea
bottom. The Baltic clam is a popular diet for Saduria entomon, an isopod
crustacean without a common name, and flounders.
2.7.5 Perch
The Perch (Perca fluviatilis) is a relative stationary fish species, only migrating
to reach their spawning location. The sexual maturity is reached at the age of 24 years for the males and 3-5 years for the females. The spawning takes place
during April to June in the Baltic Sea. During the first life year the Perch feeds
on zooplankton and then moves on to insect larvae, crustaceans and small fish.
2.7.6 Flounder
The European flounder (Platichthys flesus) is an ocean-dwelling flatfish of
European coastal waters, feeding on invertebrates, especially crustaceans,
worms, molluscs, as well as small fish. The flounder used within this thesis
were all from the Swedish coast, both from the west coast of Sweden
(Skagerrak and Kattegat) and from the Baltic Sea. The feeding habits are
somewhat different between the two coasts, e.g. flounders from the Baltic feed
on blue mussels which is not possible for the flounders on the west coast. The
time of reproduction is different as well; in Skagerrak and Kattegat it takes
place in January to April and in the Baltic Sea from May to June. This has lead
to some debate whether the flounders from these locations are of the same subspecies or not. Reproductive age for males and females are reached at two and
three years of age, respectively.
22
2.7.7 Grey seal
The Grey seals (Halichoerus grypus) in the Baltic Sea are an isolated population
and thus called (Halichoerus grypus balticus). This mammal feeds on a wide
variety of fish, e.g. sand eels, cod, flatfish but mainly herring. Grey seals are
feeding at a high trophic level and are a well-studied species [111]. It is
suffering from health effects like decreased body weight and/or blubber
thickness [112,113] and colonic ulcers [114].
23
3. Analytical methods
The analytical methods used within this thesis are well established methods for
environmental contaminant analysis. However, in some cases these methods
were modified to meet the requirements of the objectives in this thesis. Detailed
information of the methods used, are given in the separate publications (Paper
I-IV).
3.1 Samples and sampling
Samples from a number of sampling locations along the Swedish coast line
were used in this thesis. The locations are indicated on the map shown in Figure
3.1 and each of them is further presented in Paper I-IV. Additional samples
were collected to study food web distribution of OH-PBDEs, MeO-PBDEs and
PBDDs (see Chapter 4, below).
Figure 3.1 Map over the Swedish coastline with the
sampling locations marked 1-9. They are:
1.) Hornslandet
2.) Arholma
3.) Askö
4.) Kvädöfjärden
5.) Öland
6.) Abbekås
7.) Fladen
8.) Väderöarna
9.) Tjärnö
3.1.1 Algae
Brown algae, Dictyosiphon foenicolaceus (Paper I), Fucus vesiculosus (Paper
IV) from Kvädöfjärden (4 in Figure 3.1) and Pilayella littoralis from Askö (3)
and Hornslandet (1) (Paper IV) were collected in the autumn of 2006. Red
algae, Ceramium tenuicorne from Askö (3), Hornslandet (1), Arholma (2) and
Öland (Byxelkrok) (5) and Ceranium rubrum from Tjärnö (9), Polysiphonia
fucoids from Askö (3) and Abbekås (6), Polysiphonia brodari from Tjärnö (9)
and Furcellaria lumbricalis from Askö (3) and Abbekås (6) (Paper IV) were
collected between 2006 and 2009. Green algae, Cladophora glomerata from
Askö (3), Hornslandet (1) and Öland (5), Cladophora albida from Tjärnö (9)
24
and Enteromorpha intestinalis from Öland (5) (Paper IV) were collected
between 2006 and 2009. The algae were collected by hand, extensive water was
wrung out, and the samples were homogenized (Paper I and IV).
3.1.2 Cyanobacteria
Cyanobacteria (Nodularia spumigena) from Landsort Deep (Paper I) were
collected in the autumn of 2005 Aphanizomenon sp. (Chapter 4) was sampled
from Askö (3) during 2006.
3.1.3 Mussels
The blue mussels, presented in Paper I, were sampled from the background
location, Kvädöfjärden (4), as well as, from two background locations along the
west coast of Sweden, Fladen (7) and Väderöarna (8) by hand or by nets or
using scapers. Blue mussels were also collected from Askö (Paper III and
Paper IV), Kvädöfjärden, Arholma, and Abbekås (Paper IV) (Figure 3.1).
Sampling was either done with a scraper dragged along the bottom behind a
small boat, or collected by divers.
Baltic clams were sampled from the Askö area (3) (Chapter 4.2) using a scoope
that was lowered down to the sea bottom to collect sediment also containing the
Baltic clams. The sediment was removed by running water over a sieve and the
mussels handpicked.
Each mussel locations and time points were considered as one sample. The
samples were homogenised to reduce the effect of individual variations.
3.1.4 Fish
The perch and flounder, presented in Paper I and II, were sampled within the
Swedish Environmental Monitoring Program on Contaminants in Biota
(SEMPC). Perch and flounder were collected using gill nets from Kvädöfjärden
(4) located close to Swedish Baltic coastline. Flounders were also collected
from the west coast locations, Fladen (7) and Väderöarna (8). The perch were
selected by age (2-years) and all samples were collected during late summer or
autumn to ensure that the sampled individuals were well nourished and were not
reproducing. Fish muscle from the middle dorsal muscle layer (without skin and
subcutaneous fat) was used for analysis. Composite samples of >10 fishes were
prepared in order to reduce the effect of individual variations. Perch, flounder
and herring (Clupea harengus) were all collected from Askö (3) using gill nets
(Chapter 4). Blood were sampled from herrings directly after the fish had been
detangled from the gill nets. Blood from 21 herrings were drawn with a small
syringe from the blood vessel at the backbone. The fishes were numbed before
sampling and were immediately put to death by a crushing blow to the head
25
afterwards. The appropriate permit for animal experiments was obtained (No: N
147/06 and N 170/09). Heparin was added to the blood samples and the plasma
was separated from the blood cells.
3.1.5 Seal
Blood coagulate samples from 14 grey seals (Chapter 4) were collected by
personnel from the Swedish Museum of Natural History upon autopsies of the
seals. The samples were collected from seals that were found drowned in the
Baltic proper, between 1995 and 2006.
3.1.6 Sediment
Sediment core samples were collected in 2005 (Paper II) using core samplers
and were sectioned on board the sampling vessels. Top sediment was used in
the study.
3.2 Extraction methods
Organic environmental contaminants of concern are primarily lipophilic and
thus the extraction methods have been optimized for extraction of lipids and
lipid soluble compounds. Historically, Soxhlet extraction [115,116] and batch
extractions [117-119] were used. The liquid-liquid extraction method developed
by Blight and Dyer employing a solvent mixture of methanol and chloroform is
commonly used for lipid extraction. Jensen and co-workers developed a method
of equal lipid extraction efficiency for fatty aquatic organisms without using
halogenated solvents [119]. This method was later modified to give good lipid
extraction also for lean matrices by substituting acetone for 2-propanol [120].
The improvement was probably due to better extraction of phospholipids.
Further work lead to the most recent method that was optimized for extraction
of phenolic analytes in fish and blue mussels [121], changing the ratio of nhexane to diethyl ether from (9:1) to (3:1).
All biological samples within this thesis were extracted according to Jensen et
al. [119] (Paper I and II) or Jensen et al. [121] (Paper III and IV), with some
minor alterations. For example the diethyl ether was replaced by methyl-tertbutyl ether in Paper IV. In addition, the n-hexane was replaced with c-hexane
in Paper III and IV to reduce the risk of the analytical procedure.
The phenolic compounds analysed are not strictly lipophilic. Relatively few
methods have been optimized for simultaneous analysis of phenolic and neutral
compounds in tissue samples [115,121-124]. Methods developed for extraction
of phenolic and neutral compounds are e.g. liquid-liquid extraction
[117,121,125] and pressurized liquid extraction (PLE) [123,124]. PLE, also
26
called accelerated solvent extraction (ASE), use conventional solvents under
enhanced temperature and pressure. Anhydrous sodium sulfate or Hydromatrix
are often used as dehydrating agents for extraction of tissue samples using PLE.
When using anhydrous sodium sulfate low recovery for phenolic compounds
such as OH-PCBs and OH-PBDEs were reported in some studies [123,124].
The liquid-liquid extractions used in Paper I, II and IV were not evaluated for
dioxin analysis, in particular, within this thesis. However, since the dioxins are
neutral lipophillic compounds and resembles the structures of e.g. PCBs and
DDTs, it may be concluded that the same extraction methods can be used. The
extraction method used was chosen to be the same for PBDD/F as for the OHPBDE and MeO-PBDE analysis. Commonly, dioxin analyses of biotic samples
are carried out by mixing the tissues with sodium sulfate for dehydration,
placing the mixture in a glass column and extracting the analytes with hexane
and dichloromethane [126] or by Soxhlet extraction [127]. Similar
concentrations of PBDD/F in biota were reported in studies using the sodium
sulphate method [80] as in the Papers presented herein (Paper I, II and IV).
The samples for PBDD/F analysis were extracted in parallel to the samples
extracted for MeO-PBDEs and OH-PBDEs, instead of using the same samples.
This has been done to ensure that the PBDDs found are not artefacts formed
during the potassium hydroxide partitioning (see chapter 3.5.1).
3.3 Determination of extractable material and carbon content
Analytical data require some sort of normalisation to make comparison of data
possible and correct. Usually, lipophilic compounds are presented on lipid
weight (l.w.) or on wet weight (w.w.) basis. Lipid weight basis allows a better
inter-species comparison than fresh weight, especially when comparing
biomagnification. Wet weight and lipid weight have been determined
gravimetrically in all Papers. Due to the samples composition it is difficult to
compare matrices like cyanobacteria, algae, mussels and fish. These matrices
contain very different amounts of water and lipids that make comparison on a
wet weight basis particularly problematic. Further, the species analysed have
very different lipid composition (Table 3.1), implying difficulties in doing the
comparisons on a lipid basis, as well. The data in Paper II is presented on lipid
weight basis, while Paper I, III and IV is normalized on extractable organic
matter (EOM). EOM is the equivalent to lipid weight, but not only lipids are
extracted from e.g. algae. EOM includes all compounds hydrophobic enough to
be extracted with the solvents used, e.g. some pigments. Still, it needs to be
pointed out that it is difficult to determine EOM gravimetrically in e.g. algal
samples since the total weights are low. Therefore, the data presented in Paper
IV were normalized in relation to the organic carbon (OC) content in the
27
samples. Carbon content was determined by measuring the oxidation of carbon
to carbon dioxide under combustion of freeze dried samples, with elemental
analysis. Finally, it seems that the results are in general comparable for all
normalization methods used (Paper IV).
3.4 Lipid removal
Lipids were removed, in all samples (Paper I-IV), by treatment with
concentrated sulfuric acid and silica gel columns treated with sulfuric acid. The
sulfuric acid treatment is a destructive method and accordingly not suitable for
all analytes, but PCBs, PBDEs, MeO-PBDEs and PBDDs/PBDFs are not
affected, allowing sulfuric acid to be used for clean-up of these analytes. For
analysis of analytes sensitive to sulfuric acid or more lipid containing matrices a
non-destructive method such as gel permeation chromatography (GPC)
[28,122,124] or acetonitrile partitioning [128] may be recommended. GPC is
separating molecules on size, but the solvent used as well as the polarity and
planarity of the analytes also affect the separations [129].
Acetonitrile partitioning is used for lipid reduction, by dissolving aromatic
analytes like PCBs to a higher extent than lipids. The partitioning is explained
by -electrons interactions between the aromatic compounds and nitrile group
in the acetonitrile. The more lipophillic the aromatic analytes are the less
soluble they are in acetonitrile, thus to insure a good recovery of the most
lipophillic analytes the partitioning is often repeated three times. Every
treatment will solve approximately 10% of the total lipids, resulting in a 70%
lipid reduction. However, the lipid composition will affect the effectiveness of
the reduction.
The sulfuric acid clean-up procedure was originally developed for removal of
fat in biological tissue containing e.g. triglycerides. Several of the samples
included in the studies within this thesis have a slight different lipid
composition compared to e.g. tissue from fish (Table 3.1). The presence of lipid
soluble pigments (Chlorophyll) may affect the EOM determination. During the
present work it became clear that the clean-up processes applied for some of the
species/matrices were not sufficient, for example for certain algae samples.
Problems were observed as drifts in retention time and sometimes as a broad
“fat peak” in the chromatograms. The lipid composition varies slightly
depending on e.g. species, season, feed, salinity, etcetera [130-136]. Thus, the
examples of lipid compositions of a few analysed species in this thesis,
presented in Table 3.1, are generalization over time and species. It is obvious
that lipids will not behave in the same manner during extraction and clean-up,
due to their differences in chemical structure (Figure 3.2). In the future,
28
improved clean-up and lipid removal methods are required to target the lipids
within the matrices, in particular for plants like the algae samples.
Analysis of phenols (usually simple phenols) in algae samples have in many
cases been based on extraction by polar solvents such as methanol or ethyl
acetate [137,138]. These methods did not extract lipids to a major extent.
Malmvärn tried to remove the algae matrix using GPC, and although it worked
in principal, the matrix proved hard to elute resulting in re-conditioning
difficulties and thus, the GPC is not a useful tool in algae clean-up [17].
MEMBRAN LIPID
Cholesterol
STORAGE LIPID MEMBRAN LIPID MEMBRAN LIPID
triglycerol
phospholipid
glycolipid
R = alcohols
X = saccharide
PIGMENT
Chlorophyll a
STORAGE LIPID
fatty acid
STORAGE LIPID
wax
Figure 3.2. Chemical structures of some common lipids discussed within this thesis.
.
29
Table 3.1. Examples of lipid content composition in cyanobacteria, brown and green algae, mussels, fish muscle and plasma from rat and
fish.
Cyanobacteria
References:
Free fatty acids
Triacylglycerols
Waxes
a
Pigments
Phospholipids
Glycolipids
Sterols
[131,139,140]
+
+
II
III
I
+
Algae
Mussels
Brown
[130]
Green
[141]
[132]
V
III
II
n.a.
II
I
IV
I
IV
III
V
Fish muscle
Plasma
[134,135,142]
Rat
[136]
Fish
[143]
III
II
I
IV
I
(+)
I
II
IV
III
n.a.
II
n.a.
n.a.
I
n.a.
III
IV
II
III
The relative composition is given as roman numbers corresponding to the relative amounts found of each lipid group, I being in highest
abundance. A repeated number describes an equal contribution and “+” indicates presence. Only lipids present in >1% of the total lipid
content are presented.
a
including: carotene and chlorophyll
30
3.5 Separation of substance groups
Separation of substance groups is done to ensure minimum co-elution of
individual compounds upon instrumental analysis. It also facilitates the
possibility of treating different substance groups with the appropriate clean-up
procedure. A schematic description of the methods used (Paper I-IV) is shown
in Figure 3.3.
OH- and MeO-PBDEs
PBDDs
Extraction
Extraction
EOM determination
EOM determination
Separation
KOH
Clean-Up
H2SO4
Fraction 3
Back flush
OH-PBDEs
MeO-PBDEs
Separation
Charcoal Column
Derivatization
Diazomethane
Clean-up
H2SO4
Fraction 1 and 2
Planar
e.g PBDDs
Clean-up
H2SO4
Clean-up
H2SO4:SiO2-gel Columns
Non-planar
e.g. PCB
Clean-Up
H2SO4:SiO2-gel Column
Clean-up
H2SO4:SiO2-gel Column
Clean-up
SiO2-gel Column
Clean-up
SiO2-gel Column
Analysis
GC-LRMS
Analysis
GC-HRMS
Analysis
GC-LRMS
Figure 3.3. Scheme for sample clean-up for analysis of OH-PBDEs and MeO-PBDEs
(left) and PBDDs (right).
3.5.1 Separation of neutral and phenolic compounds
Neutral and phenolic compound are separated using potassium hydroxide
partitioning [144] (Paper I-IV). The phenolic compounds are re-extracted from
the potassium hydroxide phase after acidification. It has been shown that the
partitioning of phenols was not complete [121], and thus the potassium
hydroxide partitioning was repeated twice in Paper III and IV. The potassium
hydroxide may theoretically promote ring closure of ortho-OH-PBDEs forming
PBDDs. This is only possible when there is a bromine substituent in the nonhydroxylated ring, according to predioxin reactions, as reported for the
chlorinated counterparts by Jensen et al. [145]. A separate extraction was thus
carried out for PBDD/F analysis to minimize the risk of artefacts.
31
Separations of neutral and phenolic compounds can also be achieved on
Florisil® columns [122], silica gel SPE cartridges [124] or silica gel columns
with acidified mobile phases [121].
3.5.1 Separation of non-planar and planar compounds
To enable clean extracts during dioxin analysis, coplanar compounds are
separated from non-planar. Separation can either be achieved by a 2-(1pyrenyl)-ethyldimethylsilylated silica column [146,147] or as utilized in Paper
I, II and IV, by using a column with activated charcoal mixed with celite. The
non-planar compounds were washed from the charcoal column and the planar
compounds were recovered through back-flush of the column. The planar
compounds were cleaned-up using a silica gel column impregnated with sulfuric
acid. No clean-up was done of the non-planar fractions (Figure 3.3).
3.6 Derivatisation
Phenolic compounds were derivatised to ensure improved gas chromatographic
(GC) behaviour at analysis (Paper I, III and IV). Untreated phenolic
compounds will substantially interact with the stationary phase in the GC
column, leading to misshaped and wide peaks or in worst case no visible peaks
at all. The derivatisation was achieved through methylation using diazomethane,
synthesised in house from N-methyl-N-nitroso-p-toluene sulfonamide [148] and
dissolved in diethyl ether. Diazomethane is carcinogenic and explosive and thus
laboratory work requires a permit. The use of diazomethane has been approved
by the Swedish work environment authority. The derivatisation of the
commonly found natural OH-PBDEs using standards were investigated and
found to be almost complete (unpublished data).
Methylation is a common way of derivatisation leading to the formation of
stable methyl ethers. Other groups have used methyl iodine [149] or methyl
chloroformate [122], finding these as adequate, and even indicate to give a
better recovery than diazomethane. Methoxy-derivates are very stable and easy
to analyse and tolerate destructive clean-up methods (e.g. concentrated sulfuric
acid). In contrast, acetylation e.g. pentafluorobenzoyl chloride (PFBCl) [121]
and silylation [150] are unstable and instrumental analyses have to be carried
out fast. In addition, these derivatisation agents increase the molecular weight of
the analytes to a larger extent than methylation, prolonging the retention time
and also complicating identification for heavier analytes by approaching the
maximum mass range of the low resolution mass spectrometer (LRMS)
instruments in use (m/z <1000).
32
3.7 Instrumental analysis
Most analyses were carried out by gas chromatography/mass spectrometry
(GC/MS). PBDDs and PBDFs were analysed using electron ionization (EI) and
high resolution mass spectrometry (HRMS; R >10000). MeO-PBDEs were
analysed using MS in the electron capture negative ion chemical ionization
(ECNI) mode. All quantifications were done in ECNI mode employing selected
ion monitoring (SIM), scanning for the bromine ions m/z 79 and 81. Thus, the
method is not compound selective and identification depends solely on the
retention time and may therefore result in identification problems with possible
co-elutions. However, the retention time, elution order and the mass spectra of
MeO-PBDEs have been thoroughly investigated [21,151]. In addition, ECNI
fullscan was employed on mussel samples in Paper I (unpublished) to ensure
the correct identifications.
3.8 Quality Assurance/Quality Control
Solvent blank samples were analysed in parallel to the samples. In Paper I, III
and IV small amounts of PBDEs were detected in the blank samples and were
probably associated with PBDE in laboratory dust. When blank contamination
was an issue, the sample concentrations were adjusted for the blank values as in
Paper I. In Paper III and IV the obvious contamination was not deducted
from the samples but effect the limit of quantification (LOQ) (Se bellow within
this paragraph).
Surrogate standards were used to control the recovery. Generally the recoveries
of the analyses were high for the biological tissues analysed (e.g. blue mussels
and fish) in this thesis work. Lately however, problems with low recovery of
phenolic compounds in some matrices occurred. In blood samples from fish and
seals the recoveries were low, a problem that was not observed when the same
extraction method was used for human blood samples. The recoveries of the
surrogate standards were 28 ± 20% for 4-OH-BDE121 in herring plasma [152],
chapter 4.4) and 15 ± 14 % and 7 ± 9 % for 4-OH-BDE121 and 2’-OH-BDE28
in seal blood coagulate (Chapter 4.4). The recoveries of the neutral surrogate
standards were satisfactory in both in the seal blood and the herring plasma. The
recoveries were; 4-MeO-BDE121 (78 ± 4 % and 77 ± 12 %) and BDE138 or
BDE77 (77 ± 12 % and 89 ± 16 %), respectively. Hence there was a difference
between the slightly acidic OH-PBDEs and the neutral compounds.
A laboratory reference material, consisting of a composite sample of blue
mussel tissue from the west coast of Sweden bought in a supermarket, was used
for Paper IV to ensure precision in the analysis.
33
Limit of detection (LOD) and LOQ are important in trace analysis. In general,
the LOD is not critical for the OH-PBDEs and the MeO-PBDEs in Baltic biota,
since the levels of these compounds are so high. The LOD was defined as the
quantity giving rise to a signal with a signal-to-noise (S/N) ratio of 3. The LOQ
for phenols and anisoles was defined as a signal 10-fold greater than the
standard deviation of the S/N ratio. If the blank samples were contaminated, the
LOQ was defined as 3-fold greater than the background signal. For the PBDD
analyses LOQ was set equal to the LOD, since there were no interferences and a
high signal quality.
34
4. Additional results
In this chapter some unpublished data are presented to complete the picture of
the research done within this thesis. Hence OH-PBDE, MeO-PBDE and PBDD
concentrations as determined in flounder from the east and west coast of
Sweden are presented below. Further, levels of the PBDDs are presented in food
webs from the Baltic Sea. Analytical data of OH-PBDE, MeO-PBDE and
PBDD in herring and seal blood from the Baltic Sea are presented as well as in
material from the south-western Pacific Ocean around New Zealand.
4.1 Flounders
Flounders from Kvädöfjärden in the Baltic proper and west coast of Sweden,
Fladen and Väderöarna were analysed for MeO-PBDEs and PBDDs. The
analyses were done according to the methods described in Paper I.
The flounders from Baltic Sea have approximately ten times higher
concentrations of ∑MeO-PBDEs (59 ng/g EOM) than in flounders from Fladen
(0.59 ng/g EOM) and Väderöarna (0.54 ng/g EOM), respectively. The MeOPBDE congener composition presented in Figure 4.1 is similar for flounders in
Kvädöfjärden and Fladen, but the flounders from Väderöarna have a higher
contribution of 2’-MeO-BDE68. The ∑PBDDs concentration in flounders from
Kvädöfjärden was 0.025 ng/g EOM, while no PBDDs were detected in
flounders from the west coast.
15%
0% 0% 0%
Kvädöfjärden
0%
34%
6-MeO-BDE47
2'-MeO-BDE68
6-MeO-BDE85
6-MeO-BDE90
6-MeO-BDE99
2-MeO-BDE123
6-MeO-BDE137
51%
Väderöarna
Fladen
1% 0% 0%
21%
0%
39%
9%
0% 0% 3%
9%
0%
39%
79%
Figure 4.1. MeO-PBDE congener specific contributions in flounders from the Baltic Sea
(Kvädöfjärden) and from the west coast of Sweden (Fladen and Väderöarna).
35
350
∑tetraBDDs
14
12
10
8
6
4
2
0
ng/g EOM
300
ng/g EOM
250
200
150
∑triBDDs
∑diBDDs
∑monoBDDs
100
50
0
2006-06-23
2006-08-16
2007-05-09
2007-08-19
2006-10-06
2006-10-06
Cyanobacteria
Blue mussels
Blue mussels
Baltic clam
Perch
Flounder
80
∑tetraBDDs
70
ng/g C
60
ng/g C
50
40
5
∑triBDDs
4
∑diBDDs
3
∑monoBDDs
2
1
30
n.a.
0
20
10
0
2006-06-23
2006-08-16
2007-05-09
2007-08-19
2006-10-06
2006-10-06
Cyanobacteria
Blue mussels
Blue mussels
Baltic clam
Perch
Flounder
250
∑tetraBDDs
ng/g d.w.
ng/g d.w.
200
150
3
∑triBDDs
2
∑diBDDs
2
∑monoBDDs
1
1
100
n.a.
0
50
0
2006-06-23
2006-08-16
2007-05-09
2007-08-19
2006-10-06
2006-10-06
Cyanobacteria
Blue mussels
Blue mussels
Baltic clam
Perch
Flounder
Figure 4.2. PBDDs concentrations in species sampled from the Askö area during 2006
and 2007. The concentrations are presented in ng/g EOM (top), ng/g carbon content
(middle) and ng/g d.w. (bottom).
36
4.2 PBDD concentration in Askö samples taken at different trophic levels
The presences of PBDDs were studied in the cyanobacteria (Aphanizomenon
flos-aquae), baltic clam, blue mussels, flounder and perch from Askö. The
results are, presented in Figure 4.2. It is concluded that PBDDs are present in all
species, with highest concentration in the cyanobacteria. The results are
presented on extractable organic matter, carbon content and dry weight to make
comparisons as good as possible. Irrespective of the manner of normalization,
the concentrations decrease with increasing trophic level.
4.3 Samples from New Zealand
Australian marine waters have high levels of NOCs in e.g. algae [55-57]. Thus,
it is likely that the New Zealand waters also have a high potential for natural
production. This was investigated in a food web study, including biota from the
south-west Pacific Ocean at New Zealand. The study was conducted as a
comparative study to the Baltic Sea location.
The food web study from New Zealand includes analysis of fish muscle and
liver tissue and filter feeders, i.e. diloma, green lip mussels and oysters. The
samples were freeze dried and ASE extracted with dichloromethane. The
sampling and extraction was done by the National Institute of Water and
Atmospheric Research in New Zealand before shipping the samples to Sweden.
On arrival, surrogate standards were added and the samples were partitioned
with aqueous potassium hydroxide. Further clean-up and analysis of OHPBDEs and MeO-PBDEs, PBDD was done as described in Chapter 3.
The results of OH-PBDE, MeO-PBDE and PBDD levels are presented in Figure
4.3. The concentrations of OH-PBDEs are similar in fish liver and in mussels
and oysters. The OH-PBDE conger patterns are however very different with 6OH-BDE47 and 2’-OH-BDE68 dominating in the fish liver, while the filter
feeders contain penta- and hexabrominated OH-PBDEs as well. The MeOPBDEs are present at similar levels and similar patterns in the fish, mussels and
oysters. The PBDD concentrations are much higher in the filter feeding species
than in the fish. The pattern though, is similar in fish compared to the mussel
and oyster samples. The diloma samples differ from the others for all the three
substance groups. It has very high levels of OH-PBDEs compared to the other
species and a different congener pattern of MeO-PBDEs and PBDDs compared
to the other samples.
37
25
700
ng/g d.w.
ng/g d.w.
20
15
10
5
6-OH-BDE123
400
6-OH-BDE99
300
6-OH-BDE90
200
6-OH-BDE85
2'-OH-BDE68
6-OH-BDE47
0
Fish
liver
ng/g d.w.
6-OH-BDE137
500
100
0
30
600
Fish Mussels Oyster
muscle
Diloma
71
25
6-MeO-BDE137
20
2-MeO-BDE123
6-MeO-BDE99
15
6-MeO-BDE90
10
6-MeO-BDE85
5
2'-MeO-BDE68
0
6-MeO-BDE47
Fish liver
Fish
muscle
Mussels
Oyster
Diloma
10
1600
ng/g d.w.
ng/g d.w.
8
6
4
1200
800
2
400
0
0
Fish liver
Fish
muscle
∑tetraBDDs
∑triBDDs
∑diBDDs
Mussels Oyster Diloma
Figure 4.3. OH-PBDEs (top), MeO-PBDEs (middle) and PBDDs (bottom) patterns and
levels in ng/g d.w. (± standard deviation) in New Zealand food web samples. Note the
different scales on the y-axis.
38
4.4 Herring and seal blood concentrations
Individual blood plasma samples from 21 herring from the Askö area from 2007
were analysed. The most abundant brominated compounds within each
substance group are shown in Figure 4.4. The detected OH-PBDEs and MeOPBDEs congeners are presented in Table 4.1. Figure 4.4 show high
concentrations of OH-PBDEs compared to neutral substances like PBDE, MeOPBDE and PCB. However, since the recovery of the surrogate standard in the
herring blood was low (chapter 3.8), it may be misleading with recovery
corrected data. The mean concentration of 6-OH-BDE47 is 320 ng/g l.w. when
no consideration to recovery is made. The concentration of this OH-PBDE is
however still as high as commonly found in e.g. blue mussels from the Baltic
Sea.
Twelve individual seal blood coagulate samples, collected from Baltic Sea grey
seals between 1995 and 2006, were also analysed. Phenolic compounds and
PCBs dominate the seal blood as depicted in Figure 4.5. A larger number of
OH-PBDE and MeO-PBDE congeners were detected in seal blood compared to
herring blood (Table 4.1).
Table 4.1. OH-PBDEs and MeO-PBDEs detected in herring and seal blood.
OH-PBDEs
MeO-PBDE
Herring*
Seal**
Herring*
Seal**
6-OH-BDE47
6-OH-BDE47
6-MeO-BDE47
6-MeO-BDE47
2'-OH-BDE68
2'-OH-BDE68
2'-MeO-BDE68
2'-MeO-BDE68
6-OH-BDE85
6-MeO-BDE85
6-OH-BDE90
6-OH-BDE99
6-MeO-BDE90
6-MeO-BDE90
6-OH-BDE99
6-MeO-BDE99
2-OH-BDE123
2-MeO-BDE123
6-OH-BDE137
6-MeO-BDE137
* Herring samples from Askö 2007, ** seal samples from the Baltic proper, 1995-2006
39
2500
ng/g l.w.
2000
1500
1000
500
CB
-1
53
-4
7
BD
E
6M
eO
-B
DE
47
DE
47
6OH
-B
2,
4,
6tri
BP
0
Figure 4.4. Concentrations (ng/g l.w.) of 2,4,6-triBP, 6-OH-BDE47, 6-MeO-BDE47, BDE47 and CB-153 in herring plasma.
600
500
ng/g l.w.
400
300
200
100
2,
4
53
CB
-1
,6
-
t ri
BD
E47
BP
6OH
-B
DE
47
6M
eO
-B
DE
47
0
Figure 4.5. Concentrations (ng/g l.w.) of 2,4,6-triBP, 6-OH-BDE47, 6-MeO-BDE47, BDE47 and CB-153 in grey seal blood coagulates.
40
5. Discussion
5.1 Data normalization
Concentrations of POPs in biota are usually presented on wet weight (w.w.)
[31,153] or lipid weight (l.w.) [62,154] basis, while POPs concentrations in
sediment are presented on dry weight basis (d.w.) and/or extractable organic
carbon (OC) [155,156]. The OC is considered a good normalization for
sediment since hydrophobic pollutants adsorbs to the organic carbon [156].
Bierman suggested that the OC corrected data in sediment corresponds to the
lipid weight in animals [157]. Organic carbon has been indicated to be the
optimal data normalization in cyanobacteria [158]. In the algae presented in
Paper IV an evaluation of the data normalized on wet weight, extractable
organic matter (EOM), dry weight and extractable organic carbon content was
conducted. Generally, the data was comparable on EOM, d.w. and OC basis.
However, the alga species have different cell structures compared to each other
and, thus there are some differences to be noted. For example, Furcellaria and
Fucus have a more robust and hard cell structure than the other species
analysed, i.e. Ceramium, Cladophora, Pilayella, Polysiphonia and
Enteromorpha. Furcellaria and Fucus contain much less water and EOM but
have a higher d.w. and OC content. This will affect the inter-species
comparison. In addition, the cell structure of Furcellaria and Fucus may be
better at adsorption of OHCs. In conclusion, I have chosen to present the algae
species comparisons on a OC basis.
5.2 Trends
5.2.1 Temporal variations
In Paper II the time trends of MeO-PBDEs and PBDDs are investigated in
perch from the Baltic proper (Kvädöfjärden). The temporal variations do not
indicate any clear trends, instead the levels fluctuate from year to year (Figure
5.1). The fluctuations in concentration over time may possibly relate to the
primary production, i.e. with phytoplankton and algae, and thus indirectly with
temperature. A weak, but not statistically significant, correlation was indicated
between the levels of MeO-PBDEs and PBDDs and the water temperature,
depth visibility and inorganic nutrient concentrations (Paper II). This may
imply a correlation with primary production.
41
10
9
8
sPBDDs
ng/g l.w.
7
6
5
4
3
2
1
0
120
100
sM eO-PBDEs
ng/g l.w.
80
60
40
20
05
20
03
04
20
20
01
02
20
20
99
00
20
19
97
98
19
19
95
96
19
19
93
94
19
19
91
92
19
19
19
90
0
Figure 5.1. Temporal variations of concentrations (ng/g EOM) of ∑PBDDs (top) and
∑MeO-PBDEs (bottom) in perch from Kvädöfjärden, sampled in the years 1990 to 2005
(Paper II).
5.2.2 Seasonal variations
Variation of OH-PBDEs, MeO-PBDEs and PBDDs are also seen during the
summer season. The variations in concentrations of OH-PBDEs and MeOPBDEs and the indication of seasonal variation of triBDDs, presented in Paper
III, show an increasing concentration of the OH-PBDEs, MeO-PBDEs and
PBDDs from May to June and a decrease in their concentration in August
(Figure 5.2).
800
sumOH-PBDE
sumMeO-PBDE
3500
triBDD
3000
700
600
2500
500
2000
400
1500
300
1000
200
500
100
0
Neutral comounds (ng/g l.w.)
Phenolic comounds (ng/g l.w.)
4000
0
May
June
August
October
Figure 5.2. Seasonal variation of ∑OH-PBDEs, ∑MeO-PBDEs and 1,3,7-/1,3,8-triBDD in
blue mussels sampled in 2008 from the Baltic proper (Askö). Note the different y-scales
for phenolic (left) and neutral (right) compounds (Paper III).
42
The seasonal variation indicated for 1,3,7-triBDD/1,3,8-triBDD in the blue
mussels (Paper III) has been validated by GC-HRMS analysis (Figure 5.3).
The estimated concentrations of triBDD as presented in Paper III were carried
out by GC-LRMS and were pseudo quantified against the 2’-MeO-BDE68. The
estimated level of triBDD was 230 ng/g EOM in the mussels sampled in June.
The corresponding levels determined by GC-HRMS (Figure 5.3) were 120 ng/g
EOM and 250 ng/g EOM for 1,3,7-triBDD and 1,3,8-triBDD respectively,
resulting in a total concentration of 370 ng/g EOM. It is notable that the
quantifications by GC-LRMS and GC-HRMS are highly comparable.
All the PBDDs show a seasonal variation and the extent of the variation
increases with the number of bromines in the molecule (monoBDDstetraBDDs). PBDFs were also found in the samples but in considerably lower
concentrations than the PBDDs (Figure 5.3). A seasonal variation was also
observed for the PBDFs particularly for the triBDFs and to a lesser extent (a
slight increase) for the tetraBDFs. No variation was observed for the
monoBDFs.
Figure 5.3. The seasonal variation in concentration (ng/g l.w.) of PBDFs (left) and
PBDDs (right) in blue mussels from Askö sampled in 2008. The total concentrations are
given in the graph. Note the different scales in the two graphs.
The seasonal variations of OH-PBDEs, MeO-PBDEs and PBDDs are based on
few data points as a result of bad weather during the sampling season. It is thus
not possible to establish whether the highest concentrations have been found.
A seasonal variation of OH-PBDEs, MeO-PBDEs were also found in algae
simultaneously sampled with the mussel samples at Askö (Paper IV).
Indications of elevated levels of PBDDs in June were also observed. The
seasonal variation observed for both red (Ceramium tenuicorne) and green
43
(Cladophora glomerata) macroalgae (Paper IV) are shown in Figure 5.4 and
follow the same seasonal variations as presented for the blue mussels (Paper
III). The levels correlate well between the algae and mussels (Paper IV).
500
∑OH-PBDEs
ng/g OC
400
300
200
100
2008-05-28
2008-06-25
2008-08-15
Furcellaria
lumbricalis
Cladophora
glomerata
Cladophora
glomerata
Cladophora
glomerata
Ceramium
tenuicorne
Pilayella
littoralis
Cladophora
glomerata
Ceramium
tenuicorne
0
2008-10-08
25
20
∑MeO-PBDEs
ng/g OC
15
10
5
2008-05-28
2008-06-25
2008-08-15
Furcellaria
lumbricalis
Cladophora
glomerata
Cladophora
glomerata
Cladophora
glomerata
Ceramium
tenuicorne
Pilayella
littoralis
Cladophora
glomerata
Ceramium
tenuicorne
0
2008-10-08
Figure 5.4. Seasonal variation in concentrations (ng/g OC) of ΣOH-PBDEs (top) and
ΣMeO-PBDEs (bottom) in macroalgae from Askö (Paper IV). The species analysed are
specified under the bars.
44
Both the temporal and seasonal trend studies indicate that the production of OHPBDEs, MeO-PBDEs and PBDDs vary. The production of these substances
seems to be highly correlated with temperature and thus the life-cycles of some
algae and/or cyanobacteria. The peak in the seasonal variation, together with the
concentrations found in the algae (Paper IV) seem to be mostly related to the
growth season of some green, red and brown algae [159]. This is indicating
Pilayella, Ceramium and/or Cladophora as the major producers of OH-PBDEs
and PBDDs. The large cyanobacteria blooms found in the Baltic proper are
usually most abundant in July and August [7].
5.2.3 Geographical distribution
Higher levels of OH-PBDEs, MeO-PBDEs and PBDDs are found in Baltic Sea
biota compared to biota from Swedish west coast waters (Paper I and IV,
Chapter 4.1), i.e. in algae, blue mussels and flounder. The levels of OHPBDEs, MeO-PBDEs and PBDDs in blue mussels presented in Paper I are
much higher in the Baltic proper compared to the west coast of Sweden (Figure
5.5). The difference in concentration between the Baltic proper and the west
coast is far greater than the small variations that can be seen for PCBs and
PBDEs in blue mussels from the same locations [160]. The levels of OHPBDEs, MeO-PBDEs and PBDDs may reflect the difference in algae or
cyanobacteria abundance or species composition in the two coastal waters. The
abundance of red filamentous macroalgae and cyanobacteria is higher in the
Baltic proper [6,7,161] as a result of eutrophication. It may also be a result of a
higher production of natural compounds in the Baltic. Paper IV shows the
presence of OH-PBDEs, MeO-PBDEs and PBDDs in all species with no
obvious differences between red, brown and green algae (autumn samples). In
addition, these compounds were observed to have lower concentration at the
west coast than the east coast in all species sampled, i.e. Cladophora,
Polysiphonia and Ceramium (Paper IV). Thus, the difference in concentrations
of OH-PBDEs, MeO-PBDEs and PBDDs found in the mussels (Paper I) and
algae (Paper IV) between the east and west coast of Sweden is more likely
related to a higher production of NOCs in the Baltic Sea.
The comparison of the levels of OH-PBDEs, MeO-PBDEs and PBDDs in the
Baltic Sea to the worldwide geographic distribution is hard to assess. Very few
studies reports data of OH-PBDEs, MeO-PBDEs and PBDDs in the same
sample (Paper I, Paper IV, [20]). There are also very few studies of these
compounds in low trophic level organisms [59,60,79,87].
45
1000
sOH-PBDEs
sMeO-PBDEs
sPBDDs
900
800
ng/g l.w.
700
600
500
400
300
200
100
0
Baltic proper
West coast
Figure 5.5. Concentrations (ng/g l.w.) of OH-PBDEs, MeO-PBDEs and PBDDs in blue
mussels from the Baltic proper and the west coast of Sweden (Fladen and Väderöarna)
(Paper I).
OH-PBDEs have been analysed in mussels from Hudson bay, but were not
detected [60]. OH-PBDEs have also been reported in fish blood from the
Detroit river, showing concentration in the low pg/g w.w. [30]. The
concentrations of OH-PBDEs in herring plasma from the Baltic Sea (chapter
4.4.) are approximately two orders of magnitude higher than the reported
concentration from Detroit River. Routti et al. report ∑OH-PBDE in ringed seal
blood from the Baltic Sea and Svalbard with a factor 2.5 higher concentrations
in the Baltic [31]. The concentrations in the ringed seal from the Baltic [31]
were three times lower than detected in the grey seal blood in this thesis
(chapter 4.4).
Similar or lower levels of MeO-PBDEs than in the Baltic Sea have been
reported, at comparable trophic levels elsewhere, e.g. in mussels from Hudson
bay and Liaodong bay (China) and in fish from Liaodong bay [59,60].
Haglund and co-workers s have reported PBDDs in bivalve and fish from the
Baltic Sea and the west coast of Sweden, generally showing higher PBDD
concentrations in the Baltic [79].
The high levels of PBDDs found in filter feeders from New Zealand (Chapter
4.3) are much higher than what is found in blue mussels from the Baltic region
46
(Paper I). The levels of OH-PBDEs and MeO-PBDEs in the New Zealand
mussel samples (Figure 4.3) are similar, while the blue mussels from the Baltic
Sea are dominated by the MeO-PBDEs (Figure 5.5). The data in Chapter 4.3
together with the literature cited indicates that the world-wide distribution of
these compounds varies. The differences in composition probably reflect
differences in producers between areas. Far more research efforts need to be
done in this area, not the least to expand the assessments to all oceans on the
globe.
5.3 Food web distribution
Brown algae (Fucus vesiculosus and Dictyosiphon foenicolaceus) (Paper I,
IV), blue mussels (Paper I), flounder (muscle) (Chapter 4.1) and perch
(muscle) (Paper II) from Kvädöfjärden in the Baltic Sea were compared for
their content of OH-PBDEs, MeO-PBDEs and PBDDs. The analyses were done
according to the methods described in Paper I and IV. The results of this interspecies comparison are depicted in Figure 5.6. The levels of OH-PBDEs are
highest in algae, followed by the filter feeding blue mussel. The pattern is
reversed for the MeO-PBDEs and PBDDs, showing bioaccumulation to the
filter feeding blue mussel. The fish species however, contain low concentrations
of both MeO-PBDEs and PBDDs. The presence of PBDDs were further studied
in the cyanobacteria, Aphanizomenon flos-aquae, baltic clam, blue mussels,
flounder and perch from Askö, presented in Figure 4.2. The study showed
presence of PBDDs in all species, with highest concentration in the
cyanobacteria. Both comparisons conclude that although OH-PBDEs, MeOPBDEs and PBDDs are bioaccumulative, but they do not seem to biomagnify.
The rapid decrease in concentrations of the analytes found in blue mussels
(Paper III and Figure 5.3) supports the limited retention of these compounds.
Also, Paper II shows variations in retention of PBDDs and MeO-PBDEs in
perch, possibly partially explained by the metabolic stability or discriminations
in uptake of higher brominated congeners.
The bioaccumulation of MeO-PBDEs have been studied in e.g. blue fin tuna,
harbour seals, harbour porpoises, ringed seal and polar bears [32,63,64] and a
few studies describe the biomagnification potential [32,64,162]. The trophic
magnification factor (TMF) in a marine food web study from Australia
indicated biomagnification, but the TMF was lower for MeO-PBDEs than for
PBDEs [162]. Weijs et al. found the biomagnification factor (BMF) for 2′MeO-BDE 68 and 6-MeO-BDE 47 to vary between 0.1 and 5 and 0.1 and 23,
respectively [64]. Letcher and co-workers reported bioaccumulation in polar
bears, but no biomagnification from ringed seal [32].
47
1000
900
800
6-MeO-BDE137
2-MeO-BDE123
6-MeO-BDE99
ng/g EOM
700
600
500
6-MeO-BDE90
6-MeO-BDE85
2'-MeO-BDE68
400
300
6-MeO-BDE47
200
100
0
Fucus
vesiculosus
Dictosiphon
foenicolaceus
blue mussel
perch
flounder
2500
ng/g EOM
2000
6-OH-BDE137
2-OH-BDE123
6-OH-BDE99
1500
6-OH-BDE90
6-OH-BDE85
2'-OH-BDE68
1000
6-OH-BDE47
500
0
Fucus
vesiculosus
Dictosiphon
foenicolaceus
blue mussel
n.a.
n.a.
perch
flounder
180
160
ng/g EOM
140
ng/g EOM
120
100
80
60
1,2,3,8-tetraDD
0,20
0,18
0,16
0,14
0,12
0,10
0,08
0,06
0,04
0,02
0,00
40
1,2,3,7-tetraDD
1,2,4,7/1,2,4,8-tetraDD
1,3,7,9-tetraDD
1,3,6,8-tetraDD
237-triDD
1,4,7-triDD
1,3,8-triDD
1,3,7-triDD
1,8-diDD
perch
flounder
20
0
n.a.
Fucus
vesiculosus
Dictosiphon
foenicolaceus
blue mussel
perch
1,7-diDD
2,7/2,8-diDD
1,3-diDD
flounder
Figure 5.6. MeO-PBDEs (top), OH-PBDEs (middle) and PBDDs (bottom) patterns and
levels in ng/g EOM (± Standard deviation) in biota from Kvädöfjärden. Note the different
scales on the y-axis.
48
Halogenated phenolic compounds are primarily associated with wildlife and
human blood and not with muscle or lipid tissue. A comparison is made, and
shown in Figure 5.7, between algae, blue mussels, herring plasma and seal
blood (Chapter 4.4) from the Baltic proper. All species are sampled in the
waters around Askö, except for the grey seals. The herring plasma is shown
both as recovery corrected data and as non-recovery corrected data. The
retention of OH-PBDEs in herring and seal blood is notably high. The levels of
6-OH-BDE47 are much higher in the blood compare to the algae (Ceramium
tenuicorne) and mussel sample. However, when comparing the total OH-PBDE
concentration the levels are similar. This is due to a different congener pattern
in the algae samples. In the herring and seal blood, 6-OH-BDE47 is the
dominant congener, while the algae contain several congeners and mostly 6OH-BDE137. It should be noted that the algae sample was not taken the same
year as the mussel and herring blood. Also, the seal blood was taken from seals,
sampled 1995-2006, individuals collected from different locations in the Baltic
proper.
2500
6-OH-BDE47
ng/ g EOM
2000
∑OH-PBDE
1500
1000
500
0
2008-05-28
2007-05-09
2007-05-09
2007-05-09
Ceramium
tenuicorne
Blue mussel
Herring blood
Recovery
corrected
Herring blood
Not Recovery
corrected
Seal blood
Figure 5.7. Comparison of OH-PBDEs (ng/g EOM ± Stddev) in four Baltic Sea species,
sampled around Askö. Note that the time of sampling differs.
5.4. Exposure and uptake
The route of exposure of OHCs for the mussels, fish and seal presented herein
are direct uptake from the water via the gills, or by their diet. The difference
observed in congener pattern in e.g. the herring blood compared to mussels and
seal blood may indicate their route of exposure via the gills. Considering the
pKa values (Table 2.2) at least 6-OH-BDE137 and 6-OH-BDE99 are predicted
to be ionised at natural pH, and thus not likely to be taken up via the gills.
49
For the MeO-PBDEs, exposure via diet is more likely. In the perch (Paper II),
two of the congeners were not detected; 2-MeO-BDE123 and 6-MeO-BDE137.
This may possibly be explained by debromination, as seen both in vitro and in
vivo for the PBDEs in fish [163-166]. Indications of debromination processes
were also found in the seasonal variation study (Paper III). The higher
brominated MeO-PBDE congeners have a more rapid decline according to this
study, while the lower brominated, i.e. 6-MeO-BDE47 and 2’-MeO-BDE68, are
stable in their concentrations and do not decrease from June and onwards over
the sampling period. This may possibly be explained by debromination of hexaand pentabrominated methoxylated diphenylethers leading to e.g. 6-MeOBDE47 and 2’-MeO-BDE68.
5.5 Origin
It is evident, based on the high levels of OH-PBDEs, MeO-PBDEs and PBDDs
found in the Baltic Sea biota, that these compounds are preferentially natural
products. As presented herein (Paper I, III and IV), the levels of OH-PBDEs
and MeO-PBDEs, in the Baltic proper biota, far exceed that of the possible
metabolic or abiotic transformation precursors, PBDEs (Paper III and IV)
[160]. The variation in concentration of these substances (Paper II, III and IV)
gives further support for natural formation thereof. The difference in PBDF and
PBDD concentrations (Paper I and IV and Figure 5.3) also support natural
formation of PBDDs.
The possible producers suggested within this thesis and by Malmärn [17] are
firsthand filamentous macroalgae and/or cyanobacteria. The high levels of OHPBDEs and PBDDs in both algae (Paper I and IV, [20,21]) and in
cyanobacteria (Paper I, [20]) from the Baltic Sea support the production of
these compounds. As discussed above the Pilayella, Ceramium and/or
Cladophora (chapter 5.2.2) seem to be the most likely producers of OH-PBDEs
and PBDDs.
The producers of MeO-PBDEs however, are not as easily deduced. Although
the pattern is similar in algae and mussels (Paper IV), the levels found in algae
and cyanobacteria are fairly low. One possible explanation may be that the
MeO-PBDEs are methylated by bacteria outside the algae itself, as described for
other phenolic compounds [49-54]. It would thus be of interest to study algae
free from microorganisms.
No single species of algae has been determined as a major producer of the
compounds discussed herein. The difference in concentration within the same
species from the same locations again indicates that the life-cycle is important.
50
Also, studies have shown that algae under stress produce higher levels of other
brominated compounds such as PBPs [167-169]. The stress may for example be
a result of grazing or ecological changes leading to reduction or increase in sun
light [170], e.g. by change in water level.
5.6 Ecological perspective
Several worrying effects are reported for Baltic Sea biota, e.g. the large-scale
changes in biodiversity [2], decreases in body weight and/or blubber thickness
in seal [112] and Baltic herring [171], a high mortality in fish eggs [172], as
well as a massive bird death attributed to a neurological disease like thiamine
deficiency [173] and the thiamine deficiency in salmon called M74 [174], where
the salmon fry only live a few days. In addition, the constant chemical exposure
of both anthropogenic and natural compounds may add to the already stressed
ecosystem.
Although the substances discussed within this thesis do not seem to biomagnify,
the levels may still reach high levels e.g. during summer time. Animals feeding
on mussels or algae may be highly exposed to chemicals with toxic effects i.e.
the OH-PBDEs and the PBDDs during these periods. Eider duck, long-tailed
duck and flounder largely feed on blue mussels in the Baltic Sea. Their daily
exposure may be considerable resulting in potential ecotoxicological effects.
51
6. Future perspectives
The high levels of the polybrominated chemicals discussed in this thesis seem to
be linked to primary producers, such as algae and cyanobacteria, in the Baltic
Sea. Further, the occurrence and levels of these brominated chemicals may be
affected by Eutrophication. Accordingly it would be of interest to look closer
into other marine and freshwater ecosystems with a similar degree of
eutrophication as the Baltic Sea, or worse. Such environments may be e.g. the
Black Sea, the North Sea and Wadden Sea, Chesapeake Bay and the northern
Gulf of Mexico and Taihu Lake, close to Shanghai.
PBDD concentrations are presented in this thesis for two cyanobacteria species,
Aphanisomenon and Nodularia. The PBDD levels in the Aphanisomenon were
high, while the Nodularia was substantially lower. It is obviously impossible to
determine if this is a result of the different species, sampling location or time of
sampling. Studies including analysis of several samples of the two species,
sampled in the proximity of each other, will throw further light on formation
and sources of the PBDDs. In addition, there are still a few species of primary
producers that has yet to be analysed, e.g. diatoms and dinoflagellates. Further
research is required to lay out a more complete picture of PBDD, as well as OHPBDE and MeO-PBDE producers.
To enable the determination of algal producers and the route of exposure, water
samples must most likely be studied, preferably from locations close to algae
growth. Possibly this can be done under laboratory conditions. Also, studies
correlating production of natural OHCs and stressors like grazing is required.
Lastly, there is need of (eco)toxicity studies of foremost OH-PBDEs and
PBDDs. If possible correlations between observed effects in Baltic wildlife and
exposure levels to these compounds should be prioritized, still not omitting the
potential impact anthropogenic chemicals may have.
This thesis is stressing the fact that natural halogenated products play a role in
the Baltic Sea ecosystems.
52
7. Acknowledgements
Först vill jag ge ett jättetack till mina två handledare. Lillemor, du är helt
fantastisk. En mer förstående och hjälpsam handledare får man leta efter, vare
sig det gäller bebisar, analysmetoder eller handgriplig provtagning i fält. Åke,
tack för att du kämpade och fann en plats till mig på miljökemi. Jag kan inte ens
uttrycka hur kul jag har haft det under åren. Tack för allt stöd och hjälp, inte
minst med avhandlingen.
Tack till mina medförfattare, Peter, Dennis, Sören, Lena, Anders och Anna
M för allt kunnande och fantastisk hjälp, and Xitao - thank you for your kind
contributions.
Anita – du har varit en klippa när det gäller allt krångligt som ekonomi och
blanketter och har alltid tid att byta ett par ord.
Hrönn, examensarbetshandledare, rumskompis och vän – jag saknar dig och
fåglarna fortfarande. Linda, du är en alldeles speciell vän och reskamrat, jag
kommer saknar dig, Anna S, we’ll always have Canada, Johan F, tack för att
du alltid ställer upp vare sig det är på lab eller med ett skratt, Jessica, bästa
rumskompisen både borta och hemma. Emelie, tack för alla pratstunder, allt
godis och uppmuntran, Ioannis, vår underbara GC-guru vad du har fått slita.
Anna-Karin, du har ett fantastiskt humör och sätt – lycka till. Maria A, jag
saknar dig, Anna V och Hans, de må ha varit kortvarigt men gött, Hitesh, your
next. Lotta, Per, Margareta, Birgit, Maria S, Lisa, Andreas, Göran, Johan
E tack för alla trevliga stunder och till de ”nya” ansiktena; Cecilia och Dennis
lycka till. Yin, thank you for your good work and good luck with your PhD. Ett
stort tack även till alla gamla miljökemister, ingen nämnd ingen glömd, för att
ni gjort detta till en helt fantastisk arbetsmiljö.
Jag är lyckligt lottad att ha så bra vänner som stått vid min sida i vått och torrt
under alla år. Ni har alltid funnits där även när tiden tröt, Jenny, bästaste vänner
4-ever, Anna O, tack för alla tokigheter vi har gjort, Sara, underbara, Lina, du
som förstår, och alla ni andra som gjort dessa år lättare och så njutbara, Linda,
Jocke, Andreas, Susanne, Brian, Kajsa, Sandra, Helene, m.fl.
Jag vill även tacka min nya och min gamla familj. Tack, Marie, Jonas och
Marie för allt stöd och hjälp, bättre svärföräldrar finns inte, tack till alla mina
svågrar/svägerskor och barn (ingen nämn ingen glömd) för att ni förgyllt mitt
liv. Micke, du har betytt så mycket för mig under min uppväxt ja hela mitt liv.
En bättre storebror går inte att få, som till och med hjälp till med min
provtagning. Madelene och Sanny, för att ni finns. Mormor Eivor, tack för att
du hållit mitt hushåll ajour och för din underbara personlighet, jag har saknat
våra samtal över en fika.
53
Mamma och pappa, tack för allt stöd genom åren, inte minst nu under dessa
konstiga månader. Ni är underbara.
Stefan, min kärlek till dig kan inte vara större. Tack för att du stått ut med mig
under dessa år med övertid och stress och alltid mött mig med ett leende och
mat på bordet. Ser fram emot att kunna återgälda allt. Nu väntar andra äventyr...
This thesis was financially supported by the Swedish environmental protection
agency through the Swedish environmental monitoring progam on
contaminants, and by the Swedish reseasrch counsil FORMAS. Financial
support was also received from the Stockholm University's strategic marine
environmental research funds through the Baltic ecosystem adaptive
management (BEAM) program and from Ångpanneföreningen (ÅF). A grant
from the Stockholm University marine research center (SMF) have been recived
for sampling at Askö.
54
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