Micro solid-phase extraction for the analysis of per- and polyfluoroalkyl substances in environmental waters

Journal of Chromatography A 1604 (2019) 460495

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Journal of Chromatography A journal homepage: www.elsevier.com/locate/chroma

Micro solid-phase extraction for the analysis of per- and polyfluoroalkyl substances in environmental waters Thomas E. Lockwood a, Mohammad Talebi b, Andrew Minett c, Simon Mills b, Philip A. Doble a, David P. Bishop a,∗ a b c

Hyphenated Mass Spectrometry Laboratory (HyMaS), University of Technology Sydney, Broadway, New South Wales, 2007, Australia Envirolab Services Pty Ltd, Chatswood, New South Wales, 2067, Australia ePrep Pty Ltd, Mulgrave, Victoria, 3170, Australia

a r t i c l e

i n f o

Article history: Received 12 June 2019 Revised 23 August 2019 Accepted 27 August 2019 Available online 28 August 2019 Keywords: Micro-SPE PFAS Automation

a b s t r a c t Growing concern over the environmental and health impacts of per- and polyfluoroalkyl substances (PFASs) has led to the development of increasingly stringent regulatory guidelines. To meet these guidelines for the determination of PFASs in surface-water, solid-phase extraction (SPE) is commonly implemented for clean-up and pre-concentration of samples. In this paper a micro-SPE method for the cleanup and pre-concentration of PFASs from surface-water was developed. A micro-SPE packing phase was created to retain 13 long and short chain PFAS after examining combinations of four 3 μm particle size sorbents, with the optimal phase consisting of a 50:50 mixture of C18 and aminopropyl silica. Micro-SPE achieved similar results to conventional SPE methods while reducing sample preparation time to 5 min and using only 2 mL of sample. The method was validated using spiked recoveries (100 ng L−1 ) from PFAS contaminated surface-water samples with recoveries ranging from 86% to 111% and relative standard deviations below 18%. Concentrations of the PFASs in the samples ranged from below the limit of quantification to 898 ± 15 ng L−1 . Automation of sample preparation, including the micro-SPE extraction, was also demonstrated. These results show the potential for automated micro-SPE to replace conventional SPE, with the decreases in sample preparation time, sample and solvent volumes crucial for incorporation into routine analyses in commercial laboratories. © 2019 Elsevier B.V. All rights reserved.

1. Introduction Since their introduction in the late 1940s, the development and use of per- and polyfluoroalkyl substances (PFASs) has grown significantly. PFASs are a family of synthetic fluorine containing chemicals that consist of a fluorinated hydrocarbon chain bonded to a charged group. The perfluoroalkyl moiety (Cn F2n+1 ) imparts several properties that make PFASs more desirable than their hydrocarbon counterparts [1], such as a very low surface tension, hydro- and lipophobic properties, and both thermal and chemical stability due to the large number and strength of C–F bonds [2]. These properties led to the development of many different PFASs for a wide variety of commercial and industrial applications such as carpeting, non-stick cookware and fire-fighting foams [1]. Following emission, PFASs enter the local environment and contaminate soil and water sources due to high adsorption and water solubility. They are then dispersed throughout the environ-

∗

Corresponding author. E-mail address: [email protected] (D.P. Bishop).

https://doi.org/10.1016/j.chroma.2019.460495 0021-9673/© 2019 Elsevier B.V. All rights reserved.

ment through the air and migration of surface and groundwater [3]. PFASs are detected globally, even in very remote regions such as Arctic glaciers [4,5]. A number of sites in Australia are contaminated by PFASs due to the use of aqueous film-forming foams (AFFFs). In 2004 Australia ratified the Stockholm Convention on Persistent Organic Pollutants, however it has not ratified the listing of perfluorooctane sulfonate (PFOS) or any other types of PFASs, and there are no nationwide restrictions on their usage [6]. The continued use and persistent nature of these chemicals has led to concerns over the health effects of consuming water or food sourced from near these sites [7]. Animal studies, primarily performed in rodents, have shown links between PFOS and perfluorooctanoic acid (PFOA) exposure and increases in liver weight [8], behavioural and developmental changes in offspring [9,10], negative reproductive effects [11] and tumour growth [12]. Large scale studies of human exposure to PFASs by the C8 Science Panel have also highlighted possible links between PFOS and PFOA exposure and high cholesterol [13], thyroid disease [14], autoimmune disease [15] as well as testicular and kidney cancers [16]. In Australia, Victorian firefighters exposed to

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T.E. Lockwood, M. Talebi and A. Minett et al. / Journal of Chromatography A 1604 (2019) 460495

high amounts of AFFFs showed a significant increase in cancer risk [17]. In response to these concerns, the first Australian health guidance values for PFAS intake were described in June of 2016 [18,19]. In addition to recommending the total daily intake (TDI) values previously reported by the European Food Safety Authority [20], the report recommended that Food Standards Australia New Zealand (FSANZ) undertake an assessment of the toxicity of PFOS, PFOA and perfluorohexane sulfonate (PFHxS). The FSANZ report was completed in April 2017 and proposed that the TDI values be revised down from 150 and 1500 ng kg−1 d−1 to 20 and 160 ng kg−1 d−1 (70 and 560 ng L−1 in drinking water) for PFOS and PFOA [7]. Solid phase extraction (SPE) is the preferred technique for the clean-up and pre-concentration of PFAS contaminated water samples [21]. While effective, SPE is labour and time intensive, and large sample volumes are required to achieve regulatory LODs. Micro-SPE (μSPE) has been proposed as an improved technique for the extraction of PFAS from surface waters. Performing SPE under a high pressure allows the use of smaller sorbent particle sizes (<5 μm), increasing extraction efficiency and thus reducing the required sample volumes [22]. μSPE has already been demonstrated in the extraction of perfluoro carboxylic acids from complex matrices such as fish tissue (recoveries between 77 and 120%) [23] and human plasma (recoveries between 87% and 102%) [24]. The ePrep® Sample Preparation Workstation can perform automated μSPE as well as other sample preparation steps such as sample dilutions and transfers. It has been used to extract phenolic compounds in tea by Porto-Figueira et al. [25] with high precision and recoveries. Here we demonstrate the use of μSPE for the extraction of PFASs from surface water samples with LC–MS/MS analysis. The performance of the automated sample preparation platform was evaluated using the optimised μSPE method.

as much as possible. In this study PFAS-free polypropylene and polyethylene containers, lids and vial septa were used for all reagents, standards and samples. 2.1. Sample preparation Prior to μSPE extraction, 10 mL aliquots of the supplied surface water samples were acidified to approximately pH 3 with 100 μL of glacial acetic acid to aid in the retention of short chain PFASs. Samples were then spiked with 100 μL of the isotopically-enriched internal standards to produce a final concentration of 100 ng L−1 . Clean-up and extraction of the samples was performed on the eXact3 digital syringe and ePrep® sample preparation workstation using ePrep® μSPEed cartridges packed with a newly developed mixed-mode C18:aminopropyl silica (APS) phase (ePrep; Melbourne, Australia – now marketed as PFAS cartridges), shown in Fig. 1. A one-way check valve in the μSPEed cartridges allows volumes greater than the full syringe to be preconcentrated by repeatedly aspirating and dispensing. The sample preparation workflow developed in this study for these cartridges is shown in Table 1. The extract was then diluted with 100 μL of 0.1% acetic acid to improve peak shapes. Cross contamination was minimised by rinsing the syringe with a full volume (500 μL) of methanol between extractions.

2. Materials and methods Poly- and perfluoroalkyl carboxylate and sulfonate native and isotope-labelled internal standard mixtures (2 mg L−1 ) were purchased from Wellington Laboratories (Ontario, Canada). The native mixture contained the perfluoro- butanoic (PFBA), pentanoic (PFPeA), hexanoic (PFHxA), heptanoic (PFHpA), octanoic (PFOA), nonanoic (PFNA), decanoic (PFDA), undecanoic (PFUnA), dodecanoic (PFDoA) and tetradecanoic (PFTeDA) acids as well as the perfluoro- butane (PFBS), hexane (PFHxS) and octane (PFOS) sulfonates. The Structures of these compounds can be found in Supplementary Table 1. The isotope-labelled mixture consisted of [13 C4 ]PFBA, [13 C5 ]PFPeA, [1,2,3,4,6-13 C5 ]PFHxA, [1,2,3,4-13 C4 ]PFHpA, [13 C8 ]PFOA, [13 C9 ]PFNA, [1,2,3,4,5,6-13 C6 ]PFDA, [1,2,3,4,5,6,7-13 C7 ]PFUnA, [1,2-13 C2 ]PFDoA, [1,2-13 C2 ]PFTeDA, [2,3,413 C ]PFBS, [1,2,3-13 C ]PFHxS and [13 C ]PFOS. 3 3 8 Standards and stocks were prepared using LC–MS grade methanol (Merck; Darmstart, Germany) and ultra-pure water obtained from an Arium® pro water generator (Sartorius; Dandenong, Australia). Acetonitrile and isopropyl-alcohol used in the needle wash were of LC–MS grade and obtained from Honeywell via Chem-Supply (Port Adelaide, Australia). Glacial acetic acid was obtained from VWR (Tingalpa, Australia), and analytical grade sodium hydroxide and ammonium acetate were purchased from Sigma-Aldrich (Castle Hill, Australia). Samples of PFAS contaminated surface waters were provided by Envirolab Services (Chatswood, NSW, Australia) and stored in polypropylene falcon tubes at 4 °C until use. Contamination is a common issue that arises during the analysis of PFASs as they are used as polymerisation aids in the manufacture of polytetrafluoroethylene (PTFE) and other fluoropolymers. To mitigate these factors, fluoropolymers should be avoided

Fig. 1. A diagram of the ePrep® μSPEed cartridge. A one-way check valve in the cartridges allows liquids to bypass the sorbent bed during aspiration. The aspiration and dispense paths are indicated by the blue and red arrows respectively.

T.E. Lockwood, M. Talebi and A. Minett et al. / Journal of Chromatography A 1604 (2019) 460495

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Table 1 Extraction procedure for the eXact3 digital syringe.

Condition Equilibration Activation Load Wash Elute

Solvent

Volume (μl)

Dispense speed (μl min−1 )

10 mM NaOH in methanol Methanol 1% Acetic acid Acidified Sample Ultra-pure water 10 mM NaOH in methanol

250 250 250 2000 100 100

540 1080 1080 1080 1080 540

2.2. UHPLC Gradient separation of the 13 PFAS compounds (Fig. 2) and their respective internal standards was performed on a Shimadzu (Rydalmere, Australia) Nexera MP UHPLC using a Phenomonex Luna Omega 2.1 × 50 mm, 1.6 μm C18 column. Mobile phases consisted of ultra-pure water (A) and methanol (B), each with 2 mM ammonium acetate. Initial conditions of 20% B were held for 0.2 min before being raised to 70% at 2.4 min, then 95% at 5 min. The gradient was then held at 95% for 2 min before returning to the initial conditions and equilibrated for 4 min. A flow rate of 0.6 mL min−1 and column temperature of 50 °C were used throughout the run. An injection volume of 5 μL was used. A needle wash was performed to prevent carry over of strongly adsorbing compounds. The exterior of the needle was washed before and after each injection with 500 μL of the strong wash (1:1:1:1 of water, acetonitrile, methanol and isopropyl alcohol), and an internal wash of 500 μL of the strong wash then 500 μL of 20% methanol was performed during each run. An ACQUITY® PFC isolator column (Waters; Rydalmere, Australia) was installed between the solvent mixer and autosampler to mitigate PFAS contamination from the PTFE HPLC solvent lines. This column retains contaminants from the HPLC solvent lines, allowing any PFAS contamination originating from the system to elute after the analytes to prevent interference during the analysis.

Analysis of the automated extractions was performed at Envirolab, a commercial environmental testing laboratory, using a Shimadzu Shim-pack XR-ODSIII 2.0 × 50 mm, 1.6 μm column and Shimadzu Nexera X2. A flow rate, column temperature and injection volume of 0.4 mL min−1 , 40 °C and 40 μL were used respectively. The gradient separation detailed in Supplementary Table 3 was followed, again using a trap column (Shimadzu Shim-pack XR-ODS 3.0 × 30 mm, 2.2 μm) to retain any PFAS contaminants. 2.3. Mass spectrometry Detection was performed using a Shimadzu LCMS-8060 triple quadrupole mass spectrometer operated in negative ionisation and multiple reaction monitoring (MRM) modes. The interface voltage and temperature were optimised to −0.5 kV and 300 °C. Nebulising, heating and drying gas flows were set at 3, 10 and 10 L min−1 respectively, and collision gas was operated at a pressure of 270 kPa. MRM parameters (Supplementary Table 2) were optimised at −0.5 kV for transitions from the [M-H]− ion using the LabSolutions software. Where possible, both quantification and qualification ions were used for each compound, however some compounds, such as PFBA, exhibited only one significant fragment. Analysis at Envirolab was performed on a Shimadzu LCMS8050 with nebulising, heating and drying gas flows set at 1.5, 5 and 5 L min−1 respectively. The interface was operated in negative

Fig. 2. UHPLC separation of the perfluoroalkyl carboxylates and -sulfonates: PFBA (1), PFPeA (2), PFBS (3), PFHxA (4), PFHpA (5), PFHxS (6), PFOA (7), PFNA (8), PFOS (9), PFDA (10), PFUnA (11), PFDoA (12), PFTeDA (13). Mass labelled isotopes had identical retention times to their respective analytes and are therefore not shown.

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T.E. Lockwood, M. Talebi and A. Minett et al. / Journal of Chromatography A 1604 (2019) 460495 Table 2 Validation results for the instrumenta and extractionb methods (n = 6).

PFBA PFPeA PFHxA PFHpA PFOA PFNA PFDA PFUnA PFDoA PFTeDA PFBS PFHxS PFOS ∗

(n = 5),

∗∗

Linearitya

LOD (ng L−1 )a

%Recoveryb

%RSDb

0.9990 0.9998 0.9999 0.9999 0.9998 0.9998 0.9997 0.9995 0.9993 0.9992 0.9998 0.9997 0.9995

3.5 2.8 5.2 1.2 1.7 2.2 1.7 4.4 4.8 6.6 0.29 1.3 2.7

∗∗

15 10 12 3 8 7 18 12 3 6 4 12 12

101 98 101 106 97 107 96 ∗ 104 ∗ 103 ∗∗ 86 105 105 ∗ 111

(n = 4) Outliers removed after failed Grubbs tests.

ionisation mode at 300 °C. MRM parameters are detailed in Supplementary Table 4. 3. Results and discussion The LC–MS gradient separation of the 13 PFAS compounds in Fig. 2 was developed by repeated injections of a 100 ng L−1 mixed standard. Gradient parameters were optimised until acceptable resolution was obtained for all compounds, ensuring separation of isobaric fragments. A seven-point calibration curve ranging from 10 to 9 0 0 0 ng L−1 was constructed and used to determine the linearity and instrumental limits of detection (LODs) for the analytes (Table 2). The Shimadzu LabSolutions software was then used to construct a 1/C weighted calibration curve and to calculate LODs (3.3 times signal to rms noise) from the lowest concentration standard. A weighted calibration curve was chosen due to the large range of PFAS concentrations expected to be present in the samples. The μSPE packing phase was optimised by comparing the efficiency of C18, APS, pentafluorophenyl (PFP) and diol sorbent mixtures in extracting the analytes from ultra-pure water (Fig. 3). These sorbents were chosen based on their potential efficiencies and availability in the 3 μm particle size that provides the in-

Table 3 Recoveries of the isotopically enriched PFAS internal standards from spiked water. Results are reported as the mean recovery and standard deviation (n = 3). Internal Standard 13

[ C4 ]PFBA [13 C5 ]PFPeA [1,2,3,4,6-13 C5 ]PFHxA [1,2,3,4-13 C4 ]PFHpA [13 C8 ]PFOA [13 C9 ]PFNA [1,2,3,4,5,6-13 C6 ]PFDA [1,2,3,4,5,6,7-13 C7 ]PFUnA [1,2-13 C2 ]PFDoA [1,2-13 C2 ]PFTeDA [2,3,4-13 C3 ]PFBS [1,2,3-13 C3 ]PFHxS [13 C8 ]PFOS

%Recovery

%SD

50 110 123 121 107 95 75 49 28 30 102 110 78

3 1 1 1 1 2 3 1 2 5 3 4 4

creased extraction efficiency of μSPE. Reversed-phase (C18) and weak-anion exchange sorbents (APS) are well established for use in PFAS analysis [26], and both the PFP and diol sorbents were chosen for their potentially higher selectivity for the fluorinated, polar analytes [27]. The diol sorbent was unable to retain any of the compounds and the PFP phase had poor recoveries of both the short and longer chain compounds. C18 provided good retention for most compounds however APS was the only phase able to retain the short chain compound PFBA. Therefore a combination of reverse-phase C18 sorbent and APS was required for efficient extraction of both long and short chain compounds with the optimal packing phase of the μSPE cartridges determined to be a 50:50 mixture of C18:APS. Conditions for the extraction were optimised using recoveries of isotopically labelled standards from spiked ultra-pure water (Table 3). Due to the high pKa (9) of APS a strongly basic solvent such as 10 mM NaOH in methanol (∼pH 12) was required as the μSPE elution solvent to neutralise the phase and quantitatively elute analytes. Use of a high pH solvent may degrade the stationary phase in the cartridges and will limit their potential reusability, therefore for this application the cartridges were used once and discarded. Acidification of samples and sorbent during conditioning

Fig. 3. Normalised recoveries of six analytes from ultra-pure water when extracted using four different packing sorbents. A 50:50 combination of APS and C18 was found to be the most effective sorbent, providing good retention of both short and long chain compounds.

T.E. Lockwood, M. Talebi and A. Minett et al. / Journal of Chromatography A 1604 (2019) 460495

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Fig. 4. Chromatogram of sample E, perfluoroalkyl carboxylates and sulfonates are in blue and orange, respectively. Peak shouldering is visible for PFHxA (4) and PFOS (9) due to the presence of both linear and branched isomers. Peaks are labelled as in Fig. 2.

The lower recoveries observed for PFTeDA, [1,2-13 C2 ]PFTeDA, and [1,2-13 C2 ]PFDoA was hypothesised to be caused by the adsorption of the compound to both the sample container and the extraction equipment, in particular the glass syringe barrel [30]. Adsorption of analytes to sample containers is typically minimised by using the full sample volume and then rinsing the container with the elution solvent [28]. The small elution volumes used in the μSPE method make this impractical. Adsorption to the extraction equipment could be minimised by using larger elution volumes, however this would decrease the level of pre-concentration possible using the same sample volume. Use of a polypropylene syringe barrel would also reduce adsorption. Another possibility for the low recovery is strong retention of the compound by the sorbent, preventing quantitative elution. Further optimisation of the sorbent phases or sorbent mixture ratios could minimise these losses. The potential for automation of the extraction method was demonstrated on the ePrep® sample preparation workstation. A sample of surface water was spiked with 23 PFASs and masslabelled internal standards as follows: PFHxA, PFHpA, PFOA, PFNA, PFDA, PFUnDA, PFBS, PFHxS, PFOS, 4:2 FTS, 6:2 FTS, 8:2 FTS at 500 ng L−1 ; PFBA, PFPeA, PFDoDA, PFTeDA, N MeFOSAA, N EtFOSAA at 10 0 0 ng L−1 and FOSA, MeFOSA, EtFOSA, MeFOSE, EtFOSE at 50 0 0 ng L−1 (abbreviations listed in Supplementary Table 4). This was then extracted and diluted with 400 μL of ultra-pure water before being analysed using the parameters in Supplementary Tables 3 and 4. The precision was again decreased for longer chain compounds with some showing very high recoveries. These high recoveries are due to differential adsorption of analytes and inter-

improved consistency between extractions. Flow rates were optimised by performing extractions at gradually increasing loading, washing and elution flow-rates until a decrease in recovery was observed. The extraction method was validated using spiked recoveries from the PFAS contaminated water samples. Six unique samples of differing PFAS concentration and salinity were spiked (100 ng L−1 PFAS), extracted in triplicate and quantified using internal standard calibrations. The percent recovery and standard deviation for each compound over the six samples was then determined (Table 2). The instrument detection limits for all compounds were less than 10 ng L−1 . The lowest Australian guidance value for PFASs in water is 70 ng L−1 (PFOS) [7], making this method suitable for monitoring environmental waters with no additional clean-up required. A typical extraction took 5 min to perform manually, faster than conventional SPE methods such as EPA Method 537 (>30 min) [28], and achieved similar results using a 125× smaller sample volume (2 mL vs 250 mL) [26]. Conventional SPE methods using larger volumes of elution solvent do obtain a higher level of pre-concentration, however this is achieved by evaporation and reconstitution of the extract (8 mL evaporated to ∼0.5 mL and reconstituted in 1 mL for EPA Method 537), a process that can take several hours and may potentially lead to loss of more volatile PFASs such as fluorotelomer alcohols [29]. PFAS were detected in all six of the samples (see Fig. 4), with concentrations ranging from
Table 4 Quantities of PFAS detected in the six surface water samples. Results are reported as the mean ± SD in ng L−1 (n = 3).

A B C D E F

PFBA

PFPeA

PFHxA

PFHpA

PFOA

PFNA

PFDA

340 ± 40 610 ± 30 800 ± 300 225 ± 23 40 ± 6 300 ± 130

350.2 ± 1.1 11.6 ± 0.3 5.3 ± 0.3 33.4 ± 1.0 8.9 ± 0.2 ND

376.8 ± 1.5 48 ± 2 9.8 ± 0.4 76.4 ± 1.0 29.2 ± 0.7 5.1 ± 0.3

124.2 ± 1.4 3.5 ± 0.4 2.2 ± 0.2 31.40 ± 0.13 2.70 ± 0.03 ND

39.9 ± 0.4 2.5 ± 0.2 2.4 ± 0.2 65.6 ± 1.4 5.3 ± 0.5 1.52 ± 0.02

5.7 ± 0.7 6±3 3.8 ± 0.4 8.6 ± 1.0 1.6 ± 0.6 1.8 ± 1.3

4.3 7.1 3.5 7.3 4.4 2.2

ND = Not detected.

± ± ± ± ± ±

1.1 1.6 0.3 0.5 1.5 0.6

PFUnA

PFDoA

PFTeDA

15 ± 7 25 ± 12 12.6 ± 1.3 23.9 ± 1.6 ND ND

ND 20 ± 13 ND 15.6 ± 1.4 ND ND

40 ± 20 76.4 ± 0.2 50 ± 40 66 ± 2 ND 3.5 ± 0.5 30 ± 12 20.49 ± 0.16 ND 61.8 ± 1.2 ND 7.4 ± 0.4

PFBS

PFHxS

PFOS

898 ± 15 254 ± 3 3.9 ± 0.3 88.4 ± 1.1 198 ± 6 7.1 ± 1.1

860 ± 30 10.6 ± 0.7 4.9 ± 0.9 25.5 ± 1.0 582 ± 13 ND

6

T.E. Lockwood, M. Talebi and A. Minett et al. / Journal of Chromatography A 1604 (2019) 460495

Fig. 5. Recoveries and standard deviations for the automated extractions of PFAS (n = 3).

nal standards to extraction equipment following spiking. Short to medium chain PFAAs (C4-10 ) and fluorotelomer sulfonates showed good recoveries of 96 to 105% (excluding PFOS; 129%) and %RSDs of less than 14% (excluding PFOS; 29%) (Fig. 5). These results show that the automation did not affect the performance of the developed μSPE stationary phase and extraction method. Automation reduced the required sample handling, further decreasing preparation time and potentially decreasing analyst-induced errors. These factors, along with the decreased sample and solvent use, are essential for broad uptake in commercial laboratories conducting environmental PFAS analyses. 4. Conclusions This study has demonstrated the use of μSPE as an alternative to conventional SPE for the extraction of C4-10 poly- and perfluoroalkyl acids. Less sample was required (2 mL vs 250 mL), extraction times were reduced to 5 min and automation reduced the labour required. Recoveries and precision were sufficient for the majority of compounds, however further work is required to improve the recoveries of longer chain compounds (C>10 ). Losses of long chain PFASs were speculated to be due to strong sorbent retention or adsorption to containers and the glass syringe. These shortcomings may be overcome through further optimisation of the sorbent and the development of a polypropylene syringe that is suitable for the high pressures of μSPE. Declaration of Competing Interest TEL received financial support from ePrep Pty Ltd. Acknowledgements TEL is supported by an Australian Government Research Training Program Scholarship. DPB is the recipient of an Australian Research Council Discovery Early Career Researcher Award DE180100194, and PAD Australian Research Council Discovery Project Grants DP17010 0 036 and DP190102361. MT and SM received support through the Cooperative Research Centres Projects (CRC-P) grant CRCP54017 funded by the Australian Government: Department of Industry, Innovation and Science.

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