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“BioMig” - a method to evaluate the potential release of compounds from, and the formation of biofilms on polymeric materials in contact with drinking water Gang Wen, Stefan Koetzsch, Marius Vital, Thomas Egli, and Jun Ma Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.5b02539 • Publication Date (Web): 03 Sep 2015 Downloaded from http://pubs.acs.org on September 6, 2015
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Environmental Science & Technology
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“BioMig” ‐ a method to evaluate the potential release of compounds
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from, and the formation of biofilms on polymeric materials in contact
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with drinking water
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Gang Wen1,2,4, Stefan Koetzsch1,4, Marius Vital1,3, Thomas Egli1,3*, Jun Ma2
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Environmental Microbiology, P.O. Box 611, Überlandstrasse 133, CH‐8600 Dübendorf,
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Switzerland. 2
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Eawag, Swiss Federal Institute for Aquatic Science and Technology, Department of
State Key Laboratory of Urban Water Resource and Environment, Harbin Institute of Technology, Harbin 150090, People’s Republic of China.
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Institute for Biogeochemistry and Pollutant Dynamics, Swiss Federal Institute for Technology Zürich (ETH), CH‐8092 Zürich, Switzerland.
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*
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Present addresses:
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Gang Wen: School of Environmental and Municipal Engineering, Xi’an University of Architecture
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Corresponding author: Tel.:+41449233691, E‐mail address:
[email protected] and Technology, Xi’an 710055, People’s Republic of China Marius Vital: Helmholz‐Zentrum für Infektionsbiologie GmbH, Imhoffenstrasse 7, D‐38124
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The first two authors have contributed equally to this work.
Braunschweig, Germany. Thomas Egli: General‐Wille‐Strasse 194, CH‐8706 Feldmeilen, Switzerland, Phone: +41 44 923 36
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Running title: New test for biofilm formation and migration potential of plastics in water
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ABSTRACT
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In contact with water, polymeric materials (“plastics”) release compounds that can support
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suspended microbial growth and/or biofilm formation. The different methods presently used in
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the EU to test plastics take 7‐16 weeks to obtain a result. In industry, this delays material and
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product development as well as quality testing. Therefore, we developed a method package
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(“BioMig”) that allows testing with high reproducibility plastic materials in 2 weeks for their
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potential biofilm (or biomass) formation and release of carbonaceous migration products when
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in contact with water.
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“BioMig” consists of (i) an extended migration potential test (7‐times for 24 h at 60 °C) based
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on the European norm EN 12873‐1 and the German UBA (Umweltbundesamt) guideline, and (ii)
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a biomass formation potential (BFP) test (14 days at 30 °C), which is a modified version of the
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Dutch biofilm production potential test. In the migration potential test the amount of carbon
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released into water by the specimen is quantified by monitoring total and assimilable organic
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carbon over time; furthermore the modular design of the test allows to assess also additional
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parameters such as pathogen growth potential on the migration water, or toxic effects on
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microbial growth. Flow cytometry (FCM)‐based total cell counting (TCC) is used to quantify
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microbial growth in suspension and on surfaces after removal with mild sonication without
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affecting cell integrity. The BFP test allows to determine both, the planktonic (pBFP) and the
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sessile (sBFP) cell fractions. The sBFP consists of surface‐attached cells after removal (> 90%
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efficiency). Results for four standard test materials (PE‐Xa, PE‐Xc, EPDM 2%, and EPDM 20%),
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plus positive (PVC‐P) and negative (glass) controls are presented.
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FCM‐based TCC demonstrates that the release of growth‐supporting carbon and proliferation
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of surface‐attached cells stops increasing and stabilizes already after 14 days of incubation; this
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allows faster assessment of growth‐supporting properties of “plastics” with “BioMig” compared
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to established tests.
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TOC art
Migration test
Migration test
Total organic carbon
Assimilable organic carbon
Short duration 14 days
Identical test water
BioMig Easy to handle
Test battery for assessing the maximum potential of polymeric materials to support microbial growth
Identical microbial inoculum
High reproducibility
Modular
Biomass formation potential test
Biomass formation potential test
Biomass formation potential test
Planktonic cell number
Remaining dissolved organic carbon
Sessile cell number
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INTRODUCTION
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Drinking water and “plastic materials”. Drinking water suppliers take great effort to produce
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a product of high chemical and biological quality before it is delivered to the consumer. When
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water is delivered to consumers through distribution networks made of ‘plastic’ materials, water
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quality may be affected adversely by the release of carbonaceous compounds. The use of
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‘plastic’ materials in distribution networks has been increasing significantly over the last two
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decades, mainly because of their resistance to corrosion and low cost.1 Most of these organic
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polymers contain not only various (in)organic additives such as softeners, fillers, lubricants,
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antioxidants, stabilizers, or colouring agents, but often also traces of starting materials as well as
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reaction products from the polymerization process. It is well known that many of these
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(in)organic compounds can migrate into the water phase2‐4 where they can severely affect water
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quality; this can include taste and odour,2,3 toxic effects,4,5 or unwanted stimulation of microbial
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growth6‐11 (similar problems can be caused by metal pipes7,12). Of particular concern are in‐
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house plumbing systems 9‐11 where low pipe diameters lead to enhanced ratios of material
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surface area to water volume, and, inevitably, to higher concentrations of released compounds
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in the water.
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Consequently, commercial products in contact with drinking water must comply with a
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number of criteria before they can be used in drinking water installations (in the EU the
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European Food Contact Regulations;13,14 in the US the NSF/ANSI Standard15). However, current
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strategies for testing such effects differ. In the US one concentrates on migrating key
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compounds with the intention to evaluate their health effects on a toxicological basis.15 In
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contrast, EU norms/standards focus on the determination of total organic carbon (TOC)
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released,16 effects on taste and odour,17 and on properties promoting microbial growth.13,17 The
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latter aspect is especially important when distributing non‐chlorinated water as done in some
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European countries,18,19 although bacterial regrowth and formation of biofilms are also observed
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in distribution systems supplied with water containing enhanced levels of disinfectant
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residuals.20,21
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Testing migration and its microbiological effects. There is increased awareness that human
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beings are continuously exposed to chemicals migrating from package materials, containers, or
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pipes into food and water, and that little is known about extent and nature of these
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compounds.4,22,23 Here, we concentrate on one specific aspect, namely the fact that most
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polymeric materials release organic compounds that may support microbial growth, either as
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planktonic cells in the bulk water phase, and/or as biofilms on surfaces.
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Whereas the chemical and organoleptic part of testing compounds migrating from materials
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into water is harmonized in the EU,16,17 no EU standard method is yet available for testing
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synthetic polymers in contact with drinking water for their potential to support microbial growth
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and biofilm formation (Table 1). Several countries have developed their own methods and the
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three most prominent ones are listed in Table 1 (note that many countries do not have their
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own standards but rely on those established in the US or the EU). All methods have their
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weaknesses, such as lacking sensitivity, a long time required until a result is obtained, or a
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tedious procedure.
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It also becomes obvious from Table 1 that an inherent incompatibility exists in the different
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parameters and conditions applied in the tests. This includes determination of microbial growth
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(measured as oxygen consumption, ATP, slime volume, or colony forming units), exposure
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conditions (flow through systems versus batch assays with water exchange, duration between 7
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and 16 weeks), and the test water used (local tap water). As a result, data obtained with the
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different methods are not only very difficult to compare, but this methodological incompatibility
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is also in the way of reaching an EU‐wide harmonized test procedure.
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Developing “BioMig” as a faster, better testing procedure. All this calls for testing methods
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that give more information, are faster, well reproducible, but at the same time convenient to
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perform in any appropriately equipped laboratory. Based on the European norm for testing
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migration,16 we have proposed recently a new migration potential test (MP) that allows
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assessment of an additional biological parameter of polymeric materials when in contact with
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drinking water.24 It includes the determination of so‐called “assimilable organic carbon” (AOC) in
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migration test waters. This parameter was included because AOC has been demonstrated
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repeatedly to govern the extent of microbial regrowth and biofilm formation.20,21,25
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Here we combine this MP assay with a biomass formation potential (BFP) test, which we
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developed based on the biomass production potential (BPP) test from the Netherlands (Table 1).
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The new BFP test allows now a quicker assessment of the potential of a material to support i)
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planktonic microbial growth, and ii) the formation of biofilms on material surfaces. The package,
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called “BioMig”, is both easier to perform and faster (Table 1); furthermore, it yields more
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information on properties of compounds released from a material than the tests presently used.
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In addition, “BioMig” can be easily complemented with additional analyses, including for
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example toxic aspects or the determination of a pathogen growth potential (Figure 1). Here, we
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describe the basic principles of, and document typical results obtained with “BioMig” for a
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selection of reference materials. Furthermore, a number of factors that influence experimental
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results obtained with “BioMig” when testing a material for migration and biomass formation
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potential are presented as supporting information.
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EXPERIMENTAL SECTION
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Materials tested. The following polymeric materials were tested: Polyethylene peroxide
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cross‐linked (PE‐Xa), polyethylene silane cross‐linked (PE‐Xb), polyethylene electron beam cross‐
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linked (PE‐Xc), polybutylene (PB) and ethylene propylene dienemethylene with 2%, 12% and
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20% plasticizer (EPDM 2%, EPDM 12% and EPDM 20%); polyvinyl chloride‐(P) containing 30% of
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plasticizer (PVC‐P) was the positive control, and glass was employed as the negative control
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material. All test materials were produced by different manufactures. In case of EPDM’s the
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amount of plasticizer is not related to the Shore hardness of the materials. Note that PE‐Xa, PE‐
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Xb, PE‐Xc, PB, EPDM 2% and EPDM 12% are approved for drinking water. EPDM 20% is only
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approved for wastewater applications and PVC‐P is not approved at all. More information on
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materials used for experimental setup is supplied in supporting information S1.
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Material holders for testing sessile biomass formation potential (sBFP). To reliably quantify
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cells growing on the surface of polymeric materials when exposed to water, specific holders
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were designed and manufactured from inert Teflon (PTFE). The holders allow firm clamping of a
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1x1 cm piece of a material and exposure of a defined surface area to the water phase, on which
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cells can attach and develop into a biofilm. The design allows to easily remove the cells from the
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defined surface using ultrasound. The holders are described in detail in “supporting
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information” with respect to geometry (S2) and handling for detaching cells from the surface
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(S3).
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Different waters used. For rinsing and other purposes ultrapure water produced with a TKA
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water purification system (TKA GenPure, Thermo Scientific, Niederelbert, Germany) was used; it
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contained 6 °C)
25 ± 1 °C
30 ± 1 °C
Test samples
Small plates
Plates, pipes
Small plates
Pipes, plates
S/V (cm )
0.15
Flow‐through system
0.16
1
Test water
Dechlorinated drinking water
Dechlorinated drinking water
Water after slow sand filtration
Bottled mineral water (Evian)
Inoculum
River water
Dechorinated drinking water
River water
Bottled mineral water (Evian)
Water replacement
2x per week
Continuous flow, 20 L/h
1x per week
No replacement
Duration (weeks)
7
12
16
2
Parameters for determination of microbial growth
Oxygen consumption (MDOD)
Volume of surface growth (slime volume)
ATP
TCC ATP(*)
Microbial growth on the surface
Not directly measured
Slime volume
Cell detachment ATP
Cell detachment TCC, ATP(*)
Microbial growth in the water phase
Oxygen consumption (MDOD)
Not considered
ATP
TCC, ATP(*)
‐1
796 797 798 799 800 801 802
S/V: surface to volume ratio (ratio of material surface to water volume) TCC: total cell count ATP*: adenosine tri‐phosphate (but see comments in “supporting information, S14”) CFU: colony‐forming unit MDOD: mean dissolved oxygen difference BPP: biomass production potential BFP: biomass formation potential
803 804
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Table 2. Pathogen growth potential (PGP) of organic carbon compounds migrating from plastic
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materials commonly used for in‐house drinking water distribution systems, and estimated
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fraction of AOC that supports growth of E. coli O157, V. cholera or P. aeruginosa. The
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percentage of AOC consumed was calculated using the calibration factor from Vital,29,30 which
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relates cell number produced to AOC consumed. For all tests migration water M1 was used (see
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S13).
Parameters
PE‐Xb
PB
EPDM 2%
TOC (mg/L)
0.17
0.62
0.83
Natural flora
AOC (µg/L)
121±12
237±145
295±100
PGP (10 cells/mL)
1.55±0.10
8.50±0.40
6.50±0.20
AOC consumed (%)
36±1.4
88±12
40.5±1.8
PGP (10 cells/mL)
1.35±0.10
6.30±0.88
1.80±0.01
AOC consumed (%)
26.5±1.7
42.1±3.5
16.8±0.6
PGP (10 cells/mL)
0.48±1.00
1.01±2.10
0.730±0.09
AOC consumed (%)
17.8±0.4
18.8±0.6
16.9±0.3
5
P.aeruginosa
5
V. cholerae
5
E. coli O157
811
812 813 814
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Alternative situation (B)
Current situation (A)
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Migration test EN12873-1 and UBA guideline
TOC
Taste and Odour
Biofilm growth potential tests (prEN 16421:2012)
&
Identification of migrated substances
Migration test EN12873-1 or UBA guideline
&
Planktonic
Sessile
W270
No
Yes
BPP
Yes
Yes
MDOD
Yes
No
Modified migration and modified biofilm formation potential test Migration test
TOC
Taste and Odour
Identification of migrated substances
BFP test Optional
Toxicological Studies
Growth of pathogen
Taste & odour in combination with microbiology
Clarification of problem cases
Identification of nonbiodegradable substances
TOC
AOC
Planktonic Sessile Remaining DOC
Formation of disinfection by-products
815 816
Figure 1. Conceptual sketch of the current (A) and an alternative (B) strategies for testing
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polymeric materials in contact with water for migration and biofilm formation properties. Both
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strategies are based on short‐term migration tests and on long‐term biofilm formation tests.
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The proposed alternative (B) is based on a modular design of the testing procedure consisting of
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a combination of the existing EN‐migration test with some newly developed methods described
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in this work. Blue and green boxes indicate mandatory parameters; in yellow boxes some
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optional parameters that can be added. AOC, assimilable organic carbon; BFP, biomass
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formation potential; DOC, dissolved organic carbon; TOC, total organic carbon. Details on the
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German biofilm formation test W270, the Dutch biofilm production potential (BPP) test, and the
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UK oxygen consumption test (MDOD), the UBA guideline and the European norm (prEN
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16421:2012) 36 is given in Table 1.
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1.0
0.4
Negative control
0.3
EPDM 2%
0.8 0.6
0.2
0.4 0.2
TOC-AOC (mg L )
-1
0.0 0.4
-1
TOC-AOC (mg L )
0.1
PE-Xa
0.3 0.2 0.1 0.0 0.4
PE-Xc
0.0 50
EPDM 20%
40 30 20 10 0 50
Positive control
40
0.3
30 0.2 20 0.1
10
0.0
0
M1
828
M2
M3
M4
M5
M6
M1
M7
M2
M3
M4
M5
M6
M7
829
830
Figure 2. Total organic carbon (TOC, hatched columns) concentrations in the seven migration
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waters (M1 to M7) and assimilable organic carbon concentrations (AOC, empty columns, in M1,
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M3, M5, and M7) leached from different plastics, the negative (glass) and the positive control
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material (PVC‐P) obtained in a migration assay. Migration cycles were performed at 60 °C during
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24h as described in “Experimental Section” and depicted in Table S13. The values for the
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negative control were subtracted from the results obtained for the four test materials. When
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testing EPDM 20% no detectable AOC was found in M1, M3, M5 and M7.
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1.5x10
8
1.0x10
8
5.0x10
7
-2
Biomass formation potential (cells cm )
A
0.0 8 4x10 3x10
8
2x10
8
1x10
8
B
0 4x10
8
3x10
8
2x10
8
1x10
8
0
C
0
10
20
30
Time (day)
839 840
841
Figure 3.Development as a function of time of surface‐attached and suspended microbial cells in
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a ”BioMig” test performed with EPDM 2%. Growth is given as the biomass formation potential
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(BFP) in cells per cm2 of surface area of the tested material. A) planktonic cells (pBFP); B) sessile
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cells (sBFP); C) BFP, which is the sum of pBFP and sBFP (see S12).
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BFP (cells cm )
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1.0x10
9
8.0x10
8
6.0x10
8
4.0x10
8
2.0x10
8
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8.3710
pBFP sBFP BFP
A A
2.7210
8
7
4.7110
6
6.1210 3.77107
5.9210
6
30
Remaining DOC -1 (mg L )
25
8
27.24
B B
24.31
20
6.35
5 0.49
0.50
0.54
PE-Xa
PE-Xc
0 Negative control
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EPDM 2%
EPDM 20%
Positive control
849 850
Figure 4. Biomass formation potential (BFP) determined with the “BioMig” assay for four
851
polymeric materials, negative (glass) and positive (PVC‐P) control (A), and remaining DOC in the
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bulk water phase (B) after incubation for 2 weeks. pBFP, planktonic biomass formation
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potential; sBFP, sessile biomass formation potential; BFP, biomass formation potential.
854 855 856
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857 1.0 0.9
TOC
-1
AOC or TOC (mg C L )
0.8 0.7
A
0.6 0.5 0.4
AOC
0.3 0.2 0.1 0.0 1
2
3
4
5
6
7
Experiment repeats 8
-2
Biomass formation potential (cells cm )
4.0x10
pBFP sBFP BFP
8
3.5x10
B
8
3.0x10
8
2.5x10
8
2.0x10
8
1.5x10
8
1.0x10
7
5.0x10
0.0
1
2
3
4
5
Experiment repeats
858 859
Figure 5. Repeatability of “BioMig” assays exemplified for EPDM 2% as the test material. A)
860
Repeatability for the parameters TOC and AOC in the migration potential test using migration
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water M1;B) repeatability of the biomass formation potential (BFP) test for both, sessile (sBFP)
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and planktonic (pBFP) cell fraction.
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