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Competition between Naegleria fowleri and free living amoeba colonising laboratory scale and operational drinking water distribution systems Haylea C Miller, Jason Wylie, Anna Kaksonen, David Sutton, and Geoffrey J. Puzon Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b05717 • Publication Date (Web): 01 Feb 2018 Downloaded from http://pubs.acs.org on February 3, 2018

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Environmental Science & Technology

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Competition between Naegleria fowleri and free living amoeba colonising laboratory scale and

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operational drinking water distribution systems

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Haylea C. Miller1, 2, Jason T. Wylie1, Anna H. Kaksonen1, 2, David Sutton2 and Geoffrey J. Puzon1*

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1

CSIRO Land and Water, Private Bag No.5, Wembley, Western Australia 6913, Australia

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2

School of Biomedical Sciences, University of Western Australia, 35 Stirling Highway, Crawley,

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Western Australia 6009, Australia

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ABSTRACT

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Free living amoebae (FLA), including pathogenic Naegleria fowleri, can colonise and grow within

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pipe wall biofilms of drinking water distribution systems (DWDSs). Studies on the interactions

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between various FLA species in biofilms are limited. Understanding the interaction between FLA and

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the broader biofilm ecology could help better predict DWDS susceptibility to N. fowleri colonisation.

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The aim of this study was to determine if N. fowleri and other FLAs (Naegleria, Vermamoeba,

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Willaertia and Vahlkampfia spp.) co-colonise DWDS biofilm. FLAs commonly isolated from DWDSs

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(N. fowleri, V. vermiformis and N. lovaniensis) were introduced into laboratory-scale biomonitors to

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determine the impact of these amoebae on N. fowleri’s presence and viability. Over 18 months, a

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single viable amoebae (N. fowleri, N. lovaniensis, or V. vermiformis) was detected in each biofilm

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sample, with the exception of N. lovaniensis and N. fowleri, which briefly co-colonised biofilm

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following their co-inoculation. The analysis of biofilm and bulk water samples from operational

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DWDSs revealed a similar lack of co-colonisation with a single FLA detected in 99 % (n = 242) of

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samples. Interestingly, various Naegleria spp. did colonise the same DWDS locations but at different

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times. This knowledge furthers the understanding of ecological factors which enable N. fowleri to

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colonise and survive within operational DWDSs and could aid water utilities to control its occurrence.

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Keywords: Naegleria fowleri, biofilm, water, colonisation, amoeba, ecology

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INTRODUCTION

ACS Paragon Plus Environment


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Pipe wall biofilm within drinking water distribution systems (DWDSs) provides an ideal habitat for

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free living amoeba (FLA) 1, 2. Biofilm protects FLA from common water disinfectants like chlorine

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and provides dense bacterial biomass for grazing 3, 4. The role of biofilm in harbouring potential

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pathogens, including Naegleria fowleri and FLA-associated bacterial pathogens, such as Legionella

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and non-tuberculous mycobacterium (NTM) 5, is a growing issue for water utilities as managing these

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pathogens in DWDS biofilm can be difficult 4.

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N. fowleri, the causative agent of primary amoebic meningoencephalitis (PAM), has been detected in

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DWDSs in Pakistan, the USA and Australia. All three countries have reported PAM cases linked to

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DWDSs. Since 2008, 102 PAM cases were linked to nasal rinsing using tap water in Karachi,

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Pakistan, and at least 6 cases linked to DWDSs in the USA and USA territories6-11. Seven cases have

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also been reported in Queensland, Australia since 1971, linked to the domestic use of tap water, most

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recently in October 2016 12-14.

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Many factors are known to affect the ability of FLA to colonise a niche. Water origin, pipe line

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materials, ions, metals, pH, environmental temperatures, organic matter, nutrient level, biofilm

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microbial community and disinfectant residuals have been shown to influence the ability of FLA,

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including N. fowleri, to colonise and survive in DWDSs 1, 4, 15-26. However, previous research is

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limited on the effect of competitive FLA, especially Naegleria, Vermamoeba, Willaertia and

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Vahlkampfia spp., on the ability of N. fowleri to colonise and thrive in DWDSs 1, 4, 26-29. Given that

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FLA are bacterivorous, changes in viable FLA populations within DWDSs could be attributed to

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negative interactions with each other as they compete for food sources and protective niches within

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the biofilm. Under laboratory conditions Balamuthia spp. predate on N. fowleri, N. gruberi and

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Acanthamoeba spp. 30. Previous research studies have suggested biological factors may influence the

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ability of individual Naegleria spp., including N. fowleri, to colonise operational DWDSs 28.

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However, to date, FLA interactions have not been well studied. Direct negative interactions by other

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FLA could make DWDS environments unfavourable for N. fowleri colonisation and persistence.

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The aim of this study was to determine the impact of competing Naegleria, Vermamoeba, Willaertia

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and Vahlkampfia spp., on the ability of N. fowleri to colonise biofilms. Understanding the ecological

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factors and interactions that influence N. fowleri colonisation within operational DWDS biofilms is


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vital to improve the control strategies for this pathogen. To our knowledge, this is the first study to

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demonstrate that FLA competition can negatively affect the ability of N. fowleri to colonise and

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persist in drinking water biofilm. This work further aids in the understanding of how FLA can

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influence the presence and persistence of pathogenic N. fowleri in operational DWDS biofilms.

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MATERIALS AND METHODS

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Laboratory-scale biomonitors: A laboratory set up of six replicate biomonitors (KIWA,

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Netherlands), as previously described 4, were used to determine the effect of competitive FLA on the

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ability of N. fowleri to colonise and persist in biofilms. Briefly, each biomonitor contained up to 42

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glass rings, each with a surface area of 16.96 cm2, which acted as growth substrates for biofilm

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formation. Glass was selected as a lower colonisable surface versus other materials, as has been

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previously reported for laboratory and field based studies 1, 4, 26-28. The biomonitors (total volume of 25

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L per biomonitor) were operated at an average recirculation flow rate of 15 ± 1.7 L/h using bulk water

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(approximately 30 L recirculated) and biofilm sourced from operational field biomonitors described

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below with known FLA populations 1, 26-28. Field bulk water and biofilm were used in the laboratory-

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scale biomonitors in order to replicate the chemical and microbial conditions of operational DWDSs,

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including pH, water source and bacterial communities.

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After the initial set up, the biomonitors contained viable N. fowleri and V. vermiformis sourced from

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the DWDS field sites. Escherichia coli (1 mL of 108 cells/mL) was intermittently added to all

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biomonitors on a monthly basis as a food source to further promote FLA growth. At two and six

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month time points, laboratory cultured (described below) N. fowleri (5 mL of 105 cells/mL) was

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inoculated into all six biomonitors. After 7 months, laboratory cultured N. lovaniensis (5 mL of 105

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cells/mL) was inoculated into all six biomonitors, along with additional N. fowleri (5 mL of 105

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cells/mL), after which point no more FLA were inoculated. All six biomonitors were operated in an

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air conditioned laboratory at approximately 22 °C, which is the threshold temperature for the

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surveillance of N. fowleri in DWDSs in Western Australia.

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Biofilm sample collection and processing were conducted as described previously 4. Briefly, glass

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rings were removed from the biomonitors and placed in 30 mL of sterile 25 % Ringers solution in a



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sterile 50 mL tube. Glass rings were vortexed, sonicated (5 min, 30 W with a working frequency of 47

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kHz + 6 % (Bransonic, USA)) to detach the biofilm and cells were harvested by centrifugation at

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5,000 × g for 10 min. Cell pellets were resuspended in a smaller volume of the original solution and

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the cell concentrate were used for viability plating, flow cytometry and DNA extractions 4.

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Operational DWDS field sites: To determine the effect of naturally occurring FLA on the presence

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and survival of N. fowleri in operational DWDSs, five rural field sites (SK, KT, WB, P, B) along two

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regional schemes and one urban field site in one metropolitan scheme (BP) in Western Australia

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(WA) were selected. SK, KT and WB sites were located along a chlorinated DWDSs, whereas P and

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B sites were located along chloraminated DWDSs 28. The BP site was connected to an urban DWDS

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system receiving chlorine at its source (pre-treatment) and was monitored for one year. SK and KT

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sites were monitored seasonally for 3 and 1.3 years, respectively, and were selected for this study

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based on their typically low free chlorine residual (less than 0.1 mg/L) 1, 26. WB site was monitored for

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4 years and had a chlorine residual ranging from 0 mg/L to 0.55 mg/L, with the highest residuals

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detected in the winter months 1. The P and B site biomonitors were located along a chloraminated

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rural DWDS in. P and B sites had previously been monitored for 0.6 and 1 year, respectively, and had

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chloramine residuals ranging from 0 mg/L to 0.07 mg/L and 0 mg/L to 1.68 mg/L, respectively. All

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sites were selected for this study based on persistent historical detections of viable FLA, including N.

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fowleri, N. lovaniensis and V. vermiformis 1, 4, 26, 28.

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Bulk water and biofilm samples: Both bulk water and biofilm were monitored seasonally for the 6

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sites for the presence of viable and non-viable FLA, microbial cell counts, free and total chlorine or

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chloramine residuals and water temperature as previously described 1, 26. Briefly, bulk water samples

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were collected in sterile 250 mL collection bottles directly from the pipelines using a pre-sterilized

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spout after flushing the pipe for 5 min before sampling to ensure the sample represented true bulk

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water conditions. Biofilm samples were collected from the 6 KIWA biofilm monitors (described

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above) which were connected directly to the pipe lines and continuously operated with a flow rate of

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50 L/h 1, 4, 26, 27. Samples were transported to the laboratory at room temperature. Cells were

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concentrated from bulk water samples by centrifugation (5,000 × g, 10 min), and the supernatant was


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discarded. Concentrated samples were used for viability plating, flow cytometry and DNA

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extractions.

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Physical and chemical measurements: A pocket colorimeter II (Hach, U.S.) was used to measure

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free and total chlorine or monochloramine and free ammonia concentrations, according to the

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manufacturer’s protocol. Temperature was measured using the MC-87 Dual Channel Digital

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Thermometer (TPS, Australia).

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Laboratory stock culturing: Culturing methods used for laboratory FLA and bacteria including N.

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fowleri, N. lovaniensis, and Escherichia coli were as previously described 4, 27. Briefly, individual

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amoebae (either N. fowleri, N. lovaniensis or Vermamoeba vermiformis) were cultured in 75 cm2

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tissue culture flasks (Iwaki) to trophozoite phase with 10 mL of a 25 % Ringers solution (Oxoid), 100

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µL of an E. coli culture (5.39 × 108 cells/mL), and 2.5 mL of amoeba inoculum. Cultures were

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incubated at 37 °C for N. fowleri and V. vermiformis and at 30 °C for N. lovaniensis. E. coli was

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grown in 2 L of Luria-Bertani broth at 37 °C to late log phase, concentrated by centrifugation at 5,000

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× g for 10 min and resuspended in a 25 % Ringers solution.

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Microbial enumeration: Laboratory cultured FLA were enumerated using a Thoma haemocytometer

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(Lab Optik, Germany). Samples were loaded onto the haemocytometer (10 µL) and viewed under the

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microscope using phase contrast (Olympus, Japan) at 100 × magnification. The cells (trophozoites and

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cysts) present within the gridded area were counted in duplicate and calculated as cells/mL.

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Enumeration of total microbial cell concentrations in bulk water and biofilm samples were conducted

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as previously described using a Quanta flow cytometer (Beckman Coulter Quanta, U.S.) 26, 31-33. Total

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cell counts were enumerated following staining with SYBR Green 1 (Invitrogen, U.S.). Briefly, both

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field and laboratory samples were diluted with Milli-Q water to fit into the counting range of the flow

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cytometry. A 200 µL aliquot of the diluted sample was stained with 2 µL (10 ×) of SYBR Green 1

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and incubated for 15 min in the dark. Counting zones for total cell counts were selected based on

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Hoefel et al. 2003. Controls included; filtered and/or unstained field samples or Milli-Q water, and

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flow check fluorospheres. All samples were run in duplicates and average values were reported. The

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flow cytometer had a standard error of 5 % as determined by the manufacturer. Results for flow



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cytometry were calculated as either cells/mL for bulk water samples or as cells/cm2 for biofilm

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samples.

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Viability assessment: The viability of thermophilic FLA including Naegleria, Vermamoeba,

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Willaertia, Vahlkampfia, Tetramitus spp. 27, 34 was determined as previously described 26, 27. Briefly,

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cell concentrates were incubated on non-nutrient agar (NNA)- E. coli plates for at least 48 h at 42 °C

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and observed for plaques. The numbers of viable amoebae were estimated using plaque counting on

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NNA-E. coli plates under the assumption that each plaque was generated by 1 viable amoebae.

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Plaques were scraped using sterile disposable 1 µL loops and collected for DNA extraction and

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quantitative polymerase chain reaction (qPCR) for species identification. NNA was prepared by

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mixing 1 L of 25 % Ringers solution with 15 g of bacteriological agar (Agar No. 1, Oxoid England)

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and autoclaved at 121 °C for 20 min before plating 100 µL of E. coli culture (5.39 × 108 cells/mL) on

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the plates. E. coli was grown in 2 L Luria-Bertani broth (Oxoid, England) at 37 °C to late log phase,

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concentrated by centrifugation at 5,000 × g for 10 min and resuspended in 20 mL of 25 % Ringers

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solution.

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DNA extraction: DNA was extracted from both the laboratory and field biofilm samples and field

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bulk water using the following two methods. First, PowerSoil DNA Isolation kit (MO BIO

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Laboratories, USA) was used for extracting total DNA from cell pellets harvested at 21,000 × g for 5

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min from cell concentrates according to the manufacturer’s protocol. Second, the Bio-Rad InstaGene

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Matrix (BioRad, U.S.) was used according to the manufacturer’s protocol for extracting DNA from

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viable plaques as previously described 1, 4, 26-28. Positive NNA - E. coli plates were scraped using a 1

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µL sterile disposable loop and resuspended in 100 µL of InstaGene matrix. All DNA extracts were

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stored at – 20 °C until analysed by qPCR.

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FLA identification: Methods for FLA identification by qPCR melt curve analysis have previously

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been described 27 (SI Table 1). Briefly, DNA samples were analysed using a Bio-Rad iQ5 (Bio-Rad,

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U.S.) with a total reaction volume of 25 µL; containing 12.5 µL HotStar Taq Master Mix (2 ×)

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(Qiagen, U.S.), 1.25 µL of each primer (10 µM), 0.1 µL 500 µM SYTO9 dye (Molecular Probes,

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U.S), 7.9 µL sterile double distilled water and 2 µL of template DNA. Samples were run in triplicates

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with either an N. fowleri specific primer set, which only amplifies the intragenic spacer region (ITS)


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and 5.8S ribosomal DNA of N. fowleri 27, or a consensus primer which amplifies the ITS and 5.8S

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ribosomal DNA of Naegleria spp. 35 and other FLA including Tetramitus, Willaertia, Vahlkampfia,

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and Vermamoeba spp.29. The amplification efficiency, calculated by the iQ5 software, was 85.2 % for

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the specific primers and 77.2 % for the consensus primers when using N. fowleri type 5. Balamuthia

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mandrillaris was identified using species specific primers for a portion of the mitochondrial 16S

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rRNA gene 36 and the following PCR conditions modified from Booton et al. 2003: Initial denaturing

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step of 15 minutes at 95 °C, followed by 40 cycles of 95 °C 30 s, 45 °C 1 min, 72 °C 1 min, and with

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a 6 s pause at 80 °C for fluorescent dye detection. PCR for Acanthamoeba spp. was conducted using

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genus specific primers for the18S rRNA gene 37 and the following PCR conditions modified from

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Schroeder et al. 2001: Initial denaturing step of 15 minutes at 95 °C, followed by 45 cycles of 95 °C

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30 s, 60 °C 1 min, 72 °C 2 min, and with a 6 s pause at 80 °C for fluorescent dye detection. Positive

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controls (target DNA), DNA extraction method controls (Instagene Matrix or PowerSoil elution

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buffer with no DNA template) and negative controls (RNase-free H2O) were run with every PCR

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reaction. FLA identifications were further confirmed using Sanger sequencing (Macrogen, Korea).

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Statistical analysis

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The R statistical package (version 1.0.153) was used to generate box and whisker plots and perform

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statistical analysis of amoebae density and bulk water and biofilm cell count distributions across the

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laboratory biomonitors. Chi-square tests were used to compare variables between groups, including

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comparison between the viable culture based or molecular detection methods, and comparison

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between amoebae distribution in bulk water and biofilm samples. P values lower than 0.05 were

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considered statistically significant.

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RESULTS

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FLA co-colonisation under laboratory conditions

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In the laboratory biomonitors, a single viable FLA species was detected in the majority of biofilm

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samples for the first 10 months of the experiment from all six laboratory biomonitors (Table 1).

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During this time period only viable N. fowleri and V. vermiformis were detected in the same

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laboratory biomonitors at different time points, however, they were not detected within the same



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biofilm sample (i.e. the biofilm attached to one glass ring). Viable N. fowleri was displaced from the

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biomonitors by the fourth month, despite being re-inoculated into all biomonitors at 2 and 6 months.

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V. vermiformis was also displaced from the biomonitors by the end of the seventh month. Following

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N. fowleri re-inoculated and N. lovaniensis introduction at 10 months, both amoebae were detected

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co-colonising biofilm samples from three of the six biomonitors (Biomonitors 3, 5 and 6) in the first

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five days after co-inoculation (Table 1). N. fowleri and N. lovaniensis co-colonisation was again

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detected in following samples (11th month) in two of the six biomonitors (Biomonitors 2 and 6) (Table

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1). In the 13th month (four months after N. fowleri and N. lovaniensis co-inoculation), only viable N.

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lovaniensis was detected in biofilm samples from all six biomonitors. N. fowleri had been displaced

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from the biomonitors again and was not detected for the rest of the experiment (eighteen months in

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total). The detection of both viable N. fowleri and N. lovaniensis during the eleventh and twelfth

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months was the only co-colonisation of multiple Naegleria spp. seen in the biomonitors for the

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duration of the laboratory experiment.

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The concentration of viable FLA in each biomonitor differed but were within a similar range (Table 1

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and Figure 1). All biomonitors had variable means and spreads in regards to FLA densities,

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particularly biomonitors 3 and 6. This trend was also seen in the biofilm cell counts detected across

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the six biomonitors during the 18 month laboratory study (Table 2 and SI Figure 1).

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FLA co-colonisation in operational DWDSs – impact of potential competitors

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The analysis of the bulk water and biofilm samples from the operational DWDSs showed that co-

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colonisation of FLA (Naegleria, Vermamoeba, Willaertia, Vahlkampfia and Tetramitus spp.) was

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rare. A total of 278 FLA were detected by culture (n = 102) and molecular methods (n = 176) in the

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243 bulk water and biofilm samples during the long term field study at six sites (Table 3). Of those

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detections, a single viable FLA was detected in 99 % (n = 242) of field bulk water and biofilm

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samples analysed by the culture method. The most frequently detected viable FLA was N. lovaniensis

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(28 %, n = 29). Only 1 sample (< 1 %) (SK bulk water sample, 250 mL) contained multiple viable

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FLA: namely N. fowleri and V. vermiformis.

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Based on molecular analyses, 88.5 % (n = 215) of analysed samples contained a single FLA. Multiple

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non-viable FLA were detected by molecular methods in 16 % of samples (n=28) (bulk water; n = 5,


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biofilm; n = 23) across all sites. The FLA typically included Naegleria spp. with V. vermiformis. Of

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the 28 samples with multiple non-viable FLA detection, only 5 samples contained multiple non-viable

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Naegleria spp., including combinations of N. fowleri and N. andersoni, N. fowleri and N. lovaniensis,

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and N. fowleri and N. dobsoni, all of which were detected in samples from the chlorinated DWDS at

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SK, KT and WB. All other multiple non-viable detections comprised V. vermiformis and a single

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Naegleria spp.. Furthermore, FLA were more frequently detected at all sites using the molecular

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method compared to the culturing method (p < 0.01) (Table 3). The most frequently detected non-

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viable FLAs were N. fowleri and V. vermiformis (32 % and 32 %, respectively). Some FLAs were

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only detected by the molecular method, including N. andersoni and N. dobsoni, at sites SK, KT and

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WB.

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FLA co-colonisation in operational DWDSs – impact of physical and chemical conditions

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Impact of disinfection: Co-colonisation of FLA was absent across all sites, regardless of the sites

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physical or chemical conditions measured. However, FLA detections varied across the chlorine and

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chloramine treated DWDSs depending on the site and season. The chlorinated DWDS sites (SK, KT

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and WB) had similar physical and chemical conditions including water temperatures and biofilm

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densities (+ 1 log) (Table 4). Over the study FLA were less frequently detected at the KT and WB

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sites than the SK site (p < 0.01) (Table 3). SK site had a chlorine residual (< 0.1 mg/L) whereas KT

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and WB sites had fluctuating chlorine residuals (ranging from 0 mg/L to 1.57 mg/L). The

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chloraminated DWDS sites, P and B, had chloramine residuals of less than 0.07 mg/L or a residual

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ranging from 0 mg/L to 1.68 mg/L, respectively (Table 4). Despite the difference in disinfectant

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residuals FLA were detected at equal frequency at both sites (p > 0.75) (Table 3). N. lovaniensis and

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V. vermiformis were the only detected FLA at the chloramine sites. The single co-colonisation

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observed in this study was in the absence of a free chlorine residual, i.e. 0 mg/L. All sites with a

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disinfectant residual above 0.11 mg/L were negative for viable amoebae. Previous work has already

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demonstrated the relationship between the presence of N. fowleri and other FLA and a disinfectant

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residual in DWDSs1.



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The untreated underground water site BP, had no disinfectant residual detected for the duration of the

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study (Table 4) and also revealed a lack of co-colonisation of multiple viable FLA. Viable N.

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lovaniensis was the most commonly detected FLA at the site with rare detections of V. vermiformis.

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Impact of sample type: Overall, analysis of all field site samples revealed that FLA were detected

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more frequently in the biofilm samples (n = 237) compared with the bulk water (n = 41) (p < 0.01)

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(Table 3). Furthermore, all field sites had higher total cell counts present in both the bulk water and

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biofilm samples than the laboratory samples. The mean biofilm (105 to 106 cells/cm2) and bulk water

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cell counts (105 to 107 cells/mL) from the field DWDSs were relatively stable across the study

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throughout all seasons, despite disinfectant type and fluctuating seasonal temperatures (Table 4 and

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5). A decrease in the mean biofilm (1 log) and bulk water (2 log) cell counts was detected with in the

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presence of elevated disinfectant residual (Table 4 and 5).

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Impact of temperature: Temperature was seen to impact the FLA distribution in the field samples.

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Viable amoebae were detected during all seasons across the six sites, with viable amoebae detected in

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water temperatures ranging from 10 to 43 °C (Table 4), and viable N. fowleri in temperatures ranging

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from 13 to 41 °C. The BP site had the most stable average temperature (42 + 1 °C) through all seasons

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with the exception of the final winter time point when the bore was shut down (21 °C) and was

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consistently positive for viable amoebae. However, viable amoebae were not always detected in the

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presence of higher temperatures (> 25 °C).

266 267

DISCUSSION

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The results of this study indicate that viable N. fowleri does not co-colonised DWDS biofilms and

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bulk water with other Naegleria spp.. Viable N. fowleri was detected with viable V. vermiformis in a

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single DWDS bulk water sample. Occasionally, multiple Naegleria spp. were detected in the same

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field sample using molecular methods, which detect both viable and non-viable cells, however all of

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these samples only contained a single viable FLA. In addition, most sites were colonised by a single

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viable Naegleria spp. throughout the study. However, some sites (KT and SK) were able to host

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different viable Naegleria spp., (N. fowleri or N. lovaniensis) at different times. This indicates that

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while some sections of the DWDSs may favour a single Naegleria spp., different Naegleria spp., are


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able to colonise the same DWDS location at different times. This change in Naegleria spp.

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colonisation is likely due to ecological factors, but it is unclear which factor influences individual

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species colonisation. Under laboratory conditions, viable N. fowleri and N. lovaniensis were detected

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co-colonising biofilm samples upon co-inoculation into the biomonitors. However, this co-

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colonisation resulted in the displacement of N. fowleri and the domination of N. lovaniensis within

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two months in all but a single biomonitor (BM6-Table 1) and in all biomonitors in four months. The

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absence of multiple viable Naegleria spp. co-colonising distribution systems has also been seen in

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other studies, although it was not the focus of the research. Previous studies analysing the presence of

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viable amoebae in Western Australian DWDSs have also shown that DWDS biofilm is typically

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dominated by a single viable FLA including Naegleria spp. 28. Viable Naegleria spp. were detected

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along with other viable amoebae, typically Vermamoeba sp. 1, 26, 28 and in other studies 38. It is possible

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that other FLA were present at the sites, but not detected by the primers used in this study. However,

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similar trends have been shown in other molecular based studies where Naegleria, Willaertia and

289

Vahlkampfia spp. have been reported in anthropogenic and aquatic environments 39 and Naegleria

290

spp. were shown to dominate water and biofilm samples. Muchesa et al. studied the co-occurrence of

291

FLA and bacteria in hospital water networks and saw that almost 50 % of samples contained viable

292

amoebae (77 samples), including Acanthamoeba spp., V. vermiformis and N. gruberi 40. However, it is

293

not clear whether these amoebae occurred in the same samples or fluctuated in presence over time,

294

sample type and location within the building’s plumbing. Ovrutsky et al. detected multiple viable

295

amoebae within the same biofilm samples, including V. vermiformis and Flamella spp. or V.

296

vermiformis and Acanthamoeba spp. 41. They also reported the detection of two different

297

Acanthamoeba spp. present in one sample, but no detection of multiple viable Naegleria spp. was

298

reported. Interestingly, research has shown behavioural difference in amoebae strains, including

299

Willaertia magna Z503 and C2c in regards to susceptibility to Legionella pneumophila, a known

300

intracellular parasite of many FLA species 42. However, behavioural variations have not been reported

301

for strains of N. lovaniensis, N. fowleri or V. vermiformis (the three viable amoeba reported in this

302

manuscript). Different inter-amoebal interactions due to strain variations would be an interesting



303

focus of future studies as varying pathogenicity has been suggested for N. fowleri cultured in animal

304

brain passage or Vero cells compared with axenic culturing 43, 44.

305

Given that FLA are bacteriovorous, biofilms provide the ideal environment for amoebae within

306

DWDSs by providing food sources in the form of dense bacterial biomass, as well as a sheltered

307

niche, increasing resistance to disinfectants 2-4, 18, 26, which supports the increased abundance of FLA

308

in the DWDS biofilm compared to the DWDS bulk water samples (Table 3). Multiple FLA would be

309

expected to colonise niches with similar ecological and chemical conditions. Previous studies looking

310

at the presence of FLA in in-premise plumbing also showed a higher prevalence of FLA in biofilm

311

samples compared to bulk water samples 40, 41. In vitro simulations of pipe lines showed that when

312

introduced, N. fowleri quickly colonised and survived within the biofilm for more than five months in

313

an enclosed simulated pipe system 2. However, in our study, the presence of FLA including N. fowleri

314

in both the field and laboratory studies showed that colonisation was dominated by one species, and

315

far more transient and sporadic than previously thought.

316

Factors known to play an important role in the ability of amoebae including N. fowleri, to colonise

317

and thrive in DWDSs and other environments include water origin, pipe line materials, ions, metals,

318

pH, disinfectant residuals and environmental temperatures, organic matter, nutrient level 15-19, 21-23.

319

However, many of these parameters did not notably vary in the laboratory biomonitor study and at the

320

BP site (i.e. disinfectant residual and temperature), where transient and shifting FLA population were

321

still observed. Previous studies have also found that FLA population changes were not explained by

322

the environmental parameters listed above 21. Therefore, the change in FLA colonisation observed in

323

the laboratory and operational DWDSs were likely attributed to biological factors instead of physical

324

and chemical factors.

325

To date, inter-amoebae interactions have not been well studied, with the exception of Griffin et al.

326

“flagellate empty habitat hypothesis” 45, which stated that FLA communities when disrupted by

327

human activity, allowed FLA with advantageous traits to dominate over other FLA and colonise the

328

newly vacated niche. For example, higher environmental temperatures favour thermophilic amoebae,

329

like N. fowleri. However, temperature was not seen to impact FLA distribution under laboratory

330

conditions and viable N. fowleri was detected in operational DWDSs also in winter months (13 °C).


Page 12 of 23

Page 13 of 23


331

Furthermore changing FLA populations were seen even in the absence of direct human activity, as

332

detected at the BP site. However, human intervention may have played a role in the other sites

333

studied, particularly the presence of fluctuating disinfectant residuals at some sites reported here.

334

Therefore, it is likely that ecological factors like interactions with the microbial community of the

335

biofilm affect the presence and distribution of FLA populations within DWDSs.

336

Previous studies have demonstrated the benefits of multiple organisms living together within a DWDS

337

biofilm, including increased resistant to disinfectants 3, 5. However, few studies have looked at the

338

negative aspects of multiple organisms, especially N. fowleri, co-colonising biofilms. Studies of

339

Western Australian DWDSs have shown correlations between the presence of amoebae and other

340

eukaryotic organisms within the pipe wall biofilm. Puzon et al. showed that the eukaryotic

341

community of the DWDS biofilm had a significant impact on the presence of N. fowleri 28. They also

342

showed a negative correlation between the eukaryotic taxa Rotifera and N. fowleri. However, N.

343

lovaniensis and Rotifera were positively correlated. Studies have also reported the direct predation of

344

amoebae by other FLA. Balamuthia spp. and Paramecium spp. have been shown to predate on

345

amoebae including N. fowleri, N. gruberi and Acanthamoeba spp., instead of bacteria 30, 46.

346

Alternatively, negative interactions between FLA might be due to direct competition for food sources.

347

As FLA are bacteriovorous, it is likely that inter-amoebal interactions have evolved as competition for

348

food sources can be high. Previous studies have demonstrated negative interactions between

349

eukaryotes which competed for the same bacterial food sources 47. Bacterial grazers were shown to be

350

mutually sensing and responding to each other using chemical cues and antagonistic compounds that

351

detrimentally affected their competitors. Studies have also suggested that FLA-bacteria associations

352

could explain FLA distribution in DWDSs 21. Nutrients levels and food sources are known to play an

353

important role in the growth competition and biofilm colonisation by N. fowleri and other amoebae18.

354

Increasing levels of bacterial richness has been shown to be associated with the presence of N. fowleri

355

in DWDSs, therefore the biofilm composition likely effects N. fowleri colonisation 1. Furthermore,

356

individual bacterial families have been associated with the presence of different Naegleria spp. 28.

357

Food source selectivity has previously been reported for Acanthamoeba spp. and V. vermiformis 48,

358

which could explain the brief co-colonisation observed between N. fowleri and N lovaniensis in the



359

laboratory biomonitors with E. coli as an abundant food source and could have reduced inter-amoebal

360

competition for food.

361

In conclusion, colonisation of DWDS sites appears to be by individual viable Naegleria spp., and not

362

multiple viable Naegleria spp., or FLA. Viable Naegleria spp. do appear to colonise the same DWDS

363

locations but at different times, which may imply the role of bacterial food sources for the

364

colonisation of individual Naegleria spp.. Additional work to explore the link between Naegleria spp.

365

and the microbial ecology in operational DWDSs needs to be conducted to further aid water utilities

366

in monitoring and controlling potential N. fowleri risks.

367 368

AUTHOR INFORMATION

369

Corresponding Author

370

*Telephone: +61 8 9333 6174. Email: [email protected].

371 372

SUPPORTING INFORMATION

373

Additional information including the melt curves of the Consensus primers of Naegleria spp.,

374

Tetramitus spp., Willaertia spp., Vahlkampfia spp. and Vermamoeba sp., generated on the Bio-Rad

375

iQ5 software, confirmed by sequencing. Depending on the species one to three unique peaks were

376

present on the melting curves presenting different characteristics. In addition, it also contains a box –

377

whisker plot of the biofilm cell counts of the six laboratory scale biomonitors biofilm samples

378

(cells/cm2) recorded using flow cytometry over the duration of the study (standard error of 5 %).

379 380

ACKNOWLEDGEMENTS

381

The continued access and installation of the biofilm monitors were achieved with the assistance of the

382

Water Corporation of Western Australia. The Water Corporation of Western Australia and CSIRO

383

Land and Water are acknowledged for the funding. Naomi Boxall and Ka Yu Cheng from CSIRO

384

Land and Water are thanked for valuable comments on the manuscript.

385 386

REFERENCES


Page 14 of 23

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387 388 389 390 391 392 393 394 395 396 397 398 399 400 401 402 403 404 405 406 407 408 409 410 411 412 413 414 415 416 417 418 419 420 421 422 423 424 425 426 427 428 429 430 431 432 433 434 435 436


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437 438 439 440 441 442 443 444 445 446 447 448 449 450 451 452 453 454 455 456 457 458 459 460 461 462 463 464 465 466 467 468 469 470 471 472 473 474 475 476 477 478 479 480 481 482 483 484 485 486

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36. Booton, G. C.; Carmichael, J. R.; Visvesvara, G. S.; Byers, T. J.; Fuerst, P. A., Identification of Balamuthia mandrillaris by PCR assay using the mitochondrial 16S rRNA gene as a target. Journal of Clinical Microbiology 2003, 41, (1), 453-455. 37. Schroeder, J. M.; Booton, G. C.; Hay, J.; Niszl, I. A.; Seal, D. V.; Markus, M. B.; Fuerst, P. A.; Byers, T. J., Use of subgenic 18S ribosomal DNA PCR and sequencing for genus and genotype identification of acanthamoebae from humans with keratitis and from sewage sludge. Journal of Clinical Microbiology 2001, 39, (5), 1903-1911. 38. Delafont, V.; Brouke, A.; Bouchon, D.; Moulin, L.; Hechard, Y., Microbiome of free-living amoebae isolated from drinking water. Water Research 2013, 47, (19), 6958-6965. 39. Declerck, P.; Behets, J.; van Hoef, V.; Ollevier, F., Detection of Legionella spp. and some of their amoeba hosts in floating biofilms from anthropogenic and natural aquatic environments. Water Research 2007, 41, (14), 3159-3167. 40. Muchesa, P.; Leifels, M.; Jurzik, L.; Hoorzook, K. B.; Barnard, T. G.; Bartie, C., Coexistence of free-living amoebae and bacteria in selected South African hospital water distribution systems. Journal of Parasitology Research 2017, 116, (1), 155-165. 41. Ovrutsky, A. R.; Chan, E. D.; Kartalija, M.; Bai, X.; Jackson, M.; Gibbs, S.; Falkinham, J. O., 3rd; Iseman, M. D.; Reynolds, P. R.; McDonnell, G.; Thomas, V., Cooccurrence of free-living amoebae and nontuberculous Mycobacteria in hospital water networks, and preferential growth of Mycobacterium avium in Acanthamoeba lenticulata. Applied and Environmental Microbiology 2013, 79, (10), 3185-92. 42. Dey, R.; Bodennec, J.; Mameri, M. O.; Pernin, P., Free-living freshwater amoebae differ in their susceptibility to the pathogenic bacterium Legionella pneumophila. FEMS microbiology letters 2009, 290, (1), 10-7. 43. Whiteman, L. Y.; Marciano-Cabral, F., Susceptibility of pathogenic and nonpathogenic Naegleria spp. to complement-mediated lysis. Infection and immunity 1987, 55, (10), 2442-7. 44. Wong, M. M.; Karr, S. L., Jr.; Chow, C. K., Changes in the virulence of Naegleria fowleri maintained in vitro. The Journal of parasitology 1977, 63, (5), 872-8. 45. Griffin, J. L., The Pathogenic Ameboflagellate Naegleria fowleri - Environmental Isolations, Competitors, Ecological Interactions, and the Flagellate-Empty Habitat Hypothesis. Journal of Protozoology 1983, 30, (2), 403-409. 46. Matin, A.; Jeong, S. R.; Faull, J.; Rivas, A. O.; Khan, N. A., Evaluation of prokaryotic and eukaryotic cells as food source for Balamuthia mandrillaris. Archives of Microbiology 2006, 186, (4), 261-271. 47. Neidig, N.; Jousset, A.; Nunes, F.; Bonkowski, M.; Paul, R. J.; Scheu, S., Interference between bacterial feeding nematodes and amoebae relies on innate and inducible mutual toxicity. Functional Ecology 2010, 24, (5), 1133-1138. 48. Pickup, Z. L.; Pickup, R.; Parry, J. D., Effects of bacterial prey species and their concentration on growth of the amoebae Acanthamoeba castellanii and Hartmannella vermiformis. Applied and Environmental Microbiology 2007, 73, (8), 2631-2634.

526 527



Page 18 of 23

Table 1 Viable amoebae detections from the six laboratory-scale biomonitors (BM – biomonitor, V – V. vermiformis, NF – N. fowleri, NL – N. lovaniensis).

Date

BM 1 BM 2 BM 3 BM 4 BM 5 Species Amoeba/cm2* Species Amoeba/cm2* Species Amoeba/cm2* Species Amoeba/cm2* Species Amoeba/cm2* 31/10/2013 V 8.25E-01 V 8.25E-01 NF 1.18E-01 9/12/2013 V 1.18E-01 V 1.18E-01 V 2.36E-01 V 3.54E-01 V 1.06E+00 6/01/2014 V 2.95E+00 V 2.36E-01 V 2.36E-01 10/02/2014 V 1.18E-01 1.18E-01 21/03/2014 V 2.36E-01 V 2.36E-01 4/04/2014 V 3.54E+00 V 9.43E-01 V 2.36E-01 1.18E-01 1/05/2014 2.36E-01 15/09/2014 NL 1.06E+00 NL 3.54E+00 NF, NL 3.54E+00 NL 2.24E+00 NF, NL 1.18E+00 25/11/2014 NL 1.18E-01 NF, NL 4.72E-01 NL 1.89E+00 NL 1.06E+00 NL 1.06E+00 14/01/2015 NL 8.25E-01 3.54E+00 NL 7.08E-01 NL 8.25E-01 4/02/2015 NL 1.18E-01 NL 2.36E-01 3.54E-01 5/02/2015 1.18E-01 3.54E-01 NL 1.18E-01 9/02/2015 3.54E+00 NL 2.36E-01 NL 2.36E-01 17/02/2015 1.18E-01 NL 1.18E-01 24/02/2015 NL 5.90E-01 NL 2.36E-01 4.72E-01 * Amoebae densities were calculated using number of growth plaques on NNA-E. coli plates per surface area of the biofilm growth substrates.


Species NF NF V NF, NL NF, NL NL NL

BM 6 Amoeba/c

2.95E+ 3.54E-0 2.36E-0

1.18E-0 1.18E-0 3.54E+ 3.54E+ 3.54E-0

2.36E-0

Page 19 of 23


2.5 2.0 1.5 1.0 0.0

0.5

Amoebae density in biofilm (amoeba/cm2)

3.0

3.5

Amoebae density

BM1

BM2

BM3

BM4

BM5

Biomonitors

Figure 1 Cell density of viable amoebae on NNA-E. coli plates from laboratory biomonitors.


BM6


Table 2 Range and mean of cell counts (cells/cm2) of the six laboratory scale biomonitors biofilm samples recorded using flow cytometry over the duration of the study (standard error of 5 %). Biomonitor BM1 BM2 BM3 BM4 BM5 BM6

Sample Biofilm Biofilm Biofilm Biofilm Biofilm Biofilm

Range 0 - 3.97 ×106 0 - 7.16 ×105 2.18 ×104 - 1.01 ×106 1.59 ×103 - 1.07 ×106 3.57 ×104 - 8.01 ×106 5.34 ×103 - 1.05 ×107

Mean 3.65 ×105 7.37 ×104 2.70 ×105 1.43 ×105 1.26 ×106 1.26 ×106


Page 20 of 23

Page 21 of 23


Table 3 Amoebae detections in bulk and biofilm samples by either culture or molecular detection methods at each site. a represents chlorinated DWDS, b represents a chloraminated DWDS, c represents a non-treated source water of metropolitan DWDS. (TA – unknown thermophilic amoeba). Site Sample

SKa Bulk Biofilm

KTa Bulk Biofilm

N. fowleri N. lovaniensis V. vermiformis TA N. andersoni N. dobsoni Multiple Negative No Samples Total viable detections

3 0 2 2 0 0 1 14 20 7

10 0 9 20 0 0 0 18 57 39

0 0 1 1 0 0 0 6 8 2

3 3 0 0 0 0 0 11 24 6

N. fowleri N. lovaniensis V. vermiformis TA N. andersoni N. dobsoni Multiple Negative No Samples Total molecular detections

5 0 5 0 1 2 3 10 20

26 0 21 0 4 2 6 10 57

1 1 0 0 1 0 1 6 8

12 8 0 0 3 0 2 3 24

13

53

3

23

WBa Bulk Biofilm Viable detections 0 4 0 0 0 4 0 4 0 0 0 0 0 0 21 51 21 63 0 12 Molecular detections 2 11 0 1 2 17 0 0 0 6 0 1 0 7 17 34 21 63 4

36


Bulk

Pb Biofilm

Bb Biofilm

BPc Bulk Biofilm

Bulk

0 0 1 0 0 0 0 3 4 1

0 5 5 0 0 0 0 0 10 10

0 0 0 1 0 0 0 4 5 1

0 9 2 0 0 0 0 0 11 11

0 3 1 0 0 0 0 1 5 4

0 9 0 0 0 0 0 6 15 9

0 0 0 0 0 0 0 4 4

0 7 5 0 0 0 4 2 10

0 0 1 0 0 0 0 4 5

0 10 2 0 0 0 1 0 11

0 4 1 0 0 0 1 1 5

0 11 3 0 0 0 3 0 15

0

12

1

12

5

14


Page 22 of 23

Table 4 Range of physical and chemical characteristics of the bulk water at each field site. Site SK KT WB P B BP

Temperature (°C) 13 – 41 13 – 33 9 – 34 13 – 31 10 – 32 21 – 43

Turbidity (NTU) 0.47 - 0.97 0.36 - 0.9 0.34 - 1.54 0.19 - 1.68 0.16 - 59 0.26 - 1.82

Disinfectant residual (mg/L) Free Chlorine Total Chlorine Chloramine 0 – 0.09 0.06 - 0.25 N/A 0 – 1.57 N/A N/A 0 – 0.55 0.05 - 0.77 N/A N/A N/A 0 – 0.07 N/A N/A 0 – 1.68 NIL NIL NIL

Table 5 Range and mean of cell counts in bulk water samples (cells/mL) and biofilm samples (cells/cm2) (standard error of 5 %) recorded using flow cytometry for the six field sites. Site SK KT WB P B BP

Sample Bulk water Biofilm Bulk water Biofilm Bulk water Biofilm Bulk water Biofilm Bulk water Biofilm Bulk water Biofilm

Range 9.26 ×104 - 1.19 ×107 5.28 ×104 - 2.93 ×106 9.96 ×105 - 2.60 ×107 8.06 ×104 - 1.66 ×105 2.50 ×103 - 2.87 ×106 5.46 ×104 - 7.71 ×105 1.15 ×105 - 2.36 ×106 1.50 ×106 - 2.41 ×107 3.89 ×105 - 1.29 ×106 4.93 ×105 - 2.34 ×106 1.14 ×106 - 4.37 ×107 3.96 ×105 - 1.86 ×106

Mean 1.75 ×106 5.66 ×105 4.97 ×105 2.81 ×105 4.97 ×105 2.81 ×105 1.36 ×106 7.76 ×106 6.13 ×105 1.63 ×106 2.10 ×107 1.07 ×106


NH3 N/A N/A N/A 0 - 0.02 0 - 0.42 NIL

Page 23 of 23



Competition between Naegleria fowleri ... - ACS Publications

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